Dual role of MEK/ERK signaling in senescence and transformation of intestinal epithelial cells

Marie-Josée Boucher, Dominique Jean, Anne Vézina, and Nathalie Rivard

Canadian Institutes of Health Research Group on Functional Development and Physiopathology of the Digestive Tract, Département d'Anatomie et Biologie Cellulaire, Faculté de Médecine, Université de Sherbrooke, Sherbrooke, Québec, J1H 5N4, Canada

Submitted 21 October 2003 ; accepted in final form 22 December 2003


    ABSTRACT
 TOP
 ABSTRACT
 MATERIAL AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The mitogen-activated protein kinase cascade operates downstream of Ras to convey cell-surface signals to the nucleus via nuclear translocation of ERK1 and ERK2. We and others have recently demonstrated that activation of ERK1/2 by growth factors is required for proliferation of intestinal epithelial crypt cells. However, it remained to be established whether ERK1/2 activation alone was sufficient to trigger intestinal epithelial cell (IEC) proliferation. To this aim, retrovirus encoding the hemagglutinin-tagged MAPK/ERK kinase (MEK)1 wild type (wtMEK), the upstream activator of ERK1/2, or a constitutively active mutant of MEK1 (MEK1-S218D/S222D; caMEK) were used to infect nonimmortalized human normal intestinal epithelial crypt cell cultures [human intestinal epithelial cells (HIEC)] and rodent immortalized intestinal crypt cells (IEC-6). Stable expression of caMEK but not wtMEK in HIEC led to the irreversible arrest of cellular proliferation (premature senescence). Concomitant with the onset of cell-cycle arrest was the induction of the cyclin-dependent kinase inhibitors p21Cip, p53, and p16INK4A. By contrast, overexpression of caMEK in IEC-6 cells induced growth factor relaxation for DNA synthesis, promoted morphological transformation and growth in soft agar, and did not affect expression of p21Cip, p53, and p16INK4A. We provided evidences that ERK1b, an alternatively spliced isoform of ERK1, is activated and may contribute to the deregulation of contact inhibition cell growth and transformation of these cells. Constitutive activation of MEK in IECs can produce either premature senescence or forced mitogenesis depending on the integrity of a senescence program controlled by the cell cycle inhibitors p53, p16INK4A, and p21CIP.

intestinal epithelium; proliferation; cell cycle; p16INK4A; p53


THE HUMAN INTESTINAL EPITHELIUM is in a constant state of self-renewal, with complete mucosal turnover occurring every 3–5 days. Intestinal epithelial cells (IEC) are generated from a fixed stem-cell population functionally localized in the lower portion of the intestinal crypts, giving rise to four primary epithelial cell types: absorptive enterocytes, goblet cells, Paneth cells, and enteroendocrine cells (5). The differentiation of each cell type takes place as cells move either upward into the villus (absorptive, mucus, and endocrine cells) or concentrate downward at the bottom of the crypt (Paneth cells) (5). The mechanisms that direct these complex events and migration are most likely multiple and are currently poorly understood. Elucidation of these mechanisms would be invaluable not only in delineating normal cell processes leading to the differentiated phenotype but also in providing insight regarding abnormal processes, such as neoplasia formation, that can occur when these mechanisms go awry.

Mammalian cells express multiple MAP kinases that mediate the effects of extracellular signals on a wide array of biological processes. Three distinct MAPK cascades have been described, all linked to separate signal-transduction pathways resulting in the final activation of either ERK1/2, p38 {alpha}, {beta}, {delta}, or {gamma}, or JNKs 1, 2, or 3 (24). Depending on cellular context, extracellular signals are thought to elicit a specific cellular response (proliferation/differentiation/apoptosis) through the preferential activation of one of the MAPK cascades, all of which have distinct spectra of substrates (13).

In the majority of cell types, including intestinal epithelial crypt cells (2, 14, 30, 34, 53), mitogenic signals are relayed from the cytoplasm into the nucleus by nuclear translocation of the ubiquitously expressed ERK1 and ERK2, resulting in activation of a range of transcription factors such as Elk-1, c-Ets-1, and c-Ets-2 (60). The upstream activators of ERK1/2 (MEK1/2, Raf, and Ras), on the other hand, remain cytoplasmic and/or membrane-bound (49, 61). On strong and persistent agonist stimulation, the ERK-mediated signals are attenuated and even terminated through various mechanisms, including the early induction of specific MAPK phosphatases (MKPs) (10, 49, 61).

In proliferation-prone cells, all components of the signal-transduction cascade (Ras, Raf, MEK, ERK) are sequentially activated in response to serum growth factors. Ras has been shown to be crucial for cell proliferation, because expression of permanently active forms of this protein triggers the cascade and leads to fibroblast (20, 33, 63, 69) and epithelial cell transformation (52, 66). This indicates that Ras activity is sufficient to bypass growth factor requirements. Although mutant Ras is most frequently associated with human epithelial cell-derived tumors (8), the majority of Ras signal-transduction and -transformation studies have been performed in rodent immortalized fibroblasts (6). In immortal rodent lines, transformation by oncogenic Ras involves its ability to bind and activate a series of effector proteins, including Raf-1, phosphatidyl inositol 3-kinase and Ral.GDS (64, 65). Each of these proteins, in turn, activates distinct downstream targets, thereby producing different aspects of the transformed phenotypes. Although each of these effector pathways contribute to the transforming activity of Ras in immortal rodent fibroblasts, activation of the MAPK cascade is clearly sufficient (9, 12, 15, 39, 57). However, it has been previously reported that activated Raf-1, despite triggering constitutively elevated ERK1/2 activities, fails to cause morphological and growth transformation of immortal rat intestinal cells (RIE-1) (43). This observation could suggest that the relative contribution of the ERK pathway to cell transformation may be determined by genetic background. However, blockade of the MEK/ERK cascade suppresses growth of colon tumors in vivo, suggesting that the ERK activation is indeed involved in intestinal tumor progression (55).

