Endothelin-1 Attenuates ω3 Fatty Acid–Induced Apoptosis by Inhibition of Caspase 3
Abstract—Endothelin-1 (ET-1) may be involved in the induction of vascular hypertrophy in hypertension. ET-1 may also modulate vascular growth through the exertion of antiapoptotic effects. The ω3 fatty acids (ω3 FAs), which have antiproliferative effects in various cell types, may have a beneficial role in hypertension. We tested the hypothesis that ET-1 could act as a survival factor against ω3 FA–induced apoptosis and attempted to elucidate possible molecular mechanisms underlying the protective action of ET-1 on docosahexaenoic acid (DHA)-induced apoptosis. Mesenteric vascular smooth muscle cells were stimulated with DHA, a representative ω3 FA. Dose-response curves of DHA at different apoptotic stages were assessed with the use of flow cytometry: (1) very early: plasma membrane phosphatidylserine (PS) translocation; (2) early: change in mitochondrial transmembrane potential (ΔΨm); and (3) late: cell cycle analysis. Expression of the proapoptotic protein bax and the antiapoptotic protein bcl-2 was determined with Western blot assay. The activity and the expression of caspase 3, which is a critical proteolytic enzyme involved in the death-signaling pathway, were evaluated with a fluorometric immunosorbent enzyme assay and Western blot analysis, respectively. Apoptosis, which was detected with PS translocation, ΔΨm disruption, and cell cycle analysis, was increased dose dependently by DHA. DHA-induced apoptosis was attenuated through exposure to ET-1 for 1 hour before DHA in cell cycle analysis. The interference of ET-1 with DHA-induced apoptosis, as detected with cell cycle analysis, was not apparent at the membrane (PS translocation) or the mitochondrial (ΔΨm) level. The increase in bax/bcl-2 ratio in DHA-stimulated cells was not affected by ET-1. However, DHA increased both caspase 3 activity and the active forms of caspase 3 (20 and 17 kDa), resulting in enhanced DNA fragmentation as shown through Hoechst staining and fluorescence microscopy, which were attenuated by ET-1 pretreatment. In conclusion, DHA, an ω3 FA, induced apoptosis in vascular smooth muscle cells in a dose-dependent manner. ET-1 exerted important protective effects through the attenuation of DHA-induced caspase 3 activation and subsequent DNA fragmentation in the late stages of apoptosis.
We previously proposed that endothelin-1 (ET-1) may play a pathogenic role in the vascular hypertrophy observed in severe forms of experimental and human hypertension.1 2 This effect of ET-1 may be attributed to its hypertrophic and mitogenic effects.3 However, ET-1 may also promote vascular growth through the exertion of antiapoptotic effects. ET-1 attenuated apoptosis in human pericardial smooth muscle cells4 and rat aortic endothelial cells5 induced by paclitaxel (chemical trigger) and serum depravation, respectively. In desoxycorticosterone acetate (DOCA)-salt hypertensive rats, increased apoptosis detected in aorta was accentuated by ETA endothelin receptor antagonism, suggesting that ET-1 does promote vascular growth in part through the inhibition of apoptosis.6
In hypertension, a beneficial effect has been proposed for ω3 fatty acids (ω3 FAs). Dietary ω3 FAs, which are polyunsaturated FAs abundant in fish oil, attenuated the development of high blood pressure in DOCA-salt–treated rats7 and, to a lesser degree in spontaneously hypertensive rats.8 With respect to growth, although ET-1 may inhibit apoptosis,6 ω3 FAs have instead been shown to induce apoptosis in nonvascular cells9 10 and, recently, in vascular smooth muscle cells (VSMCs).11
Apoptosis, or programmed cell death, plays a critical role in both normal development and pathology in a variety of tissues.12 Programmed cell death can be followed experimentally through the monitoring of early and late-stage events typical of apoptosis. Apoptotic cell death can be clearly visualized morphologically on the basis of cell shrinkage, cell membrane blebbing, condensation of chromatin, and nuclear fragments into membrane-enclosed apoptotic bodies. These morphological changes are accompanied by biochemical changes, including elevation of cytoplasmic Ca2+ and internucleosomal DNA fragmentation. Early events in the apoptotic process also include translocation of plasma membrane phosphatidylserine (PS) from the inner membrane to the outer membrane and disruption of the mitochondrial transmembrane potential (ΔΨm).
