1 Department of Biochemistry, Faculty of Science, Charles University, Albertov 2030,128 40 Prague 2, The Czech Republic,
2 Department of Molecular Toxicology, German Cancer Research Center, Im Neuenheimer Feld 280, 69120 Heidelberg, Germany
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Abstract |
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Abbreviations: AA, aristolochic acid; AAI, 8-methoxy-6-nitro-phenanthro-(3,4-d)-1,3-dioxolo-5-carboxylic acid; AAII, 6-nitro-phenanthro-(3,4-d)-1,3-dioxolo-5-carboxylic acid; Ah receptor, aryl hydrocarbon receptor; CT-DNA, calf thymus DNA; dAp, deoxyadenosine 3'-monophosphate; dGp, deoxyguanosine 3'-monophosphate; dA-AAI, 7-(deoxyadenosin-N6-yl)aristolactam I; dA-AAII, 7-(deoxyadenosin-N6-yl)aristolactam II; dG-AAI, 7-(deoxyguanosin-N2-yl) aristolactam I; dG-AAII, 7-(deoxyguanosin-N2-yl) aristolactam II; Ks, apparent dissociation constant; PB, phenobarbital; PEI, polyethylenimine; RAL, relative adduct labeling; XO, xanthine oxidase.
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Introduction |
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Interestingly, to date only 3% of the patients treated with the slimming regimen have suffered from nephropathy. Taking into account that AA is toxic, should it not have affected more of the patients? One possible explanation for the different responses of patients may be differences in the individual activities of the enzymes catalyzing biotransformation (detoxication and/or activation) of AA. Many genes of enzymes metabolizing carcinogens are known to exist in variant forms or show polymorphisms resulting in differing activities of the gene products. These genetic variations appear to be important determinants of cancer risk (16). Therefore, screening CHN patients as well as healthy persons treated with the slimming regimen for genetic variations in genes of the enzymes involved in AA metabolism should help to find possible relationships between genotypes and nephropathy, AA-DNA adduct levels and urothelial cancer risk. Thus, the identification of the enzymes principally involved in the activation of AA in humans and detailed knowledge of their catalytic specificities is of major importance.
Recently we found that in vitro, the human cytochromes P450 1A1 and 1A2, NADPH:P450 reductase, butter milk XO, and even peroxidases were competent in activating both AAI and AAII to form the same DNA adducts found in vivo in rodents (9,10,1719) and in humans (3,4,6). The role soluble cytosolic reductases, which are known to be crucial enzymes in the reduction of many nitroaromatics (2022), play in AA activation, has not been explored as yet. Therefore, the present study was undertaken to examine their capability to activate these carcinogens.
Materials and methods
Caution
Aristolochic acids are mutagenic and carcinogenic and should be handled with care. Exposure to 32P should be avoided, by working in a confined laboratory area, with protective clothing, plexiglass shielding, Geiger counters, and body dosimeters. Wastes must be discarded according to appropriate safety procedures.
Chemicals
Chemicals were obtained from the following sources: NADH, NADPH, nuclease P1, deoxyadenosine 3'-monophosphate (dAp), deoxyguanosine 3'-monophosphate (dGp), ellipticine, hypoxanthine, cytochrome c, sodium dodecyl sulfate (SDS), dicoumarol, allopurinol, and 2-hydroxypyrimidine from Sigma Chemical Co. (St Louis, MO), bicinchoninic acid from Pierce, (Rockford, IL), Sephadex G-150 from Pharmacia (Uppsala, Sweden), menadione from Merck (Darmstadt, Germany), Affi-Gel Blue (Cibacron Blue Agarose, Porcine Blue HB, C.T. 61211 Agarose) from Bio-Rad (Richmond, CA), Sudan I from The British Drug Houses (UK), N1-methylnicotinamide from Aldrich Chemical Co., (Milwaukee, WI), and calf thymus DNA (CT-DNA) from Roche Diagnostics (Mannheim, Germany). The natural mixture consisting of 65% AAI and 34% AAII was a gift from Madaus (Cologne, Germany). AAI and AAII were isolated from the mixture by preparative HPLC; their purity was 99.7% as estimated by HPLC (23). All other chemicals were of analytical purity or better. Enzymes and chemicals for the 32P-postlabeling assay were obtained commercially from sources described previously (3,17).
