Interplay of early biochemical manifestations by cadmium insult in sertoli–germ coculture: An in vitro study

Toxicology 287 (2011) 46–53

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Interplay of early biochemical manifestations by cadmium insult in sertoli–germ coculture: An in vitro study Smita Khanna a , Pramesh C. Lakhera b , Shashi Khandelwal a,∗ a b

Immunotoxicology Lab, CSIR-Indian Institute of Toxicology Research (CSIR-IITR), Lucknow, India Department of Biotechnology, H.N.B. Garhwal University, Srinagar, Garhwal, India

a r t i c l e

i n f o

Article history: Received 8 April 2011 Received in revised form 20 May 2011 Accepted 23 May 2011 Available online 30 May 2011 Keywords: Cadmium Sertoli–germ coculture Oxidative stress Apoptosis LDH-X Tesmin

a b s t r a c t Cadmium is a common environmental and occupational hazard and its adverse effect on reproductive organ has been well documented. The present study is planned to delineate the mechanism of Cd toxicity in rat testes. Our study shows that Cd causes apoptosis in sertoli–germ cells which is governed by oxidative stress. We assayed ROS, GSH and MMP to ensure the role of oxidative stress, further confirmed it by thiol modulators. The initial biochemical response shown in sertoli–germ cells was a significant rise in intracellular calcium followed by a drastic fall in MMP and then ROS generation. The downstream events included cytochrome c release leading to caspase-3 activation and culminating in cell death via apoptosis. Furthermore Cd disrupted the spermatogenic pathway as evident by suppression in tesmin and LDH-X levels. © 2011 Elsevier Ireland Ltd. All rights reserved.

1. Introduction Cadmium (Cd), one of the most toxic heavy metals, is classified as a human carcinogen by International Agency for Research on Cancer (IARC, 1993) and is 7th in the Priority List of Hazardous Substances of Agency for Toxic Substances and Disease Registry (ATSDR, 2007). It is a broad spectrum toxicant, affecting multiple organs including the reproductive system. In females, Cd alters progesterone and estradiol synthesis (Piasek and Laskey, 1994, 1999) whereas in males, Cd exposure can lead to testosterone suppression, reduced sperm motility and testicular damage at higher doses (King et al., 1998; Liu et al., 2001; Yang et al., 2003). Effects of Cd on human fetal germ cells suggest that this metal could be deleterious to early gametogenesis, as well (Waalkes et al., 2003). In rodents, testicular toxicity by Cd is well accepted, being age and dose dependent. Higher susceptibility to Cd-induced testicular damage in mature (7- and 12-week-old) rats than in immature (3-week-old) rats was studied by Nemoto et al. (2009). Cadmium (5 ␮mol/kg or less) induces apoptosis in spermatogenic cells, inhibits spermiation and increases prevalence of prostate cancer (Waalkes et al., 1989; Zhou et al., 1999), while >10 ␮mol/kg Cd

∗ Corresponding author at: Immunotoxicolgy Lab, CSIR-Indian Institute of Toxicology Research (CSIR-IITR), 80 M.G. Marg, Lucknow 226001, U.P., India. Tel.: +91 5222627586x316; fax: +91 5222620207. E-mail address: skhandelwal [email protected] (S. Khandelwal). 0300-483X/$ – see front matter © 2011 Elsevier Ireland Ltd. All rights reserved. doi:10.1016/j.tox.2011.05.013

causes severe hemorrhagic necrosis in rats and mice and interstitial cell tumor in rats (Waalkes et al., 1989, 1999; Waalkes, 2000). Cadmium can directly inhibit primary Leydig cell testosterone levels (Laskey and Phelps, 1991) as well as follicle stimulating hormone in plasma of rats (Lafuente et al., 2000). It has also been reported to cause loss of temporal regulation in spermatocytes, suggesting testis to be more sensitive to Cd stress than other organs (Matsuura et al., 2002). Tesmin, an early marker of male germ cell differentiation has been shown to be translocated from cytoplasm to nucleus by oxidative stress inducers like cobalt chloride, diethyl maleate and Cd (Matsuura et al., 2002; Sutou et al., 2003). Clear evidence of oxidative stress, mitochondrial dysfunction and apoptosis caused by Cd in testis and other organs has been demonstrated by several groups. To name a few, involvement of reactive oxygen species (ROS) and of antioxidant enzymes in Cdinduced testicular damage (Koisume and Li, 1992; SenGupta et al., 2004), elevated lipid peroxidation in lung, brain, kidney, liver, erythrocytes and testis (Bagchi et al., 1997; Klimczak et al., 1984; Manca et al., 1991; Sugawara and Sugawara, 1984) and DNA singlestrand breaks in Leydig cells by Cd in vitro (Yang et al., 2003). Although, the testicular damage by Cd was recognized by Parizek way back in 1957 (Parizek and Zahor, 1956), the precise mechanisms underlying its toxicity still remain enigmatic. Several mechanisms of Cd-induced testicular toxicity have been proposed. A recent review (Siu et al., 2009) focused on the disruption of blood–testis barrier, the major target of Cd toxicity in testis (Tables 1–4).

