Antioxidant defenses and oxidative stress parameters in tissues of mud crab (Scylla serrata) with reference to changing salinity

Comparative Biochemistry and Physiology, Part C 151 (2010) 142–151

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Comparative Biochemistry and Physiology, Part C j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / c b p c

Antioxidant defenses and oxidative stress parameters in tissues of mud crab (Scylla serrata) with reference to changing salinity Biswaranjan Paital a, G.B.N. Chainy a,b,⁎ a b

Department of Zoology, Utkal University, Bhubaneswar-751 004, India Department of Biotechnology, Utkal University, Bhubaneswar-751 004, India

a r t i c l e

i n f o

Article history: Received 24 August 2009 Received in revised form 22 September 2009 Accepted 23 September 2009 Available online 29 September 2009 Keywords: Ascorbic acids Catalase DNA unwinding test Glutathione peroxidase Hydrogen peroxide Lipid peroxidation Glutathione reductase Protein carbonylation Salinity Scylla serrata Superoxide dismutase

a b s t r a c t The effects of salinity (10, 17 and 35 ppt) on O2 consumption, CO2 release and NH3 excretion by crabs and oxidative stress parameters and antioxidant defenses of its tissues were reported. An increase in salinity caused a decrease in O2 consumption and CO2 release and an increase in ammonia excretion by crabs. Lipid peroxidation, protein carbonyl, H2O2 levels and total antioxidant capacity of the tissues elevated significantly at 35 ppt salinity except in abdominal muscle where H2O2 content was low. Ascorbic acid content of tissues was higher at 17 ppt salinity than at 10 and 35 ppt salinities. With increasing salinity, a gradual decrease in SOD, an increase in catalase, no change in GPx and a decrease followed by an increase in GR activities were recorded for abdominal muscle. While for hepatopancreas, an increase followed by a decrease in SOD and catalase, decrease in GPx and GR activities were noticed with increasing salinity. In the case of gills, a decrease followed by an increase in SOD, a decrease in catalase and GPx and an increase in GR activities were noted when the salinity increased from 10 ppt to 35 ppt. These results suggest that salinity modulation of oxidative stress and antioxidant defenses in Scylla serrata is tissue specific. © 2009 Elsevier Inc. All rights reserved.

1. Introduction Salinity is one of the abiotic variables of estuarine ecosystem that fluctuates widely during the year and thereby, plays a significant role in the physiology of inhabiting invertebrate species (Schmidt-Nielsen, 1997). Consequently, various adaptations at biochemical, physiological and behavioral levels are observed in the estuarine inhabiting organisms to cope with the changing environmental salinity. Mud crab, an euryhaline species, generally inhabits intertidal zones and estuaries throughout the Indo-Pacific region (Chen and Chia, 1996a) including the Chilika lagoon of India (19° 28′ and 19° 54′ N and 85° 05′ and 85° 38′ E). It is an important commercial invertebrate species of the estuarine ecosystem. Several aspects of Scylla serrata are reported to be modulated by environmental salinity and temperature (Hill, 1974; Chen and Chia, 1996a,b; Hai et al., 1998; Ruscoe et al., 2004). It is also reported that larval survival, growth and development (Hill, 1974; Hamasaki 2003), oxygen uptake and ammonia excretion (Chen and Chia, 1996b) of S. serrata are considerably influenced by salinity. A seasonal variation in fat and protein content of abdominal

⁎ Corresponding author. Department of Zoology, Utkal University, Bhubaneswar-751 004, India. Fax: +91 674 2587389. E-mail address: [email protected] (G.B.N. Chainy). 1532-0456/$ – see front matter © 2009 Elsevier Inc. All rights reserved. doi:10.1016/j.cbpc.2009.09.007

muscles of the species was attributed to the changing salinity of the water (Zafar et al., 2004). Utilization of oxygen by a variety of biochemical reactions in aerobic organisms is the origin of partially reduced oxygen species that are commonly known as reactive oxygen species (ROS; such as superoxide radical, hydroxyl radical and hydrogen peroxide). The ROS being highly cytotoxic in nature produce deleterious effects on biomolecules. Cellular antioxidant defense system is one of the important biochemical strategies that give protection to cells against deleterious effects of endogenous ROS by keeping their level relatively low. Cellular antioxidant defense system comprises of both nonenzymatic small antioxidant molecules (such as reduced glutathione (GSH), ascorbic acid (AA), carotenoids etc) and a cascade of enzymes namely superoxide dismutase (SOD, EC1.15.1.1), catalase (CAT, EC 1.11.1.6) and glutathione peroxidase (GPx, EC1.11.1.9) (Halliwell and Gutteridge 2001). Antioxidant enzymes are interdependent in nature and subject to variations due to intrinsic biological cycles, ambient physico-chemical environment and anthropogenic pollutants (Sheehan and Power, 1999). There has been much interest in recent years in enzyme activities involved in metabolism of ROS in crustaceans such as prawns (Dandapat et al., 2000) and crabs (Gamble et al., 1995; Orbea et al., 2002; Kong et al., 2004, 2005, 2007, 2008; Vijayavel et al., 2004, 2005). There are reports that describe about the manifestation of

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physiological and biochemical variations especially that of the O2 consumption of euryhaline crabs at altered salinity conditions (Findley et al., 1978; Chen and Chia, 1996b; Robles et al., 2002). Since O2 uptake is directly related with the oxidative metabolism of animals, therefore, we hypothesize that changing salinity may channelize with altered oxidative stress and antioxidant defense of euryhaline crab S. serrata. In this context, it is noteworthy to mention that information on responses of antioxidant defenses of S. serrata to changing salinity is lacking. Therefore, the present study was undertaken to investigate about the effect of salinity on (i) levels of activities of enzymes of antioxidant defense system (such as superoxide dismutase, catalase, glutathione peroxidase and glutathione reductase: GR); (ii) levels of non-enzymatic small antioxidant molecule (such as ascorbic acid and non-protein sulphydriles: -SH) and total antioxidant capacity; and (iii) levels of oxidative stress parameters (such as lipid peroxidation: LPx, protein carbonylation: PC and hydrogen peroxide: H2O2) in tissues of mud crabs under laboratory conditions. Also rate of oxygen consumption and rate of release of carbon dioxide and ammonia by mud crabs were determined at different salinity points. Besides, alkaline DNA unwinding level of the tissues was also checked to detect the effect of salinity on DNA damage in the tissues of crabs. Results of the present study were not only expected to provide additional contribution to the emerging field of invertebrate oxidative stress but also will help in understanding the physiological and biochemical basis of adaptation of S. serrata to changing salinity. In addition, the possible use of the results in the future to act as biomarkers in ecotoxicological investigations and aquaculture of mud crabs cannot be ruled out. Moreover, this is the first report regarding oxidative metabolism of an estuarine invertebrate species with respect to changing salinity.

