Abstract
ABSTRACT. Tubular cell apoptosis contributes to the pathogenesis of renal injury. However, the intracellular pathways that are active in tubular epithelium are poorly understood. The lethal pathways activated by cyclosporin A (CsA), a nephrotoxin that induces caspase-dependent apoptosis in tubular epithelium, were explored. Fas expression, caspase activation, and mitochondrial injury were assessed by Western blot, flow cytometry, and microscopy in cultured murine tubular epithelial cells exposed to CsA. The influence of FasL antagonists, Bax antisense oligodeoxynucleotides, and caspase inhibitors on cell survival was explored. Tubular cells constitutively express FasL. CsA increased the expression of Fas. However, Fas had no role in CsA-induced apoptosis, as CsA did not sensitize to FasL-induced apoptosis, caspase-8 activity was not increased, and neither blocking anti-FasL antibodies nor caspase-8 inhibition prevented CsA-induced apoptosis. Apoptosis induced by CsA is associated with the translocation of Bax to the mitochondria and Bax antisense oligodeoxynucleotides protected from CsA-induced apoptosis. CsA promoted a caspase-independent release of cytochrome c and Smac/Diablo from mitochondria. CsA also led to a caspase-dependent loss of mitochondrial membrane potential. Caspase-2, caspase-3, and caspase-9 were activated, and specific caspase inhibitor prevented apoptosis and increased long-term survival. Evidence for endoplasmic reticulum stress, such as induction of GADD153, was also uncovered. However, endoplasmic reticulum-specific caspase-12 was not activated. CsA induces changes in several apoptotic pathways. However, the main lethal apoptotic pathway in CsA-exposed tubular epithelial cells involves mitochondrial injury.
Nephrotoxicity is the main adverse effect of cyclosporin A (CsA). CsA causes acute renal damage as well as a chronic tubulointerstitial nephropathy characterized by tubular atrophy, loss of tubular cells, and interstitial fibrosis (reviewed in (1,2⇓)). The mechanisms of chronic CsA nephrotoxicity seem to be multifactorial. Among the possible mechanisms, there is evidence for a direct tubular toxicity of CsA (3). An increased rate of tubular cell apoptosis was observed in human renal biopsy specimens obtained from patients with CsA nephrotoxicity (4). In addition, several independent groups have shown that CsA induces apoptosis in tubular cells in a dose- and time-dependent manner (5–7⇓⇓). Apoptosis is an active mode of cell death that promotes cell loss during both acute and chronic renal damage (8). Apoptosis involves the activation of an intracellular death program. This provides the opportunity for therapeutic intervention (8). Two main pathways for apoptosis have been defined: the extrinsic pathway that results from activation of death receptors and an intrinsic pathway that may result from mitochondrial or endoplasmic reticulum (ER) stress. Engagement of death receptors, such as Fas, leads to activation of caspase-8 and subsequent apoptosis (9). In this regard, CsA increases Fas expression in cultured tubular cells, and increased FasL and Fas expression has been reported in chronic CsA nephrotoxicity (6,10⇓). Both ligand-dependent and -independent Fas activation have been implicated in drug cytotoxicity (11,12⇓). Mitochondrial injury leads to the release of apoptosis mediators, such as cytochrome c and Smac/Diablo, and to the loss of mitochondrial transmembrane potential (13). Release of cytochrome c facilitates caspase-9 activation, subsequent activation of effector caspases, such as caspase-3, and apoptosis. More recently, ER stress has been defined as an activator of apoptosis (14). Apoptosis induced by ER stress is characterized by induction of the transcription factor GADD153 and by activation of caspase-12 (14,15⇓). We have studied the intracellular mechanisms of CsA-induced apoptosis in murine renal tubular epithelial cells and the modulatory effect of Fas system antagonism, different inhibitors of caspases, and Bax antisense oligodeoxynucleotides.
