Hepatocyte growth factor protects human endothelial cells against advanced glycation end products-induced apoposis

BBRC Biochemical and Biophysical Research Communications 344 (2006) 658–666 www.elsevier.com/locate/ybbrc

Hepatocyte growth factor protects human endothelial cells against advanced glycation end products-induced apoposis Yi Jun Zhou a

a,*

, Jia He Wang b, Jin Zhang

a

Department of Endocrinology and Metabolism, First Affiliated Hospital, China Medical University, Shenyang 110001, PR China b Department of Infectious Diseases, First Affiliated Hospital, China Medical University, Shenyang 110001, PR China Received 19 March 2006 Available online 4 April 2006

Abstract Advanced glycation end products (AGEs) form by a non-enzymatic reaction between reducing sugars and biological proteins, which play an important role in the pathogenesis of atherosclerosis. In this study, we assessed AGEs effects on human umbilical vein endothelial cells (HUVECs) growth, proliferation and apoptosis. Additionally, we investigated whether hepatocyte growth factor (HGF), an antiapoptotic factor for endothelial cells, prevents AGEs-induced apoptosis of HUVECs. HUVECs were treated with AGEs in the presence or absence of HGF. Treatment of HUVECs with AGEs changed cell morphology, decreased cell viability, and induced DNA fragmentation, leading to apoptosis. Apoptosis was induced by AGEs in a dose- and time-dependent fashion. AGEs markedly elevated Bax and decreased NF-jB, but not Bcl-2 expression. Additionally, AGEs significantly inhibited cell growth through a pro-apoptotic action involving caspase-3 and -9 activations in HUVECs. Most importantly, pretreatment with HGF protected against AGEs-induced cytotoxicity in the endothelial cells. HGF significantly promoted the expression of Bcl-2 and NF-jB, while decreasing the activities of caspase-3 and -9 without affecting Bax level. Our data suggest that AGEs induce apoptosis in endothelial cells. HGF effectively attenuate AGEs-induced endothelial cell apoptosis. These findings provide new perspectives in the role of HGF in cardiovascular disease.  2006 Elsevier Inc. All rights reserved. Keywords: Advanced glycation end products; Hepatocyte growth factor; Apoptosis; Endothelium; Caspase; Bcl-2

Cardiovascular complications are the leading cause of morbidity and mortality in patients with diabetes mellitus. Apoptosis of endothelial cells is associated with cardiovascular diseases [1,2]. Injury of endothelial cells has been postulated to be an initial trigger of the progression of atherosclerosis. Endothelial cell apoptosis is thought to play an important role in the pathogenesis of atherosclerosis. Recent study has identified that AGEs can induce endothelial cell death through the induction of apoptosis. AGEs formation may contribute to the progression of atherosclerosis, particularly in diabetes [3]. It has been well documented that AGEs progressively accumulate on the tissues and organs that develop chronic complications of diabetes mellitus, such as retinopathy, nephropathy, neuropathy, and also macrovascular disease atherosclerosis *

Corresponding author. Fax: +86 24 2334 2846. E-mail address: [email protected] (Y.J. Zhou).

0006-291X/$ - see front matter  2006 Elsevier Inc. All rights reserved. doi:10.1016/j.bbrc.2006.03.167

[4]. AGEs intervention reduce diabetes-accelerated atherosclerosis. Several growth factors have been shown to provide protection against programmed cell death [5,6]. Hepatocyte growth factor (HGF) can act as an anti-apoptotic factor for endothelial cells [7]. Therefore, we examine whether HGF prevents AGEs-induced injury of endothelial cells. HGF, also known as scatter factor, is a multifunctional cytokine possessing a wide spectrum of biological activities. It regulates cell growth, cell motility, and morphogenesis of various cell types, including endothelial cells. Moreover, HGF is a potent angiogenic factor that can induce endothelial cell proliferation and migration, without induction of vascular smooth muscle cell division [8–10]. Endothelial cell migration, proliferation, and apoptosis contribute to the pathogenesis of atherosclerosis. In addition, HGF can act as a survival factor for endothelial cells [11].

