Introduction

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Apoptosis, or programmed cell death, has been shown to be involved in many critical biologic events including embryonic development and maintenance of normal tissue functions. In addition, apoptosis is involved in various pathologic situations. Disorders involving aberrant cell accumulation such as tumor, restenosis, and benign prostatic hyperplasia are usually the result of an imbalance between proliferation and apoptosis. Apoptosis is characterized by changes in the cell membrane structure, internucleosomal fragmentation of genomic DNA, chromatin condensation, and nuclear disintegration. Mitogens and survival factors have been shown to play important roles in regulating cell proliferation and apoptosis. These factors, by first binding to membrane receptors, trigger phosphorylation of various molecular targets, which then activate the downstream signaling cascades, leading to changes in cell death-regulating proteins, and resulting in the life or the death of a cell. The signaling pathways involved in the apoptosis-preventive effects of growth factor receptors with intrinsic tyrosine kinase activities are gradually becoming known, such as activation of phosphatidylinositol 3-kinase and extracellular signal-regulated kinases 1/2 (ERK1/2) by insulin-like growth factor-1 in PC12 cells (Párrizas et al., 1997), and stimulation of nuclear factor B by insulin in Chinese hamster ovary cells overexpressing insulin receptors (Bertrand et al., 1998). As a comparison, the roles and the underlying mechanisms of G protein-coupled receptors (GPCRs) in apoptosis remain much less understood.

Endothelin (ET) is a peptide with 21-amino acid residues (Yanagisawa et al., 1988). Three distinct members of the ET family, ET-1, ET-2, and ET-3, have been identified in humans through cloning (Inoue et al., 1989). The effects of ETs on mammalian organs and cells are initiated by their binding to GPCRs found in various tissues and cells (Sokolovsky, 1992). Two types of mammalian ET receptors, ET_A and ET_B, have been characterized and purified (Wada et al., 1990; Kozuka et al., 1991), and their cDNA have been cloned (Arai et al., 1990; Sakurai et al., 1990). ET_A receptors are selective for ET-1 and ET-2, whereas ET_B receptors bind ET-1, ET-2, and ET-3 with equal affinity. ET-1 is known to activate protein kinase C (PKC), epidermal growth factor (EGF) receptor kinase, and the ERK pathway (Douglas and Ohlstein, 1997), and is a mitogen for various cells, including smooth muscle cells (Wu-Wong et al., 1994; Yoshizumi et al., 1998) and cancer cells (Bagnato et al., 1995; Pagotto et al., 1995; Nelson et al., 1997). ET-1 is thought to play important roles in various pathophysiologic conditions including cell growth disorders.

We and others have shown previously that ET-1 protects smooth muscle cells and endothelial cells from serum withdrawal- or paclitaxel-induced apoptosis (Shichiri et al., 1997, 1998; Wu-Wong et al., 1997). However, the signaling pathways leading to the antiapoptotic effect of ET-1 are still generally unknown. The goal of this study is to investigate the possible mechanism involved in the antiapoptotic effects of ET-1 in primary culture human prostatic smooth muscle cells (HPrSMC). Our results show that the ERK1/2 pathway is activated by ET-1, and that blocking this pathway abolishes the antiapoptotic effect of ET-1. The results suggest that ERK1/2 may play a role as the downstream mediator for the antiapoptotic effect of ET-1.

	Experimental Procedures

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Materials. ET-1 was obtained from American Peptide Company (Sunnyvale, CA). Other reagents were of analytical grade.

Cell Culture. Human prostatic smooth muscle cells were obtained from Clonetics (San Diego, CA) and grown in SmGM medium containing 5% fetal bovine serum (FBS). Only cells with a passage number <9 are used in this study. Cell viability was examined using the trypan blue exclusion method.

Apoptosis Detection by Enzyme-Linked Immunosorbent Assay (ELISA). Cells in 96-well plates were treated with test agents in the presence of paclitaxel or in serum-free medium (SFM) for 48 h (or as indicated) at 37°C. At the end of the incubation, cells were lysed in 200 µl of lysis buffer (catalog no. 1544675; Roche Molecular Biochemicals, Indianapolis, IN). The cell lysates were collected and centrifuged at 200g for 10 min. The samples were assayed for apoptosis using an ELISA kit according to the manufacturer's instruction (catalog no. 1544675; Roche Molecular Biochemicals). The ELISA uses monoclonal antibodies directed against DNA and histones in a quantitative sandwich enzyme-based format. The amount of histone-associated DNA fragments (mono- and oligonucleosomes) in the cell lysates was determined at A₄₀₅ in a spectrophotometer.

