Mechanism of extracellular ATP- and adenosine-induced apoptosis of cultured pulmonary artery endothelial cells

Sharon Rounds1, Winnie Lin Yee2, Doloretta D. Dawicki1, Elizabeth Harrington1, Nancy Parks1, and Michael V. Cutaia1

1 Pulmonary and Critical Care Medicine Section, Providence Veterans Affairs Medical Center, Brown University School of Medicine, Providence, Rhode Island 02908; and 2 Department of Chemistry, University of Connecticut, West Hartford, Connecticut 06117

    ABSTRACT
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Abstract
Introduction
Methods
Results
Discussion
References

Apoptosis may be important in the exacerbation of endothelial cell injury or limitation of endothelial cell proliferation. We have found that extracellular ATP (exATP) and adenosine cause endothelial apoptosis and that the development of apoptosis is linked to intracellular metabolism of adenosine [Dawicki, D. D., D. Chatterjee, J. Wyche, and S. Rounds. Am. J. Physiol. 273 (Lung Cell Mol. Physiol. 17): L485-L494, 1997]. In the present study, we investigated the mechanism of this effect. We found that exATP, adenosine, and the S-adenosyl-L-homocysteine (SAH) hydrolase inhibitor MDL-28842 caused apoptosis and decreased the ratio of S-adenosyl-L-methionine to SAH compared with untreated control cells. Using release of soluble [3H]thymidine as a measure of DNA fragmentation, we found that the effect of adenosine on soluble DNA release was potentiated by coincubation with homocysteine. These results suggest that the mechanism of exATP- and adenosine-induced endothelial cell apoptosis involves inhibition of SAH hydrolase. exATP-induced apoptosis was enhanced by an inhibitor of adenosine deaminase, whereas exogenous adenosine-induced apoptosis was partially inhibited by an adenosine deaminase inhibitor. These results suggest that adenosine deaminase may also be involved in the mechanism of adenosine-induced endothelial cell apoptosis. Adenosine and MDL-28842 caused intracellular acidosis as assessed with the fluorescent probe 2',7'-bis(2-carboxyethyl)-5(6)-carboxyfluorescein. The cell-permeant base chloroquine prevented adenosine-induced acidosis but not apoptosis. Thus, although intracellular acidosis is associated with adenosine-induced apoptosis, it is not necessary for this effect. We speculate that exATP- and adenosine-induced endothelial cell apoptosis may be due to an inhibition of methyltransferase(s) activity. Purine-induced endothelial cell apoptosis may be important in limiting endothelial cell proliferation after vascular injury.

adenosine 5'-triphosphate; S-adenosyl-L-homocysteine; S-adenosyl-L-methionine; S-adenosyl-L-homocysteine hydrolase; methyltransferase; adenosine kinase; adenosine deaminase; acidosis; sodium/hydrogen antiport; nitric oxide

    INTRODUCTION
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Abstract
Introduction
Methods
Results
Discussion
References

APOPTOSIS, OR PROGRAMMED CELL DEATH, is a potential means of exacerbation of tissue injury or control of cell replication in the recovery phase after acute tissue injury (15, 25, 27). Apoptosis has been demonstrated in lungs after acute lung injury (21). In the case of endothelial cells, apoptosis may be initiated by a variety of factors, including lipopolysaccharide (13, 14) and tumor necrosis factor (22). Dawicki et al. (7) previously demonstrated that extracellular ATP (exATP) and adenosine (Ado) cause apoptosis of cultured human and bovine pulmonary arterial cells as assessed by DNA ladder formation and ethidium bromide staining. The mechanism of this effect was dependent on the generation of Ado from exATP by ectonucleotideases and on the uptake of Ado into cells (7). Because the effects of exATP and Ado on endothelial cell apoptosis were mimicked by inhibitors of S-adenosyl-L-homocysteine (SAH) hydrolase, we speculated that inhibition of the hydrolytic reaction of this enzyme was important in the mechanism of exATP-induced apoptosis of endothelial cells. The purpose of the experiments described here was to further investigate the mechanism of exATP- and Ado-induced apoptosis of endothelial cells.

We measured the effects of exATP, Ado, and an inhibitor of SAH hydrolase on the intracellular levels of SAH and S-adenosyl-L-methionine (SAM) in endothelial cells. We also determined the effects on apoptosis of inhibitors of adenosine deaminase and adenosine kinase, other pathways for the intracellular metabolism of Ado (Fig. 1). Because intracellular acidosis has been reported as a necessary and early event in apoptosis in other cell systems (12), we determined the effects of extracellular Ado and inhibition of SAH hydrolase on endothelial cell intracellular pH (pHi) and on Na/H antiport activity. We also assessed whether intracellular acidosis was necessary for Ado-induced apoptosis.


