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PEDF induces p53-mediated apoptosis through PPAR gamma signaling in human umbilical vein endothelial cells

T.-C. Ho, S.-L. Chen, Y.-C. Yang, C.-L. Liao, H.-C. Cheng, Y.-P. Tsao
DOI: http://dx.doi.org/10.1016/j.cardiores.2007.06.032 213-223 First published online: 1 February 2008

Abstract

Objective Pigment epithelial-derived factor (PEDF) is a potent anti-angiogenic factor whose effects are partially mediated through the induction of endothelial cell apoptosis. The pathway mediating endothelial cell apoptosis has not been fully established. Here we investigated the participation of peroxisome proliferator-activated receptor δ (PPARδ) and p53 in the apoptosis of human umbilical vein endothelial cells (HUVECs).

Methods and results HUVECs pretreated with either PPARδ antagonist or PPARδ small interfering RNA (siRNA) suppressed PEDF-induced apoptosis as determined by TUNEL assay, annexin V-FITC/PI staining, and cleavage of procaspase-8, -9, -3. PEDF sequentially induced PPARδ and p53 expression as observed in immunoblotting and immunofluoresence assays. PEDF also increased the transcriptional activity of PPARδ as evident from electromobility shift assays, and p53 as determined by the phosphorylation and acetylation of p53 and the induction of Bax. The induction of p53 by PEDF was abolished by either PPARδ antagonist or PPARδ siRNA. PEDF-mediated HUVEC apoptosis and cleavage of procaspases were significantly attenuated by p53 siRNA.

Conclusions Our observations indicate that PEDF induces HUVECs apoptosis through the sequential induction of PPARδ and p53 overexpression. With the growing interest in anti-angiogenesis as a novel approach to cancer therapy, defining the mechanism of PEDF-mediated HUVEC apoptosis may facilitate the development of new therapeutics.

Keywords
  • PEDF
  • Apoptosis
  • HUVEC
  • PPAR gamma
  • p53

This article is referred to in the Editorial by Gaetano et al. (pages 195–196) in this issue.

1 Introduction

The normal vasculature is maintained by a balance between angiogenic factors and anti-angiogenic factors [1,2]. Neovascularization provides a nutrient supply route to the tumor and usually involves a shift towards angioinductive activity [2]. Anti-angiogenic agents can thus be used to fight cancer and other diseases involving neovascularization.

Pigment epithelium-derived factor (PEDF), a 50-kDa secreted glycoprotein, is widely expressed throughout the human body [3], with expression decreasing during human hepatocellular carcinoma and breast cancer progression [4–6], choroidal neovascularization in patients with age-related macular degeneration [7], and diabetic retinopathy [8]. PEDF binds to a cell membrane receptor to exhibit multifunctional activity in many cell types but its signaling mechanisms are largely unknown [9,10]. PEDF protects neurons against insults such as glutamate toxicity and oxidative damage and may be a more effective anti-angiogenic factor than angiostatin [3,11]. Systemic injection of recombinant PEDF protein is reported to prevent the development of retinal neovascularization in mice with oxygen-induced ischemic retinopathy by promoting apoptosis of vascular endothelial cells [12].

PEDF exerts anti-angiogenic activity by arresting VEGF- or bFGF-mediated endothelial cell migration [11], inhibiting capillary morphogenesis [13], and inducing endothelial cell apoptosis [12,14–16]. In human dermal microvascular cells, PEDF induces FasL expression and subsequently activation of caspase-8, which initiates the downstream apoptotic cascade [14]. In human umbilical vein endothelial cells (HUVECs), its induction of apoptosis was shown to depend on p38 MAPK activity and to involve activation of caspases-8 and-9 [16]. It remained unclear whether Fas signaling is also involved in HUVECs apoptosis and how activated p38 MAPK induced activation of multiple caspases.