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2.3. Experimental protocol The crabs were disinfected by dipping them into 17 ppt saline water with 500 ppm (parts per million) KMnO4 for 5–7 min and then they were acclimated in 17 ppt saline water without KMnO4 for 24 h in the laboratory. During this acclimatization period about 13% mortality rate was observed. Only active crabs were selected, divided into three groups containing 12 crabs each and released into three glass aquaria (75 cm × 30 cm × 30 cm) with low (10 ppt), medium (17 ppt) and high (35 ppt) salinities of artificially made sea water. Artificial sea water of 35 ppt was made according to Robinson (1954) and diluted to 17 ppt and 10 ppt with tap water supplied by Municipality Corporation, Bhubaneswar, when necessary. The aquaria had 5 cm high bed of sand that were pre-treated with KMnO4 followed by extensive washing with tap water and water level was high enough to keep the crabs in submerged condition. All aquaria were continuously aerated and covered with opaque plastic covers to make the least disturbance to the animals. Saline water was changed daily at 18.00 h and the crabs were exposed to 12 h:12 h natural light and day periods throughout the experimentation. The crabs were fed once daily at night (at 19.00 h) with fresh chick liver pieces. Temperature and salinity of water of aquaria were recorded once daily (at 8.00 h) with specific electrodes (μP based soil and water analysis kit, Esico. Co., New Delhi, India). The mean salinity and temperature of aquaria water of low, medium and high salinity during experimentation period of 21 days were 10.07 ± 0.60 ppt and 29.2 ± 2.0 °C, 17.51 ± 0.73 ppt and 29.31 ± 1.99 °C, and 35.39 ± 0.66 ppt and 29.7 ± 1.6 °C, respectively. The mortality rate of the crabs in the aquaria was around 25%, 17% and 25% at 10 ppt, 17 ppt and 35 ppt, respectively. 2.4. Measurement of O2 consumption, CO2 release and NH3 excretion

2. Materials and methods 2.1. Chemicals Thiobarbituric acid, cumene-hydroperoxide, Sephadex G-25, BSA, glutathione reductase, horse radish peroxidase, homovanilic acid, dithiobis nitrobenzoic acid, triton-X 100, Hoechst dye number 33258, tert-butyl hydroperoxide, 2, 2-Diphenyl-1-picryl hydrazyl and Tris–Cl were purchased from Sigma-Aldrich Chemical Company, USA. Proteinase-K and RNAse were obtained from Fermentas Life Sciences, USA. Dithiothretol was supplied by Biogene, USA. Sucrose, phenolnaphtaline, phenol, EDTA, riboflavin, phenylmethane sulphonyl fluoride (PMSF), hydroxylamine hydrochloride, L-methionine, oxidized and reduced glutathione, sodium azide, NADPH, chloroform and sodium molybdate were purchased from SISCO Research Laboratory, Mumbai, India. Sodium dodecyl sulphate, H2O2 and isoamyl alcohol were obtained from SD Fine chemicals, Mumbai, India. All other chemicals used were of analytical grade.

2.2. Animals Adult male mud crab (S. serrata, Forskal) at intermoult stage weighing 65.95 ± 8.49 g of body mass were collected from the Arakhakuda region of the Chilika lagoon (19° 28′ and 19° 54′ N and 85° 05′ and 85° 38′ E) of Orissa, India during May 2007 and were transported to the laboratory in gunny bags containing see weeds. The transport time from the site of collection to the laboratory was around 3 h. The salinity of Chilika lagoon varies throughout the year ranging from nearly 10 ppt in the rainy season (July to September), 17 ppt during winter (December and January) and 35 ppt during summer (April to June) with a temperature variation from 18 °C (winter) to 32 °C (summer) (Mohapatra et al., 2007; Panigrahi et al., 2007).

One group of 10 male crabs was maintained at 10 ppt salinity for three weeks after acclimatization to 17 ppt salinity in laboratory conditions as described above in order to study O2 consumption, CO2 release and NH3 excretion. The same group of crabs was then shifted to 17 ppt salinity for three weeks, followed by 35 ppt salinity for another three weeks. O2 consumption, CO2 release and NH3 excretion by the crabs were measured at each salinity point prior to shifting from one salinity to another. A respiratory chamber (RC) was designed with a cylindrical glass jar (volume 3.121 L) fitted with a hard plastic net at 7 cm height at its bottom. The RC was placed on a magnetic stirrer and a magnetic bid was kept under the plastic net to circulate the water within the jar. The upper side of the RC was equipped with an O2 electrode connected with a monitor (µP based soil and water analysis kit, Esico. Co., New Delhi, India). One crab was introduced into the chamber filled with freshly O2 saturated artificial saline water of required salinity. Prior to releasing the crabs into the RC, each was acclimated in a similar chamber for 10 min with the water of the same salinity. During experiments, the RC was sealed to prevent the diffusion of atmospheric O2 into the chamber. Fall of O2 concentration in the chamber was recorded at 15 min interval up to 60 min and the result was expressed as mg O2 consumed/100 g body mass/h. At the end of 1 h, a 500 mL water sample was drawn into an air tight dark bottle for CO2 measurement and 4 mL water into a microfuge tube for NH3 measurement. Before adding water into the RC, water samples were collected to assess initial O2, CO2 and NH3 present before the experiment and the values were deduced from the respective resulted values after the end of the experiment. Free CO2 was measured by titrating 50 mL of water sample with 0.1 mL of phenolphthalein indicator against N/10 NaOH according to APHA (1985). The end point of titration was achieved when a pink color was developed in the solution. The result was expressed as mg of CO2 released/100 g body mass/h. NH3 in the water sample was quantified according to Russell (1944). To 1.5 mL of water sample 0.1 mL of manganous salt

