Ochratoxin A Causes DNA Damage and Cytogenetic Effects but No

Brown Cancer Center, University of Louisville, Louisville, Kentucky ... investigated in male rats, the most sensitive species for OTA-induced tumor fo...
0 downloads 0 Views 393KB Size
Download PDF
Chem. Res. Toxicol. 2005, 18, 1253-1261

1253

Ochratoxin A Causes DNA Damage and Cytogenetic Effects but No DNA Adducts in Rats Angela Mally,† Gaetano Pepe,‡ Srivani Ravoori,§ Mario Fiore,‡ Ramesh C. Gupta,§ Wolfgang Dekant,†,* and Pasquale Mosesso‡ Department of Toxicology, University of Wu¨ rzburg, Germany, Dipartimento di Agrobiologia e Agrochimica, Universita` degli Studi della Tuscia, Viterbo, Italy, and Pharmacology & Toxicology/ Brown Cancer Center, University of Louisville, Louisville, Kentucky Received December 16, 2004

Ochratoxin A (OTA) is a potent nephrotoxin and renal carcinogen in rats, but the mechanism of OTA tumorigenicity is unknown. Ochratoxin A has been shown to be negative in many genetic toxicology test in vitro. However, the potential of OTA to induce genotoxic effects has not been investigated in male rats, the most sensitive species for OTA-induced tumor formation. In this study, male F344 rats were repeatedly administered OTA (0, 250, 500, 1000, and 2000 µg/kg of body wt) or the non-chlorinated analogue ochratoxin B (OTB; 2000 µg/kg of body wt) for 2 weeks (5 days/week), and DNA breakage was analyzed in target and nontarget tissues using the comet assay both in the absence and presence of formamidopyrimidine-DNA (Fpg) glycosylase. Potential DNA-adduct formation was also analyzed in the target organ kidney by 32P-postlabeling using two different solvent systems. DNA-strand breaks were evident in liver, kidney, and spleen of animals treated with OTA, and a similar degree of DNA damage was observed in rats treated with OTB, despite the lower toxicity of OTB. Moreover, the presence of DNA damage did not correlate with histopathological alterations, which were evident in the kidney but not in the liver. In liver and kidney, the extent of DNA damage was further enhanced in the presence of Fpg glycosylase, which is known to convert oxidative DNA damage into strand breaks, suggesting the presence of oxidative DNA damage. Oxidative DNA damage as a mechanism of OTA-dependent DNA damage is consistent with the absence of lipophilic DNA adducts as assessed by 32P-postlabeling analysis. No spots indicative of OTA-related DNA adducts were observed in kidney DNA extracted from OTA-treated animals by 32P-postlabeling analysis, despite the use of synthetic standard for postulated adducts. A small, but not significant, increase in the incidence of chromosomal aberrations (essentially chromatid and chromosome-type deletions) was observed in splenocytes from rats treated with OTA in vivo and subsequently cultured in vitro to express chromosomal damage. These aberrations are also compatible with oxidative DNA lesions since they are not typically caused by chemical carcinogens which form covalent DNA adducts. Together, with the lack of evidence for formation of lipophilic DNA adducts as assessed by postlabeling, these data suggest that OTA may cause genetic damage in both target and nontarget tissues independent of direct covalent binding to DNA.

Introduction

Chart 1

The mycotoxin and food contaminant ochratoxin A (OTA,1 Chart 1) is one of the most potent renal carcinogens studied by the National Cancer Institute/National Toxicology Program (NCI/NTP) to date. Little is known regarding the mechanism of tumor formation by OTA, and the genotoxic activity of OTA has been assessed in a variety of standard genetic toxicology tests. In general, only weak genotoxic effects have been observed, and the contribution of genotoxicity to renal tumor formation by * Author for correspondence: Prof. Dr. Wolfgang Dekant, Department of Toxicology, University of Wu¨rzburg, Versbacher Str. 9, 97078 Wu¨rzburg, Germany. Tel: +49-931-20148449. Fax: +49-931-20148865. E-mail: [email protected]. † University of Wu ¨ rzburg. ‡ Universita ` degli Studi della Tuscia. § University of Louisville. 1 Abbreviations: OTA, ochratoxin A; OTB, ochratoxin B; Fpg, formamidopyrimidine-DNA glycosylase; FCS, fetal calf serum; 8-OHdG, 8-oxo-7,8-dihydro-2′-deoxyguanosine; dGMP, 2′-deoxyguanosine3′-monophosphate.

OTA remains unknown. Numerous studies have shown that OTA is not mutagenic in Salmonella typhimurium both in the presence and absence of metabolic activation

10.1021/tx049650x CCC: $30.25 © 2005 American Chemical Society Published on Web 07/08/2005

1254

Chem. Res. Toxicol., Vol. 18, No. 8, 2005

systems (1-7), although mutations in bacteria after exposure to OTA have been reported in single studies using modifications of the Ames test and in the presence of mouse but not rat kidney fractions, despite the fact that rats are much more sensitive to OTA carcinogenicity than mice (1, 8, 9 ). OTA was also negative in the Escherichia coli SOS-spot test at nontoxic concentrations (10-12). In mammalian cells, conflicting results have been obtained regarding the potential of OTA to induce mutations, and positive effects were also seen in the absence of metabolic activation. OTA was not mutagenic in TK+/- mouse lymphoma cells or in the HPRT test system. In contrast, mutagenic effects of OTA have been reported in NIH 3T3 mouse fibroblasts stably transfected with human cytochrome P450s (13), although these results are difficult to interpret since studies on OTA metabolism do not suggest a role of bioactivation in OTA toxicity (14-17). In Chinese hamster ovary cells (CHO), exposure to OTA did not result in chromosomal aberrations, but a small increase in the frequency of sister chromatid exchange (SCE) was noted (1). “Unscheduled DNA synthesis” has been observed in several cell types in response to OTA treatment (18-20), and OTA has been shown to induce DNA-strand breaks independent of metabolic activation in vitro, although some of these studies used very high concentrations of OTA and it is possible that the observed DNA damage may have been a result of cytotoxicity (7, 21-24). In vivo, DNA-strand breaks have been observed in spleen, liver, and kidney of mice treated with a single dose of OTA using the alkaline elution technique (24), but little information exists regarding the genotoxic effects of OTA in the rat, the most sensitive species for OTA carcinogenicity. The aim of this study was to investigate the genotoxicity of OTA in a susceptible animal model using a dosing regimen which causes significant histological and functional alterations characteristic for OTA. Single-cell gel electrophoresis (comet assay) was used to detect DNAstrand breaks in target and nontarget tissues obtained from rats after repeated administration of OTA (0-2000 µg/kg of body wt) for 2 weeks, and the frequency of chromosomal aberrations was analyzed in splenocytes isolated from treated animals and cultured in vitro to express chromosomal damage. In addition, the genotoxic potential of ochratoxin B (OTB, Chart 1), the nonchlorinated and less toxic analogue of OTA, was also investigated. Since formation of DNA adducts by OTA is still under debate, DNA-adduct formation was also assessed in kidneys of treated animals using 32P-postlabeling and synthetic OTA-3′-dGMP adduct standard. Putative DNA adducts have previously been reported and ascribed to direct binding of OTA using this method. In contrast, DNA-binding studies using radiolabeled OTA consistently were unable to detect DNA binding of OTA with sensitivities below those of the postlabeling assays (14, 15, 17, 25).