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Cadmium has been shown to induce apoptosis in a number of cell types and tissues, both in vitro and in vivo (Friberg, 1984; Koizume et al., 1996; Pathak and Khandelwal, 2007; Tzirogiannis et al., 2003). Characteristic DNA fragmentation, TUNEL assay and histopathologically observed changes characteristic of apoptosis were found in kidney, prostate, seminal vesicles, testes, and epididymis, but not in bladder and vas deferens tissue of rats 48 h after Cd (0.03 mmol/kg) administration (Xu et al., 1996; Yan et al., 1997). Although cited in literature that low doses of Cd cause testicular dysfunction via oxidative stress and DNA damage following apoptotic pathway, the detailed signaling pathway has not been explicitly defined. The current study was therefore, designed to delineate dose and time dependent apoptogenic signaling pathway by Cd, in primary sertoli–germ coculture. The critical role of ROS and GSH, in mediating mitochondrial caspase pathway, was thoroughly evaluated. The very immediate cellular responses of calcium homeostasis, mitochondrial membrane potential and ROS were highlighted. In addition, the toxic influence of Cd on testis specific protein markers (tesmin and LDH-X) was also ascertained to understand the deleterious effects of this metal on spermatogenesis. 2. Materials and methods 2.1. Chemicals All the chemicals were of highest grade purity available. Cadmium Chloride, RNase A, DMEM-F12, antibiotic–antimycotic solution, Dulbecco’s phosphate buffered saline (PBS), fetal bovine serum (FBS), 2 ,7 -dichlorofluorescein diacetate (DCFH-DA), 4 -6-Diamidino-2-phenylindole (DAPI), collagenase type IV and all other chemicals were purchased from Sigma–Aldrich, USA. Rhodamine123 (Rh 123) and 5 -chloromethylfluorescein diacetate (CMF-DA) from Molecular Probes and propidium iodide (PI), Z-DEVD-fmk was from Calbiochem, DEVD-AFC and LDH Assay kit were purchased from Biovision, Annexin V-FITC reagent from Pharmingen (Becton Dickinson Company), anti-rabbit cytochrome c, anti-rabbit tesmin from SantaCruz and Alamar Blue from Invitrogen. 2.2. Preparation of sertoli–germ coculture Male Wistar rats were maintained in IITR animal house under standard conditions. They were housed in plastic polypropylene cages with a 12 h light/dark cycle and temperature of 25 ± 2 ◦ C. They were fed with standard rodent pellet and water ad libitum. Our animal house and breeding facility are registered with Committee for the Purpose of Control and Supervision of Experiments on Animals (CPCSEA), Government of India and CPCSEA guidelines were followed (IAEC approval obtained). Briefly, testis were dissected from 28 days old rat, de-capsulated, teased, suspended in 10 ml HBSS containing 1 mg/ml collagenase type IV and pipetted well to obtain a single cell suspension. After 30 min, the suspension was filtered through 30 ␮m nylon membrane and centrifuged at 80 × g for 10 min. The pellet collected was washed with DMEM/F12 medium and resuspended in the same. The entire procedure was carried out under aseptic conditions. The suspension obtained comprised of sertoli and germ cells (Adhikari et al., 2000). The cell density was adjusted to 1.0 × 106 cells/ml and the viability of freshly isolated cells was always over 95% (trypan blue exclusion test). 2.3. Cytotoxicity by Alamar Blue Analysis of cytotoxicity was done by Alamar Blue reduction assay using a commercial kit (Invitrogen). Cells seeded at a density of 1.0 × 104 in 96-well plate were treated with various concentrations of Cd (10, 100 and 1000 ␮M) for 6 and 24 h at 37 ◦ C in a CO2 incubator. Alamar Blue was added 6 h prior to the incubation time and absorbance was read at 530 nm on a FluoStar Omega microplate reader. 2.4. Flow cytometry analysis Flow cytometric studies were performed with a BD-LSR flow cytometer (Becton Dickinson, San Jose, CA). For each sample, data from 10,000 cells were recorded. Cell debris, characterized by a low FSC/SSC was excluded from analysis. Data was analyzed by Cell Quest software and the mean fluorescence intensity was obtained by histogram statistics. 2.4.1. Apoptotic and necrotic cell population During apoptosis phosphotidylserine (PS) externalization can be detected with annexin V, which has high affinity for PS. Using DNA specific viability dyes, such as propidium iodide (PI), it is possible to distinguish between early apoptotic, late apoptotic, and dead cells (Vermes et al., 1995).

