INTRODUCTION

TOP ABSTRACT INTRODUCTION METHODS RESULTS DISCUSSION REFERENCES

THE UNIQUE LOCATION OF ENDOTHELIAL cells at the interface of the bloodstream with different organ systems leads to exposure to a variety of stimuli that affect endothelial cell viability. Recent work suggests that changes in pH and mitochondrial function may play an important role in regulating several components of apoptotic signal transduction pathways, including caspase activation, poly(ADP-ribose) polymerase (PARP) cleavage, and DNA fragmentation in some cell types (8, 12, 15, 17, 45, 53). In contrast, the signal transduction pathways that regulate the development of apoptosis in endothelial cells are not well defined.

Extracellular acidosis exerts a protective effect on the onset of cell death in different cell lines, including neuronal, cardiac, renal, hepatic, and endothelial cells (1, 7, 36, 53). Most of this previous work has focused on the role of pH in modifying necrotic cell death. Little information is available on the role of pH in modifying apoptotic signal transduction in endothelial cells (8). Similarly, mitochondria have a crucial role in the maintenance of a critical concentration of ATP required for cell survival (6, 9, 34). Recent evidence also suggests that mitochondria have a central role in regulating the progression of apoptotic signaling (28). Disruption of mitochondrial transmembrane potential (_m) is a primary event leading to release of cytochrome c into the cytoplasm (17, 28). Release of cytochrome c is necessary for the activation of the initiator caspase-9, which occurs only in the presence of ATP (34, 37, 46). Collectively, these findings highlight the role of pH, mitochondria, and ATP supply in apoptotic cell death pathways.

We recently demonstrated that extracellular acidosis attenuates the loss of viability of human pulmonary artery endothelial cells (HPAEC) after exposure to a metabolic insult (7). On the basis of this previous work in a necrotic cell death model, we postulated that similar changes in extracellular pH or altered mitochondrial function would modify the activation of effector caspases, oligonucleosomal DNA fragmentation, and the progression to apoptosis in HPAEC. Specifically, we hypothesized that a reduction in extracellular pH or inhibition of the mitochondrial ATP pump would inhibit key events in the apoptotic pathway after exposure to a well-defined cell death stimulus. Little is known about the regulation of apoptosis pathways in endothelial cells. We used staurosporine (STS), a well-described inducer of apoptosis in mammalian cells (5), to accomplish the following objectives: 1) to determine the effect of changes in extracellular pH on caspase-3 activation, DNA fragmentation, cell morphology, and cytosolic free Ca²⁺ concentration after STS treatment and 2) to determine the effect of altered mitochondrial function and ATP supply on the development of apoptosis after STS treatment in this cell type.

	METHODS

TOP ABSTRACT INTRODUCTION METHODS RESULTS DISCUSSION REFERENCES

Endothelial cell culture. HPAEC were obtained from a commercial vendor (Clonetics) and grown in 10-cm² plates containing endothelial growth medium supplemented with 10% fetal bovine serum, as previously described (7). The cells were maintained in a humidified atmosphere (21% O₂ and 5% CO₂ at 37°C), fed endothelial growth medium containing 10% fetal bovine serum twice a week, and passaged when confluent (3-5 days). In preparation for the measurement of caspase activity, cells were seeded onto 48-well tissue culture plates incubated in a humidified atmosphere (21% O₂ and 5% CO₂ at 37°C) until ~90-95% confluence was reached. Alternatively, monolayers were seeded onto glass slides for experiments involving fluorescent imaging. All experiments were performed with monolayers matched for cell line, passage number, time to confluence, and cell density between experimental groups. No differences were observed in the results when different cell lines or passage numbers were used.

Experimental protocols. We investigated the effect of acidification by incubating monolayers for various time periods in MEM set to different pH values (7.4, 7.0, and 6.6) in the presence or absence of 1 µM STS. This treatment was followed by measurement of caspase activity or DNA fragmentation or determination of cell morphology and the percentage of apoptotic cells with fluorescent imaging. Imaging experiments involving the measurement of intracellular Ca²⁺ concentration are described in Fluorescent microscopy measurement of cytosolic free Ca²⁺ concentration. All experiments were conducted using serum-free HEPES-buffered MEM set to specific pH values for the duration of the experiment. In separate experiments, we investigated the level of caspase-3 activity under conditions of ATP depletion by incubating the monolayers with 10 µM oligomycin, an inhibitor of the mitochondrial F₀F₁-ATPase. Experiments were performed at least in triplicate in 48-well plates and repeated on three separate occasions.

Measurement of caspase activity. The assay is based on the ability of caspase-3 to hydrolyze a specific substrate linked to a fluorescent probe, rhodamine 110. Under these conditions, the substrate linked to rhodamine 110 is nonfluorescent (Z-DEVD-rhodamine 110). Cleavage of the caspase substrate from the probe leads to the generation of a measurable fluorescent signal (21, 51). Under these conditions, the magnitude of the fluorescent signal is directly proportional to the amount of caspase-3 activity in the sample. Although the acetyl-Asp-Glu-Val-Asp-CHO substrate is primarily cleaved by caspase-3, there is some cross-reactivity with other members of the caspase family (caspase-7). Therefore, this assay is a measure of "caspase-3-like" activity.

