* Departments of Pharmaceutical Sciences and
Molecular and Cell Biology, University of Connecticut, Storrs, Connecticut 06269
Received May 22, 2003; accepted June 18, 2003
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ABSTRACT |
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Key Words: silica; apoptosis; caspase 3; caspase 9; mitochondria; cathepsins.
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INTRODUCTION |
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Caspases exist as relatively inactive precursors or procaspases that are converted into their active forms by proteolytic cleavage at internal aspartic acid residues. All caspases show a high degree of specificity, with an absolute requirement for cleavage after an aspartic acid residue and a recognition sequence of at least four amino acids N-terminal to the cleavage site. This specificity is not only important in the cleavage of pro-caspases to their active form, but has been exploited in the design of highly specific and effective enzyme inhibitors (Grutter, 2000).
The mammalian caspase family contains 14 members, a subset of which participates in apoptosis, with the remainder involved in the processing of proinflammatory cytokines (Chang and Yang, 2000). Death-inducing caspases interact in a coordinated cascade to propagate cell signaling pathways leading to eventual apoptosis. Effector caspases 3, 6, and 7 are responsible for cleaving structural elements, nuclear proteins, and signaling proteins (Krammer, 1999
). Nuclear condensation is apparent with fragmentation of the nucleus and DNA and formation of apoptotic bodies. Caspase 8, coupled extrinsically to death receptors, and caspase 9, regulated intrinsically by mitochondrial dysfunction, activate caspase 3 to initiate apoptotic events.
The participation of caspase activation in silica-induced apoptosis and inflammation has also been investigated both in vitro and in vivo. Exposure in vitro of mouse macrophages (Sarih et al., 1993) or human alveolar macrophages (AMs) (Iyer et al., 1996
) has been reported to elicit apoptosis (Sarih et al., 1993
). In human AMs treated with a nonspecific (pan) caspase inhibitor, carbobenzoxy-valyl-alanyl-aspartyl-[O-methyl]-fluoromethylketone (Z-VAD-FMK), there was a decrease in silica-induced apoptosis (Iyer et al., 1996
). These authors later confirmed the involvement of caspase 3 in silica-induced apoptosis in human AM using a specific inhibitor of caspase 3, carbobenzoxy-asp-glu-val-asp-[O-methyl]-fluoromethylketone (Z-DEVD-FMK) (Iyer and Holian, 1997
). Silica also stimulated activation of caspases 1, 3, and 6 in a mouse alveolar macrophage cell line, MH-S (Chao et al., 2001
) and caspases 3 and 9-like activity in rat AMs (Shen et al., 2001
).
The intrinsic pathway of apoptosis is usually driven by alterations in inner and/or outer mitochondrial membrane permeability, which can be measured by release of intermembranous proteins (i.e., cytochrome c) or depolarization of the inner mitochondrial membrane transmembrane potential (Castedo et al., 2000). Frequently preceding the release of cytochome c is mitochondrial membrane permeabilization (MMP), also known as the mitochondrial permeability transition (MPT) (Ferri and Kroemer, 2001
). Changes in MMP can lead to the activation of caspase 9 and the subsequent activation of caspase 3 (Li et al., 1997
; Srinivasula et al., 1998
). Mechanisms that may contribute to MMP vary with apoptotic stimulus, but may include alterations in Bcl-2 proteins, reactive oxygen species, calcium, ceramide metabolites, and more recently, endolysosomal cathepsins. Lysosomal damage can induce cytochrome c release from mitochondria leading to the activation of both caspases 3 and 9 (Reiners et al., 2002
). The lysosomal cysteine protease, cathepsin B, and aspartic protease, cathepsin D, have both been implicated in the induction of apoptosis. In response to oxidative stress, cathepsin D translocation from lysosomes to the cytosol can precede cytochrome c release and loss in MMP (Roberg et al., 1999
). In this model, pepstatin A, a specific inhibitor of cathepsin D, prevented the mitochondrial dysfunction induced by oxidative stress (Roberg et al., 1999
). Similarly, lysosomal cathepsin B can induce the mitochondrial pathway (Guicciardi et al., 2000
). Silica can also elicit lysosomal injury and increase cathepsin D activity (Jajte et al., 1988
; Sjostrand and Rylander, 1984
), which may also contribute to caspase activation.