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Briefly, after treatment with various concentrations of Cd (10, 100 and 1000 ␮M) for 6 and 18 h at 37 ◦ C, the harvested cells were suspended in 200 ␮l binding buffer and was incubated with 2 ␮l annexin V-FITC and 10 ␮l PI (10 ␮g/ml) for 20 min in dark at RT and 400 ␮l PBS was then added to each sample. The FITC and PI fluorescence were measured through FL-1 filter (530 nm) and FL-2 filter (585 nm) respectively. 2.4.2. Mitochondrial membrane potential ( ) determination Rh 123, a cationic fluorophore is actively accumulated by cells in direct proportion to the mitochondrial membrane potential. For the detection of mitochondrial  , Cd treated cells were incubated with Rh 123 (5 ␮g/ml) for 60 min in dark at 37 ◦ C, harvested and suspended in PBS. The mitochondrial  was measured by fluorescence intensity (FL-1, 530 nm) of 10,000 cells on flow cytometer (Juan et al., 1994). 2.4.3. Glutathione (GSH) estimation 5 -Chloromethylfluorescein diacetate (CMF-DA) was used as a selective probe to monitor intracellular GSH (Hedley and Chow, 1994). Following Cd treatment for different time points, cells were incubated with CMF-DA (1 ␮M final conc.) for 30 min in dark at 37 ◦ C. After harvesting, the cells were suspended in 500 ␮l PBS and GSH was measured by the fluorescence intensity (FL-1, 530 nm) of 10,000 cells on flow cytometer. 2.4.4. Caspase-3 enzymatic assay Caspases get activated during apoptosis and the assay is based on detection of cleavage of substrate DEVD-AFC and the free AFC can be quantified using flow cytometer. Briefly, cells were treated with 100 ␮M Cd for 30, 1, 3, 6 and 18 h, harvested, suspended in PBS containing 0.05% Tween-20 and kept in ice for 10 min. Caspase-3 substrate DEVD-AFC (50 ␮M final concentration) was then added and further incubated for 30 min at room temperature. Cells were then collected, washed and resuspended in PBS. AFC fluorescence intensity was measured on flow cytometer (FL-1, 530 nm). Using DEVD-AFC, we were able to determine the enzyme activity on flow cytometer, with high reproducibility. 2.5. DNA condensation by DAPI 4 -6-Diamidino-2-phenylindole (DAPI) is known to form fluorescent complexes with natural double-stranded DNA, showing fluorescence specificity for AT, AU and IC clusters (Kapuscinski, 1995). When DAPI binds to DNA, its fluorescence is strongly enhanced. Briefly, following 100 ␮M Cd treatment for 18 h, cells were harvested and washed once with PBS, and then resuspended in PBS containing 0.1% Triton X and incubated for 10 min on ice. Cells were spin down and resuspended in 4% PBS buffered paraformaldehyde solution containing 10 ␮g/ml DAPI. DNA condensation was observed under fluorescence microscope. 2.6. ROS measurement Intracellular ROS generation by primary sertoli–germ coculture was analyzed by using the florescent probe 2 ,7 -dichlorofluorescein diacetate (DCFH-DA) which is deacetylated to DCFH inside the cell. ROS is able to oxidize DCFH to DCF which is proportional to the cellular oxidant production (Zamzami et al., 1995). Cells were incubated with Cd for various time points (15 min, 30 min, 1 h, 3 h, 6 h and 18 h) and further incubated with DCFH-DA (100 ␮M) for 60 min in dark at 37 ◦ C. The cells were harvested, washed and resuspended in PBS and ROS generation was measured by the fluorescence intensity on a FluoStar Omega Plate reader with Ex, 488 nm and Em, 510 nm. 2.7. Lipid peroxidation Lipid peroxidation is a well-established mechanism of cellular injury in both plants and animals, and is used as an indicator of oxidative stress in cells. Malondialdehyde (MDA) and 4-hydroxyalkenals (HAE) has been used as a measure of lipid peroxidation (Seljeskog et al., 2006). The method of Ohkawa et al. (1979) was used with slight modifications to measure MDA. Briefly, Cd treated cells were collected and suspended in 200 ␮l PBS. Hundred microliters SDS (8.1%), 700 ␮l acetic acid (20%, pH3.5) and 700 ␮l of TBA (0.8%) were added and the volume made up to 200 ␮l with milli Q. After boiling for 15 min, samples were cooled and centrifuged at 1500 × g for 15 min. Supernatant was collected and absorbance was measured at 532 nm using FluoStar Omega Plate reader to check MDA levels. 2.8. LDH-X assay LDH-X was determined by using commercial kit (BIOVISION). LDH-X is the predominant isoform present in testicular germ cells. Briefly, cells were treated with various concentrations of Cd for different time periods. After incubation, supernatant was collected and equal volume of Reaction Mixture was added and further incubated for 30 min at RT in dark. Absorbance was measured at 490–500 nm using FluoStar Omega Plate reader, the reference wavelength should be more than 600 nm.

