Assessment of the toxicity of organochlorine pesticide endosulfan in clams Ruditapes philippinarum

Ecotoxicology and Environmental Safety 93 (2013) 22–30

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Assessment of the toxicity of organochlorine pesticide endosulfan in clams Ruditapes philippinarum Yanxia Tao, Luqing Pan n, Hui Zhang, Shuangmei Tian The Key Laboratory of Mariculture, Ministry of Education, Ocean University of China, Qingdao 266003, PR China

art ic l e i nf o

a b s t r a c t

Article history: Received 5 February 2013 Received in revised form 25 March 2013 Accepted 26 March 2013 Available online 2 May 2013

This study is aimed at evaluating the effects of endosulfan in clams (Ruditapes philippinarum). For this purpose, a study was performed on clams exposed to 0.005, 0.05 and 0.5 μg/L endosulfan for 15 days. S ubsequently, the level of ethoxyresorufin-O-deethylase (EROD) activity, glutathione-S-transferase (GST) activity, glutathione (GSH) content, superoxide dismutase (SOD) activity, lipid peroxidation (LPO) and DNA strand break was determined in gills and digestive glands. Among the parameters, endosulfan caused significant changes in induction of EROD activity and oxidative stress in clams R. philippinarum. The exposure to endosulfan increased the concentration of EROD, GST, GSH, MDA and decreased the concentration of SOD. Moreover, according to the correlation analysis results, the EROD activity and GSH content in digestive gland as well as GST activity, LPO and DNA damage in both tissues had excellent correlation with endosulfan concentration. These results provided information on potential biomarkers that could be effectively applied to the biomonitoring of aquatic ecosystem in areas susceptible to persistent organochlorine compounds contamination, and also information on toxic effects. & 2013 Elsevier Inc. All rights reserved.

Keywords: Endosulfan Ruditapes philippinarum EROD Oxidative stress DNA damage Biomarker

1. Introduction Some persistent organochlorine compounds are found in the environment and can bioaccumulate in the organisms along the trophic chain. Endosulfan (EDS) is one such organochlorine (OC) compound that has been classified as highly toxic by the majority of environmental protection agencies (Sutherl et al., 2004). However, EDS has been used as a pesticide since 1995s in many parts of the world over a large variety of crop fields (Jia et al., 2009). Although its use has been restricted (UNEP-POPS-COP5, 2011), residues of this compound have been detected in fruit, salad vegetables, meat, and milk (Kumari et al., 2006; Ciscato et al., 2002; Nag and Raikwar, 2008). Endosulfan belongs to the cyclodiene group (C9H6Cl6O3S), and is composed of a mixture of isomers, namely, α-endosulfan and βendosulfan. It is considered to be toxic to all kinds of organisms and its residues have been detected in various geographical regions ranging from temperate environments to the Arctic (Xu et al., 2007; Weber et al., 2010). According to the survey, the endosulfan content in surface sediment from Xiaohai Bay, China, was 0.54 ng/g (Mu et al., 2007).

n Correspondence to: Fisheries College, Ocean University of China, Yushan Road 5, Qingdao 266003, China. Fax: +86 532 82032963. E-mail address: [email protected] (L. Pan).

0147-6513/$ - see front matter & 2013 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.ecoenv.2013.03.036

The aquatic environment is inevitably affected by environmental pollutants, including EDS. And it consequently affects the bivalves, which have been proved to be suitable bioindicators for monitoring trace toxic contaminant levels of coastal waters; i.e., Mussel Watch in the United States and RNO in France (Goldberg, 1978; RNO, 1988). Because they can take up and concentrate contaminants and exhibit a certain ability to respond to pollutants at a molecular level in the first place. So presently, attention has focused on the detection of biological responses of bivalves, which can give information on toxic effects (Anderson et al., 1999; Quinn et al., 2005). Therefore, there is an urgent demand for studies that can correlate the effects of xenobiotic in bivalves. Beauchamp and Fridovich (1971) showed that pollutants can be metabolized in two different ways in aquatic organisms. The first process is the metabolism or biotransformation through mixed function oxygenases (MFO) and glutathione S-transferase (GST) for detoxification. Many field and laboratory studies have reported that 7-ethoxyresorufin O-deethylase (EROD) is a sensitive biomarker in detecting the effects of different pollutants including ester, polycyclic aromatic hydrocarbons (PAHs), nitro aromatic hydrocarbons (NAHs), polychlorinated biphenyls (PCBs) and endosulfan (EDS) (Lemaire-Gony et al., 1995; Viarengo et al., 1997; Wassenberg et al., 2002; Salvo et al., 2012). However, only a few studies relating to EROD have been done in invertebrates. Glutathione-S-transferase (GST) is an enzyme in phase II metabolism, previously studied in Mytilus edulis. GST activity has been established as a suitable biomarker for aquatic contamination