Materials and Methods
Cells
Murine proximal tubular epithelial MCT cells were cultured in RPMI 1640 (Life Technologies, Grand Island, NY), 10% decomplemented FCS, 2 mM glutamine, 100 U/ml penicillin, and 100 μg/ml streptomycin in 5% CO2 at 37°C (16). MCT cells have been characterized extensively (17). CsA (Novartis, Barcelona, Spain) was dissolved in ethanol. Final concentration of ethanol was 0.1%, and it did not influence MCT cell apoptosis. CsA-induced apoptosis was already noted at concentrations similar to those found in the blood of patients treated with CsA (1 μg/ml, 0.83 mM), and it increased with dose and time (5). To study the mechanisms of CsA-induced apoptosis, a dose (10 μg/ml, 8.3 mM) that induced apoptosis in a significant percentage of cells in 24 h was chosen. Staurosporine and tunicamycin (both from Sigma, St. Louis, MO) were used at the concentration of 100 nM and 1 μg/ml, respectively.
Studies of Cell Death and Apoptosis
For quantification of cell death and apoptosis, 10,000 cells were seeded in 24-well plates (Costar, Cambridge, MA) in 10% FCS RPMI overnight. Thereafter, they were rested in serum-free medium for 24 h, and CsA or vehicle (ethanol, final concentration 0.1%) was added to subconfluent cells (5). The following caspase inhibitory peptides were used: the caspase-3 inhibitor Z-Asp(OMe)-Glu(OMe)-Val-DL-Asp(OMe)-fluoromethylketone (DEVD-fmk), the pancaspase inhibitor Z-Val-Ala-DL-Asp-fluoromethylketone (zVAD-fmk; Bachem, Bubendorf, Switzerland), the caspase-8 inhibitor Z-Ile-Glu(OMe)-Thr-Asp(OMe)-fluoromethylketone (IETD-fmk), the caspase-9 inhibitor Z-Leu-Glu(OMe)-His-Asp(OMe)-fluoromethylketone (LEHD-fmk), and the irreversible caspase-2 inhibitor benzyloxycarbonyl-Val-Asp-Val-Ala-Asp-fluoromethylketone (z-VDVAD-fmk; Calbiochem, San Diego, CA). The caspase inhibitory peptides were used at concentrations previously shown to protect from apoptosis in other cell systems. DEVD-fmk, zVAD-fmk, IETD-fmk, and LEHD-fmk were dissolved in DMSO. Final concentration of DMSO was 0.05%, and it did not influence MCT cell apoptosis. Peptides (200 μM) or vehicle was added to the cell cultures 3 h before addition of CsA. Cells were cultured in the presence of stimuli for an additional 24 h.
Apoptosis was characterized by morphologic and functional criteria. Nuclei of formalin-fixed cells were stained with propidium iodide in the presence of RNAse A to observe the typical morphologic changes, as described previously (18). For assessment of apoptosis by flow cytometry, adherent cells were pooled with spontaneously detached cells and stained in 100 μg/ml propidium iodide, 0.05% NP-40, and 10 μg/ml RNAse A in PBS and incubated at 4°C for >1 h. This assay permeabilizes the cells; thus, it is not based on the known ability of propidium iodide to enter dead cells. The percentage of apoptotic cells with decreased DNA staining (A0) was counted as described previously (18).
Neutralizing anti-FasL antibodies (clone MFL3; Pharmingen, San Diego, CA) were used at a concentration of 10 μg/ml (18). Recombinant human FasL (Alexis Biochemicals, Montreal, Canada) was used in the presence of a 10-fold excess of a cross-linking antibody, which by itself was devoid of lethal activity. Cross-linking of FasL restores the biologic activity of soluble FasL and simulates its presence on the cell membrane (18). Human FasL activates the murine Fas receptor (18). FasL 100 ng/ml was added with CsA, and cells were incubated for 24 h.
The long-term protective effect of caspase inhibitors was assessed in colony-forming assays. Cells were plated on six-well plates; exposed to CsA and caspase inhibitor or vehicle for 24 h; and then were trypsinized, washed, and seeded in Petri dishes in the presence of complete medium with 10% FCS to allow for their recovery. The number of colonies was estimated at 7 d by crystal violet staining.
