HGF-mediated apoptosis via p53/bax-independent pathway activating JNK1

Elizabeth A. Conner, Tadahisa Teramoto, Peter J. Wirth, Andras Kiss1, Susan Garfield and Snorri S. Thorgeirsson2

Laboratory of Experimental Carcinogenesis, National Cancer Institute, National Institutes of Health, Bethesda, MD 20892, USA


    Abstract
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 Abstract
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 Materials and methods
 Results
 Discussion
 References
 
Current studies have indicated both positive and negative roles for the hepatocyte growth factor (HGF)/c-met receptor signaling system in tumor development. Recently, we have shown that HGF has the capacity to induce both growth inhibition and programmed cell death in aflatoxin-transformed (AFLB8) rat liver epithelial cells. Using the same cell line, we have now investigated a potential mechanism for HGF-induced apoptosis. Immunoblot analysis of bcl-2 gene family member (bax, bcl-2, bclX-s/l) expression showed no correlation with HGF treatment, suggesting that HGF-mediated apoptosis is bax independent. Following HGF treatment retinoblastoma protein (pRB) was present in the hypophosphorylated state. HGF treatment increased cyclin A, cyclin G1 and nuclear transcriptional factor (NF{kappa}B) protein expression. However, electrophoretic mobility shift analysis showed that NF{kappa}B activity decreased with HGF treatment. Under these apoptotic conditions, c-Jun N-terminal kinase (JNK1) and extracellular signal-regulated kinase (ERK2) were activated with lower level activation of ERK2, while no involvement of phosphatidylinositol-3 kinase was observed. Epidermal growth factor (EGF) was not protective, and actually induced cells to undergo apoptosis to a level similar to that of HGF alone or EGF/HGF in combination. These results suggest the possibility of cross-talk between HGF/c-met and EGF/EGFR signaling pathways, and the involvement of JNK1 induction in HGF-mediated apoptotic cell death.

Abbreviations: EGF, epidermal growth factor; ERKS, extracellular signal-regulated kinase; HGF, hepatocyte growth factor; ICE, interleukin-1-converting enzyme; IGF, insulin-like growth factor; INF{gamma}, interferon gamma; JNK, c-Jun N-terminal kinase; PBS, phosphate-buffered saline; PDGF, platelet-derived growth factor; PI3K, phosphatidylinositol-3 kinase; PTK, protein tyrosine kinase.


    Introduction
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 Introduction
 Materials and methods
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Many growth factors and cytokines activate protein tyrosine kinase (PTK) signaling pathways leading ultimately to gene expression (1,2). PTK pathways have been shown to play integral roles in cell proliferation and survival. For many cells, either cell growth arrest or apoptosis will occur following growth factor or cytokine deprivation (2,3). Thus, growth factors, such as epidermal growth factor (EGF), platelet-derived growth factor (PDGF) and insulin-like growth factor (IGF), which normally induce mitogenic responses, can act as survival factors (2,46). In contrast, certain growth factors and cytokines, such as EGF and interferon gamma (INF{gamma}), can trigger cell cycle arrest and death in a variety of cells following activation of the receptor tyrosine kinase complex (2,7,8).

Hepatocyte growth factor (HGF), also a potent mitogen and survival factor for a variety of cell types, can inhibit the growth of hepatoma cell lines (912). Evidence from our laboratory demonstrated that HGF inhibits c-myc dependent carcinogenesis, and recently, has the capacity to induce programmed cell death (13,14) in vitro. However, the mechanism by which apoptosis is induced by PTK pathways and the nature of the signal transduction mediators involved remain to be determined. Recently, STK/RON, a novel PTK receptor that belongs to the same receptor subfamily as the HGF receptor, c-met, was found to mediate both cell death by apoptosis, as well as inducing cell proliferation (15). Apoptosis, in this case, was accompanied by the prolonged activation of c-Jun N-terminal kinase (JNK) with no involvement of bcl-2 family members or interleukin-1ß-converting enzyme (ICE) (15).

Activation of the JNK kinase pathway has been identified as a critical signaling pathway in mediating apoptosis. Prominent activation of JNK has been observed in cells treated with apoptotic inducers, such as TNF-{alpha} and {gamma} irradiation, as well as with the tumor suppressor, p53 (16,17). Sustained activation of JNK has been associated with concurrent inhibition of the extracellular signal-regulated kinases (ERKS), suggesting that a dynamic balance between ERK and JNK activation may be important in whether a cell survives or undergoes apoptotic cell death (16).

