Mutagenesis, Vol. 17, No. 5, 411-417,
September 2002
© 2002 UK Environmental Mutagen Society/Oxford University Press
Cytogenetic and oxidative damage induced in human lymphocytes by platinum, rhodium and palladium compounds
Dipartimento di Scienze dell’Uomo e dell’Ambiente and 1 Dipartimento di Morfologia Umana e Biologia Applicata, University of Pisa, via S. Giuseppe 22, 56126 Pisa, Italy and 2 European Commission, JRC-Ispra, Institute for Health and Consumer Protection (IHCP), ECVAM Unit, 21020-Ispra, Varese, Italy
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Abstract |
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This study of soluble compounds of platinum, palladium and rhodium investigated the genotoxic properties of (NH4)2PtCl4, PtCl2, PtCl4, (NH4)2PdCl4, PdCl2 and RhCl3 using the human lymphocyte micronucleus (MN) assay coupled with fluorescence in situ hybridization (FISH). A pancentromeric DNA probe was used to detect both centromere-positive micronuclei (C+ MN) as well as centromere-negative micronuclei (C– MN). A modified alkaline single cell gel electrophoresis (SCGE) assay was used to evaluate the possible role of oxidative damage in genotoxicity of the Pt, Pd and Rh compounds tested. Two enzymes, endonuclease III and formamidopyrimidine glycosylase, were used to recognize and subsequently cut oxidized pyrimidines and purines, respectively. A significant induction of MN by Pt and Rh compounds was observed compared with controls, while (NH4)2PdCl4 and PdCl2 displayed weak significant MN induction. The FISH technique revealed no significant difference in the frequency of C+ MN and C– MN for all compounds tested. These findings suggest that MN induction is due both to a clastogenic and an aneuploidogenic mechanism. SCGE detected an increase in the level of DNA oxidative damage for the Rh compound and for Pt(IV) which was also capable of inducing an increase in primary DNA damage at all the tested doses. This work highlights the stronger genotoxicity, likely mediated by oxidative damage induction, of Pt and Rh compounds compared with Pd salts.
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Introduction |
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This study is a part of a project aimed to screen metal compounds for genotoxicity by micronucleus (MN) assay. In particular, the aims of this investigation were: (i) to assess the genotoxic properties of some Pt, Pd and Rh metal salts using the human lymphocyte micronucleus assay (HLMA) and fluorescence in situ hybridization (FISH) technique; (ii) to test the possible role of oxidative damage in genotoxicity of Pt, Pd and Rh salts as the toxicity of some metals is also due to oxidative damage induction by production of O2 reactive species (Stohs and Bagchi, 1995
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The FISH technique has been applied to those compounds that gave clearly positive results with the HLMA, in order to detect both centromere-negative micronuclei (C– MN) due to chromosome breakage as well as centromere-positive micronuclei (C+ MN) due to malsegregation phenomena. This approach has already been employed to study the cytogenetic damage induced by some Al, Cd, Hg, Sb, Te and Tl compounds in human lymphocytes (Migliore et al., 1999). To study the oxidative damage, a modified alkaline version of the SCGE, optimized by Collins et al. (1995, 1997, 1998), using enzymes able to recognize and cut oxidized pyrimidines and purines, respectively, was applied.
There is a growing interest in the assessment of health risks posed by increased human exposure to some platinum group metals (PGM), namely platinum, rhodium and palladium. In spite of their low natural occurrence in the Earth’s crust (0.05, 0.001 and 0.0004 p.p.m. for Pt, Pd and Rh, respectively) (König and Schuster, 1994) these metals have important applications in high technology processes related to their catalytic properties and resistance to oxidation and corrosion (König and Schuster, 1994).
Of environmental significance is the fact that these metals are the three active components of automotive catalytic converters. In this context, recent work has shown the presence of 317 ng/g Pt and 74 ng/g Rh in road dust (Gómez et al., 2001), confirming that these metals can be released into the environment, representing ‘new’ potential environmental pollutants (Pietra et al., 1994). In addition, there is also growing interest in assessing exposure to PGM from an occupational and a medical angle.
