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Am J Physiol Gastrointest Liver Physiol 294: G263-G275, 2008. First published November 8, 2007; doi:10.1152/ajpgi.00267.2007
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MUCOSAL BIOLOGY

Fas Ag-FasL coupling leads to ERK1/2-mediated proliferation of gastric mucosal cells

Hanchen Li,1,2,* Xun Cai,1,2,* Xueli Fan,1,2 Brian Moquin,1 Calin Stoicov,1 and JeanMarie Houghton1,2

1Department of Medicine, Division of Gastroenterology, and 2Department of Cancer Biology, University of Massachusetts Medical School, Worcester, Massachusetts

Submitted 13 June 2007 ; accepted in final form 2 November 2007


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
When cells within the gastric mucosa progress from metaplasia to dysplasia to cancer, they acquire a Fas Ag apoptosis-resistant phenotype. It is unusual to completely abolish the pathway, suggesting other forms of Fas Ag signaling may be important or even necessary for gastric cancer to progress. Little is known about alternate signaling of the Fas Ag pathway in gastric mucosal cells. Using a cell culture model of rat gastric mucosal cells, we show that gastric mucosal cells utilize a type II signaling pathway for apoptosis. Under conditions of low receptor stimulation or under conditions where apoptosis is blocked downstream of the death-inducing signal complex, Fas Ag signaling proceeds toward proliferative signaling. Under conditions favoring proliferative signaling, cFLIP is recruited to the Fas-associated death domain-like interleukin-1β-converting enzyme at the death-inducing signal complex and activates ERK1/2. ERK1/2 in turn activates NF-{kappa}B. ERK1/2 stimulates proliferation, whereas NF-{kappa}B activation results in upregulation of the antiapoptotic protein survivin, further promoting proliferation over apoptosis. These results suggest that factors that inhibit apoptosis confer a growth advantage to the cells beyond the survival advantage of avoiding apoptosis and in effect convert the Fas Ag signaling pathway from a tumor suppressor to a tumor promoter.

tumor promoter; cell culture; alternate signaling

CELLULAR CONTEXT AND ENVIRONMENTAL conditions are crucial in determining signaling outcome of Fas Ag-Fas ligand (FasL) interaction. Although it is best known as a death receptor (50), Fas Ag has also been shown to transduce nonapoptotic signals, including those for proliferation (1, 13), differentiation (48, 60), cytokine production (6, 41, 48, 51), and axonal regeneration in neurons (30). The decision to undergo apoptosis or utilize one of these alternate signaling functions of Fas is poorly understood but appears to depend on several factors, including the cell type, position within the cell cycle, activation status of parallel signaling pathways, and environmental context (60). The exact mechanisms involved remain unclear, and these likely differ between cell types and growth conditions.

Fas Ag signaling plays little role in normal development and homeostasis of the stomach, as mice deficient in either Fas Ag (lpr mutation) or FasL (gld mutation) develop normal gastrointestinal tracts during embryogenesis. In addition, mice lacking Fas Ag signaling do not develop gastric disease in the absence of factors known to cause disease in wild-type mice, such as infection or application of carcinogens (20). Under normal conditions, gastric mucosal cells express negligible Fas Ag and no FasL (21, 22), and the mucosa is devoid of inflammatory cells, limiting exposure to exogenous ligand. With autoimmune inflammation or inflammation secondary to Helicobacter pylori infection, cytokines such as TNF-{alpha}, IL-1β, and IFN-{gamma} upregulate surface Fas Ag expression (22) and initiate a massive influx of FasL-bearing inflammatory cells, resulting in apoptosis of the gastric epithelial cells (21). If the increase in apoptosis is not balanced by a commensurate increase in proliferation, ulcer disease and/or atrophic changes result (17, 20, 24, 37, 40, 49, 61, 62). This increase in proliferation may prevent or heal mucosal damage, and it has also been linked with an increase in cancer risk (16). Although attention has been given to pathological increases in apoptotic signaling within the stomach, little is known about the impact of blocking Fas-mediated apoptosis or the potential contribution of other forms of Fas Ag signaling such as proliferative signaling to gastric disease.

A hallmark of cancer is the ability to evade apoptosis (16, 45). Many antiapoptotic mechanisms employed by cancer cells involve alterations within the Fas Ag-FasL signaling pathway and consist of mutations altering expression or function of the receptor or ligand (4, 38, 47), aberrant death-inducing signal complex (DISC) assembly and function, and increased expression of inhibitor proteins (58, 63) or downregulation of caspases (36, 55). Despite strong pressure to avoid apoptosis, tumors that are completely deficient in surface Fas Ag receptor are rare, suggesting a pressure to preserve nonapoptotic signaling functions that may be important for tumor growth and survival (45). Indeed, there has been speculation that Fas may have tumor-promoting functions (45).

Helicobacter pylori is a type I carcinogen, and infection with this bacterium is a major risk factor for gastric adenocarcinoma (23). During infection, mucosal damage progresses through several well-defined stages, including atrophy, metaplasia, and dysplasia, culminating in adenocarcinoma in a subset of patients (7). Apoptosis peaks early in infection, as high Fas Ag-expressing cells are removed (20). As infection continues, gastric mucosal cell proliferation increases and is sustained (12). During this progression, Fas Ag expressing metaplastic and dysplastic cells acquire apoptosis resistance via a variety of methods, including altered surface receptor mobility and aggregation (31, 54), mutations in the Fas Ag death domain, and expression of inhibitor proteins (31, 33, 42, 44). Absolute loss of surface receptor is not seen. We postulate that gastric mucosal cells, which acquired apoptosis resistance, may be able to use alternate forms of Fas Ag signaling to enhance proliferation, thus offering cancer cells not only a way to avoid death but an additional mechanism for growth.

