Influence of cadmium on murine thymocytes: Potentiation of apoptosis and oxidative stress

Toxicology Letters 165 (2006) 121–132

Influence of cadmium on murine thymocytes: Potentiation of apoptosis and oxidative stress Neelima Pathak, Shashi Khandelwal ∗ Industrial Toxicology Research Centre, P.O. Box 80, Mahatma Gandhi Marg, Lucknow 226001, India Received 15 December 2005; received in revised form 13 February 2006; accepted 13 February 2006 Available online 6 March 2006

Abstract Cadmium (Cd) is a well-known environmental carcinogen and a potent immunotoxicant. It induces thymocyte apoptosis in vitro. However, the mode of action is unclear. In this study, we examined the effect of Cd (10, 25 and 50 ␮M) on mitochondrial membrane potential and caspase-3 as well as oxidative stress markers in murine thymocytes. The cadmium induced apoptosis occurred in a concentration and time dependent manner. The early markers of apoptosis-loss in mitochondrial membrane potential and caspase-3 activation were evident as early as 1.5 h by 50 ␮M Cd. Enhanced reactive oxygen species (ROS) generation and glutathione (GSH) depletion were observed at 60 min, prior to the lowering of mitochondrial membrane potential. The Cd induced DNA damage as depicted by internucleosomal fragmentation on agarose and histone associated mono- and oligonucleosomes detection by ELISA, corrobated with the apoptotic DNA (sub-G1 population) and total apoptotic cells by Annexin V binding assay. The number of cells in sub-G1 population increased to 66% at 50 ␮M Cd concentration and the distribution of early and late apoptotic cells was 47% and 15%, respectively. Addition of N-acetylcysteine and pyrrolidine dithiocarbamate (thiol antioxidants) to the Cd treated cells, lowered the sub-G1 population, inhibited the ROS generation and raised the GSH levels. Buthionine sulfoximine (GSH depletor) on the other hand, enhanced both the ROS production and the sub-G1 fraction. These results clearly demonstrate the apoptogenic potential of Cd in murine thymocytes, following mitochondrial membrane depolarization, caspase activation and ROS and GSH acting as critical mediators. © 2006 Elsevier Ireland Ltd. All rights reserved. Keywords: Cadmium; Murine thymocytes; Apoptosis; Oxidative stress; Caspase-3

Abbreviations: BSO, buthionine sulfoximine; Cd, cadmium; CMF-DA, 5 -chloromethylfluorescein diacetate; DCFH-DA, 2 ,7 dichlorofluorescein diacetate; DEVD-AFC, benzyloxy-carbonyl-Asp-Glu-Val-Asp-7-amino-4-trifluoromethyl-coumarin; DEX, dexamethasone; DMSO, dimethyl sulfoxide; EDTA, ethylenediamine tetracetic acid; FBS, fetal bovine serum; FITC, fluorescein-5-isothiocyanate; GSH, glutathione; MTT, 3-(4,5-dimethyl-2-yl)-2,5-diphenyl tetrazolium bromide; NAC, n-acetyl cysteine; PBS, phosphate buffered saline; PDTC, pyrrolidine dithiocarbamate; PI, propidium iodide; Rh 123, rhodamine 123; ROS, reactive oxygen species; TE buffer, Tris–EDTA buffer;
ψ, membrane potential ∗ Corresponding author. Tel.: +91 522 262 7586; fax: +91 522 262 8227. E-mail address: skhandelwal [email protected] (S. Khandelwal). 0378-4274/$ – see front matter © 2006 Elsevier Ireland Ltd. All rights reserved. doi:10.1016/j.toxlet.2006.02.004