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Fig. 1.   Scheme of extracellular and intracellular metabolism of extracellular ATP (exATP) and adenosine. X, enzyme inhibitors; Decof, deoxycoformicin; SAH, S-adenosyl-L-homocysteine; SAM, S-adenosyl-L-methionine.

    METHODS
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Abstract
Introduction
Methods
Results
Discussion
References

Materials

Ado, ATP, allopurinol, N-acetylcysteine, DL-homocysteine, disodium chloroquine, SAH, SAM, 1-heptanesulfonic acid, and HEPES were obtained from Sigma (St. Louis, MO). NG-monomethyl-L-arginine (L-NMMA) was purchased from Calbiochem-Novabiochem (La Jolla, CA). 5-(N-methyl-N-isobutyl)-amiloride (MIA) was obtained from Research Biochemicals (Natick, MA). 2',7'-Bis(carboxyethyl)-5,6-carboxyfluorescein (BCECF), ethidium bromide, and nigericin were obtained from Molecular Probes (Eugene, OR). [methyl-3H]thymidine (20 Ci/mmol) came from NEN (Boston, MA). Terminal deoxynucleotidyltransferase (TdT) was obtained from Stratagene (La Jolla, CA), and biotinylated dUTP was from Boehringer Mannheim (Indianapolis, IN). Z-5'-fluoro-4',5'-didehydro-5-deoxyadenosine (MDL-28842) was donated by Hoechst Marion Roussel Research Institute (Hoechst Marion Roussel, Cincinnati, OH). 4-Amino-1-[5-amino-5-deoxy-1-beta -D-ribofuranosyl]-3-bromo-pyrazolo[3,4-d]pyrimidine (GP-1-515) was donated by Gensia (San Diego, CA).

Isolation and Culture of Bovine Endothelial Cells

Endothelial cells were obtained from bovine main pulmonary arteries with a standard scraping technique (8, 24) without enzymes and grown in MEM (GIBCO BRL, Grand Island, NY) containing 10% heat-inactivated fetal bovine serum (Gemini Bioproducts, Calabasas, CA), 1 mM sodium pyruvate, and penicillin-streptomycin-Fungizone (GIBCO BRL). Primary cultures reached confluence at 5-7 days and were subcultured every 5-7 days with trypsin-EDTA. Cultures were routinely refed two times per week (every 72 h) and 24 h before experiments.

Endothelial cells were identified by typical phase-contrast "cobblestone" morphology, by immunofluorescence to factor VIII antigen, and by uptake of acetylated low-density lipoprotein labeled with fluorescent 1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate (Biomedical Technologies, Stoughton, MA).

Assessment of Apoptosis

TdT staining. Bovine pulmonary artery endothelial cell (BPAEC) monolayers were grown on coverslips, rinsed with PBS, and then incubated for 24 h in 10 mM HEPES buffer, pH 7.35, containing 135 mM NaCl, 2 mM CaCl2, 2 mM MgSO4, 5 mM KCl, and 10 mM glucose [HEPES-buffered saline (HBS)] containing 100 µM MDL-28842, 100 µM or 1 mM Ado, or 100 µM or 1 mM ATP. Control cultures were incubated in HEPES buffer alone. Cells were washed once with PBS, fixed with 4% paraformaldehyde, and rendered permeable with 0.1% Triton X-100 in 0.1% sodium citrate. Cells used as a positive control were pretreated with 10 mg/ml of DNase I for 15 min. Cells were subsequently washed with TdT buffer (30 mM Tris, 140 mM cacodylic acid, 1 mM cobalt chloride, and 0.5% BSA) and incubated in TdT buffer supplemented with 0.3 units/ml of TdT and 1 nmol/ml of biotinylated dUTP at 37°C for 1 h. Cell samples serving as negative controls for staining were incubated in the absence of either TdT or biotinylated dUTP. The cells were washed once in 300 mM sodium chloride-30 mM sodium citrate and once in double-distilled H2O. Texas Red-conjugated streptavidin probe was added, and the cell samples were incubated at room temperature for 1 h. The stained cells were washed two times with double-distilled H2O, and the slides were placed facedown on a slide with Fluor-save. Cultures were examined with phase and fluorescence microscopy.