Gel electrophoresis visualization of DNA fragmentation. We quantified the amount of DNA digested into 180- to 200-kb base pairs (percent DNA fragmentation) after STS treatment, as previously described (44). Briefly, 1 × 10⁶ cells were washed twice with ice-cold phosphate-buffered saline (PBS) and placed on ice with 0.4 ml of hypotonic buffer (10 mM Tris, pH 7.5, 1 mM EDTA, and 0.2% Triton) for 15 min. The cells were then centrifuged at 13,000 g at 4°C for 20 min. The pellets containing the high-molecular-weight DNA were incubated with 0.4 ml of 10 mM EDTA, 50 mM Tris, 1% SDS, and 0.5 µg/ml proteinase K and incubated at 48°C for 18 h. The supernatant (centrifugation-resistant fraction) containing low-molecular-weight DNA was incubated with RNase A (20 µg/ml) at 37°C for 1 h. Low- and high-molecular-weight DNA were then extracted with phenol-chloroform and precipitated with 2.5 vol of ethanol. Samples were resuspended in water containing loading buffer (2.5% Ficoll, 0.025% bromphenol blue, and 0.025% xylene cyanol) and electrophoresed in a 2% agarose gel at 40 V in Tris-borate-EDTA. DNA was visualized by ultraviolet examination after incubation for 30 min with the fluorescent dye Sybergold (Molecular Probes) at a dilution of 1:10,000. A Polaroid camera was used to photograph the gel. The amount of DNA obtained from each extraction was quantified by comparison with a calibration curve constructed from known standard quantities of DNA using a fluorescent nucleic acid staining kit (PicoGreen, Molecular Probes). The protocol for this assay was obtained from the manufacturer. The samples were read on a microplate reader with the wavelength settings set to read at the optimal wavelengths for PicoGreen (480-nm excitation and 520-nm emission). The percent DNA fragmentation was calculated as percentage of total DNA (supernatant + pellet) recovered as low-molecular-weight DNA in the supernatant.

Fluorescent microscopy measurement of cell viability. Nuclear morphology was assessed using an inverted microscope (Olympus IX70) equipped with a xenon light source (75 W). After an experimental treatment, monolayers grown on glass slides or in six-well tissue culture plates were incubated for 5-10 min at 37°C with Hoechst-33342 (final concentration 10 µg/ml) and propidium iodide (PI, final concentration 20 µg/ml) (29, 36). Hoechst-stained cells were visualized under epifluorescence illumination using a 360-nm excitation and 510-nm barrier filter. PI-stained cells were viewed using a 540-nm excitation and 510-nm barrier filter. Cells were counted in five ×20 fields per well. A minimum of 200 cells were counted for each condition in each experiment. Brightly fluorescent-staining nuclei showing a highly condensed mass of chromatin or lobulated condensed chromatin typical of apoptotic bodies were scored as apoptotic cells compared with more diffusely stained normal nuclei. Cells were scored as necrotic on the basis of the absence of these typical apoptotic nuclear changes and evidence of PI uptake, indicating loss of plasma membrane integrity. In experiments using the intracellular Ca²⁺ chelator 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA)-AM, monolayers were pretreated with 10 µM BAPTA-AM for 1 h before STS treatment. The Ca²⁺ chelator was loaded into cells in the presence of 64 µM Pluronic 127, as previously described (40).

Measurement of cellular ATP content. ATP content in control and treated monolayers was determined using the CellTiter-Glo luminescent assay (Promega) according to the manufacturer's instructions. This assay produces a luminescent signal through the mono-oxygenation of luciferin catalyzed by luciferase in the presence of Mg²⁺, ATP, and molecular oxygen to form oxyluciferin, AMP, inorganic phosphate, and carbon dioxide, and a luminescent signal that is proportional to the amount of ATP present is generated. HPAEC monolayers grown to ~90% confluence in 96-well plates were incubated in pH 7.4 MEM at 37°C under the following conditions: control ± glucose, control with oligomycin ± glucose, STS ± glucose, and STS with oligomycin ± glucose for 6 h. After the plate was allowed to equilibrate to room temperature for 30 min, background luminescence was determined on a microplate spectrofluorometer (SPECTRAMax Gemini XS, Molecular Devices) using the luminescent measurement mode. Next, a volume of CellTiter-Glo reagent equivalent to the volume of each well of the 96-well plate was added to each well. The plate was mixed for 2 min and allowed to incubate at room temperature for 10 min before measurement of the luminescent signal in the microplate spectrofluorometer set for an end-point reading. The emission fluorescent signal was captured over the full range of the instrument (250-850 nm). Each experimental condition was run in triplicate; n = 1 represents the average of these triplicate readings. Background luminescence was subtracted from each signal obtained from each well before final data analysis. Results are expressed as the percentage of the signal obtained from the control monolayers in MEM containing glucose.

Fluorescent microscopy measurement of cytosolic free Ca²+ concentration. Cytosolic free Ca²⁺ concentration was monitored in monolayers using fluo 3-AM (Molecular Probes), as previously described (50). In preliminary experiments, we optimized the dose, image exposure time, and other experimental variables noted below. Nearly confluent monolayers plated in 24-well tissue culture plates were incubated for 15 min at room temperature (27°C) in HEPES-buffered MEM containing 2 µM fluo 3 in the presence of 64 µM Pluronic 127 and 2.5 mM probenecid to maximize uptake and to inhibit efflux of the probe, respectively. The monolayers were then incubated for an additional 20 min in the same medium at 37°C to increase hydrolysis of the ester portion of the probe in the cytosol. Monolayers were assessed by light microscopy before and after probe loading to ensure normal morphology and the absence of toxicity. Monolayers were positioned on the stage of an inverted microscope (Olympus IX70), where baseline images were obtained before initiation of the experiment by the addition of reagents to the tissue culture wells. Images were obtained using separate populations of cells at the different time points (10 s and 30 and 60 min) to avoid potential problems related to photobleaching or loss of viability in the treated cells with repeated imaging over time. Images were acquired after a change of the medium with the addition of medium containing STS at a specified pH or the addition of a similar volume of normal medium to control monolayers. Images were obtained using a 4-s exposure time with the following light filters: 495 ± 25 nm excitation and 540 ± 25 nm emission on the same imaging equipment used for the viability studies (20× objective, xenon light source, 150 W). No significant change in background extracellular fluorescence or loss of intracellular signal strength was noted during experiments lasting up to 60 min in control monolayers, suggesting little probe leakage or photobleaching, respectively, under these conditions. Fluo 3 is a nonratiometric probe that is weakly fluorescent when not bound to Ca²⁺. The signal strength of the probe, which represents an indirect measurement of intracellular Ca²⁺ concentration, was quantitated at the different time points using imaging software (MetaMorph, Universal Imaging, West Chester, PA). The experiments designed to investigate the effect of altered extracellular pH on intracellular Ca²⁺ concentration after STS treatment were confined to extracellular pH 7.0-7.4 to avoid complications related to the effect of altered pH on probe signal strength (33).

