Negative Regulation of beta -Catenin Signaling by Tyrosine Phosphatase SHP-1 in Intestinal Epithelial Cells*

Cathia DuchesneDagger §, Stéphanie CharlandDagger §, Claude Asselin, Clara Nahmias||, and Nathalie RivardDagger **

From the Dagger  Département d'Anatomie et Biologie Cellulaire, Faculté de Médecine, Université de Sherbrooke, Sherbrooke, Québec J1H 5N4, Canada and the || Department of Cell Biology, Institut Cochin, 75014 Paris, France

Received for publication, January 14, 2003

    ABSTRACT
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Protein-tyrosine phosphatase SHP-1 is expressed at high levels in hematopoietic cells and at moderate levels in many other cell types including epithelial cells. Although SHP-1 has been shown to be a negative regulator of multiple signaling pathways in hematopoietic cells, very little is known about the biological role of SHP-1 in epithelial cells. In order to elucidate the mechanism(s) responsible for the loss of proliferative potential once committed intestinal epithelial cells begin to differentiate, the role and regulation of SHP-1 were analyzed in both intact epithelium as well as in well established intestinal cell models recapitulating the crypt-villus axis in vitro. Results show that SHP-1 was expressed in the nuclei of all intestinal epithelial cell models as well as in epithelial cells of intact human fetal jejunum and colon. Expression and phosphatase activity levels of SHP-1 were much more elevated in confluent growth-arrested intestinal epithelial cells and in differentiated enterocytes as well. Overexpression of SHP-1 in intestinal epithelial crypt cells significantly inhibited dhfr, c-myc, and cyclin D1 gene expression but did not interfere with c-fos gene expression. In contrast, a mutated inactive form of SHP-1 had no effect on these genes. SHP-1 expression significantly decreased beta -catenin/TCF-dependent transcription in intestinal epithelial crypt cells. Immunoprecipitation experiments revealed that beta -catenin is one of the main binding partners and a substrate for SHP-1. Taken together, our results indicate that SHP-1 may be involved in the regulation of beta -catenin transcriptional function and in the negative control of intestinal epithelial cell proliferation.

    INTRODUCTION
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INTRODUCTION
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Protein phosphorylation and dephosphorylation of tyrosyl residues are important events involved in the regulation of cell growth and differentiation. Protein-tyrosine phosphatases (PTPs)1 act both as positive and negative regulators of signal transduction (1). Numerous PTPs have been identified to date, including the extensively studied SHP-1 (initially designated as SHPTP-1, SHP, HCP, and PTP1C) and SHP-2 (initially designated SHPTP-2, Syp, PTP2C, and PTP1D). Whereas SHP-1 is highly expressed in hematopoietic cells and moderately in many other cell types such as epithelial cells, SHP-2 has a more widespread distribution (2-4). Both proteins have similar structures, comprising two tandem SH2 domains at the N terminus, a single central catalytic domain and a C-terminal domain. The SH2 domains recruit SHP-1 and SHP-2 to tyrosine-phosphorylated molecules, enabling dephosphorylation to be performed by the catalytic domain. Understanding the biological roles of SHP-1 has been vastly invigorated by the discovery that loss-of-function mutations in the gene encoding SHP-1 are responsible for the profound immunological dysfunction observed in mice homozygous for the motheaten (me/me) or allelic viable motheaten (mev/mev) mutations (5, 6). These mice express either no SHP-1 or a catalytically defective SHP-1 protein consequent to splice site mutations in the SHP-1 gene (6). These defects, in turn, engender severe hematopoietic disruption as exemplified by an enormous expansion and tissue accumulation of myeloid/monocytic cells that lead to patchy dermatitis, extramedullary hematopoiesis, splenomegaly, and hemorrhagic pneumonitis resulting in death at about 2-3 (me/me) or 9-12 (mev/mev) weeks. Lymphocyte ontogeny and function are also dramatically altered. Cells isolated from me and mev mice have enabled the identification of SHP-1 target proteins in hematopoietic cells (7, 8). SHP-1 has been shown to negatively regulate downstream signaling of the erythropoietin receptor and to dephosphorylate several target molecules such as c-Kit, the granulocyte/macrophage colony-stimulating factor receptor, the B and T cell antigen receptor, the adapter protein SLP-76, the cytosolic tyrosine kinase ZAP-70, and the lymphoid-specific Src family kinase Lck (7, 9-13). Very little is known on the other hand about the biological roles of SHP-1 in epithelial cells, although the existence of an epithelium-specific isoform of SHP-1 (the epithelial and hematopoietic variants differ in the sequence of 4 amino acids at the N terminus) is suggestive of specific function(s) in these cells (14). Recently, Keilhack et al. (15) have shown that SHP-1 is an important downstream regulator of ROS receptor signaling in epididymal epithelium (15). Furthermore, recent evidence indicates that SHP-1 associates with and dephosphorylates p120 catenin in EGF-stimulated A431 cells (16), suggesting a role for this PTP in the regulation of catenin function and cadherin-mediated epithelial cell-cell adhesion.

The intestinal epithelium remains a model of choice to study regulation of signal transduction pathways during cell proliferation and differentiation largely due to its constant differentiating system with a rapid and orderly turnover of cells (17). Cell differentiation begins with a sudden loss of proliferative ability, a process characterized by marked changes in cell ultrastructure and by the expression of several newly acquired end products, including the expression of the gut disaccharidase sucrase-isomaltase (17, 18). Committed intestinal epithelial cells withdraw from the cell cycle to differentiate during the G1 phase. Hence, molecules that stimulate or inhibit G1 phase progression are thereby likely candidates for controlling cell cycle and differentiation in developing tissue. The Wnt/beta -catenin signaling pathway plays a central role in the regulation of gastrointestinal proliferation, and mutations in this pathway have been detected in over 80% of colorectal cancers (19, 20). Indeed, beta -catenin plays an essential role in intestinal epithelial cells, not only as a cadherin-associated complex but also as a signaling molecule in the nucleus. Specifically, beta -catenin has been reported to transactivate gene expression through binding with its counterpart in intestinal cells, TCF-4 (21, 22). Downstream targets of the beta -catenin-TCF complex include cyclin D1, c-Myc, peroxisome proliferator-activated receptor delta , and E-cadherin (23-27). In order to mobilize into the nucleus and function as a signaling molecule, it is reasonable to assume that beta -catenin must first dissociate from the cadherin complexes and be stabilized in the cytoplasm. Increased cytoplasmic pools of beta -catenin can be prompted by several mechanisms such as down-regulation of E-cadherin (28), point mutation of beta -catenin itself (29), or by inhibition of the degradation pathway mediated by adenomatous polyposis coli, axin, glycogen synthase kinase-3beta , and beta -TrCP (30, 31).

