Organotin-Induced Caspase Activation and Apoptosis in Human

Chem. Res. Toxicol. 2001, 14, 791-798

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Articles Organotin-Induced Caspase Activation and Apoptosis in Human Peripheral Blood Lymphocytes He´le`ne Stridh,*,†,‡ Ian Cotgreave,† Malin Mu¨ller,†,‡ Sten Orrenius,† and Dulceaydee Gigliotti§ Institute of Environmental Medicine, Department of Medicine, Division of Respiratory Medicine, Microbiology and Tumorbiology Centre and Department of Microbiology, Pathology and Immunology, Division of Biomedical Laboratory Technology, Karolinska Institutet, Stockholm, Sweden Received July 19, 2000

In the present study, we show that the immunotoxicant, tributyltin (TBT), induces a dosedependent activation of caspases followed by typical apoptotic morphology in resting human peripheral blood lymphocytes. TBT also caused an early loss of mitochondrial membrane potential (∆Ψm) and release of cytochrome c, suggesting that apoptosis was triggered by the mitochondrial pathway. When CD4+ T-cells were sorted from peripheral blood and exposed to TBT for 30 min, caspase activation and apoptosis were induced. Interestingly, in the sorted CD8+ T-cell population, caspase activation was not observed until 2 h of TBT exposure, suggesting that these cells were more resistant toward TBT. Moreover, a time-dependent induction of caspase activity was also detected in CD3-stimulated peripheral blood lymphocytes. This caspase activation was not associated with cytochrome c release or loss of mitochondrial ∆Ψ and did not lead to apoptotic morphology, although it did lead to both PARP and DFF cleavage. We also noticed a concomitant induction of Hsp27, and it awaits to be seen if this chaperone may interfere with the processing of nuclear protein substrates downstream from these primary caspase-3 substrates. Moreover, no increase in caspase activation or induction of apoptosis was observed after TBT treatment in these cells. Instead, the cells were directed toward necrotic deletion. Taken together, these data suggest that TBT-induced deletion of peripheral lymphocytes is likely to be a component in the overall risk for immunotoxic responses in exposed humans.

Introduction Organotin compounds are important organometallic chemicals, which are used in a variety of technical applications. Of the trialkyltins, particularly tributyltin (TBT)1 is applied as an active component in wood preservatives, disinfectants, and antifoulant paints (1, 2). Consequently, these compounds are now present in the environment, exerting cytotoxic effects to a broad range of species including mammals (3, 4). Several in vivo studies have shown that low doses of TBT target the * To whom correspondence and should be addressed. Phone: +46 8 5177 6183. Fax: +46 8 5177 54 51. E-mail: [email protected]. † Institute of Environmental Medicine. ‡ Department of Medicine, Division of Respiratory Medicine. § Microbiology and Tumorbiology Centre and Department of Microbiology, Pathology and Immunology, Division of Biomedical Laboratory Technology. 1 Abbreviations: huPBL, human peripheral blood lymphocytes; TBT, tributyltin; TPT, triphenyltin; DBT, dibutyltin; DEVD-AMC, Ac-AspGlu-Val-Asp-(7-amino-4-methylcoumarin); MPT, mitochondrial permeability transition; (∆Ψm) mitochondrial membrane potential; NP-40, octylphenoxy polyethoxy ethanol; CHAPS, (3-cholamidopropyldimethylammonio)-1-propane sulfonate; TMRE, tetramethylrhodamine ethylester; FACS, fluorescence activated cell sorter; G3PDH; glyceraldehyde3-phosphate -dehydrogenase; PARP, poly (ADP-ribose)polymerase; DFF/ICAD, DNA fragmentation factor/inhibitor of caspase-activated inhibitor of caspase-activated DNase.