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2.9. Protein levels by western blot: cytochrome c and tesmin To prepare the fractions, cells were allowed to swell in ice cold mitochondria buffer (250 mM sucrose, 20 mM HEPES, 10 mM KCl, 1.5 mM MgCl2 , 1 mM EDTA, 1 mM EGTA, 1 mM dithiothreitol, 1 mM phenylmethylsulfonyl fluoride and protease inhibitor mixture, pH 7.4) for 20 min on ice. The cells were homogenized by syringe and centrifuged at 12,000 × g for 30 min at 4 ◦ C to separate heavy organelles and cytosol (Lemarieˇı et al., 2004). The supernatant representing the cytosolic fraction was analyzed for cytochrome c and tesmin. Fifty micrograms protein sample was separated on a 12%SDS–PAGE and was subsequently transferred on to a nitrocellulose membrane. Membrane was then blocked with blocking solution (Sigma) in PBST buffer (PBS buffer containing 0.1% Tween-20) for 4 h at RT, probed with anticytochrome c (1:2000) and anti–tesmin antibody (1:2000) overnight at 4 ◦ C, After 3–5 washings of 5 min each with wash buffer (PBS and 0.05% Tween-20), blots were incubated with HRP-conjugated secondary antibody (1:7000) in blocking solution for 2 h at RT. Proteins were visualized by enhanced chemiluminescent detection system according to manufacturer’s protocol (MILLIPORE). 2.10. Statistical analysis Significance of mean of different parameters between the treatment groups were analyzed using one-way analysis of variance (ANOVA) after ascertaining the homogeneity of variance between the treatments. Pair wise comparisons were done by calculating the least significant difference.

Fig. 1. Effect of Cd on cell viability: freshly isolated sertoli–germ cells (1.0 × 104 ) were treated with Cd (10, 100, 1000 ␮M) for 6 and 24 h. Alamar blue was added 6 h prior to completion time and absorbance was measured at 530 nm. Each point represents mean ± S.D. (n = 3). ***P < 0.001, **P < 0.01, *P < 0.05 as compared to control, using one-way ANOVA.

vations, 100 ␮M Cd exhibiting ∼50% cell loss, was the preferred dose to monitor some of the apoptotic events.

3. Results 3.2. Cd induces DNA damage 3.1. Effect of Cd on cell survival Loss in cell viability of sertoli–germ coculture was evident as early as 6 h by 100 and 1000 ␮M Cd as shown in Fig. 1. On increasing the exposure time, 100 ␮M Cd showed a time dependent loss in cell viability i.e. from 20% at 6 h it became 50% at 24 h. 1000 ␮M Cd the highest dose, did not result in increased cytotoxicity, as the effect was almost similar to 100 ␮M Cd dose at 24 h. Based on these obser-

On Cd exposure, the sertoli–germ coculture undergoes apoptosis. Two techniques were employed to measure DNA damage, the Annexin V binding assay and DAPI staining. The basal level of dead cells was less than 4.0% throughout the experiment. All the three Cd (10, 100 and 1000 ␮M) concentrations caused a dose and time dependent induction of apoptosis at 6 and 18 h (Fig. 2A and B). Even the lowest Cd dose initiated mild apoptosis at 6 h increas-

Fig. 2. Effect of Cd on apoptosis: apoptotic and necrotic cell distribution: freshly isolated sertoli–germ cells (0.2 × 106 ) were treated with Cd (10, 100, 1000 ␮M) for 6 and 18 h. Annexin V and PI (10 ␮g/ml) was added for 20 min in dark and FITC and PI fluorescence was measured using flow cytometer with FL-1 and FL-2 filters, respectively. Results were expressed as (A) bar chart showing cell apoptosis and necrosis at 6 and 18 h. Each bar represents mean ± S.D. (n = 3). ***P < 0.001, **P < 0.01, *P < 0.05 as compared to control, using one-way ANOVA and (B) dot plots representing one of the three independent experiments. LL: live cells (Annexin V− /PI− ), LR: early/primary apoptotic cells (Annexin V+ /PI− ), UR: late/secondary apoptotic cells (Annexin V+ /PI+ ), and UL: necrotic cells (Annexin V− /PI+ ). (C) DNA condensation by DAPI: to freshly isolated sertoli–germ cells (1.0 × 106 ) treated with 100 ␮M Cd for 18 h at 37 ◦ C, DAPI (10 ␮g/ml) was added and the cells observed under fluorescence microscope.

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Table 1 Effect of Cd on ROS, GSH and MMP: freshly isolated sertoli–germ coculture (1.0 × 106 ) was incubated with 10 ␮M, 100 ␮M and 1000 ␮M Cd for various time points. ROS (by DCFH-DA): DCFH-DA (100 ␮M) was added and cells were incubated for 60 min in dark at 37 ◦ C. DCF fluorescence was measured on a plate reader with Ex: 488 nm and Em: 510 nm. GSH (by CMFDA): CMF-DA (1 ␮M) was added and cells were incubated for 30 min. CMF fluorescence was measured using flow cytometer with FL-1 filter. MMP (by Rhodamine 123): Rh123 was added, cells were incubated for 60 min and the fluorescence was measured using a flow cytometer with FL-1 filter. Data represents mean ± S.D. from three independent experiments. ***P < .001,**P < 0.01,*P < 0.05 as compared to control, using one-way ANOVA. 10 ␮M Cd