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was added followed by 1 mL of 25% alkaline phenol and 0.5 mL of hypochlorite solution. The mixture was gently rotated and boiled for 5 min, cooled and was centrifuged at 200 g for 1 min at 25 °C. Absorbance of the color of solution was measured at 625 nm. NH3 concentration was calculated from its standard curve and the results were expressed as mg of NH3 excreted/100 g body mass of crab/h. 2.5. Tissue collection and sub cellular fractionation Animals were sacrificed by removing their carapace from their abdomen with a jerk and the hepatopancrea, gills and abdominal muscle tissues were dissected out quickly. Tissues were washed in ice-cold normal saline (0.67%, w/v), blotted, flash frozen in liquid nitrogen and stored at −80 °C. A 10% or 20% (w/v) homogenate of tissues were prepared at 4 °C in the homogenizing buffer (50 mM Tris–Cl, 1 mM EDTA, I mM DTT, 0.5 mM sucrose, 150 mM KCl and 1 mM PMSF, pH 7.8) with the help of a Potter–Elvejhem type, motor driven glass Teflon homogenizer at 250 rpm speed with 7–8 up and down strokes whereas due to rigidity, gill tissues were homogenized in pre-cooled mortar and pestle. The crude homogenates were centrifuged at 1,000g for 10 min at 4 °C in a cooling centrifuge (REMI, Model C-24) to sediment nuclei and tissue debris. The supernatant fractions (post-nuclear fraction: PNF) were centrifuged at 10,000g for 10 min at 4 °C to obtain the clear supernatant which was referred as post-mitochondrial fraction (PMF). 2.6. Biochemical determinations Protein estimation in PNF and PMF samples were made according to the method of Bradford (1976) using bovine serum albumin (BSA) as a standard. 2.6.1. Measurement of lipid peroxidation For LPx estimation, post-nuclear fractions were diluted with a cold KCl (1.15%; w/v) solution and the level of thiobarbituric acid reactive substances (TBARS) was measured according to the method of Ohkawa et al. (1979). Results were calculated from the molar extinction coefficient of TBARS as 1.56 × 105 M− 1 cm− 1 and were expressed as nmoles of TBARS formed per mg protein. To reduce the autoperoxidation of lipids during incubation, butylated hydroxytoluene (0.02%, w/v) was added to samples prior to assay. 2.6.2. Measurement of protein carbonylation Protein carbonylation content was measured in post-nuclear fractions of tissue samples according to the method of Levine et al. (1994). Carbonyl content was calculated from its molar absorption coefficient as 22,000 M− 1 cm− 1 and results were expressed as nmol protein carbonyl per mg protein. 2.6.3. Measurement of hydrogen peroxide Hydrogen peroxide (H2O2) content in tissue samples were measured spectrofluorimetrically according to Anguelov and Chichovska (2004). In brief, tissue samples were homogenized in 50 mM phosphate buffer containing 2 mM EDTA, pH 7.6 (PBE) at 4 °C. The crude homogenate (10%, w/v) was centrifuged at 1,000 g for 10 min at 4 °C to obtain PNF. The samples were pre-incubated for 1 h at 25 °C in the assay mixture (0.6 mL) containing 2 U/mL horseradish peroxidase (HRP) and, 200 µM homovanillic acid (HVA) in PBE. Then the mixture was added with 0.9 mL of PBE and 0.375 mL glycine-NaOH (0.1 M) buffer, pH 12. The HRP mediated dimer product of HVA by H2O2 was measured with a fluorescence spectrophotometer (Hitachi, Japan, Model F 2500). Instrument specifications during measurement were 5 nm of both extinction and emission slit widths, 400 PMT voltages and 312 nm and 420 nm of extinction and emission wavelengths, respectively. Pure H2O2 was taken as standard and the results were expressed as µg of H2O2 per g of wet tissue. To calculate the internal

loss of H2O2 during sample preparation, muscle, gill and hepatopancreas of a group of five crabs were processed as described above and precipitated immediately with 5% trichloro acetic acid and centrifuged at 1, 000 rpm for 5 min to collect the supernatant. The samples were then neutralized with tris base and H2O2 was measured as described above. A loss of 2.12 ± 0.11, 1.51 ± 0.07 and 1.56 ± 0.12 fold of H2O2 was observed in TCA precipitated sample of muscle, hepatopancreas and gills, respectively. Therefore, the observed data obtained for nonprecipitated samples were multiplied with the corresponding factors for calculation of actual H2O2 content in the tissues. 2.7. Determination of antioxidant enzyme activities SOD, GPx and GR activities were measured in sephadex G-25 column elutes of PMF of tissue samples. In brief, 200 µl of PMF sample was loaded to a pre-cooled Sephadex G-25 column prepared in a 1 mL syringe and equilibrated with the homogenizing buffer. The column was centrifuged at 2000 rpm for 2 min at 4 °C to remove the buffer prior to sample loading. After loading of the sample, the syringe was put in a cleaned cool centrifuge tube and centrifuged at 2000 rpm for 2 min at 4 °C and enzyme activities were assayed in the elute. Activities of all enzymes were measured at 25 °C. 2.7.1. Superoxide dismutase SOD activity in samples was measured according to the method of Das et al. (2000). The principle of the assay is based on the estimation of superoxide scavenging capacity of samples. Superoxide radicals were generated in the assay system by photoreduction of riboflavin. In brief, the assay mixture (1.58 mL) contained hydroxylamine hydrochloride (0.47 mM), L-methionine (0.9 mM), Triton-X 100 (0.026%), and riboflavin (2.5 µM) in 100 mM Tris buffer, pH 8. The reaction was started by exposing the mixture to two 20 W fluorescent lamps (fitted parallel to each other inside an aluminum coated wooden chamber) for 10 min at 25 °C. The nitrite produced by superoxide radicals was fixed by adding 1 mL of Griess reagent and the intensity of the color formed was measured at 540 nm against an appropriate blank. Enzyme activity was expressed in U (unit) per mg protein where one unit is defined as the amount of protein that causes 50% inhibition of nitrite production. Final SOD activity was calculated by subtracting the value obtained for boiled (95 °C for 30 min) samples from the corresponding unboiled samples. 2.7.2. Catalase CAT activity was measured in the samples according to the method of Aebi (1974) by monitoring the decrease in absorbance of H2O2 at 240 nm. Prior to the assay, triton-X 100 (1%, v/v) and ethanol (1%) were added to the samples to increase the observable CAT activity by releasing it from peroxisomes and to prevent formation of the inactive complex (complex-II) of CAT with H2O2, respectively (Cohen et al., 1970). In brief, fall of absorbance of 25 mM H2O2 was recorded by an UV-VIS spectrophotometer at 240 nm (Varian, CARY 100) in the assay volume of 3 mL phosphate buffer (50 mM, pH 7). Enzyme activity was calculated from the extinction coefficient of H2O2 as 43.6 M− 1 cm− 1 and was expressed as nKat per mg protein. (One Kat is defined as one mole of H2O2 consumed per second per mg protein). 2.7.3. Glutathione peroxidase GPx activity was measured according to the method of Paglia and Valentine (1967) with cumene-hydroperoxide as the substrate. In brief, 1 mL of the assay system contained 50 mM phosphate buffer (pH 7.6), 30 mM glutathione, 30 mM sodium azide, 4.5 mM NADPH, 10 U/mL glutathione reductase and 7.5 mM cumene-hydroperoxide. All the reagents were prepared in phosphate buffer. Utilization of NADPH by glutathione system was recorded at 340 nm and results were calculated from the extinction coefficient of NADPH as

