e n v i r o n m e n t a l t o x i c o l o g y a n d p h a r m a c o l o g y 3 6 ( 2 0 1 3 ) 956–963

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Oxidative stress and apoptosis was induced bybio-insecticide spinosad in the liver ofOreochromis niloticus

Petek Pinera,∗, Nevin Ünerb

a Kahramanmaras Sütcü Imam University, Faculty of Education, Division of Science Education, Avsar Campus,Kahramanmaras, Turkeyb University of C ukurova, Faculty of Science and Letters, Department of Biology, 01330 Balcalı, Adana, Turkey

a r t i c l e i n f o

Article history:

Received 28 January 2013

Received in revised form

13 August 2013

Accepted 15 August 2013

Available online 28 August 2013

Keywords:

Spinosad

a b s t r a c t

This study was conducted to investigate acute toxic effects of spinosad on Glutathione-

related oxidative stress markers, lipid peroxidation, heat shock proteins, apoptosis in the

liver of Oreochromis niloticus selected as a model organism. The fish were exposed to sub-

lethal spinosad concentrations (25, 50, 75 mg/L) for 24–48–72 h. tGSH, GSH, GSSG, and TBARS

contents, GSH/GSSG ratio, and GPx, GR, GST and caspase enzyme activities were measured

using spectrophotometrical methods, and Hsp70 content was measured by ELISA technique.

The results demonstrated that spinosad exposure caused significant alterations in the GSH-

related oxidative stress markers, and also caused increases in lipid peroxidation and stress

proteins with inducing ROS generation in the liver. Apoptosis initiated with the induction of

Oxidative stress

Hsp70

Apoptosis

Liver

Oreochromis niloticus

caspase-3 and Hsp70 could not protect the liver cells. Our results indicated that GSH-related

antioxidant system tried to protect the liver cells from spinosad-induced hepatotoxicity

however, the oxidative stress resulting from induction of ROS generation induced apoptosis

in the liver of O. niloticus.

© 2013 Elsevier B.V. All rights reserved.

receptor activation, mitochondrial apoptotic signaling, drug

1. Introduction

Oxidative stress is defined as an imbalance between produc-tion of free radicals and reactive metabolites, and their elimi-nation by protective mechanisms, referred to as antioxidants(El Golli-Bennour and Bacha, 2011). Glutathione (GSH), themajor water-soluble antioxidant, is known to protect cellularmembranes and lipoproteins from peroxidation (Rana et al.,2002). GSH is involved in many cellular functions including

antioxidant defense via direct interaction with reactive oxy-gen species (ROS) or via activities of detoxification enzymeslike GSH peroxidases (GPx; EC 1.11.1.9) and GSH-S-transferases

∗ Corresponding author at: Kahramanmaras Sütcü Imam University, FCampus, Kahramanmaras, Turkey. Tel.: +90 344 2801333; fax: +90 344 2

E-mail addresses: ppiner@ksu.edu.tr, petekpiner@gmail.com (P. Pin1382-6689/$ – see front matter © 2013 Elsevier B.V. All rights reserved.http://dx.doi.org/10.1016/j.etap.2013.08.009

(GST; EC 2.5.1.18) (Dickinson and Forman, 2002). Cellular GSHis predominantly present in the reduced thiol form which isthe biologically active form. Under oxidizing conditions, oxi-dation of GSH to its disulphide, oxidized glutathione (GSSG),results in a decreased GSH/GSSG ratio (Meister and Anderson,1983). At normal physiological circumstances, GSSG is reducedto GSH by glutathione reductase (GR; EC 1.6.2.4) (Lu, 1999).

