Heat stress prevents mitochondrial injury in ATP-depleted renal epithelial cells

F. Li1, H. P. Mao2, K. L. Ruchalski1, Y. H. Wang3, W. Choy1, J. H. Schwartz1, and S. C. Borkan1

1 Renal Section, Department of Medicine, Boston Medical Center, Boston University, Boston 02118-2518; 2 Department of Pathology, Tufts University and New England Medical Center, Boston, Massachusetts 02111-1533; and 3 Department of Nephrology, First Affiliated Hospital, Zhongshan University, GuangZhou, China 510080


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

The events that precipitate cell death and the stress proteins responsible for cytoprotection during ATP depletion remain elusive. We hypothesize that exposure to metabolic inhibitors damages mitochondria, allowing proapoptotic proteins to leak into the cytosol, and suggest that heat stress-induced hsp72 accumulation prevents mitochondrial membrane injury. To test these hypotheses, renal epithelial cells were transiently ATP depleted with sodium cyanide and 2-deoxy-D-glucose in the absence of medium dextrose. Recovery from ATP depletion was associated with the release into the cytosol of cytochrome c and apoptosis-inducing factor (AIF), proapoptotic proteins that localize to the intermitochondrial membrane space. Concomitant with mitochondrial cytochrome c leak, a seven- to eightfold increase in caspase 3 activity was observed. In controls, state III mitochondrial respiration was reduced by 30% after transient exposure to metabolic inhibitors. Prior heat stress preserved mitochondrial ATP production and significantly reduced both cytochrome c release and caspase 3 activation. Despite less cytochrome c release, prior heat stress increased binding between cytochrome c and hsp72. The present study demonstrates that mitochondrial injury accompanies exposure to metabolic inhibitors. By reducing outer mitochondrial membrane injury and by complexing with cytochrome c, hsp72 could inhibit caspase activation and subsequent apoptosis.

hsp72; cytochrome c; caspase 3; apoptosis-inducing factor; mitochondrial membrane potential


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

APOPTOSIS HAS EMERGED as an important cause of cell death in renal epithelial cells subjected to stress. Observations in both in vivo and in vitro models demonstrate that ischemia (38, 47), oxidant stress (16), hypoxia (45), ultraviolet (UV) irradiation (25), urinary tract obstruction (11), glomerulonephritis (58), and kidney transplantation (12) cause apoptosis. Exposure to metabolic inhibitors, an in vitro model of ischemia, also induces apoptosis in renal epithelial cells (29, 45, 53). Prior heat stress sufficient to induce hsp72, a cytoprotectant protein, reduces apoptosis in ATP-depleted renal cells (53). However, the biochemical pathways that mediate ischemia-induced apoptosis and the mechanism(s) of cytoprotection by heat stress proteins (HSPs) have not been clarified (46).

The mitochondrion has recently been implicated as the primary regulator of apoptosis after diverse forms of cell injury (21, 45, 50, 51, 59). Disruption of the inner mitochondrial membrane dissipates the -220 mV proton gradient responsible for the mitochondrial membrane voltage potential (Delta Psi m,) (13, 43, 48, 57). In contrast, changes in the permeability of the outer membrane release proteins that are normally restricted to the intermitochondrial membrane space into the cytosol that can cause apoptosis (2, 24, 49). These two events may be independent of one another (5). Proapoptotic mitochondrial proteins in the intermembranous space include (but are not limited to) cytochrome c, several caspases, and apoptosis-inducing factor (AIF) (14, 24, 46, 50). On release of mitochondrial cytochrome c into the cytosol, the "apoptosome" is formed. This multimeric complex consists of procaspase 3, caspase 9, apoptosis-activating factor (APAF)-1, and cytochrome c (1, 4). The apoptosome activates caspase 9, initiating the self-amplifying apoptotic enzyme cascade (1). In contrast to cytochrome c, mitochondrial AIF migrates to the nucleus, causing DNA fragmentation and partial chromatin condensation (14, 49).

In the present study, we tested the hypothesis that exposure to cyanide and 2-deoxy-D-glucose in the absence of exogenous dextrose induces apoptosis in renal epithelial cells by injuring the outer mitochondrial membrane. Furthermore, we evaluated the possibility that the induction of HSPs, including hsp72, decreases apoptosis by ameliorating outer mitochondrial membrane injury and inhibiting subsequent caspase activation. The present studies show that inhibition of electron transport causes a rapid loss of mitochondrial membrane potential associated with the redistribution of both mitochondrial cytochrome c and AIF to the cytosolic compartment. Loss of mitochondrial cytochrome c and AIF is accompanied by a marked fall in maximal mitochondrial ATP production and activation of caspase 3. In contrast, the accumulation of hsp72 is associated with preserved mitochondrial function, a decrement in cytochrome c release after exposure to metabolic inhibitors, and significantly less activation of caspase 3. We describe for the first time an interaction between cytochrome c and hsp72 and suggest two mechanisms by which hsp72 might inhibit apoptosis.


    METHODS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Materials. All reagents were obtained from Sigma Chemical (St. Louis, MO) unless otherwise indicated.

Cell culture. Renal epithelial cells from the opossum kidney (OK) were obtained from the American Type Culture Collection (ATCC CRL-1840) and were grown in DMEM (GIBCO BRL, Grand Island, NY) supplemented with 10% FCS. Cells were used within 72 h of achieving confluence.

ATP depletion and induction of hsp72. To induce ATP depletion, cells were incubated for 1 h at 37°C in glucose-free medium (DMEM; GIBCO BRL no. 23800-014) that contained sodium cyanide and 2-deoxy-D-glucose (5 mM each) as previously described (55). Fresh DMEM containing 10 mM glucose (without metabolic inhibitors) was used to initiate recovery. Parallel medium changes were made in controls using glucose-containing DMEM. To induce hsp72, OK cells were heated to 42.5 ± 0.5°C for 45 min in a temperature-regulated incubator followed by incubation at 37°C for 16-18 h (55).

Delta Psi m. Delta Psi m was assessed in viable OK cells with the use of 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazol-carbocyanine iodide (JC-1; Molecular Probes, Eugene, OR), a lipophilic, cationic dye, according to the manufacturer's protocol. Cells were plated on coverslips coated with rat tail collagen in 12-well plates in DMEM supplemented with 10% FCS. Subconfluent monolayers of cells were washed with PBS three times and then incubated with DMEM containing cyanide (5 mM) and 2-deoxy-D-glucose (5 mM) in glucose-free DMEM at 37°C. JC-1 (10 µg/ml) was added during the final 10 min of exposure to the metabolic inhibitors (for 30, 60, and 90 min in the absence of recovery). To intentionally disrupt Delta Psi m, 1 µM carbonyl cyanide p-(trifluoromethoxy)phenylhydrazone (FCCP) was added to cells for 10 min before addition of JC-1. The cells were rinsed twice, and then coverslips were mounted on glass slides. Dual-emission images were simultaneously obtained by using an inverted laser scanning confocal microscope (Zeiss LSM 510, Thornwood, NY) at 475 nm (excitation) and 525 and 590 nm (emission) in 1-µm-thick sections.

