Departments of * Pharmaceutical Sciences, Molecular and Cell Biology, and
Chemical Engineering, University of Connecticut, Storrs, Connecticut 06269
Received November 17, 2003; accepted February 26, 2004
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ABSTRACT |
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Key Words: silica; apoptosis; caspase; lysosome; cathepsin; sphingomyelinase.
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INTRODUCTION |
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Recently, the molecular mechanisms contributing to silica-induced apoptosis have begun to be elucidated. That is, silica-induced apoptosis requires the activation of specific caspases (Iyer and Holian, 1997; Thibodeau et al., 2003
). However, the cellular and biochemical mechanisms underlying this activation remain unclear (McCabe, 2003
). It is important that an in-depth understanding of how silica induces caspase activation and subsequent apoptosis is realized, since these upstream pathways may provide novel targets for therapeutic intervention.
Apoptosis is typically regulated by caspases, which exist as precursor zymogens that are inactive or weakly active until they undergo cleavage after aspartic acid residues at one or more sites (Chang and Yang, 2000). This activation of caspases is governed by a coordinated hierarchy of initiator caspases (e.g., caspases 8 and 9) that activate effector caspases (e.g., caspase 3). Effector caspases are responsible for the biochemical and morphological hallmarks of apoptosis via proteolytic cleavage of homeostatic and structural proteins such as poly (ADP-ribose) polymerase (PARP), lamins, fodrin, and cytokeratin 18 (Cohen, 1997
). The hierarchical cleavage maturation of effector caspases by initiator caspases is induced by distinct intrinsic (intracellular) and extrinsic (extracellular) cellular pathways. Extrinsic signals leading to effector caspase activation are mediated by members of the tumor necrosis factor (TNF) receptor superfamily of plasma membrane death receptors (i.e., TNF receptor 1 and Fas) and the subsequent activation of caspase 8 (Ferri and Kroemer, 2001
). Intrinsic signal transduction pathways are initiated by perturbations of intracellular homeostasis typically affecting the mitochondria, which lead to the activation of caspase 9 (Ferri and Kroemer, 2001
).
The intrinsic pathway of apoptosis is usually driven by alterations in mitochondrial membrane permeability that is measurable as depolarization of the inner mitochondrial membrane transmembrane potential (Castedo et al., 2000). During mitochondrial permeabilization, the intermembranous space can release cytochrome c, which translocates to the cytosol initiating the formation of the apoptosome, a multiprotein complex capable of recruiting and activating caspase 9. Silica-induced effector caspase activation has been demonstrated to require, in part, alterations in mitochondrial integrity. Following silica exposure, cells demonstrate mitochondrial depolarization and the activation of caspase 9 (Thibodeau et al., 2003
). Participation of mitochondria in silica-induced caspase activation is evidenced by the inhibition of caspase 3 activation by either cyclosporin A, an inhibitor of the mitochondrial transition core complex, or by the caspase 9 inhibitor carbobenzoxy-leu-glu-[O-methyl]-his-asp-[O-methyl]-fluoromethylketone (Z-LEHD-FMK; Thibodeau et al., 2003
).
It is widely accepted that perturbations in the mitochondria contribute to apoptosis. However, it is not clear if injury to other cellular organelles (i.e., nucleus, endoplasmic reticulum, golgi apparatus, and lysosomes) can contribute indirectly to caspase activation as well (Ferri and Kroemer, 2001). Much evidence has accumulated to support a role for lysosomal enzymes and lysosomal permeability in apoptosis. Indeed, a role for the acidic compartment of the lysosome is evident in TNF-
induced caspase activation and apoptosis (Monney et al., 1998
). Included in this acidic compartment are cathepsins and acidic sphingomyelinases, the lysosomal enzymes implicated in apoptosis. Cathepsin B contributes to TNF-
induced apoptosis (Guicciardi et al., 2000
), whereas cathepsin D contributes to apoptosis induced by oxidative stress (Kagedal et al., 2001a
), TNF-
(Demoz et al., 2002
), Fas (Deiss et al., 1996
), or free radical injury (Ollinger, 2000
). Lysosomal cathepsin D activity has also been suggested to contribute to silica-induced mitochondrial depolarization and caspase activation (Thibodeau et al., 2003
). Acidic sphingomyelinase is a lysosomal hydrolase that has been suggested to also contribute to apoptosis induced by a variety of stimuli including Fas ligation (Lin et al., 2000
).