A decrease in cellular GSH concentration has been reportedto be an early event in the apoptotic cascade induced by death

aculty of Education, Division of Science Education, 46100 Avsar801302.er).

exposure and oxidative stress (Circu and Aw, 2008). Cellularcaspases belong to a highly conserved family of cysteine pro-teases that cleave aspartate residues of caspase substrates

p h a r

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(seciHsa2

fsc8o4rstsot9ssonalr1w2

aslpa(beMtldscosi

e n v i r o n m e n t a l t o x i c o l o g y a n d

nd are the main players in the execution phase of apoptosisNicholson, 1999). Caspases are constitutively expressed in theytosol as inactive zymogen monomers and are activated bypoptotic signals such as ROS via proteolysis at internal sitesSalvesen and Abrams, 2004). Caspase-3 is the most commonly

easured marker for determination of pesticide-induced apo-tosis (Pena-Llopis et al., 2003; Ramachandiran et al., 2007;hoi et al., 2010; Chatterjee et al., 2011).

Various studies demonstrated that heat shock proteinHsp)-related cytoprotection can be attributed partly to theuppression of apoptosis (Samali and Orrenius, 1998; Creaght al., 2000; Pandey et al., 2000). Hsps function as molecularhaperones in regulating cellular homeostasis and promot-ng cell survival (Sreedhar and Csermely, 2004). Hsps includingsp70 family were highly conserved proteins whose expres-

ions were induced by different kinds of stress factors, suchs heat, oxidative stress, or anticancer drugs (Garrido et al.,001).

Spinosad is an insecticide product derived via fermentationrom a naturally occurring soil actinomycete, Saccharopolysporapinosa (Mertz and Yao, 1990). Spinosad contains two insecti-idal factors, spinosyns A and D, present in an approximately5:15% ratio in the final product (Sparks et al., 1999). Thectanol/water partition coefficient is pH-dependent, 4.01 and.53 at pH 7, expressed as log PKow for spinosyn-A and -D,espectively (WHO, 2005). Spinosad is soluble in water, andoluble in common organic solvents such as acetone, acetoni-rile, methanol, and toluene (Kollman, 2012). Degradation ofpinosad in the environment occurs through a combinationf routes, primarily photodegradation and microbial degrada-ion. The half-life of spinosad degraded by soil photolysis is–10 days. The half-life is less than 1 day for aqueous photoly-is and 1.6–16 days for leaf surface photolysis. The half-life ofpinosad degraded by aerobic soil metabolism in the absencef light is 9–17 days (Thompson et al., 2000). Hydrolysis doesot contribute significantly to degradation, as spinosad is rel-tively stable in water at pH 5–7 and has a half-life of ateast 200 days at pH 9 (Schoonover and Larson, 1995). It waseported that spinosad concentrations were found as 0.54 and.20 �g/mL in water samples obtained from different rivers inhere extensive agricultural practices in Spain (Vega et al.,

005).It affects nicotinic acetylcholine (nAChR) and gamma

mino butyric acid (GABA) receptor sites in the insect nervousystem (Salgado et al., 1997; Watson, 2001). Spinosad exhibitsow mammalian toxicity and a highly favorable environmentalrofile (Cleveland et al., 2001), thus approved for use in organicgriculture by numerous international certification programsCleveland, 2007; Racke, 2007). However an increasing num-er of studies have showed that spinosad caused in vivo toxicffects in mammals (Stebbins et al., 2002; Yano et al., 2002;ansour et al., 2007, 2008a,b) An in vitro study also showed

hat spinosad caused oxidative stress in two mammalian cel-ular models (Perez-Pertejo et al., 2008). A recent study alsoemonstrated that spinosad affects antioxidant system andtimulates oxidative stress, DNA fragmentation, structural

hromosomal aberrations and hence apoptosis in the liverf rats (Aboul-Enein et al., 2012). Previously we reported thatpinosad altered GSH-related antioxidant system and Hsp70n the fish brain (Piner and Üner, 2012a) and also inhibited

m a c o l o g y 3 6 ( 2 0 1 3 ) 956–963 957

acetylcholinesterase (AChE) enzyme activity in the fish tis-sues (Piner and Üner, 2012b). During our literature review, wecould not find any published study related to acute effectsof spinosad on oxidative stress and apoptosis in the liver offish. As a continuation of our previous studies, the effects ofspinosad on GSH-related oxidative stress markers, lipid per-oxidation, Hsps and apoptosis were studied in the liver of O.niloticus.

