Vanadate induces DNA strand breaks in cultured human fibroblasts at doses relevant to occupational exposure

Mutation Research 519 (2002) 25–35

Vanadate induces DNA strand breaks in cultured human fibroblasts at doses relevant to occupational exposure Sabine Ivancsits∗ , Alexander Pilger, Elisabeth Diem, Andreas Schaffer, Hugo W. Rüdiger Division of Occupational Medicine, University Hospital/AKH, University of Vienna, Waehringer Guertel 18-20, A-1090 Vienna, Austria Received 30 October 2001; received in revised form 4 May 2002; accepted 6 May 2002

Abstract To study possible genotoxic effects of occupational exposure to vanadium pentoxide, we determined DNA strand breaks (with alkaline comet assay), 8-hydroxy-2 deoxyguanosine (8-OHdG) and the frequency of sister chromatid exchange (SCE) in whole blood leukocytes or lymphocytes of 49 male workers employed in a vanadium factory in comparison to 12 non-exposed controls. In addition, vanadate has been tested in vitro to induce DNA strand breaks in whole blood cells, isolated lymphocytes and cultured human fibroblasts of healthy donors at concentrations comparable to the observed levels of vanadium in vivo. To investigate the impact of vanadate on the repair of damaged DNA, co-exposure to UV or bleomycin was used in fibroblasts, and DNA migration in the alkaline and neutral comet assay was determined. Although, exposed workers showed a significant vanadium uptake (serum: median 5.38 ␮g/l, range 2.18–46.35 ␮g/l) no increase in cytogenetic effects or oxidative DNA damage in leukocytes could be demonstrated. This was consistent with the observation that in vitro exposure of whole blood leukocytes and lymphocytes to vanadate caused no significant changes in DNA strand breaks below concentrations of 1 ␮M (50 ␮g/l). In contrast, vanadate clearly induced DNA fragmentation in cultured fibroblasts at relevant concentrations. Combined exposure of fibroblasts to vanadate/UV or vanadate/bleomycin resulted in non-repairable DNA double strand breaks (DSBs) as seen in the neutral comet assay. We conclude that exposure of human fibroblasts to vanadate effectively causes DNA strand breaks, and co-exposure of cells to other genotoxic agents may result in persistent DNA damage. © 2002 Elsevier Science B.V. All rights reserved. Keywords: Vanadium; Comet assay; Oxidative DNA damage; Sister chromatid exchange

1. Introduction Vanadium is the first element of the fifth group in the periodic system of elements existing in oxidation states ranging from −1 to +5, preferentially +3, +4 (vanadyl) and +5 (vanadate). The latter is the most common and the quadrivalent, the most stable form. The most frequently used vanadium compound ∗ Corresponding author. Tel.: +43-1-40400-4022; fax: +43-1-4088011. E-mail address: [email protected] (S. Ivancsits).

is vanadium pentoxide (V2 O5 , TLV = 0.05 mg/m3 ), a yellowish to red-brown crystalline powder, which is sparingly soluble in water. Due to its hardness and its ability to form alloys, vanadium is frequently found as a constituent of steel used for tools or orthopedic implants. Vanadium salts serve as catalysts for the production of sulfuric acid, semiconductors, photographic developers, and yellow pigments in ceramics. Occupational exposure occurs during vanadium production and cleaning of oil- or gas-fired boilers, as vanadium is a natural component of certain Mexican and Venezuelan oils. Exposure to