Y. Tao et al. / Ecotoxicology and Environmental Safety 93 (2013) 22–30

using bivalves as sentinel organisms. An estuarine sediment quality report showed that GST activities were most probably due to contamination and could be used as biomarker (Fossi Tankoua et al., 2013). Another metabolic process is antioxidant enzyme metabolism after biotransformation. Some reactive oxygen species (ROS) including superoxide radicals (O2−), hydroxyl radicals and hydrogen peroxide will be produced by aquatic organisms for detoxification and excretion of lipophilic chemicals after biotransformation (Livingstone et al., 1992). Many studies have indicated that active oxygen is more dangerous to organisms comparing with the environment pollutant itself, and hence antioxidant systems are important for eliminating the active oxygen. Superoxide dismutase (SOD) is crucial in preventing the formation of lipid peroxidation by catalyzing the disproportionation of the lipid peroxidation initiator and the transformation of superoxide radicals (O2−) to H2O2 and O2. The enzyme plays an important role in protecting organisms from oxidative stress. Most studies on antioxidants as biomarkers for the aquatic environment have been carried out on fish (Fenet et al., 1996; Van der Oost et al., 1996; Telli Karakoc et al., 1997) and on marine invertebrates (Porte et al., 1991; Livingstone et al., 1992, 1995; Labrot et al., 1996). Organisms can adapt to increasing ROS production by upregulating antioxidant defenses, such as the activities of antioxidant enzymes (Livingstone, 2003). But failure of antioxidant defenses to detoxify excess ROS production can lead to significant oxidative damage including enzyme inactivation, protein degradation, DNA damage and lipid peroxidation (LPO) (Halliwell and Gutteridge, 1999). In particular, cytotoxicity as brought by LPO was noted in cases where antioxidant defenses were the lowest, which provided support for the use of these parameters as biomarkers for toxicity (Di Giulio et al., 1993; Regoli and Principato, 1995; Sole et al., 1995; Telli Karakoc et al., 1997). Meanwhile, reactive molecules can interact with the genetic material and cause DNA damage as a toxic terminal of the induction of a cascade of cellular events. DNA integrity reduction represents the genotoxicity which may have a long-term effect on the sustainability of a particular population. DNA strand breakage in bivalves using alkaline unwinding has been approved as an effective biomarker to assess the genotoxicity of pollutants (Ching et al., 2001). The use of biochemical biomarkers of environmental contamination allows a sensitive assessment of the xenobiotic effects in aquatic organisms in order to detect early alterations in the environment prior to any irreversible harm being caused to the ecosystem. In our research, in order to select the biochemical biomarkers and investigate detoxification mechanism against EDS, we used R. philippinarum as a model organism and studied the effects of different EDS concentrations on enzyme activities of EROD, GST, SOD, GSH contents, LPO levels and the DNA in digestive gland and gills.

2. Materials and methods 2.1. Chemicals Endosulfan of 97.5% purity consisting of α and β isomers (7:3) and ultrapure reduced glutathione (GSH) was purchased from Amresco (American). 1-Chloro-2,4dinitrobenzene (CDNB), 5, 5′, dithiobis-2-acid nitrobenzoic (DTNB), disodium salt of reduced form -nicotinamideadenine dinucleotide phosphate (NADPH), pyrogallic acid, O7-ethylresorufin (ERF) were purchased from Fluka (USA). All other chemicals and solvents were of analytical grade. 2.2. Animals and treatments Healthy R. philippinarum (shell length 4.03 7 7 0.21 cm) were collected from Red Island (Yellow Sea, Qingdao, China) and acclimated in tanks containing aerated sand-filtered seawater (salinity 31‰, pH 8.1) at 127 7 0.5 1C for two weeks before the exposure test. During the acclimated period, the water in each tank was