Material and Methods Chemicals. OTA and OTB were obtained from SigmaAldrich, Taufkirchen, Germany (Lot-No. 38H4120) or purchased from Prof. Peter Mantle, Imperial College of Sciences, London, U.K. The purity of OTA and OTB was >99.9%, as assessed by HPLC with UV and fluorescence detection. Aristolochic acid (I) sodium salt was purchased from Sigma-Aldrich, Taufkirchen, Germany (Lot.-No. 073K1607). Fetal calf serum (FCS) and RPMI medium was purchased from (Gibco BRL). Other chemi-

Mally et al. cals were obtained from Sigma-Aldrich, Taufkirchen, Germany, in the highest purity available. Animal Treatment. Male F344 Fisher rats (8-9 weeks) were purchased from Harlan-Winkelmann, Borchen, Germany. Animals were housed in macrolon cages and allowed free access to standard laboratory chow (Altromin) and tap water. Room temperatures were maintained at 21 ( 2 °C with a relative humidity of 55 ( 10% and a day/night cycle of 12 h. Following a week of acclimatization, rats (3/group) were treated with OTA (250, 500, 1000, and 2000 µg/kg of body wt) or OTB (2000 µg/kg of body wt) in corn oil by oral gavage for 2 weeks (5 days per week). Control rats received equal volumes of corn oil. Rats were sacrificed by CO2 asphyxiation and cervical dislocation 72 h after the last dose. The time-point for sacrifice was selected based on the slow elimination of OTA (t1/2 in rats ∼230 h) and previous studies which reported high DNA-adduct levels between 48 and 72 h after administration of a single dose of OTA (26, 27). Livers, kidneys, and spleens were removed; aliquots of tissues were placed on ice and immediately used to prepare cell suspensions for the comet assay. The remaining tissues were aliquoted, flashfrozen in liquid nitrogen, and stored at -80 °C until further analysis. To obtain bone marrow cells, the femurs were removed and cleaned of surrounding tissue. The bone was cut at the proximal end and irrigated with FCS using a 1 mL syringe. Blood samples were obtained by cardiac puncture. In a separate experiment, two female Sprague Dawley rats (300-310 g) were administered aristolochic acid (10 mg/kg of body wt dissolved in a volume of 2 mL of H2O) by oral gavage on 5 consecutive days. Animals were sacrificed 4 h after administration of the final dose by CO2 asphyxiation and cervical dislocation; kidneys were excised, flash-frozen in liquid nitrogen, and stored at -80 °C until further analysis. These samples served as positive controls for 32P-postlabeling analysis. Preparation of Metaphases for Cytogenetic Analysis from Splenocytes. Approximately one-third of the organ was placed in RPMI 1640 medium supplemented with 20% FCS. Spleens were squeezed through a 10 mL syringe without the needle to obtain cell homogenates which were then filtered through sterile cell strainers (Falcon) with a pore size of 100 µm to obtain cell suspensions in 5 mL RPMI 1640 medium. The cell suspension obtained was carefully layered over Histopaque1077 (Sigma Chemical, St. Louis, MO) and centrifuged at 400g for 25 min at room temperature. Splenocytes were removed from the gradient with a sterile pipet, washed twice with PBS, and counted using a haemocytometer. Cells were seeded at 6 × 106 cells per T-75 culture flask in RPMI medium supplemented with 20% FCS (heat inactivated), 200 mM l-glutamine, and 20 mM Hepes. Concanavalin A (Sigma) was added to a final concentration of 2 µg/mL culture medium. Cultures were grown for 54 h at 37 °C. Colcemid (Gibco BRL) was added 3 h before harvesting at a final concentration of 0.2 µg/mL. At the end of treatment, cells were brought into suspension with trypsin. The cell suspension was centrifuged and resuspended in hypotonic solution (KCl, 0.075 M), fixed in freshly prepared methanolacetic acid (3:1 v/v), and washed three times. Air-dried slides were prepared from the cell suspension and stained in 3% Giemsa. A blind scoring of metaphases was performed using coded slides. Where possible, a minimum of 100 metaphases were scored per animal. Chromosomal aberrations were classified as chromatid-type gaps, chromatid-type breaks, chromatid-type exchanges, chromosome-type gaps, chromosome-type breaks, chromosome-type exchanges, and isolocus events, which include isochromatid and isolocus breaks when these cannot be distinguished, as described by Savage (28). Preparation of Cell Suspensions from Organs for the Comet Assay. Tissues were dissected and processed immediately after harvesting. For preparation of cell suspensions, tissues (one-third of liver lobe, two-thirds of spleen, one-third of kidney) were rinsed with cold PBS and placed into a Petri dish containing 3 mL of PBS at +4 °C. Tissues were minced with scissors, and cell homogenates were filtered through sterile