2. Materials and methods

2.1. Toxicity tests

Juvenile O. niloticus specimens were acclimatized to laboratoryconditions for a month before the experiments. Experimen-tal tanks contained 130 L of dechlorinated and gently aeratedtap water. A commercial spinosad preparation called Laser(480 g/L active ingredient) was used for toxicity tests. Inactiveingredients in commercial formulation were ignored as usual.In preliminary tests, fish were exposed to seven different(5–25–50–75–100–125–150 mg/L) concentrations of spinosad for96 h. Mortality was observed in the concentrations higherthan 100 mg/L. Fish were exposed to three; 25 mg/L, 50 mg/L,75 mg/L different concentrations of spinosad in a static-renewal system for 24–48–72 h, in main toxicity test. Bothexperimental and control fish were kept in similar conditions.No mortality was observed during the toxicity tests (Piner andÜner, 2012a). Liver tissues were dissected out quickly on anice cold plate, divided into two parts, washed in physiologicalsaline solution (0.59% NaCl), weighed and frozen at −85 ◦C. Allchemicals were obtained from Sigma, Chemical Co. St. (USA)and Merck & Co. Inc. (USA).

2.2. Homogenization of liver samples

One part of liver samples were homogenized with 20(v/w) volumes of ice-cold phosphate buffered saline (PBS,136 mmol/L NaCl, 1.5 mmol/L KH2PO4, 0.9 mmol/L CaCl2,8.1 mmol/L Na2HPO4, 2.7 mmol/L KCl, and 0.24 mmol/L MgCl2with 2.5 mmol/L adenosine triphosphate, pH 7.4) containing2.5 mM ATP using a stainless-steel homogenizer (UltraTurraxT-25). After homogenization of samples, they were centrifugedat 16,000 × g for 20 min at 4 ◦C. Supernatants were usedfor determining protein, Hsp70, tGSH, GSSG, thiobarbituricacid reactive substances (TBARS) contents and GPx, GR, GSTenzyme activities. Protein and Hsp70 contents were measuredusing a microplate reader (Varian Cary 50 MPR). tGSH, GSSGcontents and GSH-related enzyme activities were measuredby using a spectrophotometer (Shimadzu UV-Mini 12).

2.3. Biochemical analysis

2.3.1. Measurements of protein contentsLiver samples were analyzed using Bradford reagent (Sigma) to

determine protein contents by the method of Bradford (1976).Absorbance was measured at 595 nm using microplate readerand converted to �g/mL by using bovine serum albumin as astandard.

d p h a r m a c o l o g y 3 6 ( 2 0 1 3 ) 956–963

Table 1 – Effects of spinosad on tGSH and GSH levels(�M/mg protein).

tGSH

Control 25 mg/L 50 mg/L 75 mg/L

24 h 0.59 ± 0.02bx 0.49 ± 0.02bx 0.60 ± 0.04bxy 0.91 ± 0.03ax

48 h 0.63 ± 0.04abx 0.57 ± 0.04bx 0.70 ± 0.06ay 0.72 ± 0.06ay

72 h 0.56 ± 0.01abx 0.62 ± 0.01bx 0.56 ± 0.03abx 0.47 ± 0.03az

GSH

Control 25 mg/L 50 mg/L 75 mg/L

24 h 0.51 ± 0.02ax 0.41 ± 0.02ax 0.51 ± 0.04ax 0.83 ± 0.04bx

48 h 0.56 ± 0.04abx 0.47 ± 0.04ax 0.60 ± 0.06bx 0.62 ± 0.05by

72 h 0.49 ± 0.01abx 0.54 ± 0.02bx 0.48 ± 0.03abx 0.40 ± 0.02az

Values are expressed as mean ± standard error.Letters a and b show the differences between groups at the sameduration, and letters x and y show the differences between groups