In addition to lysosomal enzymes, it appears that lysosomal permeability may be a prerequisite for lysosome-induced apoptosis. Lysosomal permeability has been shown to occur in models of apoptosis induced by TNF- (Werneburg et al., 2002
), Fas (Brunk and Svensson, 1999
), oxidative stress (Antunes et al., 2001
), or sphingosine (Kagedal et al., 2001b
). Although silica can elicit lysosome injury (Nadler and Goldfischer, 1970
), it is unclear if silica can influence lysosome-mediated pathways that contribute to apoptosis. We tested the hypothesis that changes in lysosomal integrity precede silica-induced apoptosis and that apoptosis is, in part, dependent upon the lysosomal acidic compartment and the lysosomal enzymes cathepsin D and acidic sphingomyelinase.
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MATERIALS AND METHODS |
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For neutralization of lysosomal acidity, cells were pretreated for 1 h with 10 mM ammonium chloride (Sigma Chemical Co.) and then concomitantly with silica for 6 h. For inhibition of the aspartic protease cathepsin D, cells were pretreated for 16 h with 1 µM pepstatin A (Sigma Chemical Co.) or DMSO (0.25%) and then concomitantly with silica for 6 h. For inhibition of acidic sphingomyelinase activity, cells were pretreated for 2.5 h with desipramine (50 µM, Sigma Chemical Co.) and then concomitantly with silica. To elucidate the role of oxidative stress and free radical injury, cells were pretreated for 1 h with reduced glutathione (GSH, 1 and 10 mM), N-acetylcysteine (NAC, 1.25 and 5 mM), pyrrolidine dithiocarbamate (PDTC, 0.011 mM), vitamin E (1 µM5 mM), or Nw-nitro-L-arginine methyl ester (L-NAME, 50 µM1 mM; Sigma Chemical Co.), and then concomitantly with silica for 6 h.
Aluminum lactate treatment of silica particles. In some experiments, -quartz silica was pretreated with aluminum lactate to blunt surface active sites and decrease cytotoxicity (Fubini and Hubbard, 2003
). Treatment with aluminum lactate was performed as previously described (Begin et al., 1986
). Briefly, silica was suspended (5 mg/ml) in either sterile 1% aluminum lactate (Aldrich, Milwaukee, WI) or deionized water for 12 h at room temperature with continuous rocking. Following overnight treatment, the particles were washed once in sterile deionized water by centrifugation and then washed twice in RPMI 1640 by centrifugation. After washing, the particles were resuspended in RPMI 1640 for exposure to cells. For scanning electron microscopy (SEM), particles treated with either aluminum lactate or deionized water were mixed with ethanol to reduce surface tension and then placed on a microscope slide. An SEM stub with double-sided carbon tape was then pressed into the sample, and the resulting stub coated with Au-Pd in an evaporative sputter coater. Samples were examined at 20 kV under the environmental scanning electron microscope (Philips ESEM 2020; Eindhoven, The Netherlands).
Western immunoblot analysis for caspase cleavage. Following treatments, the adherent and floating cells were collected and cell extracts were prepared in phosphate-buffered saline (pH 7.6) with 1% Nonidet P-40, 0.5% sodium deoxycholate, and 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 Chemical Co.). Extracts were incubated on ice (1 h), centrifuged at 10,000 x g for 10 min at 4°C, and the supernatants were stored at 80°C. Samples were boiled under reducing conditions, resolved on a 12.5% SDS-polyacrylamide gel, and electrotransferred to nitrocellulose. Blots were blocked in TBS/0.1% Tween 20/5% (w/v) lowfat milk for 1 h at room temperature. Immunodetection of specific caspases was conducted with rabbit anticaspase 3 (Cell Signaling Technology, Beverly, MA) or rabbit anticaspase 9 (Cell Signaling Technology) followed by a peroxidase-conjugated goat antirabbit secondary antibody (Santa Cruz Biotechnology, Inc., Santa Cruz, CA). Normalization of samples was performed by loading similar amounts of protein (extracts equivalent to 3 x 104 cells/µl) and by reprobing the blots with goat antiactin antibody (Santa Cruz Biotechnology, Inc.) followed by peroxidase-conjugated donkey antigoat antibody (Santa Cruz Biotechnology, Inc.). Detection of immunopositive bands was performed using luminol reagents (Santa Cruz Biotechnology, Inc.) and a Kodak Image Station 440CF (Eastman Kodak, Rochester, NY).