DNA Damage and Cytogenetic Effects of OTA cell strainers (Falcon) with a pore-size of 100 µm to obtain single-cell suspensions. Cell suspensions were then centrifuged at 500 rpm for 10 min at 4 °C and finally re-suspended in 800 µL (liver, spleen) or 1000 µL (kidney) cold (+4 °C) PBS. Bone marrow cells were prepared by repeatedly aspirating the extracted bone marrow. The cell suspension was then centrifuged at 1000 rpm for 10 min at +4 °C and re-suspended in 8 mL cold PBS. An aliquot of heparinized whole blood (10 µL) was mixed directly with 190 µL of low-melting agarose (LMA) kept at 37 °C to prepare slides for the comet assay. Single-Cell Gel Electrophoresis. Comet slides were prepared using the protocol for the alkaline “comet assay” of Singh et al. (29). Briefly, 10 µL of cell suspension was mixed with 65 µL of 0.7% (w/v) low-melting-point agarose (Bio-Rad Lab.) and sandwiched between a lower layer of 1% (w/v) normal-meltingpoint agarose (Bio-Rad Lab.) and an upper layer of 0.7% (w/v) low-melting-point agarose on microscope slides (Carlo Erba). Duplicate slides were prepared from each individual treatment. The slides were then immersed in a lysing solution (2.5 M NaCl, 100 mM Na2EDTA, and 10 mM Tris, pH 10) containing 10% DMSO and 1% Triton X100 (ICN Biomedicals, Inc.) overnight at 4 °C. Upon completion of lysis, slides were placed in a horizontal gel electrophoresis tank with fresh alkaline electrophoresis buffer (300 mM NaOH and 1 mM Na2EDTA, pH g 13) and left in the solution for 25 min at 4 °C to allow the DNA to unwind and to express the alkali-labile sites. Electrophoresis was carried out at 4 °C for 25 min, 30 V (1 V/cm), and 300 mA, using a Bio-Rad power supply. After electrophoresis, the slides were immersed in 0.3 M sodium acetate in ethanol for 30 min. Microgels were then dehydrated in absolute ethanol for 2 h and immersed for 5 min in 70% ethanol. Slides were air-dried at room temperature. Immediately before scoring, slides were stained with 12 µg/mL ethidium bromide (Boehringer Mannheim, Germany) and examined at 400× magnification with an automatic image analyzer (Comet Assay III; Perceptive Instruments, U.K.) connected to a fluorescence microscope (Eclipse E400; Nikon). To evaluate the amount of DNA damage, computergenerated tail moment values and percentage of migrated DNA were used. For each individual treatment, a total of 100 cells were scored from two different slides. Formamidopyrimidine-DNA Glycosylase (Fpg) Digestion. To obtain information on the presence of oxidized DNA bases in the different organs of treated animals, a digestion using E. coli formamidopyrimidine-DNA glycosylase was included in the protocol for the comet assay to express lesions as DNA single-strand breaks. Besides its high sensitivity to detect 8-oxo-7,8-dihydro-2′-deoxyguanosine (8-OH-dG), 2,6-diamino-4hydroxy-5-formamidopyrimidine (FapyGua), and 4,6-diamino5-formamidopyrimidine (FapyAde), Fpg is also known to recognize abasic sites (AP sites) and ring-opened N-7 guanine adducts (30-33). Following cell lysis, slides were washed 3 × 5 min in enzyme buffer (40 mM Hepes, 100 mM KCl, 0.5 mM EDTA, and bovine serum albumin 0.2 mg/mL at pH 8.0), covered with 100 µL of Fpg solution (0.5 µg/mL Fpg in 10 mM Tris, pH 7.5, 1 mM EDTA pH 8.0, and 50 mM NaCl), sealed with a coverslip, and incubated for 30 min at 37 °C in a moist chamber. For DNA unwinding and electrophoresis, slides were further processed as described above. Flow-Cytometry Analysis of DNA Content. For flowcytometric evaluation of ploidy status, medullary and cortical regions were carefully dissected and kidney cells were prepared as described above (preparation of cell suspensions from organs for comet assay). Cells were fixed in ice-cold (-20 °C) PBS/ methanol (1:1 v/v) and stained with propidium iodide (20 µg/ mL) immediately before analysis. A FACStar Plus flow cytometer (Becton Dickinson) equipped with a 5W Innova 90 Coherent laser with 488-nm wavelength excitation light was used. DNA histograms were analyzed using WinMDI 2.5 software. A minimum of 5000 events were analyzed for each sample. Data obtained were plotted as percentage of cells with DNA content less than 2n (2n).

Chem. Res. Toxicol., Vol. 18, No. 8, 2005 1255 Preparation of 3′-dGMP-OTA Adduct Standards by Photoirradiation. 3′-dGMP-OTA adducts were prepared by photochemical reaction as described by us and others (17, 34). Briefly, a solution of OTA (500 µM) and 3′-dGMP (20 mM) in a mixture (3:1) of potassium phosphate buffer (0.1 M, pH 7.4) and DMSO was irradiated for 8 min with a mercury high-pressure lamp (Heraeus TQ 150) through a cutoff filter (Schott WG 305, l > 305 nm) under aerobic conditions. Formation of the previously described C-C8 and O-C8 adducts of OTA-3′-dGMP was confirmed by LC/MS/MS. LC/MS/MS analysis was performed on a Agilent 1100 series LC coupled to an API 3000 triple quadrupole mass spectrometer (Applied Biosystems, Darmstadt, Germany) (17). The API 3000 mass spectrometer was operated with a Turbo Ion Spray source in the negative ion mode with a voltage of -4000 V. Spectral data were recorded in the multiple reaction monitoring mode (MRM). On the basis of the ESspectra of the C-C8 and O-C8 adducts of OTA-3′-dGMP (34), the following m/z transitions representing loss of 3′-monophosphate-deoxyribose were analyzed: m/z 713-517 (C-C8-adduct of OTA-3′-dGMP) and 747-551 (O-C8-adduct of OTA-3′dGMP). Analysis of DNA Adducts by 32P-Postlabeling. Tissue samples from kidneys of OTA- and aristolochic acid-treated animals were collected at terminal sacrifice, immediately frozen in liquid nitrogen, and shipped on dry ice for postlabeling analysis. A blinded postlabeling analysis was performed with respect to sample status. DNA was isolated from kidneys of treated animals by a solvent extraction procedure described in detail previously (35). Briefly, crude nuclei were isolated from frozen tissues prior to digestion with RNAses and proteinase K, followed by sequential extraction with phenol, phenol/Sevag (chloroform/isoamyl alcohol, 24:1), and Sevag. The DNA was recovered by precipitation with ethanol in the presence of sodium chloride. The conditions for DNA digestion, nuclease P1 enrichment, and 32P-postlabeling have been described in detail elsewhere (35). Briefly, aliquots of DNA (10 µg) were digested at 37 °C for 5 h with micrococcal nuclease (375 mU; Sigma) and spleen phosphodiesterase (10 mU; Calbiochem) in a reaction mixture containing 20 mM sodium succinate and 10 mM calcium chloride, pH 6.0, and subsequently treated with nuclease P1 (8 U) for 45 min at 37 °C. DNA digests were then labeled with 80 µCi [γ-32P]ATP using T4 polynucleotide kinase (Amersham) in a total volume of 15 µL and resolved on polyethyleneimine (PEI)-cellulose by multidimensional TLC, using two different conditions for separation of potential DNA adducts. Solvent system A (LFU/SPU) was identical to that previously used to analyze OTA-derived DNA adducts (26, 27, 34) and consisted of the following: D1 opposite D3, 2.3M sodium phosphate, pH 5.8, onto a 5 cm Whatman 17 wick; D3, 4.8 M lithium formate/ 7.8 M urea, pH 3.5; D4 perpendicular to D3, 0.6 M sodium phosphate/5.95 M urea, pH 6.4; D5 in D4, 1.7 M sodium phosphate, pH 6.0, onto a 5 cm Whatman 1 wick. Solvent system B (LFU/LTU) consisted of the following: D1 opposite D3, 2.3 M sodium phosphate, pH 5.8, onto a 5 cm Whatman 17 wick; D3, 4 M lithium formate/7 M urea, pH 3.5; D4 perpendicular to D3, 0.8 M lithium chloride/0.5 M Tris HCl/7 M urea, pH 8.0; D5 in D4, 1.7 M sodium phosphate, pH 6.0, onto a 5 cm Whatman 1 wick. Adducts were visualized by intensifying screen-enhanced autoradiography, and total radioactivity was determined using the Packard Instant Imager. All samples were digested, labeled, and separated in duplicate. Statistical Analyses. Statistical analysis was performed using ANOVA followed by Dunnett’s test. For chromosomal aberration analyses in splenocytes, a modified chi-squared statistical method was employed to compare treated and control groups. The degree of heterogeneity within each group was first calculated, and where this was significant, it was taken into account in the comparisons between groups. Variance ratios or chi-squared values are taken to show the significance of any difference between each treatment group and the control.