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1. Introduction Cadmium (Cd) is a common environmental contaminant having a long biological half-life and multiorgan toxicity. In addition to hepatic and renal toxicity, Cd is a potent immunotoxicant both in mammalian and nonmammalian species. Thymus, an important primary lymphoid organ, is the target organ of Cd induced immunotoxicity. In an early in vivo study, Morselt et al. (1988) observed thymic damage and modulation of proliferative rate in Cd exposed rat thymocytes. A marked loss of thymus weight, T-cell depletion and thymic atrophy are also reported under in vivo conditions (Borgman et al., 1986; Mackova et al., 1996; Liu et al., 1999). Recently, Lafuente et al. (2004) demonstrated that Cd, when given to rats in drinking water showed a differential effect on the blood lymphocyte phenotyping. Apoptosis is an important mechanism in T-cell development and differentiation. Modulation of apoptosis results in dysfunction of T-cell dependent immune responses. Many immunotoxins such as cadmium, tributyltin, lead, nickel, deltamethrin, 2,3,7,8tetrachlorobenzo-p-dioxin (TCDD),
9 -tetra hydrocannabinol and arsenic have been reported to act through induction of thymocyte apoptosis (Enan et al., 1996; Fujimaki et al., 2000; Okada et al., 2000; De la Fuente et al., 2002; Mckallip et al., 2002; Camacho et al., 2004). Cadmium has been shown to induce apoptosis in mice thymocytes and in other immune cell lines (El Azzouzi et al., 1994; Tsangaris and Tzortzatou-Stathopoulou, 1998; Fujimaki et al., 2000). A number of in vitro experiments have demonstrated potentiation of murine thymocyte apoptosis by Cd. Sustained increase of intracellular Ca2+ preceding caspase-3 activation and DNA fragmentation was shown by Shen et al. (2001) and high susceptibility of CD8+ thymocytes followed by a marked decrease in CD4+ /CD8+ ratio was observed by Dong et al. (2001). Reactive oxygen species are involved in apoptosis as well as in cell proliferation (Mates and Sanchez-Jimenez, 2000). ROS may induce cell death directly or act as intracellular messengers during cell death induced by various other kinds of stimuli. Cadmium is reported to cause oxidative stress, i.e. increased lipid peroxidation, ROS generation and alterations in glutathione (GSH) and related enzymes, under both in vitro and in vivo conditions (Stohs and Bagchi, 1995; Hart et al., 1999; Thevenod et al., 2000; De la Fuente et al., 2002). The role of ROS in Cd induced apoptosis (Pulido and Parrish, 2003; Lemarie et al., 2004) is further supported by activation of redox sensitive AP-1 transcription factor and alteration in GSH metabolism prior to apoptosis (Hart et al., 1999). Immune cells, because of major production of

ROS, are highly sensitive to oxidative stress. ROS plays a vital role in their immune function, acting as intracellular signals and oxidant–antioxidant balance is important for maintenance of the immune response. Based on the apoptogenic potential of Cd and the crucial role of ROS in immune function, the current study was designed to investigate the Cd induced apoptogenic signalling pathway in murine thymocytes. The involvement of ROS and GSH, specifically in mediating mitochondrial caspase pathway, was thoroughly evaluated. In the present study, we demonstrate Cd induced apoptosis as early as 6 h and ROS generation and GSH depletion precede mitochondrial membrane depolarization and caspase-3 activation. 2. Materials and methods 2.1. Chemicals All the chemicals were of highest grade purity available. Cadmium chloride (CdCl2 ), RNase A, RPMI 1640, antibiotic–antimycotic solution, Dulbecco’s phosphate buffered saline (PBS), fetal bovine serum (FBS), agarose, 3(4,5-dimethyl-2-yl)-2,5-diphenyl tetrazolium bromide (MTT), 2 ,7 -dichlorofluorescein diacetate (DCFH-DA) and all other chemicals were purchased from Sigma–Aldrich, USA. Rhodamine 123 (Rh 123) and 5 -chloromethylfluorescein diacetate (CMF-DA) from Molecular Probes and propidium iodide (PI) was from Calbiochem. Cell Death Detection Sandwich ELISA kit was purchased from Roche, Germany, caspase-3 fluorometric protease assay kit from Chemicon, USA, and Annexin V-FITC Apoptosis Detection kit from Pharmingen (Becton Dickinson Company). 2.2. Preparation of thymocyte suspension Thymus was dissected from male BALB/c mice (4–6 weeks old) and single cell suspension prepared under aseptic conditions. The suspension was passed through 100 ␮M stainless steel mesh and suspended in complete cell culture medium (RPMI 1640 containing HEPES and 2 mM glutamine, supplemented with 10% FBS and 1% antibiotic–antimycotic solution). The cell density was adjusted to ca. 1.5 × 106 cells/ml and the viability of the freshly isolated cells was always over 95% (trypan blue exclusion test). 2.3. Determination of cell survival The number of live cells after treatment with various Cd concentrations (1, 10, 25, 50 and 100 ␮M) were determined by the MTT reduction method of Mosmann (1983). The relative survival rate was calculated as the percentage of control cells. The cells (1 × 104 ) were seeded in 96-well plate and incubated with Cd for 24 h at 37 ◦ C in a CO2 incubator. Ten microliters of MTT (5 mg/ml PBS) was added, 4 h prior to completion of