Data analysis. Values for the caspase assay are means ± SE expressed in arbitrary RFU. DNA fragmentation values are means ± SE expressed as the percentage of total DNA recovered as low-molecular-weight DNA in the supernatant from separate experiments. Differences between treatment groups for caspase activity, DNA fragmentation, and ATP content were analyzed with an analysis of variance. Mean values were then compared with a post hoc multiple comparison procedure (Bonferroni). Data for ATP content after different treatments are presented as the percentage of the ATP content of control monolayers incubated in media containing glucose. In the experiments involving imaging of Ca²⁺ uptake with fluo 3 after STS treatment, intracellular probe signal strength was quantified in RFU for comparison among different experimental conditions at a given time point. This procedure involved using 4-6 microscopic fields, each containing 60-75 cells/field, for each condition. Differences between means were considered significant if P < 0.01 or at the level appropriate for the number of comparisons of interest in each experiment.

Materials. HPAEC and cell culture reagents, including MEM, HEPES, 0.5% trypsin, and EDTA, were purchased from Clonetics (San Diego, CA). STS, Trizma base, boric acid, probenecid, and agarose were obtained from Sigma (St. Louis, MO). The molecular weight DNA standard was obtained from Roche (Mannheim, Germany). Fluoroprobes (Hoechst-33342, PI, fluo 3, Sybergold, and PicoGreen) and the caspase assay kit (EnzChek caspase-3 assay kit, E-13184) were obtained from Molecular Probes (Eugene, OR). The caspase-3 inhibitor was a component of the caspase assay kit. The pancaspase inhibitor N-benzoylcarbonyl-Val-Ala-Asp-fluoromethyl ketone (Z-VAD-fmk) was obtained from Biomol (Plymouth Meeting, PA). The ATP assay kit (CellTiter-Glo luminescent assay) was obtained from Promega.

	RESULTS

TOP ABSTRACT INTRODUCTION METHODS RESULTS DISCUSSION REFERENCES

We induced apoptosis in HPAEC by incubating them with 0.1 and 1 µM STS in HEPES-buffered MEM at pH 7.4 for 6 and 24 h, respectively. In accord with previous reports in other cell types (5), STS demonstrated a time-dependent pattern of caspase-3 activation in HPAEC (Fig. 1). Caspase-3 activity was detected as early as 2 h after STS treatment (data not shown) and continued to increase up to 6 h after treatment. Caspase activity consistently reached a peak level after treatment with 1 µM STS for 6 h compared with the level observed in untreated control monolayers. A significant increase in caspase-3 activity was also observed with the lower dose of STS (0.1 µM), suggesting a dose-related pattern of caspase activation (data not shown). Caspase activity was markedly reduced at 24 h after treatment with 1 µM STS, suggesting that the time course for induction of apoptosis in HPAEC is relatively rapid and complete at 24 h after STS exposure. Incubation of monolayers with 1 µM STS for 6 h followed by addition of a specific caspase-3 inhibitor, acetyl-Asp-Glu-Val-Asp-CHO (5 µM), at the start of the assay completely inhibited the caspase activation after STS treatment. These results demonstrate the specificity of the fluorogenic assay for activation of caspase-3 after STS exposure.

View larger version (13K): [in this window] [in a new window]   Fig. 1.   Time-dependent caspase-3 activation in human pulmonary artery endothelial cells after staurosporine (STS) treatment. Caspase-3 activity was monitored in pH 7.4 medium under the following conditions: untreated monolayers incubated in MEM for 6 h (control), monolayers incubated with 1 µM STS in the presence of caspase-3 inhibitor acetyl-Asp-Glu-Val-Asp-CHO (5 µM; STS 1 w/Inhib), monolayers incubated with 1 µM STS for 24 h (STS 1, 24H), and monolayers exposed to 1 µM STS for 6 h (STS 1, 6H). Values are means ± SE expressed in relative fluorescent units (RFU); n = 3. * P < 0.0001 vs. control; ** P < 0.0001 vs. STS 1, 6H.