Here, we show that gastric mucosal cells possess two distinct Fas signaling pathways: a type II mitochondrial-dependent apoptotic pathway and a mitochondrial-independent proliferative pathway requiring assembly of the DISC and involving ERK1/2 and NF-{kappa}B activation. The apoptotic and proliferative signals occur in parallel, although with different initiation thresholds. Proximal portions of the pathway are shared with bifurcation at the DISC. Inhibition of apoptosis greatly increases the proliferative response. These findings suggest that, as gastric cells acquire mechanisms to evade Fas apoptosis, they may maintain Fas Ag proliferative signaling, which can contribute to tumor promotion. Defining the molecules involved in the propagation of proliferative signals at the DISC may provide targets for anticancer therapy.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Plasmids and reagents. Mouse Fas Ag cDNA was synthesized by RT-PCR (Superscript One-Step RT-PCR system, Platinum Taq DNA polymerase; Invitrogen, Carlsbad, CA) using the sense primer 5'-GAAGATCTGCAGACATGCTGTGGATCTGGGCTGTC-3' and antisense primer 5'-CTCGAATTCTCACTCCAGACATTGTCCTTCATTTTC-3' from crude template RNA (Trizol reagent; Invitrogen) prepared from an 8-wk-old male C57BL/6 mouse thymus. The 1-kb fragment was inserted into the BglII/EcoRI site of pMSCV-puro (BD Clonetech) to generate the plasmid pMSCVpuro-mFasAg. Rat Bcl-2 cDNA was amplified from the RNA of rat liver using sense primer 5'-GTACCTGCAGCTTCTTTCCCCGGAAGG-3' and antisense primer 5'-GCAGGTCTGCTGACCTCACT-3' and inserted into EcoRI site of pIRES plasmid (BD Clonetech). pCMV-iKBM was purchased from BD Clonetech, and pCMV-drop (empty vector) was generated by deletion of the iKBM sequence using HindIII/BamHI digestion followed by plasmid ligation.

The NF-{kappa}B luciferase vector was a kind gift from Dr. Ian Whitehead (University of Medicine and Dentistry of New Jersey), pcDNA3 was purchased from Invitrogen, and pcDNA3-cFLIPL was a kind gift from Dr. Michael J. Lenardo (Laboratory of Immunology, NIAID, National Institutes of Health, Bethesda, MD). HSV-TK promoter-driven Renilla expression plasmid was purchased from Promega (Madison, WI). The soluble recombinant FasL was purchased from Alexis (San Diego, CA), in which the extracellular domain is fused at the NH2 terminus to a linker peptide (26 aa) and a FLAG-tag. It binds to human, mouse, and rat Fas Ag.

Generation and characterization of cell lines. AGS cells (a human gastric adenocarcinoma cell line) were purchased from American Type Culture Collection and grown according to company protocol. RGM-1 cells (RIKEN Cell Bank, Tsukuba Science City, Japan) have been previously described (54). Cells were transfected via electroporation (960 µF, 300 V) with pMSCVpuro-mFasAg or empty vector and selected in 2.5 µg/ml puromycin (Calbiochem) for 2 wk. Receptor abundance of single cell clones were verified by RT-PCR and standardized against GAPDH. The rat GAPDH sense primer was 5'-TCTTCACCACCATGGAGAA-3', and the antisense primer was 5'-ACTGTGGTCATGAGTCCTT-3'; this gave a 231-bp product. Mouse Fas Ag sense primer was 5'-TGGAAACAAACTGCACCCTGAC-3', and antisense primer was 5'-TGCCCTCCTTGATGTTATTTTCTC-3', giving a 417-bp product. PCR products were resolved on 1.0% agarose gel with ethidium bromide. Membrane surface receptor expression of AGS cells and RGM-1 cell lines were confirmed by FACS analysis, and 25 RGM-1 clones were categorized as empty vector control (puro) or low-, moderate-, or high-expressing clones. Results have been confirmed in at least two additional lines for each category and are reported for one representative of each group [low (F10), moderate (F14), high (F21), and empty control (puro) cell lines].

Cell lines were stably transfected with a second vector: pMSCVneoBcl-2, pCMV-iKBM, pCDNA3-cFLIPL, or empty vector, and stable clones were selected.

Growth assays. Cell lines were grown with or without FasL (6.26–50 ng/ml) and with or without 10 µM VAD-fmk (broad caspase inhibitor) or 10 µM IETD-fmk (caspase 8 inhibitor) before the following assays. For 5-bromo-2'-deoxyuridine (BrdU) incorporation assay, G0-synchronized cultures were labeled with 13 µM BrdU in the last 4 h of a 24-h cultivation. Incorporated BrdU was evaluated by FITC-conjugated anti-BrdU staining and flow cytometry analysis. For cell growth curves, triplicate six-well plates were seeded with 1 x 105 G0-synchronized cells and subjected to growth conditions as indicated. Trypan blue-negative cells were counted daily with a hemocytometer. Apoptosis was evaluated with the annexin V-FITC apoptosis detection kit (EMD Biosciences, La Jolla, CA) according to the manufacturer's instruction. Mitochondrial depolarization was measured with 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolylcarbocyanine iodide. This substance exhibits potential dependent accumulation within mitochondria and is detected as a fluorescence emission shift from green (~529 nm) to red (~590 nm) secondary to the concentration-dependent formation of red fluorescent J-aggregates. Mitochondrial depolarization was detected as a decrease in the red-to-green fluorescence intensity ratio, dependent only on the membrane potential.

Protein immunoblotting. Proteins were extracted from cells in RIPA buffer (20 mM MOPS, 150 mM NaCl, 1 mM EDTA, 1% Nonidet P-40, 0.1% SDS, protease inhibitors including aprotinin, leupeptin, pepstatin A, PMSF, and phosphotase inhibitors), and protein concentration was measured by the bicinchoninic acid protein assay reagent (Pierce). Protein extracts (50 µg) were separated by 10% SDS-PAGE, transferred to an Immobilon-P membrane (Millipore, Bedford, MA), and incubated with specific primary antibodies, incubated with appropriate secondary antibodies, and analyzed with the SuperSignal chemiluminescent reagent (Amersham). Antibodies used included those directed against phospho-MEK, MEK, phospho-ERK, ERK, phospho-I{kappa}B{alpha}, I{kappa}B{alpha} (Cell Signaling Technology, Beverly, MA), survivin, Bcl-2 (Santa Cruz), and β-tubulin (Sigma).