Pure PGM and their alloys are extensively used as materials for melting tubes, in laboratory instruments, in jewellery and in spinning jets in synthetic fibre production. In medicine, while Pt and Pd are components of precious and semi-precious alloys used in dental restoration (Wataha and Hanks, 1996), Pt compounds are used as chemotherapeutic drugs as a part of standard treatments.
Despite their increasing use, the health effects of Pt, Pd and Rh remain little known. No information is available to date on long-term low level exposure to Pt or its compounds, thus preventing prediction of the environmental and health impacts of Pt emitted from mobile sources (Pietra et al., 1994).
The well-known effects of Pt and its compounds are those due to chronic industrial exposure to soluble salts of Pt, which can lead to sensitization, with further induction of conjunctivitis, asthma and contact dermatitis (platinosis). In contrast, Pt has been reported as being non-toxic and non-allergenic in its metallic state (Renner and Schmuckler, 1991). Knowledge concerning the genotoxicity of Pt, Pd and Rh compounds is fairly good for Pt, mainly in relation to Pt complex compounds used as antitumor agents (IARC, 1987), while little or no information is available for Pd and Rh compounds. Inorganic Pt compounds but not Pt salts are mutagenic in bacteria, Drosophila melanogaster and mammalian cells (Richardson and Gangotti, 1994). Mutations and cell death can be introduced by binding of Pt to DNA, which prevents cell replication (Choudhury et al., 2000). The genotoxicity of Pd salts seems to be low in bacteria and mammalian cells (Gebel et al., 1997).
In spite of this evidence, scientific information on the mutagenicity and carcinogenic potential of PGM in mammals is poor, making the assessment of health risk arising from exposure to PGM compounds impossible.
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Materials and methods |
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Chemicals
Metal salts used were (NH4)2PtCl4, PtCl2, PtCl4, (NH4)2PdCl4, PdCl2 and RhCl3 (Alfachem, Cologno Monzese, Italy). Unless otherwise indicated, all other chemicals and reagents were obtained from Sigma (St Louis, MO).
In order to assess metal contamination due to the addition of Pt, Pd or Rh compounds to culture medium, 33 metal impurities were determined in the 10–2 M mother solutions of such compounds by inductively coupled plasma mass spectrometry (model ELAN 6000; Perkin-Elmer/Sciex, Tornhill, Canada) or graphite furnace atomic absorption spectroscopy (model SIMAA 6000; Perkin-Elmer, Norwalk, CT) (Minoia et al., 1990).
The analysis of 33 metal impurities in the Pt, Pd and Rh solutions used in our assays showed that 22 elements (Al, Au, Ba, Bi, Cd, Ce, Co, Cs, Cu, Ge, Ir, La, Nb, Pb, Pd, Rh, Se, Sn, Sr, Th, U and W) were in concentrations ranging from 1.3 (Al) to 0.0007 µg/l (Cs). Another 11 elements (Ag, As, Be, Ga, Hg, Mn, Mo, Sb, Te, Tl and Zn) were below the detection limit, in any case <1 µg/l (results not shown).
Metal salts were dissolved in bi-distilled water at a concentration of 10–3 M, aliquoted and stored at 4°C until cell treatment. In the MN assay mitomycin C (MMC) was used as a positive standard clastogenic mutagen at a dose of 0.51 µM, while for FISH analysis we also employed griseofulvin (GF) at a dose of 43 µM, because of its known aneuploidogenic activity. In both cases water was used as the negative control.
For each metal compound tested three doses at which a significant increase in MN was obtained were selected for the Comet assay. Hydrogen peroxide was used as a positive control for the Comet assay at doses of 50, 100, 150 and 200 µM.
Cell culture and chemical treatment
Cytogenetic analyses were carried out on whole blood obtained from a young, healthy, non-smoking, male donor. Lymphocytes were stimulated with phytohemagglutinin (PHA) and cultured in a standard fashion for 72 h for the MN assay. Briefly, heparinized peripheral blood samples were cultured in duplicate at 37°C for up to 72 h in RPMI 1640 (Gibco BRL, Milan, Italy) supplemented with 20% fetal bovine serum (Gibco BRL), 1.5% PHA (Gibco BRL) and 1% penicillin/streptomycin (Gibco BRL). Each culture was performed in duplicate. Treatments with metals were performed 24 h after PHA stimulation. For each metal salt a wider range of doses was exploited, starting from an ineffective dose to a toxic one. Final experiments comprised at least three doses, with the highest concentration showing a significant reduction in the proportion of binucleate cells in the cultures. To block the cytokinesis of interphase cells, cytochalasin B (final concentration 6 µg/ml) was added at 44 h to all tubes.