ATP production rate. State III mitochondrial respiration, the maximal capacity for mitochondria to generate ATP, was estimated by use of an assay developed in our laboratory (6, 55). In this assay, cells are selectively permeabilized with digitonin (0.15 mg/ml) in the presence of metabolic substrates (5 mM glutamate, 1 mM butyrate, 5 mM malate, and 600 nmol ADP) for 10 min at 37°C. ATP production was measured with luciferase (Analytical Luminescence Laboratory, San Diego, CA) in a luminometer (Turner Designs, Sunnyvale, CA). The results are expressed in nanomoles of ATP per milligram of protein.

Immunoblot analysis and coimmunoprecipitation. Harvested cells were resuspended in cell lysis buffer (containing 150 mM NaCl, 10 mM Tris · HCl, 5 mM EDTA, 1 mM EGTA, and 1% Triton X-100) and protease inhibitors (5 µM AEBSF-HCl, 10 nM leupeptin, 1.5 nM aprotinin, 10 nM E-64, and 5 µM EDTA; pH 7.40; Calbiochem-Novabiochem, San Diego, CA). The cells were sonicated and then centrifuged at 10,000 g for 10 min at 4°C. Antigens in the supernatant were detected by immunoblot with the use of commercially available monoclonal antibodies directed against cytochrome c (Research Diagnostics, Flanders, NJ; catalog no. RDI-cytoC12-abm), hsp72 (Amersham, Arlington Heights, IL), and AIF (Santa Cruz Biotechnology, Santa Cruz, CA). Specific protein bands were detected with an anti-IgG antibody coupled to a horseradish peroxidase-based enzyme-linked chemiluminescence system (Lumigolow; Kirkegaard and Perry, Gaithersberg, MD). After digitization of the image of each immunoblot (Hewlett-Packard, Desk Scan II), band densities were quantified by use of NIH Image Quant software. Cytosolic protein fractions were obtained by incubation of cells with 0.15 mg/ml digitonin for 10 min at 4°C before immunoprecipitation (IP). Samples were then incubated overnight at 4°C with a polyclonal rabbit antibody directed against cytochrome c (1-2 µg · mg protein-1 · ml buffer-1; Santa Cruz Biotechnology) or hsp72 (1-2 µg · mg protein-1 · ml buffer-1; Stress Gen). The IP buffer contained 150 mM NaCl, 10 mM Tris · HCl, 5 mM EDTA, 1 mM EGTA, 1% Triton X-100, 0.5% NP-40, and protease inhibitors at pH 7.4. To prevent the potential release of proteins attached to hsp72 during isolation, apyrase (10 U/ml), a compound that causes ATP hydrolysis, was added. The absence of ATP prevents the dissociation of HSP70 from its bound ligands during sample preparation (53), permitting coimmunoprecipitation studies to be performed (54). After protein separation by 12% SDS gel electrophoresis, the immunoblots were probed with one or more specific antibodies.

Cellular distribution of cytochrome c and AIF. To visualize mitochondria, subconfluent, live cells grown on glass coverslips were incubated with Mitotracker Green-FM (600 nM; Molecular Probes), a mitochondrial specific marker, for 30 min at 37°C. Cells were then fixed in methanol (4°C for 20 min), and routine immunohistochemistry was performed with an anti-cytochrome c antibody (cytoC12-abm; Research Diagnostics) that was detected with a Cy3-conjugated antibody (Jackson ImmunoResearch Laboratories, West Grove, PA). Confocal microscopy was used to localize cytochrome c and Mitotracker Green.

To quantify "cytosolic" and "mitochondrial" pools of cytochrome c and AIF, immunoblot analysis was performed in samples of cells exposed to either digitonin or SDS, respectively. Exposure to digitonin, a selective detergent, permeabilizes the plasma membrane without disrupting mitochondria (45) or their ability to generate ATP (6). In contrast, SDS solubilizes virtually all cell and organelle membranes, permitting cytochrome c (and AIF) within mitochondria to be detected (45). After removal of the medium, cell monolayers were incubated for 10 min at 4°C in 100-mm2 dishes with 2 ml of an intracellular-like buffer containing 120 mM KCl, 5 mM KH2PO4, 10 mM HEPES, 2 mM EGTA, 1% bovine serum albumin, 0.15 mg/ml digitonin, and a mixture of protease inhibitors (described above) and then harvested with a rubber policeman as previously described (55). Samples were centrifuged at 14,000 g for 5 min at 4°C. The supernatant was designated as the "digitonin-soluble" protein fraction. The pellet was incubated in 2% SDS at 4°C for 5 min, sonicated, and then centrifuged at 14,000 g for 5 min. This supernatant was designated as the "SDS-soluble" protein fraction. The content of cytochrome c and AIF was examined in both fractions by immunoblot analysis.

Caspase 3 activity. Caspase 3 enzyme activity was measured with the use of a fluorometric assay according to the manufacturer's protocol (ApoAlert caspase 3 fluorescence assay; Clontech Laboratories, Palo Alto, CA). OK cells grown on 60-mm2 dishes were lysed with 200 µl of chilled cell lysis buffer (Clontech Laboratories) on ice for 10 min. After centrifugation (12,000 rpm × 3 min, 4°C), 50 µl of supernatant, 50 µl of 2× reaction buffer containing 1 mM 1,4-dithiothreitol, and 5 µl of caspase substrate (DEVD-AFC; 50 µM) were added to each well of a 96-well plate. After incubation at 37°C for 1 h, fluorescence was determined with the use of a plate reader with a 400-nm excitation filter and 505-nm emission filter (Spectra Max Gemini, Molecular Devices, Sunnyvale, CA). To confirm assay specificity, parallel reactions were performed either without substrate or in the presence of a specific caspase 3 inhibitor (1 µl DEVD-CHO; Clontech Labs). The role of hsp72 per se in caspase 3 activation was evaluated by adding 1 µg of purified human hsp72 (Stress Gen, SPP-755, Victoria, BC, Canada) to the assay mixture and then repeating measurements of caspase 3 activity. This concentration of hsp72 exceeds the concentration of this protein detected in cells subjected to heat stress (44).

Protein assay. Protein concentrations were determined with a colorometric dye-binding assay [bicinchoninic acid (BCA) assay; Pierce, Rockford, IL]. Results are expressed in milligrams of protein per milliliter.