Poly(ADP-ribose) polymerase cleavage assay. Cells were washed with ice-cold PBS containing protease inhibitors (1 mM PMSF and 0.5 mg/ml each of leupeptin and apoprotinin), resuspended in a reducing loading buffer (62.5 mM Tris, pH 6.8, 6 M urea, 10% glycerol, 2% SDS, 0.003% bromphenol blue, 5% 2-mercaptoethanol), sonicated on ice, resuspended, and incubated at 65°C for 15 min. Cells (1.5 x 105) were loaded on a 10% SDS polyacrylamide gel and analyzed by Western blot using the poly(ADP-ribose) polymerase (PARP) cleavage detection kit (Calbiochem).

DISC immunoprecipitation and analysis. Cells were treated with or without 50 ng/ml FasL fused to a FLAG-tag and protein supernatant prepared according to standard protocol and incubated with 40 µl of EZview red anti-flag M2 affinity gel (Sigma, St. Louis, MO) overnight at 4°C to precipitate DISCs according to protocol (54). Beads were collected by centrifugation, and the FLAG-fusion protein was eluted with 3x FLAG peptide (Sigma), separated by 10% SDS-PAGE, transferred onto PVDF membrane, and incubated with anti-cFLIP antibody (eBioscience, San Diego, CA) anti-Fas Ag antibody or anti-caspase 8 [Fas-associated death domain (FADD)-like interleukin-1β-converting enzyme (FLICE)] antibody (Santa Cruz) followed by horseradish peroxidase-conjugated secondary antibody (Amersham Biosciences). The immune complexes were visualized with the enhanced chemiluminescence reagent (Amersham).

Transient transfection and NF-{kappa}B reporter gene assays. Transient transfections were performed using Superfect (Qiagen, Valencia, CA) according to the manufacturer's protocol. Equal numbers of cells were seeded, grown to 60% confluence, and transfected with 0.005 µg of HSV-TK promoter-driven Renilla luciferase expression plasmid as internal control and 0.5 µg of NF-{kappa}B-driven firefly luciferase plasmid or 0.5 µg of the empty vector. Cells were stimulated the following day with FasL as indicated, washed, and frozen at –80°C for 30 min, transferred to room temperature, and incubated with 250 µl of 1x passive lysis buffer (Promega) for 20 min with constant shaking. Fifty microliters of the cell lysate were assayed in a Monolight luminometer (Monolight 3010 BD Pharmingen). Results are presented as relative promoter activity and are representative of triplicate experiments.

Nuclear extract preparation and EMSAs. Nuclear extract was prepared by using NE-PERTM nuclear and cytoplasmic extraction reagents (Pierce Biotechnology, Rockford, IL). Wild-type (5'-AGTTGAGGGGACTTTCCCAGGC-3') and mutant (5'-AGTTGAGGCGACTTTCCCAGGC-3') oligonucleotides were purchased from Santa Cruz. Wild-type oligonucleotides were end-labeled with T4 polynucleotide kinase (Promega, Madison, WI) and {gamma}-32P. Nuclear extract (10 µg) was mixed with poly(dI-dC) (Amersham) in a 20-µl reaction volume containing 10 mM HEPES (pH 7.9), 50 mM KCl, 1 mM EDTA, 1 mM DTT, 0.5% Nonidet P-40, and 5% glycerol and incubated at room temperature for 15 min; 100,000 cpm of [{gamma}-32P]dATP oligonucleotides were added and incubated for an additional 10 min. For competition analyses, 100-fold unlabeled wild-type or mutant oligo and NF-{kappa}B p65 or p50 antibody (Santa Cruz) were incubated with nuclear extract before the addition of radiolabeled oligonucleotides. Complexes were loaded onto a 5% nondenatured polyacrylamide gel and separated at 250 V for 1.5 h at 4°C. The gel was dried and visualized by autoradiography.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Human gastric adenocarcinoma cells express Fas Ag and proliferate in response to FasL. We predict that transformed gastric mucosal cells acquired apoptosis resistance, unmasking the ability to use alternate forms of Fas Ag signaling to enhance proliferation. This could offer cancer cells not only a way to avoid death but an additional mechanism for growth. To verify the clinical relevance of investigating the Fas Ag proliferative pathway, we first evaluated Fas signaling in human gastric adenocarcinoma cells. AGS cells, derived from a human gastric adenocarcinoma, express moderate levels of Fas Ag on the cell surface, as determined by FACS analysis (Fig. 1A). Exposure to 12.5 ng/ml FasL reliably and predictably increases proliferation of the AGS cells over control levels (Fig. 1B), confirming the presence of a Fas Ag-FasL proliferative signaling pathway in human gastric cancer and a relative resistance to apoptosis. To explore the signaling pathway further, we chose to use a nontransformed gastric mucosal cell line that could be expected to have signaling properties more closely approximating normal gastric mucosa.


Figure 1
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  		Fig. 1. Fas Ag expression in a human adenocarcinoma cell line (AGS) and a rat gastric mucosal cell line (RGM-1). A: AGS expresses surface Fas Ag by FACS analysis. B: AGS cells proliferate in response to 12.5 ng/ml Fas ligand (FasL). RGM-1 cells were stably transfected with a plasmid expressing mouse Fas Ag, and stable clones were isolated and categorized into 3 groups: low (F10), moderate (F14), and high expressing (F21). C: expression of Fas Ag by RT-PCR. D: expression of Fas Ag by FACS analysis. Empty vector (puro) is used as a negative control.