We have previously shown that HGF-mediated apoptosis in transformed rat liver epithelial cells was associated with constitutive expression of c-myc and 1 kb bax-{alpha} transcripts (13). Neither c-myc nor bax expression was modulated by either HGF treatment or p53 expression (13). Here, we show that HGF mediates a bax-independent apoptotic pathway which decreases NF{kappa}B activity, activates JNK1 and potentially involves cyclin G1. Both HGF- and EGF-induced JNK1 activation showed similar kinetics, suggesting the possibility of cross-talk between HGF and EGF receptor signaling pathways.


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Human recombinant HGF was a generous gift of Dr George Vande Woude (NCI-FDRDC, ABL-Basic Research Program, Frederick, MD).

Derivation of cell lines
The RLE {Phi} 13 cells and the clonal, aflatoxin B1-transformed variant, AFLB8, have been extensively characterized (1821), and were maintained in Ham's modified F-12 medium (Biofluids, Rockville, MD) supplemented with 10% defined FBS and 50 µg/ml gentamicin at 37°C, in a humidified atmosphere of 5% CO2 and 95% air unless otherwise stated.

Treatment of cells and preparation of cellular lysates
Cells were seeded at a density of 2.6x103 cells/cm2 in 100 mm dishes. Twenty-four hours after plating, cells were equilibrated in either serum-reduced media (2%) or serum-free media for up to 24 h, and then treated with HGF (50 ng/ml), EGF (10 ng/ml) or Wortmannin (3 or 10 µM) for the times and conditions indicated. The concentrations of HGF and EGF were determined from our previous report on the apoptotic effects of HGF (13) and earlier characterization of the RLE cell lines. The doses of Wortmannin were determined from previous studies by colleagues examining PI3K activity in normal RLE cells (T.Teramoto and S.S.Thorgeirsson, unpublished data). After treatment, cells were washed twice with phosphate-buffered saline (PBS) and harvested by scraping with a cell scraper in RIPA buffer (1x PBS, 1% NP-40, 0.5% sodium deoxycholate and 0.1% SDS) containing inhibitors (1 mM sodium orthovanadate, 20 mM sodium pyrophosphate, 1 mM phenylmethylsulfonylfluoride, 10 µg/ml each leupeptin and aprotinin). Whole cell lysates were collected by centrifugation at 15 000 g at 4°C, and either used directly for SDS/PAGE or stored at –20°C until electrophoresis.

SDS PAGE and immunoblot analysis
Aliquots of whole cell lysate proteins (40 µg) were diluted 1:4 with 5x SDS sample buffer, heated in boiling water bath for 5 min, and separated by SDS/PAGE in either 10 or 14% polyacrylamide SDS gels (Novex, San Diego, CA). Proteins were then transferred electrophoretically to nitrocellulose membranes (Novex) in 25 mM Tris base, 192 mM glycine and 20% methanol at 100 V for 2 h. Immunoblot analysis was performed after incubating membranes in 3% non-fat milk in 20 mM Tris buffered saline, pH 7.5, containing 0.1% Tween-20 (TBST) for 1 h. Membranes were incubated with the primary antibodies diluted in TBST either overnight at 4°C or for 1 h at room temperature. The appropriate horseradish peroxidase-conjugated secondary antibody was diluted 1:10 000 in 3% non-fat milk in TBST and incubated with the membrane for 1 h at room temperature. Proteins were visualized by the ECL detection system (Amersham, Piscataway, NJ). Primary antibodies were as follows: Bax (I-19) SC-930, BclX-s/l (S-18) SC-634, Bcl-2 (N-19) SC-492, Rb (C-15) SC-50, p27 SC-527, cyclin A (C-19) SC-596, and NF{kappa}B p65 (C-20) SC-372-G and were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Cyclin G1 polyclonal rabbit antibody was made against a synthetic 24 amino acid peptide from the deduced cyclin G1 sequence (22).

Trypan blue exclusion test
Twenty-four hours after seeding in 100 mm dishes (1.2x104 cells/dish), the cultures were exposed to serum-reduced media or media containing 50 ng/ml HGF. At 24, 48, 72 and 96 h after treatment, the medium was removed, and the cultures washed twice with PBS and trypsinized. Cell viability was determined with a haemocytometer and trypan blue exclusion using a 0.4% solution of trypan blue in PBS.