In the next set of experiments, we examined the effect of altering extracellular pH on caspase-3 activation after STS treatment (Fig. 2). Control monolayers incubated for 6 h in acidic media (pH 7.0 or 6.6) demonstrated the same level of caspase-3 activity as control monolayers incubated for the same time period in pH 7.4 medium. Inspection of these monolayers using phase-contrast microscopy after the 6-h incubation demonstrated no significant difference in the appearance of monolayers incubated in pH 7.4 vs. 7.0 or 6.6 medium. All monolayers were intact with a normal appearance, demonstrating that a reduction in medium pH had no overt effect on monolayer morphology and viability during this treatment period. Monolayers incubated in 1 µM STS for 6 h in pH 7.4 medium demonstrated a two- to threefold increase in caspase activity compared with control monolayers in pH 7.4 medium. The magnitude of caspase-3 activation after STS treatment was dependent on the pH of the medium. There was a progressive decrease in the magnitude of caspase activation after incubation in 1 µM STS for 6 h in pH 7.0 and 6.6 media, respectively. The level of caspase activation observed in pH 6.6 medium was not significantly different from that observed in the control monolayers in pH 6.6 and 7.4 media. Maximal caspase activation was consistently observed in pH 7.4 medium after STS exposure.

View larger version (18K): [in this window] [in a new window]   Fig. 2.   Effect of medium acidification on STS-induced caspase-3 activation. Caspase-3 activity was measured after 6 h of STS incubation under the following conditions: untreated monolayers incubated in pH 7.4 medium (control pH 7.4), monolayers treated with 1 µM STS in pH 7.4 medium (STS pH 7.4), untreated monolayers incubated in pH 7.0 medium (control pH 7.0), monolayers treated with 1 µM STS in pH 7.0 medium (STS pH 7.0), untreated monolayers incubated in pH 6.6 medium (control pH 6.6), and monolayers treated with 1 µM STS in pH 6.6 medium (STS pH 6.6). Values are means ± SE expressed in RFU; n = 3. * P < 0.001 vs. STS pH 7.4. # P < 0.001 vs. control at the same pH.

The effect of STS treatment on DNA ladder formation and percent DNA fragmentation in media set to different pH values is illustrated in Fig. 3 and Table 1. A small degree of DNA fragmentation occurred in control monolayers incubated in pH 7.4, 7.0, or 6.6 medium for 6 h, suggesting a minimal degree of apoptosis under these conditions (Table 1). In contrast, incubation with 1 µM STS was associated with the appearance of visible oligonucleosomal fragments of DNA after only 2 h of incubation with STS at all medium pH values (data not shown). A sample gel electrophoresis of DNA extracted from monolayers after 6 h of exposure to 1 µM STS in media set to different pH values is shown in Fig. 3. After 6 h of treatment with 1 µM STS in pH 7.4 medium, a typical DNA ladder consistent with apoptosis is clearly visible. The peak in caspase-3 activity after 6 h of STS treatment (Figs. 1 and 2) corresponded closely to this same time point at which maximal DNA fragmentation was observed. Lowering of medium pH progressively lowered the percentage of DNA recovered in the low-molecular-weight DNA fraction after STS treatment compared with samples incubated in pH 7.4 medium. A summary of the effect of media acidification on the magnitude of DNA fragmentation is shown in Table 1. These data represent the averaged results of all gel electrophoresis experiments performed on monolayers incubated with STS at different medium pH values. These results confirm the same pH-dependent pattern of DNA fragmentation as observed with caspase-3 activation (Fig. 2). The percentage of DNA fragmentation was maximal in pH 7.4 medium (37.9%) and demonstrated a progressive reduction in magnitude with each decrease in medium pH. Pretreatment with the irreversible caspase inhibitor Z-VAD-fmk (10 µM) abolished the DNA fragmentation that occurred after STS treatment at all medium pH values (Fig. 4), demonstrating the link between caspase activation and DNA fragmentation under these conditions.

View larger version (128K): [in this window] [in a new window]   Fig. 3.   Effect of medium acidification on DNA fragmentation after STS treatment. High- and low-molecular-weight DNA from monolayers incubated for 6 h were subjected to gel electrophoresis. M, molecular weight marker; lanes 1 and 2, high- and low-molecular-weight DNA of control monolayers in pH 7.4 medium; lanes 3, 5, and 7, high-molecular-weight DNA of monolayers treated with 1 µM STS in pH 7.4, 7.0, and 6.6 media, respectively; lanes 4, 6, and 8, low-molecular-weight DNA of monolayers treated with 1 µM STS in pH 7.4, 7.0, and 6.6 media, respectively. DNA fragmentation is expressed as percentage of total DNA recovered as low-molecular-weight DNA.

View larger version (137K): [in this window] [in a new window]   Fig. 4.   Effect of N-benzoylcarbonyl-Val-Ala-Asp-fluoromethyl ketone (Z-VAD-fmk) on percent DNA fragmentation after STS treatment in media with different pH values. High- and low-molecular-weight DNA from monolayers incubated for 6 h were subjected to gel electrophoresis. M, molecular weight marker; lanes 1, 3, and 5, high-molecular-weight DNA of monolayers treated with 1 µM STS + 10 µM Z-VAD-fmk in pH 7.4, 7.0, and 6.6 media, respectively; lanes 2, 4, and 6, low-molecular-weight DNA of monolayers treated with 1 µM STS + 10 µM Z-VAD-fmk in pH 7.4, 7.0, and 6.6 media, respectively. DNA fragmentation is expressed as percentage of total DNA recovered as low-molecular-weight DNA (see METHODS).

We used monolayers loaded with both Hoechst-33342 and PI to determine the effect of STS treatment at different extracellular pH values on cell morphology and the percentage of apoptotic cells (Fig. 5). STS-treated monolayers in pH 7.4-6.6 media demonstrated a larger number of cells with the morphological features characteristic of apoptosis than did control monolayers (Fig. 5A). None of these STS-treated monolayers demonstrated PI uptake (data not shown). The largest percentage (40%) of apoptotic cells was observed after STS treatment in pH 7.4 medium. These data demonstrate that the primary mode of cell death under these conditions was apoptosis. There was a progressive decrease in the percentage of apoptotic cells with each decrease in medium pH in the STS-treated cells; only 9% of cells demonstrated apoptotic cell morphology at pH 6.6 with no evidence of necrosis (Fig. 5B). These results are similar to the progressive decrease in the magnitude of caspase activation and percent DNA fragmentation observed after STS treatment in media set to different pH values (Fig. 2, Table 1).