Binucleated lymphocytes were harvested after 72 h culture; the treatment time was 48 h. Cells were treated with 0.075 M KCl to lyse erythrocytes, prefixed in methanol, washed twice with fixative (methanol:acetic acid 5:1) and then dropped onto clean glass slides. The air-dried slides were stained in 4% Giemsa solution or hybridized within 1 week of preparation.
FISH analysis
For FISH analysis a digoxigenin-labeled 
-satellite DNA probe specific for the centromeres of all human chromosomes (Oncor) was used. Prewarmed slides were denatured in 70% formamide, 2x SSC (saline sodium citrate buffer), pH 7.0, at 70°C for 2 min, followed by dehydration via an ethanol series. After heating at 70°C for 5 min, probes were placed on the denatured slide and incubated overnight at 37°C in a moist chamber. Post-hybridization washes were performed, first in a solution of 2x SSC, pH 7.0, at 72°C for 5 min and then in 4x SSC, 0.05% Tween 20 (SSCW) for 5 min at room temperature. To minimize the background, slides were preincubated for 10 min at 37°C in 4x SSC, with 5% non-fat dry milk as immunological buffer (IB). For detection of the digoxigenin-labeled probe, anti-digoxigenin (Boehringer, Italy), TRITC-conjugated anti-mouse and TRITC-conjugated anti-rabbit antibodies were diluted in IB and alternately incubated for 30 min at 37°C. Each incubation step was followed by three 2 min washes in SSCW at 37°C. After dehydrating through an ethanol series, slides were counterstained with 0.5 µg/ml DAPI dissolved in glycerol/DABCO antifade solution.
Slide scoring and statistical analysis
Giemsa stained slides were scored blind for MN analysis under an optical microscope (final magnification 400x). The criteria for MN acceptance listed by Fenech (1993) were followed. The results are expressed as the average number of micronucleated cells ± SD from two observations of 1000 cells on two different slides from two culture tubes (according to Migliore et al., 1999). MN frequency was expressed as the number of micronucleated binucleate cells (MNBN, containing one or more MN) per 1000 cells. The ratio of percent binucleate to mononucleate cells was used as a parameter of cell proliferation in culture. For FISH analysis, preparations were analyzed on a fluorescence microscope equipped with a triple bandpass filter for simultaneous visualization of TRITC, FITC and DAPI fluorescence. A sufficient number of lymphocytes were scored in order to record 50 MN for each experimental point. MN were analyzed for the presence of the fluorescent signal by considering a TRITC-labeled MN as centromere-positive MN (C+ MN) and a non-labeled MN as centromere-negative MN (C– MN). Data were analyzed by Fisher’s exact test to determine the significant difference between each treatment and the control for MNBN. Fisher’s exact test was also used to analyze the results obtained with the FISH technique: percentage of C+ MN (or C– MN) in treated cultures versus percentage of C+ MN (or C– MN) in control cultures.
Comet assay (SCGE)
Human leukocytes were isolated using the procedure described by Green et al.(1992). Whole blood (0.5 ml) was centrifuged twice in lysing buffer (Na4Cl, KHCO3, Na2EDTA) and resuspended in RPMI 1640 medium. The SCGE assay was performed basically according to Singh et al.(1988), with some modifications (Klaude et al., 1996).
Following their isolation, the cells were mixed with 0.4% trypan blue solution and, after 15 min, they were counted and checked for viability.
Leukocytes resuspended in RPMI 1640 were treated with the test substances for 2 h at 37°C immediately before the start of the assay, as suggested by Hartmann and Speit (1994). The 2 h treatment makes sure that DNA primary damage cannot be hidden by excision repair, as human leukocytes require 4 h to repair strand breaks (Collins et al., 1995).