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vanadium dusts may cause irritation of the skin and the respiratory tract [1,2] and may lead to a greenish-black discoloration of the tongue [3]. Vanadium compounds, especially insoluble vanadium oxides, are hardly absorbed from the intestinal tract [4], but easily from the lung [5]. The distribution of vanadium via blood circulation is rapid and the highest concentrations of vanadium initially appear in the kidney, liver and lungs. About 90% of plasma vanadium is transported via binding to transferrin or albumin [6]. The principal route of elimination of absorbed vanadium is through the kidneys. The kinetics of vanadium elimination via urine is biphasic with an initial rapid rate of elimination (10–20 h) and a longer terminal phase (40–50 days) [7,8] and can be used as biological monitor in occupationally exposed workers. Below pH 3.5, vanadium mainly exists as vanadyl (VO2+ ), while in alkaline solutions the ortho-vanadate form is predominant, which is chemically similar to phosphates. In neutral solutions, vanadium occurs in its pentavalent form as H2 VO4 − , which is the most toxic, as it easily enters the cell, where it is reduced to vanadyl (+4). Vanadyl interacts with cellular molecules, and an involvement in free radical generating processes has been postulated [9]. In the general population, food is the major source of vanadium exposure. Vanadium is not essential for mammals and the daily uptake of vanadium is small (<10 ␮g per day). The highest amounts of vanadium can be found in black pepper, dill seed, mushrooms, parsley and shellfish [10,11]. In general, seafood contains higher concentrations of vanadium compared with terrestrial sources of food. High concentrations of 1–8 ppm vanadium [12] are detected in tobacco smoke. In medicine, vanadium compounds (e.g. sodium meta-vanadate) have been administered for the treatment of anemia, tuberculosis, diabetes, and syphilis in doses of 1–8 mg [13]. Vanadyl sulfate (VOSO4 ) is a common food supplement in weight training athletes at doses up to 60 mg per day. Vanadium displays several metabolic and pharmacological actions. It is an inhibitor of the Na/K/ATPase, interferes with phosphate-containing enzymes [14] and mimics the action of insulin by increasing glucose transport and improving glucose metabolism in in vitro and in animal experiments [15]. The toxicity of vanadium compounds depends on the route of administration and increases with higher valencies.

Investigations on vanadium-exposed workers on blood count, hematological and clinical laboratory parameters, did not reveal any significant differences between exposed individuals and controls, although an increase in inflammatory cells in the nasal mucosa of exposed workers could be detected [16]. In addition, several studies have demonstrated severe respiratory symptoms and irritations of the bronchial tract in vanadium-exposed workers and boilermakers [17–19]. Although, vanadium interferes with mitosis and chromosome distribution [20] and may generate reactive oxygen species due to its redox potential, it is only a weak mutagen [21] and there is no evidence for carcinogenic effects. Cytogenetic studies on mammalian cells have demonstrated that vanadium compounds increase the frequency of micronuclei, sister chromatid exchanges (SCEs) and induce DNA single strand breaks (SSBs) and DNA–protein cross-links [22–26]. An investigation of patients with joint prostheses made of titanium–aluminium–vanadium alloys [27] revealed a significant elevation of SCE values compared to the control population, indicating an in vivo genotoxic action of vanadium. Only a few studies exist about the genotoxic action of vanadium compounds. Most of them are in vitro experiments, carried out at metal concentrations far higher than in vivo exposure data. Our work is the first study directly relating in vivo with in vitro effects of vanadium. The aim of the study was to evaluate the in vivo genotoxic hazards resulting from longterm vanadium exposure by means of SCE test and alkaline comet assay. In addition, 8-hydroxy-2 deoxyguanosine (8-OHdG), a well-established marker for oxidative stress, was determined in lymphocytes of vanadium-exposed workers. In vitro exposure was carried out with whole blood cells, isolated lymphocytes and cultured human fibroblasts at vanadium concentrations determined in the in vivo investigation. Genotoxicity was assessed in vitro using the alkaline and neutral comet assay as well as 8-OHdG analysis. 2. Materials and methods 2.1. Subjects for in vivo study Forty-nine male workers exposed to vanadium pentoxide and 12 non-exposed controls from the same factory were studied for the grade of DNA fragmen-

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Table 1 Demographic data of the examined workers (mean ± S.D. and ranges) Non-exposed

Exposed

Number of probands Age in years (range) Years of exposure (range) Body mass index (BMI) (range)

12 50.1 ± 7.3 (38–59) – 26.9 ± 3.1 (22.8–31.8)

49 42.4 ± 7 (27–57) 12.4 ± 7.9 (0.5–31) 26.2 ± 2.5 (20.3–34.7)