958 e n v i r o n m e n t a l t o x i c o l o g y a n

2.3.2. Measurements of Hsp70 contentsThe Hsp70 contents were measured with an indirect noncom-petitive enzyme-linked immunosorbent assay (ELISA). 96-wellimmunoassay/radio immunoassays highbinding microtiterplates (COSTAR, Cambridge, MA, USA) were used. Mono-clonal anti-heat shock protein 70 Clone BRM 22 (Sigma;1/1000 dilution) and anti-mouse IgG (whole molecule)-peroxidase conjugate (Sigma; 1/2000 dilution) were used asprimary and secondary antibody, respectively. After incuba-tion period with freshly prepared Sigma, Fast OPD substrate(o-phenylenediamine dihydrochloride), reaction was stoppedwith 50 �L 3 M H2SO4. Absorbance was measured at 492 nmand converted to ng Hsp70/�g total protein using standard(Sigma; Hsp70 from bovine brain) (De-Boeck et al., 2003).

2.3.3. Measurements of GSH, GSSG contents andGSH-related enzymesSupernants were mixed with 10% 5-sulfosalicylic acid (ice-cold, 1/0.5, v/v) for the determination of tGHS and GSSGcontents. Samples were centrifuged at 9500 × g for 5 min at4 ◦C. The tGSH and GSSG contents in the supernatant weredetermined using 5,5′-dithiobis-(2-nitrobenzoic acid) (DTNB)in the presence of GR (Anderson, 1985). The GSSG was mea-sured after trapping the reduced fraction with 2-vinylpyridine(Griffith, 1980). The samples were analyzed by using the samemethod for tGSH (Pena-Llopis et al., 2001).

GPx activity was analyzed at 37 ◦C and 340 nm throughthe GSH/NADPH/GR system (Beutler, 1984). GR activity wasassayed by monitoring the oxidation of NADPH by GSSG at37 ◦C and 340 nm (Carlberg and Mannervik, 1975). GST activitywas determined by monitoring the changes in absorbance at340 nm, which reflects the rate of 1-chloro-2,4-dinitrobenzene(CDNB) conjugation with GSH at 30 ◦C for 2 min (Habig et al.,1974).

2.3.4. Measurements of TBARS contentsSamples were vortexed in a medium containing 8.1% ofSDS, 20% of acetic acid and 0.8% of TBA (pH 3.4) andincubated at 95 ◦C. After 30 min, they were cooled, mixedwith n-butanol/pyridine (14:1) and centrifuged at 1073 × g.TBARS contents were determined by measuring the pinkcolor at 532 nm. Absorbance were converted to TBARScontents by using the standard graph prepared with 1,1′,3,3′

tetramethoxypropane (malonaldehyde bis(dimethyl acetal))(Ohkawa et al., 1979).

2.3.5. Measurements of caspase-3 enzyme activityLiver samples were homogenized with 10 volumes of ice-coldlysis buffer which is in Sigma, caspase-3 colorimetric assaykit using a homogenizer (UltraTurrax T-25). Homogenateswere centrifuged at 13,000 × g for 15 min at 4 ◦C. Caspase-3enzyme activity was measured in 96-well plates using theSigma, caspase-3 colorimetric assay kit. The hydrolysis of thepeptide substrate acetyl-Asp-Glu-Val-Asp p-nitroanilide (Ac-

DEVD-pNA) to release pNA was measured at 405 nm by usingmicroplate reader (Varian Cary 50 MPR). The pNA calibrationcurve was used for determination of caspase-3 enzyme activ-ity (Sigma, Caspase 3 Assay Kit, Technical Bulletin).

at the same concentration. Data shown different letters are signifi-cantly different at the p < 0.05 level (N = 6).

2.3.6. Statistical analysesAll data were expressed as mean ± standard error. Statisticaldifferences between the treatment and control groups weredetermined by the analysis of variance (ANOVA) using theDuncan in SPSS 16.0 (SPSS Inc., Chicago, IL, USA).