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renewed completely once daily and the clams were fed with dried powder of Spirulina platensis (30 mg for each individual per day). After acclimatization, clams were transferred to plastic aquaria (50
40
30 cm), experimental conditions (salinity, pH, temperature, feeding and photoperiod) were maintained as mentioned above. In treatment aquaria, clams were exposed to different endosulfan concentrations (0.005, 0.05 and 0.5 μg/ L). There were triplicates for each level and 30 clams in each aquarium. One third of the water was renewed every morning, and seawater containing the same concentrations of endosulfan was added to maintain the concentrations of endosulfan during the experiments. Endosulfan was first dissolved in acetone. The final acetone concentration was 0.001% in all tanks including the control ones (the acetone test has been done in a preliminary experiment with the result that there was no influence on clams of acetone). The exposure concentrations of endosulfan were based on the concentration of endosulfan in the coastal seawater, surface sediments in China, as well as endosulfan solubility (22–25 1C). During the experimental period (15 days), there was no mortality of clams at all concentrations of endosulfan and the control groups. Clams were sampled 1, 3, 6, 10, and 15 days after the end of the acclimatization period. The gills and the digestive gland of each clam were used for the further assay. Ten to fifteen clams were sampled for each day and concentration, including controls. And the tissues were dissected, immediately frozen in liquid nitrogen and then kept at −80 1C. 2.3. Measurement of biomarkers of R. philippinarum 2.3.1. Sample preparation Gills and digestive glands were placed into phosphate buffer (0.125 M, pH 7.7, containing Na2EDTA 0.05 M, 2–4 1C), and homogenized (3 min duration) in an ice bath. After centrifugation at 3000
g for 10 min and 13,000
g for 30 min at 4 1C, the supernatant was separated and measured for GSH contents and EROD, GST and SOD activities. MDA was quantified in digestive gland and gill. The tissues were placed in Tris– acetate buffer (50 mM, pH 7.4), homogenized on ice and then centrifuged at 3000
g for 10 min at 4 1C. The supernatant was assayed for MDA contents. DNA extraction: DNA was extracted from gills and digestive glands by mashing the tissues with plastic sticks in 3 mL TE buffer (50 mM Tris, 100 mM EDTA, pH 8.0). 0.45 mL lysate was transferred into 1.5 mL Eppendorf tubes. Then 0.05 mL of 10% SDS and 3 mL of 20 g/L proteinase K were added and the mixture was incubated for 5 h at 55 1C. An equal volume of buffered phenol/chloroform/isoamyl alcohol (PCI) (25:24:1, v/v/v, pH 8.0) was then added to the sample. The sample was gently mixed for 15 min and then centrifuged for 10 min at 13,000 rpm at 4 1C. The aqueous layer was transferred to a new tube and digested with 0.8 mL RNase (10 mg/mL) for 30 min at 37 1C and the digesta was extracted twice by PCI. DNA was precipitated from the resulting aqueous layer by adding 2 volumes of absolute ethanol and 1/10 volume of 3 M sodium acetate, pH 5.2. The samples were allowed to settle for 2 h at −20 1C, and then centrifuged for 15 min at 13,000 rpm. The resulting pellet was rinsed with 1 mL 70% ethanol and air-dried, then dissolved in 1 mL of TE buffer (10 mM Tris, 1 mM EDTA, pH 8.0). The amount of DNA was quantified using a UV/visible spectrophotometer (Ultro spec 2100 pro, Amersham Biosciences, Sweden). Soluble protein in enzyme extracts was quantified with a commercial kit (BioRad) based on Coomassie blue, using bovine serum albumin standards (Bradford, 1976).