Virtually all studies examining MEK/ERK signaling in intestinal systems have used rodent immortal or human tumorderived cell lines harboring unknown genetic alterations. Hence, the current study analyzed whether MEK/ERK activation alone is sufficient to trigger human cell proliferation and transformation, taking advantage of the recent generation of nonimmortalized human normal intestinal epithelial crypt cell cultures. The human normal intestinal epithelial crypt cell cultures [human IECs (HIEC)] human diploid cell line, derived by Perreault and Beaulieu (46) from normal fetal (14–18 wk of gestation) intestinal tissue, has a finite lifetime and thus provides a new avenue for investigating, in human cells, the potential implication of the MEK/ERK signaling pathway in IEC proliferation and transformation. The role of this pathway was also further investigated in IEC-6 cells, an established rat intestinal cell line that is immortalized but reported to display properties of normal epithelial cells (50, 53).


    MATERIAL AND METHODS
 TOP
 ABSTRACT
 MATERIAL AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Materials. [{gamma}-32P]ATP, [methyl-3H]-thymidine, and the enhanced chemiluminescence (ECL) immunodetection system were obtained from Amersham-Pharmacia Biotech (Baie d'Urfé, QC, Canada). Monoclonal antibodies against phosphorylated and active forms of ERK1/2 were from New England Biolaboratories (Mississauga, ON, Canada). Antibodies for the detection of cyclin D1/D2 (C-17), cyclin E (M-20), cdk4 (C-22), cdk2 (M2), p16INK4A (F-12), p27Kip1 (C-19), p21Cip/Waf (C-19), p53 (DO-1), ERK1 (C-16), and HA tag were from Santa Cruz Biotechnologies (Santa Cruz, CA). A second antibody for the detection of p16INK4A (Ab-8) was also used and purchased from Neomarkers (Fremont, ON, Canada). pRB antibody was obtained from Pharmingen (Mississauga, ON, Canada). The recombinant ERK2 protein was purchased from Upstate Biotechnology (Lake Placid, NY). Secondary antibodies used in immunofluorescence were from Chemicon (Mississauga, ON, Canada), whereas antibodies used for Western blot analysis were purchased from Sigma-Aldrich (Oakville, ON, Canada). Insulin was from Connaught Novo Laboratories (Willowdale, ON, Canada). All other materials were obtained from Sigma-Aldrich unless stated otherwise.

Cell culture. Nonimmortalized HIEC-6 were cultured in DMEM (Invitrogen, Burlington, ON, Canada) supplemented with 4 mM glutamine, 20 mM HEPES, 50 U/ml penicillin, 50 µg/ml streptomycin, 0.2 IU/ml insulin, and 5% FBS. HIEC cells were originally generated from normal fetal human small intestine at midgestation (46); they express typical features of the lower adult crypt region and appear unable to differentiate. The lifespan of this normal cell line is limited to 22–25 passages. The rat intestinal epithelial crypt cell line IEC-6 (50) was obtained from A. Quaroni (Cornell University, Ithaca, NY) and cultured in plastic dishes in DMEM containing 5% FBS, as described previously (53).

Virus production and retroviral infections. Wild-type (wt)MEK and constitutively active mutant of MEK1 (caMEK)-encoding cDNA were cloned after excision from pECE, filling with Klenow, and subcloning blunt downstream of the cytomegalovirus promoter of retroviral vector pLXIN. wtMEK, caMEK, and empty pLXIN vectors were used to produce viruses in HEK293T cells in cotransfection with helper amphotropic DNA vector (pAmpho; kindly provided by Dr. C. Asselin, Université de Sherbrooke). Cells (5–8 x 106) were transfected 4 h by the lipofection method using Lipofectamine 2000 (Invitrogen). Cell culture media (OptiMEM, Invitrogen) were collected 48 h after infection and filtered through Millex-HA filters (25 mm, 0.45 µm; Millipore, Bedford, MA). Subconfluent HIEC cells were infected with the viral suspension containing 4 µg/ml of polybrene (Hexadimethrine bromide H-9268, Sigma-Aldrich Canada, Oakville, ON, Canada) for 1 h under agitation at 37°C. The medium was then changed and selection for Geneticin (0.25 mg/ml; Bio Media Canada, Drummondville, QC, Canada) resistance was applied 24 h later for 10–15 days to obtain stable populations. All experiments shown were performed within 3 wk following selection.

Protein expression and immunoblotting. Cells were lysed in SDS sample buffer (62.5 mM Tris·HCl; pH 6.8, 2.3% SDS, 10% glycerol, 5% {beta}-mercaptoethanol, 0.005% bromophenol blue). Proteins (10–50 µg) from whole cell lysates were separated by SDS-PAGE in 7.5 or 10% gels. Proteins were detected immunologically following electrotransfer onto nitrocellulose membranes (Amersham-Pharmacia Biotechnology). Protein and molecular weight markers (BioRad, Mississauga, ON, Canada) were revealed by Ponceau Red staining. Membranes were blocked in PBS containing 5% powdered milk and 0.05% Tween-20 for 1 h at 25°C. Membranes were then incubated overnight at 4°C with primary antibodies in blocking solution, then with horseradish peroxidase-conjugated goat anti-mouse or anti-rabbit (1:1,000) IgG for 1 h. Blots were visualized using the Amersham ECL system. Protein concentrations were measured using a modified Lowry procedure with bovine serum albumin as standard (47).

DNA synthesis reinitiation. Subconfluent IEC-6/pLXIN, IEC-6/wtMEK, and IEC-6/caMEK cells were serum starved for 48 h in DMEM. Cells were then stimulated for 16 h with 0.1, 0.5, 1, and 5% FBS in fresh DMEM medium. At the end of the incubation, 1 µCi of [methyl-3H]-thymidine was added for an additional 6 h. Cells were subsequently fixed and washed three times with ice-cold trichloroacetic acid (5%), harvested with 0.1 N NaOH, and incorporated radio-activity was counted.

Growth assay. All experiments were performed starting with HIEC and IEC-6 cell populations at 14 days postselection and subsequently plated for growth assay in 12-well plates at a concentration of 7.5 x 104 cells/well for HIEC and in six-well plates at a concentration of 2 x 105 cells/well for IEC-6. Cell growth was measured after 5 days for HIEC cell populations and after 2, 5, 10, 15, 20, 25, and 30 days for IEC-6 cell populations using a cell particle counter.