Fluorescence microscopy
Cells (2x103 cells/well) were plated on 2-well chamber slides (NUC, Naperville, IL) and then treated with 50 ng/ml HGF in 2% FBS media for the time indicated. Cells were washed with PBS, stained for 30 min at 37°C with 5 µg propidium iodide in PBS containing 0.1% Triton X-100, 0.1 M EDTA and 25 U/ml Rnace-It (Stratagene, La Jolla, CA), washed again and mounted with Aqu-polymount (Polysciences, Warrington, PA). Confocal images were collected with a Bio-Rad (Hercules, CA) MRC 1024 scan head mounted on a Nikon Optiphot microscope with a x20 planapochromat lens. Excitation was provided by a krypton–argon gas laser. Hard copy prints were produced with a Codonics NP-1600 printer.

FACS analysis
Cells were washed with PBS, pH 7.4, treated with 0.04% trypsin and 1% EDTA in PBS for 5 min at 37°C and fixed for 60 min at 4°C with ethanol. Cells were then treated with RNase and stained with 0.1 mg/ml propidium iodide containing 0.1% Triton X-100 and 0.1 M EDTA. Single cells were gated on FL-2 width and FL-2 area dot plot using a FACScan flow cytometer, and further analyzed to measure subdiploid/apoptotic population (Cellquest) and cell cycle distribution (MODFIT). Apoptosis was confirmed by microscopic observation of cells stained with propidium iodine (13).

Kinase assays
Cells were seeded at 2x104 cells/cm2 into 100 mm dishes. Twenty-four hours after plating, cells were equilibrated in serum-free media for up to 24 h. HGF (50 ng/ml) was added and cells incubated for 5, 15 and 30 min. After treatment, cells were washed 3 times with buffer A (137 mM NaCl, 20 mM Tris–HCl, pH 7.4, 1 mM CaCl2, 1 mM MgCl2 and 0.1 mM sodium orthovanadate), and harvested by scraping with a cell scraper in buffer A containing 1% NP-40 and 1 mM PMSF. Whole cell lysates were collected by centrifugation at 15 000 g, normalized for protein content and incubated overnight at 4°C with either anti-PI3K (06–195; Upstate Biotechnology, Lake Placid, NY), JNK1 (15701A; PharMingen, San Diego, CA) or ERK2 (SC-154; Santa Cruz) antibodies. The immune complexes were absorbed onto Protein A or G-Sepharose depending on the assay performed and washed. The PI3K assay was performed directly on the beads. The reaction was carried out for 10 min at 37°C in a buffer containing 10 mM Tris–HCl, pH 7.4, 150 mM NaCl, 5 mM EDTA, 20 µg phosphatidylinositol (PI) and 5 µl of [{gamma}-32P]ATP working stock solution (0.88 mM ATP containing 30 µCi of [32P]ATP, 3000 Ci/mmol and 20 mM MgCl2). The reaction was stopped by the addition of 20 µl 6 N HCl. Lipids were extracted and analyzed by thin-layer chromatography. ERK2 and JNK1 kinase assays were initiated by 1.5 mg/ml myelin basic protein (Sigma, St Louis, MO) or GST-ATF-2 fusion protein (Santa Cruz Biotechnology), respectively, and 5[{gamma}-32P]ATP. The reaction was terminated by addition of 5x Laemmli sample buffer. After boiling for 5 min, samples were electrophoresed on 10% SDS–polyacrylamide gel and visualized by autoradiography.

Electrophoretic mobility shift assay
Nuclear extracts were prepared as described previously (23). Oligonucleotides for NF{kappa}B and all antibodies were purchased from Santa Cruz Biotechnology. Nuclear protein (5 µg) and antibodies (1 mg) were added in a standard buffer (10 mM Tris, pH 7.5; 50 mM NaCl; 1 mM MgCl2; 20% glycerol) and pre-incubated at room temperature for 15 min. The 32P-end-labelled oligonucleotides (1x105 c.p.m.) and 2 µg of poly(dI-dC) were then added and incubated at room temperature for an additional 15 min. DNA complexes were resolved by gel electrophoresis on 5% polyacrylamide/1x TBE. The gels were dried and exposed to Kodak (Rochester, NY) X-AR film at –80°C for ~12 h.