Measurement of DNA fragmentation. Following treatments, the 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 phosphate-buffered saline with 0.5 mM EDTA), combined with nonadherent cells, centrifuged for 6 min (150 x g at 4°C), washed, and centrifuged again. Cells were resuspended, fixed, and permeabilized overnight in 70% ethanol at 4°C. Cells were again centrifuged, resuspended in phosphate-buffered saline (pH 7.4) containing 0.1% Triton X-100, 200 µg/ml RNAase A (Sigma Chemical Co.), and 20 µg/ml propidium iodide (Sigma Chemical Co.), and incubated for 30 min at 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 were expressed as a percentage of total gated cells.
Measurement of mitochondrial depolarization. Following treatments, the cells were evaluated by flow cytometry for mitochondrial staining with the lipophilic, cationic dye tetramethylrhodamine ethyl ester (TMRE). The adherent cells were gently detached from tissue culture plates with phosphate-buffered saline (pH 7.4) containing 0.5 mM EDTA, combined with nonadherent cells, centrifuged for 6 min (150 x g at 4°C), and resuspended in PBS. Cells were stained with 200 nM TMRE for 15 min at 37°C in the dark. Positive control cells were stained in the presence of carbonyl cyanide trifluoromethoxyphenylhydrazone (10 µM FCCP, Sigma Chemical Co.), a protonophore causing mitochondrial depolarization in approximately 70% 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). Cells were gated to exclude silica particles and small-sized cell debris (low forward scatter). Cells with decreased red (FL2) fluorescence (mitochondrial depolarization) were expressed as a percentage of total gated cells.
Percentage cytotoxicity (LDH release). Cells were treated with -quartz and evaluated for cell death using the Cytotox 96 Nonradioactive Cytotoxicity Assay (Promega, Madison, WI), according to the manufacturer's instructions. 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 PBS were incubated with the provided LDH substrate for 30 min at room temperature, followed by the 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).
Measurement of reactive oxygen species. Reactive oxygen species (ROS) were measured with dichlorodihydrofluorescein diacetate (H2DCFDA; Molecular Probes, Eugene, OR). H2DCFDA is a cell-permeable probe that diffuses through the cell membrane to be enzymatically trapped when hydrolyzed by intracellular esterases to the nonfluorescent dichlorofluorescein (DCFH). H2DCFDA is colorless and nonfluorescent until both the acetate is hydrolyzed and it is subsequently oxidized to fluorescent dichlorofluorescein in the presence of hydrogen peroxide (H2O2), peroxyl radicals (HOO·), nitric oxide (NO), and peroxynitrate anion (ONOO). Stock H2DCFDA was prepared at 50 mM in DMSO and stored at 20°C in the dark. In a 96-well plate, cells were pretreated with 5 µM H2DCFDA for 0.5 h, then concomitantly treated for up to 6 h with RPMI 1640 with or without either 50 µg/cm2 silica or 300 µM H2O2. Dichlorofluorescein fluorescence was determined at excitation wavelength 485 nm and emission wavelength 530 nm using a Cytofluor 4000 (Perceptive Biosystems, Framingham, MA).