mammalian immune system, thereby causing immunosuppression by an elimination of cortical thymocytes (5-9). The toxicity of the organotins has been previously studied in experimental animal systems. The di- and the tributyltin compounds have been shown to cause atrophy both of the thymus and of the thymus-dependent areas of the spleen and lymph nodes in rats. Thus, the weight and overall morphology of rat lymphoid tissues, as well as peripheral lymphocyte counts and total serum immunoglobulin concentrations, have all been shown to be greatly affected by oral exposure to TBT (8-10). The TBT-induced atrophy of the rat thymus was associated with a selective elimination of cortical thymocytes, resulting in a marked depletion of small-sized, nonproliferating cells (11). Signs of extensive cell destruction within the tissue were not observed. Consequently, the T-cell-dependent immune functions in rats fed with TBT were suppressed. The mechanisms of the immunotoxicity of TBT are still not clear. Some studies suggest that it is the major metabolite of TBT, dibutyltin (DBT),1 which is responsible for the thymic toxicity. The primary effect of DBT is thought to interrupt the maturation of the developing

10.1021/tx000156c CCC: $20.00 © 2001 American Chemical Society Published on Web 06/26/2001

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thymocytes by inhibiting their binding to thymic epithelial cells (7, 12). However, several studies have recently shown that low concentrations of trisubstituted organotins are potent inducers of apoptosis in rat thymocytes (13, 14), as well as in neuronal PC12 cells (15), suggesting apoptosis as a potential mechanism for the thymocyte depletion observed in vivo. Despite the large amount of toxicity data, studies concerning the effect of the organotins in human peripheral lymphocytes and mature T-cells are still lacking. We have previously shown that TBT and triphenyltin (TPT)1 activate the caspases and induce apoptosis in the human leukemia T-cell lines, Jurkat and HUT-78 (16, 17). TBTinduced apoptosis was initiated at the mitochondrial level, by the induction of mitochondrial permeability transition (MPT)1 and release of cytochrome c (16). In the present study, we have investigated the sensitivity of human peripheral blood lymphocytes (huPBL)1 to the cytotoxic effects of TBT. Resting and CD3-activated huPBL from healthy individuals, as well as their sorted CD4+ and CD8+ T-cell subsets, were prepared and exposed to TBT for different time-periods. Differences related to the activation state of the T-cells as well as, between CD4+ and CD8+ T-cell subsets, suggest that complex mechanisms are involved in the regulation of apoptosis in huPBL. The data are discussed in terms of the potential immunotoxic role of TBT in exposed humans.

Materials and Methods Reagents. Tri-n-butyltin chloride was purchased from Merck (Germany), DEVD-AMC from Peptide Institute, Inc. (Japan). All other chemicals were from Sigma Chemicals (St. Louis, MO, USA). Tri-n-butyltin chloride was diluted in DMSO and prepared freshly prior to experiments. Ficoll-Paque (Pharmacia Biotech, Sweden). PhiPhilux-G1D2 (OncoImmunin, Inc., USA), Tetramethylrhodamine ethylester (Molecular Probes, Netherlands). Separation of Mononuclear Cells and Fractionation Techniques. Mononuclear cells were obtained by separation of buffy coats from healthy individuals over Ficoll-Paque gradients. Cells were resuspended in complete medium (RPMI1640 containing 5% FCS, Gibco, BRL, Life Technologies, U.K.) to a concentration of 40 × 106/mL and applied on a nylon wool column, according to standard protocols. Nylon wool-enriched T-cells were triple-stained with monoclonal antibodies to CD4, CD8, and CD3 and directly labeled with Phycoerythrin-C5 (PECy5), Phycoerythrin (PE), and Fluorescein isothiocyanate (FITC), respectively (Dakopatts, Denmark), for further fractionation by use of a cell sorter (FACS Vantage, Becton Dickinson, USA). CD4+ and CD8+ T-cells were sorted for double-staining with CD3-FITC. T-Cell Activation. A total of 2 × 106 nylon wool-enriched T-cells, resupended in complete media, was stimulated with CD3 (3 µg/mL in PBS) (Immunotech Coulter, USA) in precoated 24 well tissue culture plates (Nunc, Denmark) for up to 300 h. T-cell activation was assessed by flow cytometry by double staining for CD69-PE (Immunotech Coulter) and CD3-FITC (Dakopatts, Denmark) and single staining for Fas expression (Fas-FITC) (PharMingen, USA) (not shown). CD69 is an early antigen expressed by activated T-cells. Morphological Evaluation of Cell Death. Cultures were stained with a mixture of chromatin dyes: the membranepermeant dye H-33342 (500 ng/mL) and the membrane-impermeant dye propidium iodide (500 nM) (Molecular Probes, Netherlands). Stained cells were examined by fluorescence microscopy (Wetzlar, Germany). Necrotic cells (damaged plasma membrane and noncondensed nuclei) and apoptotic cells (condensed or fragmented nuclei) were scored manually. At least 200 cells per slide were scored.