Soluble DNA. The assay is similar to that described by Cui et al. (5) and Zheng et al. (31). Endothelial cells were grown in six-well plates. Before confluence was reached, the cells were radiolabeled overnight with [3H]thymidine (0.05 µCi/ml) in medium-fetal bovine serum at 37°C in 95% air-5% CO2. The cells were then rinsed two times with HBS and incubated with various agents in HBS for 24 h at 37°C. Treatments were performed in duplicate on each plate. Adherent and nonadherent cells were harvested with a rubber policeman; then the cultures were centrifuged (2,000 g at 4°C for 10 min) to pellet both adherent and nonadherent cells. The resulting supernatant was saved for counting [culture supernatant counts/min (cpm)]. The cell pellet was lysed in 500 µl of ice-cold 25 mM sodium acetate buffer, pH 6.6, for 1.5 h or in 10 mM Tris · HCl, pH 7.5, containing 0.2% Triton X-100 and 2 mM EDTA for 30 min at 4°C and then centrifuged for 20 min at 4°C at 12,000 g. The lysate supernatant was removed and counted. The lysate pellet (consisting of large DNA fragments and intact chromatin) was solubilized with 1 N NaOH (100 µl) and counted (pellet cpm). Radioactivity (in cpm) in the culture supernatant (200 µl out of 5 ml), cell lysate supernatant (100 µl out of 500 µl), and insoluble cell pellet (100 µl) was determined with Optifluor scintillation fluid (Packard, Meriden, CT) and a TriCarb liquid scintillation analyzer (Packard). Soluble DNA (in percent) was calculated with the following formula
<FR><NU><AR><R><C>[(culture supernatant cpm × 25) </C></R><R><C> + (lysate supernatant cpm × 5)] × 100</C></R></AR></NU><DE><AR><R><C>(culture supernatant cpm × 25) </C></R><R><C> + (lysate supernatant cpm × 5) + (pellet cpm)</C></R></AR></DE></FR>
Dawicki et al. (7) previously demonstrated that exATP-induced soluble DNA release, as measured by this assay, is inhibited by the endonuclease inhibitor Zn2+ and is not associated with cell necrosis as assessed by 51Cr release.

Measurement of SAH and SAM

Endothelial cell monolayers in 100-mm plates were incubated in HEPES buffer, pH 7.4, for 1 h with 1 mM Ado, 1 mM exATP, or 100 µM MDL-28842 or with buffer alone. Cultures were washed two times with ice-cold PBS, and the cells were harvested by scraping, resuspended in PBS, and spun at 1,000 g for 10 min at 4°C. The cell pellet was extracted with ice-cold 0.2 N perchloric acid with stirring on ice for 10 min, and the resulting cell extract was centrifuged at 12,000 g in a microfuge at 5°C. Analysis of SAH and SAM in the supernatant was performed with Hewlett-Packard HPLC with an ODS Hypersil column (4.6 mm × 100 mm, particle size 5 µm). The mobile phase consisted of 6% methanol, 94% 40 mM NH4H2PO4, and 6 mM 1-heptanesulfonic acid sodium salt. The pH was adjusted to 4.2, and the mobile phase was filtered and degassed under vacuum through a 0.45-µm Millipore membrane filter. Isocratic elution was carried out at ambient temperature and a flow rate of 1 ml/min. A 50-µl aliquot of supernatant was injected into the column. Absorbance at 254 nm was monitored, and peaks were identified with authentic standards, SAH eluting at 10.9 min and SAM eluting at 14.5 min. Sensitivity of the assay in our hands is 10 pmol. Results are expressed as picomoles per milligram of protein. Protein was determined on the cell pellet with the Bio-Rad method.