1256

Chem. Res. Toxicol., Vol. 18, No. 8, 2005

Figure 1. DNA damage observed in liver (a), kidney (b), and spleen (c) of rats after repeated administration of OTA (0, 250, 500, 1000, and 2000 µg/kg of body wt) and OTB (2000 µg/kg of body wt) for 2 weeks (5 d/week) using the alkaline single-cell gel electrophorhesis. Data are presented as mean tail moment obtained from scoring a total of 100 cells per individual animal (n ) 3/dose group). Statistical analysis was performed by ANOVA followed by Dunnett’s test. Significant changes compared to controls are indicated as *, p < 0.05; **, p < 0.01; and ***, p < 0.001.

Results A small but significant increase in DNA breakage was observed in the kidney, liver, and spleen of OTA-treated animals using the alkaline comet assay (Figure 1). In both liver and spleen, this effect was dose-dependent. In contrast, the extent of DNA damage in the kidney was more pronounced in the 250 and 500 µg/kg dose groups, which may indicate that, at higher doses, cells may have been lost due to toxicity by apoptosis, resulting in lower numbers of cells with DNA damage observed in the comet assay. In support of this, apoptotic cells (either degenerating while attached to the basement membrane or detached into the tubule lumen) were frequently observed in kidneys of rats treated with the higher doses of OTA

Mally et al.

along with a high incidence of abnormally enlarged nuclei and mitotic cells, while no changes were evident in livers (36). Interestingly, treatment with OTB (2000 µg/kg of body wt), which is considered as nonmutagenic and less toxic than OTA and which did not induce histopathological changes (36), also resulted in an increase in mean tail moment in kidney, liver, and spleen (Figure 1). In bone marrow, a significant increase in DNA fragmentation was only observed in animals treated with 500 µg/ kg of body wt of OTA (data not shown). Only minor changes were seen in peripheral lymphocytes, and these were not considered to be biologically relevant (data no shown). To further characterize the OTA-induced DNA damage, single-cell gel electrophoresis was also performed in the presence of formamidopyrimidine-glycosylase (FGP protein), which converts oxidative DNA base modifications, particularly 8-oxo-7,8-dihydro-2′-deoxyguanosine (8-OHdG), to strand breaks. Under these conditions, DNA damage was significantly increased in hepatocytes and peripheral lymphocytes isolated from both OTA- and OTB-treated animals (Figure 2a,c). In the kidney, the target organ of OTA toxicity and carcinogenicity, a significant increase over control was evident in high-dose animals only (Figure 2b). However, a much higher level of oxidative DNA damage was already evident in kidneys from control animals as compared to other organs/tissues. In both spleen and bone marrow, the extent of DNA damage was not enhanced in the presence of Fpg glycosylase (data not shown). In general, heavily damaged cells resembling apoptotic cells were not observed by single-cell gel electrophoresis. Representative images of DNA breakage observed in controls and OTA-treated animals are shown in Figure 3. Chromosomal aberrations were investigated in splenocytes isolated from animals treated with OTA and OTB in vivo and subsequently cultured in vitro to express cytogenetic damage. Although the spleen is not the target for OTA carcinogenicity, lymphocytes isolated from spleen present a powerful and sensitive tool to study cytogenetic effects of compounds, as splenocytes rest in the G0 phase of the cell cycle and are therefore exposed to the test agent or its metabolites for the whole length of the treatment without selection as in the case of proliferating cells (e.g., kidney cells), which can lead to underestimation of genetic damage. Slight increases in chromosomal aberrations were observed at the 250, 1000, and 2000 µg/kg of body wt dose levels, although this was not doserelated and did not reach statistical significance due to the heterogeneity of group responses (Table 1). Chromosomal damage essentially consisted of chromatid deletions (Figure 4), which are not representative of chemically induced direct DNA damage but are also compatible with oxidative damage. Exchange-type aberrations, which are indicative of bulky DNA adducts as could be expected for OTA, were not observed. Mitotic indices of cultured splenocytes were not significantly affected at any of the dose levels employed. Consistent with the histopathological observations (36), flow-cytometric analysis of kidney cells revealed a dosedependent increase in the incidence of hyperdiploid cells (>4n DNA content) in the renal medulla, but not in cortical regions of kidneys obtained from OTA-treated animals (Figure 5). Previous analysis of potential DNA adducts of the same samples using both 32P-postlabeling and LC/MS/MS

DNA Damage and Cytogenetic Effects of OTA

Chem. Res. Toxicol., Vol. 18, No. 8, 2005 1257

Figure 3. Representative comet image showing undamaged DNA (control kidney (a)) and kidney DNA obtained from OTAtreated rats (2 mg/kg of body wt) in the absence (b) and presence (c) of Fpg glycosylase. Figure 2. Dose-dependent increase in mean tail moment in the presence of Fpg glycosylase indicative of oxidative DNA damage in liver (a), kidney (b), and peripheral lymphocytes (c) of rats treated with OTA (0, 250, 500, 1000 and 2000 µg/kg of body wt) and OTB for 2 weeks (5 d/week). Statistical analysis was performed by ANOVA followed by Dunnett’s test. Significant changes compared to controls are indicated as *, p < 0.05; **, p < 0.01; and ***, p < 0.001.

analysis (method development based on the synthetic carbon-bonded OTA-C8-deoxyguanosine) did not result in “spots” which coeluted with the synthetic standard in multidimensional TLC or in a signal when monitoring specific transitions for OTA-C8-deoxyguanosine by LC/ MS/MS (17). Therefore, these data did not support the presence of DNA adducts from OTA in kidney of OTAtreated rats or in DNA reacted with OTA under conditions reported to induce DNA adducts in vitro (37, 38). To reassess possible DNA binding by postlabeling, kidney DNA samples from rats treated with OTA for 2 weeks were sent to a laboratory with a longstanding experience in the application of the postlabeling technique to detect DNA adducts formed from a number of different chemicals. Analysis was performed blinded to sample status using two different solvent systems for separation of lipophilic adducts, and a mixture of OTA-3′-dGMP

adducts formed by photoirradiation of OTA in the presence of 3′-dGMP as adduct standard. In agreement with our previous negative results, lipophilic DNA adducts related to OTA treatment were not observed in this analysis (Figure 6). Several background spots, most likely due to endogenous “I compounds”, were detected in DNA from both control and OTA-treated animals. However, these adducts were not increased in response to OTA treatment. In contrast, analysis of kidney DNA extracted from rats treated orally with five daily doses of aristolochic acid (10 mg/kg of body wt) produced adduct maps containing several adducts.