Statistical analysis. Data are expressed as means ± SE. Comparison of two groups was performed using a two-tailed Student's t-test. Results involving more than one group were compared using analysis of variance (ANOVA) and were then analyzed with the Fishers post hoc test. A result was considered significant if P value < 0.05.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

To determine the effect of exposure to cyanide and 2-deoxy-D-glucose in the absence of exogenous dextrose on Delta Psi m, live cells were incubated with JC-1 (Fig. 1). In control cells (Fig. 1A), mitochondria demonstrated a heterogeneous pattern of fluorescence. Many mitochondria exhibited red fluorescence, demonstrating JC-1 aggregation in response to a normal Delta Psi m. Some mitochondria exhibited green fluorescence, demonstrating the presence of JC-1 monomers, a finding consistent with a low Delta Psi m. In other control cells, a white pseudocolor (representing colocalization of JC-1 monomers and aggregates) was observed. Higher magnification confirmed the heterogeneous pattern of Delta Psi m within a single cell (Fig. 1A, inset). After 30-min exposure to metabolic inhibitors (without recovery), most JC-1 was present in the monomeric (green) form (Fig. 1B). At higher magnification, most mitochondria were depolarized (Fig. 1B, inset). Green JC-1 monomers also predominated after 60 and 90 min of exposure to metabolic inhibitors without recovery (data not shown). Incubation with FCCP, a mitochondrial uncoupling agent (data not shown), produced similar increases in the monomeric form of JC-1 as did exposure to metabolic inhibitors. Prior heat stress (Fig. 1C) did not have an independent effect on Delta Psi m, since the pattern of JC-1 staining was similar to that observed in control cells. During exposure to metabolic inhibitors, most mitochondria in previously heated cells were depolarized (Fig. 1D).


View larger version (88K):
[in this window]
[in a new window]
 
Fig. 1.   Mitochondrial membrane potential during ATP depletion. Viable cells loaded with 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazol-carbocyanine iodide (JC-1) were examined by laser scanning confocal microscopy. In control cells (A), numerous red JC-1 aggregates were precipitated by a high mitochondrial membrane voltage potential (Delta Psi m). Green fluorescence (indicating the presence of JC-1 monomers indicative of low Delta Psi m) and areas of white pseudofluorescence (representing JC-1 monomers and JC-1 aggregates in close proximity) were visible. At higher magnification, similar heterogeneity in JC-1 staining is seen (inset). After 30-min exposure to metabolic inhibitors (B and inset), loss of Delta Psi m (decreased red fluorescence) was evident. In previously heated cells, the pattern of JC-1 fluorescence was similar to control under normal conditions (C) as well as after exposure to metabolic inhibitors (D). The bar indicates 5 µM.

If the mitochondria were a target for injury by metabolic inhibitors, then organelle dysfunction would be expected. To assess mitochondrial function, the rate of ATP production was examined in selectively permeabilized OK cells under conditions that estimate maximal (state III) mitochondrial respiration. Compared with control, the maximal rate of ATP production fell by ~50% after 60 min of exposure to metabolic inhibitors (Fig. 2). In this experiment, cyanide and 2-deoxy-D-glucose were removed during the measurements of mitochondrial respiration. Prior heat stress prevented the mitochondrial dysfunction associated with exposure to metabolic inhibitors (P < 0.05; ATP deplete vs. heat stress + ATP deplete; n = 6). The observation that previously heated cells exposed to metabolic inhibitors exhibited similar rates of respiration as did controls suggests that these agents had little if any residual effect on mitochondrial metabolism. Prior heat stress alone (43°C followed by 16 h of recovery at 37°C) did not significantly alter mitochondrial ATP production compared with controls (P > 0.05).


View larger version (30K):
[in this window]
[in a new window]
 
Fig. 2.   ATP production (state III mitochondrial respiration). Maximal mitochondrial ATP production rate was measured in control, previously heat-stressed, and ATP-depleted renal cells over 10 min at 37°C in the presence of ADP and metabolic substrates (see METHODS). Data are expressed as mean values ± SE; n = 6 for each group. * P < 0.05 vs. control; dagger  P < 0.05 vs. ATP deplete by ANOVA.

Because exposure to metabolic inhibitors induced mitochondrial depolarization and impaired oxidative phosphorylation, release of cytochrome c was examined by two complementary techniques. First, cytochrome c (detected with a primary antibody to cytochrome c and a secondary antibody coupled to Cy3) and Mitotracker Green (a fluorescent probe that is highly concentrated within mitochondria) were localized in renal epithelial cells by use of immunohistochemistry. In control cells, cytochrome c and Mitotracker colocalized, represented as yellow-orange fluorescence (Fig. 3A). After 60 min of exposure to metabolic inhibitors, cytochrome c and Mitotracker no longer colocalized in many cells (Fig. 3B). By immunofluorescence, cytochrome c (red) was not observed in the cytosol of cells exposed to metabolic inhibitors. This may be due to the translocation of a relatively small amount of cytochrome c into a large compartment (i.e., the cytosol). To quantify cytochrome c release, immunoblot analysis was performed on two cell fractions: a cytosolic fraction (normally containing no immunoreactive cytochrome c) obtained after exposure of intact cells to digitonin, a plasma membrane-selective detergent, and a whole cell protein fraction (including mitochondria) solubilized with SDS, a nonselective detergent. In control cells, leak of cytochrome c (14 kDa) into the cytosol could be observed after 60 min of exposure to cyanide and 2-deoxy-D-glucose in the absence of dextrose (Fig. 4A). Cytosolic cytochrome c increased 30 min after removal of the metabolic inhibitors (ATP repletion). In contrast, prior heat stress decreased the translocation of cytochrome c. In both control and previously heated cells, exposure to metabolic inhibitors or recovery caused minimal changes in the amount of residual mitochondrial cytochrome c detected in the whole cell fraction (SDS). To quantify the leak of cytochrome c during and after exposure to metabolic inhibitors, data from several studies were analyzed by densitometry (Fig. 4B). In these studies, prior heat stress significantly reduced the leak of cytochrome c associated with exposure to metabolic inhibitors (P < 0.05, n = 3).


View larger version (32K):
[in this window]
[in a new window]
 
Fig. 3.   A and B: localization of cytochrome c and mitotracker during ATP depletion. Control (A) and ATP-depleted (B) renal cells were incubated with Mitotracker Green FM (green) for 30 min at 37°C. Cells were then fixed in methanol, and routine immunohistochemistry was performed with an anti-cytochrome c antibody detected with a Cy3-conjugated secondary antibody (red). Confocal microscopy was used to colocalize cytochrome c and Mitotracker Green. Overlap of the 2 markers appears as pseudocolor (yellow-orange). The 2 markers were highly colocalized in control but not in ATP-depleted cells.