    Results
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 Abstract
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 Materials and methods
 Results
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HGF induces growth inhibition and cell death in AFLB8 cells
We have previously reported that HGF exerts an inhibitory effect on the growth of aflatoxin transformed rat liver epithelial cells (AFLB8) and was capable of inducing apoptosis (13). Growth inhibition was confirmed by western immunoblot analysis which demonstrated that pRb was predominantly in the hypophosphorylated state in AFLB8 cells after treatment with 50 ng/ml HGF for 96 h (Figure 1AGo). In contrast, in normal RLE cells pRb is phosphorylated under the same conditions (Figure 1AGo). To determine the time course for induction of apoptosis, AFLB8 cells were incubated in the presence or absence of 50 ng/ml HGF. Within 72 h of treatment with HGF, 50% of the cells died which increased to nearly 60% after 96 h (Figure 1BGo). HGF-induced death was accompanied by chromatin condensation and fragmented nuclei, hallmark features of apoptosis (Figure 1C and DGo).



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Fig. 1. Demonstration of growth inhibition and apoptotic cell death. (A) Representative western immunoblot showing pRb phosphorylation status after HGF treatment for 96 h in RLE and AFLB8 cells. Whole cell lysates were used for western immunoblot analysis as described in Materials and methods. (B) Viability assay. AFLB8 cells were treated with or without HGF at 50 ng/ml for the times indicated. The percentage viability was assessed by trypan blue exclusion and plotted as mean ± SD. (C and D) Typical PI staining of AFLB8 cells with and without HGF treatment at 50 ng/ml for 72 h. Magnification x320.

 
HGF-mediated apoptosis is not regulated by bax, bcl-2 and bclX
To examine whether members of the bcl-2 gene family are involved in the control of HGF-mediated apoptosis, we examined the expression of bcl-2 gene family proteins by western immunoblot analysis (Figure 2Go). Bax, the proapoptotic member, was expressed in higher amounts in the AFLB8 cells than in the normal RLE cells, but did not increase with treatment. These results confirmed our previous finding that the 1 kb bax transcript was not modulated by either increased p53 expression or HGF treatment (13). Bcl-2, one of the anti-apoptotic protein partners for bax, was down-regulated in the normal RLE cells, but slightly increased in AFLB8 cells following HGF treatment. BclX-L, another anti-apoptotic protein partner for bax, was unchanged with HGF treatment in both normal and transformed cells. BclX-S was not detected (data not shown) in either cell line.



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Fig. 2. Western immunoblot analysis of bcl-2 family members during apoptosis by HGF in RLE and AFLB8 cells. Cells treated with HGF at 50 ng/ml HGF for 96 h were subjected to lysis and western blot analysis, and probed with antibodies specific for proteins shown on the right-hand side of the panel. Each experiment was repeated at least twice and a representative blot is shown above.

 
Regulation of other apoptotic genes
Recent observations strongly implicate cyclin gene expression as a mechanism for inducing cell death. Cyclin A, implicated in c-myc-induced apoptosis (2426), was increased in the AFLB8 cells treated with HGF. Constitutive expression of cyclin A in normal RLE cells was very low and not affected by HGF treatment (Figure 3Go). Cyclin G1, important in p53 mediated apoptosis in erythroleukemic cells (27) and a potential mediator of G2/M cell cycle arrest (23), was increased in AFLB8 cells undergoing HGF-mediated apoptosis, but was unaffected in RLE cells. The mitotic inhibitor, p27, which functions as a negative regulator of G1 progression and has been postulated to function as a mediator of growth factor-induced G1 arrest (28), did not change with HGF treatment in either cell line, although constitutive expression of p27 was much higher in AFLB8 cells than in RLE cells. Recently, the nuclear factor-kappa B (NF{kappa}B) rel family of transcriptional factors have been shown to be regulators of apoptosis (29,30). HGF increased NF{kappa}B protein expression in the AFLB8 cells, but had no effect on NF{kappa}B expression in normal RLE cells. However, NF{kappa}B activity in the AFLB8 cells was down-regulated after HGF treatment (Figure 3BGo). These data suggested that a lowered level of NF{kappa}B activity might reduce the protective effects of HGF and contribute to apoptosis in AFLB8 cells.



