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Corinne and Ferrier, Vincent and Devaux, Alain Comparative Evaluation of
Genotoxicity of Captan in Amphibian Larvae (Xenopus laevis and Pleurodeles
waltl) Using the Comet Assay and the Micronucleus Test. (2006) Environmental
Toxicology, vol. 21 (n°3). pp. 264-277. ISSN 1520-4081
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Comparative Evaluation of Genotoxicity of
Captan in Amphibian Larvae (Xenopus laevis and
Pleurodeles waltl) Using the Comet Assay and
the Micronucleus Test
F. Mouchet,1L. Gauthier,2C. Mailhes,3V. Ferrier,4A. Devaux5,6 1
Laboratoire d’Ecologie des Hydrosyste`mes—LEH, UMR CNRS UPS 5177, 29 Rue Jeanne Marvig, 31055 Toulouse, France
2
Laboratoire d’Ecologie des Hydrosyste`mes—LEH, UMR CNRS UPS 5177, Centre Universitaire de Formation et de Recherche Jean-Franc¸ois Champollion,
Campus d’Albi, place de Verdun, 81012 Albi cedex 9, France 3
IRIT–ENSEEIHT–Te´SA, 2 rue Camichel, BP 7122, F-31071 Toulouse cedex 7, France 4
Centre de Biologie du De´veloppement, Universite´ Paul Sabatier, CNRS–UMR 5547, 118 route de Narbonne, 31062 Toulouse, France
5
Laboratoire des Sciences de l’Environnement, Ecole Nationale des Travaux Publics de l’Etat, 69518 Vaulx-en-Velin, France
6
INRA, De´partement d’Hydrobiologie et Faune Sauvage, Campus de Beaulieu, 35042 Rennes cedex, France
ABSTRACT: Captan (N-trichloromethylthio-4-cyclohexene-1,2-dicarboximide) is a fungicide used to in-hibit the growth of many types of fungi on plants used as foodstuffs. The toxic and genotoxic potentials of captan were evaluated with the micronucleus test (MNT; AFNOR, 2000) and the comet assay (CA) using amphibian larvae (Xenopus laevis and Pleurodeles waltl). Acute toxicity results showed that captan was toxic (1) to Xenopus larvae exposed to from 2 mg/L to 125 or 62.5g/L, depending on the nature of the water [reconstituted water containing mineral salts or mineral water (MW; Volvic1, Danone, France)] and (2) to Pleurodeles exposed to from 2 mg/L to 125g/L in both types of water. The MNT results obtained in MW showed that captan (62.5g/L) was genotoxic to Xenopus but not genotoxic to Pleurodeles at all concentrations tested. CA established that the genotoxicity of captan to Xenopus and Pleurodeles larvae depended on the concentration, the exposure times, and the comet parameters (tail DNA, TEM, OTM, and TL). The CA and MNT results were compared for their ability to detect DNA damage at the concentra-tions of captan and the exposure times applied. CA showed captan to be genotoxic from the first day of exposure. In amphibians, CA appears to be a sensitive and suitable method for detecting genotoxicity such as that caused by captan.
Keywords: captan; amphibian; Xenopus laevis; Pleurodeles waltl; acute toxicity; genotoxicity; comet assay; micronucleus test
INTRODUCTION
The wide use of pesticides in agriculture has made them ubiquitous in the environment. The agronomic benefits of pesticides must be weighed against their potential effects on ecosystems and human health, an area of increasing con-cern in industrialized countries. Like all pollutants, pesti-cides tend to find their way into the aquatic compartment and can exhibit toxic and genotoxic activity in freshwater. Genotoxic pollutants can affect aquatic ecosystems directly, but in water they can also affect nonaquatic species via food chains or simply via drinking water.
Captan (133-06-2, N-trichloromethylthio-4-cyclohexene-1,2-dicarboximide, C9H8Cl3NO2S) is an organochlorine fungicide of the dicarboximide chemical family (U.S. EPA, 1999). Introduced in the early 1950s (Newsome et al., 1992), it is widely used as an antimicrobial pesticide to inhibit the growth of many types of fungi on plants used as foodstuffs (Fisher et al., 1992) and as a treatment for seed and ornamental sites (U.S. EPA, 1999). More than 160 registered products are reported to contain the active ingre-dient captan (U.S. EPA, 1999). Captan has been classified as a B2 (probable human) carcinogen (U.S. EPA, 1999).
The available information in the literature about the toxic and genotoxic effects of captan on aquatic species is more limited than is that about pesticides generally. The aim of the present work was to evaluate the acute toxicity and genotoxicity of captan in amphibian larvae in con-trolled laboratory conditions.
Among the numerous biomarkers of genotoxicity (DNA adducts, DNA strand breaks, chromosomal aberrations, sis-ter chromatid exchange, micronuclei, etc.) used to evaluate the genetic damage resulting from exposure to environmen-tal pollutants, the present testing for captan was carried out to evaluate two end points: induction of micronuclei (micronucleus test) and of primary DNA damage (comet assay) in amphibian models (Xenopus laevis and Pleuro-deles waltl larvae).
The micronucleus test (MNT) detects the induction of micronuclei, which are the consequence of chromosome fragmentation and/or malfunction of the mitotic apparatus. Thus, clastogenic compounds and spindle poisons both lead to an increased number of micronucleated cells. In aquatic vertebrates, the MNT has been carried out in different fish species (Das and Nanda, 1986; Metcalfe, 1988; Carrasco et al., 1990). Amphibians have also proved to be valuable biological models for the study of micronucleus induction in erythrocytes (Jaylet et al., 1986; Krauter et al., 1987; Jay-let and Zoll, 1990; Zoll et al., 1990; Gauthier, 1996; Ferrier et al., 1998; AFNOR, 2000; Djomo et al., 2000; Bekaert et al., 2002). The sensitivity and reliability of the MNT in detecting chromosomic/genomic mutations make it a good method to use in the analysis of the potential cytogenetic damage caused by pure substances (Grinfeld et al., 1986; Jaylet et al., 1986; Fernandez and Jaylet, 1987; Fernandez
et al., 1989; Fernandez et al., 1993; Gauthier, 1996; Mou-chet et al., 2005a).