Animal experiments
Gavage with aristolochic acid I and a natural mixture of both acids (AA) dissolved in 0.15 mM NaCl (10 mg/kg body weight) was administered to six male Wistar rats (100150 g) once a day for four consecutive days. Ten male Wistar rats were injected i.p. with Sudan I in maize oil (20 mg Sudan I/kg body weight) once a day for three consecutive days or with ellipticine in maize oil/dimethyl sulfoxide (1:1, v/v) (40 mg ellipticine/kg body weight) once a day for two consecutive days. Pretreatment of rats with phenobarbital (PB) and ethanol or acetone was carried out by procedures described by Hodek et al. (25) and Yang et al. (26), respectively. Control rats obtained the solvents only. Rats were placed in cages in temperature- and humidity-controlled rooms. Standardized diet and water were provided ad libitum. Animals were killed 24 h after the last treatment by cervical dislocation (27). Liver and kidney of animals were excised immediately after sacrifice, quickly frozen in liquid nitrogen, and stored at 80°C until the isolation of cytosolic and microsomal fractions.
Preparation of cytosolic fractions
Liver and renal fractions (cytosol and microsomes) were prepared by differential centrifugation as described previously (27,28). The 105 000 g supernatant was taken as cytosol and used for studies presented in the paper. All tissue fractions were stored at 80°C. Each cytosolic preparation was analyzed for specific enzyme activities. The assays used were as follows.
DT-diaphorase, XO and aldehyde oxidase assays
DT-diaphorase activity was measured essentially as described by Ernster (29). The standard assay system contained 25 mM TrisHCl (pH 7.4), 0.2% Tween 20, 0.07% bovine serum albumin, 400 mM NADH (or NADPH) and 100 mM menadione (2-methyl-1,4-naphthoquinone) dissolved in methanol. The enzyme activity was determined by following the oxidation of NADH (NADPH) spectrophotometrically at 340 nm on a Hewlett-Packard 8453 diode array spectrophotometer. One unit of activity is defined as the amount of enzyme catalyzing the oxidation of 1 µmol of NADH (molar absorption coefficient =6.27 mM/cm). The activities of XO in cytosolic fractions was measured as described by Ichikawa et al. (30) using hypoxanthine as a substrate. The aldehyde oxidase activity was assayed as described by Felsted and coworkers (31) using N1-methylnicotinamide as a substrate. Protein concentration was assessed using the bicinchoninic acid protein assay with serum albumin as a standard (32).
Isolation of DT-diaphorase
DT-diaphorase was isolated as described earlier (33). Liver cytosol from Sudan I-treated rats was used. Briefly, proteins of cytosol (20 ml, 24 mg/ml) were fractionated with ammonium sulfate and the fraction of 3090% saturation containing most of the DT-diaphorase activity was dialyzed against 2000 ml of 150 mM KCl in 50 mM Tris (pH 7.4). The dialyzed enzyme preparation was chromatographed on a Sephadex G-150 column and DT-diaphorase was eluted with the same buffer. Pooled fractions containing the DT-diaphorase activity were applied onto a column of Affi-Gel Blue and non-DT-diaphorase proteins were eluted with the same buffer and subsequently with a gradient of NaCl (03.5 M) in this buffer. DT-diaphorase was eluted from Affi-Gel Blue with 20 mM Tris buffer (pH 10.0) containing 1 mM NADH. In order to remove residual protein impurities, the DT-diaphorase sample was applied onto a Sephadex G-150 column and re-chromatographed. The eluate was concentrated by ultrafiltration and stored at 80°C.