Western Blot
Cell samples were homogenized in lysis buffer (50 mM Tris-HCl, 150 mM NaCl, 2 mM EDTA, 2 mM EGTA, 0.2% Triton X-100, 0.3% NP-40, 0.1 mM PMSF, and 1 μg/ml pepstatin A) then separated by 12% SDS-PAGE under reducing conditions. After electrophoresis, samples were transferred to polyvinylidene fluoride membranes (Millipore Corp., Bedford, MA), blocked with 5% skim milk in PBS/0.5% (vol/vol) Tween 20 for 1 h, washed with PBS/Tween, and incubated with rabbit polyclonal anti-Fas (1:500; Santa Cruz Biotechnology, Santa Cruz, CA), rabbit polyclonal anti-FasL (1:500; Santa Cruz Biotechnology), rabbit polyclonal anti-caspase-9 (1:1000; Cell Signaling, Hertfordshire, UK), rabbit polyclonal anti-caspase-12 (1:1000; Cell Signaling), rabbit polyclonal anti-cleaved caspase-3 (1:1000; Cell Signaling), rabbit polyclonal anti-GADD153 (1:500; Santa Cruz Biotechnology), mouse monoclonal anti-FAS-Associated Death Domain (FADD) (1:500; MBL, Nagoya, Japan), or rabbit polyclonal anti-Bax (1:500; Santa Cruz Biotechnology). Antibodies were diluted in 5% milk PBS/Tween. Blots were washed with PBS/Tween and subsequently incubated with appropriate horseradish peroxidase-conjugated secondary antibody (1:2000; Amersham, Aylesbury, UK). After washing with PBS/Tween, the blots were developed with the chemiluminescence method following the manufacturer’s instructions (Amersham). Blots were then probed with mouse monoclonal anti-tubulin antibody (1:2000; Sigma), and levels of expression were corrected for minor differences in loading (18).
Flow Cytometry Analysis of Fas Expression
For cytofluorography, cells were cultured in the presence of control medium or CsA. After washing the culture with PBS, adherent cells were detached with 2.2 mM EDTA and 0.2% BSA in PBS (18). Single-cell (5 × 105) suspensions were incubated in PBS/BSA for 30 min at 4°C with 20 μg/ml Jo-2 or 20 μg/ml of a control Ig. FITC hamster IgG (diluted 1:100) was used as a secondary antibody (all from Pharmingen). For analysis, dead cells and debris were excluded by selective gating based on anterior and right-angle scatter. At least 10,000 events were collected from each sample, and data were displayed on a logarithmic scale of increasing green-fluorescence intensity. Mean cell fluorescence was calculated using LYSIS II software.
Caspase Activity Assay
Caspase-3 and caspase-2 activity (MBL) and caspase-8 activity (Sigma) were measured following the manufacturer’s instructions. In brief, cell extracts (70 μg of protein) were incubated in half-area 96-well plates at 37°C with 200 μM DEVD-pNA peptide or Ac-IETD-pNA peptide in a total volume of 100 μl. The pNA light emission was quantified using a spectrophotometer plate reader at 405 nm. Comparison of the absorbance of pNA from an apoptotic sample with an uninduced control allows determination of the fold increase in caspase activity.
Examination of Mitochondrial Transmembrane Potential
Changes in mitochondrial transmembrane potential (ΔΨm) were determined by staining the cells with JC-1 (Molecular Probes, Leiden, The Netherlands) before flow cytometry analysis (19). Data analysis was performed using Lysis II software by measuring both the green (530 ± 15 nm) and the red (585 ± 21 nm) JC-1 fluorescence. The loss in ΔΨm is seen as a shift to lower JC-1 red fluorescence accompanied by an increase in JC-1 green fluorescence. At least 10,000 events were collected per sample.
Assay of Cytochrome c Release from Mitochondria
Release of cytochrome c from mitochondria to cytosol was measured by Western blot. Cells (5 × 106) were harvested, washed once with ice-cold PBS, and gently lysed for 6 min in ice with 100 μl of lysis buffer (250 mM sucrose, 80 mM KCl, 500 μg/ml digitonin, 1 mM dithiothreitol, 0.1 mM PMSF, and protease inhibitors, in PBS). Lysates were centrifuged at 12,000 × g at 4°C for 5 min to obtain the supernatants (cytosolic extract free of mitochondria) and the pellets (fraction that contains the mitochondria). Supernatants (50 μg) and pellets (50 μg) were electrophoresed on 15% polyacrylamide gels and then analyzed by Western blot as described above. Rabbit polyclonal anti-cytochrome c (1:500; Santa Cruz Biotechnology), rat monoclonal anti-Smac/DIABLO (1:500, Alexis Biochemicals), and rabbit polyclonal anti-Bax (1:500; Santa Cruz Biotechnology) were used. The mitochondrial enzyme cytochrome oxidase subunit IV (1:500; Molecular Probes) is not released from mitochondria during apoptosis and was used as control for the technique.