Soft-agar growth assay. Anchorage-independent growth was evaluated as described by Diaz et al. (18). Briefly, cells were plated at 2 x 105 cells/well in complete DMEM containing 0.6% agar in six-well plates. Medium was added (500 µl/well) twice a week to maintain constant humidity. After 16 days, colonies were photographed under light microscopy. Images of colonies were also taken after staining with 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide (MTT; 0.5 mg/ml) for 3 h at 37°C.

Electron microscopy. Cell cultures were rinsed with PBS, prefixed for 15 min with a 1:1 mixture of culture medium (DMEM-BRL) and freshly prepared 2.8% glutaraldehyde in cacodylate buffer (0.1 M cacodylate-7.5% sucrose), then fixed for 30 min with 2.8% glutaraldehyde at room temperature. After two rinses, cultures were postfixed for 60 min with 2% osmium tetroxide in cacodylate buffer. The cells were then dehydrated under increasing ethanol concentrations (40, 70, 90, 95, and 100%, all 3 times each), then covered twice for 3 h with a thin layer of Araldite 502 resin (for ethanol substitution). Finally, the resin was allowed to polymerize at 60°C for 48 h. Specimens were detached from the plastic vessels, inverted in embedding molds, covered with Araldite 502 and repolymerized at 60°C for 48 h. Thin sections, prepared by ultramicrotomy, were contrasted with lead citrate and uranyl acetate and observed in a blind fashion on a JEOL 100 CX transmission electron microscope. All reagents were purchased from Electron Microscopy Sciences (Cedarlane, Hornby, ON, Canada).

Expression vectors and reporter constructs. Plasmid thymidine-luciferase, which contains a high-affinity E2F binding site in the thymidine kinase promoter (16), was purchased from Clontech (BD Biosciences Clontech, Palo Alto, CA). The pRL-SV40 Renilla luciferase reporter vector was obtained from Promega (Nepean, ON, Canada). The expression vectors for HA-tagged MEK1 (wtMEK) and caMEK (in which the Raf1-dependent regulatory phosphorylation sites S218 and S222 were substituted by aspartic residues) (12) were kindly provided by Dr J. Pouysségur (Université de Nice, France). The HA-wtMEK and HA-caMEK were subcloned into the retroviral expression vector pLXIN (Clontech).

Transient transfections and luciferase assays. Subconfluent HIEC were seeded in six-well plates and cotransfected by lipofection (Lipofectamine 2000, Invitrogen) with 0.1 µg of thymidine kinase-luciferase reporter, 0.25 µg of the relevant expression vector (pECE) containing the epitope-tagged hyperactivated MEK-1 (S218D/S222D), or wtMEK-1. The pRL-SV40 Renilla luciferase vector (Promega) was used as a control for transfection efficiency. Luciferase activity was measured 2 days later according to the Promega protocol.

Detection of senescence-associated {beta}-galactosidase activity. HIEC cells were infected with retrovirus encoding wtMEK or caMEK. Two weeks after selection, cells were seeded in 12-well plates. Senescence-associated {beta}-galactosidase activity was detected using a commercial kit from Cell Signaling Technology (Mississauga, ON, Canada). Briefly, cells were fixed in 2% formaldehyde/0.2% glutaraldehyde. After cells were washed, they were incubated overnight in the staining solution [5 mM potassium ferrocyanide, 5 mM potassium ferricyanide, 1 mg X-gal in dymethylformamide, 40 mM citric acid (pH 6), 0.15 M NaCl, and 2 mM MgCl2]at37°C. Development of a blue color was visualized under a microscope.

RT-PCR analysis. RNA was purified from cultured cells using the TRIzol reagent (Invitrogen). For amplification, starting from 5 µg of RNA as template, single-stranded cDNA was synthesized from total RNA using an oligo(dT) primer and RT-AMV (Promega). Aliquots of the diluted cDNA preparations (5 µl) were then used as templates for 100-µl PCR reactions with the primers described by Yung et al. (68): ERK1–850-S: TACCTACAGTCTCTGCCCTCTAAA and ERK1b-CT-AS: CTGGGGGCAAAGACAGT. Reactions were performed using 2.5 units of Taq polymerase (Qiagen, Mississauga, ON, Canada). Parameters for DNA amplification were 94°C for 30 s, annealing temperature (55°C) for 30 s, and 72°C for 30 s. Oligonucleotide primers used for DNA amplification were synthesized by Invitrogen. DNA amplification products were analyzed by gel electrophoresis on a polyacrylamide gel stained with ethidium bromide.

Immunofluorescence microscopy. HIEC cells grown on sterile glass coverslips were washed twice with ice-cold PBS. Cultures were then fixed in paraformaldehyde 3% for 15 min at room temperature, permeabilized with 0.1% Triton X-100 in PBS for 10 min, and blocked with PBS/BSA 2% (20 min at room temperature). Cells were immunostained for 1 h with p16INK4A antibody followed by a 30-min incubation with FITC-conjugated secondary antibody, both at room temperature. Negative controls (no primary antibody) were included in all experiments. Quantification of cell fluorescence staining intensity was performed using the Metamorph Meta Imaging Series software v. 5.0 (Universal Imaging, Downington, PA).

Data presentation and statistical analysis. Assays were performed in either duplicate or triplicate. Luciferase results were analyzed by the Student's t-test and were considered statistically significant at P < 0.05. Typical Western blots shown are representative of three independent experiments. Densitometric analyses were performed by using Scion Image 4.02 (Scion, Frederick, MA). Representative results of in situ indirect immunofluorescence from three independent experiments are shown.


    RESULTS
 TOP
 ABSTRACT
 MATERIAL AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Stable expression of wtMEK and caMEK in HIEC and IEC-6 cells. The biological consequences of HA-tagged wtMEK and caMEK overexpression were examined in cultured HIEC and rat (IEC-6) IECs after viral infection of their respective cDNA cloned in the retroviral vector pLXIN. The percentage of retrovirally transduced cells ranged between 60 and 80%, as estimated by parallel infections using viruses expressing the green fluorescent protein gene product. The pLXIN retroviral vector coexpressed a G418 resistance gene that allowed selection of pure populations of transduced cells within 10 days. As shown in Fig. 1A, the selected populations expressed a high level of hemagglutinin (HA)-tagged wtMEK and HA-caMEK, as determined by Western blot analysis. Immunofluorescence studies confirmed that wtMEK and caMEK were cytoplasmic enzymes (data not shown).