Con

ROS

15 min 30 min 1h 3h 6h 18 h

1686 1624 1669 1667 1995 1964

± ± ± ± ± ±

196 73 123 82 93 164

GSH

15 min 30 min 1h 3h 6h 18 h

209 239 244 226 253 239

± ± ± ± ± ±

MMP

15 min 30 min 1h 3h 6h 18 h

135 130 139 147 146 140

± ± ± ± ± ±

100 ␮M Cd

1836 1670 2344 1843 2678 2123

± ± ± ± ± ±

121 187 127* 88 173 73

1912 2383 2933 2992 3684 2444

± ± ± ± ± ±

84 99** 300** 242*** 479** 316

11 6 10 18 21 5

207 218 233 190 174 213

± ± ± ± ± ±

20 21 19 13* 17** 9

275 207 180 158 116 195

± ± ± ± ± ±

12 6 10 19 14 5

120 94 119 102 159 141

± ± ± ± ± ±

14 24 11 12 22 15

103 81 93 78 124 71

± ± ± ± ± ±

1000 ␮M Cd 1708 2293 3143 3277 4489 1909

± ± ± ± ± ±

83 124** 434*** 292*** 594*** 300

14 3* 9* 8** 13*** 2 **

213 233 140 132 86 157

± ± ± ± ± ±

18 15 27** 3** 23*** 16***

3 18* 12** 12** 8* 13*

94 73 69 73 130 97

± ± ± ± ± ±

15* 14** 19*** 12** 13 17

Table 2 Effect of NAC and BSO on ROS, GSH and apoptosis: freshly isolated sertoli–germ coculture (1.0 × 106 ) was treated with Cd (100 ␮M) at 37 ◦ C. BSO (500 ␮M) was added 20 h and NAC (10 mM) 10 min prior to Cd treatment. 1, 3 and 18 h for ROS, GSH and apoptosis, respectively. Data represents mean ± S.D. (n = 3). ***P < 0.001, **P < 0.01, *P < 0.05 as compared to Cd treatment, using one-way ANOVA.

ROS GSH % Apoptosis

Con

100 ␮M Cd

100 ␮M Cd + NAC

100 ␮M Cd + BSO

1933 ± 107 234 ± 21 2.08 ± 0.6

3836 ± 102 104 ± 12 32.31 ± 2.7

2037 ± 224** 199 ± 35** 9.9 ± 0.8***

5534 ± 877** 55 ± 24* 59.16 ± 3.5***

ing to ∼10% at 18 h. The apoptotic cells ranged from ∼6% to 30% (10–1000 ␮M Cd) at 6 h. On prolonging the exposure duration to 18 h, apoptotic cells appeared in the range of ∼10% to ∼55%. The necrotic cell population remained <3%. Results indicate the mode of cell death by apoptosis and not by necrosis. DNA condensation measured by DAPI staining (Fig. 2C) revealed significant DNA damage in almost 30% of sertoli–germ cells. These two techniques clearly support evidence of apoptosis by Cd. Consequent to establishing sertoli–germ coculture apoptosis by Cd, we investigated participation of caspase-3 and alterations in mitochondrial membrane potential ( ) along with release of mitochondrial apoptotic protein such as cytochrome c. In addition, the initial kinetics of calcium, MMP and ROS was monitored and the role of redox markers was re-established.

3.3. Cd causes mitochondrial membrane depolarization, release of cytochrome c and caspase-3 activation Mitochondrial participation in apoptosis has been confirmed by several researchers. A decrease in mitochondrial  , an early marker of apoptosis, was observed in a dose and time dependent fashion, in sertoli–germ coculture. The earliest loss in  was observed at 15 min by all the three Cd (10, 100 and 1000 ␮M) doses (Table 1) which maximized at 3 h and continued till 18 h, by the two higher Cd concentrations, even when significant apoptosis (∼36 and 55%) persisted. The release of cytochrome c from mitochondria was detected very early. Western blot analysis revealed increased expression of cytochrome c at 30 min tapering with time (Fig. 3B). Prolonged Cd treatment did not further release this protein, although the loss in mitochondrial potential continued till 3 h.

The next step was to ascertain involvement of caspase-3 during apoptosis. This event is subsequent to mitochondrial depolarization and release of cytochrome c. Once released into the cytosol, cytochrome c binds to its adaptor molecule, Apaf-1, which oligomerizes and then activates pro-caspase-9. Once activated, initiator caspases are responsible for cleaving and activating effector pro-caspases. Caspase-9 can signal downstream and activate procaspase-3 and -7. Effector caspases, in turn, cleave various proteins leading to morphological and biochemical features characteristic of apoptosis (Robertson and Orrenius, 2000). We find significant activation of caspase-3 at 6 h increasing 2-folds at 18 h (Fig. 4A). The steady activation of effector caspase correlates well with the rise in apoptotic population from 6 to 18 h. Caspase-3 inhibitor Z-DEVD-Fmk failed to completely inhibit apoptosis, suggesting caspase independent pathways, as well (Fig. 4C).