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the incubation time, followed by centrifugation at 1200 × g for 10 min. After removal of supernatant, 100 ␮l of dimethyl sulfoxide (DMSO) was added and the absorbance read at 530 nm after 5 min, in a microplate reader (Synergy HT of BIO-TEK, USA). 2.4. Assessment of apoptosis Flow cytometry was used to assess the membrane and nuclear events during apoptosis. The membrane events were analysed by measuring the binding of FITC-Annexin V protein to the phospholipid phosphatidylserine present on the external surface of the apoptotic cell membrane. The assay was performed with a two colour analysis of FITC-labelled Annexin V binding and PI uptake using the Annexin VFITC Apoptosis Detection kit. Positioning of quadrants on Annexin V/PI dot plots was performed and live cells (Annexin V− /PI− ), early/primary apoptotic cells (Annexin V+ /PI− ), late/secondary apoptotic cells (Annexin V+ /PI+ ) and necrotic cells (Annexin V− /PI+ ) were distinguished (Vermes et al., 1995). Therefore, the total apoptotic proportion included the percentage of cells with fluorescence Annexin V+ /PI− and Annexin V+ /PI+ . Briefly, 1.5 × 106 cells/ml were incubated with Cd (10, 25 and 50 ␮M) for 6 and 18 h. Harvested cells were resuspended in 1 ml binding buffer (1×). An aliquot of 100 ␮l was incubated with 5 ␮l Annexin V-FITC and 10 ␮l PI for 15 min in dark at room temperature and 400 ␮l binding buffer (1×) was added to each sample. The fluorescein5-isothiocyanate (FITC) and PI fluorescence were measured through FL-1 filter (530 nm) and FL-2 filter (585 nm), respectively, on BD-LSR flow cytometer using Cell Quest software and 10,000 events were acquired. The nuclear events including hypodiploid DNA were determined by cell cycle studies. A population of 1.5 × 106 cells/ml was incubated with Cd (10, 25 and 50 ␮M) for 3, 6 and 18 h. Harvested cells were washed with PBS and fixed by dropby-drop addition of ice cold 70% ethanol and stored at 4 ◦ C overnight. The fixed cells were harvested, washed with PBS and suspended in 1 ml PBS. Two hundred microliters of phosphate citrate buffer (pH 7.8) was added and the cells incubated for 60 min at room temperature. After centrifugation, the cells were resuspended in 0.5 ml of PI stain (10 mg PI, 0.1 ml Triton X-100 and 3.7 mg ethylenediamine tetracetic acid (EDTA) in 100 ml PBS) and 0.5 ml of RNase A (50 ␮g/ml) and further incubated for 30 min in dark. The PI fluorescence was measured through a FL-2 filter (585 nm) on flow cytometer and 10,000 events were acquired (Darzynkiewicz et al., 1992).

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Tris–HCl, pH 7.5, 20 mM EDTA and 0.5% Triton X-100) on ice for 30 min. The DNA in lysed solution was extracted with phenol/chloroform and precipitated with 3 M sodium acetate (pH 5.2) and cold ethanol. After repeated washings, the DNA was dissolved in TE buffer (10 mM Tris–HCl, pH 8.0, and 1 mM EDTA). The purity of DNA at 260 and 280 nm absorbance ratio was between 1.7 and 1.9. DNA (2 ␮g) was then loaded on 0.7% agarose gel and electrophoresis carried out. The bands were visualized by ethidium bromide staining under UV light. The DNA fragmentation (histone associated mono- and oligonucleosomes) was measured using Cell Death Detection Sandwich ELISA kit (Roche, Germany). Briefly, 1.5 × 106 cells/ml were incubated with Cd (10, 25 and 50 ␮M) for 6 and 18 h at 37 ◦ C in a CO2 incubator, out of which 1 × 105 cells were transferred to a clear tube. After centrifugation at 200 × g for 10 min, the cells were suspended in incubation buffer for 30 min at 15–25 ◦ C for lysis. Following centrifugation at 20,000 × g for 10 min, the supernatant was diluted 10-fold with incubation buffer and the nucleosomes in the sample were measured by ELISA. Modular microtitre plates were incubated overnight with 100 ␮l of the antihistone antibody (coating solution) under cold conditions. After aspirating the coating solution, 100 ␮l of incubation buffer (blocking solution) was added and allowed to stand for 30 min at 15–25 ◦ C. The plates were then washed with 200 ␮l of washing buffer and after removing the buffer carefully, the sample solution (100 ␮l) was added into each well and incubated for 90 min at 15–25 ◦ C. After washing the plates, 100 ␮l of conjugate solution (anti-DNA-peroxidase) was added to each well and the cells were further incubated for 90 min at 15–25 ◦ C, followed by washing. One hundred microliters of ABTS (22 -azido-di-[3-ethylbenzthiazoline sulfonate]) was then added and incubated for 10 min and the absorbance measured at 405 nm. 2.6. Measurement of caspase-3 activity Caspase-3 activity was measured using a commercial kit (Chemicon). Thymocytes (3.0 × 106 cells/ml) were incubated with Cd (10, 25 and 50 ␮M) for 1.5, 3 and 6 h at 37 ◦ C in a CO2 incubator. The cells were scraped and lysed on ice for 10 min using cell lysis buffer. The reaction buffer and benzyloxy-carbonyl-Asp-Glu-Val-Asp-7-amino-4trifluoromethyl-coumarin (DEVD-AFC) substrate were then added and further incubated at 37 ◦ C for 2 h. The resultant fluorescence was measured, at excitation and emission wavelengths of 400 and 505 nm, respectively, on a microplate reader.

