We used the SCGE assay to study DNA migration in whole-blood leukocytes from 52 vanadium production workers exposed to V₂O₅ by inhalation and from 52 nonexposed controls (jail wardens). Additionally, we analyzed lymphocytes of 24 workers and 23 controls using CBMN Cyt experiments. We collected data concerning age, weight, height, and smoking status with a questionnaire (Table 1).

Workers are exposed to vanadium dust during the entire shift (8 hr) and are required to wear protective masks and gloves. The study was approved by the Ethics Committee of the Medical University of Vienna. After obtaining informed consent, we collected blood (2 ×10 mL) in heparin and EDTA tubes (Vacutainer-Systems, Becton Dickinson, Plymouth, UK). We stored blood samples at 4°C and transported them to the Institute of Cancer Research at the Medical University of Vienna. Whole blood was centrifuged at 623 ×g for 10 min at 14°C (Sigma Laboratory Centrifuge 4K15; Sigma Chemical Co., St. Louis, MO, USA) and plasma was collected, aliquoted, and stored at −80°C. We conducted experiments between October 2004 and May 2005.

We determined vanadium concentrations in plasma using graphite furnace atomic absorption spectrometry, as described by Ivancsits et al. (2002), with Zeeman background correction at a temperature of 2,450°C. The calibration curves for vanadium were in the range of 0–40 μg/L, and the detection limit was 0.3 μg/μL (5.9 nmol/L).

We determined pyridoxal phosphate (the active form of vitamin B₆) in plasma by HPLC using commercial assays (Immundiagnostic AG, Bensheim, Germany) as described by Majchrzak et al. (2006). We measured vitamin B₁₂ and folate in plasma using commercial radioimmunoassays (IUL Instruments GmbH, Königswinter, Germany) (Majchrzak et al. 2006).

We used the protocol of Buschini et al. (2002) for leukocyte isolation: EDTA-anticoagulated blood was maintained in erythrocyte-lysis buffer (155 mM NH₄Cl, 5 mM KHCO₃, 0.005 mM Na₂EDTA, pH 7.4) at 37°C for 5 min; centrifuged (200 ×g, 5 min at 4°C); washed twice in phosphate-buffered saline (PBS; pH 7.4); and resuspended in RPMI 1640 (Sigma-Aldrich Chemie Gmbh, Munich, Germany) without serum (Buschini et al. 2002). We used a modified version of the protocol for BLEO sensitivity and DNA repair measurements (Schmezer et al. 2001). Leukocytes were treated with 10 μg/mL BLEO sulfate (Nippon Kayaku Co. Ltd., Tokyo, Japan) at 37°C for 30 min. To terminate the treatment, cells were washed twice in PBS (pH 7.4). To measure DNA repair capacity, we incubated a second batch of BLEO-treated cells for 15 min (37°C) before lysis and electrophoresis. Cell viability was monitored with trypan blue (Lindl and Bauer 1994). Viability of all samples was ≥ 90%.

We performed the SCGE assay according to the guidelines of Tice et al. (2000). Briefly, cell pellets were mixed with 60 μL 0.5% low-melting-point agarose (Invitrogen Life Technologies Ltd., Paisley, Scotland), transferred to precoated (1.5% normal-melting-point agarose) glass slides, and sealed with a coverslip. The slides were placed on ice for 5 min to allow solidification of the agarose. After removing the coverslip, we immersed the slides in lysis solution (2.5 M NaCl, 100 mM Na₂EDTA, 10 mM Tris, 1% Triton X, 10% dimethyl sulfoxide, pH 10.0) and kept them at 4°C for ≥ 1 hr. To prevent DNA damage, lysis and all subsequent steps were conducted under red light. After lysis, we placed the slides on a horizontal gel electrophoresis unit (C.B.S. Scientific, Solana Beach, CA, USA) and allowed the DNA to unwind at 4°C in alkaline electrophoresis buffer (300 mM NaOH, 1 mM Na₂EDTA, pH ≥ 12.5) for 20 min. Electrophoresis was performed at 25 V and 300 mA for 20 min; then the slides were covered with neutralization buffer (0.4 M Trizma base, pH 7.5, 4°C) for 2 ×8 min and air dried. The coded slides were stained with ethidium bromide (20 μg/mL) and evaluated under a fluorescence microscope (Nikon 027012; Nikon, Tokyo, Japan). We used an automated analysis system (Helma and Uhl 2000) to acquire images, compute the integrated intensity profile for each cell, estimate the comet cell components, and evaluate derived parameters. For each experimental end point we analyzed three cultures, and we measured 50 randomly captured cells from each slide. To quantify DNA damage, tail lengths and tail moments were determined.

We used a modified version of the protocol published by Collins et al. (1993) to measure endogenous formation of oxidized DNA bases. Formamidopyrimidine glycosylase (FPG) and endonuclease III (ENDO III) were provided by K. Angelis (Czech Academy of Sciences, Prague, Czech Republic).

After lysis of the cells, the slides were washed in enzyme buffer (0.1 M KCl, 40 mM HEPES, 0.5 mM Na₂EDTA, 0.2 mg/mL bovine serum albumin, 4°C, pH 8) for 2 ×8 min. Subsequently, agarose-embedded cells were covered with 50 μL enzyme buffer or with enzyme solutions (1.0 μg/mL). Gels were sealed with a coverslip and incubated at 37°C in the dark for 45 min (ENDO III) or 30 min (FPG). Subsequently, the slides were transferred into alkaline buffer for unwinding, and electrophoresis was performed.

