When hypertonicity is imposed with sufficient intensity and acuteness, cells die. Here we investigated the cellular pathways involved in death using a cell line derived from renal epithelium. We found that hypertonicity rapidly induced activation of an intrinsic cell death pathway—release of cytochrome c and activation of caspase-3 and caspase-9—and an extrinsic pathway—activation of caspase-8. Likewise, a lysosomal pathway of cell death characterized by partial lysosomal rupture and release of cathepsin B from lysosomes to the cytosol was also activated. Relationships among the pathways were examined using specific inhibitors. Caspase inhibitors did not affect cathepsin B release into the cytosol by hypertonicity. In addition, cathepsin B inhibitors and caspase inhibitors did not affect hypertonicity-induced cytochrome c release, suggesting that the three pathways were independently activated. Combined inhibition of caspases and cathepsin B conferred significantly more protection from hypertonicity-induced cell death than inhibition of caspase or cathepsin B alone, indicating that all the three pathways contributed to the hypertonicity-induced cell death. Similar pattern of sensitivity to the inhibitors was observed in two other cell lines derived from renal epithelia. We conclude that multiple cell death pathways are independently activated early in response to lethal hypertonic stress in renal epithelial cells.
- cytochrome c
- cathepsin B
the interstitial fluid of mammalian renal medulla is hypertonic due to hyperosmotic accumulation of NaCl. The NaCl concentration often exceeds 1,000 mosmol/kgH2O in rodents, providing the osmotic gradient for urinary concentration (1). While the renal medullary cells function normally under extreme and variable hypertonicity, cultured cells die when exposed to hypertonicity depending on dose (the intensity of hypertonicity) and acuteness (abrupt switch is more effective than gradual increase). Both apoptotic and necrotic cell death has been observed in hypertonicity-induced cell death from the same renal epithelial cells (25, 32). While the underlying cellular mechanism is not fully understood, there have been several proposals over the years. First is activation of death-inducing plasma membrane receptor such as TNF-α receptor. Both ligand-independent and -dependent mechanisms have been proposed (13, 24). Although TNF receptor activation leads to cell death by way of caspase-8 activation (30), this has not been directly demonstrated in hypertonicity-induced cell death. Second is apoptosis initiated by mitochondrial dysfunction. Genetic studies in yeast have shown that the antioxidant function of mitochondria is required for adaptation to hypertonicity (22). In mammalian renal epithelial cells, mitochondrial dysfunction was observed in correlation with hypertonicity-induced cell death (17). However, cytochrome c release was not observed in these studies, raising questions over the relevance of the mitochondrial dysfunction in hypertonicity-induced cell death. Third is lysosomal release of cathepsin. In certain fibroblasts, cell death in response to hypertonicity is blunted by an inhibitor of cathepsin, suggesting that lysosomal release of cathepsin contributes to cell death (20). While cathepsin release can explain hypertonicity-induced necrosis, lysosomal membrane permeabilization in response to hypertonicity has not been demonstrated.
We are interested in cellular mechanisms involved in the hypertonicity-induced cell death because this knowledge will be essential for our efforts in understanding how the renal medullary cells adapt to extreme local hypertonicity. The goal of this study was to examine cellular pathways involved in hypertonicity-induced cell death in renal epithelial cell lines. We found that multiple cellular pathways of cell death were activated early, i.e., within minutes of exposure to deadly hypertonicity: cathepsin release from lysosome, cytochrome c release from mitochondria, and activation of caspase. Our data demonstrate that each of these pathways contributes to cell death in response to hypertonicity.1
Mouse inner medullary collecting duct cells (mIMCD3) were maintained in DMEM/F-12 supplemented with 10% FBS as described previously (23). Madin-Darby canine kidney (MDCK) cells were maintained in minimum essential medium supplemented with 10% FBS, 0.1 mM nonessential amino acids, and 1 mM sodium pyruvate. Immortalized mouse cortical collecting duct (mpkCCDc14) principal cells were grown in defined medium as described previously (2). Medium osmolality was raised by adding NaCl to various concentrations as indicated and measured using a Vapor pressure osmometer (Wescor, Logan, UT).