In the present study, beta -catenin is revealed as one of the key binding partners for SHP-1 in human intestinal crypt cells and a substrate for SHP-1 in these cells. Furthermore, our results indicate that SHP-1 negatively regulates beta -catenin-dependent transcriptional activity resulting in the inhibition of cyclin D1 and c-myc gene expression, the activation of which represents one of the earliest cell cycle-regulated events occurring during the transition from G0/G1 to S phase.

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INTRODUCTION
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DISCUSSION
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Animals and Human Specimens-- Fifteen SHP-1 mutant mev/mev mice (C57aBL/6J-Hcph mev (homozygous males) and 12 control animals (C57BL/6J) were purchased from The Jackson Laboratory (Bar Harbor, ME). Mice were fed Purina chow ad libitum and kept in a controlled temperature and light cycle environment (20 °C; 12 h light, 12 h darkness). All studies were conducted in agreement with the principles and procedures outlined in the Canadian Guidelines for Care and Use of Experimental Animals. Tissues from human fetuses varying in age from 18 to 20 weeks of gestation (post-fertilization fetal ages were estimated according to Streeter (32)) were obtained from normal elective pregnancy terminations. No tissue was collected from cases associated with a known fetal abnormality or fetal death. Studies were approved by the Institutional Human Subject Review Board.

Indirect Immunofluorescence in Human Intestine-- Segments of human fetal small intestine were rinsed with 0.15 M NaCl, cut into small fragments, embedded in optimum cutting temperature compound, and quickly frozen in liquid nitrogen. Frozen sections 2-3 µm thick were spread on silane-coated glass slides and air-dried for 1 h at room temperature before storage at -80 °C (33-35). For indirect immunofluorescence, sections were fixed with 2% formaldehyde in PBS (pH 7.4; 45 min, 4 °C) prior to detection of 4,6-diamidino-2-phenylindole (Vector Laboratories, Burlingame, CA) and SHP-1 (a kind gift from Dr. A. Veillette, University of Montreal, Quebec, Canada) (1:250, 2 h, room temperature). Secondary antibodies consisted of goat anti-rabbit IgG-fluorescein isothiocyanate from Roche Diagnostics. Negative controls (no primary antibody) were included in all experiments.

Cell Culture-- Human intestinal epithelial cells (HIEC) were cultured as described previously (36) in Dulbecco's modified Eagle's medium (DMEM; Invitrogen) supplemented with 4 mM glutamine, 20 mM HEPES, 50 units/ml penicillin, 50 µg/ml streptomycin, 5 ng/ml recombinant human EGF (all obtained from Invitrogen), 0.2 IU/ml insulin (Connaught Novo Laboratories, Willowdale, Ontario, Canada), and 5% fetal bovine serum (FBS). The rat intestinal epithelial crypt cell line IEC-6 (37) and the Caco-2/15 cell line were obtained from A. Quaroni (Cornell University, Ithaca, NY). This clone of the parent Caco-2 cell line (HTB 37; American Type Culture Collection, Manassas, VA) has been extensively characterized elsewhere (34, 35, 38-39) and was originally selected as expressing the highest levels of sucrase-isomaltase among 16 clones obtained by random cloning. Both cell lines were cultured in DMEM containing 10% FBS, as described previously (34). Primary cultures of human differentiated enterocytes (PCDE) prepared from fetal small intestines ranging from 18 to 20 weeks of age were cultured as described above for HIEC. When tested after 5-7 days, these primary cultures remained well preserved, and both goblet and absorptive cells exhibited all the main characteristics of intact villus intestinal cells (40). Human embryonic kidney 293 cells (American Type Culture Collection) were cultured in DMEM containing 10% FBS.

Immunofluorescence Microscopy on Cultured Cells-- HIEC, Caco-2/15, and IEC-6 cells grown on sterile glass coverslips were washed twice with ice-cold PBS, fixed in methanol/acetone (30-70%) for 15 min at -20 °C, permeabilized with 0.1% Triton X-100 in PBS for 10 min, and blocked with PBS/bovine serum albumin 2% (20 min at room temperature). Cells were then immunostained for 1 h with primary antibody for the detection of SHP-1 or beta -catenin and 30 min with the secondary conjugated antibody. Negative controls (no primary antibody) were included in all experiments.

Expression of Proteins in Intestinal Tissue-- After mice were sacrificed by cervical dislocation, the jejunum and colon were rapidly removed and the mucosae scraped and homogenized (1 mg of tissue/50 µl) in Triton buffer (150 mM NaCl, 1 mM EDTA, 40 mM Tris-HCl, pH 7.6, 1% Triton X-100, 0.2 mM orthovanadate) supplemented with protease inhibitors (0.2 mM phenylmethylsulfonyl fluoride (PMSF), 20 µg/ml leupeptin, 2 µg/ml pepstatin, 20 µg/ml aprotinin). Homogenates were cleared by centrifugation (13,000 × g, 10 min). One-half of the homogenates was mixed in Laemmli's buffer, boiled for 5 min, and frozen until preparation of Western blots; the other half was stored at -80 °C for beta -catenin immunoprecipitation. Proteins were determined by the modified Lowry procedure described by Peterson (41).