View larger version (34K):
[in this window]
[in a new window]
 
Fig. 4.   A and B: effect of ATP depletion and recovery on cytochrome c localization. To evaluate the cytosolic and mitochondrial pools of cytochrome c, immunoblot analysis was performed on whole cell lysates after solubilization with either a plasma membrane selective (digitonin) or a nonselective (SDS) detergent. The amount of total protein loaded into each lane is indicated. A single immunoblot representative of 3 studies is shown (A). Densitometric analysis obtained from 3 immunoblot studies (expressed as %total cell cytochrome c released) was used to quantify protein translocation from the "mitochondrial" to the "cytosolic" compartment (B) at baseline immediately after ATP depletion and after 30 min of recovery in control (open bars) and previously heated cells (solid bars). * P < 0.05 vs. control; n = 3.

To confirm injury to the mitochondrial membrane, the translocation of AIF (57 kDa) was also examined after exposure to metabolic inhibitors. In control cells, virtually no AIF was detected in the protein fraction obtained by exposing cells to digitonin (Fig. 5). In contrast to normal cells at baseline, progressively larger amounts of AIF were detected in the cytosolic protein fraction after the removal of metabolic inhibitors. The major immunoreactive band present in the protein fraction obtained after exposure to SDS confirmed the localization of AIF at the expected molecular mass. A single minor band (~40-45 kDa) was also detected in the SDS protein fraction. Although not characterized, this minor band has been previously detected by others (15). Similar to cytochrome c, most AIF remained in the mitochondrial compartment even in injured cells.


View larger version (38K):
[in this window]
[in a new window]
 
Fig. 5.   Apoptosis-inducing factor (AIF) release after ATP depletion. Translocation of AIF in renal cells was evaluated with the use of mitochondrial membrane selective and nonselective detergent solubilization as described in legend to Fig 4. One of four representative immunoblots is shown. The identity of the minor band (40-45 kDa) in the SDS protein fraction has not been evaluated.

The release of intermitochondrial membrane proteins into the cytosol could result from nonspecific mitochondrial swelling/rupture to selective opening of a channel in the outer membrane (5, 51) or to both. To distinguish between these possibilities, cells were preincubated with bongkrekic acid (50 µM × 2 h), an inhibitor of the membrane permeability transition pore (MPT; Refs. 5 and 10). Exposure to bongkrekic acid alone did not cause AIF release (data not shown). This MPT inhibitor significantly reduced AIF leak (Fig. 6) during ATP depletion as well as during both time points of recovery (P < 0.05 vs. control).


View larger version (21K):
[in this window]
[in a new window]
 
Fig. 6.   Effect of a membrane permeability transition pore (MPT) inhibitor on AIF release. Release of mitochondrial AIF into the cytosol was evaluated after 60 min of exposure to metabolic inhibitors as well as during recovery (15 and 30 min) in the presence (vs. the absence) of bongkrekic acid (BA; 50 µM). Cytosolic samples were obtained by exposing cells to digitonin (as described in METHODS). Densitometric analysis of the data is expressed as the percent decrease in AIF release (mean ± SE) in cells exposed to BA compared with control (* P < 0.05, n = 3-4 experiments at each time point).

To determine whether the leak of cytochrome c in ATP-depleted cells activated the apoptotic pathway, caspase 3 activity was measured. This assay did not require digitonin. This is important in that detergent exposure could increase the release of cytochrome c and/or AIF from mitochondria injured by exposure to metabolic inhibitors. Compared with control cells, exposure to metabolic inhibitors resulted in a seven- to eightfold increase in caspase 3 activity (P < 0.5, n = 3; Fig. 7). Prior heat stress significantly ameliorated the activation of caspase 3 caused by exposure to metabolic inhibitors (P < 0.05, n = 3). Purified hsp72 (1 µg) added to the caspase 3 reaction mixture (in vitro) did not alter the activity of caspase 3 in cell lysates obtained from cells exposed to metabolic inhibitors (data not shown). In contrast, the addition of a specific inhibitor of caspase 3 completely abolished caspase activity, confirming the specificity of this assay (data not shown).


View larger version (17K):
[in this window]
[in a new window]
 
Fig. 7.   Caspase 3 activation. Caspase 3 activity was measured in samples obtained from whole cell lysates derived from control (diamond ) and previously heated (black-triangle) renal cells at baseline immediately after 60 min of ATP depletion and at 15 and 30 min of recovery with the use of an in vitro fluorometric assay. Data are expressed relative to the baseline caspase 3 activity and represent the mean ± SE values of 3 separate experiments. Each sample was assayed in triplicate. Background activity measured in the absence of the specific caspase substrate was subtracted from each sample. * P < 0.05 vs. control.

Potent suppression of caspase 3 despite measurable cytochrome c release in heat-stressed cells suggested that an additional mechanism might inhibit apoptosis. To explore the possibility that hsp72 might prevent caspase activation by binding cytosolic cytochrome c, coimmunoprecipitation of the two proteins was investigated. In control cells, interaction between hsp72 and cytochrome c was observed immediately after 60 min of exposure to metabolic inhibitors and to a lesser degree during recovery (Fig. 8). Despite the reduction in cytosolic cytochrome c in previously heated cells (Fig. 8B), the interaction between these two proteins after exposure to metabolic inhibitors was markedly increased (Fig. 8A). Immunoblot analysis confirmed the adequacy of heat stress in increasing the content of hsp72 (Fig. 8C). Persistent nonspecific bands precluded interpretation of studies in which hsp72 was immunoprecipitated and the blots were probed for cytochrome c (data not shown).


View larger version (59K):
[in this window]
[in a new window]
 
Fig. 8.   Hsp72 interacts with cytochrome c. Interaction between hsp72 and cytochrome c was examined in samples obtained from control and previously heated opossum kidney (OK) cells by coimmunoprecipitation (IP). Cytosolic samples were obtained by selective permeabilization of the plasma membrane (as described in legend to Fig 4). After IP with a rabbit anti-cytochrome c antibody, hsp72 was detected by use of a mouse monoclonal antibody (A). Hsp72 was identified with the use of commercially available purified protein (top left). After IP with a rabbit anti-cytochrome c antibody, cytochrome c was detected with a mouse monoclonal antibody (B). Cytochrome c was identified by use of purified cytochrome c (middle left). Hsp72 content was assessed after IP and immunoblot (IB) analysis with the use of a mouse monoclonal antibody (C). Each sample contained 100 µg of total protein separated by 12% SDS-PAGE.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Historically, the mitochondrion has been overlooked as an early mediator of apoptosis (30, 51). This is largely due to the fact that major changes in mitochondrial membrane integrity and organelle function occur well before the classic morphological signs of cell death are evident (30, 51). Subsequent investigations have shown that mitochondrial products are rate limiting for activating caspase and endonuclease in cell free systems, that stabilization of mitochondrial membranes prevents apoptosis, and that selective overexpression of mitochondrial protective proteins (e.g., Bcl family members) ameliorates apoptosis (reviewed in Ref 51). These observations emphasize the role of mitochondria in regulating apoptosis.