Cell viability assay.
To assess cell death in hypertonic conditions, cells were switched to fresh medium containing additional NaCl for the indicated period. In some experiments, cells were pretreated with caspase and/or cathepsin inhibitors for 1 h before hypertonic exposure. Cell viability was determined by measuring metabolic activity using 2,3-Bis(2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide (XTT, Invitrogen, Carlsbad, CA). To assess cell death, culture medium was collected at the end of treatment, and the adhering cells were lysed in culture medium containing 1% Triton X-100. The amount of lactate dehydrogenase (LDH) in culture medium and cell lysates was quantified using the LDH cytotoxicity detection kit (Clontech, Mountain View, CA). The percentage of LDH release was calculated.
Caspase activity assay.
The catalytic activity of the caspases was measured using a colorimetric assay kit (Calbiochem, La Jolla, CA) according to the manufacturer's instructions. Briefly, cells were lysed in caspase lysis buffer (50 mM HEPES, pH 7.4, 100 mM NaCl, 0.1% CHAPS, 0.1 mM EDTA, 1 mM dithiothreitol). The supernatant was collected after centrifugation at 15,000 g for 20 min at 4°C. Caspase-3, -8, and -9 activity was measured by adding one volume of peptide substrate [200 μM acetyl-Asp-Glu-Val-Asp p-nitroanilide (Ac-DEVD-pNA), acetyl-Ile-Glu-Thr-Asp pNA (Ac-IETD-pNA), and acetyl-Leu-Glu-His-Asp pNA (Ac-LEHD-pNA), respectively] in caspase reaction buffer (50 mM HEPES pH 7.4, 100 mM NaCl, 0.1% CHAPS, 0.1 mM EDTA, 10 mM dithiothreitol, 10% glycerol) containing 50 μg of cell lysates at 37°C. The released p-nitroaniline was measured at 405 nm at indicated time intervals with a Spectra Max 340 PC plate reader.
Measurement of cytosolic enzyme activity.
To measure cytosolic enzyme activity, cells were treated with an extraction buffer (250 mM sucrose, 20 mM HEPES, 10 mM KCl, 1.5 mM MgCl2, 1 mM EDTA, 1 mM EGTA, and 1 mM pefabloc, pH 7.5) containing 25 μg/ml digitonin for 15 min on ice. The digitonin concentration and treatment times were optimized to result in the total release of cytosolic LDH activity without disruption of lysosomes as previously described (20). Cathepsin B activity was measured by adding one volume of cytosolic extract in cathepsin reaction buffer (50 mM sodium acetate, 4 mM EDTA, 8 mM DTT, 1 mM pefabloc, pH 6.0) containing 50 μM Z-Arg-Arg-7-amido-4-methylcoumarin hydrochloride (AMC) followed by incubation for 1 h at 37°C. The liberated AMC was measured at 360 nm excitation and 460 nm emission on fluorescence reader. β-N-acetyl-glucosaminidase (NAG) activity was estimated by incubation with three volumes of 0.2 M sodium citrate buffer, pH 4.5, containing 0.3 mg/ml 4-methylumbelliferyl N-acetyl-β-d-glucosaminide for 60 min at 37°C. The liberated methylumbelliferyl was measured by fluorometer with excitation at 356 nm and emission at 444 nm as previously described (15). The amount of protein in each sample was used as an internal standard with which protease activity was normalized.
Detection of cytochrome c release into the cytosol.
Cells were washed with PBS, then treated with an extraction buffer (250 mM sucrose, 20 mM HEPES, 10 mM KCl, 1.5 mM MgCl2, 1 mM EDTA, 1 mM EGTA, and 1 mM pefabloc, pH 7.5) containing 25 μg/ml digitonin for 15 min on ice with constant rotation to extract the cytosol without disrupting the mitochondrial membrane as previously described (10). The supernatant was then collected and centrifuged at 18,000 g at 4°C for 20 min. The supernatants were subjected to 15% SDS-polyacrylamide gels (PAGE).
Western blot analysis.