Smoking habits Non-smokers Smokers Ex-smokers

8 3 1

13 27 9

tation, amount of 8-OHdG and frequency of SCE in whole blood leukocytes or lymphocytes. Subject demographics are presented in Table 1. All workers came from the same working area and used, at least occasionally, protective masks during work (70% always, 30% occasionally). Forty-two percent of the workers considered vanadium dust exposure at the working place as high, 52% as medium and 6% as low. The occurrence of green or black tongue (n = 10) was the only significant symptom related to vanadium pentoxide exposure. Unspecific symptoms included cough (n = 33), irritation of the eyes (n = 31), rhinitis (n = 25) and irritation of the throat (n = 24). Blood and urine samples were collected prior to work-shift, according to Hauser et al. [28], who figured out, that in biomonitoring studies sample collection prior to shift should be preferred to sampling at the end of shift. Data collection was performed by questionnaire, which included information on years of employment, age, body-weight, stature, smoking habits and diet. 2.2. Assessment of vanadium exposure Vanadium in serum (VS ) was determined by graphite furnace atomic absorption spectrometry (GF-AAS Perkin-Elmer 5100-ZL spectrophotometer) with Zeeman background correction at a temperature of 2450 ◦ C. The calibration curves for vanadium ranged from 0 to 5 ␮g/l and from 0 to 40 ␮g/l, respectively. The detection limit in serum was 0.3 ␮g/l (5.9 nmol/l). Urinary vanadium (VU ) was analyzed with GF-AAS at a temperature of 2450 ◦ C and quantified using calibration curves ranged 0–5 and 0–40 ␮g/l, respectively. The detection limits in urine was 0.2 ␮g/l (3.9 nmol/l).

Blood and urinary samples were taken from each subject in the morning. Blood was centrifuged 10 min at 800 × g in Vacationer glass tubes (Becton Dickinson Europe, Myelin, France) and collected serum was stored at +4 ◦ C until analysis. Urinary specimens were acidified and stored in plastic tubes (Sarstedt, Nümbrecht, Germany) at −20 ◦ C until analysis. 2.3. Cell culture and in vitro vanadate treatment Heparinized blood was collected from a healthy donor (female, age 26, smoker) by venepuncture. Either whole blood (2 ml) or isolated (Ficoll Paque© density gradient centrifugation, according to manufacturer’s instructions), non-stimulated lymphocytes (diluted in 5 ml RPMI 1640) were incubated with ortho-vanadate (0.5–10 ␮M, Na3 VO4 ) for 22 h at 37 ◦ C on a shaking water bath. Then, alkaline comet assay analysis was performed. Human diploid fibroblasts were initiated from a skin biopsy from the upper arm of a healthy donor and cultivated in Dulbecco’s modified Eagles medium (DMEM, Gibco, Vienna, Austria) supplemented with 10% fetal calf serum (FCS), 20 mM Hepes buffer, 40 ␮g/ml neomycin, 2 mM l-glutamine, 100 IU/ml penicillin and 100 ␮g/ml streptomycin (Gibco, Vienna, Austria) in the presence of a 5% CO2 atmosphere. Cells were grown in 175 cm2 culture flasks and were supplied with fresh culture medium every 48 h. At confluence cells were treated with different vanadate concentrations (0.5–10 ␮M) in RPMI 1640 medium for 22 h at usual cultivation conditions. To prevent complexion effects, exposure was performed without phenol red or additives. After incubation cells

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were detached by trypsin. Cell viability was determined by trypan blue exclusion assay, where treated and untreated cells showed a viability of 90 and 95%, respectively. 2.4. DNA isolation and digestion for 8-OHdG analysis DNA isolation from whole blood and cultured cells was performed with Qiagen genomic DNA-Kit (R. and P. Margaritella GmbH, Vienna, Austria) according to the manufacturer’s instructions with several modifications. Briefly, in case of cultured fibroblasts, cells were suspended in cold PBS containing 1 mM of desferoxamine mesylate (DFMA) and lysed by addition of 2 ml lysis buffer (320 mM saccharose, 5 mM MgCl2 , 10 mM Tris–HCl, 1% Triton-X-100 pH 7.5) and 6 ml Chelex-100 treated bidistilled water. The tubes were inverted several times followed by incubation on ice for 5 min. The lysed cells were centrifuged (1300 × g, 4 ◦ C, 15 min), and the supernatant was discarded. The remaining pellet was re-suspended in 1 ml lysis buffer, 3 ml bidistilled water and DFMA in a final concentration of 2 mM. After short vortexing and incubation on ice for 5 min, the lysate was centrifuged (1300 × g, 4 ◦ C, 15 min). The supernatant was discarded and the nuclei were re-suspended in 5 ml general lysis buffer (800 mM guanidinium/HCl, 30 mM EDTA, 30 mM Tris–HCl, 5% Tween-20, 0.5% Triton-X-100 pH 8) with 6 mM DFMA. Subsequently DNase-free RNase A [29] (200 ␮g/ml, Sigma) and Protease (400 ␮g/ml) were added and the nuclei were incubated at 50 ◦ C for 1 h. After addition of NaCl yielding a final concentration of 2 mM, DNA was precipitated by addition of 3.5 ml isopropanol. Finally, DNA was transferred with a plastic rod into 1.5 ml polypropylene tubes containing 300 ␮l of ethanol (70%). DNA was precipitated and stored at −20 ◦ C until analysis. DNA digestion and analysis were performed according to Shigenaga et al. [30] with slight modifications. DNA in 300 ␮l 70% ethanol was pelleted, the supernatant was discarded, and the DNA pellet was dried under a slight stream of nitrogen (1–2 min) and dissolved in 180 ␮l of sodium acetate buffer (20 mM pH 5.5). After denaturation of DNA by incubation at 95 ◦ C for 3 min and addition of 100 ␮M DFMA the DNA was incubated with 10 units of nuclease