The comet assay (CA), first described by O¨ stling and Johanson (1984) and further developed by Singh et al. (1988) and Olive et al. (1990), is a sensitive method widely used for the detection of DNA damage in individual cells (double- and single-strand breaks, alkali-labile sites, exci-sion repair sites, and crosslinks) induced by a variety of genotoxic agents such as industrial chemicals, biocides, agrochemicals, and pharmaceuticals (for general review articles, see McKelvey et al., 1993; Fairbairn et al., 1995; Anderson and Plewa, 1998; Rojas et al., 1999; Speit and Hartmann, 1999; Tice et al., 2000; Hartmann et al., 2003). Numerous studies have demonstrated its capacity to detect low levels of DNA damage, its requirement for few cells, its low cost, and its speed of execution and analysis for assessing the genotoxicity of chemicals toward fish (Bel-paeme et al., 1998; Devaux et al., 1998; Mitchelmore and Chipman, 1998; Larno et al., 2001; Flammarion et al., 2002; Winter et al., 2004), mussels (Nacci et al., 1996; Pavlica et al., 2001), and amphibians (Ralph et al., 1996; Clements et al., 1997; Ralph and Petras, 1998a, 1998b; Mouchet et al., 2005a, 2005b, 2006).
In the present work, the sensitivity of both tests was compared. The amphibiansXenopus laevis and Pleurodeles waltl were used because of their ecotoxicological rele-vance, which has already been well established in a large array of studies on biomarkers such as induction of bio-transformation enzyme activity (Bekaert et al., 2002; Gauthier et al., 2003), micronuclei (Ferrier et al., 1998), and DNA adduct formation (Bekaert et al., 2002), comet induction following DNA damage (Mouchet et al., 2005a, 2005b; Mouchet et al., 2006), and teratogenic malforma-tions (Chenon et al., 2002).
MATERIALS AND METHODS
Amphibian (Xenopus and Pleurodeles)
Maintenance and Breeding
Sexually mature Xenopus and Pleurodeles were provided by the Developmental Biology Department of Paul Sabatier University (Toulouse, France). To induce spawning, Xeno-pus males were injected with 400 IU and females with 700 IU of human chorionic gonadotropin. No hormonal induc-tion was needed forPleurodeles because of the natural lay-ing period. Amphibian males and females were then placed together in normal tap water filtered through active char-coal at 228C 6 28C. Twelve hours later for Xenopus, the pairs were separated, and viable eggs were maintained in an aquarium also containing normal tap water filtered through active charcoal at 208C–228C until they reached a development stage appropriate for experimentation (usually 3 weeks). For Pleurodeles, after a few days, viable eggs
were maintained in a bowl containing 5–6 L of normal tap water filtered through active charcoal at 188C until the swimming stage. They were then transferred to an aquar-ium containing the same type of water at 208C 6 18C until they reached a development stage appropriate for experi-mentation (about 6 weeks after laying). Both species were fed daily—Xenopus larvae on dehydrated aquarium fish food and youngPleurodeles larvae first on freshly hatched artemia (Artemia spec.), which was then switched to thawed chironoma (Chironomus) larvae. Each day one third of the water was renewed before feeding.
Exposure Conditions
The experimental exposure conditions, basically the same for the MNT and CA, are described in the French Standard AFNOR (French National Organization for Quality Regula-tion) NF T90-325 (AFNOR, 2000). The amphibian models were exposed to either reconstituted water [RW; normal tap water filtered through active charcoal to which nutritive salts (294 mg/L CaCl22H2O, 123.25 mg/L MgSO47H2O, 64.75 mg/L NaHCO3, 5.75 mg/L KCl) were added] or min-eral water (MW; 9.9 mg/L Ca2þ, 6.1 mg/L Mg2þ, 9.4 mg/L Na2þ, 5.7 mg/L Kþ, 8.4 mg/L Cl, 6.3 mg/L NO3, 6.9 mg/L SO42, 65.3 mg/L HCO32, 30.0 mg/L Si, pH 7; Vol-vic, Danone, France) at 228C 6 0.58C for Xenopus and 208C 6 18C for Pleurodeles. Exposure began on larvae at stage 50 of the Xenopus development table (Nieuwkoop and Faber, 1956), characterized by the hind limb bud being longer than broad and constricted at the base. For Pleuro-deles, exposure began at stage 53 of the development table (Galien and Durocher, 1957), when the hind limbs present four well-formed digits with an outline of the fifth. The lar-vae were taken from the same hatch to reduce interanimal variability. Control (negative and positive) and experimen-tal groups of 20 (100 mL/Larvae) were exposed in 5-L glass flasks to the control medium (negative and positive con-trols) or the test medium (different concentrations of cap-tan), respectively. The negative control was either RW or MW. Positive controls were systematically performed in each experiment in order to check that the responsiveness of the amphibian larvae and the experiment was correct. The positive control for the MNT was benzo[a]pyrene (B[a]P, [50-32-8], purity:96.0%, Sigma France) at 0.125 mg/L (AFNOR, 2000). For the CA, the positive control was methyl methanesulfonate (MMs, [66-27-3], purity 99%, Sigma France) at 1.56 mg/L (Mouchet et al., 2005a). Cap-tan and B[a]P were dissolved in dimethyl sulfoxide (DMSO, [67-68-5], Sigma France) at a final concentration of 0.05%—previously shown to be nongenotoxic and non-toxic toXenopus (data not published)—before being added to the water. The concentrations of captan were 2000, 1000, 500, 250, 125, 65.5, 31.25, and 12.25g/L. Amphib-ian larvae were submitted to a 12 h light:12 h dark cycle. Xenopus were fed every day on dehydrated aquarium fish
food and Pleurodeles on thawed chironoma larvae. The flasks were partially immersed in water baths to maintain the water temperature at 228C 6 0.58C for Xenopus and 208C 6 18C for Pleurodeles. The media in the control and exposed flasks were renewed daily.
Acute Toxicity
Acute toxicity to the larvae exposed to captan in RW and MW was examined for 12 days by visual inspection to determine death (severe toxicity) and abnormal behavior, reduced size, and diminished food intake (weak toxicity). Recording the data daily about toxicity in treated larvae in MW ensured that the genotoxicity tests were performed under conditions that were not acutely toxic.