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6.22 mM− 1 cm− 1 and expressed as nmol of NADPH oxidized per min per mg protein. 2.7.4. Glutathione reductase GR activity was measured according to the method of Massey and Williams (1965) in which the rate of conversion of GSSG to GSH was estimated by monitoring oxidation of NADPH in the assay system at 340 nm. In brief, 1 mL of the assay system contained phosphate buffer (50 mM, pH 7.6), 120 mM oxidized glutathione and 4.5 mM NADPH. The enzyme activity was calculated from the extinction coefficient of NADPH. The results were expressed in nmol of NADPH oxidized per min per mg protein. 2.8. Alkaline DNA unwinding test Alkaline DNA unwinding assay was performed fluorimetrically according to Shugart (1988a,b). In brief, DNA was isolated from tissue samples by preparing a 10% (w/v) homogenate (50 mg tissue) in lysis buffer (50 mM tris,100 mM EDTA, 0.5% SDS, pH 8) with the help of a hand homogenizer in 1.5 mL microfuge tubes at room temperature. RNAse (10 µg/mL) was added to it and was incubated for 1 h followed by proteinase-K (100 µg/mL) treatment for another 1 h. The samples were then mixed with equal volumes of phenol and centrifuged at 12,000 g for 10 min at 4 °C. The aqueous phase was transferred to a new tube and an equal volume of phenol:chloroform:isoamyl alcohol (25:24:1) was added to it and again centrifuged at 12,000 × g at 4 °C for 10 min. The aqueous phase was extracted with an equal volume of chloroform followed by centrifugation at 12,000 g for 10 min at 4 °C. The aqueous phase was again transferred to another new tube and added with a double volume of absolute alcohol and sodium acetate (0.3 M) and kept at − 40 °C for 30 min. The sample was then centrifuged at 12,000 g for 10 min at 4 °C to obtain DNA pellet. The pellet was washed with 75% ethanol at 12,000 g at 4 °C and dissolved in 20 µL of Tris (10 mM) EDTA (1 mM) buffer, pH 8. DNA was quantified by taking absorbance of the sample at 260 nm at proper dilution with double distilled water. Around 5 µg of DNA was mixed with 1 µg of Hoechst dye (no. 33258) for DNA unwinding assay. Briefly, for each sample, three sets of incubation were performed at 4 °C, 38 °C and 80 °C with 50 µL of 0.05 N NaOH solution for 15 min. The mixture was then neutralized with 50 µl of 0.05 N HCl solution. Then 5 µL of SDS (0.2%,w/v) containing 2 mM EDTA was added, mixed and followed by 100 μL of Hoechst dye (1 µg) and kept for 15 min at room temperature. The volume of reaction mixture was adjusted to 3 mL with 0.2 M PB, pH 6.9. Fluorescence of the sample was measured at 360 nm excitation and 450 nm emission wavelengths and 5 nm for both the slit widths in a fluorescence spectrophotometer (Hitachi, Japan, Model F-2500). The results were calculated from the fluorescence value of three incubations of each sample and expressed as F-value, where F-value = (F 38 °C–F 80 °C) / (F 4 °C–F 80 °C) and F indicated fluorescence of sample at the respective temperatures. The F-value closer to 1.0 or less than 1.0 indicates more intact DNA or more unwound DNA, respectively. 2.9. Measurement of small antioxidant molecules and total antioxidant capacity 2.9.1. Ascorbic acid 1 mL of the post-mitochondrial fraction samples was precipitated with ice-cold TCA (5%, w/v) immediately after fractionation. TCA precipitated samples were centrifuged at 10,000 g for 10 min at 4 °C to obtain the supernatant. Ascorbic acid was estimated according to the method of Mitsui and Ohta (1961) by monitoring the reduction of phosphomolybdate by ascorbic acid. Briefly, 1 mL of assay mixture contained 0.5 mL sample, 0.2 mL Na-molybdate (0.66%), 0.2 mL H2SO4 (0.05 N) and 0.1 Na-phosphate (0.025 mM) and was kept at 60 °C water bath for 40 min. It was then cooled under running water and

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centrifuged. Absorbance of the clear supernatant was taken at 660 nm. AA was taken as standard and the results were expressed in ng of ascorbic acid per g wet tissue mass. 2.9.2. Non-protein -SH group Non-protein -SH content was determined according to the method of Sedlak and Lindsay (1968). In brief, 1.0 mL of the post-mitochondrial fractions of samples were precipitated with ice-cold TCA (5%, w/v) immediately after fractionation and centrifuged at 10,000g for 10 min at 4 °C. To 0.5 mL of sample, 80 µL of saturated Tris base was added to make the pH of the solution between 8 and 9. To it, 50 µL of 0.01 M dithiobis nitrobenzoic acid (DTNB) was added and the color product was measured at 412 nm against a suitable blank. Results were calculated from the standard curve of GSH and expressed in nmol of -SH content per g of wet tissue mass. 2.9.3. Determination of total antioxidant capacity Tissue samples were homogenized in 50 mM phosphate buffer containing 2 mM EDTA, pH 7.6 (PBE) for measurement of total antioxidant capacity of tissues in the form of DPPH (2, 2-diphenyl-1picryl hydrazyl) scavenging activity. The crude homogenate (10%, w/v) was centrifuged at 1000 g for 10 min to obtain PNF. DPPH scavenging assay was performed according to Marxen et al. (2007) in the aqueous phase of PNF. The results were expressed as % of decrease in absorbance by 100 µL of PNF from a control set (without sample) at 515 nm. The calculation was performed as % of inhibition = (Abs control−Abs of sample)/Abs control × 100 (Sing et al., 2002). 2.10. Statistical analysis Results were expressed as mean± standard deviation. Means of O2, CO2 and NH3 measurements were compared and analyzed for differences using repeated ANOVA (StatPages.org.). Means of biochemical estimations were compared and analyzed by one way ANOVA analysis followed by Duncan's new multiple range test. Difference among the means was considered significant at P ≤ 0.05 levels. 3. Results 3.1. Oxygen consumption, carbon dioxide release and ammonia excretion Table 1 shows changes in oxygen consumption, carbon dioxide release and ammonia excretion by the crabs when they were stepwise shifted from 10 ppt to 17 ppt and from 17 ppt to 35 ppt salinity levels. Significant reductions (18% and 22%) in oxygen consumption by the crabs were recorded when the crabs were transformed from 10 ppt to 17 ppt salinity and from 17 ppt to 35 ppt salinity levels, respectively. Although release of carbon dioxide by the crabs decreased to 64% and

Table 1 Effect of salinity on O2 consumption, CO2 release and NH3 excretion of mud crab Scylla serrata. Parameters

10 ppt group

17 ppt group

35 ppt group

O2 consumption (mg/100 g body mass/h) CO2 released (mg/100 g body mass/h) NH3 excreted (mg/100 g body mass/h)

15.13 ± 3.03

12.46 ± 2.37

9.75 ± 0.52

63.49 ± 14.50

22.58 ± 7.90

14.37 ± 3.19

0.55 ± 0.39

4.02 ± 0.96

6.04 ± 1.42

Each value represents the mean ± SD of 10 individuals. Individual data for each parameter for different salinities were subjected to repeated ANOVA. Results of which indicated very strong evidence against null hypothesis among the means at P value < 0.001 level.