2.3.2. Measurement of biochemical biomarkers Mixed function oxidase activity (EROD) was determined according to the modified method of Pohl and Fouts (1980). The reaction mixture contained 100 μL supernatant, 10 μL 0.2 mM O7-Ethylresorufin, 10 μL 6 mM NADPH and 1.88 mL phosphate buffer (0.125 M, pH 7.7, containing Na2EDTA 0.05 M, 2–4 1C), allowed to proceed for 10 min at RT, and stopped by the addition of 0.5 mL carbinol. Incubation vials were centrifuged to remove precipitated microsomal protein, and supernatants were transferred to vials for measurement of resorufin concentrations in a luminescence spectrometer (Model LS55, Perkin-Elmer of U.K.) at an excitation wavelength of 560 nm and an emission wavelength of 580 nm. Resorufin was identified and concentrations were calculated by comparison to retention times and responses of resorufin standards. Blanks corresponded to t ¼0 min and quantification was achieved with standard additions of resorufin. EROD activity was expressed as nmol resorufin/min/mg microsomal protein. GST was purified by a modification of the method of Habig et al. (1974). There action mixture contained 200 μL supernatant, 2 mL phosphate buffer (0.125 M, pH 7.7, containing Na2EDTA, 0.05 M, 2–4 1C), H2O 400 μL, 200 μL 15 mM 1-chloro-2,4dinitrobenzene (CDNB) dissolved in 95% ethanol and 200 μL 15 mM of reduced glutathione (GSH ). GST activity was determined following the conjugation of GSH with CDNB at 340 nm (a1¼ 9.6 mM−1). A unit of GST activity was defined as the amount of glutathione conjugate formed using 1 nM GSH and CDNB/m in per mg protein (nM 2, 4-dinitrophenyl glutathione/mg protein/min). SOD activity was measured by a modification of the method of Marklund and Marklund (1974). This assay is based on the ability of SOD to inhibit the autooxidation of pyrogallol (50 mM) in 50 mM Tris–HCl buffer (pH 8.3). The reaction mixture contained 4.5 mL Tris–HCl buffer, 100 μL supernatant and 10 μL pyrogallol.

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Oxidation of pyrogallol was monitored by measuring absorbance at 325 nm. 1 U of SOD activity was defined as 50% inhibit ion of the oxidation process (U/mg protein/min). GSH content was measured using spectrofluorometric assay by slight modifications of the method of Hissin and Hilf (1976). Approximately 0.1 mL of each tissue supernatant was neutralized with 2.8 mL 1 M phosphate buffer (pH 8.0) and 0.1 mL 1 mg/mL ortho-opthalaldehyde. Then the mixture was incubated at room temperature for 20 min and measured using spectrofluorometer. The standard curve, obtained from known concentrations of GSH standards, was determined from the absorbance at 340 nm. LPO levels were expressed by MDA contents. MDA contents were evaluated by an improved thiobarbituric method (Wills, 1987). Briefly, 50 μL of tissue homogenate was diluted to 1 mL with distilled water and mixed with 500 μL 20% trichloroacetic acid containing 1 mM FeSO4, 1 mL of 0.67% thiobarbituric acid was added and the mixture was kept at 90 1C for 10 min. After centrifugation at 3000
g for 5 min, 1 mL aliquot was withdrawn and mixed with 2 mL distilled water. Absorbance was measured at 530 nm. MDA contents were expressed as nmol TBARS/mg protein/min. The alkaline unwinding assay used in the study was adapted from Ching et al. (2001). In the assay, the rate of transition of double-stranded DNA (dsDNA) to single-stranded DNA (ssDNA) under pre-defined alkaline denaturing conditions was proportional to the number of breaks in the phosphodiester backbone, and thus was used as a measure of DNA integrity (Daniel et al., 1985). The amounts of various types of DNA were quantified by measuring the varying degrees of fluorescence resulting from a DNA-binding dye, bisbenzamide. After reaction with the dye, the fluorescence of dsDNA is double than that of the ssDNA (Cesarone et al., 1979). The DNA sample was diluted and divided into three equal portions for fluorescence determination of dsDNA, ssDNA and alkaline unwound DNA (auDNA). The fluorescence of the initial DNA or dsDNA was determined by placing 100 mL of the DNA sample, 100 mL of 25 mM NaCl and 2 mL of 0.5% SDS in a prechilled test tube, followed by the addition of 3 mL of 0.2 M potassium phosphate (pH 6.9), and 3 mL of bisbenzamide (1 mg/mL). The contents were mixed and allowed to react in darkness for 15 min. The fluorescence of the sample was measured using a spectrofluorimeter (Molecular Spectroscopy LS 55 from Instruments, P.E., MA, USA) with an excitation wavelength of 360 nm and an emission wavelength of 450 nm. The fluorescence of ssDNA was determined as above but using a DNA sample that had already been boiled at 80 1C for 30 min to completely unwind the DNA. auDNA samples were made by subjecting the initial DNA samples to alkaline treatment, i.e., 50 mL of 50 mM NaOH was rapidly mixed with 100 mL of DNA sample in a prechilled test tube and incubated on ice in darkness for 30 min, followed by a rapid addition and mixing of 50 mL of 50 mM HCl and 2 mL of 0.5% SDS; then the mixture was forcefully passed through a 21 G needle several times.