Cells were washed with PBS and lysed in 50 mM Tris, pH 7.5, 150 mM NaCl, 1 mM EDTA, 1% Triton X-100, 1 mM dithiothreitol, and protease inhibitor for 30 min at 4°C. Lysates were centrifuged for 10 min at 15,000 g to remove insoluble materials. Supernatants were separated on SDS-PAGE and transferred to a polyvinylidene difluoride membrane. After nonspecific binding sites were blocked with 5% nonfat milk at room temperature for 1 h, membranes were incubated with the primary antibodies overnight at 4°C. Immune complexes were detected with horseradish peroxidase-conjugated secondary antibodies, using enhanced chemiluminescence (Amersham Pharmacia, Piscataway, NJ) with exposure to light-sensitive film (Amersham Pharmacia).
Immunocytochemistry and lysosomal staining.
The cells were grown on glass coverslips and fixed with 4% paraformaldehyde in PBS (pH 7.4) for 20 min at 4°C. Cells were permeabilized with 0.3% Triton-X 100 in PBS for 30 min and blocked with PBS containing 3% goat serum and 1% bovine serum albumin for 1 h at room temperature. After incubation with mouse anti-cathepsin B, rabbit anti-Lamp2, mouse anti-cytochrome c, and rabbit anti-cytochrome-c oxidase subunit IV (COX IV) overnight at 4°C, the cells were washed with PBS and treated with goat anti-mouse or anti-rabbit Alexa Fluor 488-conjugated and Alexa Fluor 568-conjugated secondary antibodies for 1 h. Cells were washed with PBS and incubated in 0.1 μg/ml Hoechst for 30 min. After being washed with PBS, coverslips were mounted onto microscope slides. Images were recorded using a Zeiss LSM410 confocal fluorescence microscope. Alternatively, to examine lysosomal membrane integrity, cells were stained with 500 nM LysoTracker Red DND 99 for indicated times at 37°C and then washed several times in PBS to reduce background. The cells were viewed on glass coverslips under a Zeiss LSM410 confocal fluorescence microscope.
All data are expressed as means ± SD. Differences between groups were compared by using unpaired Student's t-test, and P < 0.05 was considered statistically significant.
Hypertonicity induces lysosomal membrane permeabilization and cathepsin B release in mIMCD3 cells.
Previous study has shown that hypertonicity induces cytosolic activation of cathepsin B in immortalized murine embryonic fibroblast (MEF) cells (21). To define the role of cathepsins in hypertonicity-induced cell death, we decided to examine the effect of cathepsin inhibitors on hypertonicity-induced cell death in mIMCD3 cells. To establish experimental conditions, hypertonicity-induced cell death was monitored in hypertonic conditions obtained by addition of NaCl from 150 mM (N150) to 225 mM (N225) using LDH and XTT assays. The osmolality of the basal medium was 300 mosmol/kgH2O. The final osmolality of hypertonic media was 560 (N150), 600 (N175), 650 (N200), and 690 (N225) mosmol/kgH2O. The osmolality in the renal medulla changes gradually in vivo. We and other investigators have found that cells tolerate much greater hyperosmolality when the osmolality is raised gradually compared with when the osmolality is raised abruptly (6, 14). The shift is due to increased expression of protective genes: naïve cells, i.e., cells in isotonic conditions (not stressed), are more sensitive to stress because of lower levels of protective proteins. Since differences in the concentration, rather than the repertoire, of protective proteins are responsible for the differences in sensitivity, underlying mechanisms of cell death appear similar whether the osmotic stress was imposed slowly or gradually. Thus, we employed abrupt changes in osmolality rather than gradual changes for simplicity.
Dramatic cell death occurred between N150 and N200 in 24 h (Fig. 1A). Cell viability had decreased significantly after 3 h of exposure to N225 and declined markedly after 18 h (Fig. 1B). When mIMCD3 cells were preincubated with cathepsin B inhibitors, CA-074 Me and Z-FA-FMK, before exposure to hypertonicity (N225), cell death was significantly reduced (Fig. 1C). However, cathepsin D inhibitor (pepstatin A) or cathepsin L inhibitor (Z-FF-FMK) was without effects. These data indicate that cathepsin B, but not D or L, contributed to cell death.