3.4. Cd induces ROS generation and GSH depletion, role of thiol modulators Significant ROS generation was actuated at 30 min reaching maximum at 6 h. This effect being time and concentration related (Table 1). There was a moderate increase in ROS even by the lowest Cd dose. At 30 min, 100 ␮M Cd resulted in a 1.2-fold rise in ROS becoming 2.0-folds at 6 h and with 1000 ␮M Cd, a still higher ROS production was evident at all time points. At 18 h, there was no further increase, implying that it is an early response caused by Cd insult. In the case of GSH, the earliest discernible decline in GSH was at 30 min by the median dose. Upto 6 h, the GSH levels continued to decrease in accordance to dose and time (Table 1). Ten micromolar Cd caused a mild GSH decline, whereas 1000 ␮M Cd exhibited a 2.5-

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Fig. 3. Effect of Cd on Cytochrome c: freshly isolated sertoli–germ coculture (3.0 × 106 ) was treated with 100 ␮M Cd for various time points at 37 ◦ C. Expression was analyzed by (A) western blot and (B) bar chart representing fold change. Data represents mean ± S.D. (n = 3). ***P < 0.001, **P < 0.01, *P < 0.05 as compared to control, using one-way ANOVA.

fold change. On prolonging the exposure time to 18 h, sertoli–germ coculture failed to exhibit any further depletion of GSH. Interplay of early biochemical response by Cd insult, as depicted in Fig. 5 displayed earliest rise in intracellular calcium, followed by mitochondrial membrane depolarization leading to ROS generation. We next attempted to delineate whether ROS and GSH modulations were responsible for the downstream events leading to cell death. For this, thiol modulators NAC and BSO were incorporated. N-Acetyl-cysteine (NAC) is an effective scavenger of free radicals, as well as, a major contributor in maintenance of cellular glutathione status (Aruoma et al., 1989). Buthionine sulfoximine (BSO) depletes intracellular GSH by inhibiting ␥-glutamyl cysteine synthetase activity (Armstrong and Jones, 2002). Cells were pretreated with NAC (10 mM) 10 min and BSO (500 ␮M) for 20 h, prior

to the addition of Cd and further incubated for 18 h to evaluate the degree of apoptosis. The period selected for ROS and GSH evaluation were 1 and 3 h respectively, based on our observations (Table 1). As depicted in Table 2, NAC effectively restored the GSH levels, lowered ROS and inhibited apoptosis. On the other hand, BSO enhanced the DCF fluorescence, further depleted GSH and apoptosis was 2folds higher than that of Cd treated cells. These results indicate the vital role played by ROS and GSH in modulating apoptosis. 3.5. Cd induces lipid peroxidation Lipid peroxidation is also an indicator of oxidative stress. Substantial rise in malondialdehyde was seen only at 6 and 18 h on Cd exposure (Table 3). The two higher Cd concentrations exhibited

Fig. 4. Effect of Cd on Caspase 3 activation: sertoli–germ coculture was treated with 100 ␮M Cd for different time intervals. Cells were then harvested, suspended in PBS. (A) Caspase 3 activation: DEVD-AFC (50 ␮M) was added for 30 min at RT. Cells collected, washed and resuspended in PBS. AFC fluorescence intensity was measured on flow cytometer (FL-1, 530 nm). Fluorescence results were expressed as (A) bar chart. Each bar represents mean ± S.D. (n = 3). ***P < 0.001, **P < 0.01, *P < 0.05 as compared to control, using one-way ANOVA (B) representative histogram. (C) Caspase 3 inhibition and apoptosis: cells were pretreated with caspase 3 inhibitor (Z-DEVD-FMK) for 1 h followed by Cd exposure for 18 h. Cells were collected, washed and resuspended in PBS. The FITC and PI fluorescence was measured using flow cytometer with FL-1 and FL-2 filters, respectively.

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Fig. 5. Effect of Cd on calcium, MMP and ROS at early time points: the sertoligerm coculture was preincubated with respective dyes for 30 min and treated with 100 ␮M Cd for 60 min. Each point represents mean ± S.D. (n = 3). ***P < 0.001, **P < 0.05, *P < 0.01 as compared to control, using one-way ANOVA.

almost similar response causing a 6-fold rise in MDA at 6 h, while at 18 h, the highest dose further raised MDA by 8-folds. 3.6. Cd reduces germ cell markers, LDH-X activity and tesmin LDH-X enzyme is a marker for normal spermatozoa metabolism. Dose dependent marked reduction in LDH-X activity was apparent only from 6 h (Table 4). The highest Cd concentration, exhibited almost 50% reduction which continued upto 48 h. At 72 h, the median dose i.e. 100 ␮M Cd suppressed LDH-X activity further i.e. 1.2-folds at 48 h to 1.8-folds at 72 h. Tesmin, an early marker of male germ cell differentiation was also affected by Cd. A time dependent suppression of cytosolic tesmin was observed by 100 ␮M Cd (Fig. 6A and B). Significant reduction in tesmin levels at 1hr and beyond, demonstrated interference of Cd in spermatogenic pathway. 4. Discussion It is well established that testicular oxidative stress is commonly induced under varied environmental conditions, leading to male infertility, illustrating the vulnerability of testis towards Cd - an oxidative stress inducer (Bagchi et al., 2000; Tremellen, 2008). This is likely to involve disruption of the blood-testis barrier via specific signal transduction pathways and signaling molecules (Siu et al., 2009). Although, the susceptibility of testis towards Cd lies in the