The fluorescence of the alkaline unwound DNA sample was measured as described above. The ratio of double-stranded DNA to total DNA (F value) was determined as follows: F value ¼ (auDNA−ssDNA)/(dsDNA−ssDNA).

2.4. Statistical analysis All data presented are the mean values of three independent sets of experiments. Each value was presented as means 7 standard deviation (S.D.). Statistical analysis were carried out by one-way ANOVA using the Dunnet's test to evaluate whether the means were significantly different, taking P o0.05 as minimal significant. Statistical computations and correlation analysis were performed with SPSS 13.0 for Windows (SPSS Inc.). The correlation was showed as the Pearson ratio.

3. Results 3.1. Effects of EDS exposure on clam R. philippinarum 3.1.1. EROD activity The EROD responses of the gill and digestive gland are shown in Fig. 1. The results showed that the EROD activities in digestive gland of 0.05 and 0.5 μg/L treatment groups increased and became significantly different from control values at day 1 (Po 0.05 and Po 0.01, respectively). The EROD activities remained high and significantly different from control values until the end of the experiment. While significant effects were found on the EROD activity in gills only in 0.05 μg/L treatment group at day 1 and 0.5 μg/L treatment group at day 6 and day 10.

3.1.2. GST activity As shown in Fig. 2, in 0.005 μg/L EDS group increased first and peaked at day 6 (gills) and day 10 (digestive glands). The GST activity in 0.05 and 0.5 μg/L EDS groups increased significantly (P o0.05) at first and then remained constant from day 6 until the end of the experiments.

Fig. 1. The activities of EROD of gills and digestive glands of R. philippinarum exposed to EDS (0.005, 0.05 and 0.5 μg/L) for 1, 3, 6, 10 and 15 days. Values are presented as the mean 7S.D. (n¼ 3). Statistical analysis was carried out by one-way ANOVA using the Dunnet's test. Significant differences from control in the same time of sampling are indicated with an asterisk at Po 0.05, and with two asterisks at P o 0.01.

Y. Tao et al. / Ecotoxicology and Environmental Safety 93 (2013) 22–30

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Fig. 2. The activities of GST of gills and digestive glands of R. philippinarum exposed to EDS (0.005, 0.05 and 0.5 μg/L) for 1, 3, 6, 10 and 15 days. Values are presented as the mean 7 S.D. (n¼ 3). Statistical analysis was carried out by one-way ANOVA using the Dunnet's test. Significant differences from control in the same time of sampling are indicated with an asterisk at Po 0.05, and with two asterisks at Po 0.01.

Fig. 3. The content of GSH of gills and digestive glands of R. philippinarum exposed to EDS (0.005, 0.05 and 0.5 μg/L) for 1, 3, 6, 10 and 15 days. Values are presented as the mean 7 S.D. (n¼ 3). Significant differences from control in the same time of sampling are indicated with an asterisk at Po 0.05, and with two asterisks at Po 0.01.

3.1.3. GSH content The level of GSH in digestive gland was induced by EDS at first 1 day then declined gradually with exposure period prolonged (Fig. 3). In the gill, GSH content of 0.005 and 0.05 μg/L EDS increased and peaked at day 6 and day 3 respectively, and

remained high and significantly different from control values until the end of the experiment (Po 0.05 or Po 0.01); The GSH content of 0.5 μg/L EDS exposure group reached the highest level at day 1 and then decreased at day 3 and remained constant from day 6 until the end of the experiments.

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Fig. 4. The activities of SOD of gills and digestive gland of R. philippinarum exposed to EDS (0.005, 0.05 and 0.5 μg/L) for 1, 3, 6, 10 and 15 days. Values are presented as the mean 7S.D. (n¼ 3). Statistical analysis was carried out by one-way ANOVA using the Dunnet's test. Significant differences from control in the same time of sampling are indicated with an asterisk at Po 0.05, and with two asterisks at P o 0.01.