P1 (Sigma) for 10 min at 65 ◦ C. Subsequently, the solution was adjusted to pH 8 by addition of 20 ␮l Tris–HCl buffer (1 M, pH 8.5) and the nucleotides were incubated with 4 units of alkaline phosphatase (Boehringer-Mannheim, Germany) at 37 ◦ C for 1 h. After removal of enzymes by centrifugation (12 000 × g, 10–15 min, 4 ◦ C) in 0.4 ml vials with 10 000 NMGG filter units (UFC3 TGC NB, Millipore Ltd., Watford, UK), the nucleosides were subjected to analysis. 2.5. HPLC analysis of 8-OHdG The nucleoside mixture (80 ␮l) was injected into a reversed phase C 18 column (4 ␮m, EcoCART, Superspher 100 RP-18 end capped, 125 mm × 3 mm, Merck GmbH, Darmstadt, Germany) with guard column (5 ␮m, LiChroCART, LiChrospher 100 RP-18 end capped, 4 mm × 4 mm) and analyzed by a high performance liquid chromatography (HPLC) system (Hewlett-Packard 1050-series) with UV absorbance detection and electrochemical detection (Hewlett-Packard Model 1049 amperometric electrochemical detector with a silver/silver chloride reference electrode) coupled in series. Separation was performed at a flow rate of 0.5 ml/min with 50 mM potassium phosphate pH 5.5/1% methanol/0.5% tetrahydrofuran in case of calf thymus DNA and 50 mM potassium phosphate pH 5.5/1.5% acetonitrile/0.1% tetrahydrofuran in case of cellular DNA. The system was monitored by a computer (Hewlett-Packard, Vectra) with adequate software (HPLC-Chemstation, Revision 4.02). Nucleosides were measured by UV-detection at 270 nm. The 8-OHdG was detected at 0.6 V. Amounts of 8-OHdG and dG were determined using calibration curves of these two nucleosides (Wako Chemicals GmbH, Neuss, Germany). The rate of 8-OHdG formation was calculated by relating the amount of 8-OHdG to the amount of 105 dG. 2.6. Analysis of sister chromatid exchanges Lymphocyte cultures were prepared with 0.4 ml heparinized whole blood in 5 ml of chromosome medium 1A (Gibco, Vienna, Austria), containing phytohemagglutinin and 75 ␮g 5-bromodeoxyuridine (Serva, Heidelberg, Germany). After the cells were incubated at 37 ◦ C for 70 h, 0.5 ␮g colcemid (Gibco,