Peroxisome proliferator-activated receptor gamma (PPARδ) is a ligand-dependent transcription factor belonging to the nuclear hormone receptor super-family [17]. PPARδ is expressed at low levels in many tissues, including the smooth muscle and endothelial cells of the vessel wall [18]. Exposure of endothelial cells to 15d-PGJ2, a natural ligand of PPARδ, as well as PPARδ overexpression achieved by transient plasmid transfection, each induce caspase-mediated endothelial cell apoptosis [19]. p53 is a well documented pro-apoptotic molecule. It mediates apoptosis primarily via the mitochondrial pathway that induces the release of potent apoptotic activators like cytochrome c [20]. Adenovirus-mediated p53 overexpression in HUVECs has been shown to induce HUVEC apoptosis and inhibit capillary-like differentiation in vitro [21,22]. However, the involvement of PPARδ and p53 in the PEDF-mediated apoptosis has not been studied.

To further our understanding of the mechanism of PEDF-induced apoptosis, we investigated the role of PPARδ and p53 in apoptosis. Our present findings suggest that PEDF activates PPARδ, which in turn leads to overexpression of p53 and apoptosis of HUVECs.

2 Materials and methods

2.1 Cell culture and treatment

HUVECs (Cascade Biologics, Inc., Portland, OR, USA) were grown in Medium 200 with Low Serum Growth Supplement (LSGS Kit, supplement contains 1.9% fetal bovine serum, 3 ng/ml bFGF, 10 μg/ml heparin, 1 μg/ml hydrocortisone, and 10 ng/ml EGF). Culture plates were coated with 2% gelatin. Cells (passages 4−8) were cultured at 37 °C in a humidified atmosphere of 5% CO2. The recombinant PEDF derived from E. coli was prepared as described previously [23]. Treatments with PEDF (200 ng/ml, unless differently specified), PPARδ inhibitors or caspase inhibitors (Calbiochem) were performed on cells (5×105/ml) seeded in LSGS medium.

2.2 Evaluation of apoptosis

After treatment, the cells were fixed in 4% (w/v) paraformaldehyde for 16 h at 4 °C and then stained using TdT-mediated dUTP nick-end labeling (TUNEL)-based kit (Roche Molecular Biochemicals) following the manufacturer's instructions. Cell number was monitored by counterstaining with 4',6-diamidino-2-phenylindole (DAPI). The percentage of TUNEL-positive nuclei was calculated in six randomly selected fields of the three different chambers (∼7200 cells). Specimens were examined and photographed on a Zeiss microscope equipped with phase-contrast and epifluorescence optics. Pictures were recorded on Zeiss software. The percentage of HUVECs apoptosis was also confirmed using the Annexin V-FITC Apoptosis Detection kit (Roche). Stained cells were analyzed by flow cytometry (FACScaliber; Beckman).

2.3 Western blot analysis

Cells were scraped into lysis buffer (150 μL/35 mm well) containing 20 mM HEPES (pH 7.4), 1% SDS, 150 mM NaCl, 1 mM EGTA, 5 mM β-glycerophosphate, 10 μg/mL leupeptin, and 10 μg/mL aprotinin. Total cell lysate was separated into cytoplasmic and nuclear fractions using the NE-PER Nuclear and Cytoplasmic Extraction Kit (Pierce). Samples containing 20 μg of protein were analyzed by 12% SDS-PAGE and then were electrotransferred to PVDF membranes (Immobilon-P; Millipore) and processed for immunoblot analysis. Antibodies used in this study were for active p38 (Promega), p38/SAPK2, acetyl-p53 (Lys373), caspase-8, caspase-9, and cytochrome c (Upstate Biotechnology), PPARα, PPARβ, and PPARδ (Santa Cruz Biotechnology), p53 (Chemicon), phospho-p53 (Ser15 and Ser20), acetylated-p53 (Lys382), and eNOS (Cell Signaling Technology), cleaved caspase-3 (Abcam), β-actin (Sigma). Proteins of interest were detected using the appropriate IgG-HRP secondary antibody (Santa Cruz Biotechnology) and ECL reagent (Amersham). X-ray films were scanned on the Model GS-700 Imaging Densitometer (Bio-Rad Laboratories) and analyzed using Labworks 4.0 software. For quantification, blots of at least three independent experiments were used.