Incubations
The deaerated and argon-purged incubation mixtures contained in a final volume of 0.75 ml: 50 mM TrisHCl buffer (pH 7.4) containing 0.2% Tween 20, 1 mM cofactors of cytosolic enzymes (NADH or NADPH or hypoxanthine or 2-hydroxypyrimidine), 1 mg cytosolic protein, 0.5 mM AAI or AAII as sodium salts dissolved in water and 1 mg calf thymus DNA (4 mM dNp). The reaction was initiated by adding cofactor. Control incubations were carried out either without activating system (cytosol) or with activating system and AAI and AAII, but without DNA or with activating system and DNA but without AAs. Incubations with purified DT-diaphorase contained in a final volume of 0.75 ml: 50 mM TrisHCl buffer (pH 7.4) containing 0.2% Tween 20, 1 mM NADH or NADPH, 0.10.5 mM AAI or AAII as sodium salts dissolved in water, 1 mg of calf thymus DNA (4 mM) and 0.080.4 units of DT-diaphorase instead of cytosolic fractions. After incubation of all reaction mixtures (37°C, 60 min), the mixtures were extracted twice with ethyl acetate (2 x 2 ml). DNA was isolated from the residual water phase by the phenol/chloroform extraction method as described earlier (17,19,27,28). The content of DNA was determined spectrophotometrically (34).
Inhibition studies
The following chemicals were used to inhibit the activation of AAs in rat cytosolic fractions: dicoumarol, allopurinol and menadione for DT-diaphorase, XO and aldehyde oxidase inhibition, respectively (20,22). Inhibitors dissolved in 7.5 µl of methanol, to yield final concentrations of 110 µM, depending on the chemical, were added to the incubation mixtures. An equal volume of methanol alone was added to the control incubations. The reaction mixtures were incubated at 37°C for 60 min. Then mixtures were extracted by ethyl acetate and DNA was isolated by procedures described above.
32P-postlabeling analysis
The nuclease P1 enrichment version (35) and the 1-butanol extraction-mediated enrichment procedure (36) were used. DNA samples (12.5 µg) were digested with micrococcal nuclease (750 mU) and spleen phosphodiesterase (12.5 mU) in digestion buffer (20 mM sodium succinate, 8 mM CaCl2, pH 6.0) for 3 h at 37°C in a total volume of 12.5 µl. Digests (2.5 µl) were removed and diluted 1:1500 to determine the amount of normal nucleotides. In the nuclease P1 version digests (10 µl) were enriched for adducts by incubation with 5 µg (5 U) of nuclease P1 in 3 µl of a buffer containing 0.8 M sodium acetate (pH 5.0), 2 mM ZnCl2 for 30 min at 37°C. The reaction was stopped by adding 3 µl of 427 mM Tris base. The extraction with 1-butanol to enrich the adducts was carried out as described earlier (36). Labeling mix (4 µl) consisting of 400 mM bicine (pH 9.5), 300 mM dithiothreitol, 200 mM MgCl2, 10 mM spermidine, 100 µCi [-32P]ATP (15 pmol), 0.5 µl 90 µM ATP and 10 U T4 polynucleotide kinase was added. After incubation for 30 min at room temperature, 20 µl were applied to a polyethylenimine (PEI)-coated cellulose TLC plate (Macherey-Nagel, Düren, Germany) and chromatographed as described (37) except that D3 and D4 were adjusted to pH 4.0 and 9.1 for better resolution. To determine the amount of normal nucleotides 5 µl of the 1:1500 dilution of digests were mixed with 2.5 µl of Tris buffer (10 mM, pH 9.0) and 2.5 µl of labeling mix (see above) and incubated for 30 min at room temperature. The labeling mixture was diluted by mixing 4 µl with 750 µl of 10 mM Tris buffer (pH 9.0). This solution (5 µl) was applied to a PEI-cellulose TLC plate and run in 0.28 M (NH4)2SO4, 50 mM NaH2PO4 (pH 6.5). Adducts and normal nucleotides were detected and quantitated by an Instant imager (Packard). Count rates of adducted fractions were determined from triplicate maps after subtraction of count rates from adjacent blank areas. Excess [
-32P]ATP after the postlabeling reaction was confirmed. Adduct levels were calculated in units of relative adduct labeling (RAL) which is the ratio of c.p.m. of adducted nucleotides to c.p.m. of total nucleotides in the assay. Enzymatic synthesis of reference compounds, dAp-AAI, dGp-AAI, dAp-AAII and dGp-AAII and their 32P-postlabeling were carried out as described earlier (9).