DNA Synthesis. Cells in 96-well plates were cultured in growth medium until at ~70% confluency. Cells then were cultured in 0.2 ml/well SFM for 48 h and treated with FBS or ET-1 in the presence of 0.5 µCi/well [³H]thymidine for another 48 h. After the incubation, each well was washed with 0.2 ml of PBS, and then incubated with 0.2 ml of ice-cold 10% trichloroacetic acid for 30 min at 4°C. Each well was then washed again with 0.2 ml of 10% trichloroacetic acid. Materials not soluble in trichloroacetic acid were dissolved in 0.1 N NaOH for scintillation counting.

Immunocytostaining and Confocal Microscopy. Cells were grown in two-chamber slides in SmGM medium containing 5% FBS. Cells were then put into SFM for 48 h and then stimulated with or without ET-1 or 5% FBS for different periods of time. Afterward, cells were washed with PBS for 30 s, fixed with a fixing solution (0.1% glutaldehyde, 2% formalin, 80 mM PIPES, 5 mM EGTA, 1 mM MgCl₂, 0.5% Triton X-100) for 7 min, washed again with PBS for 5 min, and placed in methanol at 20°C. The slides were transferred into ice-cold acetone at 20°C for 7 min, rinsed with PBS for three times, and incubated with PBS plus 10% donkey serum for 30 min at room temperature. The slides were then incubated with an anti-phosphorylated ERK1/2 monoclonal antibody derived from mice (1000-fold dilution; New England Biolabs, Beverly, MA) in PBS with 2% donkey serum for 24 h at 4°C. After the incubation, slides were rinsed with PBS for three times and then incubated with Cy3-conjugated donkey anti-mouse IgG (100-fold dilution; Jackson ImmunoResearch Lab, West Grove, PA) for 30 min at 37°C in a humidified chamber, followed by another three rinsing with PBS. The slides were mounted, and pictures were taken using a Bio-Rad (Richmond, CA) MRC1000 confocal microscope linked to an image analyzer.

Mitogen-Activated Protein Kinase (MAPK) Activity Assay. Cells were plated at a density of 5 × 10⁴ cells/ml in 6-cm dishes. After different treatments, cells were washed with PBS twice and lysed in 0.3 ml of buffer A (10 mM Tris, pH 7.4, 150 mM NaCl, 2 mM EGTA, 2 mM DTT, 1 mM sodium orthovanadate, 1 mM phenylmethylsulfonyl fluoride, 10 µg/ml leupeptin, 10 µg/ml aprotinin, 2 µg/ml pepstatin A). Cells were homogenized using a microultrasonic cell disrupter (Kontes, Vineland, NJ) and then centrifuged at 42,000g for 20 min. The supernatant was collected. Protein content was determined by the Bio-Rad dye-binding protein assay. The MAPK activity was determined using an assay kit (catalog no. RPNS84; Amersham Pharmacia Biotech, Piscataway, NJ) according to the manufacturer's instructions. Briefly, 15 µl of the cell lysate was mixed with 15 µl of a reaction mixture containing [-³³P]ATP (1 µCi/reaction) and a synthetic peptide (NH₂-KRELVEPLTPAGEAPNQALLR-COOH) as the substrate for ERK1/2. The mixture was incubated at 30°C for 30 min. The reaction was terminated by adding 10 µl of a stop solution. The sample (30 µl) was applied to a phosphocellulose membrane. The membrane was washed extensively in 75 mM phosphoric acid and then in water. The radioactivity on the membrane was determined by a liquid scintillation counter.