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Fig. 3. (A) Western immunoblot of other apoptotic genes. Cells treated with HGF at 50 ng/ml HGF for 96 h were subjected to lysis and western immunoblot analysis, and probed with antibodies specific for proteins shown on the right-hand side of the panel. Each experiment was repeated at least twice and a representative blot is shown above. (B) NF{kappa}B transcriptional activity in AFLB8 cells after 0, 8 and 24 h treatment with/without HGF (50 ng/ml). The gel shift assay was performed with 5 µg nuclear protein as described in Materials and methods.

 
Apoptotic signal mediated by HGF overrides inhibition of PI3K
It has been shown previously that Wortmannin, a selective inhibitor of PI3K, blocks growth factor-induced cell survival (31). We therefore tested the effect of Wortmannin on HGF-mediated apoptosis. Quantitative analysis by FACS (Figure 4AGo) showed a two-fold increase in the percentage of apoptotic AFLB8 cells after HGF treatment. In the presence of 3 µM Wortmannin alone, apoptosis was markedly decreased. In contrast, the combination of HGF and Wortmannin resulted in the same level of apoptosis as observed with HGF alone. These results suggest that PI3K does not deliver a survival signal in these cells.



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Fig. 4. (A) Effect of Wortmannin on HGF-mediated apoptosis in AFLB8 cells. Cells serum deprived (2%) for 24 h were treated with 50 ng/ml HGF, 3 µM Wortmannin or the combination for 96 h. Quantitation of apoptotic cells was determined by flow cytometry as described in Materials and methods. Each bar represents the mean ± SD of a representative experiment. (B) Effect of Wortmannin on HGF-mediated cell scattering. Colonies of serum-deprived AFLB8 cells (a) incubated for 24 h; (b and c) pre-incubated with 3 and 10 µM Wortmannin, respectively, for 30 min; (d) incubated with 50 ng/ml HGF for 24 h; (e and f) pre-incubated for 30 min with 3 and 10 µM Wortmannin, respectively, followed by stimulation with 50 ng/ml HGF for 24 h. (C) Effect of HGF on PI3K activity in RLE and AFLB8 cells. Cells serum starved for 24 h were stimulated with HGF at 50 ng/ml HGF for the times indicated. Equivalent amounts of whole cell lysate were immunoprecipitated and immune complexes assayed for PI3K activity as described in Materials and methods. The position of migration of phosphatidylinositol 3-phosphate [PI(3)P] is indicated. (D) Effect of Wortmannin on PI3K activity in RLE and AFLB8 cells. Cells were stimulated with or without HGF for 30 min. The immune complexes were pre-incubated with 3 (W3) or 10 (W10) µM Wortmannin for 30 min prior to performing the kinase assay.

 
Wortmannin has also been shown to block HGFmediated scattering, suggesting that PI3K may play an important role in this event (33). To evaluate the effect of Wortmannin on HGF-induced cell scattering, colonies of serum-deprived AFLB8 cells were pre-incubated for 30 min at 37°C with Wortmannin at 3 or 10 µM, and then stimulated with 50 ng/ml HGF. AFLB8 cells, following HGF stimulation, were scattered by 24 h (Figure 4BGod); whereas, AFLB8 cells in the presence of Wortmannin remained as tight colonies (Figure 4BGoe and f). Figure 4BGob and c show the effect of Wortmannin alone at 3 and 10 µM, respectively.

To evaluate the effectiveness of HGF as an inducer of PI3K activity, serum-starved AFLB8 cells were stimulated with HGF for 5, 15 and 30 min. Figure 4CGo shows the amount of activity measured in the PI3K immunoprecipitates from control, and HGF-treated AFLB8 and RLE cells, respectively. PI3K activity was not affected by HGF treatment in either cell line. However, Wortmannin was effective in inhibiting PI3K activity at all doses examined in both cell lines (Figure 4DGo).