2.4 Semi-quantitative reverse transcriptase (RT)-PCR

Total RNA was extracted from HUVECs with TRIzol reagent (Invitrogen). Synthesis of cDNA was performed with 1 μg of total RNA at 50 °C for 50 min, using oligo (dT) primers and reverse transcriptase (Superscript III, Invitrogen). The amplification mixture (final volume, 25 μl) contained 1×Taq polymerase buffer, 0.2 mM dNTPs, 1.5 mM MgCl2, 1 μM primer pair, and 0.5 U of Taq DNA polymerase (Life Technologies). cDNA was equalized in an 18–22 cycle amplification reaction with PPARδ primers 5′-caggagcagagcaaagaggtg-3′ (forward) and 5′-caaactcaaacttgggctcca-3′ (reverse), yielding a 300-bp product. The number of cycles for the PPARδ primer set (denaturation, 20 s, 94 °C; annealing, 30 s, 57 °C; and polymerization, 40 s, 72 °C) was chosen to be in the linear range of amplification.

2.5 Transfection studies

The sequences of PPARδ1 siRNA and control pGL3 siRNA duplexes were synthesized (Dharmacon) as previously described [24]. p53 siRNA was purchased from Santa Cruz Biotechnology. For the transfection procedure, cells were grown to 70% confluence, and siRNA was transfected using TransIT-TKO Transfection Reagent (Mirus Corporation). The final concentration of siRNA was 200 nM. By 24 h after siRNA transfection, cells were resuspended in new culture media, and treated with PEDF.

2.6 PPARδ transcriptional activity assay

After treatment, nuclear extracts were collected using a NucBuster protein extraction kit (Novagen), protein concentration was determination by Micro BCA Protein Assay Reagent Kit (Pierce), and the PPARδ transcriptional activity was measured using NoShiftTM II PPAR Transcription Factor Assay Kit (a modified electromobility shift assay; Novagen) as specified by the manufacturer. The specificity of protein binding was established using TransCruzTM Gel Shift PPAR specific and mutant oligonucleotides (Santa Cruz Biotechnology). Luminescence was measured by microplate luminometer (Molecular Devices).

2.7 Immunocytochemistry

Cells were plated on 2% gelatin-coated coverslips in LSGS medium. After treatment, cells were fixed with 4% paraformaldehyde and then treated at 4 °C with methanol for 10 min, and blocked with 1% goat serum and 5% BSA for 1 h. Cells were stained with antibodies to PPARδ (1:500, Santa Cruz Biotechnology) or p53 (1:1000, Chemicon), incubated with FITC-conjugated goat anti-mouse IgG antibody (1:600 dilution; Santa Cruz Biotechnology) for 1 h, and viewed with an Olympus epi-fluorescence microscope.

2.8 Statistical analysis

Data are expressed as mean±standard deviation (SD) of three independent experiments, each done in triplicate (n=3–4 dishes). The Mann–Whitney U test was used to determine statistically significant differences. P values<0.05 were considered significant.

3 Results

3.1 PPARδ antagonist abolishes PEDF-induced HUVEC apoptosis and caspase activation

To examine the involvement of PPARδ in the signaling of PEDF-induced HUVEC apoptosis, cells were pretreated with PPARδ antagonists (GW9662 or T0070907; 10 μM, 1 h) and then exposed to PEDF for 24 h. The apoptotic cells were assayed by TUNEL staining. As shown in Fig. 1A, both PPARδ antagonists can markedly inhibit PEDF-induced apoptosis. Pretreatment with the broad spectrum caspase inhibitor, zVAD-fmk (20 μM, 1 h), inhibited the PEDF effect, indicating caspase mediation. Quantification of apoptosis by annexin V-conjugated FITC and propidium iodide (PI) staining and fluorescence activated cell sorting (FACS) showed that treatment of HUVECs with PEDF for 24 h increased late apoptotic cells from 3.3±0.6% to 17.2±1.9% and pretreatment with GW9662 suppressed PEDF-induced apoptosis to ∼4.8±0.9% (Fig. 1B, upper right). Thus, inhibition of PPARδ activity attenuated PEDF-induced HUVEC apoptosis.