Immunostaining
Cells were plated onto Labtek slides in RPMI-10%. After 24 h, the medium was changed to RPMI-0%, and then cells were incubated from 1 to 24 h with the indicated stimuli. Then cells were fixed in 4% paraformaldehyde and permeabilized in 0.2% Triton X-10 in PBS for 10 min each. After washing in PBS, cells were incubated overnight at 4°C with rabbit polyclonal anti-cytochrome c (1:100; Santa Cruz Biotechnology) and rabbit polyclonal anti-Bax (1:100; Santa Cruz Biotechnology), followed by 1 h of incubation with appropriate FITC secondary antibody (1:200; Sigma). Cell nuclei were counterstained with DAPI or propidium iodide.
Bax Antisense Oligonucleotide
A phosphothionate-modified antisense oligodeoxynucleotide (ODN) 5′-TGCTCCCCGGACCCGTCCAT-3′ directed against the translation initiation region of mouse Bax mRNA and a control scrambled ODN 5′-TCGTCCGGCCACCGCTCACT-3′, which has little complementarity with Bax mRNA but the same composition as the antisense ODN, were synthesized commercially (Metabion GmbH, Germany) (20). Cells were treated with 10 μg/ml CsA in the presence or absence of Bax antisense ODN (20 μg/ml) or scramble ODN (20 μg/ml) (20).
Statistical Analyses
Results are expressed as mean ± SD. Significance at the P < 0.05 level was assessed by nonparametric Mann-Whitney test for two groups of data and Kruskal-Wallis for three of more groups by means of the SigmaStat statistical software (Jandel, San Rafael, CA).
Results
Upregulation of Fas Is Not Associated with Fas-Related Apoptosis
MCT constitutively express FasL and the Fas receptor (Figure 1). Whereas FasL expression was unchanged (Figure 1A), CsA upregulated Fas expression in a dose-dependent manner (Figure 1B). Flow cytometry confirmed that an increased amount of Fas was indeed exposed in the cell membrane (Figure 1C). Thus, CsA could theoretically lead to autocrine Fas engagement by FasL. However, neutralizing anti-FasL antibodies did not decrease apoptosis induced by CsA, and CsA did not sensitize to death induced by exogenous FasL (Figure 1D). Still, CsA could lead to ligand-independent Fas oligomerization and activation of caspase-8 (12). However, no activation of caspase-8 was detected, and the caspase-8 inhibitor IETD did not prevent CsA-induced apoptosis (Figure 1, E and F). Multiple mechanisms regulate the cell response to the presence of lethal cytokines. FADD is an adaptor protein that plays a critical role in signal transduction from the Fas receptor (reviewed in (9)). CsA decreased FADD expression in a dose-dependent manner (Figure 1G).
Figure 1. Absence of a role for Fas upregulation in cyclosporin A (CsA)-induced apoptosis. (A) MCT cells express FasL (Western blot), and CsA does not modulate FasL expression. (B) CsA upregulates Fas in a dose-dependent manner (Western blot). (C) CsA (10 μg/ml) upregulates cell surface Fas in MCT cells (flow cytometry). (D) Neither recombinant FasL nor anti-FasL blocking antibodies modulate CsA-induced apoptosis. (E) Lack of activation of caspase-8 after incubation with CsA. (F) Inhibition of caspase-8 by IETD did not prevent CsA-induced apoptosis. (G) CsA decreases FAS-Associated Death Domain expression (Western blot). Results were assessed at 24 h. Apoptosis assays are expressed as mean ± SD of four independent experiments. Western blots are representative of three independent experiments.
CsA Induces Bax-Dependent Apoptosis
Bax is a critical mediator of mitochondrial injury in the course of apoptosis. CsA induced Bax translocation to the mitochondria, where it formed aggregates (Figure 2, A and B). Mitotracker co-staining confirmed the mitochondrial localization of Bax (not shown). Bax translocation to the mitochondria was not prevented by caspase inhibitors (Figure 2B). Bax antisense ODN but not scrambled ODN decreased Bax protein expression (Figure 2C) and protected from CsA-induced apoptosis (Figure 2D).