Stridh et al. Caspase Activity. Caspase activity (caspase-3 and -7) was measured by the assay of DEVD-AMC1 cleavage, modified from (18). Cells were pelleted and frozen on microtiter plates at 1 × 106 cells/25 µL of PBS. Fifty microliters of buffer (100 mM HEPES, 10% sucrose, 5 mM dithiothreitol, 10-4% octylphenoxy polyethoxy ethanol [NP-40]2 and 0.1% 3-[(3-cholomidopropyl) dimethylammonio] propane-1-sulfonic acid [CHAPS]2, at pH 7.25) was added to each well along with the fluorogenic peptide substrate DEVD-AMC (50 µmol/L). The cleavage of DEVD-AMC was monitored in a Fluoroscan II plate reader (Labsystems, Sweden) using 355 nm excitation and 460 nm emission wavelengths. Fluorescence was measured every 70 s during a 30minute period and the fluorescence units were converted to picomoles of AMC using a standard curve generated with free AMC. Data from duplicate samples were then analyzed by linear regression. Caspase activity in intact cells was measured by using the cell-permeable substrate PhiPhilux-G1D2 (OncoImmunin, Inc.), containing the consensus sequence DEVDG. Resting or CD3-activated lymphocytes were washed and incubated with the substrate (10 µM) for 1 h at 37 °C/5% CO2, according to the manufacture instructions. Fluorescence was measured in the FL1 channel of a flow cytometer (FACScalibur, Becton Dickinson). Release of Cytochrome c. Cytosolic fractions from enriched peripheral T-cells were isolated by a quick cell lysis method with digitonin (19). Under vortexing, lysis buffer (9.4 µg of digitonin/ 106 cells, 500 mM sucrose in PBS: 2 mM NaH2PO4, 16 mM Na2HPO4, 150 mM NaCl, pH 7.6) was quickly added to a cell suspension of 4 × 106 cells in PBS. Heavy organelles and cell debris were pelleted for 60 s at 14000g at 4 °C. The cytosolic fraction (supernatant) was analyzed for cytochrome c by SDSPAGE and immunoblot with a specific anti-cytochrome c antibody (a generous gift from Dr. Ronald Jemmerson, University of Minnesota, Medical School, Minneapolis, MN, USA; 1:2500). Western Blotting. Sample extracts (20 µg/mL) were loaded onto a 15% SDS-polyacrylamide gel and electrophoresed at 130 V for 2 h and transferred to nitrocellulose filters for another 2 h at 100 V. Membranes were blocked overnight in high salt buffer (50 mM Tris base, 500 mM NaCl, 0.05% Tween 20, pH 7.6) containing 1% bovine serum albumin and 5% dried nonfat milk. The membranes were then probed with intermittent stripping with antibodies to cytochrome c (1:2500), DFF, PARP (Biomol Research Laboratories), inducible Hsp70 (Hsp72) and Hsp27 (Stressgen Biotechnology Corp., USA), G3PDH (Trevigen, USA), followed by peroxidase labeled anti-mouse antibodies (1:10.000) and visualized by ECL (Amersham, U.K.).