2.5. DNA fragmentation and cell death detection

2.7. Mitochondrial membrane potential analysis

The internucleosomal fragmentation pattern (DNA ladder) was carried out by Agarose gel electrophoresis. Two milliliters (1.5 × 106 thymocytes/ml) was incubated with Cd (10, 25 and 50 ␮M) for 6 and 18 h at 37 ◦ C in a CO2 incubator. At the end of incubation, cells were pelleted by centrifugation at 200 × g for 10 min and the pellet was lysed with 0.5 ml lysis buffer (10 mM

Rhodamine 123 uptake by mitochondria is directly proportional to its membrane potential. 1.5 × 106 cells/ml were exposed to 10, 25 and 50 ␮M Cd concentration for 60 min, 1.5, 3, 6 and 18 h and were incubated with Rh 123 (5 ␮g/ml final concentration) for 60 min in dark at 37 ◦ C. The cells were harvested and suspended in PBS. The mitochondrial membrane

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potential was measured by the fluorescence intensity (FL-1, 530 nm) of 10,000 cells on flow cytometer (Bai et al., 1999). 2.8. Reactive oxygen species (ROS) measurement The generation of ROS was detected by DCF fluorescence. 1.5 × 106 cells/ml were incubated with 10, 25 and 50 ␮M Cd for 60 min, 1.5, 3, 6 and 18 h. DCFH-DA (100 ␮M final concentration) was added simultaneously to the medium. The cells were harvested and suspended in PBS. ROS generation was measured by the fluorescence intensity (FL-1, 530 nm) of 10,000 cells on flow cytometer (Wang et al., 1996). 2.9. Glutathione measurement 5-Chloromethylfluorescein diacetate was used to monitor the cellular level of GSH. 1.5 × 106 cells/ml were incubated with 10, 25 and 50 ␮M Cd for 60 min, 1.5, 3, 6 and 18 h. The cells were incubated with CMF diacetate (1 ␮M final concentration) for 30 min in dark at 37 ◦ C. After harvesting, the cells were suspended in PBS and GSH was measured by the fluorescence intensity (FL-1, 530 nm) of 10,000 cells on flow cytometer (Okada et al., 2000). 2.10. Statistical analysis Significance of mean of different parameters between the treatment groups were analysed 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.

3. Results 3.1. Effect of Cd on cell survival As shown in Fig. 1, a concentration dependent loss in cell viability was observed in thymocytes exposed to

Fig. 1. Effect of Cd on viability in murine thymocytes. Freshly isolated thymocytes (1.5 × 104 ) were treated with Cd (1–100 ␮M) for 24 h. Absorbance was measured at 530 nm. Each point represents mean ± S.D. (n = 3). *** p < 0.001 and ** p < 0.01 as compared to control, using one-way ANOVA.

Fig. 2. Effect of Cd on DNA fragmentation by (0.7%) agarose gel electrophoresis. Freshly isolated thymocytes (3 × 106 ) were treated with Cd (10–50 ␮M) for 6 and 18 h.