Figure 2. Critical role of Bax in CsA-induced apoptosis. (A) Bax is translocated to the mitochondria, where it forms aggregates (arrow; confocal microscopy: Bax in green propidium iodide in orange). (B) Bax translocation to the mitochondria is accompanied by release of proapoptotic factors, such as cytochrome c and Smac/Diablo, from mitochondria to cytosol in a caspase-independent manner (Western blot at 24 h). Staurosporine (STS) was used as a positive control. (C) Bax antisense oligodeoxynucleotide (ODN) decrease Bax protein expression. (D) Bax antisense ODN prevent CsA-induced apoptosis at 24 h. Apoptosis assays are expressed as mean ± SD of four independent experiments *P < 0.05 versus CsA alone. Western blots are representative of three independent experiments. Magnification, ×40 in A.
Mitochondrial Injury
Exposure to CsA resulted in the release of proapoptotic factors, such as cytochrome c and Smac/Diablo, from the mitochondria (Figures 2B and 3A⇓). Cytochrome c release was noted as early as 6 h after CsA addition. Caspase inhibitors did not prevent cytochrome c release, thus placing this event upstream of caspase activation. CsA also induced mitochondrial injury, as evidenced by loss of mitochondrial transmembrane potential (Figure 3B). However, inhibition of caspases by zVAD prevented the loss of mitochondrial membrane potential (Figure 3B). More specific, caspase-2 seems to be involved in this effect, as inhibition of caspase-2 also prevented the loss of mitochondrial membrane potential (vehicle and CsA/VDVAD: 10% of cells with loss of potential at 24 h versus 27% among CsA-treated cells).
Figure 3. CsA induces mitochondrial injury. (A) Cytochrome c is released from mitochondria. When cytochrome c is released from the mitochondria, the labeling becomes diffuse and yields a light blue color (fluorescence microscopy: cytochrome c in green, DAPI in blue). Cells treated with 100 mM STS for 6 h were used as a positive control (inset). (B) Loss of mitochondrial membrane potential is seen as a shift to lower JC-1 red fluorescence, and it is prevented by caspase inhibition with zVAD. Cells were incubated for 24 h. Results are representative of three independent experiments. Magnification, ×40 in A.
Activation of Caspases and Effect of Caspase Inhibitors
Caspases-2, -3, and -9 are activated in a time-dependent manner in the course of CsA-induced apoptosis (Figure 4, A and B). Peak caspase-2 and -9 activation preceded peak caspase-3 activation. Caspase-3 activation was inhibited by the pancaspase inhibitor zVAD and by the caspase-3 inhibitor DEVD. In addition, the caspase-9 inhibitor LEHD and the caspase-2 inhibitor VDVAD inhibited caspase-3 activation, thus placing caspase-2 and -9 activation upstream of caspase-3 activation (Figure 4, C and D). Inhibition of caspase-2, caspase-9, or caspase-3 prevented apoptosis induced by CsA (Figure 4E). In addition, inhibition of caspase-9 or caspase-3 prolonged cell survival in long-term (7 d) assays (Figure 4F).
Figure 4. CsA activates caspases-2, -3, and -9. (A) Incubation with CsA resulted in the appearance of active caspase-9 and -3 fragments (Western blot). Tunicamycin was used as a positive control. (B) Activation of caspases-2 and -3: time course (activity assay). Peak caspase-2 activation precedes peak caspase-3 activation. (C) Cleavage of caspase-3 could be inhibited by using caspase inhibitors (Western blot): active caspase-3 fragments were determined at 24 h. (D) Activation of caspase-3 could be inhibited by treating the cells with inhibitors of caspases before the addition of CsA (activity assay): caspase-3 activity was measured after a 24-h incubation with CsA. (E) Effect of inhibitors of caspases on apoptosis. Apoptosis was studied at 24 h. Mean ± SD of four independent experiments; *P < 0.05 versus CsA alone. (F) Inhibitors of caspases improved prolonged (7 d) cell survival. Results are representative of three independent experiments. ZVAD, pancaspase inhibitor; DEVD, caspase-3 inhibitor; LEHD, caspase-9 inhibitor; VDVAD, caspase-2 inhibitor. Inhibitors of caspases were used at 200 μM.
ER Stress
CsA induced the expression of GADD153, a transcription factor that is a marker of ER stress (Figure 5A). However, ER-specific caspase-12 (14) was not activated (Figure 5B). Tunicamycin, a stimulus that induces ER-mediated apoptosis of renal tubular epithelium (14), resulted in GADD153 induction and activation of caspase-12 (Figure 5).