3. Results

3.1. tGSH, GSH, GSSG levels and GSH/GSSG ratio

tGSH and GSH contents of the liver are shown in Table 1. Thedata indicated that exposure to spinosad caused significantincreases in tGSH (54%) and GSH (62%) for 24 h of 75 mg/Lspinosad exposure when compared to respective controls inthe liver. Additionally, tGSH and GSH contents decreased in atime-dependent manner for 75 mg/L spinosad exposure. Therewere no significant changes in tGSH and GSH contents of theliver for 48–72 h of all spinosad exposures compared to respec-tive controls.

Table 2 shows results obtained with GSSG levels andGSH/GSSG ratio. Exposure to spinosad induced a significantincrease in GSSG levels of liver for 75 mg/L spinosad at 24 h(30%) and for all applied spinosad concentrations at 48 h (30%,30% and 43% for 25, 50, and 75 mg/L spinosad, respectively). Onthe contrary, the exposure to 25 mg/L spinosad caused a sig-nificant decrease in GSH/GSSG ratio (21%) at 24 h, but 75 mg/Lspinosad caused a significant increase in GSH/GSSG ratio(18%) at the same period. Spinosad treatments of 25–75 mg/Ldecreased GSH/GSSG ratio (31% and 20%) following 48 h ofexposure. All tested spinosad concentrations had no signifi-cant effect on GSSG levels and GSH/GSSG at 72 h of exposure.

3.2. GPx, GR, GST enzyme activities

Spinosad caused significant increases in the GPx activities at

24 and 48 h of exposures. Exposure to 50 and 75 mg/L spinosadcaused a significant elevation in GPx activities (90% and 66%)at 24 h. Similar to this findings, all tested concentrations ofthe spinosad significantly increased GPx activities (45%, 82%

e n v i r o n m e n t a l t o x i c o l o g y a n d p h a r m a c o l o g y 3 6 ( 2 0 1 3 ) 956–963 959

Table 2 – Effects of spinosad on GSSG levels (�M/mg protein) and GSH/GSSG ratio.

GSSG

Control 25 mg/L 50 mg/L 75 mg/L

24 h 0.073 ± 0.005ax 0.080 ± 0.006abxy 0.084 ± 0.004abxy 0.095 ± 0.005bx

48 h 0.071 ± 0.004ax 0.093 ± 0.005bx 0.093 ± 0.005bx 0.102 ± 0.005bx

72 h 0.075 ± 0.004ax 0.078 ± 0.005ay 0.074 ± 0.005ay 0.075 ± 0.006ay

GSH/GSSG

Control 25 mg/L 50 mg/L 75 mg/L

24 h 8.16 ± 0.359ax 6.37 ± 0.695cy 7.07 ± 0.352acx 9.63 ± 0.425bx

48 h 8.80 ± 0.579ax 6.07 ± 0.428by 7.41 ± 0.501abx 6.99 ± 0.471by

72 h 7.49 ± 0.241ax 8.10 ± 0.626ax 7.50 ± 0.438ax 6.54 ± 0.567ay

Values are expressed as mean ± standard error.duratfferen

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Letters a, b and c show the differences between groups at the same

same concentration. Data shown different letters are significantly di

nd 77% for 25, 50, and 75 mg/L spinosad, respectively) at 48 hf exposure. There were no significant alterations in GR andST enzyme activities at 24 h. However spinosad treatmentf 50 mg/L increased GR activity (41%) following 48 h of expo-ure and spinosad treatments of 25 and 50 mg/L increased GRctivities (60% and 100%) following 72 h of exposures. Spinosadreatments of 50 and 75 mg/L significantly elevated GST spe-ific activity (19%) after 48 h of exposures. In contrast, spinosadreatment of 50 mg/L decreased GST activity (23%) following2 h of exposure (Table 3).

.3. TBARS levels

here were no significant alterations in TBARS levels at 24 and

2 h of spinosad exposures. However, spinosad treatment of5 mg/L increased TBARS levels (128%) following 48 h of expo-ure (Table 4).