HGF activates JNK1 in AFLB8 cells
Recently it has been shown that activation of JNK1 is critical for apoptosis mediated by the RTK, RON (15). Apoptosis induced by growth factor deprivation and ceramide treatment have also been shown to involve the activation of JNK (32). To determine whether JNK1 is activated during HGF-mediated apoptosis, we measured JNK1 activity after HGF stimulation in AFLB8 and RLE cells. As shown in Figure 5Go, JNK1 activity was increased in the AFLB8 cells within 5 min after HGF treatment and was maintained up to 15 min. In contrast, HGF had little to no effect in JNK1 activity in RLE cells. ERK2, whose activation is involved in the prevention of apoptosis, was activated within 5 min of HGF stimulation in both AFLB8 and RLE cells. HGF-mediated ERK2 activation was greater in the RLE than in the AFLB8 cells. Interestingly, EGF was an effective inducer of both JNK1 and ERK2 activities in both AFLB8 and RLE cells. These results are consistent with our recent finding that TGF-{alpha} induces programmed cell death in RLE cells (22). A similar pattern of ERK2 activation by TGF-{alpha} was also reported (T.Teramoto and S.S.Thorgeirsson, in preparation).



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Fig. 5. Time course of JNK1 and ERK2 activation after HGF and EGF stimulation in AFLB8 and RLE cells. Cells were collected at different time points after stimulation (0, 5, 15 and 30 min). JNK1 and ERK2 were immunoprecipitated from equal amounts of cell lysates. ATF2 and MBP phosphorylation were examined after addition of [{gamma}32]ATP and detected after SDS–PAGE by autoradiograghy.

 
EGF is not protective against HGF-mediated apoptosis
EGF, similar to HGF, is a cytokine that has the ability to induce DNA synthesis in primary rat hepatocytes in culture (34). To determine the effect of EGF on the AFLB8 cells, individual and combined treatments of EGF (10 ng/ml) and HGF were administered for up to 96 h. HGF-treated cells displayed extensive, flattened lamellipodial processes, whereas EGF-treated cells expressed long, thin processes and rarely exhibited flattened lamellipodia (data not shown). Other morphological alterations which were characteristic of HGF-mediated apoptosis, such as membrane blebbing, cell shrinkage and detachment, were also observed with EGF treatment (Figure 6BGo). Condensed chromatin and fragmented nuclei, hallmark features of apoptosis, are shown in Figure 6CGo. Quantitative analysis by FACS (Figure 6AGo) showed that EGF was capable of apoptosing AFLB8 cells to the same degree (two-fold) as HGF. The combined treatment of HGF and EGF was not additive, supporting the hypothesis that EGF/EGF receptor and HGF/c-met pathways co-operate at the cellular level.



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Fig. 6. (A) EGF induces apoptosis in AFLB8 cells. Cells serum-deprived (2%) for 24 h were treated with 50 ng/ml HGF, 10 ng/ml EGF or both for 96 h. Quantitation of apoptotic cells was determined by flow cytometry as described in Materials and methods. Each bar represents the mean ± SD of a representative experiment. Phase contrast (B) and propidium iodide staining (C) of AFLB8 cells treated with EGF (10 ng/ml) for 96 h.

 

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Although the role of PTKs in mediating apoptosis has been recently described, the specific mechanisms have not been clearly identified. Our results indicate that HGF-mediated apoptosis in chemically transformed rat liver epithelial cells involves a JNK1 pathway, perhaps including cyclin G1 and the down-regulation of NF{kappa}B activity, which is not dependent on p53 (13) or bcl-2 family members. Furthermore, EGF was also capable of inducing apoptosis in these cells, suggesting that EGF and HGF pathways may converge at the cellular level.

Using the same cell line which we found previously to be growth-inhibited and to undergo apoptosis upon stimulation with HGF (13), we show that pRB is expressed predominantly in the unphosphorylated state. These results were indicative of cells blocked in G1 of the cell cycle (35). In normal RLE cells, which are proliferating, pRb appears to be phosphorylated following HGF stimulation, thus releasing E2F, and allowing it to transactivate genes that regulate DNA replication and cell cycle progression. Since growth inhibition was observed in AFLB8 cells following HGF treatment, the potential role of mitotic inhibitors in regulating cell cycle progression in response to HGF was investigated. TGF-ß prevents pRb phosphorylation and induces G1 arrest mediated, in part, by the mitotic inhibitor, p27, which functions as a negative regulator of G1 progression (35). p27 protein expression was not modulated by HGF treatment in the AFLB8 cells, suggesting that the mechanism(s) involved in HGF-induced G1 arrest in AFLB8 cells is different than that described for TGF-ß. Preliminary results also suggest that p21, a p53 transcriptionally regulated mitotic inhibitor whose levels are elevated following DNA damage late in G1 (36), is similarly not involved. It is important to note, however, that although HGF-mediated apoptosis appears to coincide with growth inhibition in AFLB8 cells, EGF at a lower dose (10 ng/ml) than HGF (50 ng/ml) does not induce growth arrest despite being capable of inducing cell death in these cells. In addition, spontaneously transformed RLE cells (C4T) which were growth-inhibited by HGF did not undergo programmed cell death (13). The potential role of mitotic inhibitors in contributing to HGF G1 arrest needs to be further pursued.