Measurement of pHi

These methods have been previously published (6). For the fluorometric determination of pHi, cells grown on coverslips coated with CellTak (Collaborative Research) were washed two times with HEPES-buffered medium (in mM: 135 NaCl, 5 KCl, 1 CaCl2, 1 MgSO4, 10 glucose, and 16 HEPES-Tris), pH 7.4, at 37°C. All of the experiments were performed in this nominally HCO-3-free, HEPES-buffered MEM to minimize the activity of other pHi regulating systems. The cells were then incubated for 40 min at room temperature with BCECF-AM (10 µM) in the HEPES-buffered MEM. After each coverslip was carefully rinsed to remove extracellular dye by passing it through an MEM solution three times, each coverslip was inserted into a cell holder and standard cuvette in the temperature-controlled (37°C) compartment of a spectrofluorimeter equipped with a magnetic stirrer (Fluorolog system, SPEX Industries). The cell holder maintained the slide in a stable position at a 30° angle to the incident light. At the beginning of each experiment, a background measurement of autofluorescence from unloaded cells was determined. This background signal was electronically subtracted from the signals measured in dye-loaded preparations. Each monolayer was excited at 440 and 490 nm at 10-s intervals while the emitted fluorescent signals were monitored at 530 nm. The output from the photomultiplier tube of the spectrofluorimeter was downloaded to a computer that averaged the emission signals at each wavelength; ratio values (490 to 440 nm) were computed from these emission signals. The ratio of emitted fluorescence signals (530 nm) permitted calculation of the resting pHi and the rate of change in pHi, which were independent of cell number, dye loading, and dye leakage or bleaching. At the start of an experiment, the fluorescent signal from the dye-loaded cells was between 100 and 200 times background. Experiments were terminated when the fluorescent signal was <10 times background. In no instance was the autofluorescence value > 1% of the minimal fluorescence value obtained at 490 nm during an entire experiment.

The fluorescence signal was calibrated at the end of each experiment with a composite calibration curve (6) of pHi assessed by the H+ equilibration method in HEPES buffer containing 140 mM KCl and the ionophore nigericin (5 µM) as previously described (6).

Measurement of Na/H Antiport Activity

Na/H antiport activity was assessed from the rate of recovery from intracellular acidosis as previously described (6). After a 5-min equilibration period after the slides were mounted in the spectrofluorimeter, the experimental protocol was initiated. Slides were discarded if a stable signal representing baseline pHi was not obtained. Acidification was accomplished by changing the cuvette volume to an Na+-free choline chloride solution containing 1 µM nigericin, with an equimolar substitution of choline for Na+. This maneuver produced an acidification of the cells because of ionophore-induced K+ exit-H+ entry into the cell. The lowering of pHi was stabilized at a new acidified value with the addition of albumin (5 mg/ml) to scavenge the ionophore. Recovery from acidosis was dependent on the activation of the Na/H antiport. Recovery from acidosis was initiated with a change of cuvette volume back to normal MEM. As previously published by Cutaia and Parks (6), the Na+ dependency of acid recovery and the sensitivity of acid recovery to the amiloride analog 5-(N-methyl-N-isobutyl)-amiloride (MIA, 100 µM) demonstrate that acid recovery under these conditions is mediated by the Na/H antiport. The acid recovery rate was determined by calculating the rate of alkalinization (Delta pH/min) over a defined pHi range (6.65-6.80) as previously described (6). The changes in pHi (Delta pHi) at a given time point during recovery were compared between control and experimentally treated cells.

Statistics

Data are expressed as means ± SE. For most experiments, groups were analyzed for significant differences with analysis of variance and Fisher's least significant difference multiple comparison test (Statview). For experiments involving only two groups, means were compared with Student's t-test. Each monolayer was considered as n = 1. Differences were considered significant at P < 0.05.

    RESULTS
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Abstract
Introduction
Methods
Results
Discussion
References

Figure 2 shows that incubation of bovine main pulmonary artery endothelial cells for 24 h with the SAH inhibitor MDL-28842 (100 µM), Ado (100 µM or 1 mM), or ATP (100 µM or 1 mM) caused apoptosis as assessed by the TdT technique. Monolayers incubated in buffer alone (control) displayed considerably less TdT staining.


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Fig. 2.   Terminal deoxynucleotidyltransferase (TdT) staining. Bovine pulmonary arterial endothelial cell (BPAEC) monolayers on coverslips were incubated for 24 h in buffer alone (control; A and B), MDL-28842 (100 µM; C and D), adenosine (1 mM, E and F; 100 µM, G and H), or ATP (1 mM, I and J; 100 µM, K and L). Monolayers were stained for apoptosis with TdT technique as described in METHODS. Arrows, representative apoptotic cells in fluorescence (A, C, E, G, I, and K) and phase (B, D, F, H, J, and L) micrographs of the same fields. Magnification, ×920.