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The anti-apoptotic effects of HGF occur through several different pathways in other cell types. The different signals that converge on mitochondria to trigger or inhibit apoptosis and their downstream effects delineate several major pathways in physiological cell death. The effectors of apoptosis are now well known to be represented by a family of intracellular cysteine proteases known as caspases [12]. A feature of apoptosis that impinges on caspases is altered mitochondrial function characterized by a reduction in the electrochemical gradient across the mitochondrial membrane and release of mitochondrial cytochrome c to cytoplasm [13], and it is inhibited by the presence of Bcl2 in these organelles [14,15]. Bcl-2 and Bax are homologous proteins that have opposing effects on cell life and death, with Bcl-2 serving to prolong cell survival and Bax acting as an accelerator of apoptosis. Bcl-2 and Bax reciprocally control apoptosis by, respectively, inhibiting or stimulating mitochondrial cytochrome c release. Cytosolic cytochrome c and Apaf-1 cooperatively activate initiator caspase-9 that triggers a caspase cascade leading to apoptosis [16]. Another regulator of apoptosis is the transcription factor, nuclear factor jB (NF-jB), that participates in many important processes in endothelial cells including apoptosis and regulation of inflammatory responses. It can play a role in promoting cell survival. Several mediators of atherosclerosis, including cytokines and growth factors, have been shown to activate NF-jB [17,18]. Regulation of NF-jB activity by HGF may provide an additional anti-apoptotic mechanism in endothelial cells. However, the molecular mechanisms of the anti-apoptotic action of HGF on endothelial cells are not fully understood. We therefore use an in vitro model to further investigate the pathways of HGF against AGEs-induced apoptosis in the endothelium. Materials and methods Reagents. Recombinant human HGF(rhHGF) was purchased from R&D Systems, Inc., Monoclonal anti-Bcl-2, polyclonal anti-Bax, and monoclonal anti-b-actin antibodies were purchased from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA, USA). Anti-NF-jB p65 isoform polyclonal antibody was from Upstate (Charlottesvile, VA). RPMI-1640 and fetal bovine serum (FBS) were purchased from Gibco-BRL Company (Gibco, NY, USA). Human serum albumin (HSA), D-glucose, dimethylsulfoxide (DMSO), 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) were from Sigma Company (Sigma, St. Louis, MO). Annexin V-FITC/PI apoptosis detection kit was obtained from Bio Vision, USA, and caspase-3 and -9 activity assay kit was obtained from BD Company. Cell culture. Human umbilical vein endothelial cells (HUVECs) were obtained by collagenase treatment of umbilical cord veins as described previously [19,20]. Briefly, umbilical veins were rinsed with sterile saline and digested with 0.25% trypsin. Cells were cultured on gelatin-coated dishes and propagated in RPMI-1640 medium supplemented with 20% (v/v) heat-inactivated FBS, 50 lg/mL endothelial cell growth factor, 90 lg/mL heparin, 100 IU/mL penicillin, and 100 lg/mL streptomycin. Cells were incubated at 37 C in a humidified atmosphere of 95% air–5% carbon dioxide. Medium was refreshed every 2–3 days. These cells were tested positive for factor VIII antigen by immunohistochemical examination. HUVECs of the third to fifth passages were used. Preparation of AGE–HSA in vitro. AGE–HSA was prepared as previously reported [21]. Briefly, 0.5 g HSA was dissolved in 10 ml of 0.5