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Table 2 Effect of salinity on lipid peroxidation (LPx), protein carbonyl (PC) and hydrogen peroxide (H2O2) and DNA unwinding value of gills and abdominal muscle (M), hepatopancreas (HP) and gills (G) of mud crab Scylla serrata. Parameters LPx (nmol TBARS/mg protein)

PC (nmol PC/mg protein)

H2O2 (μg/g wet tissue)

DNA UW (F-value)

10 ppt group M HP G M HP G M HP G M HP G

a

0.04 ± 0.01 0.90 ± 0.23a 0.13 ± 0.04a 1.82 ± 0.72a 2.46 ± 0.13a 1.69 ± 0.26a 0.91 ± 0.04a 8.19 ± 1.51a 1.22 ± 0.48a 0.76 ± 0.14 0.81 ± 0.12 0.80 ± 0.89

17 ppt group b

0.1 ± 0.02 0.89 ± 0.22a 0.18 ± 0.06b 2.15 ± 0.89a 2.67 ± 0.31a 2.19 ± 0.71b 0.97 ± 0.04b 7.11 ± 1.36a 1.28 ± 0.02a 0.88 ± 0.10 0.85 ± 0.08 0.81 ± 0.09

35 ppt group 0.24 ± 0.02c 1.32 ± 0.16b 0.28 ± 0.05c 6.08 ± 1.21b 3.29 ± 0.60b 2.60 ± 0.71b 0.80 ± 0.04c 10.05 ± 1.27b 1.58 ± 0.13b 0.85 ± 0.12 0.84 ± 0.06 0.80 ± 0.08

Each value represents the mean ± SD of 9 individuals. Superscripts (a, b and c) indicate statistical significance within the groups at P ≤ 0.05 level. Groups with different superscripts are statistically different from each other.

37% when they were stepwise transformed from 10 ppt to 17 ppt and from 17 ppt to 35 ppt salinity levels, respectively; the difference between values of 17 ppt and 35 ppt was not statistically significant. On the other hand, it was noticed that total ammonia excretion by the crabs elevated to around 630% and 50% when they were shifted from 10 ppt to 17 ppt and from 17 ppt to 35 ppt salinity levels, respectively.

magnitude of change in protein carbonyl content was different from one tissue to the other. Protein carbonyl content of abdominal muscle of the crabs at 35 ppt was 234% and 182% higher than 10 ppt and 17 ppt salinity groups, respectively. It was further noticed that the protein carbonyl content of hepatopancreas of crabs of the 35 ppt group was 53% and 23% higher than that of 10 ppt and 17 ppt groups, respectively. Protein carbonyl content of gills of the crabs was 29% and 53% higher at 17 ppt and 35 ppt groups in comparison with 10 ppt salinity group, respectively. 3.2.3. Hydrogen peroxide content A small but a significant decrease (12%) in hydrogen peroxide content of abdominal muscle of the crabs of 35 ppt salinity group was recorded in comparison with 10 ppt salinity group. On the other hand, a significant 22% and 41% increase in hydrogen peroxide content of hepatopancreas of the crabs of 35 ppt group were recorded in comparison with 10 ppt and 17 ppt groups, respectively. Similarly, 29% and 23% elevations in hydrogen peroxide content of gills of the crabs of 35 ppt group was noted in comparison with 10 ppt and 17 ppt groups, respectively. 3.2.3. DNA unwinding test The DNA unwinding value of the three tissues of crabs studied did not change in response to changing salinity. 3.3. Antioxidant enzymes

3.2. Oxidative stress parameters Table 2 depicts changes in various oxidative stress parameters in abdominal muscle, hepatopancreas and gills of adult male crabs maintained at different salinities.

Table 3 shows changes in levels of various antioxidant enzymes of abdominal muscle, hepatopancreas and gills of adult male crabs in response to different salinities.

3.2.1. Lipid peroxidation A gradual elevation in LPx value of abdominal muscle of the crabs was recorded in response to increasing salinity. Lipid peroxidation level in abdominal muscle of the crabs was 150% and 500% higher at 17 ppt and 35 ppt salinity in comparison to 10 ppt salinity, respectively. Although the LPx value of hepatopancreas of the crabs did not vary significantly between 10 ppt and 17 ppt salinity levels, its value was 45% higher at 35 ppt salinity level than that of 10 ppt and 17 ppt. In case of gills of the crabs, LPx value was 45% and 110% higher at 17 ppt and 35 ppt in comparison with 10 ppt salinity level, respectively.

3.3.1. Superoxide dismutase It was noticed that the level of SOD in abdominal muscle of the crabs decreased gradually as the level of salinity increased. The activity of the enzyme in abdominal muscle tissue of the crabs diminished 55% and 85% in 17 ppt and 35 ppt groups in comparison with 10 ppt group, respectively. In case of hepatopancreas, the enzyme activity of the crabs at 17 ppt salinity was observed to be higher (56%) than that of 10 ppt group. But a significant 22% and 51% decrease in its level were recorded in the tissues of crabs of 35 ppt groups in comparison with 10 ppt and 17 ppt groups, respectively. On the other hand, the level of the enzyme was 22% lower in gills of the crabs of 17 ppt groups in comparison with 10 ppt and 35 ppt groups, respectively.

3.2.2. Protein carbonyl content In all the three tissues viz. abdominal muscle, hepatopancreas and gills of the crabs, protein carbonyl content was significantly higher at 35 ppt salinity in comparison with 10 ppt salinity. However, the

3.3.2. Catalase The response of catalase to different salinities varied from one tissue to another. Significant 119% and 155% increments in the level of catalase of abdominal muscle of the crabs of 35 ppt group were noted

Table 3 Effect of salinity on superoxide dismutase (SOD), catalase (CAT), glutathione peroxidase (GPx) and glutathione reductase (GR) activities of abdominal muscle (M), hepatopancreas (HP) and gills (G) of mud crab Scylla serrata. Parameters SOD (U/mg protein)

CAT (nKAT/mg protein)

GPx (nmol of NADPH oxidized/min/mg protein) GR (nmol of NADPH oxidized/min/mg protein)

M HP G M HP G M HP G M HP G

10 ppt group

17 ppt group

35 ppt group

2.64 ± 0.35a 4.31 ± 0.63a 1.42 ± 0.50a 62.58 ± 20.20a 513.75 ± 89.59a 489.02 ± 61.19a 18.29 ± 4.80 140.0 ± 13.02a 65.69 ± 18.04a 0.32 ± 0.11ab 1.41 ± 0.42a 0.43 ± 0.12a

1.20 ± 0.18b 6.74 ± 0.73b 0.83 ± 0.10b 53.54 ± 19.98a 721.0 ± 93.85b 268.74 ± 38.17b 13.98 ± 5.15 205.57 ± 86.73b 45.84 ± 20.42b 0.19 ± 0.06a 1.17 ± 0.62a 0.50 ± 0.33a

0.38 ± 0.14c 3.33 ± 0.79c 1.43 ± 0.26a 136.75 ± 23.09b 535.43 ± 54.18a 69.66 ± 21.06c 13.55 ± 6.59 97.16 ± 14.41c 45.58 ± 5.47b 0.46 ± 0.23b 0.71 ± 0.26b 0.70 ± 0.13b

Each value represents the mean ± SD of 9 individuals. Superscripts (a, b and c) indicate statistical significance within the groups at P ≤ 0.05 level. Groups with different superscripts are statistically different from each other.