Apoptosis or programmed cell death involves a series of consecutive, morphologically delineated phases that occur under both pathological, as well as normal physiological conditions and is required for normal development (4,37). In AFLB8 cells apoptosis occurs within 24 h after HGF treatment, with progressive cell death involving at least 50% of these cells until a plateau is reached at 96 h, suggesting that HGF-mediated apoptosis may be limited to a subpopulation of AFLB8 cells. Although investigations are continuing to determine whether HGF induces apoptosis in other transformed RLE cell lines, mouse erythroleukemia cells transfected with STK/RON (15) upon stimulation with macrophage stimulating protein (MSP) rapidly apoptose. These data support the notion that at least two PTK receptors in the HGF receptor subfamily can mediate an apoptotic signal.

STK/RON mediated apoptosis involved activation of JNK (15). JNK1 and 2 which are members of the MAP kinase-related family, and are induced by various agents including growth factors, phorbol esters, T cell activation signaling, UV light, protein synthesis inhibitors, and various cytokines (3840). JNK activation was found as well to be a requirement for ceramide and {gamma}-radiation (16,32) induced apoptosis. In rat PC-12 cells undergoing apoptosis after NGF withdrawal, Xia et al. (17) found a sustained activation of JNK concurrent with down-regulation of ERK activity. These results which propose a dynamic balance between growth factor-activated ERK and JNK pathways may be important in determining whether a cell survives or undergoes apoptosis. We demonstrate in the HGF-treated AFLB8 cells where JNK activation far exceeds the lower level of ERK activation that such a scenario may also be operative.

The bcl-2 family of related proteins can either suppress or promote apoptosis by interacting with and functionally antagonizing each other (41). In HGF-treated AFLB8 cells undergoing apoptosis, bax and bcl-x (S/L) expression, are unchanged, while bcl-2 expression increases slightly. The absence of an effect by HGF on many of the bcl-2 family members would suggest their lack of involvement in mediating apoptosis in this model similar to the results found with STK/RON. Iwana et al. (15) found that the constitutive expression of bcl-2 was very low in the mouse erythroleukemic cells and that MSP stimulation had no effect on bcl-2 expression. It was recently reported that overexpression of bcl-2 can be inactivated by phosphorylation, thereby resulting in cell death (42). The possibility exists that bcl-2 may be similarly phosphorylated in the AFLB8 cells and, therefore, inhibited. Alternatively, HGF may exert its apoptotic effects by fostering the formation of bax–bax homodimers, without an actual change in protein levels, or that the constitutive expression of bax may sensitize these cells to apoptosis. Overexpression of bax has been shown to increase the sensitivity of cells to chemicals and enhance radiation-induced apoptosis (43), both independent/dependent of p53 (43).

Downstream of the bcl-2 family members are the interleukin-1 converting enzyme (ICE) family of proteases, recently renamed caspases. The caspases have been implicated during apoptosis following growth factor deprivation, loss of contact with extracellular matrix, Fas/TNF-induced and cytotoxic T cell killing (44,45). Western immunoblot analysis (data not shown) of whole cell lysates from AFLB8 cells treated with or without HGF were performed and revealed that ICE protein levels did not change following HGF treatment in the AFLB8 cells. These results suggest that the ICE-protease may not be involved in HGF-induced apoptosis. A similar finding was also reported in the STK/RON study (15).