Results Kinetics of TBT-Induced Caspase Activity in Resting huPBL. HuPBL were incubated with TBT (0.001-10 µM) for up to 8 h and assayed for caspase activity by DEVD-AMC cleavage. TBT exposure induced a dose-dependent induction of DEVDase activity. A TBT concentration as low as 0.1µM induced caspase activity after 6 h of exposure (Figure 1). No caspase activity was observed with TBT concentrations below 0.1 µM during the 8 h time course (data not shown). TBT at 2 µM increased caspase activity up to 30-fold that detected in untreated cells, with appreciable activity arising already after 20 min (Figure 1). On the other hand, no caspase activity was induced with 10 µM TBT, instead there was an associated breach in plasma membrane integrity, indicative of necrosis (Figure 2B). Measurement of TBT-Induced Apoptosis in Resting huPBL. To investigate whether the caspase activity occurred concurrently with other apoptotic changes, huPBL were exposed to TBT and the cells were monitored for plasma membrane blebbing and nuclear fragmentation. Approximately 55% of the cells appeared apoptotic 60 min after treatment with 2 µM of TBT

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Figure 3. TBT-induced cytochrome c release in huPBL. huPBL Were exposed to 2 µM TBT for 5 or 10 min and the cytoplasmic content of cytochrome c was determined by immunoblotting. G3PDH immunoquantitation was performed as internal loading standard. The blot is representative of two similar experiments.

Figure 1. Time course of type II caspase activation in resting human peripheral T lymphocytes (huPBL) treated with TBT. huPBL (106/mL) were treated with TBT (0.1, 4; 0.5 µM, 1; 1 µM, O; 2 µM, b; 10 µM, 9), harvested at various time points after treatment and type II caspase activity of cellular extracts was measured as the ability to cleave DEVD-AMC. Results are presented as mean ( SD (n ) 3) on all points. Displayed results are obtained from one experiment typical of four. Background DEVD-AMC cleavage (2.5 ( 0.4) have been withdrawn from presented data.

Figure 4. Time course of TBT-induced caspase activation in CD4+ and CD8+ subsets of T cells. Nylon-wool-enriched T-cells were sorted into CD4+CD3+ (b) and CD8+CD3+ (O) subpopulations and exposed to 2 µM TBT. Cell extracts were obtained at various time points after treatment and the type II caspase activity was measured by the ability to cleave DEVD-AMC. Background DEVD-AMC cleavage (3.8 ( 0.8) cleavage of both T cell subsets, in absence of TBT, have been withdrawn from presented data.

Figure 2. Time course of TBT-induced cytotoxicity in resting huPBL. huPBL were exposed to TBT (0.1, 4; 2 µM, b; 10 µM, 9), for up to 8 h and the percentages of apoptotic (A) and necrotic (B) of the total cell count, were assessed at different times by staining of the cultures with the fluorescent chromatin dyes H-33342 and propidium iodide.

(Figure 2A). Prolongation of the exposure time did not result in any additional apoptotic cells, instead the apoptotic cells underwent secondary necrosis. After 8 h, no apoptotic cells were observed upon exposure to 0.001 or 0.01 µM of TBT and the cells appeared largely unaffected (not shown). Again, no signs of apoptosis were observed, even after exposing the cells to 10 µM TBT. In this case the cells underwent necrosis (Figure 2B). Two distinct populations of cells could be microscopically distinguished to differ in their sensitivity toward the induction of apoptosis by 2 µM TBT (data not shown). Detection of TBT-Induced Cytochrome c Release in Resting huPBL. Cytosolic extracts were obtained