Despite the significant work accomplished recently in dissecting the relationship between silica-induced caspase activation and cell injury, significant gaps in knowledge remain. It is not clear which caspases are activated or the cellular pathways leading to their activation. Here we address these questions to determine the role and regulation of caspase activation in response to silica.
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MATERIALS AND METHODS |
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Exposures in vitro to silica.
Prior to treatment with particle, cells were plated in 6 or 24 well plates at approximately 2 x 105 cells/cm2 in culture media and allowed to adhere for 2 h, after which culture media was replaced. The next day, cells were stimulated in RPMI 1640 media only with or without -quartz silica (Min-U-Sil 5; Pennsylvania Glass and Sand Corp.; Pittsburgh, PA) or anatase titanium dioxide (TiO2; Sigma Chemical Co.; St. Louis, MO). For inhibition of caspases, cells were pretreated for 1 h with 50 µM Z-VAD-FMK (Enzyme Systems, Inc., Livermore, CA), 50 µM carbobenzoxy-leu-glu-[O-methyl]-his-asp-[O-methyl]-fluoromethylketone (Z-LEHD-FMK) (Enzyme Systems, Inc., Livermore, CA) or DMSO (0.25%) and then concomitantly with silica. For inhibition of mitochondrial permeability transition complex, cells were pretreated for 1 h with 10 µM cyclosporin A (Sigma, St. Louis, MO) or DMSO (0.25%) and then concomitantly with silica. For inhibition of cathepsin B and D, respectively, cells were pretreated for 16 h with 0.1 to 100 µM pepstatin A or leupeptin (Sigma, St. Louis, MO) or DMSO (0.25%) and then concomitantly with silica.
Percent cytotoxicity (lactate dehydrogenase [LDH] release).
Cells were treated with -quartz and evaluated for cell death using the Cytotox 96 Non-Radioactive Cytotoxicity Assay (Promega, Madison, WI) as described by the manufacturer. For inhibition of caspases, cells were pretreated for 1 h with 50 µM Z-VAD-FMK (Enzyme Systems, Inc., Livermore, CA) or DMSO (0.25%) and then concomitantly with silica. Following treatment, cell supernatants and intact cells were separated by centrifugation at 120 x g for 6 min at 4°C. Cell supernatants containing released LDH were saved. Cell pellets (intact cells) were incubated with the provided lysis solution for 45 min at 37°C, followed by extraction of the cell lysates by centrifugation at 120 x g for 6 min at 4°C. Cell supernatants and lysates diluted 1:5 in phosphate-buffered saline (PBS) were incubated with the provided LDH substrate for 30 min at room temperature, followed by addition of the provided stop solution. LDH activity (IU/ml) was calculated after measurement of the OD490nm. Percentage cytotoxicity was calculated as the LDH IU/ml supernatant/(LDH IU/ml supernatant + LDH IU/ml intact cells).
Flow cytometric analysis for caspase activity in vitro.
Following treatment with -quartz, the media was replaced with RPMI-1640 containing a carboxyfluorescein conjugated substrate for caspase-3-like activity (CaspaTag, 7.5 µM FAM-DEVD-FMK; Intergen Co., Purchase, NY). After 1 h incubation at 37°C, nonadherent (floaters) and adherent cells were harvested for flow cytometric analysis. Adherent cells were gently detached from tissue culture plates (using PBS with 0.5 mM EDTA), combined with nonadherent cells, washed, centrifuged at 150 x g 4°C for 5 min, resuspended in wash buffer, and fixed with paraformaldhyde. Cells staining for caspase activity were detected using a FACSCalibur flow cytometer (Becton-Dickinson, San Jose, CA). Cells were gated to exclude silica particles and small sized cell debris (low forward scatter). Cells with increased caspase activity were expressed as a percentage of total gated cells.