S. Ivancsits et al. / Mutation Research 519 (2002) 25–35

Vienna, Austria) were added and incubation was continued for another 2 h. The cells were harvested, and slides were prepared using standard techniques [31]. A total of 30 well-spread second-division metaphases from each culture sample were scored for SCE according to Vereantern et al. [32] from coded slides in duplicate by two different scorers. 2.7. In vitro UV-irradiation, bleomycin treatment and comet assay We followed the technique described by Östling and Johanson [33] with minor modifications by Singh et al. [34,35]. Fully frosted microscopic slides (CMS, Houston, USA) were used. In vivo or in vitro vanadium exposed cells (10 000–30 000, 10 ␮l whole blood) and controls were mixed with 100 ␮l low melting agarose (37 ◦ C) to form a cell suspension. The cell suspension was rapidly pipetted onto 0.5% normal melting agarose pre-coated slides, spread using a cover slip, and maintained on a cold flat tray for about 10 min to solidify. After removal of the cover slip the third layer of 0.5% low melting agarose at 37 ◦ C was added and solidified. Embedded fibroblasts were additionally exposed to UV or to bleomycin sulfate. UV irradiation was performed on ice using a germicidal lamp (G30T8, 30 W, Sankyo denki, Japan) which output predominantly contains UV C (253.7 nm). The distance between the UV lamp and the slides was 28 cm. The samples were exposed at an intensity of 8 W/m2 (measured by radiometer, Blak-ray® , Ultra-violet Products Inc., Model J225, San Gabriel, USA) for 10 min, which equals 4.8 kJ/m2 . For bleomycin treatment slides were incubated for 30 min in RPMI 1640 medium, containing 1 ␮g/ml bleomycin (1.5 × 10−3 U/ml). To study repair kinetics, cells were further incubated at 37 ◦ C for further 10 or 20 min. The slides were immersed in freshly prepared cold lysis solution (2.5 M NaCl, 100 mM Na2 EDTA, 10 mM Tris, pH 10, 1% sodium sarcosinate, 1% Triton-X-100, 10% DMSO) and lysed for 90 min at 4 ◦ C. Subsequently, the slides were drained and placed in a horizontal gel electrophoresis tank side by side, nearest the anode. The tank was filled with fresh electrophoresis buffer (1 mM Na2 EDTA, 300 mM NaOH, pH 13 in case of alkaline comet assay and 100 mM

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Tris, 300 mM sodium acetate, 500 mM sodium chloride, pH 8.5 in case of neutral comet assay) to a level approximately 0.4 cm above the slides. The slides were left in the solution for 40 min for equilibration and unwinding of the DNA before electrophoresis. Electrophoresis was performed at 25 V and 300 mA (4 ◦ C, 20 min, field strength: 0.8 V/cm). All steps were performed under dimmed light to prevent the occurrence of additional DNA damage. After electrophoresis the slides were washed three times with Tris buffer (0.4 M Tris, pH 7.5) to neutralize, then air-dried and stored until analysis. Comets were visualized by ethidium bromide staining (20 ␮g/ml, 30 s) and examined at 400× magnification using a fluorescence microscope (Axiophot, Zeiss, Germany). One thousand DNA spots from each sample were classified into five categories corresponding to the amounts of DNA in the tail according to Anderson [36] with modifications. Results were expressed as “comet tailfactors”, calculated according to Diem and Rüdiger [37]. All analyses were performed by the same investigator. Fig. 1 shows the five classification groups with the group averages and the microphotograph. Tailfactors were calculated according to the following formula: tailfactor (%) A∗ FA + B ∗ FB + C ∗ FC + D ∗ FD + E ∗ FE = 1000 where A is the number of cells classified to group A, FA the average of group A (2.5), B the number of cells classified to group B, FB the average of group B (12.5), C the number of cells classified to group C, FC the average of group C (30.0), D the number of cells classified to group D, FD the average of group D (67.5); E the number of cells classified to group E, FE the average of group E (97.5). 2.8. Statistical analysis Statistical analysis was performed with STATISTICA Version 5.0 package (Statsoft Inc., Tulsa, USA). All data are presented as mean ± standard deviation (S.D.) and median and range. Comparisons and correlations were performed by one-way ANOVA using Newman–Keuls test and by multiple regression analysis using linear regression.

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Fig. 1. Classification groups and corresponding microscopic appearance.

3. Results 3.1. Internal exposure to vanadium, cytogenetic effects and oxidative DNA damage Vanadium uptake during work is mainly due to respiration of vanadium dusts and thus depends on the respiratory volume, the kind of work, the size of particles and the usage of protective measures. Therefore, we measured vanadium concentrations in serum and urine for the assessment of internal vanadium exposure. Vanadium concentrations in serum (VS ) and urine (VU ) were significantly higher in exposed subjects (VS : 7.73 (2.18–46.3) ␮g/l versus 3.43 (1.01–12.5) ␮g/l, P = 0.005; VU , 14.57 (2.11–95.2) ␮g/g creatinine versus 1.13 (0.41–3.37) ␮g/g creatinine, P < 0.0001). VS showed a strong significant correlation with VU (r = 0.618, P < 0.0001). No correlation between

vanadium concentrations and years of employment could be demonstrated. Cytogenetic parameters and 8-OHdG in leukocyte DNA of exposed and non-exposed subjects are presented in Table 2. No significant differences in comet assay values, 8-OHdG or SCE between exposed and non-exposed probands could be found, not even when the different smoking habits were considered. Workers who claimed a green or black discoloring of the tongue did not show significantly elevated levels of vanadium in serum or urine or deviations of cytogenetic parameters or 8-OHdG as compared to workers without this symptom. A significant difference in SCE values between smokers (8.62±0.93) and non-smokers (7.67± 0.76) could be revealed in vanadium-exposed workers (P = 0.025). Ex-smoking workers (8.28 ± 0.84) showed higher SCE levels than non-smoking workers and lower levels than smokers, but these differences were not statistically significant. No correlation