In the present study, mitochondrial injury accompanied exposure to metabolic inhibitors in vitro. Delta Psi m decreased within 30 min of exposure to metabolic inhibitors and remained low up to 90 min. This observation is similar to that reported by Weinberg et al. (57) in renal epithelial cells subjected to hypoxia/reoxygenation. The observed decrement in Delta Psi m is not surprising, since ATP depletion included exposure to cyanide, a potent inhibitor of mitochondrial respiration. The loss of Delta Psi m has been attributed to dissipation of the proton gradient responsible for the voltage across the inner mitochondrial membrane that in some circumstances is associated with the opening of the MPT (51). The MPT is a regulated megachannel composed of the adenine nucleotide translocator (ANT), the voltage-dependent anion channel (VDAC), cyclophilin D, hexokinase, and other yet-unidentified proteins (43, 51) that are susceptible to stress (9, 23, 31).

In addition to disrupting Delta Psi m, exposure to metabolic inhibitors caused a 50% reduction in state III mitochondrial respiration (Fig. 2). This decrease in mitochondrial energy production is multifactoral. First, the protein gradient across the mitochondrial membrane is directly coupled to ATP production. A decrease in Delta Psi m during ATP depletion would reduce ATP production by oxidative phosphorylation. Second, loss of cytochrome c during the period of ATP depletion and early recovery would directly compromise electron transport. Third, loss of AIF, a flavoprotein (35), could adversely affect metabolic function by mitochondria. Last, Weinberg et al. (57) have recently demonstrated that hypoxia/reoxygentation of rabbit tubules in suspension damages complex I enzymes in the oxidative phosphorylation pathway proximal to cytochrome c (57). A decrement in the ability to generate ATP ultimately leads to a fall in ATP content. Both the absolute level of ATP and the duration of ATP depletion are critical determinants of whether the renal epithelial cell dies by apoptosis or necrosis (18, 29).

In the present study, it is important to note that the release of cytochrome c and AIF (Figs. 4 and 5) precedes morphological evidence of apoptosis (e.g., the formation of apoptotic bodies, chromatin condensation, and endonucleosomal DNA fragmentation) previously observed in these same cells after ATP depletion (53). In addition, substantial evidence suggests that the release of either cytochrome c or AIF is sufficient to cause apoptosis (15, 43, 49). In forming the apoptosome, cytochrome c activates caspase 9, the first step in the enzyme cascade that ultimately causes apoptosis (7, 8). In contrast, AIF induces nuclear changes that are characteristic of apoptosis by a "caspase-independent" mechanism that directly activates endonucleases and causes chromatin condensation (15, 17, 24, 49). In the present study, release of both cytochrome c and AIF into the cytosol implies that outer mitochondrial membrane permeability has been altered during ATP depletion and recovery. Loss of mitochondrial membrane integrity could result from nonspecific membrane injury or more selective opening of the MPT. To distinguish between these possibilities, cells were preincubated with bongkrekic acid, an MPT inhibitor that binds to the ANT (10). Because this agent is reportedly more specific and effective for inhibiting MPT than cyclosporine A (51), the later agent was not used in the present study. Bongkrekic acid significantly inhibited AIF release during and after exposure to metabolic inhibitors (Fig. 6). Incomplete protection of AIF release by bongkrekic acid could be due to a suboptimal drug concentration or inadequate exposure to the agent before injury. Alternatively, AIF leak could also occur by an MPT-independent mechanism precipitated by mitochondrial swelling or rupture (41).

What factors disrupt the outer mitochondrial membrane? Recent investigations have focused on the role of two proteins, BAX and Bcl2, as mediators of mitochondrial permeability transition (PT) and apoptosis (9, 23, 31, 32, 34, 45). Insults that increase the ratio of BAX to Bcl2 promote apoptosis by allowing BAX to form homodimers within the mitochondrial membrane (26, 34). The mechanism by which BAX causes membrane injury remains unclear. Some investigators suggest that BAX cooperates with the ANT, promoting MPT (9). Others suggest that BAX does not require association with components of the MPT to create an outer membrane channel (34). In contrast to BAX, Bcl2 prevents MPT opening by interfering with BAX (9, 26, 34). In renal epithelial cells, hypoxia and ATP depletion are associated with marked changes in both BAX and Bcl2. After induction of ATP depletion with a mitochondrial uncoupling agent, BAX translocates to the mitochondrial membrane (45). Translocation of BAX is associated with the release of cytochrome c into the cytosol and subsequent apoptosis (45). In a similar in vitro model, as described in the present study, marked proapoptotic changes in the ratio of Bcl2 to BAX were observed in OK cells exposed to metabolic inhibitors (53). These studies do not preclude the possibility that mitochondrial "toxins" such as reactive oxygen species contribute to membrane injury and promote the leak of proapoptotic proteins (31).

The accumulation of hsp72, either selectively or induced by heat stress, ameliorates apoptosis in a variety of experimental models (4, 19, 39, 42, 44, 53). In the present study, the accumulation of hsp72 was associated with preservation of the outer mitochondrial membrane (Fig. 4) and improved state III mitochondrial respiration (Fig. 2). This protective effect occurred despite similar reductions in Delta Psi m (Fig. 1) and ATP content during exposure to metabolic inhibitors (55). These observations suggest that both heat-stress and control cells are equally sensitive to the effects of cyanide, a potent inhibitor of electron transport that causes mitochondrial depolarization, a function of the inner mitochondrial membrane (5). In contrast, heat stress prevents the loss of proapoptotic proteins from the intermembranous mitochondrial space, suggesting that disruption of the outer membrane does not necessarily accompany mitochondrial depolarization. Preservation of state III respiration in heat-stressed cells occurred despite some loss of cytochrome c (Fig. 4B). This is likely due to the fact that renal epithelial cells have a large reserve for oxidative ATP production (52). Preservation of respiration could also be explained by an excess of available cytochrome c and incomplete release from mitochondria. The presence of oxidative reserve is supported by the observation that resting cells exhibit heterogeneity in Delta Psi m (Fig. 1) and is consistent with prior reports in nonrenal cells in which Delta Psi m was measured by lipophilic, cationic probes (27, 37).

Recent studies in our laboratory confirm that the selective overexpression of hsp72 (in the absence of heat stress) is sufficient to inhibit caspase 3 activation and improve cell survival in renal epithelial cells exposed to metabolic inhibitors (56). By protecting the mitochondrial membrane, hsp72 could inhibit apoptosis and improve cell survival. Mosser et al. (36) have shown that HSP70 prevents cytochrome c release and caspase activation after thermal injury. Although the mechanism by which hsp72 protects the mitochondrial membrane has not been characterized, Bcl2 is a likely candidate for protecting mitochondria in our model. Selective overexpression of Bcl2 ameliorates mitochondrial membrane damage in hypoxic renal epithelial cells (45). Furthermore, hsp72 immunoprecipitates with Bcl2 but not Bax in renal cells exposed to metabolic inhibitors (53). Interaction between hsp72 and Bcl2 could afford cytoprotection by restoring Bcl2 function, a role compatible with the chaperone function of this stress protein (36, 40). Regardless of the mechanism by which hsp72 protects the outer mitochondrial membrane during exposure to metabolic inhibitors, reduction of cytochrome c release would prevent apoptosome assembly and caspase activation, central events in the apoptosis pathway (3).