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in comparison with 10 ppt and 17 ppt groups, respectively. In case of hepatopancreas, a 40% increase in enzyme activity was recorded for the crabs of 17 ppt than that of 10 ppt group. But the activity of the enzyme was observed to be low (25%) in the tissue of the crabs of 35 ppt group in comparison with 17 ppt group. The enzyme activity in gills of the crabs gradually decreased as the level of salinity increased. It was 85% and 74% lower in gills of the crabs of 35 ppt group in comparison with 10 ppt and 17 ppt groups, respectively. 3.3.3. Glutathione peroxidase Glutathione peroxidase activity of the abdominal muscle of the crabs did not change significantly in response to changing salinity. A significant increase (46%) in the enzyme activity of hepatopancreas was recorded in 17 ppt group than that of crabs of 10 ppt group. However, the enzyme level was 53% lower in the tissue of the crabs of 35 ppt group than that of 17 ppt group. It was observed that the enzyme level in gills of the crabs decreased to 30% in the case of 17 ppt group in comparison with 10 ppt group and did not change further in 35 ppt group. 3.3.4. Glutathione reductase The response of glutathione reductase to changing salinity varies from one tissue to another. Although the activity of the enzyme in abdominal muscle of the crabs did not vary significantly between 10 ppt and 17 ppt groups, it was significantly high (142%) in the crabs of 35 ppt groups than that of 17 ppt group. It was noticed that the enzyme activity was 49% and 39% lower in hepatopancreas of the crabs of 35 ppt in comparison with 10 ppt and 17 ppt groups, respectively. On the other hand, the enzyme activity was 62% and 40% higher in gills of the crabs of 35 ppt group than that of 10 ppt and 17 ppt groups, respectively. 3.4. Small antioxidants and total antioxidant capacity (Table 4) 3.4.1. Free -SH group The non-protein free -SH group content of abdominal muscle of the crabs of 35 ppt group was 20% higher than that of 10 ppt group. On the other hand, a small but significant decrease in -SH group content of hepatopancreas of 17 ppt group of the crabs was recorded in comparison wtih 10 and 35 ppt groups. A 67% decrease in -SH content of gills of the crabs of 35 ppt group was recorded in comparison with 10 ppt and 17 ppt groups. 3.4.2. Ascorbic acid It was observed that the ascorbic acid content of abdominal muscle of the crabs of 17 ppt group was 764% higher than that of 10 ppt group. The value decreased to 53% in the tissue of 35 ppt group than that of 17 ppt group. Although a similar pattern was also observed in the case of hepatopancreas, the increase in the case of 17 ppt group in comparison with 10 ppt was only 45%. A significant 12% and 16% decrease in ascorbic acid content of gills of the crabs of 35 ppt group were recorded in comparison with 10 and 17 ppt groups, respectively. 3.4.3. Total antioxidant capacity Result of the present experiment depicts that all the three tissues exhibited a similar pattern. DPPH scavenging activity was recorded to be the lowest at 10 ppt which reached to a maximum value at 35 ppt salinity. A positive correlation was observed between salinity and total antioxidant capacity of tissues of crabs (figure not given). 4. Discussion A distinct tissue specific pattern was noticed for antioxidant enzymes in S. serrata that were acclimatized in the laboratory at 17 ppt of salinity. Similar tissue variation regarding activities of antioxidant enzymes was reported for several other species belonging to

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phylum Arthropoda: M. rosenbergi (Dandapat et al., 2000), Charybdis japonica (Pan and Zhang, 2006), M. malcolmsoni (Arun and Subramanian, 1998), Chasmagnathus granulata (now Neohelice granulata) (Maciel et al., 2004), and Orconectus limosus (Borkovic et al., 2008). Our results regarding tissue distribution of antioxidant enzymes in the case of S. serrata are different from that reported for another estuarine crab species C. granulata. It is reported that gills of C. granulata have higher SOD (de Oliveira et al., 2005, 2006) and catalase (Maciel et al., 2004) activities than hepatopancreas. On the other hand, GR activity in gills of C. granulata was found to be low in comparison to hepatopancreas (Maciel et al., 2004). It is noticed that catalase and GPx activities in hepatopancreas of crayfish (Orconectus limosus) were high in comparison to gills and muscle (Borkovic et al., 2008). The same authors noted lower SOD and GR activities in hepatopancreas of the same species than gills and muscle tissues (Borkovic et al., 2008). This suggests that antioxidant defense strategies employed by tissues of crustaceans not only vary among themselves but also vary from one species to other. In the present study, crabs were acclimatized at 17 ppt of salinity in the laboratory before they were shifted to low i.e. 10 ppt or high i.e. 35 ppt salinity. It was observed that oxygen consumption by the crabs increased (around 30%) at 10 ppt than that of 17 ppt. On the other hand, oxygen consumption by the crabs decreased (around 30%) at 35 ppt in comparison with 17 ppt. Therefore, the possibility of experiencing a transition state of internal organs of crabs from normoxia to hypoxia (at 35 ppt) or to hyperoxia (at 10 ppt) could not be excluded as a consequence of decrease or increase in oxygen consumption rate, respectively. Our results are in good agreement with earlier results for other euryhaline crab species which exhibited elevated (Engel and Eggert, 1974; Engel et al., 1975; Sabourin, 1983) or reduced (Findley et al., 1978; Chen and Chia, 1996b; Robles et al., 2002) oxygen consumption rate when they were acclimated to low or high salinities. Increased consumption of oxygen by adult S. serrata at low salinity suggests that a higher energy demand by the organism may be due to active pumping of ions from environment to hemolymph. It is now a well established fact that most euryhaline crab species are hyperosmoregulators at low salinities, actively pumping ions from sea water into hemolymph (Mantel and Farmer, 1983; Lucu, 1990). Reduced rate of oxygen consumption and CO2 production by crabs at high salinity indicate low oxidative metabolism of the organism to regulate its hyperionic state. Ammonia, amino acids and urea are the three principal end products of protein metabolism that are released to the environment through gills in decapods (Regnault, 1987). Chen and Chia (1996a) reported that a shift in nitrogen excretion pattern from ammoniotelism to ureotelism takes place in S. serrata when crabs were moved from lower salinity to higher salinity. Chen and Chia (1996a) observed a decrease in ammonia-N release at high salinity. On the contrary, we have noted an increase in ammonia release by crabs at higher salinity. The discrepancies between our results and that of Chen and Chia (1996a) may be due to different experimental designing. Chen and Chia (1996a) have kept crabs in different salinities for 7 days only in their experiments. The duration of exposure to higher salinity may not be sufficient enough for crabs to acclimatize to salinities and may be one of the reasons for change in their nitrogen excretion pattern from ammonotelic to ureotelic. The increase release of ammonia at higher salinity may be attributed to high protein catabolism. The hepatopancreas in crustaceans is a major organ which is mainly associated with diverse metabolic activities ranging from supply of nutrition to ovary, to digestion and absorption (Sureshkumar and Kurup, 1999; Sang and Fotedar, 2004; Comoglio et al., 2008). It can be considered metabolically more active than gills and muscle tissue. It is difficult to explain the physiological basis for the observed decrease in activities of antioxidant enzymes in hepatopancreas of mud crabs at 10 ppt without any change in oxidative stress markers. It is possible that production of ROS in hepatopancreas is reduced at 10 ppt salinity