Western immunoblot analysis for caspase cleavage in vitro.
After stimulation with either -quartz or TiO2, adherent and floating cells were collected and cell extracts were prepared in PBS pH 7.6 with 1% Nonidet P-40, 0.5% sodium deoxycholate, 0.1% SDS containing freshly added protease inhibitors: 2 mM phenylmethanesulfonyl fluoride (PMSF), 10 µg/ml apoprotinin, 10 µg/ml pepstatin, and 10 µg/ml leupeptin hemisulfate (Sigma, St. Louis, MO). Extracts were incubated on ice (1 h), centrifuged at 10,000 x g for 10 min 4°C, and the supernatants electrophoresed immediately or stored at -80°C. Samples were resolved on a 12.5% or a 15% SDSpolyacrylamide gel and electrotransfered to nitrocellulose. Blots were blocked in TBS/0.1% Tween 20/5% (w/v) low fat milk for 1 h at room temperature (RT). Immunodetection of specific caspases was conducted with rabbit anti-caspase 3 (Cell Signaling Technology; Beverly, MA), rabbit anti-caspase 9 (Cell Signaling Technology; Beverly, MA), or rabbit anti-caspase 8 (Santa Cruz Biotechnology, Inc; Santa Cruz, CA; NeoMarkers Inc., Freemont, CA) and a peroxidase-conjugated goat anti-rabbit secondary antibody (Santa Cruz Biotechnology, Inc; Santa Cruz, CA). Normalization of samples was performed by loading similar amounts of protein (extracts equivalent to 1 to 2 x 104 cells/µl) and by reprobing the blots with goat anti-actin antibody (Santa Cruz Biotechnology, Inc; Santa Cruz, CA) followed by peroxidase-conjugated donkey anti-goat antibody (Santa Cruz Biotechnology, Inc; Santa Cruz, CA). Detection of immunopositive bands was performed using luminol reagents (Santa Cruz Biotechnology, Inc; Santa Cruz, CA) and a Kodak Image Station 440CF.
Measure of apoptosis in vitro.
After stimulation with either silica or TiO2 particles, cells were analyzed for DNA fragmentation into oligonucleosomes by flow cytometry using cell cycle analysis for subdiploid DNA content (Lecoeur, 2002). Adherent cells were gently detached from tissue culture plates (using PBS with 0.5 mM EDTA), combined with nonadherent cells, centrifuged 6 min at 150 x g 4°C, washed, and again centrifuged. Cells were resuspended, fixed, and permeabilized overnight in 70% ethanol at 4°C. Cells were again centrifuged, resuspended in PBS with 0.1% Triton X-100 containing 200 µg/ml RNAase A (Sigma Chemical Co., St. Louis, MO) and 20 µg/ml propidium iodide (Sigma Chemical Co., St. Louis, MO), and incubated 30 min 37°C in the dark. Cell cycle analysis was performed using a FACSCalibur flow cytometer (Becton Dickinson, San Jose, CA). Cells were gated to exclude silica particles and cellular debris having small size (low forward scatter), and the subdiploid cells expressed as a percentage of total gated cells.
Determination of mitochondrial permeabilization.
Following treatment with -quartz, the cells were evaluated by flow cytometry for mitochondrial staining with the lipophilic, cationic dye tetremethylrhodamine ethyl ester (TMRE). The adherent cells were gently detached from tissue culture plates (PBS with 0.5 mM EDTA), combined with nonadherent cells, centrifuged 6 min at 150 x g 4°C, and resuspended in PBS. Cells were stained with 200 nM TMRE and incubated 15 min 37°C in the dark. Positive control cells were stained in the presence of carbonyl cyanide trifluoromethoxyphenylhydrazone (10 µM FCCP; Sigma, St. Louis, MO), a protonophore causing mitochondrial depolarization in approximately 70% of the cells (data not shown). After incubation, cells were immediately placed on ice and evaluated for fluorescence using a using a FACSCalibur flow cytometer (Becton-Dickinson, San Jose, CA). Cells were gated to exclude silica particles and small-sized cell debris (low forward scatter). Cells with decreased FL2 fluorescence (mitochondrial depolarization) were expressed as a percentage of total gated cells.