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Fig. 1. Expression and kinase activity of wild-type (wt) MAP/ERK kinase (MEK) and constitutively active (ca) mutant of MEK1 in human intestinal epithelial cells (HIEC) and IEC-6 cells. HIEC and IEC-6 cell populations stably expressing pLXIN, wtMEK, or caMEK were harvested 2 wk postselection. A: equal amounts of whole cell lysates were separated by 10% SDS-PAGE, and proteins were analyzed by Western blotting with specific antibodies to hemagglutinin (HA) tag. B: cell extracts (400 µg) were immunoprecipitated (Ip) with a specific antibody to HA. Levels of immunoprecipitated HA-wtMEK or HA-caMEK were analyzed by Western blotting. Kinase activity of HA-wtMEK and HA-caMEK is demonstrated by the phosphorylation of recombinant ERK2.

 

We next compared the kinase activities of both wtMEK and caMEK. Exogenous MEK activities were measured following immunoprecipitation with antibodies against HA tag and using recombinant ERK2 protein as substrate. Figure 1B shows that caMEK-expressing HIEC and IEC-6 cells exhibited ~30-fold more MEK activity compared with wtMEK-expressing HIEC and IEC-6 cells after normalization of expression levels.

Effect of wtMEK and caMEK expression on HIEC and IEC-6 cell proliferation. HIEC and IEC-6 cells are highly dependent on growth factor addition for reinitiation of DNA synthesis and cell proliferation. Therefore, expression of any active gene product that lies along the growth factor-signaling pathway is expected to relax the growth factor requirement for DNA synthesis and cell proliferation. The proliferative properties of these cell populations were first monitored by growth curves. HIEC and IEC-6 cells transduced for wtMEK or caMEK or empty vector (pLXIN) were plated at low density, and cell numbers were counted. As expected, control (pLXIN)- and wtMEK-expressing HIEC (Fig. 2A) cell populations grew steadily and reached confluence by day 5 postseeding. By contrast, caMEK-expressing HIEC cells did not proliferate and stopped accumulating well before reaching confluence (Fig. 2A). In addition, the state of pRb hyperphosphorylation was also analyzed, which, in the late G1 phase, is a landmark for cells passing the restriction point and entering S phase (41). In serum-deprived wtMEK-expressing HIEC, pRb was exclusively found in its hypophosphorylated state (Fig. 2B), whereas pRb inactivation (hyperphosphorylated form) became apparent 20 h after serum stimulation. In contrast, pRb hyperphosphorylation was never detected in caMEK-expressing HIEC cells, indicating that these cells remained arrested in G0/G1 phase and thus did not progress into the cell cycle (Fig. 2B). To further evaluate the impact of strong activation of MEK/ERK signaling on HIEC cell proliferation, previously characterized expression constructs for wtMEK and caMEK were transiently transfected with a plasmid construction containing the E2F-responsive element of the thymidine kinase promoter linked to a luciferase reporter gene (see MATERIALS AND METHODS), which represents a sensitive reporter of growth factor-induced cell cycle progression and S-phase entry (16). Ectopic expression of caMEK significantly inhibited thymidine kinase gene expression by 50% (Fig. 2C). In marked contrast, wtMEK expression had no significant effect on thymidine kinase expression compared with controls (empty vector). Taken together, these results indicate that strong and persistent activation of MEK/ERK signaling in nonimmortalized human intestinal crypt cells induces cell cycle arrest.



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Fig. 2. Effect of wtMEK and caMEK expression on HIEC and IEC-6 cell proliferation. A and B: cells were cultured as described in MATERIALS AND METHODS for 5 (HIEC) or 2, 5, 10, 15, 20, 25, and 30 days (IEC-6), and cell growth was measured by cell counting. Values are the means ± SE of 3 separate experiments performed in quadruplicate. C: HIEC cell populations and were serum-starved for 36 h, and then stimulated with 5% FBS for 4 and 20 h. Equal amounts of whole cell lysates were separated by 7.5% SDS-PAGE, and proteins were analyzed by Western blotting with specific antibodies to pRb (Rb-P: hyperphosphorylated Rb). D: IEC-6 cell populations were harvested 2, 5, 15, and 20 days after seeding. Cell extracts (60 µg) were separated by 7.5% SDS-PAGE, and proteins were analyzed by Western blot for pRb expression. E: Subconfluent HIEC were cotransfected with 0.1 µg of thymidine kinase-luciferase and 0.25 µg of the relevant expression vector (pECE) in the presence or absence of wtMEK or caMEK. Luciferase activity was measured 48 h thereafter according to the Promega protocol. The increase in luciferase activity was calculated relative to control levels of thymidine kinase-luciferase obtained in pECE-transfected cells, which was set at 1. Results are the means ± SE of at least 3 separate experiments. Significantly different from control (pECE; at *P < 0.05; Student's t-test). F: reinitiation of DNA synthesis in response to increasing concentrations of FBS was measured as described in MATERIALS AND METHODS. Each point represents the mean of triplicate values. Results are expressed as fold stimulation of thymidine incorporation obtained with 0% FBS in wtMEK-expressing cells.

 

In IEC-6, there were no significant changes detected in the growth rate of caMEK-expressing cells compared with pLXIN- and wtMEK-expressing cells during the initial exponential growth in medium containing 5% serum. However, a significant change in saturation density was clearly apparent in caMEK-expressing cells that continued to proliferate after reaching confluency, in contrast to pLXIN- and wtMEK-expressing cells, which became contact-inhibited at day 5 postseeding (Fig. 2D). Moreover, there was no Rb phosphorylation detected in postconfluent pLXIN- and wtMEK-expressing cells, indicating that these cells were contact-inhibited and arrested in the G1 phase. By contrast, in the cell populations expressing caMEK, a significant proportion of cells was permanently cycling after confluency as judged by the detection of hyperphosphorylated pRb (Fig. 2E).