Immunoprecipitation. A mixture of 0.5 ml of lysis buffer (50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1% Nonidet P40, 0.5% sodium deoxycholate) with 100 µl of a protein G-agarose bead slurry (~50%) and anti-ERK1/2 antibodies (5 µg/each) derived from rabbits (Santa Cruz Biotechnology, Santa Cruz, CA) was incubated for 24 h at 4°C to form agarose-bound immune complexes. Cells in 10-cm dishes were lysed with 1 ml of lysis buffer and then centrifuged at 25,000g for 20 min at 4°C. The supernatant (0.5 ml) was incubated with agarose beads containing the preformed anti-ERK1/2 antibody-protein G complexes for 2 h at 4°C. The agarose beads were collected by centrifugation at 12,000g for 20 s and washed twice with wash buffer (50 mM Tris-HCl, pH 7.5, 0.1% Nonidet P40, 0.05% sodium deoxycholate, 1 ml/sample/wash). To determine ERK1/2 activities, beads were resuspended in 60 µl of buffer A and assayed as described above.

SDS/PAGE and Western Blot Analysis. Samples (20 µl/sample) were resolved by SDS/PAGE using a 12% gel (Novex, San Diego, CA), and proteins were transferred electrophoretically to a polyvinylidene difluoride membrane (Immobilon-P, 0.45-µm pore size; Millipore, Burlington, MA) for Western blotting. The membrane was blotted for 1 h at 25°C with nonfat dry milk (5%) in Tris-buffered saline/Tween 20 (TBST; 10 mM Tris, pH 8.0, 0.15 M NaCl, 0.1% Tween 20), and then incubated with antiphosphorylated ERK1/2 antibodies derived from rabbit (Santa Cruz Biotechnology) in TBST for 1 h at 25°C. The membrane was washed with TBST and incubated with a horseradish peroxidase-labeled anti-rabbit antibody for 1 h at 25°C. The paper was then incubated with detection reagent containing luminol in an alkaline buffer. The specific bands were visualized by exposing the paper to blue light-sensitive autoradiography films.

	Results

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Suppression of Apoptosis in HPrSMC by ET-1. Consistent with our previous findings (Wu-Wong et al., 1997), Fig. 1A shows that paclitaxel (1 µM), a tubulin-binding agent, induced apoptosis in HPrSMC, which was partially blocked by 10 nM ET-1 (a 47% decrease). The incidence of apoptosis was assayed by an ELISA that measures mono- and oligonucleosomes in cell lysates. The presence of mono- and oligonucleosomes, an indication for DNA fragmentation, is one of the characteristics of cells undergoing apoptosis (Bonfoco et al., 1995).

View larger version (23K): [in this window] [in a new window]   Fig. 1.   Effect of ET-1 on apoptosis and proliferation. A, paclitaxel-induced apoptosis; cells in 96-well plates were fed with SmGM medium containing 5% FBS 24 h before the experiments. Cells were then treated with 1 µM paclitaxel in the presence or absence of 10 nM ET-1 for 48 h at 37°C. Apoptosis was measured by an ELISA that detects histone-associated DNA fragments. Each value represents mean ± S.D. of three determinations. B, serum withdrawal-induced apoptosis; cells cultured in 96-well plates were serum-starved in the presence or absence of ET-1 (concentrations as indicated) for 48 h at 37°C. Apoptosis was measured as in A. Each value represents mean ± S.D. of three determinations. C, DNA synthesis; cells in 96-well plates were incubated in SFM for 48 h and then treated with or without ET-1 or FBS (concentrations as indicated) for another 48 h in the presence of [3H]thymidine (0.5 µCi/well). Each value represents the mean ± S.D. of four determinations. The statistical significance was determined by the paired Student's t test.

Effects of Kinase Inhibitors on Apoptosis. To investigate whether a protein kinase is involved in the antiapoptotic effect of ET-1, we took advantage of the availability of pharmacologic tools for kinases that were shown previously to be activated by ET-1.