varying concentration of Cd (1–100 ␮M) over a 24 h period. The 25 ␮M Cd concentration caused ∼45% loss in cell viability. In other words, almost 55% cells remained viable as compared to ∼18% at 100 ␮M Cd. Following the exploratory experiments, all subsequent experiments were done with 10, 25 and 50 ␮M concentration of Cd, to evaluate its apoptogenic potential. The exposure time duration of Cd was not exceeded beyond 18 h, since the DNA damage as determined by total apoptotic cells using Annexin V binding assay and sub-G1 population even at 50 ␮M Cd, did not show any further change (data not shown). 3.2. Effect of Cd on DNA damage Several techniques were utilized to demonstrate Cd induced apoptotic DNA damage. An internucleosomal DNA damage as depicted by DNA ladder on agarose gel is shown in Fig. 2. Distinct DNA fragments were observed only at 18 h, the effect being dose related. At 6 h, there was no observable DNA ladder by any of the Cd concentrations tested. Dexamethasone (DEX, 10 ␮M) served as a positive control. This assay provides a qualitative assessment of apoptosis in a cell system and need to be confirmed by other sensitive methods. To quantitate the histone associated DNA fragments, Sandwich ELISA method was employed. Thymocytes when exposed to Cd (10, 25 and 50 ␮M) exhibited enhanced absorbance in a concentration and time dependent fashion (Table 1). Significant DNA fragmentation (1.2-fold) was observed at 6 h, only with the 50 ␮M Cd concentration. Later at 18 h, even the lowest concentration of Cd (10 ␮M) caused substantial increase in DNA fragmentation. DEX used as a positive control, also exhibited enhanced absorbance at 6 and 18 h. These results indicate that 50 ␮M Cd caused DNA damage as early as 6 h. This was further confirmed with

N. Pathak, S. Khandelwal / Toxicology Letters 165 (2006) 121–132 Table 1 Effect of Cd on DNA fragmentation (mono- and oligonucleosomes) Groups

O.D. 405 (6 h)

O.D. 405 (18 h)

Control Cd 10 ␮M Cd 25 ␮M Cd 50 ␮M DEX 10 ␮M

100.0 ± 4.6 118.0 ± 9.8 120.5 ± 2.1 124.5 ± 6.1* 120.8 ± 11.4

100.0 ± 6.5 129.0 ± 4.9** 156.5 ± 0.5*** 209.0 ± 13.1*** 148.5 ± 6.5***

Freshly isolated thymocytes (1.5 × 106 ) were treated with Cd (10–50 ␮M) for 6 and 18 h at 37 ◦ C. DEX (10 ␮M) was used as a positive control. DNA fragments were determined by Cell Death Detection ELISA. Each value represents mean ± S.D. (n = 3). * p < 0.05 as compared to control, using one-way ANOVA. ** p < 0.01 as compared to control, using one-way ANOVA. *** p < 0.001 as compared to control, using one-way ANOVA.

the flow cytometry data obtained on apoptotic DNA and the total (early + late) apoptotic cells. In cell cycle studies by flow cytometry, the fraction of hypodiploid cells gradually increased with dose and time (Fig. 3). At 6 h, only the 50 ␮M Cd concentration showed a significant increase in sub-G1 population (12.6%) when compared to the control population. Later at 18 h, with advancing Cd concentration, the number of apoptotic cells increased from 13% (10 ␮M Cd) to 66% (50 ␮M Cd). The apoptotic cells in the normal population over this period remained below 2.3%. The Annexin V binding assay measures the fluorescence generated by Annexin binding with phosphatidylserine of apoptotic cells. The basal level of dead cells as evaluated at 6 h, was less than 0.3%.

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On Cd exposure, the apoptotic cells show significant increase (Fig. 4). The apoptotic cells were raised from 3.2% to 14.5% (10–50 ␮M Cd) at 6 h. On prolonging the exposure duration to 18 h, the apoptotic cells appeared in the range of 14.4–62.2% by 10–50 ␮M Cd. The total (early + late) apoptotic cells at 6 and 18 h agreed well with the fraction of subG1 population at all concentrations of Cd. These two assays substantially supported evidence of apoptosis by Cd. Consequent to establishing significant thymocyte apoptosis by Cd, we investigated the participation of caspase-3 and changes in mitochondrial membrane potential. In addition, the role of ROS and GSH were also monitered at early time points with and without thiol modulators. 3.3. Effect of Cd on mitochondrial membrane potential and caspase-3 activity Rh 123, a lipophilic cationic fluorescent dye, is selectively taken up by mitochondria and its uptake is directly proportional to mitochondrial
ψ of cells (Scaduto and Grotyohann, 1999). Being an early marker of apoptosis, a significant decrease in mitochondrial
ψ occurred as early as 1.5 h, only at 50 ␮M Cd concentration (p < 0.05) as shown in Fig. 5. Later on at 3 and 6 h, the loss in
ψ became more prominent and was concentration related. Simultaneously, there was a dose and time dependent enhancement in caspase-3 activity (Table 2). The release of cytochrome c in the cytosol as a consequence of mitochondrial transmembrane depolarization is reported to activate caspase-3 and 9 in several cell types (Habeebu

Fig. 3. Effect of Cd on apoptosis. Freshly isolated thymocytes (1.5 × 106 ) were treated with Cd (10–50 ␮M) for 18 h. DEX (10 ␮M) was used as a positive control. PI fluorescence was measured using a flow cytometer with FL-2 filter. Results were expressed as histogram representing the percentage of sub-G1 population (A, 18 h) and the percentage of apoptotic cells obtained from the histogram statistics (B). Each value represents mean ± S.D. (n = 3). *** p < 0.001 and ** p < 0.01 as compared to control, using one-way ANOVA.