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mol/L sodium phosphate buffer (pH 7.4) with 3.0 g D-glucose, 1000 lg penicillin, and 500 lg gentamicin. The samples were filter-sterilized by using a 0.22-lm Millipore filter and incubated at 37 C for 120 days under sterile and dark conditions, dialyzed against phosphate-buffered saline (PBS, pH 7.4) in order to eliminate unconjugated glucose. While non-glycated HSA was prepared simultaneously as the same method the solution lacked glucose. AGE–HSA was identified by fluorescence spectrophotometer. Cell treatment. Endothelial cells in growth medium were equally seeded into 24-well culture plates. The growth medium was changed to serum free medium when cells were grown to approximately 90% confluence. For dose-dependent experiments, AGE–HSA was added at concentrations of 100, 200, and 400 mg/L. HSA alone (final concentration 200 mg/L) was used to treat control cells. Cell lysates were collected after 48 h of AGE– HSA exposure to investigate the presence of apoptosis. For the time course study of apoptosis, a dose of 200 mg/L was used, and apoptosis was examined after 12, 24, 36, and 48 h of AGE–HSA exposure. The protective effect of HGF on AGEs-induced apoptosis was tested by pretreatment with either 50 or 100 ng/mL HGF. A pretreatment for 12, 24, and 48 h was performed with HGF (100 ng/mL). Cell lysates were collected after 48 h of AGE–HSA exposure. Cell viability. Mitochondrial dehydrogenase activity was determined by the cleavage of MTT to purple formazan as an index of cell viability. Confluent HUVECs in 96-well plates were treated with various concentrations of AGE–HSA (100–400 mg/L) for the indicated time periods and then stimulated with various concentrations of HGF for 48 h. Each group contains eight parallel wells. After the incubation, 5 mg/mL MTT was added to the cell culture. After a 4-h reaction period, the precipitates were dissolved with 200 ll DMSO and the absorbance (A value) of each well was measured at 590 nm using an automatic microtiter plate reader. Wright’s–Giemsa staining. After cells were cultured to 90% of confluence, HUVECs were treated with AGE–HSA in the presence or absence of HGF. Adherent cells still present on their culturing support and apoptotic cells detached from their support (floating cells) were analysed. An aliquot of the culture medium containing floating cells was centrifuged on glass slides at 700 rpm for 10 min with low acceleration using a Shandron cyto centrifuge (Shandron, Pittsburg, USA). Adherent cells still present on their culturing support and floating cells collected on the glass slides were fixed with methanol, stained with Wright’s–Giemsa, and examined for their morphology. Confluency was quantified by counting cells based on light microscopy. Hoechst 33258 staining. Characteristic breakdown of the nucleus during apoptosis comprises collapse and fragmentation of the chromatin, degradation of the nuclear envelope, and nuclear blebbing, resulting in the formation of micronuclei. Visualization of morphological features of apoptosis in the nucleus of the cells such as chromatin condensation and nuclear fragmentation was determined by this assay. In short, HUVECs cultured on glass coverslips were fixed and permeabilized as above. The cells were stained with Hoechst 33258 (Molecular Probes Inc., Eugene, USA) with a dilution of 1:600 (stock solution: 1 mg/mL) for 5 min in dark. The samples were observed under a fluorescence microscope. Five hundred cells were counted from each coverslip in turns, and the results were confirmed by visualizing the apoptotic nuclei. There were five coverslips in each group, otherwise it will be mentioned in the Figure legends. Measurement of DNA fragmentation. DNA fragmentation that occurs in apoptosis produces DNA strand breaks. The fragmented DNA released into medium was measured using a cell death detection assay kit (Roche Diagnostics). Confluent HUVECs in 24-well plates were treated with AGE–HSA in the presence or absence of HGF. After incubation, the supernatant was transferred into streptavidin-coated microtiter plates. A mixture of anti-histone-biotin and anti-DNA-peroxidase was added and incubated for 2 h, and then 2,2-azino-di-[3-ethylbenzthianzoline sulfate] was added as a substrate. The content of fragmented DNA in each well was measured at 590 nm using a BioKinetics Reader EL340 (Bio-Tek Instruments Inc.). Flow cytometry analysis. Apoptosis or necrosis was determined by flow cytometer using the Annexin V-FITC apoptosis kit according to the manufacturer’s instructions. Briefly, about 1 · 104 cells were suspended in 100 ll Annexin V binding buffer and incubated with 10 ll Annexin V