Genotoxicity Assays
Genotoxicity was only assayed in MW. At the end of expo-sure (12 days for the MNT and 1, 2, 4, 8, or 12 days for the CA), a blood sample was obtained from each anaesthetized larva by cardiac puncture with heparinized micropipettes (20% solution at 5000 IU/mL). For a given test (MNT or CA), larvae from the same exposure group were used. Gen-otoxicity was assessed to the highest concentration that did not lead to signs of acute toxicity of the exposed larvae.
Micronucleus Test
The micronucleus test (MNT) was carried out according to the AFNOR Standard NF T90-325 procedure (AFNOR, 2000). At the end of the 12-day exposure, a single smear of blood was prepared from each animal. After fixing in meth-anol and staining with hematoxylin, the smears were screened under a microscope (oil immersion lens, 1500). The number of erythrocytes that contained one or more micronuclei was determined in a total sample of 1000 erythrocytes per larva. Slides were scored blind by only one individual.
For each group of animals, the results (number of micro-nucleated erythrocytes per thousand, MNE%) obtained for the individual larvae were arranged in increasing order of magnitude. The medians and quartiles were then calculated. The statistical method used to compare the medians was based on the recommendations of Mac Gill et al., (1978) and consists of determining the theoretical medians of sam-ples of sizen (where n 7) and their 95% confidence limits expressed byM6 1.57 IQR/ ffipn, where M is the median and IQR is the Inter-Quartile Range (upper quartile - lower quartile). Under these conditions, the difference between the theoretical medians of the test groups and the theoreti-cal median of the negative control group is significant to within 95% certainty if there is no overlap.
Comet Assay
The comet assay (CA) was performed according to the pro-cedure described by Singh et al. (1988) and subsequently adapted to Xenopus laevis (Mouchet et al., 2005a) and Pleurodeles waltl larvae. Frosted microscope slides were cleaned and precoated with freshly prepared normal melt-ing agarose [NMA; 0.8% in phosphate buffered-saline (PBS)], left at room temperature to allow the agarose to dry, and then kept overnight at 48C. After cardiac puncture, an aliquot of heparinized blood cell suspension was imme-diately diluted 50-fold in PBS. Erythrocyte viability was routinely determined using the Trypan blue exclusion test, and samples showing<90% viability were discarded (Col-lins, 2002). Diluted blood was mixed with an equal volume of fresh, low-melting-point agarose (LMA, 1% in PBS). Sixty-five milliliters of this agarose–cell suspension was spread on each precoated slide (2 per animal) and covered with a coverslip (22*50). After cooling for 7 min on ice, the coverslip was gently removed, and a third layer, consisting of 90L of LMA (1% in PBS), was added and allowed to solidify for 7 min on ice before gently removing the cover-slip. The slides were then immersed in freshly prepared ice-cold lysing solution (48C, 2.5M NaCl, 0.1M Na2EDTA, 0.01M Tris, 1% Triton X-100, and 10% DMSO, adjusted to pH 10 with NaOH) and left for 1 h. Lysis and subsequent steps were performed in the dark under a dim red light. The slides were then gently removed from the lysing solution and transferred to a horizontal electrophoresis tank contain-ing freshly prepared alkaline buffer [0.3M NaOH, 1 mM Na2EDTA (pH> 13)] for 20 min at 48C in order to allow the DNA to unwind. Electrophoresis was carried out in the same buffer at 48C for 20 min in Xenopus and for 30 min in Pleurodeles by applying an electric field of 20 V and adjusting the current to 300 mA. Finally, the slides were gently washed twice in a neutralization buffer [0.4M Tris-HCl (pH 7.5)] for 5 min before being dehydrated in abso-lute ethanol for 5 min. Slide analysis was performed using a confocal microscope (LSM 410 inverted laser scanning microscope, Zeiss) at 40 magnification, after staining the slides with 0.05 mM ethidium bromide solution.
Which specific parameter to use in comet image analysis has not been completely resolved yet. Quantitative assess-ment of DNA damage in erythrocyte nuclei was performed using Komet 4.1 image analysis software (Kinetic Imaging Ltd.) by measuring four parameters. Four parameters were chosen because of the great variability in representativeness of an individual parameter, among the 34 provided by the software depending on the molecule tested, its concentra-tion in the medium, and/or the exposure time. The parame-ters chosen were extent tail moment (ETM; defined as the tail length weighted by the percentage of tail DNA), olive tail moment (OTM; product of the amount of DNA in the tail and the mean distance of migration in the tail; Olive et al., 1990), tail DNA (percentage of DNA in the tail), and
tail length (TL; length of the tail, distance between the head and the last DNA fragment). Slides were scored blind by only one individual. Thirty randomly selected cells on each slide were analyzed. Comet images were randomly cap-tured from the center of the slide at a constant depth in the gel, avoiding the edges of the slide and overlapping figures. Comets with completely fragmented DNA (hedgehoglike figures with no apparent head) that could not be measured by the image analysis system were not taken into account. Data from two slides per animal and two animals at each concentration tested were pooled for the final processing. Pooling of two larvae with the same treatment was statisti-cally justified (Kolmogorov–Smirnov and Mann–Whitney tests). To compare the responses obtained with the MNT and the CA, to evaluate genotoxicity with the CA at each sample time, and to evaluate a dose–response relationship, it was necessary to test at least three doses for each expo-sure time. This constraint obliged us to limit to two the number of animals tested for each dose because using more would have entailed performing a second, independent electrophoresis, with its inevitable inherent variability. In thePleurodeles CA, only two concentrations were analyzed because of an insufficient number of larvae.
Among the statistical tests found in the literature, two are particularly well adapted to the CA: the Mann–Whitney U and Kolmogorov–Smirnov (KS) tests (Vapnik, 1995). Following the results of a previous study (Mouchet et al., 2005a), ultimately only the KS test was selected to be used in the present work. The KS test has higher sensitivity than the Mann–Whitney test. This makes it more suitable because it is a distribution test based on comparison of the two cumulative distribution functions of the samples tested and therefore uses the total information brought by the sam-ple measurements. In contrast, the Mann–Whitney test reduces this information by computing rank sums. ETM, OTM, tail DNA, and TL data (120 measurements per con-centration tested) were analyzed using the nonparametric Kolmogorov–Smirnov test ( ¼ 0.05).