In this study, a previously unreported interaction between hsp72 and cytochrome c was observed. The interaction was greatest in ATP-depleted cells subjected to prior heat stress (Fig. 7), despite the marked decrement in cytochrome c released by these cells (Fig. 4). In both control and heated cells, this interaction was most apparent immediately after the period of ATP depletion. The observation is consistent with function of HSP70 members as chaperones that require ATP to release bound ligands (40). It is conceivable that by binding cytosolic cytochrome c, hsp72 interferes with the assembly of the apoptosome complex, thereby reducing caspase activation. Interestingly, HSP70 has been reported to inhibit apoptosome assembly, although the proposed mechanism did not include interaction with cytochrome c (4). Binding to hsp72 could also interfere with the immunodetection of cytochrome c in the cytosol of cells exposed to metabolic inhibitors (Fig. 3B). In addition to preventing cytochrome c-dependent caspase activation, HSP70 can inhibit apoptosis by antagonizing AIF, a proapoptotic protein responsible for DNA injury (44). This finding may be important in our model, since prior heat stress significantly reduced DNA fragmentation in OK cells exposed to metabolic inhibitors (53). Other investigators have shown that HSP70 prevents the activation of proapoptotic stress kinases such as c-Jun NH2-terminal kinase (JNK) (20) and inhibits "executioner" caspases both proximal (28) and distal (22) to caspase 3. The failure of hsp72 to inhibit caspase 3 activity in vitro suggests that this cytoprotectant protein acts at an earlier step in the apoptotic cascade in our model. Given the existing data, it is likely that hsp72 modulates apoptosis at a variety of checkpoints that may differ by cell type and the nature of the stress.

In some studies, the morphological and biochemical manifestations of apoptosis have been inhibited without improving long-term cell viability (33, 45). Although the present studies were short-term (i.e., limited to 1 h post-ATP depletion), hsp72 accumulation significantly increased renal epithelial cell survival for at least 6 days after exposure to metabolic inhibitors (55). The improvement in cell survival associated with the induction of hsp72 contrasts with biochemical maneuvers that prevented caspase activation downstream of mitochondrial injury but failed to enhance viability (33, 45) and emphasizes the role of mitochondria in determining the fate of a cell. The present study demonstrates that the outer mitochondrial membrane is an early target of injury in cells exposed to metabolic inhibitors and that upregulation of HSPs including hsp72 may ameliorate apoptosis by preventing the release and possibly by inhibiting the action of proapoptotic proteins.


    ACKNOWLEDGEMENTS

This research was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grants DK-53387 (S. C. Borkan) and DK-5298 (J. H. Schwartz) and a supplemental award from the American Society of Nephrology (S. C. Borkan).


    FOOTNOTES

Address for reprint requests and other correspondence: S. C. Borkan, Evans Biomedical Research Center, Renal Section, Rm. 547, 650 Albany St., Boston, MA 02118-2518 (E-mail: sborkan{at}bu.edu).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

May 15, 2002;10.1152/ajpcell.00517.2001

Received 26 October 2001; accepted in final form 2 May 2002.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Adrain, C, and Martin SJ. The mitochondrial apoptosome: a killer unleashed by the cytochrome seas. Trends Biochem Sci 26: 390-397, 2001[ISI][Medline].

2.   Ali, ST, Coggins JR, and Jacobs HT. The study of cell-death proteins in the outer mitochondrial membrane by chemical cross-linking. Biochem J 325: 321-324, 1997[ISI][Medline].

3.   Beere, HM, and Green DR. Stress management: heat shock protein-70 and the regulation of apoptosis. Trends Cell Biol 11: 6-10, 2001[ISI][Medline].

4.   Beere, HM, Wolf BB, Cain K, Mosser DD, Mahboubi A, Kuwana T, Tailor P, Morimoto RI, Cohen GM, and Green DR. Heat-shock protein 70 inhibits apoptosis by preventing recruitment of procaspase-9 to the Apaf-1 apoptosome. Nat Cell Biol 2: 469-475, 2000[ISI][Medline].

5.   Bernardi, P, Scorrano L, Colonna R, Petronilli V, and Di Lisa F. Mitochondria and cell death. Mechanistic aspects and methodological issues. Eur J Biochem 264: 687-701, 1999[Abstract/Free Full Text].

6.   Borkan, SC, Emami A, and Schwartz JH. Heat stress protein-associated cytoprotection of inner medullary collecting duct cells from rat kidney. Am J Physiol Renal Fluid Electrolyte Physiol 265: F333-F341, 1993[Abstract/Free Full Text].

7.   Bratton, SB, and Cohen GM. Apoptotic death sensor: an organelle's alter ego? Trends Pharmacol Sci 22: 306-315, 2001[ISI][Medline].

8.   Bratton, SB, Walker G, Srinivasula SM, Sun XM, Butterworth M, Alnemri ES, and Cohen GM. Recruitment, activation and retention of caspases-9 and -3 by Apaf-1 apoptosome and associated XIAP complexes. EMBO J 20: 998-1009, 2001[Abstract/Free Full Text].

9.   Brenner, C, Cadiou H, Vieira HL, Zamzami N, Marzo I, Xie Z, Leber B, Andrews D, Duclohier H, Reed JC, and Kroemer G. Bcl-2 and Bax regulate the channel activity of the mitochondrial adenine nucleotide translocator. Oncogene 19: 329-336, 2000[ISI][Medline].

10.   Cao, G, Minami M, Pei W, Yan C, Chen D, O'Horo C, Graham SH, and Chen J. Intracellular Bax translocation after transient cerebral ischemia: implications for a role of the mitochondrial apoptotic signaling pathway in ischemic neuronal death. J Cereb Blood Flow Metab 21: 321-333, 2001[ISI][Medline].

11.   Chevalier, RL, Thornhill BA, and Wolstenholme JT. Renal cellular response to ureteral obstruction: role of maturation and angiotensin II. Am J Physiol Renal Physiol 277: F41-F47, 1999[Abstract/Free Full Text].

12.   Chien, CT, Chen CF, Hsu SM, Lee PH, and Lai MK. Protective mechanism of preconditioning hypoxia attenuates apoptosis formation during renal ischemia/reperfusion phase. Transplant Proc 31: 2012-2013, 1999[ISI][Medline].