Statistical analysis.
Data are expressed as mean ± standard error. Differences between groups were evaluated by analysis of variance using the Student-Newman-Keuls procedure to correct for multiple comparisons (GraphPad Prism, v. 3.03). A p value of less than 0.05 was considered significant.
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RESULTS |
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Silica Induces Mitochondral Permeability Transition
Activation of caspase 9 is usually reflective of perturbation of the intrinsic mitochondrial pathway of apoptotic cell death. Figure 6 demonstrates that exposure of MH-S cells at 50 µg/cm2 for either 2 (Figs. 6A
and 6C
) or 6 h (Figs. 6B
and 6D
) significantly increased the percentage of cells with depolarization of the inner mitochondrial transmembrane potential (
m). This change in MMP induced by silica exposure could be partially prevented by 10 µM cyclosporin A (Fig. 7A
), a known inhibitor of the MPT complex. Cyclosporin A (10 µM) also significantly reduced the cleavage maturation of both caspase 9 (Fig. 7B
), and caspase 3 (Fig. 7C
) indicating a role for the MPT complex in the activation of these caspases.
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DISCUSSION |
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Our work also demonstrates that cleavage activation of caspase 9 in MH-S mouse macrophages results in the detection of both p37 and p39 products. Caspase 9 can be alternatively processed to either a p37 subunit or p39 subunit, depending upon whether cleavage occurs at Asp 353 (p37) or Asp 368 (p39) residues. In mice, procaspase 9 (49 kD) is cleaved by the apoptosome at Asp 353 to form p37 (Fujita et al., 1999, 2000
; Little and Mirkes, 2002
; Srinivasula et al., 1998
), which is usually regulated by the release of mitochondrial intermembranous cytochrome c. Active caspase 9 can then cleave the effector caspase 3 (32 kD) into active p20 and p12 subunits (Bratton et al., 2001
). Amplifying the processing of caspase 9 is a feedback loop involving active caspase 3. In mice, caspase 3 can cleave procaspase 9 at the Asp 368 residue forming the active p39 product (Little and Mirkes, 2002
). In our model, silica appears to activate the autocatalytic cleavage of procaspase 9 as well as its cleavage by caspase 3. Production of the p39 cleavage product is specific to silica-treated cells, since in media- or TiO2-treated cells there is no expression of p39.
Our work not only provides new information on silica-induced caspase activation and apoptosis, but also extends previous reports. Chao et al.(2001) detected silica-induced apoptosis in MH-S cells by the cell death ELISA and caspase 1, 3, and 6 activation by cleavage of chromogenic substrates and by Western immunoblot (caspase 3 only). Inducing apoptosis in approximately 15% of macrophages exposed to silica at 50 µg/cm2 for 6 h (Fig. 1
) is comparable to reports in rat AMs of ~8% apoptotic cells after silica exposure at 30 µg/cm2 and ~ 26% apoptotic cells after silica exposure at 100 µg/cm2 silica for 9 h (Wang et al., 2002
). Our work also extends previous studies in which neither TiO2 (Iyer et al., 1996
; Zhang et al., 2002
) nor amorphous silica (Iyer et al., 1996
) elicited apoptotic cell death. We find that TiO2 exposure does not activate either caspase 3 or caspase 9. When human AMs were treated with a pan caspase inhibitor (Z-VAD-FMK), a decrease in both silica-induced apoptosis and IL-1b release was observed (Iyer et al., 1996
). These authors later confirmed the involvement of caspase 3 in silica-induced apoptosis in human AM using a caspase 3 specific inhibitor (Z-DEVD-FMK) (Iyer and Holian, 1997
). Shen et al.(2001)
also noted in rat AMs a role for caspase 3 in silica-induced apoptosis through the use of a caspase 3 inhibitor.