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% bromphenol blue, 1 mM PMSF). Proteins (40 µg) from whole cell lysates were separated by SDS-PAGE in 7.5 or 10% gels and detected immunologically following electrotransfer onto nitrocellulose membranes (Amersham Biosciences). Protein and molecular weight markers (Bio-Rad) were localized with Ponceau Red. After blocking for 1 h at 25 °C in PBS, 0.05% Tween containing 5% powdered milk, membranes were first incubated for 2-4 h at 25 °C with antibodies against either SHP-1, beta -catenin, E-cadherin and TCF-4 (BD Biosciences), actin, cyclin D1, and antiphosphotyrosine (PY99, Santa Cruz Biotechnologies, Santa Cruz, CA), pRb (Pharmingen), c-Myc (Roche Diagnostics), or sucrase-isomaltase (HSI-14 from A. Quaroni, Cornell University) in blocking solution, followed by a second incubation with horseradish peroxidase-conjugated goat anti-mouse or anti-rabbit (1:1000) IgG (both from Sigma) in blocking solution for 1 h. The blots were visualized by the Amersham ECL system (Amersham Biosciences). Protein concentrations were measured using a modified Lowry procedure with bovine serum albumin as standard (41).

Immune Complex Phosphatase Assay-- Cells were lysed on ice for 10 min with 1 ml/dish of lysis buffer (150 mM NaCl, 1 mM EDTA, 40 mM Tris-HCl, pH 7.6, 1% Triton X-100) supplemented with protease inhibitors (0.1 mM PMSF, 10 µg/ml leupeptin, 1 µg/ml pepstatin, 10 µg/ml aprotinin). Lysates (800 µg) cleared by centrifugation (10,000 × g, 10 min) were incubated for 1 h at 4 °C with protein A-Sepharose (Amersham Biosciences) that had been initially preincubated for 2 h with anti-SHP-1. Immunocomplexes were washed four times with ice-cold lysis buffer and three times with ice-cold phosphatase buffer (25 mM HEPES, pH 7.2, 50 mM NaCl, 2.5 mM EDTA, 5 mM dithiothreitol) prior to initiating the phosphatase assay according to Upstate Biotechnologies, Inc. (Lake Placid, NY), and as described previously (42). Levels of immunoprecipitated SHP-1 were analyzed by Western blotting. The phosphatase reaction was initiated by incubating the immunocomplexes at 30 °C in the presence of 750 µM of the phosphopeptide RRLIEDAEpYAARG (where pY is phosphotyrosine). After 45 min, the reaction was stopped by addition of malachite green solution. Sample absorbance was measured at 655 nm after allowing for color development. The increase in phosphatase activity (absorbance) in PCDE was calculated relative to the level observed in HIEC, which was set at 1, whereas the increase in phosphatase activity (absorbance) in differentiating Caco-2/15 cells was calculated relative to the level measured in subconfluent Caco-2/15, which was also set at 1. Specific phosphatase activity was calculated relative to the amount of immunoprecipitated SHP-1. Band intensities in Western blots were quantified by laser densitometry using an Alpha Imager 1200 documentation and analysis system (Alpha Innotech, San Leandro, CA).

Coimmunoprecipitation Experiments-- Cells were washed twice with ice-cold PBS, lysed in chilled lysis buffer (150 mM NaCl, 1 mM EDTA, 40 mM Tris-HCl, pH 7.6, 1% Triton X-100, 0.1 mM PMSF, 10 µg/ml leupeptin, 1 µg/ml pepstatin, 10 µg/ml aprotinin, 0.1 mM orthovanadate, and 40 mM beta -glycerophosphate), and lysates cleared of cellular debris by centrifugation. Primary antibodies were added to 800 µg of each cell lysate and incubated for 2 h at 4 °C under agitation. Forty µg of protein A-Sepharose (Amersham Biosciences) was subsequently added for 1 h (4 °C under agitation). Immunocomplexes were harvested by centrifugation and washed four times with ice-cold lysis buffer. Proteins were solubilized in Laemmli's buffer and separated by SDS-PAGE.

Expression Vectors and Reporter Constructs-- Plasmid DHFR-luciferase, which contains a high affinity E2F-binding site in the dihydrofolate reductase promoter (43), was a kind gift of Dr. P. Farnham (University of Wisconsin). The dhfr gene, which is required for DNA synthesis, is transcribed at the G1/S transition. The cyclin D1 reporter construct, which contains the cyclin D1 gene promoter from nucleotides -944 to +139 cloned upstream of the luciferase gene of the pXP2 reporter construct, has been described previously (44) and was kindly provided by R. Muller (Institute of Molecular Biology and Tumor Research, Philipps-University Marburg, Germany). The c-fos-luciferase reporter vector was provided by Dr. C. Czernilofsky (Bender and Co., Vienna, Austria). The pRL-SV40 Renilla luciferase and the c-Myc-luciferase reporter vectors were from Promega (Nepean, Ontario, Canada). The T cell factor (TCF) reporter construct TOPFLASH was purchased from Upstate Biotechnology, Inc. (Lake Placid, NY). The constitutive active mutant of mouse c-Src p60Y529F was a kind gift from Dr. Josée Lavoie (Université Laval, Quebec, Canada). The full-length mouse SHP-1 cDNA (M. Thomas, Howard Hughes Medical Institute, St. Louis, MO) was subcloned into pcDNAneo vector and described previously (45). Mutation of the critical cysteine 453 of the catalytic site of the molecule for serine (SHP-1C453S) was performed by site-directed mutagenesis of double-stranded DNA according to the Clontech protocol.

Transient Transfections and Luciferase Assays-- Subconfluent HIEC were seeded in 12-well plates and cotransfected by lipofection (LipofectAMINE 2000, Invitrogen) with 0.2 µg of DHFR-luciferase, c-Myc-luciferase, cyclin D1-luciferase, c-Fos-luciferase or TOPFLASH reporters and 0.2 µg of the relevant expression vector (pcDNAneo) containing SHP-1 or SHP-1C453/S. The pRL-SV40 Renilla luciferase vector (Promega, Nepean, Ontario, Canada) was used as a control for transfection efficiency. Two days after transfection, luciferase activity was measured as described previously (34, 35), according to the Promega protocol.

Electrophoretic Mobility Shift Assays-- Nuclear extracts were prepared from HEK293 cells overexpressing SHP-1 or SHP-1C/S or Caco-2/15 cells at different times of confluency, according to Stein et al. (46). Electrophoretic mobility shift assays were performed as described previously (35, 47). Samples were electrophoresed in a 4% polyacrylamide gel containing 0.5% Tris borate buffer and 2% glycerol. The high affinity TCF/LEF-1 DNA-binding site (5'-GCACCCTTTGATCTTACC-3') previously employed in Caco-2 cells (48) was used for electrophoretic mobility shift assays.

