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Mol Biol Cell. 2007 September; 18(9): 3533–3544. doi: 10.1091/mbc.E07-03-0249. | PMCID: PMC1951739 |
Copyright © 2007 by The American Society for Cell Biology Differential Regulation of Epithelial and Mesenchymal Markers by δEF1 Proteins in Epithelial–Mesenchymal Transition Induced by TGF-β  Takuya Shirakihara, Masao Saitoh, and Kohei Miyazono  Department of Molecular Pathology, Graduate School of Medicine, University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo 113-0033, Japan Carl-Henrik Heldin, Monitoring Editor Received March 16, 2007; Revised June 18, 2007; Accepted June 25, 2007. Epithelial–mesenchymal transition (EMT), a crucial event in cancer progression and embryonic development, is induced by transforming growth factor (TGF)-β in mouse mammary NMuMG epithelial cells. Id proteins have previously been reported to inhibit major features of TGF-β–induced EMT. In this study, we show that expression of the δEF1 family proteins, δEF1 (ZEB1) and SIP1, is gradually increased by TGF-β with expression profiles reciprocal to that of E-cadherin. SIP1 and δEF1 each dramatically down-regulated the transcription of E-cadherin in NMuMG cells through direct binding to the E-cadherin promoter. Silencing of the expression of both SIP1 and δEF1, but not either alone, completely abolished TGF-β–induced E-cadherin repression. However, expression of mesenchymal markers, including fibronectin, N-cadherin, and vimentin, was not affected by knockdown of SIP1 and δEF1. TGF-β–induced the expression of Ets1, which in turn activated δEF1 promoter activity. Moreover, up-regulation of SIP1 and δEF1 expression by TGF-β was suppressed by knockdown of Ets1 expression. In addition, Id2 suppressed the TGF-β– and Ets1-induced up-regulation of δEF1. Taken together, these findings suggest that the δEF1 family proteins, SIP1 and δEF1, are necessary, but not sufficient, for TGF-β–induced EMT and that Ets1 induced by TGF-β may function as an upstream transcriptional regulator of SIP1 and δEF1. Transforming growth factor (TGF)-β, a prototypical member of the TGF-β family, regulates a broad range of cellular responses, including cell proliferation, differentiation, adhesion, migration, and apoptosis ( Bierie and Moses, 2006  ). TGF-β and related factors exhibit their pleiotropic effects through binding to transmembrane serine-threonine kinase receptors type I (TβR-I) and type II (TβR-II). On ligand-induced heteromeric complex formation between TβR-I and TβR-II, TβR-I is phosphorylated and activated by TβR-II kinase and mediates specific intracellular signaling through phosphorylation of receptor-regulated Smads (R-Smads). Phosphorylated R-Smads interact with co-Smad (Smad4) and translocate into the nucleus, where they regulate transcription of target genes in cooperation with various transcription factors and transcriptional coactivators or corepressors ( Miyazawa et al., 2002 ; Miyazono et al., 2003 ; Shi and Massague, 2003 ). TGF-β has potent antiproliferative effects on a wide variety of cells, including epithelial cells, endothelial cells, and hematopoietic cells, although under certain conditions it promotes the proliferation of mesenchymal cells, including fibroblasts, chondrocytes, and osteoblasts. TGF-β also induces the deposition of extracellular matrix proteins. In early stages of tumorigenesis, TGF-β inhibits the growth of epithelial cells, and insensitivity to this growth-inhibitory effect is associated with progression of tumors ( Akhurst and Derynck, 2001 ; Derynck et al., 2001 ). Transgenic mice expressing a dominant-negative TβR-II in epidermis exhibit malignant conversion