Finally, although the processes involved in the formation and repair of strand breaks are not linear, the following evo-lution coefficient was introduced in order to establish the time-course of DNA damage between two successive expo-sure times at a given concentration,i, of captan:
ECCA¼½mðconcentration i; time 2Þ mðconcentration i; time 1Þ fmðneg.control; time 2Þ mðneg.control; time 1Þg ½mðconcentration i; time 1Þ mðneg.control; time 1Þg 100 ð1Þ wherem(concentration i, time 2), m(concentration i, time 1), m(neg.control, time 2), and m(neg.control, time 1) are
the means of the parameter computed for a given concen-tration,i, of captan, and the corresponding negative control at the two exposure times considered. The means of the pa-rameter relative to the negative control also are included in the above expression to take into account the evolution of the negative control. The evolution percentage (ECCA) can be positive (increased DNA damage between two exposure times) or negative (reduced DNA damage).
RESULTS
Acute Toxicity
Captan was severely toxic to Xenopus larvae exposed to captan concentrations of from 2000 to 125 g/L in MW (Table I). Lethality was 100% for concentrations of 2000, 1000, and 500 g/L on days 1, 2, and 4, respectively. Lethality was also 100% at a captan concentration of 250g/L in MW on day 4 and was 55% at a captan concen-tration of 125g/L on day 7. In the same way, captan was acutely toxic to Pleurodeles in MW at captan concentra-tions of from 2000 to 250 g/L (Table I). Lethality of 100% was observed for captan in MW at concentrations of 2000 and 1000 g/L on day 1 and of 75% and 45% for captan concentrations of 500g/L on day 2 and 250 g/L on day 3, respectively. No signs of acute toxicity were observed either forXenopus tadpoles exposed to lower
con-centrations (62.5, 31.25, or 15.60g/L) or for Pleurodeles larvae exposed to concentrations of 125, 62.5, 31.25, or 15.60g/L.
In RW, captan was acutely toxic toXenopus larvae at captan concentrations of from 2000 to 500g/L (Table I). Lethality of 100% was obtained at captan concentrations of 2000 and 1000g/L on day 1 and 500 g/L on day 5. A captan concentration of 250g/L was weakly toxic to lar-vae on day 8. Severe toxicity was observed inPleurodeles (Table I) exposed to captan concentrations of 2000 and 1000g/L (100% lethality) and 500 g/L (50%) on day 1. Weak toxicity was noted for a captan concentration of 250g/L in RW on day 1. Captan was not toxic to amphib-ian larvae exposed to lower concentrations (125, 62.5, 31.25, or 15.60g/L).
Micronucleus Test
MNT was performed in MW under conditions not acutely toxic to larvae. The results of theXenopus MNT showed that a captan concentration of 62.50g/L induced a signifi-cant genotoxic response (Table II). The lowest concentra-tions (15.60 and 31.25g/L) were not genotoxic to Xeno-pus larvae.
The results of thePleurodeles MNT showed no genotox-icity regardless of the concentration of captan tested: 125, 62.50, 31.25, or 15.60g/L (Table III).
TABLE I. Results of acute toxicity to Xenopus and Pleurodeles larvae exposed to captan (lg/L) in mineral water (MW) and reconstituted water (RW) for 12 days
Concentration (g/L) 2000 1000 500 250 125 62.5 31.25 15.60 X. leavis MW þþ (100%) þþ (100%) þþ (100%) þþ (100%) þþ (55%) RW þþ (100%) þþ (100%) þþ (100%) þ P. waltl MW þþ (100%) þþ (100%) þþ (75%) þþ (45%) RW þþ (100%) þþ (100%) þþ (50%) þ
%: percentage of dead (n¼ 20 larvae); No toxicity of larvae; þ weak toxicity; þþ severe toxicity.
TABLE II. Results of Xenopus micronucleus assay: frequency of micronucleated erythrocytes per 1000 cells in larvae exposed to captan (lg/L) for 12 days in mineral water
Concentration (g/L) NC PC 15.60 31.25 62.50 Lower quartile 0.5 28.5 1 1 1 Median 1 32 1 1 2 Confidence limit (0.59–1.41) (27.55–36.45) (0.59–1.41) (0.39–1.61) (1.55–2.45) Upper quartile 1.5 36 2 2.5 2 Results þ þ Number of animals 15 15 15 15 15
NC: negative control; PC: positive control.
Genotoxicity expressed as the median (number of micronucleated erythrocytes per 1000) and 95% confidence limits.
Nongenotoxic concentration compared to the negative control group (Mac Gill et al., 1956); þ genotoxic concentration compared to the negative con-trol group (Mac Gill et al., 1956).
Comet Assay
The CA was also performed in MW under conditions not acutely toxic to amphibian larvae. A captan concentration of 125g/L was not analyzed in Xenopus CA beginning on day 8 because on day 7 this concentration had been found to be acutely toxic. The lowest concentration was analyzed instead. Results of the Xenopus CA (Table IVa) showed
that captan had genotoxic effects at all concentrations tested (15.60g/L after 8 and 12 days; 31.25 and 62.5 g/L after 1, 2, 4, and 8 days; and 125g/L after 1, 2, and 4 days). The genotoxic response was strong whatever the parameter, except for 2 days of exposure to a captan concentration of 125g/L, which showed the tail DNA and OTM parameters to be nonsignificant, and for 12 days of exposure to a captan TABLE III. Results of Pleurodeles micronucleus assay: frequency of micronucleated erythrocytes
per 1000 cells in larvae exposed to captan (lg/L) for 12 days in mineral water
Concentration (g/L) NC PC 15.60 31.25 62.50 125 Lower quartile 15 153 13 15.5 13.5 12.5 Median 22 180 15 22 17 15 Confidence limit (18.35–25.65) (161.96–198.04) (12.16–17.84) (18.15–25.85) (14.16–19.84) (11.55–18.45) Upper quartile 24 197.5 20 25 20.5 21 Results þ Number of animals 15 15 15 15 15 15
NC: negative control; PC: positive control.
Genotoxicity expressed as the median (number of micronucleated erythrocytes per 1000) and 95% confidence limits.
Nongenotoxic concentration compared to the negative control group (Mac Gill et al., 1956); þ Genotoxic concentration compared to the negative con-trol group (Mac Gill et al., 1956).