Proliferation Assay in Mice-- Control and motheatenv mice were injected with 10 mM BrdUrd (1 ml/100 g body weight) 2 h prior to sacrifice. Segments of jejunum and colon were rinsed with 0.15 M NaCl, cut into small fragments, embedded in optimum cutting temperature compound, and quickly frozen in liquid nitrogen. Frozen sections 2-3 µm thick were spread on silane-coated glass slides and air-dried 1 h at room temperature before storage at -80 °C (33-35). For indirect immunofluorescence, sections were fixed according to the In Situ Cell Proliferation Kit from Roche Diagnostics.

Data Presentation and Statistical Analysis-- Luciferase assays were performed in either duplicate or triplicate, and results were analyzed by the Student's t test. Differences were considered significantly different at p < 0.05. Typical Western blots shown are representative of at least three independent experiments.

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Expression of SHP-1 in the Human Fetal Intestinal Epithelium, in the Caco-2/15 Intestinal Cell Line, and in Normal Human and Rat Intestinal Epithelial Crypt Cells-- Expression of SHP-1 was first investigated in intact fetal intestinal epithelium (20 weeks of gestation). The use of a specific antibody against SHP-1 revealed that this PTP was expressed in the nuclei of all jejunal and colonic epithelial cells (Fig. 1, A and B), with strongest nuclear staining observed in cells occupying the lower third of the villus in the jejunum (Fig. 1A, see arrows) and the mid-region of colonic crypts (Fig. 1B, see arrowheads). In the colonic cancer cell line Caco-2/15 (Fig. 1C) and in normal undifferentiated crypt-like HIEC (Fig. 1D) and IEC-6 (Fig. 1E) cells, indirect immunofluorescence revealed that SHP-1 was well expressed and primarily nuclear, while exhibiting very weak, albeit detectable, cytoplasmic staining.


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Fig. 1.   Expression of SHP-1 in the human fetal intestinal epithelium, in the Caco-2/15 intestinal cell line, and in normal human and rat intestinal epithelial crypt cells. Frozen sections of fetal jejunum (A) and colon (B) between 18 and 20 weeks of gestation were stained with antibodies to SHP-1, and nuclei were stained with 4,6-diamidino-2-phenylindole. A, the crypt-villus axis is oriented perpendicular to the figure with the crypt at the bottom. SHP-1 was mostly localized in the nuclei of villus cells (see arrows). Scale bar, 20 µm. B, in the colon, SHP-1 was mostly detected in the nuclei of cells in the mid-region of colonic crypts (see arrowheads). Dapi, 4,6-diamidino-2-phenylindole. Scale bar, 20 µm. Subconfluent Caco-2/15 (C), HIEC (D), and IEC-6 (E) cells were fixed with methanol/acetone and permeabilized with a solution of 0.1% Triton X-100 for immunofluorescence and staining for SHP-1 protein. In situ indirect immunofluorescence shown here is representative of three independent experiments. Scale bars, 20 µm.

Increased Expression and Activity of SHP-1 during Cell Cycle Arrest and Differentiation of Intestinal Epithelial Cells-- Caco-2/15 cells, which spontaneously differentiate into a small bowel phenotype after confluence (30, 34-37), were harvested at 70 (day -2) and 100% confluence (day 0) and at 6, 9, and 12 days post-confluence and analyzed by Western blotting to confirm timing of cell cycle arrest in G1 phase and induction of sucrase-isomaltase protein expression. Consistent with previous observations (34, 35, 49), decreased phosphorylation of p105Rb protein became apparent and significant at day 6 post-confluence concomitant with the onset and accumulation of sucrase-isomaltase expression (Fig. 2A). Expression and phosphatase activity levels of SHP-1 were also analyzed by Western blotting and immunoprecipitation, respectively. Although specific activity of SHP-1 reached a peak at 6 days post-confluence (Fig. 2C), SHP-1 expression was induced as soon as cells reached confluence (day 0) and progressively increased during Caco-2/15 differentiation (Fig. 2A).


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Fig. 2.   Increased expression and activity of SHP-1 during cell cycle arrest and differentiation of intestinal epithelial cells. A, Caco-2/15 cells were harvested at 70 (SC) and 100% confluence (day 0) and at 6, 9, and 12 days post-confluence. Cell extracts (40 µg) were separated by 10% SDS-PAGE and proteins analyzed by Western blotting for expression of SHP-1, pRb, and sucrase-isomaltase. B, subconfluent proliferating HIEC, primary cultures of human differentiated enterocytes (PCDE), and intestinal mucosae were lysed and proteins separated by SDS-PAGE. SHP-1 and pRb protein expression were analyzed by Western blotting. Results are representative of three independent experiments. C, subconfluent growing HIEC and PCDE as well as Caco-2/15 cells at 70 (-2) and 100% confluence (day 0) and at 3, 6, and 9 days post-confluence were harvested. Cell extracts (800 µg) were immunoprecipitated with a specific antibody to SHP-1. Levels of immunoprecipitated SHP-1 were analyzed by Western blotting. Phosphatase activity of SHP-1 was assayed by using the phosphopeptide RRLIEDAEpYAARG as substrate, according to the Upstate Biotechnologies Inc. protocol. The increase in phosphatase activity (absorbance) in PCDE was calculated relative to the level observed in HIEC that was set at 1, whereas the increase in phosphatase activity (absorbance) in differentiating Caco-2/15 cells was calculated relative to the level measured in subconfluent Caco-2/15 that was also set at 1. Specific phosphatase activity was calculated relative to the amount of immunoprecipitated SHP-1. Data shown are representative of that obtained in three independent experiments.