The effector arm of the signal transduction pathway executing the cell death program is composed of cysteine proteases belonging to the ICE/CPP32 family, recently termed caspases. Caspase 3, which is also referred to as CPP32/Yama, plays an important role as a downstream effector of the protease cascade, where various cell death pathways converge into the same pathway.13 On activation of the protease cascade, the caspase 3 proenzyme is proteolytically cleaved into p20 and p17 subunits, which then heterodimerize to form the active enzyme.
We tested the hypothesis that ET-1 could act as a survival factor against ω3 FA–induced apoptosis in VSMCs and attempted to elucidate the possible molecular mechanisms underlying the protective action of ET-1 on ω3 FA–induced apoptosis. We evaluated the ability of a representative ω3 FA, docosahexaenoic acid (DHA), to cause apoptosis in rat VSMCs in the presence and absence of ET-1 pretreatment.
DHA, propidium iodide (PI), and Hoechst 33342 were obtained from Sigma-Aldrich Canada Ltd. ET-1 was obtained from Peninsula Laboratories. A TACS Annexin V-FITC apoptosis detection kit was obtained from Genzyme. 3′3′-Dihexyloxacarbocyanine iodide [DiOC6(3)] was from Molecular Probes. The bax and bcl-2 antibodies were obtained from Santa Cruz Biotechnology. Caspase 3 antiserum was a generous gift from Dr R.-P. Sekaly (Clinical Research Institute of Montreal). A caspase 3 activity assay was purchased from Roche Diagnostics.
VSMCs derived from the mesenteric arteries of Sprague-Dawley rats at the age of 15 weeks were isolated as described in detail previously.14 Cells at passages 3 to 8 were used in all experiments. Cells were cultured in DMEM containing 10% FBS until subconfluency. Cells were kept in serum-free DMEM for 24 to 36 hours before stimulation with DHA. DHA was dissolved in ethanol, and the maximum final concentration of ethanol was 0.01% vol/vol. The effect of ET-1 on DHA-induced apoptosis was evaluated through the preincubation of cells with ET-1 (10−7 mol/L) for 1 hour before exposure to DHA.
Cell Cycle Analysis
After stimulation, cells were harvested through trypsinization, washed, and stained with propidium iodide (100 μg/mL) in the presence of permeabilizing agent Nonidet P-40 (0.3%) and RNase (20 μg/mL). The DNA content of 15 000 cells was measured through flow cytometry with the use of FACScan (Becton Dickinson). Apoptotic cells contained <2 N DNA.
Morphological Assessment of Apoptosis
The morphological features of apoptosis (eg, cell shrinkage, chromatin condensation, and DNA fragmentation) were monitored with fluorescence microscopy after the cells were stained with Hoechst 33342.
Translocation of Cell Membrane PS
Plasma membrane PS translocation, an event associated with apoptosis in which PS, a phospholipid, moves from the inner membrane to the outer membrane, was examined through the use of flow cytometry after cell staining with annexin V-FITC. After stimulation, cells were collected through trypsinization (including cells in the supernatant), washed twice with PBS, and incubated for 15 minutes at room temperature with annexin V-FITC conjugate according to the manufacturer’s protocol. The detection of PS translocation was performed with flow cytometry.
The ΔΨm results from the asymmetric distribution of protons across the inner mitochondrial membrane, giving rise to a chemical (pH) and an electrical gradient. The inner side of the inner mitochondrial transmembrane is negatively charged. As a consequence, the cationic and lipophilic fluorochrome DiOC6(3) is distributed on the inner mitochondrial matrix as a function of the Nernst equation, correlating with ΔΨm. DiOC6(3) can be used to measure variations in the ΔΨm on a per-cell basis. Cells induced to undergo apoptosis manifest an early reduction in the incorporation of ΔΨm-sensitive dye, indicating a disruption of ΔΨm. After stimulation with DHA, cells were collected through trypsinization (including cells in supernatant), washed twice with PBS, and incubated for 15 minutes at 37°C in 80 nmol/L DiOC6(3). Cells were then washed in PBS for 30 minutes at 37°C before analysis with flow cytometry.