of epithelial cells and promotion of tumor formation ( Gorska et al., 2003 ). Resistance to the antiproliferative effects of TGF-β is observed in numerous types of cancer ( Park et al., 1994 ; Heldin et al., 1997 ; Lu et al., 2006 ). The refractoriness of many carcinomas to effects of TGF-β is due to mutations in or loss of expression of receptors for it and to mutations in R-Smads and Smad4. Increase in expression of some negative regulators of TGF-β signaling, e.g., Smad7, has also been reported in certain cancers ( Kleeff et al., 1999 ; Kim et al., 2004 ). In contrast to its tumor-suppressive effects in the early stages of carcinogenesis, TGF-β also acts as a promoter of tumor cell invasion and metastasis in advanced stages of tumorigenesis ( Bierie and Moses, 2006 ). TGF-β is often overexpressed in various tumor tissues, induces migration and invasion of cancer cells and facilitates immunosuppression, angiogenesis, and deposition of extracellular matrix proteins. Blockade of TGF-β signaling thus leads to suppression of tumor cell motility, intravasation, and metastasis ( Muraoka et al., 2002 ; Azuma et al., 2005 ). Chronic exposure to TGF-β results in loss of TGF-β–mediated growth inhibition and marked changes in cell morphology ( Caulin et al., 1995 ; Portella et al., 1998 ). One mechanism by which TGF-β induces formation of spindle cell carcinomas and promotes tumor cell motility and invasion involves the epithelial-mesenchymal transition (EMT; Zavadil and Bottinger, 2005 ). During the process of embryonic development and that of wound healing and reorganization in adult tissues, epithelial cells may lose their epithelial polarity and acquire mesenchymal phenotypes ( Lee et al., 2006 ). The process of invasion of tumor cells involves the loss of cell–cell interaction together with acquisition of migratory properties and is often associated with EMT of cells. Formation of tight cell–cell adhesions is mainly dependent on the E-cadherin system in both embryonic and adult epithelial cells. Loss of E-cadherin–mediated cell–cell interaction is thus essential for the EMT that occurs during normal embryonic development as well as during invasion of tumor cells into adjacent connective tissues ( Peinado et al., 2004 ). Besides the loss of E-cadherin, EMT is characterized by the down-regulation of cytokeratins, up-regulation of mesenchymal markers including fibronectin, N-cadherin, and vimentin and acquisition of a fibroblast-like motile and invasive phenotype ( Lee et al., 2006 ; Thiery and Sleeman, 2006 ). Recent studies on the molecular mechanisms by which expression of E-cadherin is repressed in epithelial cells have revealed that several transcription factors, including the zinc-finger factors Snail and Slug, the two-handed zinc-finger factors of δEF1 family proteins (δEF1/ZEB1 and SIP1), and the basic helix-loop-helix (bHLH) factors Twist and E12/E47, are involved in this process ( Comijn et al., 2001 ; Yang et al., 2004 ; Moreno-Bueno et al., 2006 ). These transcription factors repress expression of E-cadherin and elicit EMT when overexpressed in normal epithelial Madin-Darby canine kidney (MDCK) and Eph4 cells. In addition, when overexpressed in cancer cells, these factors induce EMT with the development of metastatic properties such as migration and invasion in vitro and in vivo ( Barrallo-Gimeno and Nieto, 2005 ; Thiery and Sleeman, 2006 ). However, it is still uncertain how the expression of these transcriptional factors is regulated at the onset of EMT in cancer cells. TGF-β was first described as an inducer of EMT during development ( Miettinen et al., 1994 ) and is now thought to promote metastasis through