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Fig. 4. Effect of Cd on percent distribution of apoptotic and necrotic cells. Freshly isolated thymocytes (1.5 × 106 ) were treated with Cd (10–50 ␮M) for 6 and 18 h and cell distribution was analysed using Annexin V binding and PI uptake. The FITC and PI fluorescence was measured using flow cytometer with FL-1 and FL-2 filters, respectively. Results were expressed as 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+ ).

et al., 1998; Li et al., 2000; Gennari et al., 2003). Significant lowering of mitochondrial
ψ at 1.5 h by 50 ␮M Cd, led to a 1.9-fold increase in caspase-3 activity. By DEX, the loss in mitochondrial
ψ was 1.4-fold and

the increase in caspase-3 activity was 2.9-fold at 3 h. These results indicate that there is mitochondrial membrane depolarization, coupled with caspase-3 activation during Cd induced thymic apoptosis.

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Fig. 5. Effect of Cd on mitochondrial membrane potential. Freshly isolated thymocytes (1.5 × 106 ) were treated with Cd (10–50 ␮M) for 60 min, 1.5, 3, 6 and 18 h at 37 ◦ C. DEX (10 ␮M) was used as a positive control. Rhodamine 123 was added and the cells were incubated for 60 min. Rh 123 fluorescence was measured using a flow cytometer with FL-1 filter. Fluorescence results were expressed as representative histogram (A, 3 h) and mean fluorescence obtained from the histogram statistics (B). Each bar represents mean ± S.D. (n = 3). *** p < 0.001 and * p < 0.05 as compared to control, using one-way ANOVA.

3.4. Effect of Cd on ROS generation

3.5. Effect of Cd on intracellular GSH

DCFH-DA, a permeable dye, is cleaved to form non-fluorescent dichlorofluorescein (DCFH) in the cells, which gets oxidized to fluorescent dichlorofluorescein (DCF) by ROS. The DCF fluorescence, proportionate to the ROS levels in the cells, was monitored on flow cytometer (Fig. 6). The dose and time dependent generation of ROS continued to rise till 6 h and the 50 ␮M Cd concentration exhibited a 3.7-fold increase at 6 h. However at 18 h, when DNA damage was more pronounced, the DCF fluorescence diminished, as shown in the figure. Significant ROS generation in the thymocytes was observed as early as 60 min with the two higher Cd concentrations (25 and 50 ␮M), prior to the lowering in mitochondrial membrane potential (p < 0.05) which was seen only at 1.5 h with 50 ␮M Cd.

The intensity of CMF fluorescence was well correlated with biochemically estimated content of GSH (Chikahisa et al., 1996). The three concentrations of Cd exhibited differential effect on GSH. The 50 ␮M Cd caused significant GSH depletion (p < 0.05) at 60 min and 1.5 h. However, the two lower Cd concentrations (10 and 25 ␮M) showed a modest increase at the same time. This phenomenon is related to the role of GSH. When the cells are oxidatively challenged, GSH synthesis increases as a protective mechanism. But with the higher dose of toxicant, the GSH synthesis is unable to compete with oxidative stress, and so the levels tend to decline. This effect was visible with the various concentrations of Cd (10–50 ␮M) and is shown in Fig. 8. At later time periods, there was a dose dependent depletion of GSH. The 50 ␮M Cd concentration caused a 2.7-fold reduction in GSH levels at 6 h, which further declined with time and at 18 h, the depletion became 3.9-fold. It appears that GSH and ROS act as intracellular signals causing mitochondrial membrane permeabilization at 1.5 h and the triggering effect being translated to DNA damage evident at 6 h with 50 ␮M Cd and at 18 h with all Cd concentrations.