Western Blot Analysis of bax, bcl-2, and Caspase 3
Protein was extracted from cells with the use of lysis buffer containing (in mmol/L) sodium pyrophosphate 50, NaF 50, NaCl 50, EDTA 5, EGTA 5, Na3VO4 2, HEPES 10, and PMSF 1. Protein concentration was determined with the Bio-Rad protein assay (Bio-Rad Laboratories Inc) with BSA as a standard. Then, 15 μg total protein was separated through electrophoresis on a 15% polyacrylamide gel at 100 V for 1 hour and transferred onto a PVDF membrane in a cooling system at 100 V for 1 hour. Membranes were incubated with specific antibodies to bax and bcl-2 at dilutions of 1:2000 and 1:1600, respectively, for 1 hour at room temperature. For an evaluation of the expression of inactive (molecular mass 32 kDa) and active (20 and 17 kDa) caspase 3, membranes were incubated overnight with rabbit antiserum to human CPP32 (antibody against caspase 3) at a dilution of 1:1000.15 Signals were revealed with chemiluminescence and visualized with autoradiography.
Caspase 3 Activity
Activity of caspase 3 was determined with a fluorometric immunosorbent enzyme assay (Roche Molecular Biochemicals, Roche Diagnostics). The principle was that caspase 3 derived from cellular lysates is captured by a monoclonal antibody. The amount of activated caspase 3 was cleaved proportionally through the addition of substrate. Due to proteolytic cleavage of the substrate, free fluorescent AFC (7-amino-4-trifluoromethylcoumarin) is generated and determined fluorometrically at λmax=505 nm. Briefly, cultured VSMCs were treated with vehicle or DHA for a time period indicated in the figure legends. Cells were washed twice with ice-cold PBS, harvested, and suspended in lysis buffer. After incubation on ice for 1 minute, the homogenate was centrifuged at 4°C for 30 minutes. The clear lysate was stored at −70°C until used for assays. The caspase 3 assay was carried out according to instructions from the company.
All values are presented as mean±SEM of at least 3 independent experiments. Data were analyzed with the use of Student’s t test or 1-way ANOVA followed by a Student-Newman-Keuls test as appropriate. Differences with a value of P<0.05 were considered significant.
Cell Cycle Analysis
DHA triggered apoptosis dose dependently as detected with PI staining and flow cytometry (Figure 1⇓). A maximum of 62.3±7.5% total cell number was in the subdiploid phase (ie, the apoptotic peak) at a dose of 80 μmol/L DHA (Figure 1C⇓). ET-1 (10−7 mol/L) significantly attenuated this effect of DHA (31.6±5.3%, P<0.05).
Morphological Characterization of VSMC Apoptosis Induced by DHA
Visual inspection of DHA-treated VSMCs by fluorescent microscopy demonstrated that DHA (40 μmol/L) produced DNA fragmentation and cell condensation, which were attenuated with ET-1 (10−7 mol/L) (Figure 2⇓).
Translocation of Cell Membrane PS
As detected through cell membrane PS translocation, the annexin V-FITC assay demonstrated that DHA increased apoptosis in VSMCs in a dose-dependent manner (Figure 3C⇓) to a maximum of 56.9±2.8% total cell number with a dose of 40 μmol/L DHA after 24-hour stimulation. Pretreatment of VSMCs with ET-1 had no significant effect on PS translocation.
DHA treatment disrupted ΔΨm (Figure 4⇓) in VSMCs in a dose-dependent manner to a maximum of 78.3±2.2% total cell number with 80 μmol/L DHA. Pretreatment of VSMCs with ET-1 had no significant effect on DHA-induced disruption of mitochondrial transmembrane potential.