induction of EMT on the front-edge cells of invasive cancer. It has been reported that TGF-β induces the expression of Snail and SIP1 mRNAs in some epithelial cells ( Comijn et al., 2001 ; Nagata et al., 2006 ). Snail is rapidly and transiently up-regulated through the TGF-β-Smad signaling pathway in mouse mammary epithelial NMuMG cells ( Nagata et al., 2006 ), whereas SIP1 is up-regulated at a later phase through unknown mechanisms. In addition to these transcription factors, a microarray screen of epithelial cells identified Id genes (inhibitors of differentiation or inhibitors of DNA binding) as early targets of TGF-β. Id proteins have an HLH domain but lack the basic DNA-binding domain. Through constitutive association with bHLH E12/E47 proteins, Id proteins maintain epithelial phenotypes by repressing the function of E12/E47, which acts as a repressor of E-cadherin. Down-regulation of Id2 by TGF-β thus relieves this inhibition, permitting conversion of epithelial cells to cells with mesenchymal phenotypes ( Kondo et al., 2004 ; Kowanetz et al., 2004 ). In the present study, we investigated the regulatory mechanisms by which TGF-β elicits EMT in NMuMG cells. We show that the δEF1 family proteins, SIP1 and δEF1, are required for TGF-β–induced E-cadherin repression, but not for the expression of mesenchymal markers. Ets1 functions as an upstream component of SIP1 and δEF1 in TGF-β–mediated EMT; its expression is up-regulated by TGF-β, and it acts in cooperative manner with E47 to induce expression of SIP1 and δEF1 genes, leading to EMT. SIP1 and δEF1 are thus critical components of TGF-β–induced EMT in NMuMG cells, although they are not sufficient to induce mesenchymal phenotype in these cells. Cell Culture and Reagents Mouse mammary epithelial NMuMG cells (American Type Culture Collection, Manassas, VA) were cultured in DMEM supplemented with 4.5 g/l glucose, 10% fetal bovine serum (FBS), 10 μg/ml insulin, 100 U/ml penicillin, and 100 μg/ml streptomycin. MDCK epithelial cells were grown in DMEM in the presence of 10% FBS and antibiotics. pBabe- or pBabe-Id2–infected NMuMG cells were generated as described previously ( Kondo et al., 2004 ). All cells were grown in a 5% CO 2 atmosphere at 37°C. Recombinant human TGF-β3 was purchased from R&D Systems (Minneapolis, MN). DNA Construction Mouse E-cadherin promoters with point mutations in E-boxes (termed E1, E2, E3, E21, E13, E23, and E213) were constructed using PCR-based mutagenesis on the mouse E-cadherin promoter (−178 to + 92 base pairs) in pGL2 ( Kondo et al., 2004 ). Human δEF1 promoter (−1107 to + 55 base pairs from the translation start site) was cloned using PCR from genomic DNA of human keratinocyte HaCaT cells isolated by DNeasy (QIAGEN, Chatsworth, CA). Amplification was performed by high-fidelity Taq polymerase (LA-Taq, Takara, Kyoto, Japan) using oligonucleotides 5′-CAGAAATCCCAAAACTTGTACC-3′ (sense) and 5′-CTGCTTTCTGCGCTTACACCT-3′ (antisense). The purified PCR fragment was first cloned into pCR2.1 vector with a TA-cloning kit (Invitrogen, Carlsbad, CA), confirmed by sequencing, and then recloned into pGL3 vector (Promega, Madison, WI). The mouse SIP1 and δEF1 cDNAs were obtained from Dr. F. van Roy (Ghent University) and Dr. Y. Higashi (Osaka University), respectively. The human E47 and Ets1 cDNAs were kindly provided by Dr. C. Murre (University of California, San Diego) and Dr. N. Kamata (Hiroshima University), respectively. Adenoviral vectors encoding SIP1 or δEF1 epitope-tagged with Flag at their N-termini were constructed using the ViraPower Adenoviral Gateway Expression System (Invitrogen). Adenoviruses were produced in transfected 293A cells and