13.   Chiu, SM, and Oleinick NL. Dissociation of mitochondrial depolarization from cytochrome c release during apoptosis induced by photodynamic therapy. Br J Cancer 84: 1099-1106, 2001[ISI][Medline].

14.   Daugas, E, Nochy D, Ravagnan L, Loeffler M, Susin SA, Zamzami N, and Kroemer G. Apoptosis-inducing factor (AIF): a ubiquitous mitochondrial oxidoreductase involved in apoptosis. FEBS Lett 476: 118-123, 2000[ISI][Medline].

15.   Daugas, E, Susin SA, Zamzami N, Ferri KF, Irinopoulou T, Larochette N, Prevost MC, Leber B, Andrews D, Penninger J, and Kroemer G. Mitochondrio-nuclear translocation of AIF in apoptosis and necrosis. FASEB J 14: 729-739, 2000[Abstract/Free Full Text].

16.   Dong, Z, Venkatachalam MA, Wang J, Patel Y, Saikumar P, Semenza GL, Force T, and Nishiyama J. Upregulation of apoptosis inhibitory protein IAP-2 by hypoxia. Hif-1-independent mechanisms. J Biol Chem 276: 18702-18709, 2001[Abstract/Free Full Text].

17.   Dumont, C, Durrbach A, Bidere N, Rouleau M, Kroemer G, Bernard G, Hirsch F, Charpentier B, Susin SA, and Senik A. Caspase-independent commitment phase to apoptosis in activated blood T lymphocytes: reversibility at low apoptotic insult. Blood 96: 1030-1038, 2000[Abstract/Free Full Text].

18.   Eguchi, Y, Shimizu S, and Tsujimoto Y. Intracellular ATP levels determine cell death fate by apoptosis or necrosis. Cancer Res 57: 1835-1840, 1997[Abstract].

19.   Gabai, VL, Meriin AB, Mosser DD, Caron AW, Rits S, Shifrin VI, and Sherman MY. Hsp70 prevents activation of stress kinases. A novel pathway of cellular thermotolerance. J Biol Chem 272: 18033-18037, 1997[Abstract/Free Full Text].

20.   Gabai, VL, Yaglom JA, Volloch V, Meriin AB, Force T, Koutroumanis M, Massie B, Mosser DD, and Sherman MY. Hsp72-mediated suppression of c-Jun N-terminal kinase is implicated in development of tolerance to caspase-independent cell death. Mol Cell Biol 20: 6826-6836, 2000[Abstract/Free Full Text].

21.   Green, D, and Reed J. Mitochondria and apoptosis. Science 81: 1309-1312, 1998.

22.   Jaattela, M, Wissing D, Kokholm K, Kallunki T, and Egeblad M. Hsp70 exerts its anti-apoptotic function downstream of caspase-3-like proteases. EMBO J 17: 6124-6134, 1998[Abstract/Free Full Text].

23.   Jacotot, E, Costantini P, Laboureau E, Zamzami N, Susin SA, and Kroemer G. Mitochondrial membrane permeabilization during the apoptotic process. Ann NY Acad Sci 887: 18-30, 1999[Abstract/Free Full Text].

24.   Joza, N, Susin SA, Daugas E, Stanford WL, Cho SK, Li CY, Sasaki T, Elia AJ, Cheng HY, Ravagnan L, Ferri KF, Zamzami N, Wakeham A, Hakem R, Yoshida H, Kong YY, Mak TW, Zuniga-Pflucker JC, Kroemer G, and Penninger JM. Essential role of the mitochondrial apoptosis-inducing factor in programmed cell death. Nature 410: 549-554, 2001[ISI][Medline].

25.   Kim, JW, Chang TS, Lee JE, Huh SH, Yeon SW, Yang WS, Joe CO, Mook-Jung I, Tanzi RE, Kim TW, and Choi EJ. Negative regulation of the SAPK/JNK signaling pathway by presenilin 1. J Cell Biol 153: 457-463, 2001[Abstract/Free Full Text].

26.   Korsmeyer, SJ, Shutter JR, Veis DJ, Merry DE, and Oltvai ZN. Bcl-2/Bax: a rheostat that regulates an antioxidant pathway and cell death. Semin Cancer Biol 4: 327-332, 1993[ISI][Medline].

27.   Lemasters, JJ, Chacon E, Ohata H, Harper IS, Nieminen AL, Tesfai SA, and Herman B. Measurement of electrical potential, pH, and free calcium ion concentration in mitochondria of living cells by laser scanning confocal microscopy. Methods Enzymol 260: 428-444, 1995[ISI][Medline].

28.   Li, C, Lee J, Ko Y, Kim J, and Seo J. Heat shock protein 70 inhibits apoptosis downstream of cytochrome c release and upstream of caspase 3 activation. J Biol Chem 275: 25665-25671, 2000[Abstract/Free Full Text].

29.   Lieberthal, W, Menza SA, and Levine JS. Graded ATP depletion can cause necrosis or apoptosis of cultured mouse proximal tubular cells. Am J Physiol Renal Physiol 274: F315-F327, 1998[Abstract/Free Full Text].

30.   Loeffler, M, and Kroemer G. The mitochondrion in cell death control: certainties and incognita. Exp Cell Res 256: 19-26, 2000[ISI][Medline].

31.   Marchetti, P, Castedo M, Susin SA, Zamzami N, Hirsch T, Macho A, Haeffner A, Hirsch F, Geuskens M, and Kroemer G. Mitochondrial permeability transition is a central coordinating event of apoptosis. J Exp Med 184: 1155-1160, 1996[Abstract].

32.   Marzo, I, Brenner C, Zamzami N, Jurgensmeier JM, Susin SA, Vieira HL, Prevost MC, Xie Z, Matsuyama S, Reed JC, and Kroemer G. Bax and adenine nucleotide translocator cooperate in the mitochondrial control of apoptosis. Science 281: 2027-2031, 1998[Abstract/Free Full Text].

33.   Matsuyama, S, and Reed JC. Mitochondria-dependent apoptosis and cellular pH regulation. Cell Death Differ 7: 1155-1165, 2000[ISI][Medline].

34.   Mikhailov, V, Mikhailova M, Pulkrabek DJ, Dong Z, Venkatachalam MA, and Saikumar P. Bcl-2 prevents Bax oligomerization in the mitochondrial outer membrane. J Biol Chem 276: 18361-18374, 2001[Abstract/Free Full Text].

35.   Miramar, MD, Costantini P, Ravagnan L, Saraiva LM, Haouzi D, Brothers G, Penninger JM, Peleato ML, Kroemer G, and Susin SA. NADH oxidase activity of mitochondrial apoptosis-inducing factor. J Biol Chem 276: 16391-16398, 2001[Abstract/Free Full Text].