Cyclin A protein has been implicated in c-myc-induced apoptosis in cells under serum starvation or growth factor deprivation and in anchorage-independent cell growth (2426). In HGF-treated AFLB8 cells cyclin A expression is increased which could be the result of constitutively high c-myc expression (13). However, cyclin A can also mediate cell proliferation by associating with CDK2 p33 (4648) suggesting that the observed increase in cyclin A expression may simply be a reflection of a population of cells normally progressing through the cell cycle. Cyclin G1, which is transcriptionally activated by p53 (49) and, unlike cyclin A and other cyclin classes, has no apparent cell cycle dependency (50,51). We report an increase in cyclin G1 in AFLB8 cells under apoptotic conditions. Up-regulation of cyclin G1, as well as other p53-regulated cell cycle genes, such as p21 and gadd45, was important for the induction of apoptosis of an erythroleukemic cell line (27), while bax, fas and bcl-2 appeared not to be involved. In addition, we have found that overexpression of cyclin G1 as a fusion protein with green fluorescence protein (GFP) in a variety of cell types results in apoptotic cell death (23). The potential role of cyclin G1 as a mediator of apoptotic cell death continues to be pursued.

An important activity of growth factors which cannot be overlooked is the capacity to promote cell survival. In fact, HGF has been shown to counteract staurosporin and etoposide-induced apoptosis of a murine cell line derived from non-parenchymal liver epithelial cells (52,53). However, in normal RLE cells, HGF failed to protect against staurosporin or etoposide-induced apoptosis (data not shown), nor did it protect against TGF-ß1-induced apoptosis in the same cells (22). Phosphoinositide 3-kinase (PI3K) is a downstream effector of the HGF receptor (54), as well as other growth factor RTK receptors and is postulated to provide a possible survival pathway (55). Downstream of PI3K, Akt/protein kinase B (PKB) has been shown to be sufficient to promote cell survival upon growth factor withdrawal (56). The ability of Akt to promote survival, however, was dependent on its kinase activity and its upstream activator, PI3K (56). In our model, HGF was a poor inducer of PI3K activity in both the normal RLE and transformed AFLB8 cells and, therefore, is unlikely to transduce its survival signal through the PI3K/Akt (PKB) pathway. It is of interest to note as well that treatment of AFLB8 cells with Wortmannin did inhibit the ability of HGF to scatter these cells confirming previous reports in other cell systems (57,58). Our data would indicate that HGF scattering, but not protection is mediated by PI3K in the AFLB8 cells.

NF{kappa}B activity has also been shown to participate in cell survival. In murine hepatocytes p65 protein was shown to play a direct role in hepatocyte survival (29). Although in our study, in AFLB8 cells NF{kappa}B activity was down-regulated with HGF treatment despite an increase in protein expression. In RLE cells, which do not undergo apoptosis following HGF treatment, NF{kappa}B activity was up-regulated (data not shown). These data suggest the possibility that the lowered NF{kappa}B activity may contribute to the lack of protection by HGF and potentially advance apoptotic cell death in our system. In the report by Bellas et al. (29) inhibition of endogenous NF{kappa}B activity by micro-injection with a purified specific inhibitor or an antibody against p65 resulted in apoptotic cell death as well.

It is also of interest that EGF was capable of eliciting a similar apoptotic response in the AFLB8 cells as observed with HGF. The implication that co-operation exists between the EGF, or TGF-{alpha} and HGF pathways has been previously described in mitogenesis and liver regeneration (59). A recent study (60) provided further evidence that communication occurs between these pathways, whereby the constitutive expression of TGF-{alpha} leads to induction of c-met mRNA and protein, and the constitutive phosphorylation of c-met. The authors conclude that these changes in c-met expression and activation may be responsible for the enhanced capacity of chemically transformed rat liver epithelial cells that overexpress TGF-{alpha} to respond to the mitogenic effect of HGF (60). Since the combined treatments of HGF and EGF was not additive in AFLB8 cells, it would imply a convergence rather than co-operation between these pathways.

In summary, our work describes a potential pathway, perhaps cell specific, by which HGF/c-met signaling may elicit an apoptotic response. One hypothetical scenario would suggest that these transformed cells which are killed in a similar way by either HGF or EGF lack, or have modified survival pathways, i.e. PI3K or inhibited NF{kappa}B activity. These and other possibilities, including a role for cyclin G1 in apoptosis, continue to be investigated in an attempt to further define the role of HGF/c-met signaling in hepatocarcinogenesis.


    Notes
 
1 Present address: Semmelweis Medical University, I. Institute of Pathology, Üllói út 26, Budapest, Hungary Back

2 To whom correspondence should be addressed Email: snorri_thorgeirsson{at}nih.gov Back


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 

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Received June 2, 1998; revised November 19, 1998; accepted December 4, 1998.