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sensitivity of blood-testis barrier in vivo, the present study relates to the mechanistic action of Cd on sertoli and germ cells. Among the two cell types, sertoli and leydig cells, central to the support of spermatogenesis, the former appears more sensitive to Cd. The cytotoxicity data of 100 ␮M Cd indicates ∼50% loss in cell viability in sertoli–germ coculture as compared to ∼40% in leydig cells (data not shown). In this study, we report Cd induced mitochondrial–caspase mediated apoptosis of primary rat sertoli–germ coculture and the pivotal role of ROS. Under normal conditions, ROS is mainly generated in mitochondria and is rapidly scavenged by cellular antioxidants (Balaban et al., 2005). Cadmium is suspected to act on the cell redox system by various mechanisms. It is reported to elevate ROS generation by depleting cellular glutathione and antioxidant enzymes, such as superoxide dismutase (SOD) and catalase (CAT) (Bagchi et al., 2000; Shaikh et al., 1999), even though being a redox-inactive metal that does not catalyze a Fenton-type reaction (Ercal et al., 2001; Stohs and Bagchi, 1995). Cadmium also produces ROS by inhibiting electron transfer chain in the mitochondria (Wang et al., 2004). Mitochondria plays a crucial role in Cd induced toxicity (Pulido and Parrish, 2003). It has been proposed that Cd initially binds with protein thiols in mitochondrial membranes, which may intensively affect mitochondrial electron transport chain promoting an elevation of ROS and causing mitochondrial permeability transition pore (Belyaeva et al., 2001; Dorta et al., 2003; Shih et al., 2004). In the present investigation, we find that Cd causes mitochondrial membrane depolarization faster within 15 min resulting in further increase of ROS generation, significantly evident at 30 min (Table 1). The study also demonstrates that ROS acts as a critical mediator of apoptosis supported by mitigation of apoptosis by N-acetylcysteine and its augmentation by buthionine sulfoximine. An interplay of initial biochemical response indicated an increased intracellular calcium resulting from the disturbance of calcium homeostasis caused by cadmium. Cd2+ is known to bind to and interact with receptor proteins on the cell surface, with ion channel proteins, or with intracellular proteins controlling Ca2+ homoeostasis (Benters et al., 1997). These calcium signals may further impact the mitochondrial function causing depolarization and subsequent cell death. Cadmium induces cell death by apoptosis by either caspasedependent (Hossain et al., 2009; Wang et al., 2009) or caspaseindependent mechanisms (Kim and Soh, 2009; Mao et al., 2007; Shih et al., 2004). In our study, although we observed mitochondrial membrane depolarization as early as 15 min, the effector caspase-3 activation took longer and was seen at 3 h, illustrating the steady influence of Cd on death machinery upto 18 h. Since caspase-3 specific inhibitor, could not abrogate the cell death completely, caspase independent apoptotic pathways may also be operative. Apoptotic proteins such as apoptosis-inducing factor (AIF) and endoG (endonuclease G) are reported to be translocated from mitochondria to nucleus during Cd induced apoptosis in certain cell types (Lemarieˇı et al., 2004; Shih et al., 2004) which act directly on the nuclear machinery causing cell death. Recently, Kim and Soh (2009), demonstrated caspase independent apoptosis in testis of rats treated with Cd. Conflicting reports on the underlying mechanism of Cd-induced apoptosis may be due to variations in cell lines and/or experimental conditions. Cadmium is known to mimic estrogen, thereby raising the possibility that FasL of spermatogenic cell source might be the ligand for Fas expressed on various cell types of spermatogenic origin (Fukuzawa et al., 2003). So far, we have studied the intrinsic pathway engaging mitochondrial depolarization and caspase-3 activation, but the possibility of extrinsic pathway involving caspase-8 activation may play some role. As such, the possibility of multiple cellular responses induced by Cd cannot be ignored.

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S. Khanna et al. / Toxicology 287 (2011) 46–53

Table 3 Effect of Cd on Lipid peroxidation: freshly isolated sertoli–germ coculture (1.0 × 106 ) was treated with Cd (10, 100 and 1000 ␮M) for different time intervals. Absorbance of MDA levels were measured at 532 nm. Data represents mean ± S.D. (n = 3). ***P < 0.001, **P < 0.01, *P < 0.05 as compared to control,using one-way ANOVA. 10 ␮M Cd