Constitutively active mutants of MEK1 have been reported to induce growth factor relaxation when expressed in immortalized fibroblasts (12). Therefore, dose responses of FBS-induced reinitiation of DNA synthesis were consequently measured in populations of IEC-6 cells expressing pLXIN, wt-MEK, and caMEK. The serum dose responses from G0-arrested pLXIN- and wtMEK-expressing cells were identical (data not shown). In marked contrast to wtMEK, expression of caMEK induced a growth factor relaxation for DNA synthesis. Indeed, in cell populations expressing caMEK, a significant proportion of cells was permanently cycling in either total absence or in low concentrations of serum (0.1%), as judged by the high rates of thymidine incorporation (Fig. 2F). These results demonstrate that expression of a permanently active MEK is sufficient to disrupt growth control and allow autonomous cell cycling in immortalized intestinal epithelial crypt cells.

Effect of wtMEK and caMEK expression on HIEC and IEC-6 cell morphology. Morphological examination of HIEC cells arrested by caMEK revealed that cells remained attached to the plates for at least 1 mo (data not shown) and acquired a large and flat morphology (Fig. 3). These caMEK-expressing cells remained arrested in a metabolically active state for at least the duration of the observation period (>1 mo). No evidences of apoptosis (i.e., no observed cell detachment or nuclear fragmentation) was detected in caMEK-expressing HIEC cells 30 days after infection (data not shown). Moreover, these HIEC cells accumulated SA-{beta}-galactosidase (Fig. 3, see arrows), a biomarker for senescent cells (19), demonstrating that constitutive activation of the MEK/ERK cascade in human intestinal crypt cells produces premature senescence.



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Fig. 3. Effect of wtMEK and caMEK expression on HIEC cell morphology. Photomicrographs of HIEC cell populations containing wtMEK (1 and 2) or caMEK (3 and 4) stained for SA-{beta}-galactosidase activity (pH 6.0) at day 14 postselection. Arrows indicate accumulated SA-{beta}-galactosidase. Bars, 25 µM.

 

Because caMEK-expressing IEC-6 cells were growth factor relaxed, sensitivity of IEC-6 cells to transformation was determined by constitutively activated mutant of MEK. Few or no morphological changes were readily detectable in exponentially growing caMEK-expressing IEC-6 cells (data not shown). However, at higher cell densities, the caMEK-expressing cells formed foci in contrast to pLXIN- and wtMEK-expressing epithelioid cells (Fig. 4A, 1–3). The multilayered morphology in caMEK-expressing cells became apparent at cell densities in which pLXIN- and wtMEK-expressing cells were forming a monolayer of contact-inhibited cells (Fig. 4B, 1 and 2). Furthermore, caMEK-expressing cells but not wtMEK-expressing cells showed the ability to form colonies in soft agar (Fig. 4C). These results indicate that constitutive activation of MEK1 in IEC-6 cells leads to morphological transformation.



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Fig. 4. Effect of wtMEK and caMEK expression on IEC-6 cell morphology and transformation. A: 1–3: representative phase/contrast pictures of 10-day postconfluent wtMEK- and caMEK-expressing cells. Bars, 25 µM. B: 1 and 2: 15-day postconfluent wtMEK- and caMEK-expressing IEC-6 cells were fixed in glutaraldehyde and osmium tetroxide before epoxy embedding for electronic microscopy analysis. Magnification, x2,800. C: wtMEK- and caMEK-expressing cells were seeded in growth medium containing 0.6% agar during 16 days. 1 and 2, colonies were photographed under light microscopy. 3 and 4, images of colonies were also taken after staining with 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide (MTT; 0.5 mg/ml).

 

Effect of wtMEK and caMEK on endogenous ERK signaling. The consequences of stable expression of the active form of caMEK on activation of endogenous ERK were analyzed in resting and stimulated HIEC and IEC-6 cells. As shown in Fig. 5, A and C (lanes 1–6), serum stimulation of HIEC and IEC-6 cells exhibited rapid and maximal activation of ERK1/2 within 10 min and persisted for up to 2 h. Nonstimulated caMEK-expressing HIEC (Fig. 5A, lane 7) and IEC-6 (Fig. 5C, lane 7) cells showed a moderately increased basal level of ERK1/2 activities compared with pLXIN- and wtMEK-expressing cells. However, little or no additional phosphorylation of ERK was detected on serum stimulation of caMEK-expressing HIEC and IEC-6 cells (Fig. 5, A and C, lanes 7–9). These data indicate the possible induction of a feedback inhibition mechanism activated by caMEK in intestinal cells stimulated by growth factors, a phenomenon previously observed in rodent fibroblasts (11, 12, 15, 25).



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Fig. 5. Effect of wtMEK and caMEK on endogenous ERK signaling. A and C: HIEC and IEC-6 cells stably expressing wtMEK or caMEK were serum-starved for 36 h, then stimulated with or without (0) 5% FBS for 10 min and 2 h. B: cell lysates were prepared from asynchronously HIEC cells stably expressing wtMEK and caMEK. D: IEC-6 cells stably expressing pLXIN, wtMEK, and caMEK were harvested 2, 5 (confluence), 15, and 20 days after seeding. A–D: equal amounts of whole cell lysates were separated by SDS-PAGE, and proteins were electrotransferred onto nitrocellulose. Western blot analysis for active ERK1/2 activities and total ERK1/2 proteins was performed as described in MATERIALS AND METHODS. Arrowhead indicates an additional species with low electrophoretic mobility that could correspond to ERK1b. E, lane 1: PCR analysis was performed (without RT) with the oligonucleotide primers ERK1–850-S and ERK1-CT-AS and total RNA from intestinal mucosae as template. Lanes 2–4, RT-PCR analysis was performed with the oligonucleotide primers ERK1–850-S and ERK1-CT-AS and total RNA from IEC-6 cells expressing wtMEK and caMEK as well from intestinal mucosae as templates.