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which might have caused decrease in synthesis of antioxidant enzymes. It is interesting to note that a significant decrease in lipid peroxidation in abdominal muscle (60%) and gills (28%) and in protein carbonylation in gills (33%) was recorded at 10 ppt salinity. Observed increase in SOD activity in abdominal muscle and SOD, catalase and GPx in gills may be responsible for lowering LPx values. It contributes to the mechanism to understand that due to higher oxygen consumption by the organism at low salinity, these two tissues might have experienced higher oxygen tension and consumption which might have resulted in enhanced production of ROS. Consequently, levels of antioxidant enzyme were enhanced to counteract the ROS. In crabs, gills are primarily responsible for respiration, acid–base balance and osmotic and ionic regulation (Henry and Cameron, 1983; Henry, 1984; Lucu, 1990). Furthermore it has been noted that a battery of biochemical and physiological changes takes place in gills of euryhaline crabs in response to low salinity stress in order to maintain osmotic and ionic homeostasis (Piller et al., 1995). It has been reported that exposure of cells to hyperoxia resulted in increase generation of ROS by enzymatic and non-enzymatic mechanisms (Freeman and Crapo, 1981). The increased consumption of oxygen by gills in crabs at low salinity would have enhanced production of superoxide radicals. In order to neutralize the increase of superoxide radicals, tissue level SOD activity might have induced. The consequence of increased SOD activity would have resulted in more production of hydrogen peroxide. Hydrogen peroxide is highly toxic to cells and subsequently generates highly toxic and reactive hydroxyl radicals by classical Fenton reactions. Consequently, activities of both the enzymes elevated in gills of crabs to keep the hydrogen peroxide level at basal level. Generally, in almost all animals hydrogen peroxide is neutralized by catalase and GPx enzymes. Nevertheless, neutralization of hydrogen peroxide by muscles and gills of mud crabs was quite distinct. In the former tissue, where only GPx plays an active role in neutralizing hydrogen peroxide, both the enzymes are required for the later tissue. It is evident from the present study that enzymatic neutralization mechanism of ROS at low salinity is different among the tissues of mud crabs. It is well known that the rate of production of ROS at the mitochondrial level in many biological systems is proportional to oxygen consumption and to mitochondrial metabolic rate. But in our experiment the decrease in oxygen consumption in crabs at 35 ppt salinity have resulted in higher production of ROS. Several studies have clearly established that deprivation of oxygen to cells or tissues has resulted in generation of ROS (Vanden Hoek et al., 1997; Arroyo et al., 1987; Bolli et al., 1988; Ward, 2006). Several reasons have been described as the cause for increased generation of ROS during hypoxia such as residual oxygen level in tissues (de Oliveira et al., 2006) and switch over of biochemical reaction (Storey and Storey, 1990; Hochachka and Lutz, 2001). Limiting oxygen supply to crabs during high salinity will slow down the electron transport in the lower part of the ETC propagating ROS formation from downward electron carriers such as complex III ubiquinone (Brand 2000; Staniek and Nohl, 2000). There are several reports that confirmed increased production of ROS during hypoxia condition in marine invertebrates (Abele and Puntarulo, 2004). Among the three organs, hepatopancreas is the one most subjected to oxidative stress during transition of crabs from 17 ppt to 35 ppt salinity as evident by augmentation of LPx, PC and hydrogen peroxide levels. Lipid peroxidation and protein carbonylation are considered as consequences of oxidation of lipids and proteins by ROS (Halliwell and Gutteridge, 2001). Therefore, both parameters i.e. lipid peroxidation and protein carbonylation are considered as indices of tissue oxidative stress in almost all animals including that of estuarine invertebrates (Monserrat et al., 2007). In the present study, elevated levels of TBARS and PC contents in tissues of S. serrata at high salinity suggest induction of oxidative stress. Not much information is available in the literature on biochemical changes taking place in hepatopancreas of S. serrata in response to changing environmental factors in general and salinity in particular. A

significant decrease in SOD activity was observed in hepatopancreas of crabs at high salinity which is accompanied with reduced catalase and GPx activities may explain for the observed high hydrogen peroxide level in the tissue. GPx is also considered to remove lipid peroxides besides hydrogen peroxide (Fernandez-Diaz et al., 2006). The physiological significance of reduced activities of antioxidant enzymes at high salinity is difficult to explain at present. However, it may be attributed to low oxidative metabolism of the crabs as consumption of oxygen by crabs decreased during its exposure to high salinity. SOD activity of gills of crabs elevated when the crabs were subjected to high salinity condition. High salinity stress might have produced a large amount of ROS in gills which could have induced SOD to scavenge superoxide radicals. However, decrease in catalase activity along with GPx at high salinity might not be able to remove excessive hydrogen peroxide causing enhancement of lipid peroxide level in the tissue. Gills, the principal organ for respiration in crabs, are directly exposed to the surrounding environment. It also maintains the osmotic balance of body fluid. Several biochemical adaptations in gills of crabs to changing salinity are reported. For example, Na+–K+ ATPase (Cryptograpus angulatus: Lopez-Mananes et al., 2002) and carbonic anhydrase (Carcinus maenus: Henry et al., 2003) are reported to be influenced by environmental salinity. In comparison with hepatopancreas and gills, abdominal muscle is metabolically less active. SOD activity of the abdominal muscle decreased with high salinity. The present results provide an idea to support the fact that at high salinity, the metabolic activity of the abdominal muscle may be very low. Consequently, less amount of superoxide radicals are generated. Hence, the tissue needs low SOD activity. Decreased level of SOD activity along with enhanced level of catalase in abdominal muscle at high salinity may be the reason for reduced hydrogen peroxide level in the tissue. It is here to note that GR activity of the muscle increased in response to high salinity. GR regenerates GSH from GSSG which is formed by GPx. The biochemical adaptations of muscles of S. serrata to varying environmental salinities are relatively unknown except for a few studies which have shown change in alkaline phosphates activity (Pinoni and Lopez-Mananes, 2004) and mobilization of lipids from muscles (Luvizotto-Santos et al., 2003) of the euryhaline crab Cyrtograpus angulatus; enhanced arginine flux from muscle tissues of another crab species Callinectes sapidus (Holt and Kinsey, 2002) in response to low salinity and increased myosin ATPase activity of the flexor muscle of S. serrata in response to high salinity (Krishnamoorthy and Venkatramiah, 1969). Results of the present study give more insight to think about the molecular mechanism by which salinity modulates antioxidant enzyme levels in tissues of mud crabs. It is to note here that posttranslational regulation of SOD, catalase and GPx does not occur (Hermes-Lima, 2004). Brouwer et al. (2004) using microarrays constructed from cDNA and subtractive hybridization studies on hepatopancreas of Callinectes sapidus concluded that hypoxia resulted in down regulation of several genes. Therefore, alteration in activities of antioxidant enzymes in tissues of crabs in response to salinity must reflect alteration in their respective rate of expression and/or degradation is to be a matter of future investigation. In this study, the effect of salinity on two non-enzymatic antioxidant molecules in different tissues of mud crab is investigated. They are non-protein SH and ascorbic acid. GSH is also reported to act as an effective antioxidant in marine animal system besides acting as a substrate for GPx to neutralize hydrogen peroxide produced by SOD enzyme. Also it acts as a reductant in conjugation with electrophilic substances. Therefore, GSH level may reflect the detoxification ability of an organism (Kovacevic et al., 2008). It is here to mention that we have measured non-protein free -SH group in the samples which may not represent the true value for GSH. Nevertheless, in several studies measurement of -SH group in this fraction is used to represent GSH (Monteiro et al., 2009; Sk and Bhattacharya, 2006). A significant