Fig. 1

Effects of various inhibitors on PEDF-induced HUVEC apoptosis. (A) HUVECs were untreated (UT) or treated with PEDF (P) or PEDF solvent (S) for 24 h or pretreated with 20 μM zVAD-fmk (caspases inhibitor), 10 μM GW9662, 10 μM T0070907 (PPARδ inhibitors), 10 μM SB203580 (p38 MAPK inhibitor) or DMSO (inhibitor solvent; 0.05%) for 1 h before PEDF treatment. Subsequently, the percentage of late apoptotic cells was quantified by TUNEL assay. *P<0.05 versus untreated cells. #P<0.05 versus PEDF+DMSO-treated cells. (B) HUVECs were treated as indicated and apoptosis was quantified using the Annexin V-FITC Apoptosis Detection kit. Stained cells were analyzed by flow cytometry. Upper right indicates the percentage of late apoptotic cells (annexin V/PI-stained positive cells), and lower right indicates the percentage of early apoptotic cells (annexin V-stained positive cells). (C) HUVECs were exposed to PEDF for the indicated times. Western blotting with antibodies against the active phosphorylated forms of p38 MAPK (p-p38) was performed, then the membrane were stripped and reprobed with anti-p38 antibodies to control for loading. Results are expressed as the optical density of p-p38 normalized to p38 levels.

PEDF-mediated HUVEC apoptosis is associated with cleavage of procaspase-3, -8 and -9 [16]. Pretreatment with the caspase-8 inhibitor or caspase-9 inhibitor (20 μM, 1 h) partially inhibited PEDF-induced apoptosis (13.5±1.8% or 7.1±1.5% versus PEDF+DMSO; 17.2±1.9%). The PEDF-induced apoptosis was almost completely blocked by pretreatment with both inhibitors combined (Fig. 2A). To investigate whether Fas-FasL death cascade is involved in PEDF-induced HUVEC apoptosis, FasL neutralizing antibody (NOK-2) was employed. As shown in Fig. 2B, NOK-2, which effectively blocks PI3/Akt inhibitor (25 μM LY-294002)-induced Jurkat cell apoptosis as previously reported [25], could not protect HUVECs from PEDF-induced apoptosis. This suggests that PEDF-induced HUVEC apoptosis is Fas-independent. Western blot analysis also revealed that treatment with PEDF but not PEDF solvent for 16–24 h significantly increased levels of cleaved caspases-8, -9 and -3 (Fig. 2C). Cytochrome c (Cyt-c) release from mitochondria to the cytosol has been shown to promote caspase-9 activation during apoptosis [20]. Western blot analysis showed a significant increase in Cyt-c release after PEDF treatment for 14 h (Fig. 6A). These studies confirmed that activation of both caspase-8 and -9 is involved in PEDF-triggered apoptosis and that no increased procaspase-8 and -9 cleavages occur at 1–12 h of incubation with PEDF (data not shown). Importantly, the PEDF-induced procaspases cleavages and Cyt-c release were markedly inhibited by pretreatment with GW9662.

Fig. 6

PEDF induces and activates p53 through PPARδ. (A) PEDF-induced p53 expression and cytochrome c (Cyt-c) translocation is inhibited by PPARδ inhibitor. HUVECs were treated with PEDF or solvent or pretreated with 10 μM GW9662 for 1 h before treatment with PEDF for the indicated periods. Aliquots containing equal amounts of protein from whole cell lysates (L) or cytosolic (C) were subjected to SDS-PAGE and Western blot analysis. (B and C) Mock or siRNA-transfected HUVECs were treated with PEDF for 12 h for detection of p53 by Western blot analysis. Quantification of p53 protein bands are expressed as the optical density of p53 and normalized to β-actin levels. The value observed at mock was set to 1. *P<0.05 versus mock-treated cells. #P<0.05 versus control siRNA-treated cells. (D) Immunofluorescence detection of p53 protein in HUVECs. Cells were treated with or without PEDF for 12 h, and then stained with antibodies to p53 (FITC). DNA was visualized with DAPI stain. Shown are representative pictures from three independent experiments photographed at X 400.