TABLE IV(a). Results of Xenopus comet assay: mean tail DNA, ETM, OTM, and TL values in larvae exposed from 1 to 12 days to captan in mineral water
Exposure Time (days) Parameter Concentration of Captan (g/L) PCMMS NC 15.6 31.25 62.5 125 1 Tail DNA 1.536 0.3 Ne 2.436 0.37* 6.236 0.9* 8.136 0.94* 5.916 0.88* TEM 0.246 0.06 0.696 0.14* 2.56 0.55* 3.696 0.64* 2.166 0.52* OTM 0.466 0.08 0.66 0.1* 1.926 0.3* 2.836 0.3* 2.086 0.29* TL 14.136 1.67 23.926 1.55* 32.386 2.78* 40.046 3.35* 34.086 4.35* 2 Tail DNA 0.896 0.18 Ne 3.636 0.51* 5.086 0.7* 1.056 0.18 6.056 0.87* TEM 0.16 0.02 1.176 0.24* 2.26 0.45* 0.176 0.04* 2.066 0.44* OTM 0.196 0.04 0.976 0.15* 1.586 0.23* 0.226 0.04 26 0.28* TL 10.156 1.2 27.246 2.19* 35.026 3.49* 14.286 1.53* 30.256 2.83* 4 Tail DNA 1.256 0.24 Ne 2.586 0.48* 2.726 0.48* 2.76 0.5* 7.226 1.16* TEM 0.26 0.05 0.726 0.19* 0.896 0.22* 0.86 0.22* 3.316 0.7* OTM 0.296 0.06 0.656 0.12* 0.756 0.15* 0.696 0.15* 2.356 0.36* TL 14.186 1.59 21.36 2.06* 23.576 2.53* 22.016 2.01* 42.726 5.27* 8 Tail DNA 1.36 0.31 1.856 0.32* 5.676 0.93* 7.046 1.11* Ne 17.946 2.48* TEM 0.246 0.07 0.486 0.12* 2.516 0.56* 3.466 1.11* 11.056 2.17* OTM 0.36 0.07 0.456 0.08* 1.776 0.33* 2.256 0.39* 7.136 0.96* TL 14.576 1.97 20.586 1.85* 36.526 2.7* 38.856 3.03* 56.776 4.96* 12 Tail DNA 0.966 0.16 0.946 0.12 1.396 0.19* 1.456 0.23* Ne 186 2.03* TEM 0.156 0.03 0.156 0.03 0.296 0.05* 0.276 0.07* 10.86 1.81* OTM 0.196 0.03 0.216 0.03* 0.316 0.03* 0.316 0.04* 7.096 0.8* TL 14.356 1.42 13.726 1.21 17.696 1.34* 15.516 1.66* 55.676 5.24*
Calculated means of tail DNA (% DNA in the tail), ETM (extent tail moment), OTM (olive tail moment), and TL (tail length) followed by their 95% con-fidence limits. *Increased DNA damage, relative to the negative control, confirmed by a Kolmogorov–Smirnov test (p¼ 0.05). NC: negative control group; PCMMS: positive control group (MMS to 1.56 mg/L); Ne: concentration not explored (acute toxicity to larvae exposed to 125g/L appeared on day 7).
concentration of 15.60g/L, using the tail DNA and TEM parameters. A significant dose–effect relationship was observed with 31.25, 62.5, and 125g/L of captan after 1 and 2 days of exposure, whatever the parameter (Table IVb). After 4 days of exposure, a significant dose–effect relation-ship was only observed using the TL parameter. A signifi-cant dose–effect relationship was also obtained for captan concentrations of 15.6, 31.25 and 62.5g/L after 8 and 12 days of exposure, whatever the parameter, except TL and OTM for 31.25 and 62.5g/L after 8 and 12 days, respec-tively. Evolution coefficients (Table IVc) showed that levels of DNA damage increased or decreased according to length of exposure to a given concentration. Significantly increased DNA damage was observed for exposure to a captan concen-tration of 31.25 g/L between days 1 and 2 and between days 4 and 8. In contrast, a significant decrease was noted between days 2 and 4 and between days 8 and 12. Both these evolutions in damage were significant whatever the parame-ter. A significant increase in DNA damage was also observed with exposure to a captan concentration of 62.5 g/L between days 1 and 2 only for the TL parameter, whereas a significant decrease was noted for tail DNA; however, the effect on the ETM and OTM parameters was not significant. A significant increase in DNA damage was also recorded between days 4 and 8 of exposure with a captan
concentra-tion of 62.5g/L, whatever the parameter. In contrast, a sig-nificant decrease was noted between days 2 and 4 and between days 8 and 12 at the same concentration of captan, whatever the parameter. The negative control also showed an evolution in DNA damage. Evolution coefficients showed a significant reduction in DNA damage between days 1 and 2 and between days 8 and 12, whatever the parameter, except with TL between days 8 and 12. In contrast, a significant increase in DNA damage was seen between days 2 and 4. An increase in DNA damage was noted between days 4 and 8, but it was not significant. The global evolution between days 1 and 12 indicates a significant reduction in the values of the four parameters after captan exposure as well as in the negative group, whatever the parameter, except for the TL parameter in the negative control.
The results of the Pleurodeles CA (Table Va) showed genotoxic effects at captan concentrations of 62.5 and 125 g/L after 1 and 2 days of exposure, whatever the parameter, except with tail DNA after 2 days of exposure to 62.5g/L. After 4 days of exposure, a captan concentration of 62.5 g/L induced significant changes in the TEM/TL (significantly decreased) and OTM (significantly increased) parameters, whereas 125 g/L induced a significant decrease in the TL parameter. After 8 days of exposure, a captan concentration of 62.5 g/L induced a significant TABLE IV(b). Results of Xenopus comet assay: dose–effect relationship in larvae exposed for 1–12 days
to captan in mineral water
Exposure Time (days) Parameter
Captan Concentration (g/L) 15.6 vs. 31.25 31.25 vs. 62.5 62.5 vs. 125 1 Tail DNA Ne * * TEM * * OTM * * TL * * 2 Tail DNA Ne * * TEM * * OTM * * TL * * 4 Tail DNA Ne — — TEM — — OTM — — TL * — 8 Tail DNA * * Ne TEM * * OTM * * TL * — 12 Tail DNA * * Ne TEM * * OTM * — TL * *
*Significant dose–effect relationship, confirmed by a Kolmogorov–Smirnov test (p¼ 0.05).