We next characterized the release of cathepsin B into the cytosol in mIMCD3 cells exposed to hypertonicity. Cytosolic cathepsin B activity and protein levels increased in a dose-dependent manner in response to hypertonicity (Fig. 2A). Cathepsin B release into the cytosol was detected as early as 10 min after exposure to lethal hypertonicity (Fig. 2B). Cytosolic cathepsin B activity continued to increase over the next 6 h. When the activity of a lysosomal enzyme, NAG, was assayed in cytosolic fractions after hypertonicity exposure, the increased activity of NAG was observed 3 h after hypertonicity exposure (Fig. 2B), consistent with lysosomal permeabilization. The cellular redistribution of cathepsin B was further examined using confocal microscopy. Under isotonic conditions, cathepsin B localized with Lamp-2, a lysosome marker, in the perinuclear region (Fig. 3A). After exposure to hypertonicity, cathepsin B displayed a punctuate pattern as early as 10 min, indicating release from the lysosomes. At the same time, LysoTracker fluorescence decreased (Fig. 3B), indicating that cathepsin B release was due to lysosomal permeabilization. Taken together, these results show that lethal hypertonicity induces an early leakage of lysosomal cathepsin B into the cytosol, leading to cell death.
Hypertonicity induces activation of both extrinsic and intrinsic pathways of cell death in mIMCD3 cells.
Since hypertonicity-induced activation of caspase-8 (extrinsic) and caspase-9 and caspase-3 (intrinsic) in MDCK cells has been reported (11), we examined them in mIMCD3 cells. Caspase-8 activity increased robustly in a dose- and time-dependent manner after exposure to lethal hypertonicity (Fig. 4, A and B). Proteolytic cleavage of caspase-8 to the active 43- and 18-kDa fragments was evident from 30 min (Fig. 4C). To confirm the activation of caspase-8, truncation of a caspase-8 substrate, Bid, was examined. The level of truncated Bid increased dramatically in parallel with the cleavage of caspase-8 (Fig. 4C, bottom). These data demonstrate that caspase-8 is activated in response to lethal hypertonicity. Likewise, we observed increased activity (Fig. 4, A and B) in correlation with activating cleavage of caspase-9 and caspase-3 (Fig. 4C). We next examined the effects of caspase inhibitors (Fig. 5). Caspase-8 inhibitor Z-Ile-Glu(OMe)-Thr-Asp(OMe)-FMK (Z-IETD-FMK), caspase-9 inhibitor Z-Leu-Glu(OMe)-His-Asp(OMe)-FMK (Z-LEHD-FMK), or caspase-3 inhibitor Z-Asp(OMe)-Glu(OMe)-Val-Asp(OMe)-FMK (Z-DEVD-FMK) inhibited hypertonicity-induced cell death in a dose-dependent manner. The inhibitory effects were much greater by a pancaspase inhibitor, benzyloxycarbonyl-Val-Ala-Asp (OMe) fluoromethylketone (Z-VAD-FMK), indicating that all of the caspases contributed to cell death.
Hypertonicity induces release of mitochondrial cytochrome c in mIMCD3 cells.
The activation of caspase-9 and caspase-3 described above prompted us to examine cytochrome c as the upstream event (19, 31). We performed subcellular fractionations to obtain cytosolic and mitochondrial fractions. Each fractionation was confirmed using COX IV, an inner mitochondrial membrane protein. Cytochrome c appeared in the cytoplasm in response to hypertonicity in a dose-dependent and time-dependent manner (Fig. 6, A and B). Confocal analysis showed an early and time-dependent release of cytochrome c from mitochondria to the cytosol, followed by accumulation of cytochrome c in the perinuclear areas (Fig. 6C). On the other hand, COX IV staining became progressively punctuate to produce large clumps consistent with mitochondrial membrane permeabilization. Thus, lethal hypertonicity induces an early mitochondrial membrane permeabilization and release of cytochrome c into the cytosol.