Table 2 Effect of Cd on caspase-3 activity Groups

Cd 10 ␮M Cd 25 ␮M Cd 50 ␮M DEX 10 ␮M

Fluorescence (% of control) 1.5 h

3.0 h

6.0 h

113 175 191 178

205 310 465 298

112 175 193 152

Freshly isolated thymocytes (3 × 106 ) were treated with Cd (10–50 ␮M) for 1.5, 3 and 6 h at 37 ◦ C. DEX (10 ␮M) was used as a positive control. The enzyme activity was determined by Fluorometric Protease Assay kit. The fluorescence was measured at Ex—400 nm and Em—505 nm. The value represents mean of two independent experiments.

3.6. Effect of thiol modulators on Cd induced apoptosis To further investigate whether ROS generation is a crucial mediator in Cd induced apoptosis, thiol modulators were incorporated. Two different thiol antioxidants: N-acetyl cysteine (NAC) and pyrrolidine dithiocarbamate (PDTC) were tested. NAC can raise intracellular

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Fig. 6. Effect of Cd on generation of ROS. Freshly isolated thymocytes (1.5 × 106 ) were incubated with DCFH-DA and Cd (10–50 ␮M) for 60 min, 1.5, 3, 6 and 18 h at 37 ◦ C. DCF fluorescence was measured using a flow cytometer with FL-1 filter. Fluorescence results were expressed as representative histogram (A, 6 h) and mean fluorescence obtained from the histogram statistics (B). Each bar represents mean ± S.D. (n = 3). *** p < 0.001, ** p < 0.01 and * p < 0.05 as compared to control, using one-way ANOVA.

glutathione levels and thereby protect the cells from the effects of ROS (Aruoma et al., 1989). In addition, the –SH group of the reagent can react directly with radicals. PDTC, an antioxidant (Schreck et al., 1991), is also a potent chelator of various metals including Cd (Shimada et al., 1991). Cells were pretreated with NAC (10 mM) and PDTC (10 ␮M), 10 min prior to the addition to Cd (25 ␮M) for 6 h. For GSH depletion, buthionine sulfoximine (BSO) was included, which acts by inhibiting ␯-glutamyl cysteine synthetase activity (Armstrong and Jones, 2002). The cells were pretreated with BSO (500 ␮M) for 20 h, after which the medium was replaced, Cd (25 ␮M) was added and the cells were further incubated for 6 h. For apoptotic DNA (sub-G1 population) the cells were incubated for 18 h. The 6 h time period for the evaluation of ROS and GSH was selected based on our experiments (Figs. 6 and 7). Maximum DCF fluorescence was observed at this time point. As shown in Fig. 8, NAC

and PDTC effectively inhibited the Cd induced apoptotic DNA, NAC being more potent than PDTC. In contrast, BSO caused an elevation in the sub-G1 population. To confirm that the suppression of cell death is associated with the antioxidant effects of NAC and PDTC, the influence on ROS generation was examined (Fig. 8). Both the antioxidants lowered the DCF fluorescence induced by Cd and raised the intracellular GSH levels. On the other hand, BSO enhanced the DCF fluorescence when the cells were depleted of GSH. In addition, NAC and PDTC pretreatment abolished the depolarization effect of Cd on the mitochondrial
ψ. The lowered Rh 123 fluorescence in the Cd exposed cells was enhanced drastically in the presence of NAC and PDTC, NAC being more effective. With BSO, the mitochondrial
ψ was further inhibited. These results indicated earliest response in ROS production followed by the lowering of mitochondrial
ψ and subsequent activation of caspase-3 would then lead

Fig. 7. Effect of Cd on glutathione levels. Freshly isolated thymocytes (1.5 × 106 ) were treated with Cd (10–50 ␮M) for 60 min, 1.5, 3, 6 and 18 h at 37 ◦ C. CMF-DA was added and the cells were incubated for 30 min. CMF fluorescence was measured using a flow cytometer with FL-1 filter. Fluorescence results were expressed as representative histogram (A, 6 h) and mean fluorescence obtained from the histogram statistics (B). Each bar represents mean ± S.D. (n = 3). *** p < 0.001 and * p < 0.05 as compared to control, using one-way ANOVA.