Our work is also the first to demonstrate that processing of caspase 3 and caspase 9 following silica exposure is, in part, due to changes in inner mitochondrial membrane transmembrane potential. The intermembranous mitochondrial space can release cytochrome c, which translocates to the cytosol and complexes with apoptosis protease activating factor-1 (APAF-1) and dATP/ATP (Srinivasula, 1998; Zou, 1999
). Frequently preceding the release of cytochome c is MPT (Ferri and Kroemer, 2001
). MPT is a change in permeability across the inner mitochondrial membrane that allows solutes less than 1.5 kD to pass, leading to depolarization of mitochondria (Bernardi, 1999
; Lemasters et al., 2002
). Our results with TMRE staining of silica-exposed cells indicate induction of mitochondrial depolarization concurrent with the activation of caspase 3 and 9. Current theory presumes the MPT to be a multiprotein permeability transition core complex (PTCC) created at Hackenbrocks contact sites between the inner and outer mitochondrial membranes (Ferri and Kroemer, 2001
; Lemasters et al., 2002
). The PTCC often includes the matrix protein cyclophilin D, making these pores susceptible to inhibition by cyclosporin A. (Ferri and Kroemer, 2001
). MPT can, therefore, be regulated by cyclosporin A (CsA) or independent from the effects of CsA (He and Lemasters, 2002
). In our study, the majority (70%) of depolarized cells exposed to silica are not regulated, whereas a subpopulation (approximately 30%) has regulated mitochondrial depolarization (Fig. 7
). Our work clearly demonstrates that silica exposure initiates MPT (Fig. 6
) that leads to caspase activation (Fig. 7
). In addition, partial prevention of mitochondrial depolarization with either cyclosporin A or pepstatin A reduced both caspase 9 and caspase 3 activation (Figs. 8B
and 8C
). These data would suggest that preventing mitochondrial depolarization would alter silica-induced apoptosis. These results demonstrate the inter relationship between aspartic proteases (e.g., lysosomal cathepsin D), mitochondrial dysfunction, and caspase activation in lung alveolar macrophages exposed to silica particles (Fig. 9
). Other groups have suggested a role for death receptors in the initiation of silica-induced apoptosis (Borges et al., 2001
, 2002
). However, the extrinsic pathway of apoptotic cell death probably does not contribute significantly in our model, because we could not detect cleavage activation of caspase 8 following silica exposure at any time points preceding activation of caspase 3 (data not shown).
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In addition to a role for cathepsin D in mitochondrial dysfunction, reactive oxygen and nitrogen species are well recognized to induce the mitochondrial pathway of apoptosis (Lemasters et al., 2002), and future studies will address this in our model. Shen et al.(2001)
noted in rat AMs, a temporal pattern of events over the first 4 h of silica exposure beginning with reactive oxygen species (ROS) formation, caspase 9 and caspase 3 activation, PARP cleavage, and DNA fragmentation. Studies with other toxic particles also suggest a mitochondrial-mediated pathway of apoptosis. In the human alveolar epithelial cell line A549, asbestos elicited mitochondrial dysfunction, translocation of cytochrome c, and the activation of caspase-9-like activity, changes that did not occur with non-fibrogenic TiO2 or glass beads (Kamp et al., 2002
).
Although silica has been documented to elicit marked pulmonary inflammation and cell death, it appears that silica can also induce apoptotic cell death through changes in mitochondrial membrane integrity and caspase activation driven, in part, by lysosomal cathepsin activity. The biologic significance of silica-induced caspase activation and resulting apoptosis may be in the resolution of these inflammatory lesions through the elimination of damaged or injured cells (Shen et al., 2001) or in a proinflammatory role to attract more alveolar macrophages into the airways (Borges et al., 2001
). Additional studies will be required to better understand the mechanisms of particle-induced apoptosis and its role in cell injury.
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ACKNOWLEDGMENTS |
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NOTES |
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