View larger version (39K): [in this window] [in a new window]   Fig. 5.   Effect of medium acidification on percentage of apoptotic cells after STS treatment. A: percentage of cells showing apoptotic morphology was determined after 6 h of incubation as follows: untreated monolayers in pH 7.4 MEM (control pH 7.4, B), monolayers incubated with 1 µM STS in pH 7.4 medium (STS pH 7.4, C), monolayers incubated with 1 µM STS in pH 7.0 medium (STS pH 7.0, D), and monolayers incubated with 1 µM STS in pH 6.6 medium (STS pH 6.6, E). Arrows in C-E indicate apoptotic cells. Values are means ± SE; n = 3. * P < 0.01 vs. control pH 7.4; # P < 0.01 vs. STS pH 7.4.

Pretreatment of monolayers for 1 h with the intracellular Ca²⁺ chelator BAPTA-AM (10 µM) inhibited STS-induced apoptosis, compared with treatment with STS alone in pH 7.4 medium (Table 2). The Ca²⁺ chelator had no effect on viability of control monolayers over the same time period. This experiment was repeated several times (n = 3) with similar results. These results suggest an important role for Ca²⁺-related signaling events, such as a rise in intracellular Ca²⁺ concentration, in this cell death pathway in HPAEC. Therefore, in a related set of experiments, we determined the effect of a change in extracellular medium pH on the rise in intracellular Ca²⁺ concentration after STS treatment. STS treatment in pH 7.4 medium led to a sustained increase in intracellular Ca²⁺ concentration in cells loaded with fluo 3, indicated by an increase in the magnitude of the fluo 3 signal after addition of STS to the media. The increase in intracellular Ca²⁺ concentration above the level observed in control monolayers was observed at 30 and 60 min after STS treatment (Table 3). Untreated control monolayers incubated in pH 7.4 or 7.0 medium demonstrated no significant change in probe signal strength over the same time periods. The difference in the magnitude of the probe signal strength in the control and STS-treated cells at 60 min (vs. 30 min) most probably reflects increased loading of probe into the cells under these conditions. STS treatment in pH 7.0 medium was associated with a smaller increase in the magnitude of the fluo 3 signal over these same time intervals compared with STS treatment in pH 7.4 medium, indicating a smaller increase in intracellular Ca²⁺ concentration under these conditions (Table 3). These experiments were repeated several times (n = 3) at each time point with similar qualitative results.

In a final set of experiments, we determined the effect of altering mitochondrial function and the bioenergetic state of endothelial cells on caspase activation, DNA fragmentation, and percentage of apoptotic cells after STS treatment. Monolayers were first pretreated for 1 h with oligomycin (10 µM), an inhibitor of the mitochondrial F₀F₁-ATPase, in glucose-free medium at pH 7.4, and then with STS in glucose-free pH 7.4 medium for 6 h. Under these conditions, STS treatment failed to produce an increase in caspase-3 activity above the level observed in untreated control monolayers incubated in pH 7.4 MEM alone (Fig. 6). In addition, the typical pattern of DNA fragmentation consistent with apoptosis that was observed after STS treatment alone was not present in monolayers pretreated with oligomycin followed by treatment with STS in glucose-free medium (data not shown). Furthermore, in monolayers pretreated in this fashion, necrosis, rather than apoptosis, was the major mode of cell death (Figs. 7 and 8). This conclusion is based on several observations. First, consistent with the results related to the absence of caspase activation under these conditions (Fig. 6), the fluorescent imaging experiments confirmed that these monolayers did not demonstrate the typical morphological features of apoptosis (chromatin condensation) that were observed with STS treatment alone (Fig. 5B). Second, the majority of the cells (90%) under these conditions demonstrated PI uptake, in contrast to the absence of PI uptake observed after treatment with STS alone (Figs. 7 and 8).

View larger version (16K): [in this window] [in a new window]   Fig. 6.   Effect of inhibition of mitochondrial F0F1-ATPase on caspase-3 activation after STS treatment. Caspase-3 activity was measured in monolayers incubated for 6 h in pH 7.4 medium as follows: untreated monolayers (control), monolayers incubated with 1 µM STS (STS), and monolayers incubated with 1 µM STS + 10 µM oligomycin in glucose-free medium (STS + O + w/o Glu). Values are means ± SE; n = 3. * P < 0.001 vs. control; # P < 0.001 vs. STS.

View larger version (81K): [in this window] [in a new window]   Fig. 7.   Effect of inhibition of mitochondrial F0F1-ATPase on cell morphology and mode of cell death after STS treatment. Cell morphology of monolayers stained with Hoechst-33342 and propidium iodide (PI) after 6 h of incubation in pH 7.4 medium was assessed as follows: untreated control monolayers (control, A), monolayers pretreated with 10 µM oligomycin for 1 h in pH 7.4 medium (control + O, B), monolayers incubated with 1 µM STS (STS, C), and monolayers pretreated with 10 µM oligomycin for 1 h in pH 7.4 medium before treatment with 1 µM STS (STS + O, D). B-D: thick arrows, apoptotic cells; thin arrows, necrotic cells staining positive for PI.