Figure 5. CsA induces GADD153 expression but does not activate caspase-12. (A) GADD153 expression (Western blot). Tunicamycin was used a positive control of endoplacmic reticulum (ER) stress-mediated cell death. (B) Caspase-12 activation (Western blot). Active caspase-12 fragments are evident in tunicamycin-treated cells but not in CsA-treated cells.
Discussion
Several groups have identified apoptosis as a mechanism of CsA nephrotoxicity, both in vitro and in vivo (4–7,10⇓⇓⇓⇓). In addition, molecular events such as increased expression of Fas, caspase-3 activation, and loss of mitochondrial membrane potential have been reported in cultured tubular epithelial cells exposed to CsA (7). However, the relationship between these events is subject to speculation. Fas receptor activation may lead to loss of mitochondrial transmembrane potential and caspase-3 activation (21). In this regard, results presented in this article provide evidence against a role of Fas in apoptosis induced by CsA in tubular epithelium. Furthermore, these results indicate that Bax-mediated mitochondrial injury is the main apoptotic pathway in CsA-induced tubular cell toxicity. In addition, they provide evidence against a role of ER stress in CsA cytotoxicity.
CsA increases Fas expression in cultured tubular cells (6), and increased FasL and Fas expression have been reported in chronic CsA nephrotoxicity (10). We have confirmed that renal tubular cells express FasL constitutively (18). CsA upregulated Fas in tubular cells, with no changes in FasL expression. This raises the potential of autocrine activation of Fas or Fas activation by FasL present in the renal milieu (10). However, CsA did not sensitize to FasL-induced apoptosis, caspase-8 activity was not increased, and neither blocking anti-FasL antibodies nor caspase-8 inhibition prevented CsA-induced apoptosis. Taken together, these data strongly argue against a role for Fas in CsA-induced tubular cell apoptosis. In this regard, there are other drugs that upregulate Fas but do not result in Fas-mediated lethality (22). In contrast, Fas upregulation as a result of cytokine stimulation of tubular epithelial cells does result in enhanced sensitivity to FasL-mediated apoptosis (18). These observations are in accordance with the fact that for Fas upregulation to increase cell sensitivity to Fas-mediated apoptosis, the intracellular pathways for apoptosis should be functional (23). The first step in the Fas signal transduction pathway is recruitment of FADD. In MCT cells, CsA decreased FADD expression. This or changes in the expression or activity of other proteins implicated in Fas signal transduction may underlie the observation that Fas is not required for CsA cytotoxicity in tubular epithelial cells.
The inhibition of CsA-induced apoptosis by the broad-spectrum caspase inhibitor zVAD and by the effector caspase inhibitor DEVD provided a functional demonstration of the involvement of caspases (5). Advances in the regulation of apoptosis have identified several potential pathways for activation of the caspase cascade (24). However, no study had previously addressed which activator caspases are involved in CsA-induced apoptosis. Data presented in this article indicate the involvement of caspase-9 and caspase-2 but not of caspase-8 or -12 in CsA-induced apoptosis.
Caspase-9 is the apical caspase of apoptosis resulting from mitochondrial injury (24). Indeed, in renal tubular epithelium, CsA induces Bax aggregation and translocation of Bax to mitochondria, as well as evidence of mitochondrial injury that includes release of cytochrome c and Smac/Diablo and loss of mitochondrial membrane potential. For certain stimuli, such as death receptor stimulation or certain chemotherapeutic agents, cytochrome c release from mitochondria may occur secondary to caspase activation (25,26⇓). However, in CsA-treated tubular epithelial cells, Bax translocation and cytochrome c release were caspase-independent phenomena, placing them upstream of or parallel to caspase activation. In the mitochondria, Bax promotes the release of cytochrome c to the cytoplasm, where it contributes to the formation of the apoptosome, which leads to the activation of caspase-9 (24). Activated caspase-9, in turn, activates caspase-3. That the appearance of active caspase-9 fragments peaked before peak caspase-3 activity and that caspase-9 inhibitor LEHD prevented both the appearance of the caspase-3 active p17 fragment and the development of caspase-3 activity indicates that caspase-9 is activated upstream of caspase-3 in CsA-induced apoptosis. Both caspase-3 and -9 play a vital role in CsA-induced apoptosis. Indeed, inhibition of caspase-9 or caspase-3 activity prevented features of apoptosis and also increased long-term cell survival. The increase in long-term cell survival indicates that caspase inhibitors rescue both from apoptosis and from other eventual forms of cell death in this model. In this regard, certain forms of apoptotic cell death are not prevented by caspase inhibition; rather, caspase inhibition induces a shift from apoptosis to necrosis (27).