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(20 lg/ml) for 5 min at room temperature in the dark. Consequently, 400 ll binding buffer containing 5 ll propidium iodide (PI, 50 lg/ml) was added and incubated on ice for additional 15 min. Then the cells were analyzed with a FacsCalibuo flow cytometer (Becton–Dickinson, USA) after 1 h. Data analysis was performed with CELL Quest software (Becton–Dickinson, USA). Experiments were performed were interpreted as follows: cells that were Annexin V( )/PI( ) (lower left quadrant) were considered as living cells, the Annexin V(+)/PI( ) cells (lower right quadrant) as apoptotic cells, Annexin V(+)/PI(+) (upper right quadrant) as necrotic or advanced apoptotic cells, and Annexin V( )/PI(+) (upper left quadrant) may be bare nuclei, cells in late necrosis, or cellular debris. Western blot analysis. After treatment, briefly, cells were washed once with ice-cold phosphate-buffered saline containing 1 mM Na2VO4 and extracted with lysis buffer (50 mM Tris, pH 8.0, 150 mM NaCl, 5 mM EDTA, 5% glycerol, 1% Triton X-100, 25 mM NaF, 2 mM Na2VO4, and 10 lg/ml of each aprotinin, leupeptin, and pepstatin). The cell lysates were frozen and thawed three times and were further centrifuged at 14,000g for 10 minutes at 4 C to pellet insoluble material. The supernatant of cell extracts was analyzed for protein concentration by a DC protein assay kit based on the Lowry method (Bio-Rad, Hercules, CA). Equal amounts of protein (50 lg) from each sample were separated on 10% sodium dodecyl sulfate–polyacrylamide gels and transferred to PVDF membranes (MSI, Westborough, MA). Membranes were blocked in 5% nonfat dry milk in Tris-buffered saline containing 0.05% Tween 20 (TBST) and then incubated with either anti-Bcl-2 (1:1000 dilution), anit-Bax (1:2000 dilution) or NF-jB (1:2000 dilution) antibodies overnight at 4 C. Actin (1:2000) was used to control for equal protein loading. The immunoblots were then washed three times with TBS-T buffer, incubated with a horseradish peroxidase conjugated secondary antibody (goat anti-rabbit IgM, Santa Cruz, CA), and developed using chemiluminescent substrate (Pierce, Rockford, IL). To quantify and compare levels of proteins, the density of each band was measured by densitometry. Measurement of caspases activity. Cells were harvested and centrifuged at 1500 rpm for 10 min. Cells were washed two times with PBS (pH 7.4) and then resuspended with 50 ll lysis buffer at 4 C and incubated on ice for 10 min. All subsequent steps were performed on ice. After centrifugation, cell extracts were transferred to fresh tubes, and protein concentrations were measured. Each 50 ll cell extract containing 100 lg of protein were combined with equal volumes of 2· reaction buffer in a microplate followed by the addition of 5 ll of peptide substrates of caspase-3 or -9 (Ac-VDVAD-pNA, Ac-DEVD-pNA, and Ac-LEHD-pNA). After overnight incubation in dark at 37 C, samples were read in a microplate reader at 405 nm. Caspase-3 and -9 activities were evaluated by the absorbance ratio of treated/control samples. Statistical analysis. Each experiment was carried out in duplicate or triplicate and three or four independent experiments were performed. Results are expressed as means ± standard deviation (SD) and analyzed with SPSS 11.5 software. Results were compared using analysis of variance (ANOVA). When ANOVA showed a statistically significant difference, a group-by-group comparison was performed using a t test with Tukey’s correction for multiple comparison. Statistical significance was set at P < 0.05.

Results Effects of AGEs on cell growth, proliferation, and apoptosis in HUVECs Our initial goal was to fully characterize the apoptotic effects of AGEs on human endothelial cells. It is important to note that increments in cell number represent a balance between cell proliferation and death. We initially examined the effect of AGEs on endothelial cells growth. To evaluate the effect of AGE–HSA on the HUVECs viability by MMT assay. Cell viability with AGE–HSA

treatment was significantly decreased compared with those with HSA treatment (control). As illustrated in Figs. 1A and B, this cytotoxic effect was time and concentration-dependent. Cytotoxicity was assayed by the morphological features of cells using Wright’s–Giemsa and Hoechst 33258 staining, respectively. AGE–HSA dramatically altered the morphology and many cells started to become round and eventually detached from the plate and floated in the medium (data not shown). HUVECs undergo apoptosis. Typical morphological changes of cell apoptosis including condensation of chromatin and nuclear fragmentation were observed using Hoechst 33258 staining under fluorescence microscope (Figs. 2A and B). Flow cytometric analysis with Annexin-V FITC/PI double staining revealed that the apoptotic rate was significantly elevated by AGE–HSA treatment compared with those with HSA treatment. AGE–HSA treatment was associated with a dose- and time-dependent increase in apoptosis of HUVECs (Figs. 3A and B). Together, these data demonstrated that AGEs cause injury to HUVECs, leading to apoptotic events. Effects of HGF on AGEs-induced apoptosis in HUVECs Previous study has shown that HGF acts as an antiapoptotic factor in endothelial cells. Therefore, we next examined its effects on AGEs-induced apoptosis. Pretreatment of HUVECs with HGF prevented AGEs-HSA induced cytotoxicity (Fig. 1C). As shown in Figs. 2A and B, HUVECs pretreated with HGF showed fewer apoptotic cells than AGEs-treated cells. A significant decrease in apoptotic cells was also confirmed by DNA fragmentation ELISA. HGF significantly suppressed the release of fragmented DNA into medium (Fig. 4). Pretreatment of HUVECs with HGF for 12, 24, and 48 h before AGE– HSA treatment significantly reduced the percentage of apoptotic cells (Fig. 5A). The percentage of cells undergoing AGEs-mediated apoptosis was decreased by pretreatment with 50 or 100 ng/mL HGF (Fig. 5B). These results suggested that HGF protects HUVECs from AGEsinduced apoptosis. The mechanism of the anti-apoptotic activity of HGF was investigated and the results are described below. Effects of HGF on AGEs-induced apoptosis-related proteins Bax, Bcl-2, and NF-jB in HUVECs To further examine the molecular mechanisms of antiapoptotic role of HGF, we focused on the expression of Bax, Bcl-2, and NF-jB proteins. AGE–HSA treatment significantly increased Bax protein, in comparison with treated control, as assessed by Western blotting. However, HGF pretreatment for 12, 24, or 48 h failed to affect the AGE–HSA induced Bax up-regulation (Fig. 6A). Interestingly, Bcl-2 protein in AGE–HSA treated samples without HGF pretreatment was not significantly different