 
Surface expression of Fas Ag, in the absence of ligand, does not alter growth characteristic of gastric mucosal cells. Under normal conditions, the gastric mucosa expresses minimal Fas Ag (21, 22). With inflammation such as that which occurs with Helicobacter infection, Fas Ag surface receptor is markedly upregulated in many gastric mucosal cell subtypes, most notably in the chief and parietal cell compartments (20). Likewise, rat gastric mucosal cells (RGM-1 cell line) express low to negligible levels of surface Fas Ag (22). Marked upregulation of Fas Ag protein occurs after exposure of cultured cells to inflammatory cytokines IL-1β, TNF-{alpha}, and IFN-{gamma}, similar to in vivo findings (22), making this cell line an ideal model for studying gastric Fas signaling. To standardize surface receptor abundance and to avoid confounding cytokine-mediated cell signaling, we established rat gastric mucosal cells stably expressing discrete quantities of Fas Ag.

We isolated several Fas-expressing clones, which were grouped into four categories: those expressing essentially no (empty puro vector), low (represented by F10), moderate (represented by F14), and high (represented by F21) abundances of Fas Ag at the mRNA (Fig. 1A) and surface protein level (Fig. 1D; arrows depict shift to the right with antibody binding). The abundance of surface receptor expressed by low-expressing clones (F10) was comparable to the lowest amount, and that for the high-expressing cell (F21) was comparable with the highest amount detected in Helicobacter felis-infected mouse gastric tissue (54) as previously determined by our laboratory, likely representing the expression level of Fas Ag seen in mucous cells (low) and parietal and chief cells (high) (20, 54) in vivo during Helicobacter infection.

In the absence of ligand, Fas Ag expression alone does not alter cell growth characteristics. Cell lines were similar in morphology, similar in growth characteristics including doubling time and culture condition requirements, had similar patterns of annexin V staining and BrdU incorporation, and maintained a nontransformed phenotype (data not shown).

Expression level of Fas Ag leads to divergent growth response to ligand. The strength of stimulation of the Fas signal cascade can be regulated by the quantity of available surface receptor and the availability of ligand (reviewed in Ref. 60). To test these differences, we used the two extremes of receptor signaling: low receptor abundance (F10 cells) and low ligand stimulation (6.25–12.5 ng/ml FasL) and high receptor abundance (F21 cells) combined with high levels of ligand availability (25–50 ng/ml FasL). In response to ligand addition, low Fas Ag-expressing cells (for example, F10) showed a consistent growth advantage with higher rates of proliferation (total cell counts, Fig. 2A; BrdU incorporation, Fig. 2B) than control cells. Apoptosis was minimally increased in F10 cells from 6% to 10% by annexin V staining and showed an increase from 1.28% to 2.88% in the sub-G1 population on FACS analysis (Fig. 2B) as a result of increasing ligand exposure, blunting somewhat the effects of proliferation. Annexin V staining detects early membrane changes of apoptosis, which occur before DNA cleavage, accounting for the discordance between annexin V and sub-G1 population estimations of apoptosis. Sub-G1 estimations of apoptosis were consistent with annexin V results at 24 h (data not shown).


Figure 2
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  		Fig. 2. Gastric mucosal cells use Fas Ag for proliferative and apoptotic signaling. Shown are growth characteristics of rat gastric mucosal cell lines expressing low (F10) or high (F21) levels of Fas Ag or empty vector control (puro) cells exposed to FasL. A: cells were grown with or without 12.5 ng/ml FasL for up to 72 h, and cell viability was determined by Trypan blue exclusion. Cell counts are plotted as a percentage of control (without FasL). F10 (low Fas expression; B) and F21 (high Fas expression; C) cells were cultured with FasL at the indicated concentrations for 4 h, and apoptosis was assessed by both annexin V staining (top), which detected early apoptotic events, and cell cycle analysis of sub-G1 population (middle), which detected late apoptotic events. PI, propidium iodide. Bromodeoxyuridine (BrdU) incorporation (bottom) evaluates proliferation. Values in A are means ± SD from 3 separate experiments. Data in B and C are representative of 3 separate experiments. *P < 0.05.

 
Cells expressing high levels of Fas Ag (F21) did not proliferate in response to low-dose ligand exposure (12.5 ng/ml; Fig. 2A) but rather underwent apoptosis over the 3-day time course of the growth curve. At 72 h, only 20% of the original cell culture was viable (Fig. 2A). Cells expressing high levels of Fas Ag demonstrate an escalating dose response to ligand. With exposure to 25 ng/ml FasL, apoptosis was more rapid, with ~30% of cells apoptotic at 4 h (Fig. 2C), 60% at 24 h, and the entire culture depleted at 72 h (data not shown). Increasing ligand to 50 ng/ml resulted in nearly 100% apoptosis at 24 h (data not shown). These data suggest both receptor abundance and ligand availability modulate the signaling cascade, acting as a rheostat to adjust the susceptibility to apoptosis and the time over which it occurs. With exposure to minimal ligand, F21 cells increase BrdU incorporation (41–46%); however, this effect was decreased with quantity of ligand added (Fig. 2C) and with time of exposure (data not shown). At higher ligand levels, apoptosis depleted the high Fas Ag-expressing population and masked obvious proliferative signaling effects, suggesting concurrent apoptotic and proliferative signaling rather than an "either-or" phenomenon.

RGM-1 cells use a type II pathway for Fas signaling. Type I signaling is defined as when Fas Ag-FasL interaction results in sufficient caspase 8 activation to directly activate downstream caspases leading to apoptosis (3, 50, 60). Type II signaling involves insufficient caspase 8 activation and requires a mitochondrial amplification loop to proceed (3, 50, 60). Although it is generally thought that cells possessed either a type I or type II pathway, recent data suggest that a cell may have a preferred pathway while maintaining the ability to use the alternate pathway (14) if conditions become permissive. We tested whether gastric mucosal cells use type I or type II signaling by Bcl-2-mediated inhibition of the mitochondrial pathway and by assessing whether signaling patterns were dependent on receptor abundance.