from TBT-treated huPBL and Western blot analyses of cytochrome c levels were performed. Detection of cytochrome c in the cytosolic fraction preceded the rise in caspase activity by at least 10 min (Figure 3). Thus, release of mitochondrial cytochrome c occurred early enough to be responsible for the caspase activation observed in TBT-exposed huPBL. Effect of TBT on Purified CD4+ and CD8+ T Cell Populations. Initial observations in our experiments revealed morphological heterogeneity in the response of huPBL to TBT, suggesting differences in sensitivity to the organotin compound between the T-cell subsets. Nylon wool-enriched T-cells were FACS sorted into CD4+CD3+ and CD8+CD3+ subpopulations. Both T-cell subsets were exposed to 2 µM of TBT for up to 3 h and assayed for DEVDase activity. The CD4+ T-cells responded in a similar manner as the unsorted population (Figure 1), with caspase activity emerging at 30 min and peaking at 1 h after initial exposure (Figure 4). However, in the CD8+ T-cell population, caspase activation was detected 2 h later than in the CD4+ T-cells, suggesting that the CD8+ T-cells were more resistant to TBTinduced caspase activation and apoptosis (Figure 4). In both situations, the caspase activation was associated with the induction of apoptotic morphology (data not shown).

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Table 1. TBT-Induced Cell Death in CD3 Stimulated PBL-T time of pretreatment with CD3 (h)a time of TBT-exposure (h)b apoptotic cells (%) necrotic cells (%)

0 1 73 ( 5.2 3 ( 0.5

6 3 54 ( 4.1 15 ( 2.0

1 68 ( 3.2 4.2 ( 1.2

80 3 49 ( 3.1 12 ( 1.1

1 0 ( 0.0 5.2 ( 1.2

3 1.2 ( 0.2 51 ( 3.2

120 1 3 22 ( 2.2 35 ( 4.1 6.2 ( 0.6 12 ( 1.1

a PBL-T were pretreated with CD3 (3 µg/mL) for up to 120 h. b At indicated time points, stimulated PBL-T were exposed to TBT (2 µM) for 1 or 3 h and cell death was determined. Apoptosis and necrosis was assayed and defined as described in the Materials and Methods. Results are presented as mean ( SD (n ) 3) on all points. Displalyed results are obtained from one experiment typical of four.

Figure 5. Time course of CD3-induced caspase activation in huPBL. huPBL were stimulated in vitro with CD3 for up to 300 h. At indicated time points, huPBL were harvested before being assayed. Cellular extracts were assayed for their type II caspase activity by the ability to cleave DEVD-AMC at the indicated time points. Results are presented as mean ( SD (n ) 3) on all points. Displayed results are obtained from one experiment typical of four. Background DEVD-AMC cleavage (2.2 ( 0.3) have been withdrawn from presented data.

Effect of CD3 Stimulation on Caspase Activity in huPBL. HuPBL were stimulated in vitro with CD3 for up to 300 h. Cells were harvested at different time-points and assayed for DEVDase activity. A time-dependent and massive induction of caspase activity was observed in the CD3-stimulated cells. Caspase activation was detectd already after 24 h and, by 72 h of stimulation, it had increased 15-fold from that of unstimulated cells (Figure 5). The specific activity of caspase-3 was also assessed in intact cells by using the cell-permeable substrate PhiPhilux, which contains the consensus caspase-3 cleavage sequence DEVDG. Results presented in Figure 6A show that no cleavage of the substrate was observed in resting huPBL. However, when cells were exposed to 2 µM TBT for 30 min, substrate cleavage was detected (Figure 6A). As predicted, PhiPhilux was constitutively