Figure 3 illustrates typical records of HPLC determinations of endothelial cell SAH and SAM levels. One hour of incubation with Ado and the SAH hydrolase inhibitor MDL-28842 (18) tended to increase intracellular SAH and SAM levels compared with control levels (Table 1), although increases in SAM levels did not reach statistical significance. exATP did not increase SAH and SAM levels under these conditions. However, all three agents decreased the ratio of SAM to SAH. These data are consistent with inhibition of the hydrolytic reaction of SAH hydrolase by MDL-28842 and Ado. The decreased ratio of SAM to SAH suggests methyltransferase inhibition as well (20).


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Fig. 3.   Typical records of HPLC determinations of SAH and SAM levels. A: control. B: 1 mM adenosine. C: 100 µM MDL 28842. Ordinate, absorbance at 254 nm [in milliabsorbance units (mAU)]; abscissa, time (in min).

                              
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Table 1.   Intracellular SAH and SAM in endothelial cells

The enzyme SAH hydrolase is reversible, with an equilibrium constant of 10-6 M, and both Ado and homocysteine are potent inhibitors of the hydrolytic reaction (4, 9, 28). Because homocysteine may enhance the inhibitory effect of Ado on the hydrolytic reaction catalyzed by SAH hydrolase (4, 9, 28), we determined whether exogenous homocysteine also enhances the effects of Ado on soluble DNA release. Figure 4 shows that homocysteine plus Ado increased soluble DNA release in concentrations that were ineffective for either agent alone. These results are consistent with the idea that Ado causes apoptosis of endothelial cells via inhibition of the hydrolytic reaction of SAH hydrolase.


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Fig. 4.   DNA fragmentation in BPAECs treated with adenosine and/or homocysteine. [3H]thymidine-labeled monolayers of BPAECs were incubated with varying concentrations of adenosine and/or homocysteine for 20 h in HEPES-buffered saline. Soluble DNA was isolated and quantitated as described in METHODS. Values are means ± SE; nos. in bars, no. of monolayers/group. * P < 0.05 compared with buffer alone.

We explored the possibility that exATP and Ado might cause apoptosis through metabolism of intracellular Ado by the adenosine kinase and/or adenosine deaminase pathways (Fig. 1). Rounds et al. (23) previously showed that the adenosine kinase pathway is the major route of metabolism of extracellular Ado in cultured pulmonary arterial endothelial cells. Table 2 shows that incubation of BPAECs with the adenosine kinase inhibitor GP-1-515 (11) significantly increased soluble DNA release, unlike the adenosine deaminase inhibitor deoxycoformicin. Coincubation with GP-1-515 and exATP did not increase soluble DNA release over that with GP-1-515 alone (Table 2). Coincubation with deoxycoformicin plus exATP enhanced soluble DNA release over that caused by 1 mM exATP alone (Table 2). Because neither GP-1-515 nor deoxycoformicin inhibited exATP-induced apoptosis, these results suggest that exATP-induced apoptosis is not caused by products of the adenosine kinase or adenosine deaminase pathways.

                              
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Table 2.   Effect of inhibitors of adenosine kinase and adenosine deaminase on exATP-induced DNA fragmentation

Table 3 shows the results of similar experiments with Ado as the stimulus for soluble DNA release. In these experiments, Ado was a much more potent stimulus for soluble DNA release. The effect of Ado was not changed by the adenosine kinase inhibitor GP-1-515 over apoptosis caused by Ado or GP-1-515 alone. However, the adenosine deaminase inhibitor deoxycoformicin did blunt Ado-induced soluble DNA release. These results suggest that a product of the adenosine deaminase pathway may contribute to Ado-induced apoptosis.

                              
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Table 3.   Effect of inhibitors of adenosine kinase and adenosine deaminase on adenosine-induced DNA fragmentation

Both exATP and Ado have been reported to cause release of nitric oxide (NO) from vascular endothelial cells via purinoceptor-mediated mechanisms (10, 17). Because NO can cause apoptosis of some cells (5), we determined whether the inhibitor of NO synthesis, L-NMMA, prevented exATP-induced soluble DNA release from BPAECs. Coincubation of monolayers with 1 mM exATP and 500 µM L-NMMA did not prevent exATP-induced soluble DNA release [control, 13 ± 1% release (4); exATP, 21 ± 3% release (4), exATP+L-NMMA, 25 ± 4% release (4)]. These results suggest that the mechanism of exATP-induced apoptosis does not involve NO release.