Effective transfection with Bcl-2 was confirmed by Western blot analysis (Fig. 3A). Transfection with Bcl-2 did not change the growth characteristic or baseline level of apoptosis in any of the cell lines (Fig. 3, B and C); however, cells transfected with Bcl-2 were resistant to FasL-induced apoptosis. Four hours after the addition of 50 ng/ml FasL, a dose previously shown to induce rapid apoptosis in the F21 cell line, F21 cells became small, rounded, and detached from the plate (Fig. 3D), whereas the F21 cells transfected with Bcl-2 remained adherent and unchanged in appearance (Fig. 3E). Analysis of mitochondrial depolarization demonstrates that F21 cells rapidly depolarize the mitochondria (Fig. 3, F and H) after ligand addition, and this depolarization is successfully inhibited in cells transfected with Bcl-2 (Fig. 3, G and I). Bcl-2 inhibited Fas-mediated apoptosis in all RGM-1-derived cell lines regardless of the amount of surface Fas receptor or the quantity of ligand used. (Fig. 3J; F10, low surface receptor, and F21, high surface receptor cell lines), confirming RGM-1 cells use only a type II signaling pathway for apoptosis. Although enforced Bcl-2 expression inhibited apoptotic signaling, proliferative signaling remained intact, supporting the notion that proliferative signaling does not require mitochondrial involvement.


Figure 3
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  		Fig. 3. Gastric mucosal cells use type II apoptotic signaling pathway. A: Western blot analysis confirms low level of Bcl-2 in F21 cells and high levels in F21 cells transfected with Bcl-2. Mouse thymus is used as a positive control. B and C: RGM-1 cells expressing high levels of surface Fas Ag (F21; B) or those expressing high levels of both Fas Ag and Bcl-2 (C) have similar growth characteristics in culture. D: after 4 h of culture with 50 ng/ml FasL, cells expressing high levels of Fas Ag undergo apoptosis as evidenced by nuclear condensation, blebbing, formation of apoptotic bodies, and detachment from the plate. E: cells expressing high levels of Fas Ag and Bcl-2 are protected from FasL-induced apoptosis. Cells expressing Fas Ag have a rapid mitochondrial depolarization accompanying onset of apoptosis [compare F (without) and H (with ligand)]. Cells expressing Bcl-2 do not have mitochondrial depolarization with the addition of ligand (compare G and I). J: protection against apoptosis is plotted as percent survival 4 h after 50 ng/ml FasL addition. B–I are representative of 3 separate experiments. Values in J are means ± SD from 3 separate experiments. *P < 0.05.

 
Proliferative and apoptotic signaling diverge at the level of the DISC. Extrapolating from work done in other cell systems, there are several proposed pathways for Fas-mediated proliferative signaling (8, 25, 34, 43, 45, 46, 52, 60). These include signaling branching from FADD (28, 39, 56, 64) or, alternately, RIP (29) or DAXX (27) recruitment directly to the death domain of Fas Ag. This Fas-RIP and/or Fas-DAXX association has been proposed to activate NF-{kappa}B or result in JNK/Ask-1/MAPK-mediated AP-1 activation (5, 18, 35, 45), leading to proliferation. To test the pathway used in gastric mucosal cells, we used a dominant-negative (DN) construct, FADD-DN, that fails to recruit FLICE, effectively interrupting downstream apoptotic signaling. Transfection of our cells with FADD-DN completely blocked both proliferative (Fig. 4A) and apoptotic signaling (Fig. 4B), indicating at least that the proximal portions of these two pathways are shared and strongly suggesting that direct receptor-RIP or receptor-DAXX signaling is unlikely in this context. We show previously that cells expressing low level of surface antigen (F10 cells) proliferate when exposed to low-level ligand. In these cells, if we inhibit the mitochondrial amplification loop (see Fig. 3) or inhibit the caspase cascade with a broad caspase inhibitor (Z-VAD-fmk) or specifically inhibit caspase 8 with IETD, we do not adversely affect proliferation (Fig. 4C). Not surprisingly, when apoptosis is blocked by these agents, the level of proliferation is higher than that seen with addition of ligand alone (Fig. 4C).


Figure 4
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  		Fig. 4. Proliferative and apoptotic signaling bifurcate at the death-inducing signal complex (DISC). A: cells expressing low Fas Ag (F10) were treated with broad 10 µM caspase inhibitor (Z-VAD-fmk), transfected with dominant-negative Fas-associated death domain (FADD-DN; open bars), and/or exposed to 12.5 ng/ml FasL (solid bars), a dose previously shown to induce proliferation. Cell proliferation determined by viable cell counts are shown at 24 h as a percentage of control (untreated) cells. B: cells expressing high levels of Fas Ag (F21) were treated with 10 µM broad caspase inhibitor (Z-VAD-fmk), transfected with FADD-DN (open bars), and/or exposed to 50 ng/ml FasL (solid bars), a dose previously shown to induce apoptosis. Percentage of cells undergoing apoptosis at 24 h was assessed by annexin V staining. C: F10 cell line was grown for 3 days with 12.5 ng/ml FasL ± 10 µM VAD-fmk (broad caspase inhibitor) or 10 µM IETD-fmk (caspase 8 inhibitor), and cell counts were determined daily for 3 days. D: F21 cell line was grown for 3 days with 25 ng/ml FasL ± 10 µM VAD-fmk (broad caspase inhibitor) or 10 µM IETD-fmk (caspase 8 inhibitor), and live cells were counted daily for 3 days. Data are presented as percent change relative to control cells (without FasL). E: F21 cells were transfected with cFLIP, FADD-DN, or empty vector pcDNA3 and grown in 25 ng/ml FasL. Viable cells were counted daily for 3 days. Data are presented as percent change relative to control cells (without FasL). F: F21-cFLIP cells were grown with 50 ng/ml FasL, and DISCs were immunoprecipitated and blotted for Fas Ag and FADD-like interleukin-1β-converting enzyme (FLICE) to confirm an intact DICS and for cFLIP to reveal FasL-cFLIP interactions at the DISC. Immunoprecipitations were performed before and 2 and 5 min after ligand addition. Values in A–E are means ± SD and representative of 3 separate experiments. *P < 0.05. F is representative of 3 experiments each using 25 or 50 ng/ml FasL, which gave similar results. Results for F are shown for 50 ng/ml FasL.