Because Caco-2/15 cells are derived from a human colonic adenocarcinoma (50), it was deemed important to validate the above results in normal human intestinally derived cells. Expression and activity of SHP-1 were therefore analyzed in lysates of the following cell models: crypt-like HIEC cells which are proliferative and undifferentiated (36), and PCDE which are primary cultures of differentiated, nonproliferative villus enterocytes (40). As shown in Fig. 2B, Rb protein was exclusively detected in its hyperphosphorylated state in HIEC compared with that found in both PCDE and intestinal mucosae extract, indicating that these latter cells were arrested in G0/G1 phase. Moreover, expression (Fig. 2B) and specific phosphatase activity (Fig. 2C) of SHP-1 were significantly lower in proliferative HIEC cells compared with that found in PCDE cells and intestinal mucosae extract. Overall these data indicate that increased SHP-1 expression and phosphatase activity do correlate with cell cycle arrest and induction of differentiation in intestinal epithelial cells.

SHP-1 Negatively Controls Cell Cycle Progression of Intestinal Epithelial Cells-- An important early event in terminal differentiation of cells, especially in tissues exhibiting a rapid turnover such as the intestinal epithelium, is their withdrawal from the cell cycle (2, 49). To evaluate the role of SHP-1 in intestinal cell cycle progression, we generated a catalytically inactive SHP-1 by mutating the catalytic cysteine 453 to serine (SHP-1/C453S). This mutation completely abolished phosphatase activity of SHP-1 (45, 51). This construct was transiently transfected in HEK293 cells, and its expression was verified by Western blotting (data not shown). To evaluate the role of SHP-1 in intestinal cell cycle progression, SHP-1 wild-type and SHP-1/C453S constructs were tested on dihydrofolate reductase (dhfr) expression in subconfluent HIEC cells. The dhfr gene, which is required for DNA synthesis and is transcribed at the G1/S transition, contains E2F-dependent binding sites in its promoter. In addition, microinjection of E2F into quiescent fibroblasts provokes S phase re-entry, underscoring the importance of E2F in cell growth control (52). Therefore, the plasmid construction containing the E2F-responsive dhfr promoter linked to a luciferase reporter gene represents a sensitive reporter of cell cycle progression and S phase entry (43, 53). The results shown in Fig. 3A demonstrate that ectopic expression of wild-type SHP-1, but not SHP-1/C453S mutant, significantly inhibited dhfr gene expression by roughly 60%, suggesting that SHP-1 negatively regulates intestinal epithelial cell cycle progression. To clarify further the role of SHP-1 in cell proliferation, the effects of either enforced SHP-1 or SHP-1/C453S expression were compared on the transcriptional activity of a range of promoters such as the cyclin D1 and c-myc gene promoters. The activation of these promoters represents one of the earliest cell cycle-regulated events occurring during the G0/G1 to S phase transition (53, 54). As shown in Fig. 3, B and C, transcriptional activities of c-myc and cyclin D1 promoters were both significantly attenuated (49 and 55%, respectively) by ectopic expression of the wild-type SHP-1 mutant but not by the SHP-1/C453S mutant.


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Fig. 3.   SHP-1 inhibits dhfr, c-myc, cyclin D1 gene expression and negatively controls beta -catenin/TCF-dependent activity (TOPFLASH) in intestinal epithelial cells. Subconfluent HIEC cells were cotransfected with 0.2 µg of pcDNAneo I containing or lacking the wild-type SHP-1 or the dominant negative mutant C453S of SHP-1 (SHP-1/CS) with 0.2 µg of DHFR- (A), c-Myc- (B), cyclin D1- (C), TOPFLASH- (D), or c-Fos-luciferase (E) reporters. Two days after transfection, cells were lysed, and luciferase activity was measured. The increase in luciferase activity was calculated relative to the pcDNAneo I level that was set at 1. Results are the mean ± S.E. of at least three separate experiments. *, significantly different from control at p < 0.05 (Student's t test).

Because overexpression of the wild-type SHP-1 inhibited the expression of the two key cell cycle regulatory genes c-myc and cyclin D1, possible SHP-1 control of beta -catenin/TCF transcriptional activity was further investigated. Indeed, functional beta -catenin/TCF-binding sites have been identified in the promoters of c-myc (24) and cyclin D1 (25, 26) genes. HIEC cells were thereby transfected with TOPFLASH reporter, which directly assays beta -catenin/TCF activity (55). As shown in Fig. 3D, beta -catenin/TCF transcriptional activity was significantly inhibited (58%) by the expression of wild-type SHP-1 but not SHP-1/C453S mutant. The effect of wild-type SHP-1 was specific because overexpression of wild-type SHP-1 or SHP-1/C453S mutant did not influence c-fos gene expression, which represents a sensitive reporter of growth factor-induced transcriptional activity (56). This early gene promoter contains the well characterized serum-responsive element, whose activity is induced upon serum activation of the serum-responsive factor (57).

SHP-1 Overexpression Does Not Interfere with the DNA Binding Capacity of beta -Catenin-TCF Complex-- Consistent with a previous study (48), gel shift analysis in Caco-2/15 cells demonstrated a decrease in binding of beta -catenin-TCF complex to TCF/LEF-1-binding site with time of confluency and differentiation, correlating with the increased expression of SHP-1 (Fig. 4A). Thus, increased expression of SHP-1 could inhibit beta -catenin/TCF transcriptional activity by interfering with its DNA binding capacity. Electrophoretic mobility shift experiments were therefore performed in HEK293 cells to determine whether the DNA binding capacity of beta -catenin/TCF was affected by SHP-1 and/or SHP-1/C453S overexpression. As shown in Fig. 4B, binding of nuclear proteins to the TCF/LEF-1 DNA-binding site was not affected by using extracts prepared from either SHP-1 or SHP-1/C453S-overexpressing HEK293 cells.


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Fig. 4.   SHP-1 overexpression does not interfere with the DNA binding capacity of beta -catenin-TCF complex. Binding of nuclear proteins to a high affinity TCF/LEF-1 DNA-binding site was assessed in Caco-2/15 cells and HEK293 overexpressing SHP-1 and SHP-1/C453S. Caco-2/15 cells were harvested at confluence (day 0) and at 3, 6, and 9 days post-confluence. HEK293 cells were harvested 2 days after transfection. Total cell extracts were separated by 10% SDS-PAGE and proteins analyzed by Western blotting for expression of SHP-1. Nuclear extracts were prepared and mixed with 32P-labeled double-stranded oligonucleotides. DNA-protein complexes were separated from the free probe on a native polyacrylamide gel. Results are representative of three independent experiments.