Detection of lysosomal injury. Changes in lysosome permeability were evaluated by the translocation of acridine orange out of lysosomes. The weakly basic dye acridine orange accumulates within acidic vacuolar compartments (lysosomes) due to protonation at low pH (de Duve et al., 1974). At high concentrations within the lysosomes, acridine orange exists in a stacked form that emits red fluorescence when excited by blue light (Bradley and Wolf, 1959
). The intensity of red fluorescence is reflective of the lysosomal concentration (Rundquist et al., 1984
), which decreases in acridine orangeloaded cells upon lysosomal rupture or deprotonation of acridine orange during impairment of the proton gradient (Zdolsek et al., 1990
). In these experiments, cells were preloaded with 0.5 µg/ml acridine orange (Molecular Probes) for 15 min at 37°C, then washed three times with RPMI 1640, followed by exposure to 50 µg/cm2 silica for 1 h. After exposure to silica, the adherent cells were gently detached from tissue culture plates using phosphate-buffered saline (pH 7.4) containing 0.5 mM EDTA and combined with nonadherent cells. Cells were immediately placed on ice and evaluated for red (FL3) fluorescence using a FACSCalibur flow cytometer (Becton Dickinson). Cells were gated to exclude silica particles and small-sized cell debris (low forward scatter). Cells with decreased FL3 fluorescence were expressed as a percentage of total gated cells.
To confirm leakage of lysosomal contents into the cytosol, cells were evaluated for the translocation of FITC-conjugated dextran out of lysosomes. Dextrans localize within minutes to the lysosomal compartment through endocytic mechanisms (Thilo et al., 1995). Cells were preloaded with 1 mg/ml FITC-conjugated 20 kD dextran (Sigma Chemical Co.) for 1.5 h at 37°C, then bathed six times with RPMI 1640, followed by exposure to 50 µg/cm2 silica for 1 or 6 h. After exposure to silica, the cells were evaluated using a Leica SP2 confocal fluorescent microscope (emission 635 nm, excitation 488 nm; Leica Microsystems AG, Wetzlar, Germany).
Differential interference contrast (DIC) microscopy. Cells were plated overnight in Delta T glass-bottom culture dishes (Bioptechs Inc., Butler, PA); 15 min prior to imaging, the media was changed to CO2-independent media (Gibco BRL Life Technologies). The culture dish was then mounted in a temperature-controlled Delta T stage set to 37°C (Bioptechs Inc.) of a Zeiss Axiovert 200 M microscope (Zeiss, Göttingen, Germany). Cells were exposed to silica and imaged by DIC microscopy. Images were captured every 30 s for 6 h using an Orca ER digital camera (Hamamatsu; Bridgewater, NJ) controlled by Openlab software (Improvisions, Inc.; Portage, MI). Images were converted to Quicktime movies (Cupertino, CA) to observe the induction of apoptotic morphology in time lapse.
Statistical analysis. Data are expressed as mean ± standard error. Differences between groups were evaluated by either t-tests or analysis of variance using the Newman-Keuls procedure to correct for multiple comparisons (GraphPad Prism, version 3.03; GraphPad Software, San Diego, CA). A p value of less than 0.05 was considered significant.
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RESULTS |
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DISCUSSION |
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Although changes in lysosomal permeability are often cited as causes of necrotic (oncotic) cell death, data have accumulated suggesting that the loss of lysosomal integrity may contribute to apoptosis as well. Loss in lysosomal integrity has been suggested to contribute to apoptosis induced by oxidative stress (Brunk and Svensson, 1999), sphingosine (Kagedal et al., 2001b
), TNF-
(Werneburg et al., 2002
), Fas (Brunk and Svensson, 1999
), or lysosomal photodamage (Reiners et al., 2002
). We also noted that silica exposure elicited significant changes in the lysosome as measured by the loss of acridine orange retention and FITCdextran translocation. Increases in lysosomal permeability (Figs. 2 and 3) occurred at 1 h, prior to the detection of caspase activation (2 h) and apoptosis (6 h; Thibodeau et al., 2003
). The exact mechanism for the change in lysosomal permeability is unknown, although the change may be the result of particle interaction with lysosomal membranes (Fig. 9).