Co-chromatography on PEI-cellulose
Adduct spots detected by the 32P-postlabeling assay were excised from the thin-layer plates, extracted and co-chromatographed with reference 3',5' bisphosphate adducts as reported previously (9).
HPLC analysis of 32P-labeled 3',5'-deoxyribonucleoside bisphosphate adducts
HPLC analysis was performed essentially as described previously (9,17,38). Individual spots detected by the 32P-postlabeling assay were excised from thin layer plates and extracted (39). The dried extracts were redissolved in 100 µl of methanol/phosphate buffer (pH 3.5) 1:1 (v/v). Aliquots (50 µl) were analysed on a phenyl-modified reversed-phase column (250 x 4.6 mm, 5 µm Zorbas Phenyl; Säulentechnik Dr Knauer, Berlin, Germany) with a linear gradient of methanol (from 40 to 80% in 45 min) in aqueous 0.5 M sodium phosphate and 0.5 M phosphoric acid (pH 3.5) at a flow rate of 0.9 ml/min. Radioactivity eluting from the column was measured by monitoring Cerenkov radiation with a Berthold LB 506 C-1 flow through radioactivity monitor (500 µl cell, dwell time 6 s).
Statistical analyses
Statistical association between DT-diaphorase- and XO-linked catalytic activities in hepatic cytosolic samples and levels of individual AA-DNA adducts formed by the same cytosolic samples were determined by the Spearman correlation coefficient using version 6.12 Statistical Analysis System sofware. Spearman correlation coefficients were based on a sample size of 8. All Ps are two-tailed and considered significant at the 0.05 level.
Molecular modeling
Crystallographic coordinates for rat DT-diaphorase with bound FAD were obtained from the Protein Data Bank (40). For modeling purposes, the physiological dimer form was used (MacroMolecular file 1qrd_1.mmol). The coordinates were used without further refinement.
The modeling of the binding of AAI to the active site was performed with the program Autodock 3.0.3. (41) and Sybyl 6.6.5 (Tripos GmbH, Germany). AAI was built via fragment libraries supplied with the modeling software. The initial structure was first energy minimized to a root-mean-square force of <0.001 with the consistent valence force field (42).
The energy-minimized AAI molecule was then used as the ligand input for the Autodock 3.0.3 program and positioned near FAD on the A chain of the DT-diaphorase dimer, and extensive interpolations were carried out with the lattice spacing of 0.25 Å. The apparent dissociation constants for AAI as well as its conformations and the positions of the interaction of AAI with DT-diaphorase were obtained as the Autodock 3.0.3 output.
Results
Activation of aristolochic acids by rat renal and hepatic cytosol
The detection of specific DNA adducts (3,4,6,9,10) by 32P-postlabeling analyses has allowed us to use AA-DNA binding as a probe for metabolic activation of AA in in vitro systems.
Initially two versions of the 32P-postlabeling assay (the nuclease P1 version and extraction with 1-butanol) were used to separate and quantify the adducts. No differences in patterns of adducts were found, but enrichment of AA-DNA adducts by n-butanol extraction yielded a lower recovery for these adducts (8090%) compared with the nuclease P1 version of the 32P-postlabeling assay (9). Therefore, all further experiments were carried out with enrichment by nuclease P1.
Rat hepatic and renal cytosolic fractions were capable of activating both AAI and AAII to form adducts with CT-DNA. It is evident from Figure 2 that AAI and AAII activated by cytosol from liver or kidney generated the same major DNA adduct spots as those obtained in vivo in rats and humans and as reported previously (3,4,6,9,37). Adduct spots 1, 2, 3 and 4 formed by AAs (Figure 2
) cochromatographed on PEI-cellulose TLC plates and by reversed-phase HPLC with those of synthetic standards (9) (not shown). Thus, spot 1 was assigned to 3',5'-bisphospho-7-(deoxyguanosin-N2-yl)-aristolactam I (dG-AAI); spot 2 to 3',5'-bisphospho-7-(deoxyadenosin-N6-yl)-aristolactam I (dA-AAI); spot 3 to 3',5'-bisphospho-7-(deoxyadenosin-N6-yl)-aristolactam II (dA-AAII); and spot 4 to 3',5'-bisphospho-7-(deoxyguanosin-N2-yl)-aristolactam II (dG-AAII). These adducts are known to be generated from AAs by nitro reduction (9,43,44). Therefore, the cytosolic fractions tested in this study contain enzymes, which are capable of catalyzing the reductive activation of AAs. XO, DT-diaphorase and aldehyde oxidase are major candidates for the reductive activation of AAs in cytosol. The XO activities in hepatic and renal cytosolic fractions from ten rats, each assayed in triplicate with hypoxanthine (30) are 40 ± 8 and 10 ± 3 nmol/min and mg protein, respectively. The DT-diaphorase activities in cytosolic fractions of both organs with menadione (45) as a substrate are 790 ± 80 and 82 ± 3 nmol/min and mg protein, respectively. In contrast, the activity of another reductase, aldehyde oxidase, measured with N1-methylnicotinamide as a substrate (31) was not detectable in either cytosol under the conditions used.