Hypertonicity-induced cathepsin B release and cytochrome c release are not causatively linked.
It has been demonstrated that lysosomal cathepsin B is involved in cell death via caspase-dependent and -independent mechanisms (5, 9). We therefore tested whether hypertonicity-induced cathepsin B release was dependent on the activation of caspases. Hypertonicity-induced release of cathepsin B in the cytosol was not inhibited by pretreatment with pancaspase inhibitor Z-VAD-FMK or cathepsin B inhibitor CA-074 Me (Fig. 7), indicating that it was not dependent on caspase or cathepsin B activity. Likewise, cytochrome c release into the cytoplasm was also not inhibited by either inhibitor (Fig. 7). These data show that cathepsin B release and cytochrome c release are not causatively linked in either direction.
Lethal hypertonicity induces parallel and independent pathways of cathepsin B and caspases.
To investigate whether cathepsin B plays a role in the activation of caspases during hypertonicity-induced cell death, activity of caspase-3, caspase-8, and caspase-9 was analyzed with CA-074 Me pretreatment. The activity of these caspases increased after exposure to hypertonicity (Fig. 8A) as demonstrated above (Fig. 4). Pretreatment with CA-074 Me had no effects on the activation of the caspases (Fig. 8A), indicating that cathepsin B activity was not required.
We next tested additive effects of cathepsin B inhibitor and caspase inhibitors on hypertonicity-induced cell death. Pancaspase inhibitor Z-VAD-FMK and cathepsin B inhibitor CA-074 Me conferred significant additive protection (Fig. 8B). Furthermore, combination of caspase-8 inhibitor z-IETD-FMK, caspase-9 inhibitor z-LEHD-FMK, or caspase-3 inhibitor z-DEVD-FMK with CA-074 Me showed an additive protection of hypertonicity-induced cell death. We asked whether caspases and cathepsin B generally contribute to hypertonicity-induced cell death in other kidney cell lines. Hypertonicity-induced cell death was prevented by the pancaspase inhibitor and cathepsin B inhibitor in MDCK cells (Fig. 9), a cell line originated from renal epithelia. Effects of the two inhibitors were additive. On the other hand, in mpkCCDc14 cells, another cell line originated from renal epithelia, cell death was inhibited by the pancaspase inhibitor but not the cathepsin B inhibitor. These data suggest that caspases and cathepsin B individually contribute to hypertonicity-induced cell death in kidney cell lines.
To elucidate the cellular pathways involved in hypertonicity-induced cell death, here we investigated lysosomal and mitochondrial membrane permeability. We find that permeabilization of both lysosomal and mitochondrial membranes is an early event in hypertonicity-induced cell death in mIMCD3 cells. The mitochondrial release of cytochrome c can explain the activation of caspase-9 and caspase-3. The activation of caspase-8 suggests involvement of the extrinsic pathway of cell death. All these pathways are activated early and independent of each other in response to lethal hypertonicity.
Our data suggest that the caspase-independent pathway of cell death in response to hypertonicity involves lysosomal membrane permeabilization associated with cathepsin B activity. Cathepsins D and L do not appear to contribute significantly to this process since their inhibition did not attenuate hypertonicity-induced cell death. Cathepsin B has been reported to contribute to apoptosis via cytochrome c release in some systems, thereby acting upstream of the caspase cascade (3, 4, 9, 27). Other studies have found that cathepsin B can act as an effector protease, downstream of caspases (8, 12). Furthermore, cathepsin B is capable of executing cell death completely independent of the apoptotic machinery in WEHI-S fibrosarcoma cells (8). In our system, cathepsin B is not required for cytochrome c release. In addition, cathepsin B inhibition does not attenuate the induction of caspase-8, caspase-9, and caspase-3 activity by hypertonicity. We also find that caspase activation does not contribute to lysosomal permeabilization. In sum, the lysosomal permeabilization and cathepsin B release occur independently of the caspase activation and cytochrome c release. The mechanism by which hypertonicity induces lysosomal membrane permeability and cathepsin B release remains unknown. Previous study has shown that enhanced reactive oxygen species (ROS) generation precedes lysosomal destabilization and cell death (28). Indirect damage to the lysosomal membrane by ROS is mediated by the intralysosomal accumulation of iron (7). As hypertonicity induces ROS formation (34), similar processes might be involved in the release of cathepsin B in response to lethal hypertonicity.