Others have shown that intracellular acidosis is associated with some models of apoptosis (1, 2, 12) and that prevention of acidosis with cell-permeant bases inhibits apoptosis (12). We investigated the role of intracellular acidosis in Ado-induced endothelial cell apoptosis. We found that inhibition of the Na/H antiport by overnight incubation of BPAECs with 10 µM MIA did increase soluble DNA release [control, 39 ± 9% release (3); MIA, 55 ± 6% release (4); P < 0.05]. Cutaia and Parks (6) previously showed that this concentration of MIA is sufficient to inhibit the endothelial Na/H antiport (6). These results suggested that intracellular acidosis may cause endothelial cell apoptosis. We then studied the role of decreased pHi in Ado-induced apoptosis. Figure 5 shows that Ado and the SAH hydrolase inhibitor MDL-28842 decreased the pHi of BPAECs after 1 h and that this effect persisted for as long as 20 h. The Na/H antiport is an important mechanism in the regulation of pHi. However, Table 4 shows that neither Ado nor MDL-28842 significantly inhibited BPAEC Na/H antiport activity as assessed by the rate of recovery from intracellular acidification. We assessed the effect of the cell-permeant base chloroquine on Ado-induced DNA fragmentation. Figure 6A shows that coincubation of BPAECs with the cell-permeant base chloroquine (1 µM) did not prevent increased soluble DNA release due to 1 mM Ado. However, this concentration of chloroquine was sufficient to prevent Ado-induced intracellular acidosis (Fig. 6B). These results indicate that, although decreased pHi is associated with Ado-induced DNA fragmentation, intracellular acidosis is probably not required for Ado-induced apoptosis.


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Fig. 5.   Effect on intracellular pH (pHi) of adenosine and SAH hydrolase inhibitor MDL-28842. BPAEC monolayers were incubated with varying concentrations of adenosine or MDL-28842 for 1, 4, or 20 h. Fluorescent probe 2',7'-bis(2-carboxyethyl)-5(6)-carboxyfluorescein (BCECF) was used to measure pHi as described in METHODS. Values are means ± SE; nos. in bars, no. of experiments. * P < 0.05 compared with respective control.

                              
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Table 4.   Effect of adenosine and SAH hydrolase inhibitor MDL-28842 on Na/H antiport activity of BPAECs


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Fig. 6.   A: effect of chloroquine on adenosine (Ado)-induced DNA fragmentation. [3H]thymidine-labeled monolayers of BPAECs were incubated with Ado and/or chloroquine (Chl) for 20 h in HEPES-buffered saline. Soluble DNA was isolated and quantitated as described in METHODS. Values are means ± SE; nos. in bars, no. of experiments. * P < 0.05 compared with control. B: effect on pHi of Ado and cell-permeant base Chl. BPAEC monolayers were incubated with Ado with and without Chl for 20 h in HEPES-buffered saline. Fluorescent probe BCECF was used to measure pHi as described in METHODS. Values are means ± SE; nos. in bars, no. of experiments. * P < 0.05 compared with control.

    DISCUSSION
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Abstract
Introduction
Methods
Results
Discussion
References

Both exATP and Ado can exert several effects on endothelial cells. Both mediators may interact with endothelial cell surface purinoceptors, with a variety of effects that could modulate acute lung injury, including enhanced adhesion of polymorphonuclear neutrophils to endothelium [exATP (8)] and generation of NO [exATP (10) and Ado (17)]. Dawicki et al. (7) previously showed that uptake of Ado into cells is necessary for exATP-induced endothelial cell apoptosis because this effect was inhibited by inhibitors of ectonucleotidases and Ado transport. Furthermore, neither P1- nor P2-purinoceptor agonists and antagonists altered exATP-induced endothelial cell apoptosis (7). In the experiments reported in this study, we provide further evidence that the effects of exATP and Ado on endothelial cell apoptosis are mediated via intracellular metabolism of Ado.

We found that only 1 h of incubation of endothelial cells with Ado mimicked the effects of an inhibitor of SAH hydrolase on intracellular SAH and SAM. The enzyme SAH hydrolase has a high Michaelis-Menten constant and is reversible. Both Ado and homocysteine are capable of product inhibition and reversal of the hydrolytic reaction (4, 19, 28). In the presence of the SAH hydrolase inhibitor MDL-28842 and Ado, intracellular levels of SAH increase (Fig. 1). Because SAH accumulation may, in turn, inhibit methyltransferases that use SAM as a methyl donor, inhibition of SAH hydrolase may result in methyltransferase inhibition, intracellular accumulation of SAM, and decreased protein, DNA, and/or mRNA methylation (4, 28). The ratio of SAM to SAH has been termed the "methylation index," and a decrease in this index has been shown to be associated with impaired intracellular methylation reactions in a number of systems (20). We found that 1 h of incubation of endothelial cells with either Ado or the SAH hydrolase inhibitor MDL-28842 decreased the SAM-to-SAH ratio. exATP had similar, but less impressive, effects on the SAM-to-SAH ratio in these experiments. These results are consistent with Ado inhibition of SAH hydrolase and methyltransferase in endothelial cells. We speculate that methyltransferase inhibition may be important in Ado-induced endothelial cell apoptosis.