amplified in the same cells according to the manufacturer's protocol. Purification of adenoviruses was carried out using the Virakit for Adenovirus 5 and Recombinant Derivatives 4-Pack (Virapur, Carlsbad, CA). Antibodies Mouse monoclonal anti-Flag M2 and anti-α-tubulin antibodies were purchased from Sigma-Aldrich (St. Louis, MO), and mouse monoclonal anti-E-cadherin and anti-N-cadherin antibodies were from BD Transduction Laboratories (Lexington, KY). Rat monoclonal anti-E-cadherin antibody for immunostaining was from Zymed (San Francisco, CA). Goat polyclonal anti-fibronectin and anti-δEF1 antibodies were from Calbiochem (La Jolla, CA) and Santa Cruz Biotechnology (Santa Cruz, CA; E-20, Lot no. K0702), respectively. Transfection and Infection of DNA Transient transfection into NMuMG cells and MDCK cells was performed using Fugene 6 reagent (Roche Molecular Biochemicals, Indianapolis, IN) and Lipofectamine 2000 reagent (Invitrogen), respectively, as recommended by the manufacturers. For infection of adenoviruses, NMuMG cells were plated at a density of 1.0 × 105 cells/well in eight-well Culture Slides (BD Falcon, Bedford, MA). After 8 h, cells were infected with each adenovirus for 1 h with gentle agitation, washed once with the same media, and incubated for another 24 h. RNA Interference Transfection of short interfering RNAs (siRNAs) was performed in 12-well tissue culture plates according to the protocol recommended for HiPerFect reagent (QIAGEN). Final concentrations of the siRNAs used were 5 nM, except for 10 nM for SIP1 siRNA. At 8 h after transfection, 1 ng/ml TGF-β was added to the media, and culture was continued for an additional 24 h. The target sequences of these siRNA duplexes were as follows: mouse SIP1 (GGAAAAACGUGGUGAACUA; B-Bridge, Sunnyvale, CA), mouse δEF1 (Stealth RNAi MSS210696; Invitrogen), mouse E2A (Stealth RNAi MSS210711; Invitrogen), mouse Ets1 (Stealth RNAi MSS215651, MSS215652, and MSS215653; Invitrogen), and Negative Control (Stealth RNAi 12935-200; Invitrogen). Immunoblotting NMuMG cells were seeded at a density of 8.0 × 104 cells/well in 12-well tissue culture plates. After 12 h, cells were treated with 1 ng/ml TGF-β for 24 h. Cells were lysed in RIPA buffer solution (20 mM Tris-HCl, pH 7.5, 150 mM NaCl, 10 mM EDTA, 1 mM EGTA, 1% Nonidet P-40, 0.5% sodium deoxycholate, and 0.1% SDS). After measurement of protein concentrations with a BCA Protein Assay Kit (Pierce, Rockford, IL), equal amounts of total protein per lane were subjected to SDS gel-electrophoresis, followed by semidry transfer of the proteins to Fluoro Trans W membrane (Pall, Glen Cove, NY). Nonspecific binding of proteins to the membrane was blocked by incubation in TBS-T buffer (50 mM Tris-HCl, pH 7.4, 150 mM NaCl, and 0.1% Tween-20) containing 5% skim milk. Immunodetection was performed with the ECL blotting system (Amersham, Piscataway, NJ). Immunofluorescence Labeling To allow direct fluorescence of the actin cytoskeleton, cell were fixed in 3.7% paraformaldehyde in phosphate-buffered saline (PBS), permeabilized with 0.2% Triton X-100 in PBS for 5 min at room temperature, and subsequently stained with 0.25 mM tetramethylrhodamine isothiocyanate (TRITC)-conjugated phalloidin (Sigma-Aldrich). Immunocytochemical analyses were carried out on eight-well Culture Slides. Cell were fixed in 1:1 acetone-methanol solution and incubated with antibodies diluted with Blocking One solution (Nacalai Tesque, Tokyo, Japan) for 1 h at room temperature. The cells were then incubated with secondary antibodies and TOTO3 (Invitrogen Molecular Probes, Eugene, OR) for 1 h. Fluorescence was examined by confocal laser scanning microscopy (Carl