We isolated lymphocytes according to the protocol of Fenech (2000). Briefly, Histopaque 1077 (Sigma) was overlaid with RPMI 1640–diluted blood and centrifuged at 318 ×g at 14°C for 30 min. Subsequently, the cell layer was removed, resuspended in RPMI, washed twice, and centrifuged (318 ×g, 14°C, 10 min).

We performed the CBMN Cyt test using the cytochalasin B technique described by Fenech (2007). We determined MNs, Nbuds, and NPBs, as well as apoptotic and necrotic cells, in samples from 24 workers and 23 controls using the cytome approach (Fenech 2007). Slides were evaluated by one of the authors (V.A.E.) who was trained in the laboratory of M.F. at Commonwealth Scientific and Industrial Research Organisation (Human Nutrition, Adelaide, South Australia), which has developed the CBMN Cyt assay and was centrally involved in the standardization of scoring criteria via the International Collaborative Project on Micronucleus Frequency in Human Populations (Human Micronucleus Project 2008). For each participant, we prepared lymphocyte cultures in duplicate. Each culture contained 10⁶ cells in 750 μL RPMI 1640 supplemented with 100 U/mL penicillin, 100 μg/mL streptomycin, 10% fetal bovine serum (Sigma), 2.0 mmol/L L-glutamine, and 30 μg/mL phytohemagglutinin (Invitrogen, Carlsbad, CA, USA). Cultures were incubated in round-bottom tubes (Becton Dickinson) at 37°C in a humidified atmosphere containing 5% CO₂. Forty-four hours after mitogen stimulation, we added cytochalasin B (final concentration, 4.0 μg/mL; Sigma-Aldrich, St. Louis, MO, USA) to block cell division. The total incubation time of the cultures was 72 hr. Subsequently, the cells were harvested and used to prepare slides, which were air dried, fixed, and stained with Diff Quick (Dade Behring, Deerfield, IL, USA). From each participant, we scored 2,000 binucleated cells using a light optical microscope (Microphot-FXA; Nikon) (Fenech et al. 2003).

We performed statistical analyses using Statistica 5.0 software (StatSoft Inc., Tulsa, OK, USA). The results are presented as medians and 25th–75th percentiles. All p-values were two-tailed, and we considered differences to be significant at p ≤ 0.05. We used the Mann-Whitney U-test for comparisons between exposed subjects and controls. We used Spearman’s correlation coefficients to test univariate relationships between different variables.

The characteristics of participants are summarized in Table 1. Age and body mass index did not differ between the groups. All individuals were males and nonvegetarians. The control group included 11 smokers and the exposed group included 12. The concentrations of folate were similar in both groups, whereas levels of vitamins B₆ and B₁₂ were higher (8.1% and 31%, respectively) in the exposed group, possibly due to intake of supplements. Vanadium levels were 7-fold higher in the plasma of the exposed group.

Table 2 summarizes the results of the SCGE experiments. We found no difference of DNA migration between exposed individuals and control subjects when the assay was carried out under standard conditions (which reflect formation of single- and double-strand breaks), but we observed increased tail lengths and tail moments in all other end points in the workers. Formation of oxidized purines (detected with FPG) was elevated by 7%, and the formation of pyrimidines (by ENDO III treatment) was enhanced by 33%. Furthermore, the sensitivity of the cells toward BLEO-induced DNA damage was higher (25%) in the exposed individuals. When we monitored DNA migration after a repair phase (15 min incubation after BLEO treatment), we observed a decrease of tail lengths (50%) and tail moments (57%) in the controls, whereas in the workers we observed no reduction of tail lengths and a less pronounced decrease of tail moments (30%).

Data on the frequencies of MNs, NPBs, Nbuds, apoptosis, and necrosis rates, as well as the division indices [nuclear divison index (NDI) and nuclear divison cyto-toxicity index (NDCI)] are summarized in Table 3. The number of MNs was 2.5-fold higher in the workers, and the numbers of NPBs and Nbuds were substantially increased (7-fold and 3-fold, respectively) .

To find out whether V₂O₅ exposure and smoking cause synergistic effects, we compared the frequencies of MNs and Nbuds in exposed smokers and nonsmokers to those in the corresponding controls. In nonexposed individuals, the levels (means ± SD) of MNs, Nbuds, and NPBs were similar in smokers and nonsmokers respectively (MNs, 1.77 ± 1.62 and 1.50 ± 1.86; Nbuds, 0.33 ± 0.39 and 0.36 ± 0.32; NPBs, 0.46 ± 0.81 and 0.45 ± 0.61). In workers, the MN rates were lower in smokers (4.17 ± 2.95 and 8.08 ± 4.54), whereas the frequencies of NPBs and Nbuds were more or less identical (NPBs, 6.83 ± 3.04 and 7.33 ± 2.50; Nbuds, 3.58 ± 2.76 and 3.30 ± 1.41).

The frequencies of necrotic and apoptotic cells were elevated in the workers by 55% and 50%, respectively; for the latter parameter the difference was not significant. The last two rows of Table 3 show the nuclear division indices. The division rates were similar in both groups, regardless of whether we considered the number of viable cells only (NDI) or also dead cells (NDCI). These results indicate that metal exposure has no significant impact on the proliferative capacity of viable or total lymphocytes, respectively.