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As shown in Figure 3 the formation of AAI-DNA adducts was stimulated by NADPH, NADH and hypoxanthine, known cofactors of DT-diaphorase (46) and XO (47), respectively. Adduct levels increased about three times when NADPH, NADH or hypoxanthine were added into the incubation mixture. 2-Hydroxypyrimidine, an electron donor of the cytosolic aldehyde oxidase (48), had almost no effect neither had menadione an inhibitor of aldehyde oxidase (20). DT-diaphorase and XO are therefore responsible for reductive activation of AAI in cytosol. This assumption is also supported by the fact that AAI-DNA adduct formation was remarkably decreased by inhibitors of these two enzymes, namely by dicoumarol, an inhibitor of DT-diaphorase, and allopurinol of XO (Figure 3
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Involvement of DT-diaphorase in activation of aristolochic acids
Cytosol isolated from livers of uninduced rats and that from those of rats pretreated with compounds increasing the DT-diaphorase activity in hepatic cytosol (i.e. Sudan I, ellipticine, AAI and a natural mixture of AAI and AAII) (33) were used in the experiments. Unfortunately, cytosolic fractions, in which the XO activity is selectively induced could not be tested, because selective XO inducers are not known (49).
The DT-diaphorase and XO activities in the different cytosols are shown in Figure 4. Cytosols from rats pretreated with Sudan I (a 5-fold increase in total DNA-binding of AAI) or ellipticine were the most effective activators of AAI followed by those from rats treated with AAs, PB or ethanol. No increase in the AAI-DNA adduct formation with respect to control cytosol was determined for acetone as an inducer (Figure 4
). It is evident from results shown in Figure 4
that cytosolic fractions having higher DT-diaphorase activities exhibit an increased capacity to form AAI-DNA adducts. Indeed, the results were consistent with the correlation analysis. Total AAI-DNA adduct formation was highly correlated with DT-diaphorase activities in cytosol (r = 0.97, P = 0.01).
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To confirm the role of DT-diaphorase in the activation of AAs, the enzyme was purified and used in the experiments. Incubations of AAs with DNA, purified DT-diaphorase and its cofactors, NADPH (Figure 5) or NADH (not shown), resulted in the formation of the same pattern of DNA adducts as that determined in either cytosol (present paper) or in vivo (3,4,6,9). The identity of adducts were confirmed by cochromatography on TLC and HPLC (not shown). Control incubations carried out in parallel without the cofactor (NADH or NADPH) were free of adduct spots. An inhibitor of the enzyme, dicoumarol at 10 µM, which is 50-fold less than the AA substrate concentration, inhibited the AA-DNA adduct formation significantly, by 4050%. The DT-diaphorase-catalyzed DNA binding of AAs was shown to be dependent on the time of incubation, being linear up to 60 min (not shown) and on the concentrations of the enzyme and AAs (Figures 6 and 7
). DT-diaphorase-mediated AA-DNA adduct formation increased with increasing amounts of AAs and then reached a plateau level at higher AA concentrations. The concentrations required for half-maximal DNA binding were 0.17 mM for AAI and 0.23 mM for AAII.
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Recently, specific AA-DNA adducts were found to be associated with a unique nephropathy, Chinese herbs nephropathy, and urothelial cancer in women who had followed a weight reducing treatment consisting of Chinese herbs containing AA. Not all participants in the slimming procedure are affected by Chinese herbs nephropathy. Differences in carcinogen activation could be the reason for individual susceptibility.