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Table 2 Mean values and medians of genotoxic parameters exposed vs. non-exposed Non-exposed

Exposed

Pa

Vanadium in serum (␮g/l) Median (range)

3.43 ± 3.19 2.54 (1.01–12.5)

7.73 ± 7.86 5.38 (2.18–46.3)

0.005

Vanadium in urine (␮g/g creatinine) Median (range)

1.13 ± 0.26 0.74 (0.41–3.37)

14.57 ± 13.81 11.25 (2.11–95.2)

0.0001

Comet tailfactors (%) Median (range)

4.5 ± 1 4.5 (3.3–6.9)

3.8 ± 1.5 3.2 (2.6–9.3)

nsb

8-OHdG/105 dG Median (range)

0.72 ± 0.70 0.6 (0.17–2.58)

1.05 ± 1 0.7 (0.37–6.28)

ns

SCE/30 metaphases Median (range)

7.5 ± 1.1 7.8 (4.9–8.6)

8.3 ± 0.9 8.2 (6.2–10.8)

ns

a b

Significance between exposed and non-exposed probands. Not significant.

between cigarette consumption or years of smoking and SCE values could be demonstrated. 3.2. Effects of vanadate in vitro In the presence of vanadate (0.5–10 ␮M, which equals 25–500 ␮g/l), whole blood cells or isolated non-proliferating lymphocytes, exhibited a signif-

icant increase in DNA migration in the alkaline comet assay only at the highest vanadate concentrations (250–500 ␮g/l). Vanadate concentrations between 0.5 and 1 ␮M (25–50 ␮g/l), which represent vanadium concentrations observed in vivo, did not significantly elevate the comet tailfactor. In contrast, cultured human fibroblasts revealed a reproducible dose-dependent increase in alkaline comet

Fig. 2. Comet tailfactors in different types of cells after treatment with vanadate (0.5–10 ␮M, 24 h).

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Fig. 3. Alkaline and neutral comet assay of human fibroblasts after exposure to UV, bleomycin, vanadate, vanadate + UV and vanadate + bleomycin.

tailfactors after vanadate treatment (r = 0.9713, Fig. 2). Vanadate showed low, if any, cytotoxicity. Cell viability was approximately 90% by trypan blue exclusion even at the highest vanadate concentration. Vanadate did not increase 8-OHdG formation in cultured fibroblasts (data not shown), indicating that oxidative mechanisms are not responsible for the in vitro observed vanadate related DNA damage. For co-exposure tests a concentration of 0.5 ␮M (25 ␮g/l) of vanadate was chosen, because this was comparable to serum vanadium concentrations measured in occupationally exposed workers (in vivo data). In preliminary experiments we determined a bleomycin dose of 1 ␮g/ml and a UV flux density of 4.8 kJ/m2 to be sufficient for the induction of comet tailfactors that are found at 0.5 ␮M vanadate treatment. This was used to compare repair kinetics. Cells solely exposed to UV showed a three-fold elevation of the alkaline comet tailfactors as compared to controls. After 20 min, UV or vanadate induced damage was completely repaired and had reached baseline

levels, whereas bleomycin induced damage was only partially removed (30%). Combined treatment (vanadate + UV, vanadate + bleomycin) showed synergistic effects with respect to the formation DNA strand breaks. Only 30% of vanadate + UV produced DNA damage could be repaired, whereas vanadate+bleomycin induced lesions could not be repaired at all (Fig. 3). In neutral comet assay analysis, UV exposed as well as vanadate-treated cells revealed no differences in comet tailfactors as compared to control cells, indicating that no double strand breaks (DSBs) were induced. In contrast, incubation with either bleomycin or vanadate/UV or vanadate/bleomycin resulted in generation of non-repairable DSBs (Fig. 3).