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Table 4 Effect of salinity on free sulphydril group (-SH), ascorbic acid (AA) and total antioxidant capacity of abdominal muscle (M), hepatopancreas (HP) and gills (G) of mud crab Scylla serrata. Parameters -SH (nmol/g wet tissue) AA (ng/g wet tissue) Total antioxidant capacity⁎ (% of inhibition)

M HP G M HP G M HP G

10 ppt group

17 ppt group

35 ppt group

21.36 ± 1.21a 29.71 ± 3.83a 27.46 ± 2.76a 17.1 ± 3.45a 105.69 ± 28.11a 201.0 ± 20.17a 11.02 ± 2.68a 16.06 ± 2.62a 15.12 ± 1.25a

22.27 ± 0.88ab 23.86 ± 3.95b 27.81 ± 1.45a 147.07 ± 22.76b 154.57 ± 17.25b 211.0 ± 27.04a 13.33 ± 2.56a 20.27 ± 1.95b 16.90 ± 0.99b

25.74 ± 3.40b 29.37 ± 1.93a 09.06 ± 2.12b 69.54 ± 18.74c 71.12 ± 9.03c 177.0 ± 24.97b 16.06 ± 1.96b 23.76 ± 3.02c 21.74 ± 0.86c

⁎Data are expressed as % of inhibition of absorbance of DPPH of sample tubes from absorbance of DPPH from control tubes having no sample. Each value represents the mean ± SD of 9 individuals. Superscript (a, b and c) indicate statistical significance within the groups at P ≤ 0.05 level. Groups having different superscripts are statistically different from each other.

decrease (60%) in non-protein -SH level in gills but not in other tissues was recorded at high salinity when oxygen consumption is less and LPx level of tissue is high. It indicates that non-protein -SH might have been utilized by the tissue to remove hydrogen peroxides and other lipoperoxides during exposure of crabs to high salinity by GST enzymes. de Oliveira et al. (2005, 2006) have observed that GST activity of gills but not of hepatopancreas of C. granulata elevated in response to anoxia. It has been observed that the concentration of GSH was maximum during nocturnal periods in gills of C. granulata and is paralleled by high oxygen consumption and high LPx level (Maciel et al., 2004). Same authors have also noticed unaltered non-protein -SH group in hepatopancreas with changing oxygen consumption. The differences in response of non-protein -SH of tissues of crabs to high salinity may be due to their specific metabolic characteristics according to their functions. Ascorbic acid is a potent antioxidant which scavenges reactive radicals such as hydroxyl, perhydroxyl, peroxyl and nitric oxide (Halliwell and Gutteridge, 2001), therefore, it serves as an important protective measure against free radicals (Bendich et al., 1986; Karakoe et al., 1997). AA is believed to regenerate vitamin E from its oxidized form (Wells et al., 1992) thereby raises the antioxidant status of cells. The significant decrease in AA content in muscle and hepatopancreas of mud crabs at high as well as low salinity signifies its important role in combating stress compared with the non-protein -SH group (Table 4). A small decrease in ascorbic acid content in gills in response to high salinity suggests its obscure role in combating oxidative stress in the tissue. To the best of our knowledge there is no information available in the literature on the effect of salinity on tissue AA content of crabs. DPPH is an established reagent in evaluating antioxidant properties of several biological compounds such as amines, phenols, vitamins, plant extracts etc. (Ionita, 2005). Results of our present work revealed that DPPH radical scavenging capacity of crab tissues was maximum at high salinity in comparison with low salinity at the same temperature. Therefore, it is possible that crabs that inhabit intertidal regions may develop some physiological mechanism to counteract salinity induced stress. Interestingly, a positive correlation was observed between salinity and DPPH radical scavenging capacity for all three tissues of crabs studied (figure not given). To the best of our knowledge no report is available in the literature which shows effect of salinity on DPPH radical scavenging activity in mud crabs. The exact mechanism by which total antioxidant capacity of tissues of crabs elevated is not clear and needs further investigation. Cells, made hypertonic with elevated levels of salt like NaCl, experience double stranded DNA breakage (Kultz and Chakravarty, 2001). It is reported that exposure of marine animals to various stressful conditions causes double stranded DNA damage (Rao et al., 2001; Verlecar et al., 2008) which is contributed by increased production of ROS (Imlay and Linn, 1988; Almeida et al., 2007). Alkaline DNA unwinding test indicated that changing salinity did not

cause any damage to DNA of the tissues of crabs. It may be inferred that the magnitude of generation of ROS in S. serrata at high salinity was not able to surpass the threshold value to cause DNA damage at molecular level. In conclusion, results of the present study suggest that the magnitude of oxidative stress (measured in terms of lipid peroxidation and protein carbonylation) and status of antioxidant defense complexes (both enzymatic and non-enzymatic) of S. serrata are tissue specific and their responses to changing salinity vary from one tissue to the other. However, high salinity (35 ppt) induces more oxidative stress in the tissues of S. serrata in comparison with low salinities (10 ppt and 17 ppt).

Acknowledgements The work was supported by a financial grant of the University Grants Commission, scheme no. F. No.31-284/2005 (SR) to GBNC. BRP is extremely grateful to the University Grants Commission, New Delhi for providing a fellowship under the RFSMS scheme to the Department of Zoology, Utkal University. The authors are highly thankful to the Head of the Departments of Zoology and Biotechnology, Utkal University, India, for providing the laboratory facilities.

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