In mammalian tissues, both cytosol and microsomes contain enzymes catalyzing the reduction of nitro aromatic compounds (2022,51). We have already identified human microsomal cytochromes P450 1A1 and 1A2 as well as NADPH: cytochrome P450 reductase as enzymes generating AAI- and AAII-DNA adduct profiles identical to those found in renal tissue in humans (19). The present paper reports on the identification of cytosolic enzymes, which participate in the reductive bioactivation of AA.
AAs are bioactivated by cytosol of both rat liver and kidney, the target organ of the AA-induced toxicity, to dG and dA adducts identical to those found in humans exposed to AA (3,4). The comparison of AAI-DNA adduct levels formed by microsomes (19) and cytosol (the present paper) reveals that the cytosolic enzymatic system is ~1.6 times more efficient in the activation of this carcinogen than microsomes. If adduct levels are based on the total cytosolic or microsomal protein in the organ, then renal cytosol yields 1.84 and renal microsomes 1.12 adducts in 105 normal nucleotides. One order of magnitude higher levels were calculated in the hepatic systems, namely, 49.5 and 34.1 adducts in 105 normal nucleotides per total cytosolic and microsomal protein, respectively. In cytosolic fractions, nitroreduction of AA leading to DNA adduct formation was predominantly catalyzed by DT-diaphorase, while XO seems to have lower, but measurable, capacity to activate AA. This conclusion is supported by strong correlation between the levels of AAI-DNA adducts formed in different cytosolic samples and DT-diaphorase activities. An inhibition of DNA adduct formation by dicoumarol, the inhibitor of DT-diaphorase, provided additional evidence for the role of this enzyme in AA activation. Experiments with pure DT-diaphorase confirmed these results. Computer modeling where the AAI molecule was docked to the DT-diaphorase active site suggests that AAI is situated where the other DT-diaphorase substrates (i.e. duroquinone, ref. 52) are found in the X-ray structure, with the planar aromatic AAI rings parallel to the flavin ring. This allows for an efficient electron transfer during the reductive activation. The value of the calculated dissociation constant for the DT-diaphorase-AAI complex (16.4 µM) corresponds well to the apparent Km value determined experimentally (27.7 µM) (33).
Levels of expression and activities of DT-diaphorase in humans differ considerably among individuals, because the enzyme is influenced by several factors (smoking, drugs, environmental chemicals and genetic polymorphisms) (53,54). Rat DT-diaphorase is coinduced with aromatic hydrocarbon hydroxylase and glutathione-S-transferases by pretreatment with inducers, which activate the Ah-locus, such as 3-methylcholanthrene or 2,3,7,8-tetrachlorodibenzo[1,4]dioxin (55,56). 1,1'-Azonaphthalenes, Sudan I, Sudan III, flavonoids, Aroclor-1254, coumarins, and the antioxidants butylated hydroquinone and butylated hydroxyanisole are additional compounds efficiently inducing the enzyme (56). The precise mechanism of induction of DT-diaphorase has not been established (5557); however, the existence of multiple genes in rat and human liver suggests the presence of multiple forms of the enzyme (5658). One gene appeared to be closely linked to a gene regulating the expression of cytochromes P450 1A1 and 1A2, enzymes which also activate AAs (19), another gene is located on another chromosome. Indeed, the antiestrogens tamoxifen and hydroxytamoxifen stimulate the expression of DT-diaphorase by activating the estrogen receptor, which is different from the Ah locus (59). Because in rats DT-diaphorase activity is increased by aristolochic acids (33), DT-diaphorase in the patients who ingested AA might also be induced. Collectively, all these data suggest that variations of this enzyme and regulatory proteins controlling its expression (Ah receptor, or its associated transcription factor, the Ah receptor nuclear translocator or Arnt protein, 16), might play a role in the risk for cancer by aristolochic acids. To test this hypothesis, CHN patients and other participants in the slimming regimen will be screened for polymorphisms of the genes encoding DT-diaphorase, cytochromes P450 1A and the Ah receptor in the next phase of our work.
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