SHP-1 Associates with beta -Catenin in Intestinal Epithelial Cells-- To investigate further the regulation of beta -catenin function by SHP-1, interaction of SHP-1 with beta -catenin was tested in coimmunoprecipitation assays. Immunoprecipitations demonstrated the formation of a complex between SHP-1 and beta -catenin in intestinal crypt cells. Indeed, Fig. 5A shows that SHP-1-beta -catenin association in IEC-6 and HIEC cells was significantly diminished once cells reached confluence (day 0) and decreased even further at day 5 of post-confluence. Localization of SHP-1 and beta -catenin proteins was further analyzed in asynchronously growing subconfluent IEC-6 cells and in quiescent confluent IEC-6 cells. As illustrated in Fig. 5B, bright nuclear SHP-1 staining was evident in proliferative subconfluent (panel 1) and in quiescent confluent (panel 2) IEC-6 cells, although some SHP-1 staining was also observed in the cytoplasm of 5-day post-confluent cells (panel 3). In subconfluent growing IEC-6 cells, beta -catenin protein staining was partially localized in the nucleus but was also clearly visible at the sites of cell-cell contacts (Fig. 5B, panel 4). Nuclear localization of beta -catenin was completely altered upon confluency, as shown by the accumulation of beta -catenin staining at sites of cell-cell contact in confluent (Fig. 5B, panel 5) and post-confluent (Fig. 5B, panel 6) IEC-6 cells. These results suggest that SHP-1 binds beta -catenin in intestinal epithelial cells and that this association decreases in confluent cells possibly because of recruitment of beta -catenin to cell junctions.


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Fig. 5.   SHP-1 associates with beta -catenin in intestinal epithelial cells. A, SHP-1 and beta -catenin were immunoprecipitated from 800 µg of lysates of pre-confluent, confluent (day 0), and 5-day post-confluent IEC-6 and HIEC cells. Proteins from immunoprecipitates were solubilized in Laemmli buffer and separated by SDS-PAGE. Proteins were analyzed by Western blotting to determine the amount of SHP-1 and beta -catenin in immunoprecipitates. Blots shown are representative of three independent experiments. B, subconfluent IEC-6 cells were fixed with methanol/acetone and permeabilized with a solution of 0.1% Triton X-100 for immunofluorescence and staining for SHP-1 or beta -catenin proteins. In situ indirect immunofluorescence shown is representative of three independent experiments. Scale bar, 20 µm.

Association of SHP-1 and Dephosphorylation of beta -Catenin by SHP-1 in HEK293 Cells-- To assess whether SHP-1/beta -catenin interaction results in SHP-1-mediated beta -catenin dephosphorylation, a constitutively active mutant of c-Src (p60Y529F) was coexpressed with SHP-1 or with the catalytically inactive SHP-1/C453S mutant in HEK293. beta -Catenin is heavily tyrosine-phosphorylated in Src-transformed cells and is considered as a good substrate for c-Src (58). Overexpression of p60Y529F in HEK293 cells led to massive tyrosine phosphorylation of endogenous beta -catenin (Fig. 6, top panel, lane 2). The C453S mutant of SHP-1 had virtually no effect on p60Y529F-induced tyrosine phosphorylation of beta -catenin (Fig. 6, top panel, lane 4), whereas SHP-1 expression resulted in a strong decrease in tyrosine-phosphorylated beta -catenin (Fig. 6, top panel, lane 3). This suggests that beta -catenin may indeed be a specific target for SHP-1. In addition, the catalytically inactive mutant of SHP-1 (C453S), which has been described as a substrate-trapping mutant (4), exhibited strongly elevated binding (Fig. 6, bottom panel, lanes 3 and 4), suggesting that beta -catenin is a very efficient SHP-1 substrate.


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Fig. 6.   Association of SHP-1 and dephosphorylation of beta -catenin by SHP-1 in HEK293 cells. HEK293 cells were transfected with the empty vectors pCMV and pCDNAneo I (EV), activated c-Src alone (c-src/CA + pCDNAneo), or together with SHP-1 (c-src/CA + SHP-1) or SHP-1/C453S (c-src/CA + SHP-1/CS). Twenty four hours after transfection, beta -catenin was immunoprecipitated from 800 µg of lysates. Proteins from immunoprecipitates were solubilized in Laemmli buffer and separated by SDS-PAGE. Tyrosine phosphorylation of immunoprecipitated proteins was analyzed by immunoblotting using PY-99 antiphosphotyrosine. Proteins were analyzed by Western blotting (WB) to determine the amount of SHP-1 and beta -catenin in immunoprecipitates. Blots shown are representative of three independent experiments.

A Subset of Motheatenv Mice Exhibits a Moderate Increase in Tyrosine Phosphorylation and Expression Levels of beta -Catenin in Intestinal Epithelium-- We next tested whether impairment of SHP-1 activity in mev/mev mice could affect beta -catenin signaling by analyzing tyrosine phosphorylation as well as levels of immunoprecipitated beta -catenin in mev/mev intestinal epithelium. As shown in Fig. 7A, jejunal samples from control and mev/mev mice revealed three bands, whereas colon samples maintained the presence of only one major 94-kDa band in both wild-type and mev/mev mice. Moreover, expression levels of beta -catenin were much more elevated in the colon than in the jejunum of all mice (Fig. 7A). Of note, the colon of a substantial subset of mev/mev mice (about one-third) exhibited increased beta -catenin tyrosine phosphorylation compared with that in control (wild-type) animals, although colonic mucosae did generate a greater quantity of immunoprecipitated beta -catenin protein. In this regard, in this subset of mev/mev mice, the jejunum and colon exhibited a moderate, but consistent, increase in beta -catenin protein levels compared with that in control (wild-type) animals (Fig. 7A). There was no difference, however, in E-cadherin and TCF-4 expression levels as well as in tyrosine phosphorylation of TCF-4 between these mev/mev and control mice (data not shown). Expression levels of cyclin D1 and c-Myc proteins, two targets of beta -catenin-TCF complex, were next analyzed in this subset of mev/mev mice. As shown in Fig. 7B, there was a dramatic increase in expression of c-Myc in the jejunal and colonic epithelium of these mev/mev mice. Cyclin D1 expression was also increased in these mev/mev mice but at a much lesser scale than c-Myc. Equal protein loading of each lane was confirmed by anti-actin antibody labeling.