View larger version (15K): [in this window] [in a new window]   Fig. 8.   Percentage of apoptotic and necrotic cells after STS treatment in the presence or absence of oligomycin. Percentage of apoptotic or necrotic cells was determined using concurrent staining with Hoechst-33342 and PI in monolayers incubated in pH 7.4 medium under the following conditions: untreated monolayers (control), control monolayers pretreated with 10 µM oligomycin for 1 h (C + O), monolayers incubated with 1 µM STS for 6 h (STS), and monolayers pretreated with 10 µM oligomycin for 1 h before treatment with 1 µM STS (STS + O). Values are means ± SE; n = 3. * P < 0.0001 vs. STS; # P < 0.0001 vs. C + O.

To further investigate the role of mitochondria and ATP supply in the loss of viability after STS treatment, we performed a final set of experiments in which we correlated the mode of cell death with the cellular ATP concentration. These results are illustrated in Table 4. There was no significant loss of viability or difference in the ATP concentration between untreated control monolayers incubated in media with or without glucose for 6 h. Pretreatment with oligomycin for 1 h before the start of the 6-h incubation period in normal medium also did not significantly alter viability or ATP content compared with control monolayers. In contrast, pretreatment with oligomycin in medium without glucose followed by 6 h of incubation in medium without glucose produced a marked decrease in ATP content to ~12% of the control level in association with a significant loss of viability (Table 4). The mode of cell death under these conditions was necrotic on the basis of the absence of the typical morphological features of apoptosis in conjunction with PI uptake, indicating loss of membrane integrity.

STS-treated monolayers demonstrated a decrease in ATP content to ~84% of controls in association with induction of apoptosis at the 6-h time point (Table 4). The omission of glucose from the medium during STS treatment resulted in a further decrease in ATP content to ~80% of controls but no change in the percentage of apoptotic cells or the mode of cell death under these conditions. Pretreatment with oligomycin, as described above, followed by STS treatment for 6 h in normal medium, resulted in a further decrease in ATP content to ~54% of control, but again there was no change in the percentage of apoptotic cells or the mode of cell death. In contrast, omission of glucose in the medium during pretreatment with oligomycin, followed by STS treatment in glucose-free medium, shifted the primary mode of cell death from apoptosis to necrosis (Table 4). The ATP content of these monolayers was markedly decreased to ~2% of controls. These experiments were repeated several times (n = 3) with similar results.

	DISCUSSION

TOP ABSTRACT INTRODUCTION METHODS RESULTS DISCUSSION REFERENCES

To our knowledge, the present results are the first to elucidate the effects of altered extracellular pH, mitochondrial function, and ATP supply on caspase activation, DNA fragmentation, and cell morphology in HPAEC after exposure to a well-known inducer of apoptosis in mammalian cells. Apoptosis is rapid and complete within 24 h after STS exposure in HPAEC. Inhibition of caspase-3 activity by Ac-DVD-CHO and complete inhibition of DNA fragmentation by the pancaspase inhibitor Z-VAD-fmk demonstrate that caspase-dependent signaling events mediate STS-induced cell death in this cell line. Extracellular acidosis inhibited caspase-3 activation, DNA fragmentation, and development of apoptotic cell morphology after STS exposure. Maximal caspase-3 activation and DNA fragmentation and the largest number of apoptotic cells were always observed after STS treatment in pH 7.4 medium. These are typically "late" events in the apoptotic signal transduction pathway, suggesting the presence of one or more upstream pH-dependent steps in this model. Inhibition of the STS-induced rise in intracellular Ca²⁺ concentration by a reduction in extracellular pH suggests that altered Ca²⁺ signaling may be involved. Similarly, inhibition of mitochondrial function and the bioenergetic state (ATP supply) of endothelial cells inhibited the development of apoptosis after STS exposure. The prevalent mode of cell death under these conditions was shifted from apoptosis to necrosis. These data suggest a central role for mitochondria and ATP supply in this model of endothelial cell apoptosis.

Recent reports suggest that apoptosis has a role in pathological conditions associated with endothelial dysfunction (11, 14, 47). In contrast, many of the details of apoptotic signaling in endothelial cells are not well defined. Cells undergoing apoptosis are defined by characteristic morphological features, consisting of chromatin condensation, blebbing of plasma membrane, formation of apoptotic bodies, and generation of oligonucleosomal DNA fragments (17, 41). These features represent the end result of specific events in the apoptotic pathway. An important central event in apoptotic signal transduction is the activation of a family of cysteine proteases, related to interleukin-1-converting enzyme, termed caspases (41). Caspases cleave a variety of target proteins, leading to disruption of crucial cellular processes and structural components (41, 48), but the initial triggering event(s) is poorly defined.

We utilized STS as a proapoptotic stimulus in HPAEC to investigate the effect of altered extracellular pH and mitochondrial function on apoptotic signaling for several reasons. STS is a well-described inducer of apoptosis in many cell lines (5). This widely accepted model of apoptosis includes many components of the apoptosis signaling cascade (5, 23). All mammalian cells constitutively express the proteins required to execute the STS-induced cell death pathway. Second, we recently demonstrated that extracellular acidosis attenuates the loss of viability in HPAEC after exposure to a simulated "ischemic" insult (metabolic inhibition). This type of insult at neutral extracellular pH (pH 7.4) led to a loss of viability that was primarily a necrotic form of cell death (7). Acidosis caused by ATP hydrolysis during anaerobic metabolism is a feature of tissue ischemia and cell injury. The conventional view is that acidosis contributes to diminished cell viability. However, a growing body of evidence suggests that acidosis may actually have a protective effect on cells or organ systems in the setting of anoxia, ischemia, or ischemia-reperfusion injury (1, 36, 43). Therefore, we postulated that changes in extracellular pH might also modify apoptotic signaling events in endothelial cells. The present results extend our previous findings by demonstrating that extracellular acidosis also inhibits a well-defined apoptotic cell death pathway in HPAEC.