Comparison of two captan concentration based on calculated means of tail DNA (% DNA in the tail), ETM (extent tail moment), OTM (olive tail moment), and TL (tail length). Ne: concentration not explored (acute toxicity to larvae exposed to 125g/L appeared on day 7).
increase only for the tail DNA and TL parameters. A captan concentration of 125 g/L induced either a significant increase in the TL parameter or a significant decrease in the OTM parameter. After 12 days of exposure, captan induced a significant decrease, whatever the concentration and the parameter, except for the tail DNA, TEM, and OTM param-eters at a concentration of 125 g/L. A significant dose– effect relationship was obtained whatever the parameter and the length of exposure, except after 4 days of exposure (Table Vb). Evolution coefficients (Table Vc) showed that levels of DNA damage significantly decreased between two successive exposures, whatever the concentration of captan, except for the 62.5g/L concentration with TEM between days 4 and 8 (not significant) and the 125g/L concentra-tion with OTM between days 8 and 12 (significantly increased). Negative controls also showed evolution in DNA damage. Evolution coefficients showed a significant reduction in DNA damage between days 1 and 2 whatever the parameter and between days 4 and 8 for the TEM and TL parameters, whereas they showed a significant increase between days 4 and 8 and between days 8 and 12, whatever the parameter. The global evolution between days 1 and 12 indicates a significant reduction in the values of the four pa-rameters after captan exposure, as well as in the negative group for the tail DNA and OTM parameters unlike the TL parameter (significantly increased).
DISCUSSION
The aim of this investigation was to determine if captan induces acute toxicity and has genetic effects on amphibian larvae.
Acute Toxicity
Captan was acutely toxic to amphibian larvae depending on the concentrations and nature of the exposure water. In fish, the LC50(48 h) ranged from 37 to 1000g/L for a variety of species (Jones et al., 1996). In the present work, toxic concentrations of captan in MW ranged from 2000 to 125g/L in Xenopus and from 2000 to 250 g/L in Pleuro-deles. They seemed to be higher for amphibian larvae exposed in RW than in MW: indeed, the toxic concentra-tions in Xenopus ranged from 2 mg/L to 250 g/L; even though a concentration of 250g/L was still toxic to Pleu-rodeles in RW, toxicity appeared less severe in RW than in MW. Before renewal, the medium in the control flasks was very clean. In contrast, the captan flasks containing RW were rather turbid. This turbidity was probably caused by amphibian residues/excretion or by the suspended captan or the degradation products that may interact with mineral ions, which make up a larger proportion of RW than of TABLE IV(c). Results of Xenopus comet assay: evolution coefficients (ECCA) of (% DNA in the tail), ETM (extent tail moment), OTM (olive tail moment), and TL (tail length) in larvae exposed for 1–12 days to captan in mineral water
Exposure Time (days) Parameter
Captan Concentration (g/L) NC 0,03125 0,0625 1–2 Tail DNA 42.15 þ205.44 10.74 ETM 60.76 þ139.18 6.5a OTM 58.42 þ464.73 4.77a TL 28.18 þ74.71 þ36.26 2–4 Tail DNA þ41.46 51.55 65.04 ETM þ113.04 51.52 67.61 OTM þ53.22 54.35 62.27 TL þ39.73 58.36 62.26 4–8 Tail DNA þ3.41a þ229.24 þ291.41 ETM þ15.10a þ338.12 þ372.09 OTM þ1.53a þ315.75 þ329.63 TL þ2.70a þ208.43 þ158.80 8–12 Tail DNA 25.85 90.26 91.51 ETM 35.10 94.17 96.32 OTM 36.28 91.99 94.04 TL 1.48a 84.78 95.23 1–12 Tail DNA 37.24 52.55 89.62 ETM 37.54 70.43 94.75 OTM 58.78 14.24 91.98 TL þ1.53a 65.84 93.65
These evolution coefficients are calculated as described in eq (1). Therefore, a positive value corresponds to increased DNA damage and a negative value to decreased DNA damage between two successive exposure times.
a
MW. Likewise, Anton et al. (1993) observed a similar tur-bidity in captan media of fish bioassays. Xenopus larvae seemed to be more sensitive thanPleurodeles to the toxic-ity of captan in MW, whereas no difference in sensitivtoxic-ity appeared in RW, probably because of its greater ion con-centrations, which may limit the difference in sensitivity by inhibiting the toxic potential of captan.
Micronucleus Induction
Captan appeared to be nongenotoxic inPleurodeles and gen-otoxic at the highest concentration inXenopus. Despite the lack of cytogenetic damage related to the exposure of Pleuro-deles larvae, a significant increase in the frequency of micro-nuclei inXenopus larvae was observed at a concentration of 62.5g/L. Therefore, it appears that the amphibian models are probably different. Sensitivity of the amphibians can dif-fer, depending on how the animals are exposed to the con-taminant. However, only a limited comparison between mod-els is possible because of their different biological character-istics such as chromosomes (small, 2n¼ 36 in Xenopus, and big, 2n¼ 24 in Pleurodeles), genomic size [3.1 pg in Xeno-pus and 19.5 pg in Pleurodeles (Fernandez et al., 1993)], and the food they eat.