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Fig. 1. Effects of HGF on AGEs-induced injury of human umbilical vein endothelial cells (HUVECs). Cell viability was measured by MTT assay. (A) HUVECs were treated with the indicated concentrations of AGE–HSA for 48 h. (B) HUVECs were treated with 200 mg/L AGE–HSA for indicated time. (C) Cells were pretreated with the indicated concentrations of rHGF for 48 h prior to induction with AGE–HSA. Result are shown as percent of control. The data represent means ± SD of three determinations. **P < 0.01 as compared with AGEs-treated group.

Fig. 2. Morphological changes of HUVECs cultures. Cell were cultured to 80% of confluence and treated with AGE–HSA in the presence or absence of HGF. (A) Wright’s–Giemsa staining of the cells was performed to analyze morphology of the apoptotic cells. (B) Fluorescence photomicrograph of HUVECs stained with Hoechst 33258. Apoptotic cells were observed as blue intact round nuclei and fragmented (or condensed) nuclei.

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A

Control

100 mg/L

200 mg/L

400 mg/L

AGE-HSA

B

0h

12h

24h

36h

48h

time Fig. 3. Effects of AGEs on apoptosis in HUVECs. Apoptosis assay by flow cytometry in HUVECs detected by Annexin V-FITC/PI staining. The Annexin V(+)/PI( ) cells were considered as apoptotic cells. (A) HUVECs were treated with 100–400 mg/L AGE–HSA for 48 h. The apoptotic rates were significantly increased as the AGE–HSA concentration was increased. (B) HUVECs were treated with 200 mg/L AGE–HSA for 0–48 h. The apoptotic rates increased in a time-dependent manner.

compared with treated control. However, in the presence of HGF pretreatment, a significant increase in Bcl-2 protein was observed (Fig. 6B). Furthermore, AGE–HSA exposure reduced NF-jB production. In contrast, the reductions in NF-jB protein levels were inhibited under HGF pretreatment condition (Fig. 6C). These experiments support the conclusion that the HGF protective effect against AGEs-induced apoptosis is mediated by Bcl-2 and NF-jB up-regulation.

Effects of HGF on AGEs-induced caspase-3 and -9 activation in HUVECs As shown in Figs. 7A and B, treatment of HUVECs with AGE–HSA for 48 h showed a marked increase of caspase-3 and -9 activations. Activities of caspase-3 and -9 in HUVECs with AGE–HSA treatment showed dose-dependent up-regulation. Importantly, pretreatment of HUVECs with HGF abolished caspase-3 and -9 activity induced by AGE–HSA in a concentration-dependent manner (Figs. 7C and D). These data imply that activations of caspase3 and -9 participate in AGEs-induced apoptosis and, therefore, the reduction of caspase-3 and -9 activity by HGF may account for the cytokine’s anti-apoptotic effect. Discussion

Fig. 4. Changes in DNA fragmentation following 48 h pretreatment with 50 or 100 ng/ml HGF prior to 48 h induction with 200 mg/L AGE–HSA. The release of fragmented DNA into medium was determined by ELISA 48 h after stimulation. Results are expressed as percentage of control. The values represent means ± SD of three determinations. **P < 0.01 as compared with AGEs-treated group.