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Fig. 7.   A subset of motheatenv mice exhibits enhanced tyrosine phosphorylation and beta -catenin expression levels in intestinal epithelium. Wild-type and mev/mev mice were sacrificed, the jejunum and colon rapidly removed, and their respective mucosae scraped and homogenized as described under "Experimental Procedures." A, beta -catenin was immunoprecipitated from 800 µg of cleared mucosal lysates from jejunum and colon. Proteins from immunoprecipitates were solubilized in Laemmli buffer and separated by SDS-PAGE. Proteins were analyzed by Western blotting to determine the amount of beta -catenin in immunoprecipitates. Tyrosine phosphorylation of immunoprecipitated proteins was also analyzed by immunoblotting using PY-99 antiphosphotyrosine. B, cleared extracts were separated by 10% SDS-PAGE and proteins analyzed by Western blotting for expression of c-Myc, cyclin D1, and actin. Blots shown are representative of three independent experiments. C, wild-type and mev/mev mice were injected with BrdUrd 2 h prior to sacrifice in order to label only cells in S phase. Immunofluorescence was performed on sections of jejunum and colon (not shown) of both wild-type (left panel) and mev/mev mice (right panel). Note the similarity in the number of labeled nuclei between wild-type and mev/mev mice. Scale bar, 20 µm.

In vivo pulse labeling of cells in S phase with BrdUrd was performed to verify whether the rate of intestinal epithelial cell proliferation is perturbed in the subset of mev/mev mice in which the expression levels of cyclin D1 and c-Myc were increased. Mice were sacrificed shortly (2 h) after post-injection at a time when only actively cycling cells (S phase) were labeled. As shown in Fig. 7C, no significant difference in the rate of epithelial cell proliferation in either jejunum or colon (latter not shown) was found between mev/mev mice and their controls.

    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Whereas multiple targets for SHP-1 have been identified in hematopoietic cells (3, 7, 8), very little is known about the function of SHP-1 in epithelial cells. In this report, we suggest for the first time the involvement of SHP-1 in the negative control of intestinal epithelial cell proliferation. Indeed, increased SHP-1 expression and phosphatase activity coincide with cell cycle arrest and induction of differentiation in intestinal epithelial cells. Results show that the expression and phosphatase activity of SHP-1 are significantly increased in differentiated intestinal epithelial cells in comparison to undifferentiated cells. The nuclear localizations of SHP-1 in the lower third of the small intestinal villi and in the mid-region of colonic crypts are in essence a reflection of the distribution of cells that have ceased proliferation. Ectopic expression of SHP-1 in human intestinal crypt cells inhibits E2F-dependent transcriptional activity and decreases the expression of c-myc and cyclin D1 genes, the activation of which represents one of the earliest cell cycle-regulated events occurring during the transition from G0/G1 to S phase. In addition, we propose that SHP-1 may regulate the nuclear transcriptional function of beta -catenin by its dephosphorylation. Finally, in agreement with a functional interaction of beta -catenin and SHP-1, there is increased expression of cyclin D1 and c-Myc proteins in the jejunum and colon of mev/mev mice, whose SHP-1 activity is strongly compromised.

Generally, SHP-1 has been shown to act as a negative regulator of signal transduction in hematopoietic cells, terminating signals from a diverse range of signaling molecules including the EGF receptor, interleukin 3 receptor, c-Kit, colony stimulating factor-1 receptor, B and T cell antigen receptors, and receptor-associated JAK kinases (for review see Ref. 59). However, it appears that the role of SHP-1 is dependent on cell type (60, 61). Indeed, overexpression of the catalytically inactive mutant of SHP-1 in HEK293 cells strongly suppresses mitogen-activated pathways and results in decreased cell growth, DNA synthesis, and the transcription of early response genes (60). Furthermore, transfection of HeLa cells with inactive SHP-1 reduces STAT DNA binding induced by interferon gamma  and EGF (61). In the present study, we propose a negative role of SHP-1 in the control of the intestinal epithelial cell cycle. As yet, the molecular basis for the apparent opposite effects of SHP-1 in different cell systems has to be defined, although the phosphatase domain appears to be critical.

It is widely assumed that SHP-1 is a cytoplasmic protein (2-4, 59). However, a novel nuclear localization for SHP-1 has been demonstrated in nonhematopoietic cells (62). Therefore, SHP-1 localization differs between nonhematopoietic and hematopoietic cells, with SHP-1 protein being virtually exclusively cytoplasmic in hematopoietic cell lines. These results have implications regarding the nuclear function of SHP-1 in nonhematopoietic cells. Few nuclear tyrosine-phosphorylated proteins have been identified. One potential target is the STAT proteins, which are activated by tyrosine phosphorylation and translocate to the nucleus (61, 63). In the present study, we were able to identify beta -catenin as a binding partner and substrate for SHP-1 in epithelial cells.

Tyrosine phosphorylation of beta -catenin has been shown to correlate with tumorigenesis, cell migration, and developmental processes (64). Furthermore, various growth factors (e.g. EGF, hepatocyte growth factor, and vascular epidermal growth factor) and cytoplasmic Src kinases are known to phosphorylate the tyrosine residue(s) on beta -catenin (58, 65, 66). Experimental data suggest that tyrosine phosphorylation of beta -catenin is implicated as a means for release from the E-cadherin complex (58, 67), although there is some controversy over its direct effect on cell adhesiveness. A recent study (68) reported that treatment of NIH 3T3 fibroblasts or HCT116 cells with the tyrosine phosphatase inhibitor pervanadate increased tyrosine phosphorylation of beta -catenin and led to relocation from cell-cell interfaces to the cytoplasm but did not change its binding activity to LEF-1, nor did it enhance cyclin D1 transactivation. This is, however, at odds with another recent study that clearly points out that phosphorylation of Tyr654, a residue placed in the last armadillo repeat of beta -catenin, decreases its binding to E-cadherin but stimulates the association of beta -catenin to the basal transcription factor TATA-binding protein (TBP) (69). Interestingly, this greater association between TBP and beta -catenin correlates with a higher stimulation of beta -catenin/TCF transcriptional activity. In addition, other nuclear factors have been shown to interact with the N- and C-terminal transactivation domains of beta -catenin, including Pontin (70), Teashirt (71), Sox 17 and 13 (72), histone deacetylase (73), Brg-1 (74), and SMAD4 (75). Hence, we propose a hypothetical model of SHP-1-induced inhibition of beta -catenin/TCF transcriptional activity in intestinal epithelial cells whereby SHP-1 dephosphorylates beta -catenin leading to a decreased interaction of the beta -catenin-TCF-4 complex with TBP or other nuclear coactivators or to an increased interaction with corepressors.