Expression of Proapoptotic and Antiapoptotic Proteins
Western blotting was performed to quantify expression of the proapoptotic protein bax and the antiapoptotic protein bcl-2. As shown in Figure 5⇓ (top panels), the bax signal was greater in the cells treated with DHA than in vehicle control cells. However, no significant changes were found in expression of bcl-2 between both groups, resulting in a significant increase in bax/bcl-2 ratio (P<0.01) in DHA-treated cells compared with vehicle-treated cells. This effect of DHA was unaffected by ET-1 pretreatment.
Involvement of Caspase 3–Like Protease
Both caspase 3 activity and protein expression of active forms of caspase 3 (20 and 17 kDa) were increased after 1 hour of stimulation with DHA (40 μmol/L) (Figure 6⇓). The increased expression of active caspase 3 was abolished with ET-1 pretreatment (Figure 6⇓).
We report here the novel finding that DHA, an ω3 FA, induced apoptosis in VSMCs in a dose-dependent manner as detected on the basis of typical apoptotic morphological features, translocation of plasma membrane PS, disrupted ΔΨm, and redistribution of cells to the subdiploid phase of the cell cycle. Despite the striking ability of DHA to trigger apoptosis in rat VSMCs, ET-1 exerted important protective effects through attenuation of DHA-induced caspase 3 activation and subsequent DNA fragmentation in late-stage apoptosis.
Previous studies have indicated a possible inhibitory effect of ET-1 on apoptosis in rat vasculature where 2 ETA endothelin receptor antagonists, A-127722 and LU 135252, further enhanced apoptosis in aortas of DOCA-salt hypertensive rats.6 This suggests that here, ET-1 likely exerted its protective effects via stimulation of the ETA endothelin receptor subtype. In view of its growth-stimulating action, such an antiapoptotic function of ET-1 may enhance its ability to mediate vascular hypertrophy,1 2 particularly because the balance between proliferation and apoptosis plays such a critical role in the determination of net growth and blood vessel structure.
The mechanism via which DHA triggered VSMC apoptosis remains unclear. In vivo studies have shown that fish oil enhanced the release of nitric oxide,16 which may induce an apoptotic signal, because nitric oxide has been shown to upregulate Fas, a mediator of cell death.17 Indeed, nitric oxide has been shown to trigger apoptosis in VSMCs.18 Our pilot studies have shown that DHA decreased the mRNA level of ET-1. NG-Nitro-l-arginine methyl ester, which is a nitric oxide synthase inhibitor, could reverse this effect of DHA on ET-1 mRNA level (data not shown). This suggests a relationship among DHA, nitric oxide, and ET-1, which may play an important role in structure changes in vascular remodeling in hypertension.
It could be argued that because DHA was introduced to cells as a solubilized lipid rather than carrier bound as in vivo (eg, to albumin), the ability of DHA to trigger apoptosis may result from a nonspecific detergent effect. However, detergent action would induce necrotic rather than apoptotic cell death. Moreover, pilot experiments in our laboratory have shown that the proapoptotic action of DHA is not common to all FAs. Palmitic (C16:0) and oleic (C18:1) acids, which are saturated and monounsaturated acids, respectively, failed to induce apoptosis in VSMCs even at high micromolar concentrations (data not shown). Another potential mechanism could be related to the fact that DHA has oxidant properties. Nevertheless, regardless of the precise mechanism via which DHA triggered programmed cell death, this does not detract from the remarkable ability of ET-1 to protect VSMCs from such an effective apoptotic trigger.
In conclusion, DHA, an ω3 FA, induced apoptosis in VSMCs in a dose-dependent manner. ET-1 exerted important protective effects through the attenuation of DHA-induced caspase 3 activation and subsequent DNA fragmentation in late-stage apoptosis. The antiapoptotic action of ET-1 thus may be an important component in the balance between progrowth/antiapoptotic and antigrowth/apoptotic factors. Shifts in this balance may play a role in vascular changes in hypertension.
This work was supported by a Group Grant from the Medical Research Council of Canada (MRC) to the Multidisciplinary Research Group on Hypertension. Drs Diep and Intengan are supported by MRC postdoctoral research fellowships. The authors are grateful to André Turgeon for his excellent technical assistance.
- Received September 14, 1999.
- Revision received October 21, 1999.
- Accepted October 29, 1999.
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