3.1.4. SOD activity The effects of endosulfan on SOD activity of clam R. philippinarum are illustrated in Fig. 4. In comparison with the controls, SOD activities of 0.005 μg/L endosulfan-exposed increased significantly at day 1 (gills) and day 3 (digestive glands), but were declined at day 6. On the other hand, the treatments of 0.05 and 0.5 μg/L EDS inhibited the SOD activities significantly at the whole exposure time.

(F value) showed obvious linear relationships. Among these indexes, GST activity and LPO level showed a significant and positive correlation with the EDS concentration. While, the level of SOD activity and F value was negatively correlated with EDS exposure. There were no correlations for GSH and for EROD.

3.1.5. LPO Level Exposures to 0.005, 0.05 and 0.5 μg/L EDS caused an elevation in LPO in digestive gland of exposed clams. The results in Fig. 5 showed that the MDA contents of digestive gland all increased with the increasing sampling time, and were statistically significant from day 1 of the experiment EDS group (P o0.05 or P o0.01), and there was a positive correlation between the MDA contents in digestive gland and the EDS concentrations. Levels of MDA value in clam gills are shown in Fig. 5. The results indicated that the MDA contents of all treatment groups increased significantly (P o0.05) in comparison with the controls and showed a peak time.

Various xenobiotics are known to cause damage in aquatic organisms by stimulating the production of harmful oxyradicals via processes of redox cycling. These processes cause interference with electron transport, either inducing enzyme systems such as cytochrome P450 that mediate oxidation reactions or depleting protective antioxidants (Winston and Di Giulio, 1991). The cytochrome P450 monooxygenase system metabolizes a large number of xenobiotic compounds. The induction of the subfamily CYP1A is catalytically expressed by the activity of some specific enzymes such as the EROD (Bucheli and Fent, 1995). This induction occurs when the aquatic organisms are exposed to environmental pollutants such as dioxins, poly-chlorinated biphenyls (PCBs), polyaromatic hydrocarbons (PAHs), and others. In the case of EDS, some studies have been carried out for evaluating the effects on various fish species, and EDS strongly induced hepatic microsomal EROD activity (Ana et al., 2007; Marc et al., 2010; Salvo et al., 2012). In fact, in our study, we have observed a significant increase in EROD activity in digestive gland in the group exposed to all the EDS dose in comparison with control, suggesting that liver phase I metabolism was induced by the presence of EDS; the results showed a better dose–response relationship between EROD activity and concentrations of EDS, reflecting the susceptibility of clam EROD to EDS. However, the EROD activity in gills did not show a significant variation, which indicates that the digestive glands are more sensitive to EDS exposure than the gills. According to Rana and Shivanandappa (2010), some environmental pollutants can induce the production of ROS in organisms.

3.1.6. DNA strand breaks The results of mean F values of each individual treatment group are presented in Fig. 6. F values from the 0.05 and 0.5 μg/L treatment groups in both tissues showed marked decreases (P o0.05) after 1 day of exposure, while the 0.005 μg/L treatment group showed marked decreases in mean F values after 3 day of exposure (P o0.05). 3.2. The correlation analysis of EDS on detoxification metabolism, LPO and DNA damage indexes of tissues in clams R. philippinarum As shown in Table 1, during the EDS exposure time, the activities of GST and SOD, the LPO level and DNA damage index

4. Discussion

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Fig. 5. The content of MDA of gills and digestive glands of R. philippinarum exposed to EDS (0.005, 0.05 and 0.5 μg/L) for 1, 3, 6, 10 and 15 days. Values are presented as the mean 7 S.D. (n¼ 3). Statistical analysis was carried out by one-way ANOVA using the Dunnet's test. Significant differences from control in the same time of sampling are indicated with an asterisk at Po 0.05, and with two asterisks at Po 0.01.

Fig. 6. F value (The ratio of double-stranded DNA to total DNA) of gills and digestive gland of R. philippinarum exposed to EDS (0.005, 0.05 and 0.5 μg/L) for 1, 3, 6, 10 and 15 days. Values are presented as the mean 7 S.D. (n ¼3). Statistical analysis was carried out by one-way ANOVA using the Dunnet's test. Significant differences from control in the same time of sampling are indicated with an asterisk at Po 0.05, and with two asterisks at P o0.01.