 

The molecular manifestation of wtMEK and activated MEK expression on ERK expression and activity was further analyzed in HIEC and IEC-6 cells cultured under normal conditions with 5% serum. As shown in Fig. 5B, caMEK-expressing HIEC cells consistently showed a slight increase in ERK1/2 activity levels (1.5-fold) compared with wtMEK-expressing cells. Expression of ERK proteins was similar in both HIEC cell populations (Fig. 5B). In IEC-6 cell populations, expression and activity of ERK1/2 remained unaffected in asynchronously growing and confluent pLXIN-, wtMEK-, and caMEK-expressing cells (Fig. 5D). However, an additional species with low electrophoretic mobility was detected with the antibody recognizing the biphosphorylated and activated forms of ERK1/2 in all three IEC-6 cell populations (Fig. 5D, see arrowhead). Of note, this higher molecular mass form was prominent in confluent caMEK-expressing cells and appeared to correlate nicely with IEC-6 cell proliferation. Indeed, this band dramatically decreased as soon as pLXIN- and wtMEK-expressing cells reached confluence to almost undetectable levels in postconfluent G1 arrested cells.

In a recent study, Yung et al. (68) cloned a 46-kDa ERK isoform, termed ERK1b, which is an alternatively spliced form of ERK1, containing a 26-amino acid insertion between residues 340 and 341 of ERK1. Unlike the uniform pattern of expression of ERK1 and ERK2, ERK1b was confined to several tissues including heart, brain, lung, and kidney. However, the expression of ERK1b in the intestinal epithelium still remained to be demonstrated. RT-PCR analysis was henceforth performed to evaluate ERK1 transcripts from rat intestinal mucosae and from wtMEK- and caMEK-expressing IEC-6 cell lines. Figure 5E illustrates the size of PCR products obtained with primers derived from the sequence of ERK1. Two amplified products were obtained: the expected 274-bp band (ERK1) and a second 352-bp band in intestinal mucosae as well as in IEC-6 cell populations, indicating that ERK1b was indeed expressed in IECs. This suggests that the slower-migrating form detected with the phospho-ERK antibodies (Fig. 5D, see arrowhead) may represent ERK1b. RT-PCR analysis demonstrates that neither wtMEK nor caMEK expression affected the expression of ERK1b (Fig. 5E). Taken together, our results demonstrate that, in contrast to ERK1 and ERK2, ERK1b phosphorylation levels are significantly enhanced in caMEK-expressing IEC-6 cells, suggesting that ERK1b could contribute to the deregulation of contact inhibition cell growth and transformation of these cells.

Effect of wtMEK and caMEK on senescence-associated proteins and G1-phase regulatory proteins. Cellular senescence and transformation are normally accompanied by a series of changes that together distinguish senescence from quiescence or transformation. These changes involve altered expression of cell cycle proteins including upregulation of p53, p21, and p16 for senescence (1, 4, 37, 42, 51, 56) and for intestinal cancer, upregulation of cyclins and/or cyclin-dependent kinases (3, 45, 54, 58), and downregulation of p27Kip1 (38, 59). Interestingly, caMEK induced p53 (~4-fold induction), p21 (~3.3-fold induction), and p16 (~3.7-fold induction) expression in HIEC cells (Fig. 6A). It should be noted that these effects require constitutively active MEK activation, because HIEC cells overexpressing a wtMEK allele grow normally and display normal low levels of p53, p21, and p16. These results confirm that the cell-cycle arrest produced by activated MEK is characteristic of cellular senescence. In contrast to HIEC, there were no significant differences in expression levels of p21 in IEC-6 cells expressing wtMEK or caMEK; however, after densitometric analyses, we found a modest decrease (~30%) in p53 protein levels in caMEK-expressing cells (Fig. 6A). Finally, Western blot analysis using two different antibodies failed to detect p16 protein in wtMEK- and caMEK-expressing IEC-6 cells as well as in parental IEC-6 cell line (data not shown).



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Fig. 6. Effect of wtMEK and caMEK on senescence-associated proteins and G1-phase regulatory proteins. A: Western blot analysis: cell extracts were prepared from HIEC and IEC-6 cells stably expressing wtMEK or caMEK. Extracts (20 µg) were separated by SDS-PAGE and transferred onto nitrocellulose membranes before Western blot analysis for HA-MEK1, p21Cip, and p53. Immunofluorescence analysis: HIEC cells stably expressing wtMEK or caMEK were fixed with 3% paraformaldehyde and permeabilized with a solution of 0.1% Triton X-100 before immunofluorescence staining for p16INK4A. Bars, 10 µm. B: IEC-6 cell populations were harvested 2, 5, 10, 15, and 20 days after seeding. Cell extracts (60 µg) were separated by 10% SDS-PAGE, and proteins were analyzed by Western blotting for cyclin D1, cdk4, cdk2, cyclin E, p27Kip1, and actin expression.

 

Although stable expression of caMEK provoked cell cycle arrest and senescence in HIEC cells, it otherwise promoted IEC-6 cell proliferation and transformation. Indeed, in IEC-6 cell populations expressing caMEK, a significant proportion of cells were permanently cycling after confluency (see Fig. 2, D and E). In a first attempt to identify the "cooperating" factors that might be responsible for deregulating proliferation of caMEK-expressing IEC-6 cells, we analyzed the expression of cyclin D1, cyclin E, cdk4, and cdk2 because the progression through the G1 phase and the G1/S phase transition are orchestrated by cyclin D/Cdk4,6 and cyclin E/Cdk2 complexes. As shown in Fig. 6B, cyclin D1, cdk2, and cdk4 proteins were significantly induced by approximately twofold (n = 3) in caMEK-expressing IEC-6 cells after confluency (occurring at day 5 postseeding) compared with wtMEK-expressing cells. No significant modulation of cyclin E expression was found, however. We also analyzed p27Kip1 expression, which is known to play a pivotal role in the G1-to-S phase transition and which is also involved in growth arrest in response to cell-cell contact in various cell types (48), including IEC-6 and HIEC cells (17, 53). As shown in Fig. 6B, p27Kip1 expression was induced as soon as wtMEK-expressing cells reached confluence (day 5 postseeding), an expression that progressively increased during postconfluence. By contrast, expression of p27Kip1 was attenuated by ~39% at day 5 postseeding and by 80% at day 20 postseeding in caMEK-compared with wtMEK-expressing cells. Finally, with the exception of cyclin D1, cdk4, cdk2, and p27Kip1, we did not observe any difference in expression levels of other cell-cycle regulatory proteins including E2F1, E2F4, p107Rb, and p130Rb (data not shown).