Discussion In this study, the potential of OTA and its nonchlorinated analogue OTB to induce DNA damage has been investigated in target and nontarget tissues of male F344 rats. This rat strain is highly susceptible to renal tumor formation by OTA (1). The obtained results show that both OTA and OTB cause DNA damage in rats in both target and nontarget organs for tumor induction. Single-cell gel electrophoresis is a sensitive tool to investigate chemically induced DNA damage in vitro and

1258

Chem. Res. Toxicol., Vol. 18, No. 8, 2005

Mally et al.

Table 1. Frequencies of Chromosomal Aberrations in Splenocytes from Rats Treated in Vivo with OTA (250, 500, 1000, and 2000 µg/kg of body wt) or OTB (2000 µg/kg of body wt) in Corn Oil by Oral Gavage for 2 Weeks (5 Days per Week) and Then Cultured in Vitro to Express Chromosomal Damagea dose (µg/kg of body wt) control ochratoxin A 250 1000 2000 ochratoxin B 2000

no. of animals

no. of cells scored

3

300

8

1

0

2 2 3

200 200 300

14 17 26

0 0 0

2

143

5

0

chromatid del. exch.

total no. of aberrations

aberrant cells (%)

0

9

3.0

1 1 1

1 1 1

16 18 28

8.0 9.0 9.0

no no no

5.2 3.5 2.4

1

1

6

4.2

no

1.2

chromosome del. exch.

statistical significance

mitotic index (%) 5.2

a

Analysis of chromosomal aberrations in splenocytes from animals treated with 500 µg/kg of body wt OTA was not possible due to low cell numbers.

Figure 4. Metaphases of splenocytes isolated from rats treated with OTA in vivo and subsequently cultured in vitro to express chromosomal damage. (a) Normal metaphase and (b) metaphase showing chromatid break (obtained from a rat treated with 2000 µg/kg of body wt OTA for 2 weeks).

in vivo (39) and was applied in this study to assess genotoxicity of OTA and OTB in rats. Since OTA is slowly eliminated resulting in high organ concentrations and since high adduct levels have been reported to occur 4872 h after exposure to OTA (26, 27), DNA breakage was analyzed 72 h after administration of the final dose of OTA in this study. Using the alkaline (pH > 13) comet assay, which detects both single- and double-strand breaks, cross-links, incomplete excision-repair sites, as well as alkali labile sites (29), we observed that DNA damage was evident in liver, kidney, and spleen of animals treated with OTA for 2 weeks. These data are consistent with previous results obtained in mice, where OTA produced DNA single-strand breaks in liver, kidney, and the spleen detected by the alkaline elution procedure (24). In the present study, the increase in DNA breakage was dose-dependent in both liver and spleen, whereas more pronounced DNA damage was evident in kidneys of rats treated with lower doses of OTA. However, the gross histopathological alterations and the presence of apoptotic cells in kidneys of animals receiving high doses of OTA may provide an explanation for the bell-shaped dose response observed in the target organ of OTA carcinogenicity, as heavily damaged cells may have been lost due to cell death by apoptosis. Although the comet assay has been reported to present a suitable method for the detection of DNA fragmentation caused by apoptosis, large comets indicative of apoptotic cells where not observed despite the unambiguous increase in the incidence of apoptosis as assessed by histopathology (36). Consistent with our findings, a similar dose response for the induction of DNA damage was recently reported in CV-1 kidney cells treated with OTA in vitro (40). The fact that DNA damage induced by OTA was not restricted to

Figure 5. DNA content in cells extracted from cortical (a) and medullary (b) regions of the kidney after repeated administration of OTA (0, 250, 500, 1000, and 2000 µg/kg of body wt) and OTB (2000 µg/kg of body wt) to rats, indicating an increase in the incidence of polyploid cells in kidney medulla consistent with the histopathological findings. Statistical significance was determined by ANOVA and Dunnett’s test and is indicated as *, p < 0.05; **, p < 0.01; and ***, p < 0.001.

the kidney, the target organ of OTA carcinogenicity, clearly deserves attention. It has previously been suggested that slow elimination and renal accumulation may contribute to the organ specificity of OTA toxicity and carcinogenicity (16). However, following repeated administration of relatively high doses of OTA to rats, similar organ concentrations were observed in kidneys and livers of treated animals (36), consistent with the extent of DNA damage in these tissues. Strikingly, no histopathological changes were evident in the liver despite the high tissue concentrations and presence of DNA damage, whereas characteristic pathological alterations consisting of prominent karyomegaly, polyploidy, and increased incidence of apoptosis and mitosis were present in kidneys (36). On the basis of these observations, it must be concluded that DNA damage caused by OTA treatment in these organs is either not related to or not sufficient for carcinogenesis, and that additional or other types of

DNA Damage and Cytogenetic Effects of OTA

Chem. Res. Toxicol., Vol. 18, No. 8, 2005 1259

Figure 6. Representative chromatograms obtained by 32P-postlabeling analysis of DNA exctracted from kidneys of a control rat (a, f) and of rats administered 500 (b, g) and 2000 (c, h) µg/kg of body wt OTA, indicating lack of DNA-adduct formation by OTA. Separation of potential adducts was carried out using two different chromatographic conditions: solvent system A (LFU/SPU) (top panel (a-e)) and solvent system B (LFU/LTU) (bottom panel (f-j)). A mixture of OTA-dGMP adducts formed by photoirradiation of OTA in the presence of 3′-dGMP was used as adduct standard (d, i). Kidney DNA from rats treated with aristolochic acid (10 mg/kg of body wt) for 5 days served as positive control (e, j).

damage are required in order for tumor formation to occur. Interestingly, treatment with the non-chlorinated analogue OTB also resulted in strand breakage in liver, kidney, and spleen DNA to a similar extent as compared to the same dose of OTA. Little is known regarding the genotoxic potential of OTB, but OTB toxicity in rats is generally accepted to be much lower due to differences in toxicokinetics (41-43), and no histopathological alterations and functional changes indicative of nephrotoxicity were evident in OTB-treated animals in this study in contrast to OTA. The observations with OTB further question the biological significance and the contribution of the observed DNA damage to OTA carcinogenicity. The nature of the DNA damage by OTA was further characterized in OTA-treated animals. Analysis of the frequency of chromosomal aberrations in splenocytes isolated from rats treated with OTA and OTB in vivo indicates that the observed DNA damage was not converted into permanent genetic damage, although a small but not significant increase in chromatid and chromosome deletions was noted. This type of chromosomal aberrations, which are compatible with single- and double-strand breaks induced by oxidative stress (Figure 1c), but not with base modifications sensitive to FGP, are not representative of direct-acting genotoxins (44), suggesting that DNA damage induced by OTA and OTB is not a result of DNA-adduct formation. This is in agreement with the absence of DNA adducts in kidney DNA after high doses of OTA, as assessed by 32 P-postlabeling in this study and previous work in this lab, as well as with DNA-binding studies using radiolabeled compound with detection limits well below those of the postlabeling procedure. These studies do not support the hypothesis that OTA forms covalent adducts with DNA (14, 15, 17, 25).