One day prior to the analyses, 1% normal melting point agarose was spread on conventional slides and left to dry. On the following day, an 85 µl layer of 0.5% low melting point agarose (LMA) together with 3x105 cells (10 µl cell suspension + 75 µl LMA) were added to the slide surface. Finally, the slide was covered with a third layer of 85 µl of LMA. Slides were immersed in ice-cold freshly prepared lysing solution (2.5 M NaCl, 100 mM Na2EDTA, 10 mM Tris–HCl, 1% Triton X-100 and 10% DMSO, pH 10) to lyse the cells and allow DNA unfolding. After at least 1 h at 4°C in the dark, slides were placed on a horizontal electrophoresis unit. The unit was filled with fresh buffer (1 mM Na2EDTA, 300 mM NaOH, pH > 13) to cover the slides. The slides were allowed to set in the high pH buffer for 20 min to allow DNA unwinding and expression of alkali-labile sites. Electrophoresis was conducted for 20 min at 25 V (300 mA). Slides were then gently washed in neutralization buffer (0.4 M Tris–HCl, pH 7.5) to remove alkali and stained with 100 µl ethidium bromide (2 µg/ml). All steps described above were conducted under yellow light to prevent additional DNA damage.
Low and normal melting point agarose, Triton X-100, Na2EDTA, Tris–HCl, PBS and ethidium bromide were obtained from Sigma (St Louis, MO).
Enzyme preparation and treatment
In this study endonuclease III (endo III) was used to detect oxidized pyrimidines, and formamidopyrimidine glycosylase (fpg) to detect damaged purines, including 8-oxoguanine. The enzymes were kindly provided by Dr A.R. Collins (Aberdeen, UK).
Enzymes were diluted (2 µl of enzyme in 2 ml) and stored at –80°C in enzyme buffer (EB) (Na2 EDTA, 0.1 M KCl, 0.2 mg/ml BSA, 40 mM HEPES, pH 8). For fpg, 2 µl of enzyme were diluted in 200 µl of buffer containing 10% glycerol. Aliquots of 50 µl of enzyme solution (or buffer alone, as a control) were placed on gels and covered with 22x22 mm coverslips. Slides were put into a moist box, in order to prevent desiccation, and incubated at 37°C for 45 (endo III) or 30 min (fpg). The enzymes recognize and cut oxidized bases, converting these lesions into DNA single-strand breaks (SSBs) and producing fragments which migrate towards the anode during electrophoresis, making up the tail of the comet.
DNA migration assessment and statistical analysis
Images of 100 randomly selected cells (50 cells from each of the two replicate slides) were analyzed from each sample under a fluorescence microscope (200x) using a calibration scale considering two variables: nucleus diameter and comet length, which included the nucleus diameter plus tail length. Two independent cultures were performed per experimental point, for a total of 100 cells, and the mean was calculated.
The effects of dose, culture and experiment were evaluated by multifactor analysis of variance (MANOVA). The multiple range test was performed (P < 0.05) in order to detect differences in DNA migration among doses.
One hundred cells per point are shown as box and whisker plots, where the y-axis displays the range of DNA migration values. Each box encloses 50% of the data, with the median value of the variable displayed as a line. The top and bottom of the plot mark the ±25% limits of the variable. The lines extending from the top and bottom of each box mark the minimum and the maximum values falling within an acceptable range. Any value outside this range is displayed as an individual point. Genotoxicity was assessed in the sub-toxic range (relative viability >80%) to prevent potential artifacts due to indirect toxicity-induced DNA damage.
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Results |
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Table I
and Figure 1 shows the results for MN frequency after exposure of cells to different concentrations of Pt, Pd and Rh compounds. A significant induction of MN for each Pt compound and for the Rh salt as compared with the control (Fisher’s exact test) was observed. In particular, PtCl4 shows a significant (P < 0.001) induction of MN at a dose of 25 µM, while (NH4)2PtCl4 and PtCl2 are weaker genotoxic inducers, producing about the same effect on MN frequency at doses of 100 and 250 µM, respectively. Regarding RhCl3, a statistically significant (P < 0.001) increase in MN frequency was observed above a dose of 100 µM, not accompanied by a significant decrease in binucleated cell percentage. The two Pd salts showed quite weak but significant MN induction only at high doses [500 µM for (NH4)2PdCl4, and 100, 200 and 300 µM for PdCl2]. Data concerning FISH, applied only for those compounds clearly positive in the MN assay and at the doses that most significantly increased MN frequency, do not show any significant difference in the frequency of C+ and C– MN for all the compounds tested (Table II), even though for (NH4)2PtCl4, RhCl3 and PdCl2 a slight prevalence of C– MN was observed.