A recent study reported cytochrome c release in response to hypertonicity by renal medullary interstitial cells (35). Here we find that cytochrome c release is one of the earliest cellular responses to hypertonicity in mIMCD3 cells. Since cytochrome c release is insensitive to caspase inhibition, it does not appear to require caspase activation as reported in many other systems (29). Thus, all of the three pathways—the extrinsic and the intrinsic pathways of caspase activation, and the lysosomal pathways—are independently induced in response to lethal hypertonicity in mIMCD3 cells.
Use of the three pathways of cell death in response to lethal hypertonicity does not appear to be universal. MDCK and mpkCCDc14 cells, both of renal epithelial origin, are protected from hypertonicity-induced cell death by a pancaspase inhibitor. While MDCK cells are protected by a cathepsin B inhibitor, mpkCCDc14 cells are not. On the other hand, MEF cells derived from fibroblasts and RAW264.7 cells derived from monocytes/macrophages display complete insensitivity to either inhibitor (data not shown). It is interesting to note that the cell death pathways are used by three different cell lines originated from renal epithelia in the setting of lethal hypertonicity.
Endogenous inhibitors of hypertonicity-induced cell death have been identified. Heat shock protein 70 (Hsp70) protects WEHI cells from hypertonicity-induced cell death (20). Organic osmolytes attenuate hypertonicity-induced apoptosis by preventing dissipation of mitochondrial membrane potential in renal medullary interstitial cells (35). Prostaglandin E2, the major renal medullary prostanoid, promotes cell survival under hypertonic stress in association with reduced caspase-3 activation, and enhanced expression of Hsp70 and cellular accumulation of organic osmolytes (18).
Cellular accumulation of Hsp70 and organic osmolytes in response to hypertonic stress is promoted by the transcription factor tonicity responsive-enhancer binding protein (TonEBP) (33). TonEBP appears to be a master regulator in protection from hypertonic stress in that it regulates many additional genes that promote survival under lethal hypertonicity (14). On the other hand, the TonEBP-mediated protective pathways require gene expression, and therefore, they come into effect hours after the onset of the three cell death pathways described here. Such action of TonEBP can explain the protection of cells from lethal hypertonicity after pretreatment with moderate hypertonicity that enhances TonEBP expression and activity (6). This is relevant to the hypertonic renal medulla in vivo because it takes many hours to build the high interstitial tonicity in response to maximum vasopressin levels (26).
What might be the clinical relevance of the cell death pathways described here? They should be activated when osmoprotective molecules (the endogenous inhibitors of hypertonicity-induced cell death discussed above) are lacking. Renal medullary injury associated with nonsteroidal anti-inflammatory agents (inhibitors of cyclooxygenases) can be explained by hypertonicity-induced cell death as a result of lower prostaglandin E2 concentrations (18). Severe renal medullary atrophy in TonEBP−/− mice is preceded by massive cell death in young animals as local hypertonicity is building up (16).
This work was supported by the National Research Foundation of Korea (NRF-2011-0020163 and NRF-2012R1A1A2043693).
No conflicts of interest, financial or otherwise, are declared by the author(s).
S.Y.C. and H.M.K. conception and design of the research; S.Y.C., W.L.K., H.H.L., J.H.L., and S.S. performed the experiments; S.Y.C., W.L.K., H.H.L., J.H.L., S.S., and H.M.K. analyzed the data; S.Y.C., W.L.K., S.S., and H.M.K. interpreted the results of the experiments; S.Y.C., W.L.K., H.H.L., J.H.L., and S.S. prepared the figures; S.Y.C. drafted the manuscript; S.Y.C. and H.M.K. edited and revised the manuscript; S.Y.C., W.L.K., H.H.L., J.H.L., S.S., and H.M.K. approved the final version of the manuscript.
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