Fig. 2

The PEDF induces cleavage of procaspase-8, -9 and -3 in HUVECs. (A) Effects of caspase inhibitors on PEDF-induced apoptosis. HUVECs were treated with solvent or PEDF for 24 h or pretreated with 20 μM z-IETD-fmk (caspase-8 inhibitor; C8i) or 20 μM z-LEHD-fmk (caspase-9 inhibitor; C9i) for 1 h before PEDF treatment. Results of TUNEL assay are presented as percentage of cell counts. *P<0.05 versus PEDF+DMSO-treated cells. (B) Effects of FasL neutralizing antibody on Akt inhibitor-induced Jurkat T cell apoptosis and PEDF-induced HUVEC apoptosis. Jurkat T cells were either left untreated (UT) or treated with LY294002 (LY, 25 μM) or pretreated with NOK-2 antibody (50 μg/ml) or isotype IgG2a (50 μg/ml) for 1 h before LY treatment for additional 6 h. HUVECs were treated as indicated as described previously for 24 h. Subsequently, the percentage of annexin V-positive cells was analyzed by flow cytometry. *P<0.05 versus untreated cells. #P<0.001 versus control IgG-treated cells. (C) HUVECs were treated with PEDF or solvent or pretreated with 10 μM GW9662 (GW) for 1 h before treatment with PEDF for the indicated periods. Cellular proteins were then extracted for Western blot analysis.

3.2 PEDF induces PPARδ expression and activity

To investigate whether PEDF exposure can induce PPARδ expression, time-course analysis of PPARδ mRNA by RT-PCR was performed and revealed that the level of PPARδ mRNA was increased at 2 h, peaked at 4–6 h, and then dropped at 8 h as compared with an untreated or solvent-treated control (Fig. 3A). When HUVECs were pretreated with actinomycin D for 1.5 h or 3 h prior to PEDF exposure for an additional 6 h, the PPARδ mRNA level was suppressed (Fig. 3B), suggesting the increase of PPARδ mRNA was transcription dependent. The influence of PEDF on PPARδ protein expression assayed by immunoblotting revealed increased expression at 4−8 h and peak expression at 6–8 h as compared with solvent-treated cells (2.7±0.4 fold; after treatment for 6 h; Fig. 3C). Western blots also revealed expression of both PPARα and PPARβ in HUVECs, but PEDF treatment had no effect on their levels (Fig. 3C). Immunofluorescence analysis of PPARδ localization showed nuclear accumulation of PPARδ protein in untreated cells but increased nuclear and cytoplasmic accumulation of PPARδ protein after treatment with PEDF for 6 h (Fig. 3F). Thus, PPARδ expression was upregulated by PEDF at both the mRNA and protein levels.

Fig. 3

Effect of PEDF on mRNA and protein levels of PPARδ in HUVECs. (A) HUVECs were treated with PEDF for the indicated periods or treated with solvent for 6 h (indicated by *). (B) HUVECs were pretreated with 10 ng/ml actinomycin D for 1.5 or 3 h, and then incubated with PEDF for 6 h (B). Total RNA was extracted, and RT-PCR analysis for PPARδ was performed. Glyceraldehyde-3-phosphate dehydrogenase (G3PDH) expression was examined for normalization purposes. (C) HUVECs were treated with PEDF for the indicated periods and PPARs were detected by Western blot analysis. Cells treated with solvent for 6 h is indicated by *. PEDF induces PPARδ overexpression through the p38 MAPK in HUVECs. Cells were stimulated with or without PEDF or pretreated with 20 μM ERK inhibitor (PD098059), 10 μM JNK inhibitor (SP600125), 10 μM p38 MAPK inhibitor (SB203580), or the inhibitor solvent (DMSO) for 1 h prior to PEDF treatment for an additional 6 h. Total RNA or protein was then isolated for RT-PCR (D) or Western blot analysis (E) of PPARδ levels. The experiment was duplicated and yielded equivalent results. (F) Immunofluorescence detection of PPARδ protein in HUVECs. Cells were plated on 2% gelatin-coated coverslips and incubated in LSGS medium for 2 h, then left untreated or treated with PEDF for 6 h. Cells were then stained with antibodies to PPARδ (FITC). DNA was visualized with DAPI stain. FITC-2nd Ab indicates stained cells in the absence of a primary antibody. Shown are representative pictures from three independent experiments photographed at X 400.