cleaved in huPBL stimulated with CD3 for 69 h (Figure 6B). Despite predominant caspase activation, no apoptotic morphology was observed in these cells. The cells looked round and healthy, with an intact membrane and noncondensed nuclei (Figure 7). As previously shown in Figure 5, this condition seemed to be transient, since in cells cultured for prolonged periods (more than 69 h), caspase activity declined. The caspase activation was shown to clearly follow the expression of the early activation marker CD69 (data not shown). The expression of CD69 by CD3-activated huPBL increased from 9 to 50% between 3 and 6 h after in vitro stimulation, peaked between 22 and 44 h and subsequently declined when analyzed at 69 h. After 69 h of CD3 stimulation in vitro, 83% of the cells expressed Fas (data not shown). TBT-Induced Cell Death in CD3-Stimulated Cells. HuPBL were exposed to 2 µM of TBT at two time points after CD3 stimulation, and DEVDase activity and apoptotic morphology were determined. At the time point when CD3-dependent caspase activation was high (69 h, Figure 5), TBT-exposure did not increase this activity any further (data not shown), neither did the agent induce apoptotic morphology. Instead the cells were directed toward necrotic deletion several hours later (Table 1). However, during earlier (less than 66 h) or later time points (more than 114 h) after CD3 stimulation, when the constitutive caspase activity was less prominent, TBT was able to increase the caspase activity above the control levels (not shown) and induce an apoptotic morphology (Table 1). Effect of TBT on ∆Ψm in Resting and CD-Activated huPBL. We have previously shown that TBT induces mitochondrial permeability transition (MPT), resulting in the loss of ∆Ψm in Jurkat T-cells, in a process thought to be important for the activation of TBT-induced apoptosis (20, 21). To investigate whether TBT-induced cytochrome c release, was associated with a similar loss in ∆Ψm and induction of MPT in huPBL, we employed the

Figure 6. Determination of caspase activity in intact huPBL. huPBL were left untreated (dashed line), treated with 2 µM TBT for 30 min (A), or stimulated with CD3 for 69 h (B). After 1 h of labeling with the cell-permeable substrate PhiPhilux (10 µM), the cells were analyzed by flow cytometry (FL-1).



brane according to ∆Ψm. A loss of ∆Ψm can be monitored by flow cytometry, as a decrease in FL-2, expressed in a log scale. TBT induced a loss in ∆Ψm in resting huPBL, initiated at 7 min of TBT exposure and accomplished after 20 min, i.e., within the time frame of cytochrome c release (Figure 8A and Figure 3). CD3 stimulation of these cells did not affect the constitutive TMRE fluorescence, whereas TBT induced a similar loss of ∆Ψm as observed in resting huPBL. (Figure 8B). Interestingly, when sorted CD4+ and CD8+ T-cells were TMREstained and further treated with 2 µM TBT for 7 and 20 min, a loss in ∆Ψm was observed exclusively in CD4+ T-cells (Figure 9).

Figure 7. Morphological examination of CD3 stimulated huPBL. huPBL were stimulated with CD3 for 69 h and stained with the membrane-permeant dye H-33342 (blue fluorescence) and the membrane-impermeant dye propidium iodide (red fluorescence) for assessment of apoptosis and necrosis, respectively.

fluorescent dye, TMRE. This mitochondrial-specific fluorochrome accumulates in the inner mitochondrial mem-

Effect of CD3 Stimulation of huPBL on the Induction of anti-Apoptotic Proteins and Cleavage of Caspase 3-Dependent Substrates. Heat shock proteins, particularly Hsp27 and Hsp70, have been implicated as inhibitors of apoptosis at various levels in the triggering of caspase-dependent processes (35-37). In an attempt to investigate the mechanism underlying the insensitivity of CD3-stimulated T-cells toward the constitutive activation of apoptosis, we analyzed the expression of Hsp 27 and 70, as well as the classical apoptosis inhibitor Bcl-2, in resting and CD3-stimulated cells. The level of Bcl-2 protein was not altered during the CD3 stimulation, and a constant level of constitutively expressed Hsp70 was seen in both resting and CD3-stimulated cells (Figure 10). However, progressive induction of Hsp 27 was detected upon CD3 stimulation of the cells (Figure 10), which was tightly correlated to the kinetics of caspase activity. Interestingly, however, both PARP and DFF cleavage were detected in CD3stimulated cells after 78 h (Figure 10), despite the lack of apoptotic morphology (Figure 7).

Figure 8. Effect of TBT on ∆Ψm in resting and activated huPBL. huPBL were left untreated (A) or stimulated with CD3 for 69 h (B), before being stained with 5 nM TMRE followed by exposure to 2 µM TBT for 7 and 20 min (solid lines), before being analyzed by flow cytometry (FL-2). Control cells are represented by dashed lines.