For OTA, a role of DNA binding is also not supported by results from incubations of OTA with DNA or nucleotides in the presence of cytochromes P450s and peroxidases (with appropriate positive controls) or in rat hepatocytes (14, 15, 17). Moreover, enzymatic formation of reactive intermediates from OTA could not be demonstrated either (4, 14, 16). Biotransformation of OTA by rat cytochromes P450 occurs by oxidation of a methyl group at very low rates, but electrophiles capable of DNA binding are not formed during this reaction (4, 14). The absence of stable DNA adducts formed from OTA is also supported by results from genotoxicity testing, since the majority of bacterial mutagenicity tests were consistently negative, even in the presence of specific activation conditions, and both positive and negative results have been reported in mammalian cells (1-7). However, many of the positive responses occurred in the absence of metabolic activation in cell systems with very low capacities for biotransformation of xenobiotics (21-24). While the results from this study indicating absence of DNA-adduct formation by OTA are consistent with a large dataset as outlined above, they are in contrast to OTA-related DNA modifications that have been reported using 32P-postlabeling and have been attributed to direct binding of OTA (26, 34). High adduct levels have been reported in DNA extracted from livers and kidneys of mice following administration of a single dose (2.5 mg/ kg of body wt), even up to 16 days after treatment, although mice are known to be much less sensitive to renal tumor formation by OTA (26). However, the number of adducts and the “adduct maps” reported from the laboratory postulating DNA adducts are not consistent over a range of studies (sometimes, up to 30 adducts were described) (26, 27, 45-48), and more recently, lower numbers of adduct “spots” have been reported (34). In kidneys of rats treated with a single dose of 2 mg/kg of

1260

Chem. Res. Toxicol., Vol. 18, No. 8, 2005

body wt and sacrificed 48 h after administration, 12 distinct adduct spots were found (27). In the study presented here, rats were repeatedly administered this dose (corresponding to approximately 10 times the top dose used in the carcinogenicity bioassay, which resulted in a kidney tumor incidence of 74%) and no spots indicative of OTA-related DNA adducts were observed. Furthermore, the presence of the postulated OTA-3′dGMP adducts formed by photoirradiation of OTA (34) could not be confirmed in DNA extracted from kidneys of OTA-treated animals. In contrast, DNA extracted from animals treated with aristolochic acid, which served as positive controls, showed adduct maps consistent with the literature (49, 50). So far, all attempts to rationalize DNA-adduct formation by OTA have failed. Use of the postlabeling technique in the present study by a laboratory with extensive experience in postlabeling as well as previous attempts to demonstrate formation of the postulated OTA-C8deoxyguanosine by postlabeling and LC/MS/MS failed to detect OTA-DNA adducts despite method development using the authentic adduct standard (17). In addition, DNA binding could not be detected in several studies using radiolabeled OTA (14C and 3H) (14, 15, 17, 25). Therefore, results from the present study and literature data suggest that the DNA breakage detected using the comet assay is not caused by covalent binding of OTA to DNA, but rather by an indirect mechanism which may be mediated by reactive oxygen species. Oxidative stress has been implicated in OTA carcinogenicity (51-53). The comet assay in the presence of Fpg protein converts oxidative DNA base modifications into DNA single-strand breaks (30). When this modification of the comet assay was used, DNA damage was enhanced in liver and kidney as well as in peripheral lymphocytes. Again, this type of DNA damage was not target-organ-specific, and treatment with OTB resulted in a similar degree of DNA breakage as the same dose of OTA. These results appear to be in contrast with data obtained using a panel of oxidative stress markers, including determination of 8-OH-dG in liver and kidney DNA, which suggest that OTA does not induce overt oxidative stress conditions as caused by other compounds known to induce oxidative stress in the kidney such as iron(III)nitrilotriacetic acid (36). However, the Fpg-modified comet assay has been shown to be a very sensitive marker of oxidative DNA damage, and the results presented here indicate that the observed DNA damage may be a result of oxidative stress induced by OTA treatment. These data are in line with recent results by Kamp et al., who reported DNA-strand breakage in various cell lines and primary rat kidney cells in response to OTA treatment using the modified comet assay (40). However, it should be noted that Fpg glycosylase is not specific for 8-OH-dG and is also known to recognize other oxidized purines as well as abasic sites (AP sites) and ring-opened N-7 guanine adducts (30-33). The direct cause for the oxidative DNA damage and its contribution to OTA carcinogenicity still remain to be elucidated. OTA genotoxicity in mammalian cells was often shown to occur independent of biotransformation, and therefore, generation of reactive oxygen species mediated by cytochrome P450s seems unlikely, although de Groene et al. reported increased mutation frequencies in OTA-treated NIH/3T3 cells stably expressing human cytochromes P450 (13). Oxidative stress may also be caused by interference of OTA with calcium homeostasis

Mally et al.

and changes in intracellular calcium. These were evident in SHE cells following treatment with OTA at concentrations which also resulted in disruption of the cytoskeleton and formation of micronuclei (54). Moreover, OTA has been reported to impair calcium and cAMP homeostasis in immortalized human kidney epithelial cells (55). Taken together, the data presented here demonstrate that repeated administration of OTA causes DNA-strand breaks in target and nontarget tissues of male rats by a mechanism which may involve oxidative stress. However, the lack of target-organ-specific induction of DNA damage strongly suggests that additional events are required for renal tumor formation by OTA. The present results also do not confirm that OTA produces DNA adducts containing the OTA moiety, consistent with the results of mechanistic studies using a variety of endpoints ranging from biotransformation to genotoxicity.

Acknowledgment. Parts of this work were supported by the Fifth RTD Framework Program of the European Union, Project No. QLK1-2001-01614 and by USPHS Grant CA77114. The authors would also like to thank Prof. Heiko Ihmels and Daniela Otto, Department of Organic Chemistry, University of Siegen, Germany, for photochemical preparation of the OTA-3′-dGMP standard.