Primary DNA Damage as Comet Induction
Primary DNA damage as comet induction was significantly observed inXenopus and Pleurodeles larvae. Captan induced DNA damage inXenopus at each exposure time, whatever the concentration. InPleurodeles, the genotoxic effects of captan were less clear, depending on the exposure time, because of the probable constant balancing between the repair and damage processes in the organisms. In the same way, DNA damage was significantly reduced with all exposures in both amphibian larvae. Such observations have been reported in previous works (Mouchet et al., 2005a, 2005b, 2006). In the present work, the reduction in DNA damage observed inPleurodeles for a given exposure and in both amphibian larvae for all expo-sures could have stemmed from different cellular processes, including DNA repair, detoxication, and adaptive responses to repeated doses. Indeed, it is well established that repair proc-esses are genetically controlled (Moustacchi, 2000). Each repair system preferentially acts on a particular kind of damage (Friedberg et al., 1995). Although the mechanism underlying the genotoxic effects is not well known, several reports have suggested that captan may interact with DNA (Anderson and Rosenkranz, 1974; SCC, 1981) and induce DNA repair (Swen-berg et al., 1976; Ahmed et al., 1977; Simmon et al., 1977). Snyder (1992) indicated that captan induces a type of DNA TABLE V(a). Results of Pleurodeles comet assay: mean tail DNA, ETM, OTM, and TL values in larvae exposed
for 1–12 days to captan in mineral water
Exposure
Time (days) Parameter
Captan Concentration (g/L) PCMMS NC 62.5 125 1 Tail DNA 1.786 0.3 3.26 0.45* 2.466 0.24* 5.386 0.77* TEM 0.316 0.07 1.46 0.24* 0.856 0.12* 2.96 0.67* OTM 0.686 0.11 1.246 0.17* 0.916 0.09* 2.356 0.35* TL 16.356 2.17 39.236 2.6* 31.126 2.25* 48.386 4.85* 2 Tail DNA 0.956 0.15 1.046 0.13 1.446 0.2* 8.916 0.96* TEM 0.156 0.04 0.246 0.05* 0.436 0.08* 5.126 0.77* OTM 0.266 0.04 0.336 0.04* 0.466 0.06* 4.896 0.5* TL 13.226 1.57 20.026 2.17* 25.66 2.62* 55.486 4.78* 4 Tail DNA 1.416 0.29 1.376 0.22 1.536 0.29 8.576 1.24* TEM 0.356 0.1 0.236 0.05** 0.36 0.08 5.326 1.05* OTM 0.366 0.08 0.46 0.06* 0.426 0.07 3.976 0.5* TL 21.226 2.9 14.796 0.97** 16.386 1.37** 57.176 4.44* 8 Tail DNA 1.246 0.21 1.446 1.02* 0.996 0.14 10.476 1.64* TEM 0.176 0.04 0.226 0.09 0.196 0.04 6.036 1.55* OTM 0.326 0.05 0.316 0.04 0.286 0.04** 5.416 0.77* TL 13.226 1.43 17.936 1.43* 16.486 1.98* 52.276 3.68* 12 Tail DNA 1.466 0.24 0.746 0.11** 1.396 0.22 8.386 1.12* TEM 0.356 0.09 0.136 0.03** 0.36 0.07 4.0996 0.89* OTM 0.426 0.07 0.236 0.03** 0.446 0.06 4.326 0.56* TL 20.836 2.48 14.626 1.68** 18.686 1.4** 55.376 4.39*
Calculated means of tail DNA (% DNA in the tail), ETM (extent tail moment), OTM (olive tail moment), and TL (tail length) followed by their 95% con-fidence limits.
*Increased DNA damage, relative to the negative control, confirmed by a Kolmogorov–Smirnov test (p¼ 0.05). **Decreased DNA damage, relative to the negative control, confirmed by a Kolmogorov–Smirnov test (p¼ 0.05). NC: negative control group; PCMMS: positive control group (MMS to 1.56 mg/L).
damage recognized by the excision repair system in human fibroblasts and that captan-induced DNA damage may be at least partially repaired via a long-patch repair system. The DNA-damaging and repair-inducing character of captan has been verified using the nick translation assay (Nose and Oka-moto, 1983; Snyder and Matheson, 1985).
Analysis of the CA results reveals that negative controls often exhibited significant background levels of strand breaks, which were dependent on length of exposure. This observation is in agreement with those of previous studies (Mouchet et al., 2005a, 2005b, 2006). Some authors (Singh et al., 1991; Guecheva et al., 2001) have demonstrated that DNA strand breaks in negative controls can be a conse-quence of prior cell isolation. This explanation does not hold for our experiments, using blood samples, for which no cell isolation step was necessary. Nevertheless, cardiac puncture may stress the cells. As previously suggested by Wilson et al. (1998), in agreement with the findings of Pacifici and Davies (1991), aerobic organisms are con-stantly exposed to endogenous and exogenous oxygen radi-cals and related oxidants, and although they have antiradi-cal defense systems, a low level of oxygen radiantiradi-cals can induce oxidative stress in all cellular components, including DNA. Significant DNA damage in controls in some cell types may be related to DNA packaging and background
al-kali-labile sites, rather than to exogenous strand breaks (Mitchelmore and Chipman, 1998; Singh et al. 1989). Moreover, as suggested by Liepelt et al. (1995), variable oxygen levels in test water lead to variation in DNA integ-rity. This background in negative controls can perhaps be expected because the comet assay measures strand breaks, some of which occur as a result of DNA damage and some of which occur naturally as a consequence of DNA replica-tion and transcripreplica-tion. Moreover, a certain background level is generally observed in biological tests, for example, in the MNT. In any case, the introduction of the evolution coefficient enabled us to consider only the part of the DNA damage that was related to the treatment.
For the same biological model, with regard to the four parameters, genotoxic responses were sometimes different. This confirms the results of previous studies that measured the four parameters in the larvae of Xenopus laevis and Pleurodeles waltl that had been exposed to identified sub-stances such as methyl methanesulfonate, ethyl methanesul-fonate (EMS), B[a]P, and metals (data not published), as well as draining water from dredged sediment containing TABLE V(b). Results of Pleurodeles comet assay:
dose–effect relationships in larvae exposed for 1–12 days to captan in mineral water
Exposure
Time (days) Parameter
Captan Concentration (g/L) 62.5 vs. 125 1 Tail DNA * TEM * OTM * TL * 2 Tail DNA * TEM * OTM * TL * 4 Tail DNA — TEM — OTM — TL — 8 Tail DNA — TEM — OTM * TL * 12 Tail DNA * TEM * OTM * TL *
Significant dose–effect relationship, confirmed by a Kolmogorov–Smir-nov test (p¼ 0.05). Comparison of 62.5 to 125 g/L based on the calcu-lated means of tail DNA (% DNA in the tail), ETM (extent tail moment), OTM (olive tail moment), and TL (tail length).