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Fig. 8. Effect of NAC, PDTC and BSO on Cd induced (A) ROS (DCF fluorescence), (B) GSH (CMF fluorescence), (C) mitochondrial membrane potential (Rh 123 fluorescence) and (D) apoptotic DNA (sub-G1 population). Freshly isolated thymocytes were pretreated with NAC (10 mM) and PDTC (10 ␮M) for 10 min and BSO (500 ␮M) for 20 h prior to Cd addition. Each bar represents mean ± S.D. (n = 3). *** p < 0.001 and ** p < 0.01 as compared to Cd group, using one-way ANOVA.

the cells to apoptose. Since increased ROS generation by the two higher concentrations and GSH depletion only with 50 ␮M Cd was observed at 60 min, the former appeared to be a more sensitive indicator in manifesting Cd induced oxidative stress. 4. Discussion In this study, we report the involvement of oxidative stress in cadmium induced apoptosis in murine thymocytes. Although, several groups (Stohs and Bagchi, 1995; Ercal et al., 2001; Ikediobi et al., 2004) suggest that Cd induced oxidative stress could be partially responsible for cell damage, little information on oxidative stress by Cd in murine thymocytes is available. However, Lag et al. (2002) showed the involvement of Bax and p53 and not of oxidative stress in Cd induced apoptosis of primary epithelial lung cells. Immune cells are particularly sensitive to oxidative stress because of their higher production of ROS, which play a vital role in their normal function (Meydani et al., 1995) as important signalling molecules in the regulation of various cellular processes (Kim et al., 2001). The oxidant–antioxidant balance is critical for maintaining

the integrity and functionality of membrane lipids, cellular proteins and nucleic acids and controlling signal tranduction and gene expression in immune cells. Staal et al. (1992), in their review article, reported that “adequate levels of GSH are required for mixed lymphocyte reactions, T-cell proliferation, T and B-cell differentiation, cytotoxic T-cell activity and natural Tcell activity. Decreasing GSH by 10–40% can inhibit completely T-cell activation in vitro. Thus, an intracellular GSH deficiency in lymphocytes has profound effects on immune functions”. Other abnormalities of immune function associated with decreased GSH levels are impaired IL-2 production and IL-2 responses and a shift to TH-2 response as compared to TH-1 (Peterson, 1998). Significant alterations in ROS and GSH as early as 60 min, strongly suggest their active participation in Cd induced apoptosis in murine thymocytes at 6 and 18 h. Mitochondrial membrane plays a crucial role in Cd induced toxicity (Pulido and Parrish, 2003). Mitochondria are the main target for damage by ROS, leading to necrosis and apoptosis as shown by Melendez and Davies (1996) and Kowaltowski et al. (1998). At the same time, there are reports stating direct

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or indirect interaction of Cd with mitochondria, promoting an elevation of ROS, which may intensively affect mitochondrial electron transport chain and mitochondrial permeability transition pore (Belyaeva et al., 2001; Shih et al., 2003). Such a probability is excluded in the present investigation, since ROS generation was observed prior to mitochondrial membrane depolarization. In the present study, significant enhancement of ROS at the earliest time point, causing a collapse of mitochondrial
ψ leading to caspase-3 activation and eventually DNA fragmentation, appeared to be the pathway of Cd induced programmed cell death in murine thymocytes. However, the possibility of an alternate pathway, i.e. the caspase independent pathway, involving mitochondrial release of apoptosis inducing factor (AIF) and endonuclease G (Endo G) also exists, since their release rely mainly on oxidative stress and alteration of intracellular calcium homeostasis. Cadmium has been demonstrated to interfere with calcium by different pathways. It is quite likely, that inhibition of calcium dependent ATPases in nuclei and endoplasmic reticulum (Beyersmann and Hechtenberg, 1997) may alter nuclear calcium homeostasis and markedly increase cytoplasmic free calcium. A sustained rise in cytoplasmic calcium is found to accumulate in mitochondria, causing dissipation of mitochondrial
ψ and oxidative phosphorylation uncoupling (Gregus and Klaassen, 2001; Orrenius et al., 2003). A rapid and continued increase in intracellular calcium followed by Cd induced DNA fragmentation has been reported in mouse thymocytes by Shen et al. (2001). Recently, Lemarie et al. (2004) showed that Cd induced apoptosis in Hep3B cells was mainly by calcium and oxidative stress related impairment of mitochondria, favouring release of AIF and Endo G. It was further added that ROS may alter mitochondrial homeostasis, in part, by decreasing NF-␬B regulated expression of bcl-xL . Further studies, to correlate the GSH and ROS levels in various T subsets to the CD4+ /CD8+ ratio are relevant, to explain altered T-cell immune functions by Cd. In our earlier study (Pathak and Khandelwal, 2006) using murine splenocytes, we found splenocytes to be more susceptible to the toxic effects of Cd, than thymocytes as shown in the present investigation. At 6 h, both 25 and 50 ␮M Cd sufficiently caused splenocyte apoptosis, whereas in thymocytes, it was only the 50 ␮M concentration which exhibited a similar effect. In conclusion, the apoptogenic potential of cadmium has been clearly demonstrated in murine thymocytes

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