References (1) NTP (1989) Toxicology and carcinogenesis studies of ochratoxin A (CAS No. 303-47-9) in F344/N rats (gavage studies). Natl. Toxicol. Program Tech. Rep. Ser. 358, 1-142. (2) Wehner, F. C., Thiel, P. G., van Rensburg, S. J., and Demasius, I. P. (1978) Mutagenicity to Salmonella typhimurium of some Aspergillus and Penicillium mycotoxins. Mutat. Res. 58, 193-203. (3) Wurgler, F. E., Friederich, U., and Schlatter, J. (1991) Lack of mutagenicity of ochratoxin A and B, citrinin, patulin and cnestine in Salmonella typhimurium TA102. Mutat. Res. 261, 209-216. (4) Zepnik, H., Pahler, A., Schauer, U., and Dekant, W. (2001) Ochratoxin A-induced tumor formation: is there a role of reactive ochratoxin A metabolites? Toxicol. Sci. 59, 59-67. (5) Bendele, A. M., Neal, S. B., Oberly, T. J., Thompson, C. Z., Bewsey, B. J., Hill, L. E., Rexroat, M. A., Carlton, W. W., and Probst, G. S. (1985) Evaluation of ochratoxin A for mutagenicity in a battery of bacterial and mammalian cell assays. Food Chem. Toxicol. 23, 911-918. (6) Follmann, W., and Lucas, S. (2003) Effects of the mycotoxin ochratoxin A in a bacterial and a mammalian in vitro mutagenicity test system. Arch. Toxicol. 77, 298-304. (7) Ehrlich, V., Darroudi, F., Uhl, M., Steinkellner, H., Gann, M., Majer, B. J., Eisenbauer, M., and Knasmuller, S. (2002) Genotoxic effects of ochratoxin A in human-derived hepatoma (HepG2) cells. Food Chem. Toxicol. 40, 1085-1090. (8) Hennig, A., Fink-Gremmels, J., and Leistner, L. (1991) Mutagenicity and effects of ochratoxin A on the frequency of sister chromatid exchange after metabolic activation. IARC Sci. Publ., 255-260. (9) Obrecht-Pflumio, S., Chassat, T., Dirheimer, G., and Marzin, D. (1999) Genotoxicity of ochratoxin A by Salmonella mutagenicity test after bioactivation by mouse kidney microsomes. Mutat. Res. 446, 95-102. (10) Auffray, Y., and Boutibonnes, P. (1986) Evaluation of the genotoxic activity of some mycotoxins using Escherichia coli in the SOS spot test. Mutat. Res. 171, 79-82. (11) Krivobok, S., Olivier, P., Marzin, D. R., Seigle-Murandi, F., and Steiman, R. (1987) Study of the genotoxic potential of 17 mycotoxins with the SOS Chromotest. Mutagenesis 2, 433-439. (12) Malaveille, C., Brun, G., and Bartsch, H. (1991) Genotoxicity of ochratoxin A and structurally related compounds in Escherichia coli strains: studies on their mode of action. IARC Sci. Publ., 261-266. (13) de Groene, E. M., Hassing, I. G., Blom, M. J., Seinen, W., FinkGremmels, J., and Horbach, G. J. (1996) Development of human cytochrome P450-expressing cell lines: application in mutagenicity testing of ochratoxin A. Cancer Res. 56, 299-304.

DNA Damage and Cytogenetic Effects of OTA (14) Gautier, J., Richoz, J., Welti, D. H., Markovic, J., Gremaud, E., Guengerich, F. P., and Turesky, R. J. (2001) Metabolism of ochratoxin A: absence of formation of genotoxic derivatives by human and rat enzymes. Chem. Res. Toxicol. 14, 34-45. (15) Gross-Steinmeyer, K., Weymann, J., Hege, H. G., and Metzler, M. (2002) Metabolism and lack of DNA reactivity of the mycotoxin ochratoxin a in cultured rat and human primary hepatocytes. J. Agric. Food Chem. 50, 938-945. (16) Zepnik, H., Volkel, W., and Dekant, W. (2003) Toxicokinetics of the mycotoxin ochratoxin A in F 344 rats after oral administration. Toxicol. Appl. Pharmacol. 192, 36-44. (17) Mally, A., Zepnik, H., Wanek, P., Eder, E., Dingley, K., Ihmels, H., Vo¨lkel, W., and Dekant, W. (2004) Ochratoxin A: lack of formation of covalent DNA adducts. Chem. Res. Toxicol. 17, 234242. (18) Dorrenhaus, A., Flieger, A., Golka, K., Schulze, H., Albrecht, M., Degen, G. H., and Follmann, W. (2000) Induction of unscheduled DNA synthesis in primary human urothelial cells by the mycotoxin ochratoxin A. Toxicol. Sci. 53, 271-277. (19) Mori, H., Kawai, K., Ohbayashi, F., Kuniyasu, T., Yamazaki, M., Hamasaki, T., and Williams, G. M. (1984) Genotoxicity of a variety of mycotoxins in the hepatocyte primary culture/DNA repair test using rat and mouse hepatocytes. Cancer Res. 44, 2918-2923. (20) Dorrenhaus, A., and Follmann, W. (1997) Effects of ochratoxin A on DNA repair in cultures of rat hepatocytes and porcine urinary bladder epithelial cells. Arch. Toxicol. 71, 709-713. (21) Lebrun, S., and Follmann, W. (2002) Detection of ochratoxin A-induced DNA damage in MDCK cells by alkaline single cell gel electrophoresis (comet assay). Arch. Toxicol. 75, 734-741. (22) de Groene, E. M., Jahn, A., Horbach, G. J., and Fink-Gremmels, J. (1996) Mutagenicity and genotoxicity of the mycotoxin ochratoxin A. Environ. Toxicol. Pharmacol. 1, 21-26. (23) Stetina, R., and Votava, M. (1986) Induction of DNA single-strand breaks and DNA synthesis inhibition by patulin, ochratoxin A, citrinin, and aflatoxin B1 in cell lines CHO and AWRF. Folia Biol. (Prague) 32, 128-144. (24) Creppy, E. E., Kane, A., Dirheimer, G., Lafarge-Frayssinet, C., Mousset, S., and Frayssinet, C. (1985) Genotoxicity of ochratoxin A in mice: DNA single-strand break evaluation in spleen, liver and kidney. Toxicol. Lett. 28, 29-35. (25) Schlatter, C., Studer, R. J., and Rasonyi, T. (1996) Carcinogenicity and kinetic aspects of ochratoxin A. Food Addit. Contam., 4344. (26) Pfohl-Leszkowicz, A., Grosse, Y., Kane, A., Creppy, E. E., and Dirheimer, G. (1993) Differential DNA adduct formation and disappearance in three mouse tissues after treatment with the mycotoxin ochratoxin A. Mutat. Res. 289, 265-273. (27) Pfohl-Leskowicz, A., Bartsch, H., Azemar, B., Mohr, U., Esteve, J., and Castegnaro, M. (2002) Mesna protects rats against nephrotoxicity but not carcinogenicity induced by ochratoxin A, implicating two separate pathways. Facta Universitatis, Med. Biol. 9, 57-63. (28) Savage, J. R. (1977) Application of chromosome banding techniques to the study of primary chromosome structural changes. J. Med. Genet. 14, 362-370. (29) Singh, N. P., McCoy, M. T., Tice, R. R., and Schneider, E. L. (1988) A simple technique for quantitation of low levels of DNA damage in individual cells. Exp. Cell Res. 175, 184-191. (30) Boiteux, S., Gajewski, E., Laval, J., and Dizdaroglu, M. (1992) Substrate specificity of the Escherichia coli Fpg protein (formamidopyrimidine-DNA glycosylase): excision of purine lesions in DNA produced by ionizing radiation or photosensitization. Biochemistry 31, 106-110. (31) Tchou, J., Bodepudi, V., Shibutani, S., Antoshechkin, I., Miller, J., Grollman, A. P., and Johnson, F. (1994) Substrate specificity of Fpg protein. Recognition and cleavage of oxidatively damaged DNA. J. Biol. Chem. 269, 15318-15324. (32) Li, Q., Laval, J., and Ludlum, D. B. (1997) Fpg protein releases a ring-opened N-7 guanine adduct from DNA that has been modified by sulfur mustard. Carcinogenesis 18, 1035-1038. (33) Tudek, B., Van Zeeland, A. A., Kusmierek, J. T., and Laval, J. (1998) Activity of Escherichia coli DNA-glycosylases on DNA damaged by methylating and ethylating agents and influence of 3-substituted adenine derivatives. Mutat. Res. 407, 169-176. (34) Faucet, V., Pfohl-Leszkowicz, A., Dai, J., Castegnaro, M., and Manderville, R. A. (2004) Evidence for covalent DNA adduction by ochratoxin A following chronic exposure to rat and subacute exposure to pig. Chem. Res. Toxicol. 17, 1289-1296. (35) Gupta, R. C. (1993) 32P-postlabelling analysis of bulky aromatic adducts. IARC Sci. Publ., 11-23.