CsA also led to loss of mitochondrial transmembrane potential, another feature of mitochondrial injury. However, this form of mitochondrial injury was prevented by caspase inhibition. This suggests the existence of a positive feedback loop, in which initial mitochondrial injury leads to cytochrome c release, which, in turn, activates caspases that further damage the mitochondria and lead to the loss of mitochondrial transmembrane potential. Alternatively, caspase-2 may initiate an independent pathway of mitochondrial injury (25). In this regard, an irreversible caspase-2 inhibitor prevented both loss of mitochondrial membrane potential and features of apoptosis. The functional involvement of caspase-2, caspase-3, and Bax in CsA-induced tubular apoptosis is reminiscent of the requirement for these mediators in toxin-induced ovarian follicle loss (28). The precise relationship between the loss of mitochondrial transmembrane potential and the release of cytochrome c varies with cell type and apoptotic stimuli (29,30⇓). In this regard, stimuli such as staurosporine in HELA cells induce cytochrome c release from mitochondria, which mediates the activation of caspases, which, in turn, promote loss of mitochondrial transmembrane potential (29). By contrast, mitochondrial depolarization precedes cytochrome c release in death receptor-mediated apoptosis (30).
Recently, ER stress has emerged as an inductor of apoptosis (14,15⇓). ER stress is characterized by induction of GADD153 expression and activation of caspase-12 (14,31⇓). Certain agents, such as tunicamycin, promote ER stress and apoptosis in renal tubular cells. Indeed, caspase-12 knockout mice are protected from tunicamycin-induced acute tubular necrosis (14). The increased expression of GADD153 in CsA-treated cells raised the spectrum of the involvement of the ER. However, caspase-12 was not activated. This strongly argues against ER stress as the initiator of CsA-induced apoptosis.
Cell death pathways may be cell- and stimulus-specific. The present article contributes to the understanding of the apoptotic pathways that are active in tubular cells. CsA promoted a Bax-dependent, caspase-dependent pathway of mitochondrial injury that leads to apoptosis. The apoptotic pathways promoted by CsA are not shared by other lethal stimuli. FasL activates death receptors (18), whereas acetaminophen activates caspase-12 and acetaminophen-induced tubular cell death fails to be inhibited by the caspase-3 inhibitor DEVD (32). The unraveling of the apoptotic pathways activated in the course of tubular cell death induced by different stimuli may provide the basis for the therapeutic targeting of apoptosis in the course of acute or chronic renal injury. In addition, the mechanisms by which some lethal pathways are initially activated but do not progress to the point of inducing cell death should be studied further (Figure 6). This information may also be useful when targeting apoptosis in the treatment of neoplasia.
Figure 6. Summary of pathways. CsA results in early signs of activation of the death receptor and ER pathways for apoptosis. However, the specific caspases involved in these pathways are not activated. The molecular mechanisms that prevent these pathways from progressing should be explored further. In contrast, evidence for the different steps of activation of the mitochondrial pathway is present. Studies with inhibitors support this flow of events. In addition, inhibition of caspase-2 prevents features of mitochondrial injury.
Acknowledgments
This work was supported by grants FISSS 01/0199 and SAF 2003/884, Comunidad de Madrid (08.2/0030/2000), Sociedad Española de Nefrología, Instituto Reina Sofia de Investigaciones Nefrológicas, and EU project QLG1-CT-2002-01215. P.J. was supported by FIS (Instituto de Salud Carlos III). A.S. was supported by Conchita Rábago de Fundación Jiménez Díaz. C.L. was supported by Ministerio Español de Educación, Cultura y Deporte.
Part of this work was presented in abstract form at the 2002 Meeting of the American Society of Nephrology, Philadelphia, October 2002.
We thank Mar Gonzalez García-Parreño for technical help with confocal microscopy.
- © 2003 American Society of Nephrology
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