36.   Mosser, DD, Caron AW, Bourget L, Meriin AB, Sherman MY, Morimoto RI, and Massie B. The chaperone function of hsp70 is required for protection against stress-induced apoptosis. Mol Cell Biol 20: 7146-7159, 2000[Abstract/Free Full Text].

37.   Nicholls, DG, and Ward MW. Mitochondrial membrane potential and neuronal glutamate excitotoxicity: mortality and millivolts. Trends Neurosci 23: 166-174, 2000[ISI][Medline].

38.   Oberbauer, R, Rohrmoser M, Regele H, Muhlbacher F, and Mayer G. Apoptosis of tubular epithelial cells in donor kidney biopsies predicts early renal allograft function. J Am Soc Nephrol 10: 2006-2013, 1999[Abstract/Free Full Text].

39.   Park, KC, Kim DS, Choi HO, Kim KH, Chung JH, Eun HC, Lee JS, and Seo JS. Overexpression of HSP70 prevents ultraviolet B-induced apoptosis of a human melanoma cell line. Arch Dermatol Res 292: 482-487, 2000[ISI][Medline].

40.   Parsell, DA, and Lindquist S. The function of heat-shock proteins in stress tolerance: degradation and reactivation of damaged proteins. Annu Rev Genet 27: 437-496, 1993[ISI][Medline].

41.   Petit, PX, Goubern M, Diolez P, Susin SA, Zamzami N, and Kroemer G. Disruption of the outer mitochondrial membrane as a result of large amplitude swelling: the impact of irreversible permeability transition. FEBS Lett 426: 111-116, 1998[ISI][Medline].

42.   Polla, BS, Kantengwa S, Francois D, Salvioli S, Franceschi C, Marsac C, and Cossarizza A. Mitochondria are selective targets for the protective effects of heat shock against oxidative injury. Proc Natl Acad Sci USA 93: 6458-6463, 1996[Abstract/Free Full Text].

43.   Priault, M, Chaudhuri B, Clow A, Camougrand N, and Manon S. Investigation of bax-induced release of cytochrome c from yeast mitochondria permeability of mitochondrial membranes, role of VDAC and ATP requirement. Eur J Biochem 260: 684-691, 1999[Abstract/Free Full Text].

44.   Ravagnan, L, Gurbuxani S, Susin SA, Maisse C, Daugas E, Zamzami N, Mak T, Jaattela M, Penninger JM, Garrido C, and Kroemer G. Heat-shock protein 70 antagonizes apoptosis-inducing factor. Nat Cell Biol 3: 839-843, 2001[ISI][Medline].

45.   Saikumar, P, Dong Z, Patel Y, Hall K, Hopfer U, Weinberg JM, and Venkatachalam MA. Role of hypoxia-induced Bax translocation and cytochrome c release in reoxygenation injury. Oncogene 17: 3401-3415, 1998[ISI][Medline].

46.   Samali, A, Zhivotovsky B, Jones DP, and Orrenius S. Detection of pro-caspase-3 in cytosol and mitochondria of various tissues. FEBS Lett 431: 167-169, 1998[ISI][Medline].

47.   Schumer, M, Colombel MC, Sawczuk IS, Gobe G, Connor J, O'Toole KM, Olsson CA, Wise GJ, and Buttyan R. Morphologic, biochemical, and molecular evidence of apoptosis during the reperfusion phase after brief periods of renal ischemia. Am J Pathol 140: 831-838, 1992[Abstract].

48.   Shimizu, S, Eguchi Y, Kamiike W, Funahashi Y, Mignon A, Lacronique V, Matsuda H, and Tsujimoto Y. Bcl-2 prevents apoptotic mitochondrial dysfunction by regulating proton flux. Proc Natl Acad Sci USA 95: 1455-1459, 1998[Abstract/Free Full Text].

49.   Susin, SA, Daugas E, Ravagnan L, Samejima K, Zamzami N, Loeffler M, Costantini P, Ferri KF, Irinopoulou T, Prevost MC, Brothers G, Mak TW, Penninger J, Earnshaw WC, and Kroemer G. Two distinct pathways leading to nuclear apoptosis. J Exp Med 192: 571-580, 2000[Abstract/Free Full Text].

50.   Susin, SA, Lorenzo HK, Zamzami N, Marzo I, Snow BE, Brothers GM, Mangion J, Jacotot E, Costantini P, Loeffler M, Larochette N, Goodlett DR, Aebersold R, Siderovski DP, Penninger JM, and Kroemer G. Molecular characterization of mitochondrial apoptosis-inducing factor. Nature 397: 441-446, 1999[ISI][Medline].

51.   Susin, SA, Zamzami N, and Kroemer G. Mitochondria as regulators of apoptosis: doubt no more. Biochim Biophys Acta 1366: 151-165, 1998[ISI][Medline].

52.   Takano, T, Soltoff SP, Murdaugh S, and Mandel LJ. Intracellular respiratory dysfunction and cell injury in short-term anoxia of rabbit renal proximal tubules. J Clin Invest 76: 2377-2384, 1985[ISI][Medline].

53.   Wang, Y, Knowlton AA, Christensen TG, Shih T, and Borkan SC. Prior heat stress inhibits apoptosis in adenosine triphosphate-depleted renal tubular cells. Kidney Int 55: 2224-2235, 1999[ISI][Medline].

54.   Wang, Y, Li F, Schwartz J, Flint P, and Borkan S. c-Src and HSP72 interact in ATP-depleted renal epithelial cells. Am J Physiol Cell Physiol 281: 1667-1675, 2001.

55.   Wang, YH, and Borkan SC. Prior heat stress enhances survival of renal epithelial cells after ATP depletion. Am J Physiol Renal Fluid Electrolyte Physiol 270: F1057-F1065, 1996[Abstract/Free Full Text].

56.  Wang YH, Knowlton AA, Li FH, and Borkan SC. HSP 72 expression enhances survival in adenosine triphosphate-depleted renal epithelial cells. Cell Stress Chaperones. In Press.

57.   Weinberg, JM, Venkatachalam MA, Roeser NF, and Nissim I. Mitochondrial dysfunction during hypoxia/reoxygenation and its correction by anaerobic metabolism of citric acid cycle intermediates. Proc Natl Acad Sci USA 97: 2826-28231, 2000[Abstract/Free Full Text].

58.   Yang, B, Johnson TS, Thomas GL, Watson PF, Wagner B, and Nahas AM. Apoptosis and caspase-3 in experimental anti-glomerular basement membrane nephritis. J Am Soc Nephrol 12: 485-495, 2001[Abstract/Free Full Text].

59.   Zamzami, N, and Kroemer G. The mitochondrion in apoptosis: how Pandora's box opens. Nat Rev Mol Cell Biol 2: 67-71, 2001[ISI][Medline].


Am J Physiol Cell Physiol 283(3):C917-C926
0363-6143/02 $5.00 Copyright © 2002 the American Physiological Society