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Data concerning SCGE show oxidative DNA damage for PtCl4 and RhCl3. All other metal compounds tested were unable to induce this effect (Table III
). A significant increase in the amount of DNA migration after treating human leukocytes with PtCl4, above a dose of 25 µM, was observed (Figure 2A). A parallel statistically significant increase in oxidized purines and pyrimidines was found at the same doses (Figure 2B and C), indicating that the genotoxic potential of PtCl4 acts via an oxidative mechanism under our experimental conditions. All other Pt salts and PdCl2 gave negative or inconclusive results. RhCl3 was ineffective in inducing any increase in DNA migration at the doses tested. In contrast, a statistically significant decrease was obtained at doses of 100 and 250 µM (Figure 3A). RhCl3 induced a statistically significant increase in oxidized bases, both pyrimidines and purines, at all exposure doses (Figure 3B and C).
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A dose–effect response above the dose of 50 µM and an increase in oxidized pyrimidines and purines were found for hydrogen peroxide, a positive control known as an inducer of oxidative damage.
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Discussion |
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The data presented here provide evidence that the compounds of Pt, Pd and Rh tested are able to induce significant cytogenetic damage. Furthermore, the lack of any significant difference in the percentages of C+ MN and C– MN compared with the control, as shown by FISH analysis, indicates that all the compounds studied can act by both a clastogenic and an aneuploidogenic mechanism.
The genotoxic potential of PGM salts tested was assessed taking into account the aspect of speciation, because it is known that the toxicological effects of individual metals depend on their chemical forms (Sabbioni et al., 1985). In particular, it is known that the genotoxicity of Pt is determined by oxidation state, conformation and structure (Gebel et al., 1997). For this, Pt compounds with two different oxidation states (+2 as anionic PtCl42– or cationic Pt2+ species and +4 as the cationic Pt4+ form) and Pd salts (+2 as cationic Pd2+ and anionic PdCl42–) were tested. In addition, in order to compare the genotoxicity results, the Pt and Pd salts analyzed had the corresponding anion/cation.
All three Pt compounds tested [(NH4)2PtCl4, PtCl2 and PtCl4] were found to produce a significant increase in MN frequency compared with the control. PtCl4, which was found to induce forward mutation in V-79 cells at a rate ~7 times the control rate (Kanematsu et al., 1990), seems to exert a stronger action in terms of both cytotoxicity and MN induction compared with those of the divalent Pt salts. In fact, even at the dose of 25 µM PtCl4 induced an ~6-fold increase in MN. These findings are in agreement with the data of Gebel et al.(1997), who showed that PtCl4 induces higher frequencies of MN in human lymphocytes when compared with PtCl2. This was confirmed by SCGE, which gave negative results with (NH4)2PtCl4 and PtCl2 while PtCl4 showed a significant increase in DNA migration above a dose of 25 µM. (NH4)2PtCl4 showed a significant increase in DNA migration and oxidized purines at the only dose of 150 µM, but we consider a positive response the presence of at least two other doses able to induce a statistical effect. We cannot exclude that other mechanisms of action not detectable by SCGE, such as crosslink formation, could take place at the tested doses of Pt2+ compounds. This could lead to the formation of bulky DNA fragments unable to migrate in the electrophoresis gel under the experimental conditions used (Tice et al., 1992; Hartman and Speit, 1994). PtCl2 exhibited mutagenicity and high toxicity for strains TA98 and TA100 in the Ames test (Uno and Morita, 1993). In contrast, Pt4+ is able to induce DNA damage via both a direct action and an oxidative mechanism.
On the other hand, metal carcinogenicity is known to be mediated by a variety of different mechanisms: direct mutagenic attack on DNA is a possible primary mode of action, as also is the induction of oxidative stress or cellular immunity, inhibition of DNA metabolism and DNA repair and the formation of DNA and/or protein crosslinks (Snow, 1992).