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Figure 9. Effect of TBT on ∆Ψm in resting and CD4+ and CD8+ T cells. CD4+ T cells were stained with 5 nM TMRE and treated with 2 µM TBT for 7 min (A) and 20 min (B) (solid lines), before being analyzed by flow cytometry (FL-2). Control cells are represented by dashed lines. CD8+ T cells were identically stained with TMRE, followed by TBT- treatment for 7 min (C) and 20 min (D) (solid lines), before analysis.

Figure 10. Induction of anti-apoptotic proteins in CD3 stimulated huPBL. huPBL were CD3 stimulated and the expression of the apoptosis-associated proteins Hsp70, Hsp27, Bcl-2, PARP, and DFF45/ICAD was determined after 0, 24, 78, and 172 h by immunoblotting. G3PDH immunoquantitation was performed as internal loading standard. The blot is representative of two similar experiments.

Discussion In the present study, we have observed that TBT is highly toxic to huPBL from healthy individuals. The

effects of TBT on caspase activation, and the induction of apoptosis or necrosis will vary, depending on dose, time of exposure and if huPBL are resting or CD3 activated. These effects are also highly dependent on which subset of T-cells is exposed to the immunotoxicant. We have previously shown that Jurkat T-cells exposed to 2 µM of TBT were induced to activate caspases after 50 min (17). Thus, when compared to Jurkat T-cells, resting huPBL were slightly more sensitive to TBTinduction of caspase activation and apoptosis. As a part of the molecular mechanism of pro-apoptotic events, we have shown that TBT-induced caspase activation in huPBL was preceded by an early release of mitochondrial cytochrome c, with similar kinetics to those previously observed in Jurkat T-cells (16). The present data suggest that TBT may target the mitochondria in huPBL, inducing MPT and releasing cytochrome c in a similar manner as in Jurkat T-cells. Tributyltin has also been shown to be a potent inducer of apoptosis in rat thymocytes (13, 14), with similar kinetics as observed in human Jurkat T-cells and huPBL. Thus, it appears that the mitochondrion may also be the primary target for initiation of TBT-induced apoptosis in rat thymocytes, but this awaits experimental proof. Therefore, it now appears that TBT is an extremely potent mitochondrial poison that may target a conserved part of the organelle, thereby exerting cytotoxic effects in immune cells over a broad range of animal species. A feature of an apoptosisresistant phenotype is the expression of anti-apoptotic members of the Bcl-2 family of proteins, such as Bcl-2 and Bcl-xL (22-24). One proposed role for Bcl-2 as an