 
Cells expressing high levels of Fas Ag are sensitive to FasL-induced apoptosis. In these cells, caspase inactivation protects cells from a major portion of apoptotic signaling but does not protect entirely (data not shown) against apoptosis. Residual apoptosis signaling obscures any increase in proliferation in this cell population (Fig. 4D).

On the basis of these findings, the site of bifurcation of the proliferative and apoptotic signaling pathway is at the DISC. The DISC consists of FasL coupled with membrane-bound Fas Ag and recruited FADD and FLICE (procaspase 8). Additional adaptor proteins may dock at the DISC (5, 34), but their nature and function are poorly understood at present. We next tested the involvement of FLICE in signal decision. FLICE binds at the DISC as a dimer; FLICE cleavage and activation (activated caspase 8) then initiate apoptotic signaling. In the case of gastric mucosal cells and other type II signaling cells, caspase 8 activation is not sufficient to initiate apoptosis; rather, it requires a mitochondrial amplification loop to proceed. Nevertheless, caspase 8 activation at the DISC is crucial in initiating this loop. cFLIP (FLICE-like inhibitory protein) competes with FLICE, forming FLIP-FLICE heterodimers that bind at the DISC and render FLICE noncleavable. FLIP is normally present at low concentrations in RGM-1 cells (data not shown). FLAG-tagged FasL was used to initiate the DISC and pull down complexes for analysis. Assembly of the DISC was confirmed by identifying the key proteins FasL (used to precipitate the complex), Fas Ag, FLICE, and FLIP (Fig. 4F). cFLIP does not bind to Fas Ag or FADD in the absence of ligand. Rapidly after ligand addition, cFLIP is recruited and can be isolated in abundance from immunoprecipitated DISCs (Fig. 4F). Overexpression of cFLIP prevented apoptotic signaling, although it allowed unopposed Fas-proliferative signaling as evidenced by the increase in proliferation over baseline levels in the F21-cFLIP cells exposed to FasL (Fig. 4E). These data support that DISC assembly is required for proliferative signaling, and the apoptotic and proliferative pathways proceed in parallel.

MEK/ERK is activated by Fas Ag-FasL in a dose-dependent fashion. Members of the MAPK family have been implicated in Fas-mediated proliferative signaling in cells of the immune system. We found no evidence of p38 or JNK activation (data not shown) in our system. However, within 5 min of FasL exposure, phospho-MEK and phospho-ERK1/2 were detected (Fig. 5, A and B). ERK1/2 activation was dependent on both ligand concentration and receptor abundance, being much greater in the F21 than in the F10 cell line. In empty vector control cells (expressing only nominal Fas Ag), 12.5 ng/ml of ligand induced phosphorylation of MEK but did not lead to ERK phosphorylation. Higher levels of ligand (50 ng/ml) were able to induce only minimal phosphorylation of ERK in these cells (data not shown). F10 cells expressing low levels of Fas Ag had measurable and reproducible (albeit low) phospho-ERK detected with minimal ligand exposure (Fig. 5A). The highest activation of the ERK1/2 pathway was seen in F21 cells expressing high levels of surface receptor (12.5 ng/ml FasL; Fig. 5B) and exposed to high ligand levels (50 ng/ml, data not shown).


Figure 5
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  		Fig. 5. ERK is phosphorylated by FasL-Fas Ag interaction and requires FADD binding. Cell lines expressing empty vector (puro) or low (F10; A) or high (F21; B) Fas Ag were exposed to 12.5 ng/ml FasL for indicated times, and phospho-MEK, MEK, phospho-ERK1/2, and ERK1/2 levels were evaluated by Western blot. C: F21 cell line was transfected with empty vector (pcDNA3), FADD-DN, or cFLIP and grown with or without 12.5 ng/ml FasL for 10 min, and phospho-ERK1/2 and ERK1/2 levels were evaluated by Western blot.

 
Next, we asked whether ERK phosphorylation is dependent on functional FADD. Transfection of cells with FADD-DN prevented the phosphorylation of ERK1/2, confirming that FADD is upstream of ERK activation and that ERK activation requires an intact DISC. Cells transfected with FLIP had an increased proliferative response and were resistant to Fas-mediated apoptosis (Fig. 4E) concomitant with enhanced levels of ERK phosphorylation (Fig. 5C), supporting that FLIP may have a role in ERK activation through its binding to FADD, possibly through recruitment of adaptor proteins such as Raf-1 to the DISC (data not shown).

ERK1/2 phosphorylation is necessary for Fas-mediated proliferation and has antiapoptotic function. The MEK inhibitor U-0126 effectively blocks FasL-induced activation of ERK in both low (F10) and high (F21) Fas Ag-expressing cell lines (Fig. 6, A and D) and inhibits FasL-induced proliferation and BrdU incorporation, although it has little effect on cell growth under normal conditions (Fig. 6, B and C). F10 cells do not have significant apoptosis with the addition of ligand. This did not change with MEK inhibition at early time points (Fig. 6C) or with longer exposure time or higher ligand dose (data not shown). Cells expressing abundant Fas Ag on their surface have higher levels of phospho-MEK, which was effectively inhibited by U-0126. In these cells, MEK inhibition prevented BrdU incorporation, similar to that shown in the F10 cell line, and additionally increased the apoptotic response to ligand from 29 to 37% at 4 h (Fig. 6E). Together, these data suggest a role for MEK/ERK1/2 in proliferative signaling and a role in prosurvival/antiapoptosis at higher levels of receptor stimulation.