And the excessive ROS may lead to oxidative damage of DNA, proteins and lipids (Halliwell and Gutteridge, 1999). Fortunately, enzymatic (e.g., SOD, GST and GPx) and non-enzymatic (e.g., GSH,

ascorbate, carotenoids, and free proline) antioxidant defense systems in living organisms can provide cellular protection against oxidative stress. Salvo et al. (2012) reported that GST, SOD and GPx

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Table 1 The correlationships of EDS on detoxification metabolism and DNA damage indexes of tissues in clams R. philippinarum. Tissue

Index

Pearson correlations (r) Exposure time 1d

Gill

3d

EROD 0.674 0.841 0.870n GST 0.860n GSH 0.785n 0.474 SOD −0.634 −0.752 LPO 0.798nn 0.901n F value −0.871n −0.856n

6d

10 d

15 d

0.762 0.737 0.685 0.727n 0.879n 0.909n 0.127 0.292 0.340 −0.989n −0.980n −0.985n 0.881n 0.802n 0.891n −0.794nn −0.794nn −0.891nn

Digestive gland EROD 0.976n 0.979n 0.975n 0.977n 0.981n GST 0.884n 0.983n 0.905n 0.955n 0.946n GSH 0.798nn 0.880nn 0.870n 0.878n 0.882n SOD −0.731 −0.677 −0.976n −0.790nn −0.883n LPO 0.969n 0.990nn 0.696 0.712 0.610 F value −0.865n −0.898nn −0.794nn −0.883n −0.788n Note: Statistical significance was denoted by respective control group.

n

P o0.05,

nn

Po 0.01 versus the

activities in juvenile common carp (Cyprinus carpio) were increased by EDS (1 mg/L) for 15 days. In the freshwater cladoceran Daphnia magna, SOD activity was equally inhibited at EDS exposure. In the study of Salvo et al. (2012), a significant increase in the total GSH concentration was observed in the liver of EDS exposed juvenile common carp Cyprinus carpio. Glutathione-S-transferase (GST) plays a key role in cellular detoxification of various xenobiotic chemicals. The enzymes protect cells against toxicants, neutralizing them and rendering the product more water soluble (Lamoureus and Rueness, 1987). Increase in the activity of GST has been used as a marker of exposure to pesticides (Machala et al., 1997). In the present study, the level of GST in tissues of clams R. philippinarum was found to be higher than that in controls. Similar results have been reported in the crayfish Procambarus clarkii exposed to cadmium (Almar et al., 1987), in the field crab Paratelphusa hydrodromus exposed to endosulfan (Yadwad, 1989), and in the mussel Mytilus edulis exposed to sediments contaminated with pollutants (Shehan et al., 1995). The elevated GST activity level noted by us in R. philippinarum might reflect the formation of GSH and endosulfan complexes as a means of detoxification/elimination. Such a mechanism has been previously noted in M. malcolmsonii exposed to heavy metals (Kabila, 1996; Yamuna, 1997). Although the activation of such a mechanism probably conferred cytoprotection against endosulfan-induced celular stress, the toxic effect of endosulfan was apparently not fully neutralized, since alterations were noted in various biochemical parameters. GSH level was instantly induced in the clam exposed to EDS even in the lowest concentration in this study. It is probably because EDS induced the synthesis of GSH as the oxidative stress response. The SOD activity of digestive glands and gills increased in short time in the lowest concentration of EDS, which demonstrating that organisms have increased the ability of eliminating the O−2 and the H2O2 coming from the metabolite of SOD and other complicated biotransformation in short time. The constant level of SOD in low concentrations occurred after 6 days of exposure, indicating that there was a balance between O−2 and detoxification of the major antioxidant enzymes. The SOD activity can be restrained in 0.05 and 0.5 μg/L EDS groups, so in high concentration EDS exposure conditions, large numbers of O−2 and the H2O2 have been accumulated which restrained the antioxidant enzyme