The effect of altered pH on apoptotic signal transduction events is controversial. Apoptosis has been reported to be associated with intracellular acidification in some models of apoptosis. For example, a drop in intracellular pH has been observed in interleukin-2-dependent cytotoxic T lymphocytes (38), in Jurkat T lymphoblasts after cycloheximide treatment or Fas ligation (15), and in HL-60 cells after treatment with the proapoptotic agent etoposide (2). In contrast, other investigators have demonstrated that a low extracellular pH inhibits PARP cleavage, DNA fragmentation, and apoptosis in different models of apoptosis, including STS (8, 45, 53). These findings suggest that the relationship between altered pH and specific events in the apoptotic signaling cascade may be cell type dependent.

A reduction in extracellular pH inhibited caspase-3 activity and DNA fragmentation and significantly reduced the number of apoptotic cells after STS exposure. These findings are in agreement with previous work demonstrating that a low extracellular pH inhibited DNA fragmentation, PARP cleavage, and chromatin condensation in STS- and etoposide-induced apoptosis in the ML-1 cell line (45), suggesting the presence of pH-sensitive step(s) upstream of caspase-3 activation. An inhibitory effect of low extracellular pH has also been reported in neuronal cells and fibroblasts using a serum deprivation-induced model of apoptosis (53). Protection against apoptotic cell death by acidosis was also observed in bovine aortic endothelial cells (8), but under experimental conditions very different from those in the present study. In this study, the endothelial cells were kept in serum-free medium and exposed to hypercarbic acidosis (pH 7.0) over a prolonged time period (7 days). Moreover, prolonged exposure to hypercarbic acidosis in bovine aortic endothelial cells induced an increase in vascular endothelial growth factor and basic fibroblast growth factor mRNA expression and secretion (8). Therefore, the protective effect of acidosis on apoptosis in this latter study may have been related to the inhibitory effect of vascular endothelial growth factor and basic fibroblast growth factor on apoptotic signal transduction, as previously demonstrated (18, 19).

In contrast to the present results, a low intracellular pH is associated with apoptosis in some cell lines (12, 16, 52). For example, a low pH stimulates caspase activity in BAF3 cells (12). However, in this latter study, the cells were incubated with nigericin at a low extracellular pH. Nigericin is an ionophore that is capable of altering _m and, thereby, inducing release of cytochrome c, potentially leading to caspase activation. Therefore, these results do not clearly demonstrate a direct relationship between caspase activation and low pH. Moreover, several reports have found that the permeable pancaspase inhibitor Z-VAD-fmk prevented the decrease in intracellular pH associated with apoptosis, suggesting that intracellular acidification is downstream of caspase-3 activation (52). In addition, Z-VAD-fmk inhibits initiator and effector caspase activity. Therefore, this inhibitor cannot be used to establish which caspase is sensitive to variations in pH. Thus, although previous reports suggested that acidification is a feature of programmed cell death in some cell lines, additional work is required to determine the precise role of altered pH on apoptotic signaling events in different cell lines. The present results demonstrate that pH 7.0 inhibits chemically induced apoptosis in a specific endothelial cell line, suggesting that the effects of altered extracellular pH may be cell type dependent.

The mechanism(s) by which acidosis modulates apoptotic signal transduction has not been clearly defined. A low extracellular pH inhibits several aspects of apoptotic signaling, such as PARP cleavage and DNA fragmentation, in different models of apoptosis, including STS (8, 45, 53). Other possible pH-sensitive loci have received little attention. Previous work has demonstrated that pH 7.4 is optimal for caspase-3 activity. Changes in pH might alter caspase-3 activity via a direct effect on enzyme activity or, indirectly, via an effect on one or more caspase substrates (48). The present results suggest that one potential site for the effect of acidosis is at the level of caspase-3 activation or upstream of caspase-3 in the signaling cascade in HPAEC, as recently demonstrated for other cell lines (45). The details of how altered pH modulates caspase-3 activity require additional work.

Other components of the STS cell death pathway may be pH dependent. Changes in intracellular Ca²⁺ concentration are involved in signal transduction in apoptotic and necrotic cell death pathways in many cell types. For example, in several models of ischemia, the cytoprotective effect of acidosis has been attributed to a decrease in Ca²⁺ influx in neuronal cells and cardiomyocytes (1). The opposite relationship was observed in chick (27) and rat cardiomyocytes (13). The role of intracellular Ca²⁺ concentration in apoptotic signaling is similarly complex. A rise in intracellular Ca²⁺ concentration triggers the apoptotic cascade in some cell types (30, 39, 40), but this is not a universal phenomenon (2, 26, 31). Little is known about the role of pH in modifying Ca²⁺ signaling events in apoptosis in endothelial cells.