Figure 6
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  		Fig. 6. ERK activation is necessary for Fas-mediated proliferation. A: F10 cells were grown for 10 min under control conditions (lane 1, C), with 12.5 ng/ml FasL (lane 2, Fas), with the MEK inhibitor U-0126 (lane 3, U), or with both 12.5 ng/ml FasL and U-0126 (lane 4, U+F), and levels of phospho-MEK, MEK, phospho-ERK1/2, and ERK1/2 were determined by Western blot. B: cells were grown under conditions in A; viable cells, enumerated by Trypan blue exclusion, were recorded daily for 72 h. C: BrdU incorporation (top) and apoptosis (bottom; via annexin V staining) under the conditions indicated were measured at 4 h. Numbers indicate percentage of BrdU incorporation (top) or percentage of apoptotic cells (bottom). D: F21 cells were grown for 10 min under control conditions (lane 1), with 12.5 ng/ml FasL (lane 2), with the MEK inhibitor U-0126 (lane 3), or with both 12.5 ng/ml FasL and U-0126 (lane 4), and levels of phospho-MEK, MEK, phospho-ERK, and ERK were determined by Western blot. E: apoptosis under the conditions indicated was measured at 4 h. Numbers within box indicate percentage of apoptotic cells. All results are representative of 3 experiments.

 
Fas Ag-FasL activates NF-{kappa}B via ERK1/2. Activation of Fas Ag can induce NF-{kappa}B in specific cell lines and may be independent of cytotoxic signaling function (43). With the use of a combination of gel shift assays (Fig. 7A), luciferase reporter construct assays (Fig. 7, B and D) and quantification of phosphorylated I{kappa}B{alpha} (Fig. 7, C and E), we assessed activation of NF-{kappa}B in response to Fas Ag-FasL interaction. NF-{kappa}B activation was proportional to the amount of surface receptor abundance (Fig. 7, B and D), with minimal activation over base line in F10 cells and a significant four- to sixfold increase in NF-{kappa}B activity in the F21 cells. NF-{kappa}B activation occurred rapidly and preceded apoptosis.


Figure 7
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  		Fig. 7. NF-{kappa}B is activated by Fas Ag ligation. F10 or F21 cell lines were grown with 12.5 ng/ml FasL for the times indicated. A: NF-{kappa}B EMSA. Arrows denote supershifted bands. B and D: F10 (B) and F21 (D) cell lines were transfected with luciferase reporter construct and grown for the indicated times with 12.5 ng/ml FasL. Values, reported in relative light units (RLU), are means ± SD of 3 experiments. *P ≤ 0.05. C and E: Western blot analysis of F10 (C) and F21 (E) cells grown for the indicated times with 12.5 ng/ml FasL and blotted for the indicated proteins.

 
Inhibition of NF-{kappa}B activation through chemical block (ALLN) or via I{kappa}B{alpha} blunted Fas Ag-FasL-mediated BrdU incorporation and proliferation (Fig. 8, A and B) and substantially increased susceptibility to Fas-mediated apoptosis as measured by annexin V staining (Fig. 8C) and caspase activity (shown by PARP cleavage in Fig. 8G), suggesting that, in gastric mucosal cells, NF-{kappa}B has antiapoptotic functions. There are several suggested candidates for inducing Fas Ag-mediated NF-{kappa}B activation, including the MAPK cascade (32). In our system, ERK1/2 is phosphorylated within 5 min of Fas Ag-FasL ligation in direct relation to receptor abundance. Inhibition of ERK1/2 phosphorylation by the specific inhibitor U-0126 (Fig. 8D), but not other members of the MAPK family (data not shown), prevented NF-{kappa}B activation and increased apoptosis linking ERK1/2 activation and NF-{kappa}B activity. Additionally, survivin, an antiapoptotic protein, was rapidly and robustly induced in gastric mucosal cells after Fas stimulation (Fig. 8E), modulating the apoptotic response. Inhibition of ERK (data not shown) or NF-{kappa}B activation (Fig. 8F) prevented survivin expression along with an increase in caspase activity, as shown by PARP cleavage (Fig. 8G), and restored apoptosis sensitivity. We did not detect any alterations in Bcl-2 expression or XIAP expression with inhibition of NF-{kappa}B.


Figure 8
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  		Fig. 8. NF-{kappa}B induces survivin and opposes Fas-mediated apoptosis. A: F10 cells were transfected with empty vector (F10-CMV) or with an NF-{kappa}B inhibitor (F10 CMV-iKBM) and grown with or without 12.5 ng/ml FasL for 4 h. BrdU incorporation was measured and is shown as percentage of positive cells. B: cell growth curve under the same conditions as in A. C: F21 cells were transfected with empty vector (F21-CMV) or with an NF-{kappa}B inhibitor (F21-CMV-iKBM) and grown with or without FasL for 4 h. Apoptosis was determined by annexin V staining. D: F21 cells were transfected with the NF-{kappa}B reporter construct and treated with MEKK inhibitor U-0126 ± FasL as indicated. Inhibition of MEKK/ERK inhibits FasL-induced NF-{kappa}B activation. E: survivin expression (green) in F21 cells. F21 cells were transfected with empty vector (F21-CMV) or with the NF-{kappa}B inhibitor (F21-CMV-iKBM) and grown with or without FasL. Adherent cells were tested, and apoptotic floating cells were discarded. F: Western blot of survivin expression in cells collected in E. Survivin is constitutively expressed and upregulated with FasL exposure. G: inhibition of NF-{kappa}B prevents induction of survivin and is associated with increase in caspase activity as shown by poly(ADP-ribose) polymerase (PARP) cleavage.