Table 3 – Effects of spinosad on GSH-related enzymes activities

GPx

Control 25 mg/L

24 h 0.33 ± 0.008cx 0.44 ± 0.031bcx

48 h 0.35 ± 0.010cx 0.51 ± 0.035bx

72 h 0.32 ± 0.014ax 0.42 ± 0.046ax

GR

Control 25 mg/L

24 h 0.012 ± 0.0004abx 0.011 ± 0.0004a

48 h 0.012 ± 0.0003bx 0.012 ± 0.0001b

72 h 0.012 ± 0.0002cx 0.020 ± 0.0001b

GST

Control 25 mg/L

24 h 33.23 ± 1.28ax 26.84 ± 0.83ay

48 h 34.85 ± 2.49bx 40.64 ± 2.56abx

72 h 34.24 ± 1.26ax 39.34 ± 1.71ax

Values are expressed as mean ± standard error.Letters a, b and c show the differences between groups at the same duratsame concentration. Data shown different letters are significantly differen

ion, and letters x and y show the differences between groups at thet at the p < 0.05 level (N = 6).

3.4. Hsp70 levels

All tested spinosad concentrations had no significant effectson Hsp70 levels at 24 and 48 h of exposures. However, spinosadexposure of 50 and 75 mg/L significantly augmented Hsp70levels (84% and 172%) after 72 h of exposures (Table 4).

3.5. Caspase-3 enzyme activity

Table 4 shows that caspase-3 enzyme activity did not exhibitsignificant changes in fish exposed to spinosad at 24 h. How-ever caspase-3 enzyme activities were significantly increasedby all tested spinosad concentrations (28%, 29% and 51% for25, 50, and 75 mg/L spinosad, respectively) following 48 h of

exposure. Likewise, spinosad exposure of 25 and 75 mg/L sig-nificantly augmented caspase-3 enzyme activities (25% and30%) after 72 h of exposures.

(U/mg protein).

50 mg/L 75 mg/L

0.63 ± 0.082ax 0.55 ± 0.033abx

0.64 ± 0.061ax 0.62 ± 0.050abx

0.37 ± 0.020ay 0.34 ± 0.015ay

50 mg/L 75 mg/Lby 0.011 ± 0.0003bz 0.013 ± 0.0004ay

y 0.017 ± 0.0001ax 0.013 ± 0.0003by

x 0.014 ± 0.0001cy 0.024 ± 0.0002ax

50 mg/L 75 mg/L

27.46 ± 1.58ay 29.58 ± 2.49ay

41.55 ± 2.81ax 41.72 ± 5.42ax

26.12 ± 1.32by 39.11 ± 2.28ax

ion, and letters x and y show the differences between groups at thet at the p < 0.05 level (N = 6).

960 e n v i r o n m e n t a l t o x i c o l o g y a n d p h a r m a c o l o g y 3 6 ( 2 0 1 3 ) 956–963

Table 4 – Effects of spinosad on Hsp70 (ng/�g protein) and TBARS (nmol/mg protein) levels and caspase-3 enzymeactivity (�U/mg protein).