A recent report indicated that PEDF induces p38 MAPK-dependent HUVEC apoptosis [16]. As shown in Fig. 1A, pretreatment with the p38 MAPK inhibitor, SB203580 markedly abolished PEDF-induced HUVEC apoptosis. In addition, PEDF stimulated a ∼3 fold increase in p38 MAPK phosphorylation for intervals ranging between 15 and 25 min (Fig. 1C). Therefore, our results confirmed the involvement of p38 MAPK. To investigate the role of the p38 MAPK signaling in PPARδ expression in HUVECs, SB203580 was also tested. RT-PCR analysis revealed that pretreatment with SB203580 completely blocked PEDF-induced PPARδ expression, whereas pretreatment with PD098059 (ERK inhibitor), SP600125 (JNK inhibitor), or DMSO had no such effect (Fig. 3D). Western blot analysis also revealed that pretreatment with SB203580 (but not PD098059 and SP600125) significantly blocked PEDF induction of PPARδ (Fig. 3E). Therefore, we suggest that PEDF induces p38 MAPK activation to upregulate PPARδ expression in HUVECs.

To investigate whether the PPARδ transcriptional activity could be increased after PEDF induction of PPARδ expression, a modified electromobility shift assay was employed to investigate the interaction between PPARδ and its responsive DNA. We found that the exposure of cells to varying concentrations of PEDF for 6 h induced a dose-dependent increase of DNA binding of PPARδ (Fig. 4) suggesting enhancement of PPARδ transcriptional activity. To identify the PPARδ isoform responsible for this effect, HUVECs pretreated with the PPARδ specific inhibitor GW9662 (10 μM, 1 h) before PEDF treatment. The suppression of the DNA binding effect indicates the major binding factor is PPARδ.

Fig. 4

PEDF induced PPARδ transcriptional activity. HUVECs were exposed to solvent or varying concentrations of PEDF for 6 h or pretreated with 10 μM GW9662 or DMSO for 1 h before PEDF treatment. Nuclear proteins were then extracted, aliquots containing equal amounts of protein were subjected to PPARδ transcriptional activity assay. *P<0.02 versus untreated or solvent treated cells. #P<0.05 versus DMSO+ PEDF treated cells.

3.3 PPARδ induction is critical for PEDF-mediated apoptosis

The above results suggest that PPARδ mediates PEDF-induced apoptosis (Figs. 1 and 2B). To further establish this notion, HUVECs were transfected with PPARδ-specific siRNA before PEDF treatment to prevent PPARδ expression. RT-PCR and Western blotting verified that transfection with PPARδ siRNA (but not control siRNA with a sequence unrelated to PPARδ) abolished PEDF induction of PPARδ (Fig. 5A and B). Importantly, compared with control siRNA, PPARδ siRNA significantly reduced PEDF-induced cleavages of procaspases (Fig. 5B) and apoptosis (Fig. 5C). This confirms that PPARδ expression is required for PEDF-mediated apoptosis.

Fig. 5

PPARδ siRNA silences the PEDF induction of PPARδ in HUVECs and reduces the PEDF-induced apoptosis. (A) HUVECs were transfected with PPARδ siRNA or control siRNA for 24 h, treated with PEDF for additional 6 h, and then processed for total RNA isolation and RT-PCR analysis. “Mock” indicates cells that were treated with transfection reagent. (B) Cells were treated by PEDF for an additional 6 h and 24 h and cell lysate was then isolated for Western blot analysis of PPARδ and cleaved caspases, respectively. (C) Mock or siRNA-transfected HUVECs were treated with PEDF for an additional 24 h and quantified by TUNEL assay. *P<0.05 versus mock. #P<0.05 versus control siRNA.

3.4 PEDF induces p53 gene expression and activates p53-mediated transcription through PPARδ

PPARδ can upregulate p53 gene expression [26]. We performed Western blot analysis to find out whether p53 protein level is increased by PEDF treatment. p53 levels were increased at 6 h after PEDF exposure and peaked at 10–14 h. Moreover, GW9662 pretreatment blocked p53 induction of PEDF (Fig. 6A). Transfection of HUVECs with PPARδ-specific siRNA attenuated PEDF-induced p53 overexpression (Fig. 6B and C), confirming the involvement of PPARδ in p53 induction. Immunofluorescence analysis of p53 localization showed nuclear accumulation of p53 protein before PEDF treatment and increased nuclear and cytoplasmic accumulation as compared with untreated control after treatment for 12 h (Fig. 6D). Thus increase in both PPARδ level and activity is essential for the induction of p53 expression. Endothelial nitric oxide synthase (eNOS) activation can be a candidate mechanism mediating endothelial apoptosis [27]. However, PEDF exposure did not change the levels of eNOS protein (Fig. 6A).”