    DISCUSSION
 TOP
 ABSTRACT
 MATERIAL AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
It has been previously demonstrated that activation of the MEK-ERK pathway is required for S-phase entry (2), p27Kip1 downregulation (53), and proliferation (14, 30, 34, 53) of IECs. Herein, we describe the consequences of stable expression of constitutively active MEK1 mutant on growth factor-dependent signaling events in immortalized (IEC-6) and nonimmortalized (HIEC) IECs. First, expression of the constitutively active form of MEK1 induced senescence in nonimmortalized human intestinal cells, whereas it was found to be sufficient to mimic growth factor signals and to deregulate growth control of immortal rodent IEC-6 cells. These conclusions are based on the fact that expression of caMEK 1) inhibited HIEC cell proliferation but induced growth factor relaxation for DNA synthesis in IEC-6 cells; 2) prevented cell cycle entry (e.g., hyperphosphorylation of pRb; thymidine kinase gene expression) of HIEC in response to serum but was sufficient for cell-cycle progression in an IEC-6 cell population deprived of growth factors; 3) induced the accumulation of SA-{beta}-galactosidase, a biomarker for senescent cells, in HIEC but promoted morphological transformation and growth in soft agar in IEC-6 cells; and 4-enhanced the expression levels of cell cycle inhibitors p21, p53, and p16 in HIEC but not in IEC-6 cells.

The results presented herewith clearly demonstrate that MEK activation can produce two precisely opposite outcomes in IECs: cell cycle arrest or forced mitogenesis. This may be dependent on the integrity of the senescence program controlled by p21 and mostly by p53 and p16. It has recently been shown that oncogenic Ras requires immortalizing changes to promote oncogenic transformation (37, 56). Indeed, because immortal rodent cell lines have already lost aspects of the senescence program, they are transformed by oncogenic Ras alone (56). Recently, Lin et al. (37) demonstrated that constitutive activation of MEK1 arrests primary murine fibroblasts but forces uncontrolled mitogenesis and transformation in cells lacking p53 or p16. The present data are in total agreement with these studies performed in fibroblasts. Indeed, expression of activated MEK in nonimmortalized IECs (HIEC) resulted in induced expression of p21, p53, and p16 and in senescence, whereas expression of the same mutant in rodent immortalized IEC-6 cells resulted in growth factor relaxation and transformation. No induction of p21 and p53 expression was observed by activated MEK, and we were unable to detect p16 expression in IEC-6 cells with two different antibodies. The failure to detect p16 expression in IEC-6 cells is not surprising, because it has been reported that several nontumorigenic immortalized cell lines also lack functional p16 protein (29, 51). In fact, its silencing is now recognized as the second most common molecular defect in immortalization and cancer (27). The Ink4a/Arf locus, which encodes the tumor suppressors p16Ink4a (p16) and p14Arf [alternative reading frame (ARF); the mouse homolog is called p19ARF], is often methylated in a broad range of common human solid tumors, including carcinomas of the colon and breast. A considerable body of circumstantial evidence supports the notion that such methylation is relatively specific and results in functionally significant gene inactivation. Recently, it has been demonstrated that p16INK4A methylation exists in the immortal IEC-18 cell line, an IEC line close to the IEC-6 cell line (26). In addition to methylation, frequent point mutations of p16 and homozygous deletion of the locus were noted in a variety of tumor cell lines (31). p53 Protein levels were barely detected in either control or wt-MEK- and caMEK-expressing IEC-6 cells. Moucadel et al. (40) recently reported that p53 is active in IEC-6 cells. Under normal cell conditions, p53 is maintained at low levels because it has a very short half-life. However, in the presence of DNA damage or oncogenic stresses, p53 accumulates rapidly through posttranscriptional mechanisms and gains full potential as a transcriptional activator (56). It has been shown that abnormal proliferation induced by oncogenes such as Ras results in deregulated E2F1 activity, which induces ARF expression. This ARF expression, in turn, stabilizes p53 and leads to cell-cycle arrest or apoptosis unless there is occurrence of a second lesion such as a mutation in ARF itself or p53 (7, 44). Methylation of the p16 and/or Arf promoters in the Ink4a/Arf locus has been found in about one-half of colon carcinomas and adenomas (26), suggesting that the locus may suppress early stages of development of this tumor type. Furthermore, it has been reported (22) that ARF is epigenetically inactivated in several colorectal cell lines and its expression is restored by treatment with demethylating agents. Therefore, one could speculate that the failure to induce p19ARF expression in caMEK-expressing IEC-6 cells may explain the failure to induce p53. Restriction digesting-based methylation analysis and methylation-specific PCR are required to determine the methylation status of the p16 and ARF promoters in IEC-6 cells. Therefore, our results confirm the view that premature senescence may act as a fail-safe mechanism to limit the transforming potential of excessive Ras/MAPK signaling not only in fibroblasts but also in epithelial cells, the same cell type from which the majority of Ras mutation-positive cancers arise (8).

In a first attempt to identify the factors possibly responsible for deregulating proliferation of IEC-6 cells expressing activated MEK1, we have analyzed the expression of other cell-cycle regulatory proteins and found an upregulation of cyclin D1, cdk4, and cdk2 expression and a strong reduction of p27Kip1 levels in caMEK-expressing cells. Abnormalities in the expression of these cell-cycle regulatory proteins have been previously reported in tumors of the small bowel (3) and colorectal carcinoma (38, 45, 54, 58, 59). Regarding p27Kip1, the result is in agreement with our previous data showing that MEK/ERK cascade is required for p27 downregulation and S-phase entry in fibroblasts and IEC-6 cells (53). Moreover, it has been reported that reduced expression of p27Kip1 is a predictive factor of a poor prognosis for patients with breast and colorectal cancers (38, 59). Taken together, it is therefore likely that upregulation of cyclin D1, cdk4, and cdk2 expression, reduction of p27Kip1 levels, and the noninduction of p16, p21, and p53 in caMEK-expressing cells may all contribute in maintaining a significant proportion of pRb in an hyperphosphorylated state after confluency. These actions are complementary, because they all cooperate toward the induction of cyclin-dependent kinases and in the stimulation of cell proliferation.