TABLE V(c). Results of Pleurodeles comet assay: evolution coefficients (ECCA) of % DNA in the tail, ETM (extent tail moment), OTM (olive tail moment), and TL (tail length) in larvae exposed for 1–12 days to captan in mineral water
Exposure
Time (days) Parameter
Captan Concentration (g/L) NC 62.5 125 1–2 Tail DNA 46.82 93.64 27.04 TEM 52.97 91.50 47.80 OTM 62.40 86.41 13.42 TL 19.16 70.23 16.14 2–4 Tail DNA þ48.68 145.93 75.29 TEM þ141.34 236.87 119.44 OTM þ39.50 46.11 69.83 TL þ60.53 194.36 139.03 4–8 Tail DNA 12.16a 598.59 303.88 TEM 51.20 140.07a 135.71 OTM 11.26a 126.42 153.16 TL 37.70 173.41 167.59 8–12 Tail DNA þ18.23 449.23 71.52 TEM þ104.06 536.23 339.15 OTM þ32.91 þ1661.53 159.43 TL þ57.63 231.80 165.81 1–12 Tail DNA 17.89 150.85 110.46 TEM þ13.01a 120.34 108.67 OTM 38.14 134.07 91.74 TL þ27.42 127.17 114.56
These evolution coefficients were calculated as described in eq (1). Therefore, a positive value corresponds to increased DNA damage between two successive exposure times and a decreased value to decreased damage.
a
Evolution of DNA damage between two successive exposure times not significant (Kolmogorov–Smirnov test,p¼ 0.05).
metal mixtures (Mouchet et al., 2005b). This result empha-sizes the importance of the number of parameters used in evaluating pollutant genotoxicity. Indeed, at least two pa-rameters must be used simultaneously. Likewise, Belpaeme et al. (1998) showed that the use of tail length as a discrimi-nating parameter in thein vivo CA on tissues of marine flat-fish did not provide consistent evidence of the genotoxic effects of EMS, whereas tail DNA did. Finally, Tomsu et al. (2002) found no significant correlations between head and tail parameters in sperm of patients undergoing treat-ment prior toin vitro fertilization.
Comparison between CA and MNT
The genetic results of the present evaluation showed a sig-nificant increase in the frequency of micronuclei in Xeno-pus larvae exposed to a captan concentration of only 62.5 g/L, whereas significant primary DNA damage was found whatever the concentration. Moreover, even though captan did not lead to the induction of any micronuclei in Pleuro-deles at any concentration, it did induce significant primary DNA damage depending on the concentration and the pa-rameter studied. This finding suggests a lack of clastogenic and/or aneugenic effects related to exposure to this particu-lar pesticide exposure in particu-larvae. Indeed, the genetic end points are different. Thein vivo MNT in amphibians detects chromosomal and/or genomic mutations (chromosomal damage and/or alteration of mitotic spindles), whereas alka-line CA detects primary DNA damage, expressed as single-and double-strsingle-and breaks, alkali-labile sites expressed as single-strand breaks, and single-strand breaks associated with incompletely repaired excision (sites present at the time of cell lysis). Captan exposure led to significant DNA damage measured through both the CA and the MNT in Xenopus, probably because of their clastogenic properties rather than their aneugenic ones. This result suggests that a captan concentration of 62.5 g/L induced primary DNA lesions and that at least a fraction of them could give rise to some fixed mutations, assuming that single-strand breaks could induce chromosomal damage resulting in micronu-cleus formation. The lowest concentration of captan exhib-ited a significant genotoxic response assessed by the CA, whereas no positive response was obtained through the MNT after 12 days of exposure at the same concentrations. In addition to DNA strand breaks, captan may also induce DNA base modifications and DNA–protein crosslinking in human fibroblasts (Snyder, 1992). Captan has been reported to bind to histones, thus altering their ability to stabilize DNA structure (Couch and Siegel, 1977). The ability of captan to denature proteins suggests that exposure to it might induce the stress response (Jones et al., 1996). Cap-tan induced lipid peroxidation with severe cytotoxicity in isolated rat hepatocytes (Suzuki et al., 1997). It is metabol-ically activated by microsomal cytochrome P450 (Suzuki
et al., 2004). Their metabolites might generate free radicals that would react with membrane lipids and induce oxidative breakdown of membrane structure. This mechanism could be the cause of DNA damage revealed by the CA in the present study. In this way, the CA applied to amphibian lar-vae for assessing genotoxicity of captan seems to be a more sensitive method than the normalized MNT (AFNOR, 2000). This might be attributed to the generation of repaira-ble damage that does not persist after one mitotic cycle. So it is not surprising that low concentrations of genotoxicant can induce single-strand breaks, as revealed by the CA, without necessarily inducing micronucleus formation because of repair mechanisms. Genotoxic responses to cap-tan can be measured from the first day of exposure with the CA, in comparison with the 12-day exposure required by the MNT according to the standardized procedure (AFNOR, 2000). Taking into account such results and because of their different end points and response times, both tests carried out on amphibian larvae could be pro-posed in a complementary manner in a battery test system.
The half-life of captan is about 3 h in water at a pH of 7.07 because of nonenzymatic hydrolysis (Wolfe et al., 1976), suggesting that the toxic and genotoxic effects that were measured would not only be associated with the intact parent compound. These effects would result from a combi-nation of parent plus metabolic or transformation products. The first step in the metabolism of captan is the cleavage of its NS bond to form tetrahydrophthalimide (THPI) and a derivative of the trichloromethylthio (TMS) side chain. THPI and the TMS side chain are each further metabolized through independent pathways. Four metabolites are formed from the TMS side chain, including the highly reac-tive species thiophosgene. Much of the toxicity of captan is attributed to thiophosgene, which denatures proteins by reacting with the amino and thiol groups of cysteine, form-ing a 2-thiazolidinethione rform-ing and by reactform-ing with other amino groups to form thiourea derivatives (Jones et al., 1996).
CONCLUSIONS
Summing up, captan could induce adverse effects in aquatic species. Nevertheless, the MNT showed no increase in cytogenetic damage inPleurodeles larvae , but it did show increased damage inXenopus tadpoles exposed to a captan concentration of 62.5g/L. Moreover, genotoxic response was determined according to primary DNA damage. Geno-toxic response may have resulted from the parent captan or its metabolites, with no distinction made between them.
Given that the induction of DNA damage measured in the two bioassay stems from different mechanisms and that they have different response times, the CA could be proposed for use with amphibians as a relevant complemen-tary method for the assessment of genotoxicity of pure
substances and complex contaminated matrices for biomo-nitoring studies.
This work was supported by the French Agency for the Envi-ronment and Energy Resources (ADEME). We are especially grateful to Marie Charveron and Caroline Baudouin from the Pierre Fabre Laboratories in Toulouse for use of the Komet soft-ware. We are also very indebted to Peter Winterton for reviewing the English version of the manuscript.
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