Chem. Res. Toxicol., Vol. 18, No. 8, 2005 1261 (36) Mally, A., Vo¨lkel, W., Amberg, A., Kurz, M., Wanek, P., Eder, E., Hard, G., and Dekant, W. (2005) Functional, biochemical, and pathological effects of repeated administration of ochratoxin A to rats. Chem. Res. Toxicol. 18, 1242-1252. (37) Obrecht-Pflumio, S., and Dirheimer, G. (2001) Horseradish peroxidase mediates DNA and deoxyguanosine 3′-monophosphate adduct formation in the presence of ochratoxin A. Arch. Toxicol. 75, 583-590. (38) Obrecht-Pflumio, S., and Dirheimer, G. (2000) In vitro DNA and dGMP adducts formation caused by ochratoxin A. Chem.-Biol. Interact. 127, 29-44. (39) Ostling, O., Johanson, K. J., Blomquist, E., and Hagelqvist, E. (1987) DNA damage in clinical radiation therapy studied by microelectrophoresis in single tumour cells. A preliminary report. Acta Oncol. 26, 45-48. (40) Kamp, H. G., Eisenbrand, G., Schlatter, J., Wurth, K., and Janzowski, C. (2005) Ochratoxin A: induction of (oxidative) DNA damage, cytotoxicity and apoptosis in mammalian cell lines and primary cells. Toxicology 206, 413-425. (41) Mally, A., Keim-Heusler, H., Amberg, A., Kurz, M., Zepnik, H., Mantle, P., Vo¨lkel, W., Hard, G., and Dekant, W. (2005) Biotransformation and nephrotoxicity of ochratoxin B in rats. Toxicol. Appl. Pharmacol., in press. (42) Breitholtz-Emanuelsson, A., Fuchs, R., Hult, K., and Appelgren, L. E. (1992) Synthesis of 14C-ochratoxin A and 14C-ochratoxin B and a comparative study of their distribution in rats using whole body autoradiography. Pharmacol. Toxicol. 70, 255-261. (43) Hagelberg, S., Hult, K., and Fuchs, R. (1989) Toxicokinetics of ochratoxin A in several species and its plasma-binding properties. J. Appl. Toxicol. 9, 91-96. (44) Duncam, A. M., and Evans, H. J. (1983) The exchange hypothesis for the formation of chromatid aberrations: an experimental test using bleomycin. Mutat. Res. 107, 307-313. (45) Castegnaro, M., Mohr, U., Pfohl-Leszkowicz, A., Esteve, J., Steinmann, J., Tillmann, T., Michelon, J., and Bartsch, H. (1998) Sex- and strain-specific induction of renal tumors by ochratoxin A in rats correlates with DNA adduction. Int. J. Cancer 77, 7075. (46) Petkova-Bocharova, T., Stoichev, I. I., Chernozemsky, I. N., Castegnaro, M., and Pfohl-Leszkowicz, A. (1998) Formation of DNA adducts in tissues of mouse progeny through transplacental contamination and/or lactation after administration of a single dose of ochratoxin A to the pregnant mother. Environ. Mol. Mutagen. 32, 155-162. (47) Grosse, Y., Baudrimont, I., Castegnaro, M., Betbeder, A. M., Creppy, E. E., Dirheimer, G., and Pfohl-Leszkowicz, A. (1995) Formation of ochratoxin A metabolites and DNA-adducts in monkey kidney cells. Chem.-Biol. Interact. 95, 175-187. (48) Obrecht-Pflumio, S., Grosse, Y., Pfohl-Leszkowicz, A., and Dirheimer, G. (1996) Protection by indomethacin and aspirin against genotoxicity of ochratoxin A, particularly in the urinary bladder and kidney. Arch. Toxicol. 70, 244-248. (49) Pfau, W., Schmeiser, H. H., and Wiessler, M. (1990) 32Ppostlabelling analysis of the DNA adducts formed by aristolochic acid I and II. Carcinogenesis 11, 1627-1633. (50) Arlt, V. M., Pfohl-Leszkowicz, A., Cosyns, J., and Schmeiser, H. H. (2001) Analyses of DNA adducts formed by ochratoxin A and aristolochic acid in patients with Chinese herbs nephropathy. Mutat. Res. 494, 143-150. (51) Rahimtula, A. D., Bereziat, J. C., Bussacchini-Griot, V., and Bartsch, H. (1988) Lipid peroxidation as a possible cause of ochratoxin A toxicity. Biochem. Pharmacol. 37, 4469-4477. (52) Schaaf, G. J., Nijmeijer, S. M., Maas, R. F., Roestenberg, P., de Groene, E. M., and Fink-Gremmels, J. (2002) The role of oxidative stress in the ochratoxin A-mediated toxicity in proximal tubular cells. Biochim. Biophys. Acta 1588, 149-158. (53) Gautier, J. C., Holzhaeuser, D., Markovic, J., Gremaud, E., Schilter, B., and Turesky, R. J. (2001) Oxidative damage and stress response from ochratoxin a exposure in rats. Free Radical Biol. Med. 30, 1089-1098. (54) Dopp, E., Muller, J., Hahnel, C., and Schiffmann, D. (1999) Induction of genotoxic effects and modulation of the intracellular calcium level in syrian hamster embryo (SHE) fibroblasts caused by ochratoxin A. Food Chem. Toxicol. 37, 713-721. (55) Benesic, A., Mildenberger, S., and Gekle, M. (2000) Nephritogenic ochratoxin A interferes with hormonal signalling in immortalized human kidney epithelial cells. Pflugers Arch. 439, 278-287.

TX049650X