View larger version (43K): [in this window] [in a new window]   Fig. 2.   Effect of kinase inhibitors on apoptosis. A and B, cells in 96-well plates were treated with various kinase inhibitors for 48 h in the presence or absence of 5% FBS. Wortmannin (Wort), 1 µM; bisindolylmaleimide (Bisindo), 5 µM; tyrphostin AG490, 5 µM; AG1478, 1 µM; genistein, 10 µg/ml; PD98059 (PD), 80 µM. C and D, cells in 96-well plates were cultured in SFM for 48 h in the presence or absence of various kinase inhibitors with or without 100 nM ET-1. Concentrations for inhibitors: as indicated or PD98059, 80 µM. Apoptosis was measured by an ELISA that detects histone-associated DNA fragments. Each value represents mean ± S.D. of four determinations. In all experiments, cells were fed with SmGM medium containing 5% FBS 24 h before the treatments. In C, each inhibitor was tested in an independent experiment. Data were first converted to percentage of control (no ET-1) and then compiled into C.

Activation of ERK1/2 by ET-1. We then examined the effect of ET-1 on ERK1/2 by various approaches. Figure 3A shows that both ET-1 (10 nM) and bFGF (10 ng/ml) induced the phosphorylation of ERK1/2. Very little phosphorylation of ERK1/2 was observed in control cells that were serum-starved for 48 h. In Fig. 3B, the ERK1/2 activity was determined using a synthetic peptide (see Experimental Procedures) as the substrate after ERK1/2 was immunoprecipitated by anti-ERK1/2 antibodies. The result shows that ET-1 at 10 nM stimulated the activity of ERK1/2 by 20-fold. As a comparison, bFGF at 10 ng/ml stimulated the activity of ERK1/2 by 13-fold. In Fig. 3C, the ERK1/2 activity was determined in cellular extracts directly (without immunoprecipitation by anti-ERK1/2 antibodies) using the same assay. Similar to that shown in Fig. 3B, ET-1 exhibited a profound effect on stimulating the activity of ERK1/2. As expected, PD98059 inhibited the effect of ET-1 by >75%. Figure 3D shows the translocation of ERK1/2 from cytoplasm to nucleus on ET-1 stimulation. Cells after serum withdrawal (top) showed very faint staining with the anti-phosphorylated ERK1/2 antibody, suggesting that the MAPK activity was low. When ET-1 was added to cells, positive staining in the nucleus increased greatly at 5 and 15 min (middle and bottom). At 30 min, the nuclear staining was less than that at 5 and 15 min, but still more intense than that at time 0 (data not shown). Similar results were obtained when FBS (5%) was added to cells after serum withdrawal (data not shown). These results obtained from different approaches indicate that ET-1 exhibits a profound effect on activating ERK1/2 in these cells. Furthermore, in combination with the control studies described in Experimental Procedures, the MAPK assay using the synthetic peptide as the substrate proves to be an accurate and useful method for determining the ERK1/2 activity quantitatively.

View larger version (62K): [in this window] [in a new window]   Fig. 3.   Effects of ET-1 on ERK1/2. A, phosphorylation of ERK1/2. Cells were serum-starved for 48 h at 37°C and then treated with or without ET-1 (10 nM) or bFGF (10 ng/ml) for 5 min. Cellular extracts were prepared and analyzed by SDS/PAGE and Western blotting using an anti-phosphorylated ERK1/2 antibody as described in Experimental Procedures. B, determination of ERK1/2 activity after immunoprecipitation. Cells were treated as in A. ERK1/2 were immunoprecipitated and then assayed for the kinase activity as described in Experimental Procedures. Radioactivity, but not nanomoles per milligram per minute, was reported for the ERK activity because the protein content of ERK1/2 could not be determined accurately in the immunoprecipitates. C, determination of ERK1/2 activity in cellular extracts. Cells were serum-starved as in A. Cells were then treated with 80 µM PD98059 for 1 h before stimulated with 10 nM ET-1 for 5 min. Cellular extracts were prepared and assayed for the ERK1/2 activity directly without immunoprecipitation. D, ERK1/2 translocation. Cells in chamber slides were serum-starved for 48 h at 37°C and then treated with ET-1 (10 nM) for different time periods. Cells were then fixed and immunostained as described in Experimental Procedures.