Recently, it was reported that HGF might play an important role as a member of endothelium-specific growth factors locally released to counteract endothelial cell dysfunction. HGF level might be elevated in response to hypertension, acute myocardial infraction, and diabetes mellitus with hypertensive complication [22,23]. Recent studies demonstrate that HGF inhibits endothelial cells from undergoing apoptosis when serum and other growth factors are withdrawn [24,25]. Others have shown that HGF can attenuate high D-glucose-induced apoptosis and necrosis [26]. In addition, recombinant HGF-induced

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apoptosis has important implications for the maintenance of endothelial cell function. The AGEs concept proposes that chemical modification and crosslinking of tissue proteins, lipids and DNA affect their structure, function, and turnover, contributing to a

Fig. 5. The protection of HGF in AGEs-induced apoptosis. Apoptosis was measured by flow cytometry with Annexin V-FITC/PI double staining. (A) HGF pretreatment (100 ng/mL) for 48, 24, and 12 h before AGE–HSA exposure (200 mg/L) prevented AGEs-induced apoptosis. **P < 0.01 (AGE–HSA treated group versus control group). ##P < 0.01 (HGF-pretreatment for 48 and 24 h group versus no pretreatment group). # P < 0.05 (12 h HGF pretreatment group versus no pretreatment group). (B) The protective effect of HGF is dose-dependent. **P < 0.01 (AGE– HSA treated group versus control group). #P < 0.05 (50 ng/ml HGF pretreatment versus no pretreatment group). ##P < 0.01 (100 ng/ml HGF pretreatment versus no pretreatment group).

angiogenesis can attenuate the tissue damage and cell death associated with ischemic injury [27]. These studies suggest that HGF acts as a protective factor for endothelium. Understanding how HGF can inhibit endothelial cell

c Fig. 6. Effect of AGEs with and without HGF pretreatment on the Bax, Bcl-2 and NF-jB expression. The expression of Bax, Bcl-2, and NF-jB protein levels was measured by Western blot analysis using specific antibodies. The protein bands of Bax, Bcl-2, and NF-jB are indicated. b-Actin expression was used as an internal loading control. The quantitative densitometric scanning results were shown at bottom of panel. (A) AGE–HSA treatment (200 mg/L) for 48 h increased Bax protein levels (**P < 0.01). (B) HGF pretreatment for 48 h (***P < 0.001) and 24 h (**P < 0.01) increased Bcl-2 levels, compared to non-pretreated cells. (C) AGE–HSA treatment (200 mg/L) for 48 h decreased NF-jB protein levels (#P < 0.05). HGF pretreatment for 48 h (**P < 0.01) and 24 h (*P < 0.05) preserved NF-jB levels, compared to non-pretreated cells. Immunoblots are representative of three independent experiments.

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Fig. 7. Effect of HGF on AGEs-induced increase of caspase-3 and -9 activity of HUVECs. (A,B) Concentration-dependency of AGEs-induced caspase-3 and -9 activity. *P < 0.05, **P < 0.01, ***P < 0.001 as compared with the control group. (C,D) HUVECs were pretreated with the indicated concentrations of HGF for 48 h prior to induction with 200 ng/ml AGE–HSA for 48 h. Pretreatment with HGF inhibited AGEs-activated caspase-3 and -9. #P < 0.05, ##P < 0.01 as compared with non-pretreated group. Results shown are representative of at least four independent experiments.

gradual decline in tissue function and to the pathogenesis of diabetic complications. It is now thought that AGEs and advanced lipoxidation end-products contribute to accelerated micro- and macrovasculopathy observed in diabetes. Previous study has identified that AGEs play an important role in diabetes-associated atherosclerosis [28]. In this in vitro study, we examine whether HGF inhibits AGEs-induced endothelial injury that finally results in cell death through apoptotic programs. Preliminary results showed that HGF protects against morphological changes, decreases in cell viability and cell death involving both necrotic and apoptotic events resulting from treatment of HUVECs with AGEs. To our knowledge this is the first report of HGF rescued endothelial cells from death induced by AGEs. The mechanisms for AGEs-induced apoptosis and for HGF protection are still a matter of investigation. In current study, we found that AGEs-induced apoptosis involved the activation of caspase-3 and -9. Additionally, our data showed that the expression of pro-apoptotic factor Bax is greatly upregulated in endothelial cells. Caspases appear to be important for the progression of apoptotic cell death. After delivery of death signals to cells in culture, Bax protein moves to the mitochondria and other membrane sites and triggers a catastrophic change of mitochondrial function, with the subsequent release of cytochrome c. Cytochrome c is necessary for caspase-9 activation [15]. Caspase-9 can function as an initiator caspase when