 

   DISCUSSION
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There is increasing evidence for dual Fas signaling in several different cell systems and the signaling outcome appears to depend on several factors, including the threshold of activation (32) and the interplay of environmental factors. By way of example, hepatocytes are normally highly sensitive to Fas-mediated apoptosis; however, after partial hepatectomy, they become resistant to apoptosis and instead proliferate in response to FasL (8) as part of a regenerative response. Likewise, fibroblasts that are normally sensitive to Fas apoptosis proliferate in response to FasL under conditions of low receptor abundance and serum deprivation (2), reminiscent of conditions found during wound repair. These data become germane when considering signaling within premalignant and malignant tissues as one extrapolates data from wound healing to signaling in malignant tissues. Many cancers arise within areas of chronic inflammation and persistent tissue regeneration. The similar tissue environment seen within physiological regeneration and cancer may have similar effects on driving Fas Ag signaling outcomes. Although the factors responsible for the switch in Fas signaling outcome during wound repair and regeneration have not been elucidated, there are several candidate scenarios, with recent studies suggesting the majority of the regulation occurs on the cytoplasmic side and involves availability and activity of proteins of the DISC. For example, IFN-{gamma} dramatically increases the expression of major histocompatibility complex class II throughout the body, not only on bone marrow-derived cells but on cells of the gastrointestinal tract. Our group (54) has shown that coexpression of major histocompatibility complex class II inhibits lateral migration of the Fas receptor and oligomerization of complexes within lipid rafts and prevents apoptotic signaling. Other mechanisms for Fas apoptosis resistance include regulation of inhibitor of apoptosis proteins or FLIP and, less frequently, decreasing surface receptor expression. The loss of Fas-mediated apoptosis appears integral to tumor formation and progression; however, total loss of surface receptor is extremely unusual, supporting the notion that there is strong pressure for pathway preservation to utilize alternate signaling functions.

Within the luminal gastrointestinal tract, several chronic inflammatory disorders are associated with cancer, including chronic Helicobacter infection and gastric adenocarcinoma, long-standing inflammatory bowel disease and colon cancer, and esophageal adenocarcinoma arising in reflux-induced metaplastic tissue. With regard to gastric cancer, the inflammatory milieu most closely associated with progression of disease (10, 11) is responsible for the upregulation and activation of the Fas signaling cascade, which has been shown to be a central mediator of tissue damage (17, 20, 21, 24, 37, 40, 49, 61, 62), leading to atrophy and the loss of specialized cells, which precede adenocarcinoma. With time, overall levels of apoptosis within the gastric mucosa decline (20), and a population of proliferating cells arises that are apoptosis resistant, although these cells still express surface receptor (45). These observations suggest a common environment driving two separate signaling decisions, albeit in opposite directions, or, as our data suggest, that Fas apoptotic signaling is redirected toward proliferative signaling.

We show that gastric mucosal cells are able to use the Fas pathway for both apoptosis and proliferation. Furthermore, the proliferative pathway can be unmasked or augmented when cells gain apoptosis resistance, which is a common early event in gastric carcinogenesis (31). In gastric mucosal cells, both Fas apoptotic and proliferative signaling are initiated at the receptor, and signaling bifurcates at the DISC. Although it has been suggested that type I and type II signaling may be preferred but not absolute pathways, we found no evidence that gastric mucosal cells could signal directly (type I signaling) regardless of the abundance of receptor or the level of stimulation with ligand. In the rat gastric mucosal cells, a mitochondrial amplification loop triggers the caspase cascade, resulting in apoptosis (type II pathway), whereas activation of MEK/ERK1/2 at the DISC precedes proliferation. ERK activation is required for Fas-mediated BrdU incorporation and is responsible for the growth advantage seen in low Fas-expressing cells. ERK activation, if sufficient, also activates NF-{kappa}B. NF-{kappa}B does not appear necessary for proliferation because inhibition does not significantly affect Fas-induced BrdU incorporation. In this situation, NF-{kappa}B activation functions to oppose apoptosis through the upregulation of the antiapoptotic protein, survivin, thus protecting cells to a degree from apoptosis and favoring proliferation over apoptosis.

The ERK pathway has been described for preventing cell death induced by Fas in lymphocytes and constitutes a major regulator of T-cell death. The influence of ERK in other cell types has diverse outcomes, with the majority of reports implicating antiapoptotic effects (18, 25, 57). In these situations, ERK activity is induced by competing signaling pathways and not directly by Fas stimulation. Our findings show that direct ERK phosphorylation by Fas ligation is necessary for the proliferation signaling of gastric mucosal cells and allows regulation of the pathway independent of exogenous factors.

Unlike colon cancer, where the sequence of mutations driving transformation are known and consist of logical alterations in growth, adhesion, and cell death pathways, the mechanism of signaling changes leading to gastric cancer is unclear. Gastric cancers harbor many of the mutations found in other gastrointestinal tumors such as alterations in p53 (26), adenomatous polyposis coli gene (53) deleted in colorectal cancer gene (53), VEGF (53), cyclooxygenase-2 (53), and β-catenin (53); however, these mutations appear to accumulate after transformation. In addition, although they may give growth and metastatic advantage, they cannot, by temporal occurrence, be instrumental in initiating gastric cancer. Previous work from our laboratory (20, 22) demonstrated that alterations in Fas Ag signaling are involved in gastric cancer progression, which occurs as an early event. Apoptotic signaling is attenuated through a variety of mechanisms (20, 54), and alterations that switch Fas from a tumor suppressor (through induction of apoptosis) to a tumor promoter (rechanneling signaling to proliferation) may provide the early growth alterations necessary for malignant transformation.


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This study was supported by the American Cancer Society Institutional Research Grant IRG 93-033 and National Cancer Institute Grant RO1 CA-113564 to J. Houghton.


   FOOTNOTES
 

Address for reprint requests and other correspondence: J. Houghton, Dept. of Medicine, Univ. of Massachusetts Medical School, LRB 209, 364 Plantation St., Worcester MA 01605 (e-mail: jeanmarie.houghton{at}umassmed.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

* H. Li and X. Cai contributed equally to this work. Back


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294/1/G263    most recent
00267.2007v1
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