Activity of ERK1/2 after Paclitaxel Treatment and Serum Withdrawal. To investigate the effect of ET-1 and paclitaxel treatment on the ERK1/2 pathway, the activity of ERK1/2 after paclitaxel treatment in the presence or absence of ET-1 was examined by using the quantitative MAPK assay with the synthetic peptide as the substrate. It is important to note that to keep the experimental conditions similar to the apoptosis studies noted previously, cells were fed with SmGM containing 5% FBS 24 h before the treatment. Figure 4A shows that when cells were treated with paclitaxel, the ERK1/2 activity decreased to 30% of control (the activity at time 0) at 1 h and stayed at that level for 8 h. When ET-1 was added together with paclitaxel, the ERK1/2 activity was maintained at a level similar to that at time 0 for up to 4 h. Even at 8 h, the ERK1/2 activity in the presence of ET-1 was 51% higher than that in the absence of ET-1. Figure 4B shows the results from a more detailed examination of the ERK1/2 activity in the first hour of treatment with paclitaxel and ET-1. The ERK1/2 activity at time 0 was 0.57 ± 0.01 nmol/mg/min. On treatment with paclitaxel, the ERK1/2 activity was slightly increased by 39% at 5 min and then decreased to 45% of control at 20 min, a 55% decrease in the activity compared with that at time 0. The addition of ET-1 (10 nM) in the presence or absence of paclitaxel stimulated the activity of ERK1/2 in a transient manner, reaching a peak at 2.47 ± 0.05 nmol/mg/min after 5 min of incubation at 37°C and declining to nearly the level at time 0 within 20 min. When ET-1 was added with paclitaxel, the ERK1/2 activity was maintained at ~2-fold of that in the presence of paclitaxel without ET-1 at 1 h after the treatment.

View larger version (21K): [in this window] [in a new window]   Fig. 4.   Determination of the ERK1/2 activity after paclitaxel treatment. Cells were fed with SmGM medium containing 5% FBS 24 h before the experiments. Cells were then treated with paclitaxel (1 µM) for different time periods in the presence or absence of 10 nM ET-1 without a medium change. Cellular extracts were prepared, and the ERK1/2 activity was determined as described in Experimental Procedures. Each value represents mean ± S.D. of three determinations.

View larger version (16K): [in this window] [in a new window]   Fig. 5.   Determination of the ERK1/2 activity after serum withdrawal. Cells were fed with SmGM medium containing 5% FBS 24 h before being serum-starved for different time periods in the presence or absence of 10 nM ET-1 (A) or 5% FBS (B) with or without 80 µM PD98059 (PD). Cellular extracts were prepared, and the ERK1/2 activity was determined as described in Experimental Procedures. Each value represents mean ± S.D. of three determinations.

	Discussion

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The role that GPCRs play in apoptosis is controversial. Ligands for GPCRs have been shown to be both antiapoptotic and proapoptotic. For example, sphingosine-1-phosphate, a bioactive lipid that binds to the GPCR Edg-1, is shown to suppress apoptosis in HL-60 and PC12 cells (Van Brocklyn et al., 1998). On the other hand, activation of angiotension II type 2 receptor promotes apoptosis in a rat pheochromocytoma cell line and a mouse fibroblast cell line involving the dephosphorylation of MAPKs (Yamada et al., 1996). Regarding ET, we and others (Shichiri et al. 1997, 1998) have shown that ET is antiapoptotic in smooth muscle cells, fibroblasts, and endothelial cells, whereas Okazawa et al. (1998) reports that ET induces apoptosis in A375 human melanoma cells. It is possible that different GPCRs in different cells and tissues play opposite roles in promoting or inhibiting the survival of cells. Studies on the involvement of ET-1 in regulating apoptosis pathways are still in the early phase. Very little is known about how ETs modulate apoptosis and whether the anti- versus proapoptosis effects of ETs in different cells are mediated by the same or different pathways. The role of the ET system in regulating apoptosis remains an interesting and potentially important area for future research.