4. Discussion In the present study, vanadate has been shown to induce DNA strand breaks in human fibroblast cells in a dose-dependent manner. Whole blood cells and

S. Ivancsits et al. / Mutation Research 519 (2002) 25–35

isolated lymphocytes were found to be less sensitive than fibroblasts with respect to vanadium generated effects observed in the alkaline comet assay. Occupationally exposed workers showed serum concentrations of vanadium up to 46 ␮g/l, but this was not associated with an increase in the comet tailfactor, frequency of sister chromatid exchanges or the level of oxidative DNA damage. The lack of cytogenetic consequences of occupational exposure to vanadium in whole blood leukocytes might be explained by the results of our in vitro tests. Exposure of whole blood or isolated lymphocytes to vanadate up to 1 ␮M (50 ␮g/l), which was the range of serum vanadium levels of the examined workers, did not lead to an increase in DNA strand breaks. This is consistent with the absence of effects in the comet assay in vivo. We found a significant increase in the comet tailfactor in whole blood cells and in lymphocytes at vanadate concentrations >5 ␮M, which is in accordance with results by Rojas et al. [25], who showed a dose-dependent response of effects observed in the alkaline comet assay in whole blood leukocytes treated with vanadium pentoxide (0.3–3000 ␮M). The vanadate-induced effects in the alkaline comet assay were 5–10-fold higher in cultured fibroblasts than in lymphocytes. This is possibly due to the fact that in contrast to unstimulated lymphocytes, fibroblasts are proliferating. Proliferating cells might be preferentially damaged and there is evidence that proliferating lymphocytes show a higher amount of DNA fragmentation in the comet assay than quiescent lymphocytes [38]. Differences in chromatin structure and cell cycle profile may also influence DNA migration in the comet assay. Moreover, cell sensitivity to vanadate and, perhaps, mechanism of action might differ depending on cell type. Altamirano-Lozano et al. showed that intraperitonal injection of vanadium pentoxide solutions in mice resulted in organ specific differences of DNA damage, being highest in liver, heart and kiney [23,24]. Beyond that, Foresti et al. [39] reported that meta-vanadate could damage the ability of mouse erythroleukemia cells to undergo erythroid differentiation, which appeared to depend on the cell cycle-related efficiency of DNA repair systems. Taken together, the absence of genotoxic effects in leukocytes of vanadium pentoxide-exposed workers examined in our study does not exclude possible vanadium induced DNA alterations in other cell types of these probands.

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The alkaline comet assay is a sensitive method for evaluation of DNA single and DSBs and alkali labile sites in cellular DNA. In combination with the neutral comet assay it is possible to discriminate between induced SSBs and DSBs of DNA, and therefore to uncover possible mechanisms of genotoxic actions. Vanadate-induced DNA damage was rapidly repaired (5–10 min) in vitro and the tailfactors remained unaffected in the neutral comet assay. From this we conclude that predominately DNA SSBs were produced, which is in accordance with data of Rojas et al. [25]. Vanadate (VO4 3− ) is a phosphate analogue and thus may interfere with phosphate containing enzymes, being involved in DNA repair mechanisms. Therefore, we tested the impact of vanadate on DNA repair kinetics of UV (physically genotoxic exposure) and bleomycin (radiomimetic exposure) treated fibroblasts. We observed a significant elevation of DNA migration in the alkaline comet assay accompanied by the occurence of persistent DSB after exposure to vanadate in combination with UV-light or bleomycin, as compared to vanadate treatment alone. This indicates that vanadate is genotoxic per se and may also act as an indirect genotoxic agent by converting repairable SSBs into non-repairable DSBs. This interaction may take place at unwinded unprotected DNA strands during the DNA repair process. Vanadate may therefore intensify DNA damage from additional exposures to genotoxic agents. The interindividual variability of vanadium-induced effects in the comet assay was not evaluated in this study. Therefore, differences in sensitivity to vanadium between individual cell lines or donors remain to be established. However, the lack of correlation between vanadium concentrations in serum or urine and the determined comet tailfactors does not support an influence of vanadium on DNA damage in leukocytes at the reported range of internal exposure. This is accentuated by the absence of correlation among all endpoints examined. In vanadium-exposed workers, smokers showed a significant increase in SCE compared to non-smokers, which is in good accordance with other studies reporting the effect of tobacco smoking on incidences of SCE [40,41]. As the control group (8 non-smokers, 3 smokers, 1 ex-smoker) and exposed group (13 non-smokers, 27 smokers, 9 ex-smokers) were not matched with respect to smoking habits and

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