The consequence of this increased lysosomal permeability may be the release of lysosomal enzymes into the cytoplasm to initiate apoptotic pathways. For example, cathepsins translocate from lysosomes to the cytosol during apoptosis induced by TNF- (Werneburg et al., 2002
) or oxidative stress (Kagedal et al., 2001a
). The translocation of cathepsins is suspected to be an important initiating event of apoptosis (Reiners et al., 2002
; Roberg et al., 1999
). However, the requirement for lysosomal permeability and cathepsin translocation has been questioned in other models. For example, TNF-
induced apoptosis demonstrated a lack of lysosomal permeability and translocation of cathepsin D, despite a requirement for cathepsin D activity during apoptosis (Demoz et al., 2002
). It is evident that not all cells undergoing lysosomal permeability proceed to DNA fragmentation at the same time. The percentage of cells undergoing DNA fragmentation increases with increasing time (Thibodeau et al., 2003
). For example, 25 and 70% of cells undergo DNA fragmentation at 12 and 24 h, respectively (50 µg/cm2).
To characterize candidate lysosomal acidic hydrolases in our model, cells were pretreated with NH4Cl, the cathepsin D inhibitor pepstatin A, or the acidic sphingomyelinase inhibitor desipramine. Alkalinization of lysosomes with NH4Cl can decrease the activity of lysosomal enzymes (Seglen, 1983). NH4Cl dissociates to NH3, which can diffuse into acidic compartments to become protonated as NH4+ (de Duve et al., 1974
; Ohkuma and Poole, 1978
). Proton trapping and limited back diffusion of the protonated base creates a proton leak (de Duve et al., 1974
) that increases vacuolar pH toward neutrality. The normal pH of macrophage lysosomes is about 4.5, but, upon the addition of 10 mM NH4Cl, lysosomal pH can increase to over 6 (Ohkuma and Poole, 1978
). By increasing the pH, the activity of lysosomal acidic enzymes can be selectively decreased (Seglen, 1983
). Our work demonstrates that pretreatment of cells with NH4Cl inhibits silica-induced measurements of apoptosis (Fig. 4). Interestingly, NH4Cl inhibited only the formation of the caspase 9 p37 subunit and not the caspase 9 p39 subunit (Fig. 4A). The p37 subunit of caspase 9 is a product of the mitochondria-derived apoptosome, whereas the p39 subunit is a product of cleavage by caspase 3 (a positive feedback loop). Perhaps exposure of cells to NH4Cl also initiates caspase 3 activation by other pathways (e.g., apoptosome independent).
We also showed that the cathepsin D inhibitor pepstatin A reduces the DNA fragmentation characteristic of apoptosis (Fig. 5C) as well as cleavage activation of caspase 9 (Fig. 5A) and caspase 3 (Fig. 5B). Although cathepsins are not believed to directly activate procaspases (Stoka et al., 2001), a role for both cathepsin B and cathepsin D in apoptosis has been suggested (Guicciardi et al., 2000
; Roberg et al., 1999
). Previously, we demonstrated a role for cathepsin D activity in mitochondrial depolarization and caspase activation induced by silica. These changes were specific to pepstatin A since leupeptin, an inhibitor of cathepsin B, did not elicit these effects (Thibodeau et al., 2003
).
To identify a role for acidic sphingomyelinase activity in our model of apoptosis, cells were pretreated with desipramine, an inhibitor of acidic sphingomyelinase. Desipramine is a cationic amphiphilic drug that selectively decreases the activity of acidic sphingomyelinase without affecting other acidic hydrolases (i.e., acid lipase, arylsulfatases A and B, and hexominidases; Albouz et al., 1981; Hurwitz et al., 1994
). Desipramine reduced silica-induced cleavage activation of caspase 9 (Fig. 6A) as well as cleavage of caspase 3 (Fig. 6B). Desipramine also significantly reduced the percentage of cells undergoing apoptotic DNA fragmentation (Fig. 6C). Acidic sphingomyelinase has been suggested to contribute to apoptosis through the degradation of sphingomyelin and the generation of ceramide, an important mediator of apoptosis (Pettus et al., 2002
).