In experiments designed to test the role of changes in intracellular Ca²⁺ concentration in the cell death pathway, we found that chelation of intracellular Ca²⁺ inhibited STS-induced apoptosis in HPAEC, suggesting that an increase in intracellular Ca²⁺ concentration is an important signaling event in the cell death pathway, as noted in other cell types (30, 39, 40). The STS-induced rise in intracellular Ca²⁺ concentration in the present experiments was sustained (Table 3), as previously noted (30, 40). The increase in intracellular Ca²⁺ concentration occurred before any biochemical, molecular, or morphological evidence of apoptosis (caspase activation, DNA fragmentation, or chromatin condensation) could be detected, suggesting that this rise in intracellular Ca²⁺ concentration is an early event in the cell death pathway. The decrease in the STS-induced rise in intracellular Ca²⁺ concentration in acidic medium (Table 3) demonstrates that a reduction in extracellular pH modifies Ca²⁺ signaling events in this cell death pathway. These findings suggest that one mechanism for the inhibitory effect of acidosis on STS-induced apoptosis may involve altered Ca²⁺ signaling, as noted in other cell types under different circumstances (49).

Alterations in cell volume or intracellular ion homeostasis could potentially contribute to the inhibitory effect of acidosis on STS-induced apoptosis. A decrease in extracellular pH could induce a change in volume (cell swelling), which might modify signaling events. Recent work found that a degree of acidosis (pH 6.6) similar to that in the present experiments induced only small changes (3-4%) in endothelial cell volume in HEPES-buffered medium (3). These small changes in cell volume are unlikely to have a major effect on this cell death pathway.

Additional mechanisms that might also contribute to the inhibitory effect of acidosis deserve mention. These mechanisms focus on other possible pH-sensitive steps in the pathway, including mitochondrial components, such as cytochrome c release, PARP cleavage, or opening of the mitochondrial permeability transition pore (4, 43), protonation of undefined enzymes or substrates of the signaling cascade leading to allosteric effects on protein conformation and the activity of these molecules (42), changes in the degree of phosphorylation of key substrates (22), or direct effects of altered pH on gene expression of pathway components (20, 32). Thus H⁺ may be exerting an effect on apoptotic signaling at more than one locus. To our knowledge, these possibilities have not been actively investigated in endothelial cells. In addition, cell type and/or experimental model-based differences in responses limit broad generalizations about the effect of pH on apoptotic signaling at this time (25, 45). These findings highlight the need for additional studies designed to define the pH-dependent step(s) in apoptotic signaling in different cell types.

Mitochondria play a central role in apoptotic signal transduction in many cell lines (28, 54). Disruption of _m is followed by release of cytochrome c into the cytosol. In the presence of ATP, cytochrome c triggers complex formation between caspase-9 and Apaf-1, leading to the activation of caspase-3 (37). Chromatin condensation, nuclear fragmentation, and transport of larger molecules across the nuclear membrane are also events that are ATP dependent (24, 34, 46). These findings suggest that apoptosis is an active energy-dependent process that requires functioning mitochondria and the availability of a critical concentration of ATP (9, 34, 46). Depletion of ATP by >50% in Jurkat cells before exposure to an apoptotic stimulus shifted the mode of cell death from apoptosis to necrosis (34). In oxidant-stressed human umbilical vein endothelial cells, a threshold ATP concentration of 25% of the basal level was required for induction of apoptosis (35).

In agreement with these latter findings, our results demonstrate that inhibition of the mitochondrial F₀F₁-ATPase (oligomycin) in glucose-free medium not only inhibited caspase-3 activation and DNA fragmentation after STS exposure but shifted the cell death pathway from apoptosis to necrosis (Fig. 6, Table 4). These results suggest that the STS-induced apoptosis pathway in HPAEC contains an ATP-dependent step(s) upstream of caspase-3 activation. These findings link the change in the mode of cell death with cellular ATP content of HPAEC. Disruption of mitochondrial function by pretreatment with oligomycin before STS exposure produced a modest reduction in ATP content (~80% of control) but was not associated with a change in the mode of cell death. In contrast, oligomycin pretreatment followed by STS treatment in a glucose-free medium produced the largest reduction in ATP content (~2% of control) and a change in the mode of cell death from apoptosis to necrosis. Similar to findings in another endothelial cell line (35), these findings suggest that HPAEC require a minimum critical concentration of ATP for effective completion of apoptotic signal transduction.

Additional work is required to precisely define this critical ATP concentration. The findings in the control monolayers and STS-treated monolayers pretreated with oligomycin and incubated in glucose-free medium provide a clue (Table 4). In the control monolayers not treated with STS, necrotic cell death was observed in conjunction with a drop in ATP content to ~12% of control, whereas apoptosis proceeded normally in the STS-treated cells when the ATP concentration was ~50% of control. These findings suggest that the critical value of ATP required for completion of apoptotic signaling is 12-50% of the normal cellular ATP concentration. Overall, our findings suggest that the specific source of ATP is not as important as a rate-limiting step in signaling compared with the magnitude of ATP depletion induced by mitochondrial F₀F₁-ATPase inhibition and the absence of glucose. Another insight from these findings is that the critical threshold ATP concentration below which apoptotic signaling is inhibited appears to be lower in endothelial cells than in other cell types, e.g., Jurkat cells (34). The physiological relevance of a lower ATP threshold for inhibition of apoptosis under pathophysiological circumstances in endothelial cells remains to be determined.

In conclusion, these results highlight the role of extracellular pH, mitochondria, and ATP supply in regulating signaling events in a specific model of endothelial cell apoptosis. These results suggest the possibility of modulating cell death via interventions that modify these factors. Questions remain as to where these regulatory factors act in the cell death pathway. Apoptosis has been observed in a variety of human disorders, including cardiovascular and pulmonary diseases (10, 47), and disorders involving remodeling of the vasculature and the endothelium (11, 14). Nevertheless, the role of apoptosis in diseases involving endothelial cells remains poorly defined. A greater understanding of the factors that regulate the signaling events in endothelial cell death pathways may lead to the design of new treatments for diseases involving the vasculature and the endothelium.