Con 1h 3h 6h 18 h

0.057 0.068 0.059 0.060

± ± ± ±

0.002 0.01 0.006 0.007

0.057 0.060 0.133 0.109

± ± ± ±

100 ␮M Cd

0.006 0.002 0.011 0.008

0.069 0.074 0.62 0.21

± ± ± ±

0.011 0.008 0.033*** 0.023*

1000 ␮M Cd 0.06 0.07 0.59 0.81

± ± ± ±

0.003 0.008 0.037*** 0.013***

Table 4 Effect of Cd on LDH-X: the sertoli -germ coculture was treated with Cd (10, 100 and 1000 ␮M) for different time interval. After incubation, equal volume of Reaction Mixture was added to the supernatant for 30 min at RT in dark and absorbance measured at 490–500 nm. Data represents mean ± S.D. (n = 3). ***P < 0.001, **P < 0.01, *P < 0.05 as compared to Cd treatment, using one-way ANOVA. 10 ␮M Cd

Con 6h 24 h 48 h 72 h

1.67 1.60 1.63 1.70

± ± ± ±

0.1 0.21 0.14 0.05

1.48 1.3 1.56 1.5

± ± ± ±

100 ␮M Cd

0.058* 0.15 0.15 0.08

1.37 1.29 1.28 0.92

± ± ± ±

0.06** 0.19 0.13** 0.03***

1000 ␮M Cd 0.72 0.821 0.83 1.51

± ± ± ±

0.03*** 0.13** 0.12*** 0.05

Fig. 6. Effect of Cd Tesmin: the sertoli–germ coculture was treated with 100 ␮M Cd for different time periods. (A) Cytosolic levels of tesmin were measured by western blotting. (B) Quantitative data is shown as bar chart. Each bar represents mean ± S.D. (n = 3). ***P < 0.001, **P < 0.01, *P < 0.05 as compared to Cd treatment, using one-way ANOVA.

High levels of lipid peroxides by Cd (6–8-folds) in our study are in agreement with previous reports of Cd and LPO (Yadav and Khandelwal, 2008). Peroxidation of sperm membrane lipids is generally considered as the first marking point of germ cell damage induced by reactive oxygen intermediates. Significant decrease of ∼1.8-folds in LDH activity (mainly of LDH-X), a consequence of enhanced lipid peroxidation after exposure to Cd may be due to disintegration of the mitochondrial membrane ultrastructure which in turn affects the membrane bound LDH-X function. LDH-X, a unique isoenzyme of lactate dehydrogenase, in inner mitochondrial membrane of spermatogenic cells of mature and developing testis, plays an important role in transferring hydrogen from cytoplasm to mitochondria by redox coupling ␣-hydroxy acid/␣-keto acid related to spermatozoal metabolism (Gu et al., 1989). This could be one of the contributory factors leading to reduced male sperm concentration and sperm motility on Cd exposure (Benoff et al., 2009). Further, decrease in cytosolic tesmin, which is a specific protein for germ cell differentiation was also observed, which agrees well with the study of Matsuura et al. (2002) demonstrating nucleocytoplasmic shuttling of tesmin in response to stress and Cd treatment in Cos-1 cell line. 5. Conclusion To sum up, our investigation demonstrates significant increases in calcium, ROS and MDA levels, accompanied with substantial decrease in GSH in sertoli–germ coculture exposed to low doses of Cd. Also, intrinsic pathway engaging mitochondrial depolarization, cytochrome c release and caspase-3 activation appears to be one of the underlying mechanisms of Cd induced testicular dam-

age, ROS acting as the critical mediator of cell death. Furthermore Cd disrupted the spermatogenic pathway as evident by suppression in tesmin and LDH-X levels. Conflict of interest The authors declare that there are no conflicts of interest. Acknowledgements We are grateful to Director, IITR for his keen interest. One of us (S. Khanna) is thankful to Council of Scientific and Industrial Research (CSIR), New Delhi for the award of Senior Research Fellowship. We are also thankful to Dr. Y. Shukla for providing flow cytometer facility. Financial assistance of CSIR Supra Institutional Project (SIP-08) is gratefully acknowledged. References Adhikari, N., Sinha, N., Saxena, D.K., 2000. Effect of lead on Sertoli–germ cell coculture of rat. Toxicol. Lett. 116, 45–49. ATSDR, 2007. The 2007 CERCLA Priority List of Hazardous Substances. Agency for Toxic Substance and Disease Registry, Department of Health and Human Services, Atlanta, GA. Armstrong, J.S., Jones, D.P., 2002. Glutathione depletion enforced the mitochondrial permeability transition and causes cell death in Bcl-2 over expressing HL60 cells. FASEB J. 16, 1263–1265. Aruoma, O.I., Halliwell, B., Hoey, B.M., Butlar, J., 1989. The antioxidant action of N-acetylcysteine, its reaction with hydrogen peroxide, hydroxyl radical, superoxide and hypochlorous acid. Free Radic. Biol. Med. 6, 593–597. Bagchi, D., Joshi, S.S., Bagchi, M., Balmoori, J., Benner, E.J., Kuszynski, C.A., Stohs, S.J., 2000. Cadmium and chromium induced oxidative stress. DNA damage and apoptotic cell death in cultured human chronic myelogenous leukemic K562

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