It was previously reported that, although necessary for Ras transformation, Raf/ERK cascade alone is not sufficient to cause transformation of RIE-1, an immortalized rat IEC line (43). This observation could suggest that the relative contribution of the ERK pathway to cell transformation may be determined by genetic background. However, blockade of MEK by the highly potent and selective inhibitor of MEK, PD-184352, suppresses growth of colon tumors in vivo, suggesting that the MEK activation is indeed involved in intestinal tumor progression (55). Whereas some studies (28, 35, 36) have reported that ERK1/2 activities are elevated in intestinal tumors, others (21, 62) have shown no modulation or inhibition. In the present study, basal activity of ERK1/2 was slightly to moderately enhanced in both cell populations expressing the activated form of MEK1 (caMEK). However, stimulation of ERK activities in response to serum was not observed in either cell populations expressing caMEK. One plausible explanation is that cells permanently stimulated by autoactive MEK1 are desensitized via multiple mechanisms including the previously reported ERK-mediated feedback inhibition of MEK (11) and possible increased basal levels of MAPK phosphatase (10, 25, 49). Our preliminary results indicate that the MKP-1, -2, and -3 are not induced in caMEK-expressing cells. Hence, an alternative and yet unknown MAPK phosphatase is likely responsible for the prevention of ERK induction by serum growth factors in our cells. A plausible candidate is PTP-ER (closely related to PTP-SL), which was found to play a key role in Ras1 signaling in Drosophila (32). It should be noted, however, that transduction of activated MEK in IEC-6 cells did not initially enhance ERK1 and ERK2 activities in the presence of serum, although after 5–6 wk, these cells had elevated ERK1/2 activities as well as a slightly more refractile and fibroblastic appearance (data not shown). These disparate observations suggest that selective pressures rather than direct signaling may determine the level of ERK activity in transformed cells and that these selective pressures may vary depending on cellular and genetic backgrounds.

In our view, the most interesting and novel finding of this study is the detection, by the antibody recognizing phosphorylated and activated forms of ERK, of a band with a molecular mass of 46 kDa in our IEC-6 cell populations. We believe that this 46-kDa band is in fact ERK1b, an alternatively spliced isoform of ERK1, which has recently been cloned and characterized by Yung et al. (67, 68) because 1) RT-PCR analyses performed with specific primers derived from the sequence of ERK1 and also used by Seger's group (68) revealed the expression of an ERK1b transcript (352 pb) in intestinal epithelium and IEC-6 cell lines; 2) under most circumstances, the kinetics of phosphorylation (activation) of this 46-kDa were similar to ERK1 and ERK2 (see in serum-stimulated cells and in asynchronously growing cells); and 3) phosphorylation levels of the 46-kDa band were markedly enhanced in IEC-6 cells expressing the constitutively activated mutant form of MEK1. Finally, as previously reported (68), the 46-kDa band was also recognized by Abs directed against the COOH terminus of ERK1 (weaker staining, see Fig. 5D).

Of particular note is that phosphorylation levels of ERK1b appeared to correlate very closely with IEC-6 cell proliferation. Indeed, in contrast to ERK1 and ERK2, ERK1b activity dramatically decreased as soon as pLXIN- and wtMEK-expressing cells reached confluence, to almost undetectable levels in postconfluent G1-arrested cells. In addition, ERK1b was prominent in confluent caMEK-expressing cells, which continued to proliferate well after confluence, indicating that ERK1b may contribute to the deregulation of contact inhibition cell growth and transformation of IEC-6 cells. Hence, it appears that in IEC-6 cells (Fig. 5D) as well as in other transformed cells (25), constitutive activation of the upstream components of the ERK cascade is not accompanied by a comparable activation of ERK1 and ERK2, at least initially. This suggests that these two latter isoforms may be under tight downregulation. However, ERK1b appears to escape this tight downregulation in IEC-6 cells and appears to be the major responsive ERK isoform. Interestingly, in Ras-transformed Rat1 cells, there was higher expression of ERK1b, which was also more responsive than ERK1 and ERK2 to various extracellular treatments in transformed cells (68). These observations are consistent with the fact that ERK1b is less sensitive to phosphatases (67). Hence, ERK1b may transmit exogenous signal under conditions in which ERK1 and ERK2 are downregulated on persistent stimulation.

Unfortunately, our RT-PCR and Western blot analysis never permitted us to detect Erk1b expression in HIEC cells. Seger's group (68) has already reported that this ERK isoform is abundant mainly in rat tissues, and much lesser amounts were detected in humans. Furthermore, expression of Erk1b protein was reported only in cancer cells and never in normal human cells. Nevertheless, we predict that even if Erk1b is stimulated by constitutively active MEK in these cells, G1 arrest will still occur because integrity of the HIEC senescence program remains intact (upregulation of p53, and p16 expression will still occur).

In summary, our data demonstrate that the constitutive activation of MEK in IECs can produce either premature senescence or forced mitogenesis depending on the integrity of the senescence program controlled by p53 and p16. In this view, mutations that deregulate the MAPK cascade provide an initial proliferative advantage but also accelerate senescence. However, cells acquiring mutations in p53 or at the INK4A/ARF locus escape senescence, thereby revealing the full mitogenic potential of the MAPK cascade. Consistent with this scenario, ras mutations often precede mutations in p53 or INK4A during tumor development (23). Finally, our findings are consistent with ERK1b being the major ERK isoform that mediates MEK1-induced IEC transformation.


    ACKNOWLEDGMENTS
 
We thank Pierre Pothier for the critical reading of the manuscript. N. Rivard is a recipient of a Canadian Research Chair in Signaling and Digestive Physiopathology. M.-J. Boucher is a student scholar from the Fonds pour la Recherche en Santé du Québec.

GRANTS

This research was supported by Canadian Institutes of Health Research Grants MT-14405 and GR-15186.


    FOOTNOTES
 

Address for reprint requests and other correspondence: N. Rivard, Département d'Anatomie et de Biologie Cellulaire, Faculté de Médecine, Université de Sherbrooke, Sherbrooke, QC, J1H 5N4, Canada (E-mail: Nathalie.Rivard{at}USherbrooke.ca).

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.


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