Hsp70

Control 25 mg/L 50 mg/L 75 mg/L

24 h 18.41 ± 0.42ax 18.96 ± 0.59ax 19.82 ± 1.11ay 18.25 ± 0.28ay

48 h 17.80 ± 1.81ax 17.03 ± 0.73axy 16.02 ± 0.37az 18.46 ± 1.04ay

72 h 16.50 ± 1.64cx 15.54 ± 0.74cy 30.51 ± 1.47bx 45.02 ± 0.88ax

TBARS

Control 25 mg/L 50 mg/L 75 mg/L

24 h 0.37 ± 0.049ax 0.33 ± 0.023ax 0.47 ± 0.075ax 0.34 ± 0.022az

48 h 0.35 ± 0.015bx 0.39 ± 0.106bx 0.48 ± 0.083bx 0.80 ± 0.071ax

72 h 0.50 ± 0.050abx 0.37 ± 0.041abx 0.30 ± 0.068bx 0.55 ± 0.065ay

Caspase-3

Control 25 mg/L 50 mg/L 75 mg/L

24 h 7.32 ± 0.38ax 6.56 ± 0.65ay 7.71 ± 0.52ay 7.75 ± 0.24ay

48 h 7.33 ± 0.17cx 9.40 ± 0.71bx 9.48 ± 0.78bx 11.09 ± 0.29ax

72 h 6.25 ± 0.16bx 7.77 ± 0.22ay 7.32 ± 0.28aby 8.14 ± 0.21ay

Values are expressed as mean ± standard error.uratifferen

Letters a, b and c show the differences between groups at the same dsame concentration. Data shown different letters are significantly di

4. Discussion

Pesticide producers are continuously replacing the older gen-eration pesticides with an array of newly developed pesticides.Spinosad the natural insect control products was classifiedby Environmental Protection Agency (EPA) as a “reduced-risk” compound because of its favorable environmental andtoxicological profile (EPA, 1997). The pesticides have beenshown to induce production of ROS by altering the balancebetween the oxidants/prooxidants and antioxidants throughpromoting lipid peroxidation and depleting the antioxidativecellular reserves (both the enzymatic and non enzymatic)leading to a condition of oxidative stress (Agrawal and Sharma,2010). Aboul-Enein et al. (2012) reported that spinosad alteredantioxidant system and induced apoptosis with initiatingoxidative stress in rat liver. Our findings also showed thatspinosad causes oxidative stress and induces apoptosis in theliver of O. niloticus.

Glutathione is an important naturally occurring antioxi-dant which prevents free radical damage. GSH is consumedby GSH-related enzymes and GSH converted to GSSG underoxidative stress conditions (Hayes and McLellan, 1999). Weobserved an initial adaptive response to the oxidative stressby tGSH, GSH, GSSG levels and GSH/GSSH ratio. Aboul-Eneinet al. (2012) reported that spinosad exposure stimulated oxida-tive stress by reducing GSH level in the liver of rats. Changesin GSSG and GSH/GSSG ratio have been used as markers ofoxidative stress status in biological systems (Ji, 1999). In thisstudy, a significant increase in GSSG and a significant decreasein GSH/GSSG ratio were observed in all spinosad treatmentsat 48 h which was indicated an increased-free-radical produc-

tion. Similar to our results, spinosad caused oxidative stress byaltering GSH-redox cycle and lipid peroxidation in two mam-malian cellular models (Perez-Pertejo et al., 2008).

on, and letters x,y and z show the differences between groups at thet at the p < 0.05 level (N = 6).

GSH is central to the cellular antioxidant defenses and actsas an essential cofactor for antioxidant enzymes includingGPx, GR and GST (Mascio et al., 1991; Hayes et al., 2005). In thecurrent study we showed that spinosad induced an antioxi-dant response with a significant stimulation of GSH-relatedenzymes. The biological function of GPx is to remove hydro-gen peroxide (H2O2) and to reduce lipid hydroperoxides totheir corresponding alcohols (Arthur, 2000). In rats, spinosadexposure caused significant induction in GPx enzyme activityin rat liver (Aboul-Enein et al., 2012). Similarly, higher levelsof spinosad concentrations used in this study increased GPxactivity that could be related to the elimination of H2O2 andthen an enhanced GSSG production. The results also indicatedthat the observed induction in GR activity depended on theincreases in GSSG levels and decreases in the GSH/GSSG ratios.Likewise, Perez-Pertejo et al. (2008) reported that spinosadcaused rises in GPx and GR activities in two mammalian cel-lular models. Based on our findings, we showed that spinosadsignificantly stimulated GSH-related oxidative stress markers.