mitochondrial dysfunction is the primary event in apoptosis, whereas it serves to amplify the apoptotic signaling of other initiator caspases under conditions in which disruption of mitochondria is a late event [29,30]. Cytochrome c and Apaf-1 cooperatively activate initiator caspase-9 that triggers caspase cascade and activates caspase-3. Caspase3 in turn can amplify the signal by cleaving initiator-caspases and leading to apoptosis by cleaving key intracellular targets. NF-jB activation has been linked to atherosclerosis [17,18]. Upon activation, NF-jB dimmers translocate to the nucleus and bind target genes and stimulate transcription. It regulates the activation of many genes in endothelial cells including those that regulate cell survival, apoptosis, and inflammation. In the present paper, our observations demonstrated that the apoptotic action of AGEs involves the suppression of NF-jB expression. NF-jB plays a survival role in HGF anti-apoptotic action in endothelial cells. The mechanism of HGF-induced NFjB upregulation is not clear. But several proteins are known to be intrinsically correlated. Overexpression of Bcl-2 was reported to enhance NF-jB activity. On the other hand, NF-jB was reported to increase the Bcl-2 expression by activation of Bcl-2 gene [31]. Importantly, the present study also demonstrated that HGF significantly increased the antiapoptotic proto-oncogene Bcl-2 protein without affecting Bax protein and attenuated the AGEs-induced caspase-3 and -9 activation. Bcl-2

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is an anti-apoptotic protein that maintains the integrity of mitochondria. Bcl-2 has also been suggested to exert antiapoptotic activity by inhibiting the translocation of Bax upstream of mitochondria and blocking the release of cytochrome c from mitochondria, thereby inhibiting caspase-3 and -9 activation in human endothelial cells. Anti-apoptotic action of HGF through Bcl-2 induction may be effective against not only AGEs conditions, but also other stimulation involved in high glucose. HGF attenuated high D-glucose-induced endothelial cell death through up-regulated Bcl-2, whereas HGF failed to affect antiapoptotic protein Bcl-XL[32]. Overexpressed Bcl-2 attenuated caspase-3 (-like) activation induced by high D-glucose [33]. It has also been reported that HGF can protect cell death through the phosphorylation of bad via phosphatidylinositol 3-kinase and increase Bcl-xL. HGF protected against hypoxia/reoxygenation-induced apoptosis in murine lung endothelial cell by up-regulating the expression of the Bcl-XL and FLICE-like inhibiting protein, an inhibitor of caspase-8 [34]. However, the potential unique mechanism of HGF is the ability of direct association between Bcl-2 and c-met (specific receptor of HGF) via bag-1 protein. The bag-1 protein has been reported to interact with the Bcl-2 protein and to cooperate with the Bcl-2 protein to suppress apoptosis [35]. Of importance, the bag-1 protein appears to inhibit cell death by binding to Bcl-2, the raf1 protein kinase, and c-met. The cooperative activation of these Bcl-2–related genes may also participate in the prevention of cell death by HGF, although further studies are necessary. In conclusion, the results of the current study demonstrate that AGEs can induce endothelial cell apoptosis in vitro and that this phenomenon could be completely prevented by HGF pretreatment. In addition, we propose that the upregulating Bcl-2, NF-jB expression, and inhibiting caspase-3 and -9 activation by HGF could be responsible for its protection from AGEs-induced apoptosis in the endothelial cells. These findings extend our understanding of anti-apoptotic role for endothelial cells of HGF in atherosclerosis, particularly in the diabetic context, and provide clinical investigators with an increasing number of therapeutic options to explore as part of the approach to reduce that burden of cardiovascular disease in diabetes.

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Acknowledgments The authors are grateful to Dr. Lily and Dr. Yang Peng for their excellent technical assistance and advice.

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