Phosphorylation and acetylation of p53 are important for its promoter targeting and influence on cell fate [28,29]. To confirm that the PEDF-induced p53 is transcriptionally active, we investigated posttranslational modifications of p53. As shown in Fig. 7, PEDF stimulation caused increase in p53 acetylation (K373 and K382) and phosphorylation (S15 and S20) in a time-dependent manner, in parallel with PEDF induced increase in the expression of total p53, indicating the overexpressed p53 is modified. Since Bax is a p53 target gene and a marker of p53-mediated proapoptotic effects [28], we investigated whether Bax is induced by PEDF stimulation. As shown in Fig. 7, PEDF treatment induced Bax expression accompanying p53 overexpression.

Fig. 7

Effects of PEDF on the acetylation and phosphorylation of p53 and the expression of Bax in HUVECs. Cells were treated with PEDF alone, or pretreated with 10 μM GW9662 for 1 h and then stimulated by PEDF for the indicated time periods. Cells were harvested and subjected to Western blot analysis with acetyl-or phospho-specific antibodies of p53. PEDF-induced p53 and Bax expression was confirmed by probing membranes with total p53 or Bax antibody, respectively. Equal protein loading was confirmed by reprobing the membranes with β-actin antibody.

Taken together, our results indicate that PEDF upregulates p53 protein expression and transcriptional activity in HUVECs and these PEDF effects are mediated by PPARδ.

3.5 PEDF triggers p53-dependent HUVEC apoptosis

To verify that PEDF-induced p53 overexpression is required for PEDF-mediated apoptosis, HUVECs were transfected with p53-specific siRNA before PEDF treatment to prevent p53 expression. RT-PCR and Western blotting verified that transfection with p53 siRNA abolished PEDF induction of p53 (Fig. 8A and B). Western blot analysis also revealed that the ability of PEDF to induce procaspase 8, 9 and 3 cleavage was significantly suppressed in p53 siRNA-transfected HUVECs as compared with mock or control siRNA-transfected cells (Fig. 8B). Quantitative analysis of the numbers of TUNEL-positive cells revealed that PEDF-induced apoptosis of p53 siRNA-transfected HUVECs was significantly lower than control siRNA-transfected cells (Fig. 8C, P<0.01). Thus, p53 level also controls PEDF-induced apoptosis.

Fig. 8

PEDF does not significantly induce cleavage of multiple caspases and apoptosis in p53 siRNA-expressing HUVECs. (A) HUVECs were transfected with p53 siRNA or control siRNA for 24 h, induced by PEDF for additional 8 h, and then processed for total RNA isolation and RT-PCR analysis. (B) Cells were treated by PEDF for an additional 12 h and 24 h and cell lysate was then isolated for Western blot analysis of p53 and cleaved caspases, respectively. (C) Mock or siRNA-transfected HUVECs were treated with PEDF for an additional 24 h and quantified by TUNEL assay. *P<0.05 versus mock. #P<0.01 versus control siRNA.

4 Discussion

Induction of apoptosis of proliferating endothelial cells is an important PEDF anti-angiogenic activity. However, apoptosis regulation by PEDF signaling remains poorly understood. Here we used HUVECs to investigate proapoptotic inducers of PEDF signaling. We demonstrated that PEDF sequentially induces PPARδ and p53 expressions. Blocking any one of these effects caused attenuation of apoptosis. Use of proapoptotic inducers of endothelial cells may provide a new therapeutic avenue to restrict pathological neovascularization.

Our present study is the first to report the participation of PPARδ in PEDF signaling. We found that PEDF increase the expression (Fig. 3) and transcriptional activity (Fig. 4) of PPARδ in HUVECs. To establish the essential role of PPARδ in PEDF effect, we employed PPARδ-specific siRNA and PPARδ antagonist to inhibit PPARδ signaling and demonstrated the prevention of apoptosis (Figs. 1 and 5C) and caspases activation (Figs. 2C and 5B). Interestingly, the capability of PPARδ to induce endothelial apoptosis has been demonstrated in a previous report that exposure to PPARδ agonists or transfection-mediated overexpression of PPARδ can stimulate endothelial apoptosis [19]. The connection between PEDF and PPARδ is less clear. However, a recent report indicated that PEDF receptor is a phospholipase A2 (PLA2)-linked cell membrane receptor [10], while PLA2 can enhance the formation of PPARδ ligands to activate PPARδ in vascular endothelial cells [24]. These observations provide possible signaling mechanism between PEDF and PPARδ and support our model that PEDF signals through PPARδ.