SHP-1 could be targeted to nuclear beta -catenin by recognizing the consensus sequence for ligands of the N-terminal SH2 domain of SHP-1, i.e. hXY(P)XXh (where h = hydrophobic and X = any amino acid). Indeed, three tyrosine residues in the sequence of human beta -catenin partially resemble the consensus sequence (16). Tyrosine 333 (Tyr-Thr-Tyr333-Glu-Lys-Leu), 604 (Leu-Leu-Tyr604-Ser-Pro-Ile), and 654 (Ala-Thr-Tyr654-Ala-Ala-Ala) in their phosphorylated forms are therefore candidates for SHP-1 interaction sites in beta -catenin. Interestingly, one of the phosphorylated beta -catenin tyrosine residues previously mapped was Tyr654 (58). One could therefore speculate that binding of SHP-1 to Tyr654 of beta -catenin results in the dephosphorylation of beta -catenin on Tyr654 (or other phosphotyrosines) eventually leading to dissociation of TBP or other coactivators from the beta -catenin-TCF complex. However, further studies are required at this point to identify the phosphorylated tyrosine on beta -catenin and its roles in beta -catenin function in intestinal epithelial cells and to clarify the mechanism by which SHP-1 specifically inhibits beta -catenin/TCF transcriptional activity.

The observation that most of the mev/mev mice did not exhibit any alterations in intestinal epithelium might suggest that the residual activity of SHP-1 in these mev/mev mice (5, 6) is probably sufficient to prevent intestinal disorders. However, it is of note that a moderate increased expression levels of beta -catenin in the jejunal and colonic mucosa of a subset of mev/mev mice was reproducibly observed in comparison to that observed in control mice. In contrast to the relatively well known degrading effects of serine phosphorylation on beta -catenin, the effects of its tyrosine phosphorylation are not well characterized beyond an effect on cell adhesiveness. An attractive possibility is that tyrosine phosphorylation may increase the entry of beta -catenin into the nucleus by increasing its stabilized pools in the cytoplasm. However, preliminary experiments demonstrate that SHP-1 overexpression in HEK293 cells did not result in dysregulation of beta -catenin stability.2 Although we did not observe an altered localization of beta -catenin in mev/mev mice (data not shown), one cannot rule out the possibility that an increased tyrosine phosphorylation may induce subtle changes in stability and subcellular localization, which are sufficient to affect its transactivating function. In fact, our data did demonstrate an increased expression of both cyclin D1 protein and even more predominantly of c-Myc in a subset of mev/mev mice. However, any significant increase in proliferation of gut epithelial cells was observed in mev/mev mice suggesting that the increased expression of cyclin D1 and c-Myc was not sufficient for S phase entry in these cells.

In conclusion, our data indicate that increased SHP-1 expression and activity coincide with cell cycle arrest and induction of differentiation of intestinal epithelial cells. The observed association of SHP-1 with beta -catenin in intestinal epithelial cells and its dephosphorylation by SHP-1 provide evidence that SHP-1 may be involved in the regulation of the beta -catenin/TCF signaling pathway and therefore modulate proliferation of these cells. Interestingly, down-regulation of the beta -catenin/TCF pathway is associated with the promotion of a more differentiated phenotype in colonic epithelial cells (48). Although further studies are needed to pinpoint the role of tyrosine phosphorylation in beta -catenin nuclear function and signaling, our study provides novel fundamental insights into the function of SHP-1 in the control of intestinal epithelial cell proliferation and in the early events of intestinal epithelial cell differentiation.

    ACKNOWLEDGEMENTS

We thank A. Vézina for technical assistance and P. Pothier for the critical reading of the manuscript. We also thank Drs. C. Poulin and F. Jacot, obstetricians from the Département de la Santé Communautaire du Centre Universitaire de Santé de l'Estrie, for invaluable collaboration in providing the tissue specimens used in this study.

    FOOTNOTES

* This work was supported in part by a grant from the Natural Sciences and Engineering Research Council of Canada.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.

§ Both authors contributed equally to this work.

Student scholar from the Fonds pour la Recherche en Santé du Québec.

** Recipient of a Canadian Research Chair in Signaling and Digestive Physiopathology. To whom correspondence should be addressed: Dépt. of d'Anatomie et de Biologie Cellulaire, Faculté de Médecine, Université de Sherbrooke, Sherbrooke, Quebec J1H 5N4, Canada. Tel.: 819-564-5271; Fax: 819-564-5320; E-mail: Nathalie.Rivard@USherbrooke.ca.

Published, JBC Papers in Press, February 5, 2003, DOI 10.1074/jbc.M300425200

2 C. Duchesne and N. Rivard, unpublished results.

    ABBREVIATIONS

The abbreviations used are: PTP, protein-tyrosine phosphatase; APC, adenomatous polyposis coli; BrdUrd, bromodeoxyuridine; dhfr, dihydrofolate reductase; EGF, epidermal growth factor; HEK, human embryonic kidney; HIEC, human intestinal epithelial cells; IEC, intestinal epithelial cells; PCDE, primary cultures of differentiated enterocytes; pRb, retinoblastoma protein; TBP, TATA-binding protein; TCF, T cell factor; DMEM, Dulbecco's modified Eagle's medium; FBS, fetal bovine serum; PBS, phosphate-buffered saline; PMSF, phenylmethylsulfonyl fluoride.

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