The mechanisms by which silica may activate lysosomal enzyme pathways or how lysosomal enzymes contribute to silica-induced apoptosis remain unclear. A direct role for these enzymes in caspase cleavage is unlikely since lysosomal lysates fail to activate recombinant caspase 3, caspase 8, or caspase 9 (Stoka et al., 2001). A more direct relationship may exist between acidic sphingomyelinase and apoptosis. Acidic sphingomyelinase can generate ceramide, an intracellular signaling agent capable of signaling apoptosis through mitochondria. Also, acidic sphingomyelinase activity and ceramide generation contribute to the activation of the lysosomal proform of cathepsin D (Heinrich et al., 1999
). It is unclear if acid hydrolases such as cathepsin D act directly on mitochondrial membranes or indirectly via other mediators (e.g., the Bcl-2 family member Bid). Others have suggested that lysosomal enzymes can cleave the full-length, inactive Bcl-2related Bid protein to a proapoptotic truncated protein (tBid), promoting mitochondrial depolarization and cytochrome c release (Heinrich et al., 2004
; Reiners et al., 2002
). However, this latter mechanism does not appear to be pertinent to our model. Full-length Bid was not cleaved to proapoptotic tBid despite decreases in the expression of the proform (Western immunoblot analysis, data not shown). This confirms work by Shukla et al. (2003)
in which no apparent tBid was observed in primary rat pleural mesothelial cells cultured with another inorganic particle, crocidolite asbestos.
To determine if apoptosis and lysosomal permeability are events that depend on the surface-reactive sites of -quartz silica, we evaluated the effect of aluminum lactate pretreatment of silica on measurements of apoptosis. We found that such pretreatment reduced apoptosis as measured by mitochondrial depolarization (Figs. 7A and 7B), caspase activation (Figs. 7C and 7D), and apoptotic DNA fragmentation (Fig. 7E). Our work confirms and extends reports by others. Studies in vivo demonstrated that aluminum treatment of silica prevented silicotic inflammation (Begin et al., 1986
; Duffin et al., 2001
). Reduction of pulmonary inflammation by aluminum is specific to silica particles, because aluminum did not prevent bacteria-induced inflammation (Brown et al., 1989
). Studies in vitro also suggested that aluminum pretreatment reduced the cytotoxicity of silica, as measured by the release of LDH (data not shown; Schins et al., 2002
). The mechanism by which aluminum reduces silica toxicity is unclear. Aluminum has been suggested to substitute for silicon in the silica framework where it influences surface acidity (Fubini et al., 1995
) and the protonation of surface silanol groups. Furthermore, aluminum reduces the ability of silica to generate hydroxyl radicals in the presence of hydrogen peroxide (Schins et al., 2002
). These effects are believed to alter quartz interaction with cell membranes (Fubini, 1998
).
We also speculated that this pretreatment of particles would suppress lysosomal permeability. Although a moderate decrease was noted (Fig. 9B), lysosomal permeability still occurred early (1 h, Fig. 9) and prior to the detection of caspase activation (2 h) and apoptosis (6 h; Thibodeau et al., 2003).
To explore another pathway contributing to silica-induced apoptosis, we studied the role of ROS. However, ROS were not detected after a 6-h exposure of cells to silica (Fig. 10). Also, administration of the antioxidants GSH, NAC, PDTC, vitamin E, and L-NAME did not inhibit silica-induced apoptotic events (Fig. 10 and data not shown). Rather than ROS, silica-induced apoptosis in our model may be more dependent upon particle interactions with cellular biomolecules due to hydrogen bonding, surface charge, or hydrophilicity/hydrophobicity, characteristics that may potentially be modified by aluminum treatment. Although these data question the importance of ROS and free radicals in our model of apoptosis, others have noted that ROS production was elevated by silica (Deshpande et al., 2002; Porter et al., 2002
) and appeared to influence apoptosis (Shen et al., 2001
) and cell activation (Albrecht et al., 2002
; Shukla et al., 2001
).
This study suggests that lysosomal enzyme activity and the lysosomal acidic compartment contribute to silica-induced apoptosis preceded by a loss in lysosomal membrane integrity (Fig. 11). Additional research is needed to elucidate the cellular mechanisms underlying the contributions of lysosomes to the intrinsic pathway of silica-induced apoptosis.
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SUPPLEMENTARY DATA |
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ACKNOWLEDGMENTS |
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NOTES |
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1 To whom correspondence should be addressed at Department of Pharmaceutical Sciences, University of Connecticut, 372 Fairfield Road, U-2092, Storrs, CT 06269. Fax: (860) 486-4998. E-mail: Andrea.Hubbard{at}uconn.edu.
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