Homocysteine has been reported to cause lysis of cultured endothelial cells, possibly by an oxidant-mediated mechanism (26). Homocysteine also potentiates Ado inhibition of SAH hydrolase (4, 28). Homocysteine enhanced tumor necrosis factor-alpha -induced cytotoxicity of L929 cells, whereas the combination of Ado and homocysteine increased intracellular SAH and decreased labile protein methylation (3). We found that homocysteine potentiated Ado-induced endothelial cell apoptosis. These results are consistent with the hypothesis that Ado inhibition of SAH hydrolase plays a role in Ado-induced endothelial cell apoptosis.

We also investigated the effects of inhibitors of adenosine kinase and adenosine deaminase on exATP- and Ado-induced apoptosis of endothelial cells. Rounds et al. (23) reported that exogenous Ado is metabolized by both of these enzymes in cultured endothelial cells. Wakade et al. (29) found that inhibition of adenosine kinase prevents and inhibition of adenosine deaminase potentiates Ado-induced apoptosis of chick neuronal cells. These investigators interpreted their results as being consistent with apoptosis because of adenosine kinase action causing excessive phosphorylation to adenine nucleotides and subsequent depletion of phosphoribosyl pyrophosphate. We found that the inhibitor of adenosine kinase, GP-1-515 (11), itself increased soluble DNA release from endothelial cells and that neither exATP- nor Ado-induced apoptosis was inhibited by GP-1-515. We interpret these results as not supporting a role for adenosine kinase in exATP- or Ado-induced apoptosis of cultured endothelial cells.

The products of the adenosine deaminase pathway include oxidants generated by xanthine oxidase (Fig. 1). Because oxidants may be important in apoptosis in some models (15, 30), we investigated the effects of an inhibitor of adenosine deaminase, deoxycoformicin, on exATP- and Ado-induced apoptosis. We found that deoxycoformicin enhanced exATP-induced apoptosis. In a previous studies, Dawicki et al. (7) determined that the products of adenosine deaminase, hypoxanthine and inosine, did not cause endothelial cell apoptosis and that the xanthine oxidase inhibitor allopurinol did not prevent exATP-induced apoptosis. These results do not support a role for adenosine deaminase in the mechanism of exATP-induced apoptosis.

In contrast, deoxycoformicin blunted soluble DNA release caused by exogenous Ado. Thus, unlike exATP, adenosine deaminase action may contribute to the mechanism of Ado-induced apoptosis. However, coincubation of endothelial cells with Ado and hypoxanthine did not potentiate effects of Ado on apoptosis (data not shown). We do not know why adenosine deaminase activity might be important in Ado-induced apoptosis but not in that due to exATP. These results suggest that the mechanisms of exATP- and Ado-induced apoptosis are not identical.

In other experiments, we demonstrated that 5 µM deoxycoformicin was sufficient to prevent incorporation of radiolabel from exogenous [3H]Ado into intracellular nucleosides (data not shown). Thus 5 µM deoxycoformicin was effective in inhibiting endothelial cell adenosine deaminase. Firestein et al. (11) reported that the IC50 for adenosine kinase inhibition by GP-1-515 was ~200 nM in cultured human aortic endothelial cells and ~90 nM in bovine microvascular endothelial cells and that 100 µM GP-1-515 was sufficient to maximally inhibit polymorphonuclear neutrophil adhesion. Thus the concentrations of deoxycoformicin and GP-1-515 used in these experiments were sufficient to effectively inhibit cultured endothelial cell adenosine kinase and adenosine deaminase. Because specificity is a potential problem with all pharmacological inhibitors, we cannot exclude another action of either GP-1-515 or deoxycoformicin as the cause of the effects on endothelial soluble DNA release.

Both exATP and Ado have been reported to stimulate endothelial cell generation of NO (10, 17), and NO may cause apoptosis in some cell systems (5). We found that an inhibitor of NO generation did not prevent exATP-induced apoptosis. Thus it is not likely that NO is important in this model of endothelial cell apoptosis.