Concerning the Pd compounds studied, both of them appear to be weak inducers of cytogenetic damage. However, PdCl2 is unable to give rise to any increase in DNA migration at the doses tested. A decrease in DNA migration was observed at 100 µM in the absence of endo III (primary DNA damage), as well as at doses of 100, 200 and 300 µM in the presence of endo III. As already stated, some metals can produce crosslinks (Snow, 1992), and mutagenic and/or carcinogenic compounds characterized by a crosslinking action are known to be poor inducers of DNA migration (Tice et al., 1992; Hartman and Speit, 1994; Frenzilli et al., 2000). Some of them, such as active aldehydes, are even able to induce a reduction in DNA migration at high doses (Frenzilli et al., 2000). It is thus possible that crosslink formation might be partially responsible for the lack of an effect obtained with PdCl2. The decreased DNA migration observed in the presence of endoIII at all doses might also represent the result of an enzyme–metal interaction that may interfere with the action of the enzyme itself on spontaneously oxidized pyrimidines, whose presence is demonstrated by control strand break levels.
Bunger et al. (1996) showed both a cyto- and a genotoxic activity for some Pt, Rh and Pd metal salts. In particular, they found that spontaneous mutation rates increased by a factor of 3–20 when the four strains tested were exposed to Pt complexes, as assayed by the Ames test. The Rh compounds proved to be considerably less mutagenic, while Pd showed a non-mutagenic potential, in agreement with Uno and Morita (1993). The reason for the weak Pd genotoxicity might be that inorganic Pd2+ complexes seem to be more labile than their corresponding Pt2+ counterparts (Gebel et al., 1997).
The MN assay coupled with FISH analysis revealed that RhCl3 can act as both a clastogen and aneuploidogen in the dose range 100–1000 µM. In a recent study Sadiq et al. (2000) showed that a Rh3+ complex induced chromosomal aberrations of all types in cultured human lymphocytes, exerting its clastogenic effects without the need for metabolic activation, in a radiomimetic S phase-independent way. The mutagenicity of RhCl3 was studied in V-79 cells. At 300 µM a 4-fold mutation rate compared with the control was observed (Kanematsu et al., 1990). The same compound administered in drinking water to mice at concentrations of 5 mg/l led to the development of lymphomas and leukemia (Schäfer et al., 1999).
The mutagenic potential of Pt and Rh complexes appears to be based on a variety of mechanisms that damage DNA (Bunger et al., 1996). Our current knowledge on mechanisms of metal genotoxicity suggests that there is no unique mechanism which could account for the genotoxic potential of all the diverse metals and their compounds, although the two main actions seem to be enhanced formation of reactive oxygen species, resulting in lipid peroxidation and DNA damage, and interference with DNA repair and/or DNA replication processes (Hartwig, 1995).
The present work shows the capability of Pt, Pd and Rh compounds to act by different mechanisms. In this context, the aneuploidogenic effect was proved by FISH assay and the clastogenic effect by both FISH and SCGE assays, while possible crosslink formation was drawn from SCGE data.
Concerning the occurrence of oxidative damage, PtCl4 displays an increase in oxidized purines, associated with a clastogenic effect that was revealed in the absence of enzymes, as shown by SCGE. In the case of RhCl3, oxidative damage seemed to be prevalent. However, the decrease in DNA migration observed at doses of 100 and 250 µM in the absence of enzymes suggests that a possible alkylating mechanism could occur. We can hypothesize that at the basal level oxidative damage is likely hidden by crosslink formation, but it is highlighted by the use of endoIII and fpg. On the other hand, Pd compounds exert only weak effects when measured with the MN assay and no significant effect with the other methods employed, so we cannot make conclusive statements about their possible mechanism of action.
We can conclude that a better understanding of metal genotoxicity requires the simultaneous use of a wide spectrum of tests in order to obtain evidence of the different modes of action following exposure to metal compounds.
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Acknowledgments |
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The authors are grateful to Leonardo Cocchi and Mariacarla Iorio for their excellent technical assistance. This work is a contribution to contract no. 15469-1999 11F1ED ISP ES.
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Notes |
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3 To whom correspondence should be addressed. Tel: +39 050 836223; Fax: +39 050 551290; Email: l.migliore{at}geog.unipi.it
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Received on January 30, 2002; accepted on May 20, 2002.
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