activities in reverse. The detoxification function of antioxidant system has been inhibited in this time. The imbalance between pro and antioxidant systems will cause the formation of toxic hydroxyl radicals, with direct consequences on cell integrity and cell function itself (Dorval and Hontela, 2003). Enhanced lipid peroxidation in aquatic animals was responsive to toxicants (Thomas and Wofford, 1993; Roméo and GnassiaBarelli, 1997) and was a direct indicator of oxidative stress and hence quantification of MDA is the way to evaluate the LPO level. In our studies, the MDA contents were significantly higher than controls, indicating that the antioxidant enzymes system could not wholly eliminate O−2 . Production of LPO induced by EDS has been reported previously by other researchers. Hincal et al. (1995) demonstrated that either a single dose of 30 mg/kg or repeated doses of 5 and 15 mg/kg/d EDS induced LPO in brain and liver of rat. Similarly, Pandey et al. (2001) and Atif et al. (2005) observed elevated levels of LPO in gills, liver and kidney of C. punctatus exposed to 5 μg/L EDS for 24 h. Also, Dorval and Hontela (2003) found elevation in LPO levels in cultured adrenocortical cells of O. mykiss exposed to 10−7, 10−6 and10−4 M EDS for 60 min. Moreover, some of the reactive intermediates can directly bind to DNA forming DNA adducts, while others, comprising free radicals and ROS cause oxidative damage of DNA (Shugart, 1999). Pandey et al. (2006) studied the effect of EDS on Channa punctatus and found serious DNA strand breaks in the gill and kidney. Another research showed that endosulfan induced DNA fracture and migration and caused DNA damage in grass carp liver cell (Wu et al., 2012). In our results, The F value in the gill and digestive gland of the 0.005 μg/L EDS group decreased until day 6 and then showed a gradual increase trend until the end of the experiment, which implied that the reactive intermediates may be maintained at a comparative persistent level, initiating the DNA repair system. While the F value in both tissues of the 0.05 and 0.5 μg/L EDS treatment groups showed a decrease until day 3 and became constant gradually. We presume that DNA damage in the 0.005 μg/L EDS group could be repaired as the repair system was initiated, while in the 0.05 and 0.5 μg/L EDS groups the repair systems were overwhelmed. The result also showed that all the basal levels of the studied biomarkers in digestive glands were much higher than in gills, which indicating that the gill's detoxification function is weaker than the digestive gland. Similarly, Pan et al. (2009) found that SOD activity in the digestive gland of scallop Chlamys farreri was much higher than in gill. In the green-lipped mussel (Perna viridis), GSH contents and the activity of SOD in the hepatopancreas were higher than in gills (Cheung et al., 2004). The difference of basal values of the biomarkers might be due to the difference of the functions in the two tissues. Gills are in direct contact with water (environmental contaminants or other stressors) and the digestive gland is a storage organ important in the metabolic process of contaminants bound to particulate matter or to feeding processes (Maria and Bebianno, 2011). At present, the research on biomarkers based on shellfish mainly includes organization detoxification enzyme activity and biological macromolecules damage effects (Company et al., 2008; Binelli et al., 2009), showing that the toxicity effect indexes are closely related to the concentration of environment contaminant. But whether these indicators can be used as biomarkers requires further evaluation and validation. Cheung et al. (2001) suggested that there should be a clear dose–effect relationship between the biomarker changes and the concentration of environment contaminant. Oost et al. (2003) made the same point of view about biomarker evaluation standard. In our research, according to the correlation analysis, the EROD activity, antioxidant system and DNA damage index (F value) showed obvious linear relationships. So the EROD, GST activity, LPO and DNA damage may be suitable biomarkers to evaluate the toxicity of EDS.

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5. Conclusion It is deduced that EDS could be biotransformed and subsequently caused changes in EROD and GST activities and antioxidants damage on R. philippinarum. Because of the sensitivity to low EDS concentrations, EROD activity, antioxidant enzymes, especially GST activity could be potential indicators to assess the toxicity of environmental pollutants in organisms. Besides, there was a significant dose-dependent in LPO and DNA strand break. We conclude that the EROD, GST activity, LPO and DNA damage may be suitable biomarkers to evaluate the toxicity of EDS and that R. philippinarum is a suitable sentinel species for biomonitoring EDS toxicity. However, the metabolism process and toxicity mechanism of EDS are not clear. The exploration of the relationship among the indexes in the present work provides a mechanistic explanation for mechanisms of endosulfan toxicity and may represent an investigative area of endosulfan toxicity to organisms. The authors suggest that further studies are needed to explore the toxic mechanism of endosulfan.

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