ET-1 activates various kinases, including PKC, EGF receptor kinase, and MAPK. These kinases all have been shown to be involved in apoptosis signaling. For example, that PKC plays a role in apoptosis has been shown in HL-60 cells. Treating HL-60 cells with 12-O-tetradecanoylphorbol-13-acetate, a PKC activator, results in the inhibition of apoptosis (Stadheim and Kucera, 1998). Inhibition of EGF receptor kinase leads to the induction of apoptosis in human carcinoma cell line HN5 (Modjtahedi et al., 1998). Furthermore, activation of ERK1/2 is shown to be antiapoptotic (Párrizas et al., 1997; Stadheim and Kucera, 1998; Yan and Green, 1998). Our results show that inhibitors of PKC and JAK do not significantly affect apoptosis induced by serum withdrawal in HPrSMC. As a comparison, PD98059, an inhibitor of MEK-1, induces apoptosis in cells cultured in medium containing 5% FBS, potentiates apoptosis induced by serum withdrawal, and completely abolishes the antiapoptotic effect of ET-1. Previously Shichiri et al. (1998) have shown that in TGR-1 fibroblasts, PD98059 itself does not induce apoptosis but is able to abolish the antiapoptotic effect of ET-1, suggesting that ERK1/2 seems to mediate the antiapoptotic effect of ET-1 in those cells, a finding that is consistent with our observation. Furthermore, we show that although the ERK1/2 activity decreases rapidly after serum withdrawal or paclitaxel treatment in HPrSMC, the addition of ET-1 prevents the rapid fall in the ERK1/2 activity and sustains the ERK1/2 activity at a higher level for at least 4 h after serum withdrawal or paclitaxel treatment. The effect of ET-1 on ERK1/2 is also substantially inhibited by PD98059. Taken together, these data show that ERK1/2 is important for the survival of HPrSMC and suggest a role of ERK1/2 in the antiapoptotic effect of ET-1. Because cells used in this study are primary culture human cells with passage numbers <9, it is difficult to transfect these cells with dominant-negative mutants of ERK1/2 to confirm the observation made by using the kinase inhibitors. However, we feel that the observation is particularly important, because the cell system used in this study is of human origin and is in the natural state without artificial manipulation.

The antiapoptotic effect of ET-1 under serum withdrawal is extremely potent, with an approximate EC₅₀ value of 0.1 nM, which is ~10-fold less than that in paclitaxel-induced apoptosis studies with cells cultured in medium containing 5% FBS (Wu-Wong et al., 1997). However, it is important to note that the EC₅₀ value is consistent with those observed in other ET-1-evoked biologic responses, especially under conditions with a low concentration of serum albumin or other proteins. The primary reason for this difference in the EC₅₀ values in the presence or absence of serum albumin can be explained based on our previous finding that ET receptor ligands, including ET-1, -3, and some receptor antagonists, exhibit a high degree of binding to serum albumin and other human plasma proteins (Wu-Wong et al., 1998). The addition of increasing doses of plasma or serum albumin can incrementally decrease the potency of ET-1 and vice versa. Therefore, it is expected that the EC₅₀ in the absence of serum will be less than that in the presence of serum.

During this study, we have never observed a complete inhibition of apoptosis by ET-1. As a comparison, addition of 5% FBS completely blocked apoptosis. To test the possibility that degradation of ET-1 was responsible for the lack of complete inhibition, we have conducted a series of experiments in which ET-1 was added every 4 h during the 48-h incubation period after serum withdrawal. A partial inhibition of apoptosis was still observed. It is possible that FBS, but not ET-1, is able to activate an ERK-independent pathway that is also involved in apoptosis signaling. Another possibility may be linked to the fact that although ET-1 has a profound effect on activating ERK1/2, the ERK1/2 activity in the presence of ET-1 is maintained at a less active state than that in the presence of FBS (Fig. 5). Also, it is interesting to note that ET-1 does not stimulate cell growth in these cells (Fig. 1C), although ET-1 is a mitogenic factor for human pericardial smooth muscle cells (Wu-Wong et al., 1994). It appears that the effect of ET-1 on activating ERK1/2 in the prostatic smooth muscle cells is linked specifically to apoptosis, but not to cell proliferation. Previously, it has been shown that activation of ERK1/2 is required, but not sufficient, for cell growth in some cell types (Post et al., 1996; Hügl et al., 1998). Our observation in this study is consistent with that notion.

In conclusion, our results strongly suggest that ERK1/2 is involved in ET-1 antiapoptotic signaling. However, we can- not rule out the possibility that other, yet-to-be-identified, pathways may be involved. It will be necessary to examine proteins involved in regulating apoptosis, such as bcl-2, bax, and bad, to develop a better understanding of apoptosis signaling pathways involving GPCRs.