anti-apoptotic mediator involves the inhibition of cytochrome c release from the mitochondria (24, 25). We have previously shown that Bcl-2 and Bcl-xL had no significant effect in the kinetics and extent of TBT-induced apoptosis in Jurkat cells (20). Indeed, a number of studies have shown that the basal expression of the Bcl-2 protein is higher in the CD8+ subset than in the CD4+ T-cells (26, 27). Thus, increased expression of Bcl-2 could underlie a possible protective mechanism in resting CD8+ T-cells against mitochondrially triggered apoptosis. Selective sensitivities of CD4+ and CD8+ cells toward induction of apoptosis have been reported in response to stimulation of the Fas/FasL apoptotic pathway during different pathological conditions (28, 29). Data addressing other apoptotic pathways than Fas/FasL are, however, inadequate. The reason for the different sensitivities between the CD4+ and CD8+ T-cells toward TBT-induced cytoxicity is, thus, presently obscure. We are tempted to speculate that the difference in TBT-induced apoptosis observed in CD8+ and CD4+ T-cells could be related to the complex, intrinsic death machinery of perforin and granzymes which is present in CD8+ T-cells. A more restricted regulatory mechanism for effector cytotoxic functions could explain the resistance of CD8+ T-cells to caspase activation and apoptosis. In addition to the differential responses of the T-cell subsets to a pro-apoptotic stimulus such as TBT, another important point raised from our results is the importance of the activation state of the T-cells at the time of exposure. T lymphocytes already engaged in an immunological response must proliferate and differentiate into effector cells. The need to control and avoid premature death of cells participating in an immune response is certainly of physiological relevance. Thus, the caspase activation detected in stimulated T-cells might be an intrinsic part of the cellular activation machinery. This condition seems to be transient since caspase activity declined in cells which were cultured for a prolonged period. The mechanism of caspase-3 activation in CD3stimulated huPBLs remains obscure, but our data, demonstrating intact mitochondrial membrane potential and lack of cytosolic cytochrome c, seems to suggest a mechanism exclusive of the involvement of mitochondrial cytochrome c release or apoptosome activation. Previous studies have indicated that caspase activation in human T-cells does not necessarily lead to cell death, arguing for a checkpoint in the apoptotic pathway (3032). Recent data has confirmed caspase activation in nonapoptotic T cells and, moreover, demonstrated that caspases may be required for T cell proliferation in vitro (33, 34). To further define at which level of the apoptotic process such checkpoint(s) might occur we investigated the processing of several caspase-3-dependent substrates associated with nuclear events, as well as the expression of several antiapoptotic proteins, in CD3-stimulated cells. We clearly show that both PARP and DFF45 (Figure 10) were cleaved in association with maximal caspase-3 activity (Figure 5). These data strongly indicate that the apoptotic checkpoint occurs at a level downstream from these important nuclear effector molecules. It is thus not surprising that we detected no correlation with the expression of either Bcl2 or Hsp70, which were both constitutively expressed throughout the activation process and which are known to interfere with caspase activation primarily at the mitochondrial (24, 25) and apoptosomal levels (36), respectively. This again supports

the supposition that mitochondrially driven activation of the apoptosome complex is not involved in CD3-dependent activation of huPBL as the high expression of Bcl2 and Hsp70 would constitute a natural blockade of this process. Additionally, it may be speculated that the treatment of CD3-activated cells at the point of maximal caspase activity with TBT fails to further induce activation of caspase 3, despite cytochrome c release, due to Hsp72-dependent inhibition of apoptosomal recruitment of pro-caspase 9 (36). In contrast, however, we noted a progressive induction of Hsp27 in response to CD3 stimulation of huPBLs (Figure 10). This chaperone has been shown to prevent apoptosis by interaction with cytochrome c release from the mitochondrion (35, 37). However, as the blockade in induction of apoptosis appears localized to the events after the cleavage of DFF45 and PARP, it awaits to be seen if HSP27 is able to interfere with the processing of nuclear proteins downstream of these. In summary, TBT induces apoptosis in human peripheral lymphocytes at similar concentrations to those effective with rat thymocytes and human leukemia T-cell lines, suggesting that TBT might interact with the cells via an evolutionarily conserved mechanism. This mechanism appears to involve release of mitochondrial cytochrome c, resulting in the activation of caspases in huPBL. The CD8+ subset of T cells was more resistant to TBT exposure than the CD4+ subset. Moreover, activated T cells were not prompted to undergo TBTinduced apoptosis, instead, the cells were forced to a necrotic death after a prolonged exposure period. The mechanism involved in the prevention of apoptosis during constitutive expression of caspase 3 during activation and during TBT treatment of the cells appears to lie downstream from the cleavage of DFF45 and PARP and might involve the induction and activity of HSP27. Thus, our data have shown that TBT, at toxicologically relevant exposure concentrations, dictates a definite risk to both resting human peripheral blood cells, by the induction of apoptosis, and to activated T-cells, by the induction of necrosis.

Acknowledgment. We thank Dr. Bengt Fadeel and Prof. Hans Wigzell for critically reading the manuscript. This work was supported by grants from the Swedish Medical Research Council (S.O. and I.C.).

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Organotin-Induced Caspase Activation and Apoptosis in Human

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