These studies of the mechanism of exATP- and Ado-induced apoptosis are heavily dependent on the use of pharmacological inhibitors that could have other effects. However, in conjunction with previously reported results (7), the pattern is most consistent with a mechanism mediated via SAH hydrolase.

The literature reports a variety of effects of pHi on apoptosis. Perez-Sala et al. (19) reported that inhibition of Na/H antiport and intracellular acidification potentiated apoptosis, whereas Zhu and Loh (32) reported that inhibition of Na/H antiport and intracellular acidification were protective against apoptosis in HL-60 cells. We found that the Na/H antiport inhibitor MIA increased soluble DNA release, suggesting that decreased pHi might play a role in endothelial cell apoptosis. Furthermore, Barry et al. (2) reported that etoposide-induced apoptosis is associated with intracellular acidification in HL-60 cells. Gottlieb et al. (12) found that apoptosis in Jurkat cells (caused by anti-Fas IgM, cycloheximide, or ultraviolet irradiation) is preceded by a decreased pHi and that prevention of acidosis by the cell-permeant base chloroquine blunted apoptosis. We found that exogenous Ado and the SAH hydrolase inhibitor both caused intracellular acidification in cultured endothelial cells. This acidification was not due to inhibition of Na/H antiport activity as assessed by the rate of acid recovery under conditions where the Na/H antiport is the major mechanism of acid recovery. Thus endothelial cells possessed a functional Na/H antiport even in the presence of exogenous Ado and MDL-28842. However, we cannot exclude alteration of the pH sensitivity of the antiport as a cause of acidification, as has been suggested in the case of interleukin-2 withdrawal-induced apoptosis of the lymphocytic cell line CTLL-2 (16). Other possible causes of apoptosis-associated acidification are increased metabolic acid production via glycolysis or inhibited mitochondrial respiration. Further experiments are needed to determine the cause of Ado- and MDL-28842-induced intracellular acidosis.

We found that prevention of Ado-induced acidosis by concommitant incubation with the cell-permeant base chloroquine did not prevent apoptosis. Thus intracellular acidification is not necessary for Ado-induced apoptosis. These results are consistent with those reported by Li and Eastman (16) in CTLL-2 cells. The relationship between intracellular acidosis and apoptosis is not clear. In some systems, acidosis appears to be necessary for apoptosis, perhaps via activation of endonuclease(s) (1). Our results suggest that acidosis is an associated, but not necessary, condition for Ado-induced apoptosis.

In summary, the results of these experiments support the hypothesis that Ado-induced apoptosis of endothelial cells is due to intracellular metabolism of Ado and/or inhibition of SAH hydrolase. We speculate that inhibition of methyltransferase(s) may be important in Ado-induced endothelial cell apoptosis. Intracellular acidosis is associated with apoptosis in this model but does not appear to be causative. Increased tissue concentrations of purines may occur after cytolytic injuries such as those due to rhabdomyolysis or hemolysis. We speculate that Ado-induced endothelial cell apoptosis may be an important means of endothelial cell injury.

    ACKNOWLEDGEMENTS

We thank Dr. Ekkehard H. W. Bohme (Hoechst Marion Roussel Research Institute, Hoechst Marion Roussel, Cincinnati. OH) for the kind donation of MDL-28842 and Dr. Gary S. Firestein (Gensia, San Diego, CA) for the kind donation of GP-1-515 used in these studies. We are grateful to Drs. Ming Y. W. Chu, Bai-Chuan Pan, and Paul Calabresi (Division of Clinical Pharmacology, Department of Medicine, Brown University School of Medicine, Providence, RI) for generous assistance and guidance with the high-performance liquid chromatography determinations.

    FOOTNOTES

This work was supported by a Merit Review Grant from the Department of Veterans Affairs; the Dean of Brown University School of Medicine; the Cystic Fibrosis Foundation; and National Heart, Lung, and Blood Institute Grant HL-34009.

Preliminary results were presented at the annual meeting of the American Thoracic Society in May 1997 and have been published in abstract form (Am. J. Respir. Crit. Care Med. 155: A122, 1997).

Address for reprint requests: S. Rounds, Pulmonary and Critical Care Medicine Section, Providence VA Medical Center, 830 Chalkstone Ave., Providence, RI 02908.

Received 29 September 1997; accepted in final form 20 April 1998.

    REFERENCES
Top
Abstract
Introduction
Methods
Results
Discussion
References

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