Recently, PEDF-mediated HUVEC apoptosis was reported to be associated with activation of p38 MAPK [16]. We further found that treatment with the p38 MAPK inhibitor extensively blocks PPARδ mRNA and protein expression (Fig. 3) indicating that p38 MAPK is upstream to PPARδ in PEDF signaling. The mechanism of p38 MAPK-mediated PPARδ expression remains elusive. It has been found that early growth-response factor-1 (Egr-1) acts as transcriptional activator of PPARδ1 gene [30]. Whether p38 MAPK can regulate Egr-1 activity to promote PPARδ expression in HUVECs awaits further investigation.

Our study is the first to demonstrate that p53 is a mediator of PEDF-induced apoptosis (Fig. 8) and to reveal that PEDF upregulates p53 expression via PPARδ (Fig. 6A and B), identifying p53 as a major target in PPARδ-mediated apoptosis. The demonstration that PPARδ induces p53 expression by binding directly to the promoter region of p53 gene in human MCF7 breast cancer cells [26] suggested that PEDF might induce a similar transcriptional response in HUVECs. However, our results cannot rule out that PPARδ may trigger entry into a proapoptotic state by repressing the expression of survival/anti-apoptotic genes in HUVECs. In that report, association of nuclear receptor corepressor complexes with PPARδ caused NF-κB promoter silencing [31]. NF-κB expressed in ECs is considered important for resistance to many apoptotic stimuli [32]. The potential role of NF-κB in PEDF-mediated cytotoxicity can be tested experimentally.

Previous reports show that PEDF generates anti-angiogenic signals by activating the Fas-FasL death cascade. This effect is prevented in ECs by treatment with neutralizing antibodies against FasL or is absent in mutant mice defective for Fas or FasL [14,33]. Fas-mediated signaling mainly induces procaspase-8 autoproteolytic cleavage, although it may subsequently cause activation of procaspase-9 [34]. Our finding revealed that caspase-9 inhibitor provides more extensive protection than caspase-8 inhibitor from PEDF-induced apoptosis (Fig. 2A). In addition, we found that the FasL neutralizing antibody could not protect HUVECs from PEDF-induced apoptosis (Fig. 2B). These suggest that PEDF-induced apoptosis in HUVECs is Fas-independent and occurs primarily through activation of procaspase-9. In support of this notion, PEDF was found to inhibit ocular angiogenesis in mice deficient in Fas or FasL [35] and induces caspase-9 activation in HUVECs [16]. However, our data do not rule out differences in Fas signaling between HUVECs and ECs from other sources.

Expression of PEDF (an important multifunctional factor) decreases during progression of many diseases. We are the first to show that PPARδ and p53 acts as an apoptotic activator in PEDF-mediated signaling. This finding may contribute in part therapeutic solutions for conditions such as cancer. In addition, the pleiotropic effects of PPARδ and p53 on different cell types may help to explain the multiple biological functions of PEDF.

Acknowledgements

We thank Ju-Yun Wu for technical support. This study was supported by grants from the National Science Council, Taiwan (NSC 95-2314-B-195-009-MY3, NSC 96-3112-B-195-001) and Mackay Memorial Hospital (MMH- E - 96006).

References

  1. [1]
  2. [2]
  3. [3]
  4. [4]
  5. [5]
  6. [6]
  7. [7]
  8. [8]
  9. [9]
  10. [10]
  11. [11]
  12. [12]
  13. [13]
  14. [14]
  15. [15]
  16. [16]
  17. [17]
  18. [18]
  19. [19]
  20. [20]
  21. [21]
  22. [22]
  23. [23]
  24. [24]
  25. [25]
  26. [26]
  27. [27]
  28. [28]
  29. [29]
  30. [30]
  31. [31]
  32. [32]
  33. [33]
  34. [34]
  35. [35]
View Abstract