Oxidative damage primarily occurs through production ofROS and subsequently react with biological molecules as wellas causing damage to membranes and other tissues (Banerjeeet al., 1999). Pesticides can cause lipid peroxidation and alter-nations in membrane fluidity (Zama et al., 2007; Aly et al.,2010; Elsharkawy et al., 2012). Lipid peroxidation has beensuggested as one of the molecular mechanisms involved inpesticide-induced toxicity (Abdollahi et al., 2004). The resultsof the present study showed that the lipid peroxidation ini-tially was not increased by spinosad due to the stimulationof GSH related antioxidant system. However, this system didnot protect membrane lipids from peroxidation and spinosad

created oxidative stress by rising the lipid peroxidation in theliver at 48 h. Organic peroxides can be detoxified by the activityof GST (Halliwell and Gutteridge, 1999). It was also suggestedthat activation of GST could be related to scavenging of lipid

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eroxides which were produced by spinosad exposure. Previ-usly we showed that spinosad caused no lipid peroxidationnd inhibited GST enzyme activity in the brain of O. niloticusPiner and Üner, 2012a). GST is a detoxifying enzyme that cat-lyzes the conjugation of a variety of electrophilic substrateso GSH (Hayes et al., 2005). Conjugation of spinosad with GSH,ither directly or after O-or N-demethylation was determinedn rats (FAO/WHO, 2001). In the present study, increases in GSTctivity may also be related to the conjugation of GSH withpinosad.

ROS can have diverse effects on cell growth and are capa-le of directing cells to undergo apoptosis (Ray et al., 1999;inkel and Holbrook, 2000). Oxidative stress might be a com-on mediator of apoptosis (Esteve et al., 1999; Circu and

w, 2010). Activation of caspase-3 is a critical event in thexecution phase of apoptosis involving pesticide exposureKitazawa et al., 2003; Ramachandiran et al., 2007). Aboul-nein et al. (2012) reported that spinosad caused DNA damagen liver, and the formation of a DNA ladder, hallmark of cellsndergoing apoptosis. Elevated GSH levels afforded protectiongainst stress-induced apoptosis (Watson et al., 1997; Friesent al., 2004). Studies have provided evidence for a role forhe GSH/GSSG redox status in cell apoptosis in various cellypes (Pias and Aw, 2002; Pias et al., 2003). In this study, itas suggested that augmented GSH initially prevented the

aspase activation in liver cells. However spinosad increasedSSG levels and decreased GSH/GSSG ratio due to increasedOS production; and then caused caspase-3 activation and

nitiated apoptosis. As a result of this finding, GSH-relatedntioxidant system did not produce an effective protectionnder the severe oxidative conditions.

Under various stress conditions the synthesis of stress-nducible Hsp70 enhances the ability of stressed cells to cope

ith increased concentrations of unfolded or denatured pro-eins (Garrido et al., 2001). It was reported that free-radicalsaused protein oxidation (Berlett and Stadtman, 1997) andxidative modification of proteins can lead to diminishedpecific protein functions which may ultimately result inell death (Stadtman, 1992; Dean et al., 1997). In this study,nchanged Hsp70 levels may be associated with the protec-ion of cell proteins from oxidation by GSH-related antioxidantystem. Increased oxidative conditions may be the cause ofncreased protein oxidation and then Hsp70 expression in theiver. However Hsp70 was not sufficient for the protection ofiver cells from caspase activation and following apoptosis.

. Conclusion

his study demonstrated that spinosad caused oxidativetress in the liver of O. niloticus with increasing lipid per-xidation, Hsp70 and also the activities of GSH-dependentntioxidant system by inducing ROS generation. It was alsoound that spinosad triggered apoptosis with activation ofaspase-3 enzyme however, Hsp70 induction did not protectiver cells from the apoptosis. Our results indicated that the

ighest level of spinosad used in this study had significantffects on tGSH, GSH, GSSG levels, GSH/GSSG ratio and GPxctivity in short term exposure; on Hsp70 level and caspase-3ctivity in long term exposure. As a conclusion, GSH-related

m a c o l o g y 3 6 ( 2 0 1 3 ) 956–963 961

antioxidant system initially tried to protect liver cells fromoxidative stress and apoptosis however, this system did notachieve an effective protection under the influence of intenseoxidative conditions.

Conflict of interest statement

There are no conflicts of interest.

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