Zeiss, Thornwood, NY). Extraction of RNAs and Quantitative RT-PCR Total RNAs were extracted from NMuMG cells using the RNeasy Mini Kit (QIAGEN). First-strand cDNAs were generated by Oligo (dT) priming using Superscript III Reverse Transcriptase (Invitrogen) following the manufacturer's instructions. Quantitative RT-PCR analyses were performed using the ABI PRISM 7500 Fast Real-Time PCR System (Applied Biosystems, Foster City, CA) and Power SYBR Green (Applied Biosciences, Foster City, CA). Luciferase Assays Cells were seeded in duplicate in 12-well tissue culture plates, followed by transient transfection with various combinations of promoter-reporter constructs and expression plasmids as required. Luciferase activity in cell lysates was determined with a dual luciferase reporter assay system (Promega) using a luminometer (AutoLumat LB953, EG&G Berthold, Natick, MA). Luciferase activity was normalized to sea-pansy luciferase activity of cotransfected phRL-TK plasmid (Promega). Electrophoretic Mobility Shift Assay The E-box2/1 WT probe covers the region from −84 to −73 of the mouse E-cadherin promoter. Double-stranded oligonucleotides were labeled with [γ- 32P]ATP and T4 polynucleotide kinase. Preparation of nuclear extracts was performed as previously described ( Kobayashi et al., 2007 ). Briefly, cells were infected with Flag-tagged SIP1 adenovirus, washed with PBS, collected in buffer A (10 mM HEPES-KOH, pH 7.9, 1.5 mM MgCl 2, 10 mM KCl, 0.5 mM dithiothreitol, 1 mM phenylmethylsulfonyl fluoride, and 50 U/ml aprotinin), and incubated on ice for 10 min. After centrifugation, the precipitate was washed with buffer A and suspended in buffer C (20 mM HEPES-KOH, pH 7.9, 420 mM NaCl, 1.5 mM MgCl 2, 0.2 mM EDTA, 0.5 mM dithiothreitol, 1 mM phenylmethylsulfonyl fluoride, and 50 U/ml aprotinin). The suspension was incubated on ice for 20 min, and the supernatant obtained by centrifugation was used as a nuclear fraction. DNA-binding assay (20 μl final volume) was carried out for 30 min at 4°C, with 5 μg of NMuMG nuclear proteins, poly(dI-dC) [poly(deoxyinosinic-deoxycytidylic acid)], and 32P-labeled double-stranded oligonucleotide in binding buffer (15 mM Tris-HCl, pH 7.5, 75 mM NaCl, 1.5 mM EDTA, 1.5 mM dithiothreitol, 7.5% glycerol, 0.3% Nonidet P-40, and 1 mg/ml bovine serum albumin). For supershift experiments, the extracts were incubated with anti-Flag M2 antibody and were loaded onto a 6% polyacrylamide gel prepared in 0.25× TBE buffer. After electrophoresis, gels were dried, exposed to imaging plates, and analyzed with the BAS-5000 system (FujiFilm, Tokyo, Japan). Cell Motility Assay NMuMG cells transfected with siRNAs were seeded at 1.0 × 105 cells/well in 6-well tissue culture plates. After 12 h, wounds were incised by scratching the cell monolayers using 200-μl pipette tips, and then 1 ng/ml TGF-β was added to the media. Photographs were taken under phase-contrast microscopy immediately after incision and after 24 h of ligand stimulation. The assays were independently performed in triplicate. The area of migrating cells was estimated by counting the number of pixels after the photographs had been converted to Photoshop data (Adobe, San Jose, CA). TGF-β Promotes EMT TGF-β has been reported to induce EMT in NMuMG cells ( Miettinen et al., 1994 ; Piek et al., 1999 ; Kondo et al., 2004 ). As previously reported, treatment with TGF-β dramatically altered the morphological phenotypes of NMuMG cells from cobblestone-like to spindle shapes (A). It also induced actin fiber formation typical of transdifferentiation and elicited the so-called cadherin switch, i.e., down-regulation of E-cadherin and up-regulation of N-cadherin (B). Immunoblotting of whole-cell lysates from NMuMG cells revealed that treatment with TGF-β for 24 h resulted in down-regulation of E-cadherin expression with concomitant up-regulation of representative mesenchymal markers, i.e., N-cadherin and fibronectin (C). Analyses by semiquantitative RT-PCR revealed that levels of E-cadherin mRNA gradually decreased and reached a minimum level at 12 h after TGF-β treatment (D). To determine whether certain transcription factors are regulated by TGF-β in a manner reciprocal to that of the reduction in E-cadherin expression, we examined the expression of SIP1, δEF1, Snail, Twist, and E12/47 mRNAs after TGF-β treatment. SIP1 and δEF1 mRNA levels were gradually increased by TGF-β, with profiles of expression reciprocal to that of E-cadherin (D). Greater than 10-fold expression of Snail mRNA was induced by 1 h after TGF-β treatment (E). Expression of Twist could not be detected, whereas that of E12/E47 was not altered after TGF-β treatment (data not shown and Kondo et al., 2004 ). Expression of mRNA for Id2, a negative regulator of TGF-β–induced EMT ( Kondo et al., 2004 ; Kowanetz et al., 2004 ), decreased rapidly by 4 h after treatment with TGF-β (D). These findings demonstrated that, among these factors, increases in SIP1 and δEF1 by TGF-β appear to strongly correlate with the reduction in E-cadherin in NMuMG cells. SIP1 and δEF1, But Not Snail, Repress E-Cadherin Expression Because expression of SIP1, δEF1, and Snail mRNAs was up-regulated by TGF-β in NMuMG cells, we next examined the effects of these factors on E-cadherin promoter activity. Transfection of ALK5TD, a constitutively active form of TβR-I, and E47 repressed E-cadherin promoter activity in NMuMG cells (A). Overexpression of either SIP1 or δEF1 strongly repressed E-cadherin promoter activity to the level equal to that suppressed by E47 in NMuMG cells. Protein levels of transfected SIP1 determined by immunoblotting were much less than those of transfected E47 (data not shown). Snail and Slug each repressed E-cadherin promoter activity in MDCK cells (A and data not shown). However, neither Snail, Slug, nor Twist exhibited significant effects on E-cadherin promoter activity in NMuMG cells, suggesting that Snail and Slug function in a cell context–dependent manner, as previously reported ( Barrallo-Gimeno and Nieto, 2005 ). SIP1 was originally identified as a Smad-interacting protein by yeast two-hybrid screening and was shown to contain the Smad-binding domain (SBD) at its N-terminus ( Verschueren et al., 1999 ). The degree of amino acid sequence similarity in the SBD between SIP1 and δEF1 is ~40%, and interaction of δEF1 with Smad2/3 was weaker than that of SIP1 (data not shown and Postigo, 2003 ), suggesting the possibility that the SBD is not necessary for repression of E-cadherin promoter activity. To test this, we prepared a SIP1 deletion mutant lacking SBD (SIP1-ΔSBD) and measured E-cadherin promoter activity after transient transfection of NMuMG cells. Similar to wild-type SIP1, the SIP1-ΔSBD mutant repressed E-cadherin promoter activity (B). Binding of the wild-type SIP1 to Smad2/3 was observed only in the presence of TGF-β, whereas that of SIP1-ΔSBD was not observed even in the presence of TGF-β (data not shown). These findings suggest that the interactions with Smads through the SBD are not required for repression by SIP1 and δEF1 of E-cadherin promoter activity. Consistent with the results of E-cadherin reporter assays, overexpression of either SIP1 or δEF1 by adenoviruses repressed the expression of E-cadherin mRNA as well as that of E-cadherin protein (, C and D). These findings suggest that SIP1 and δEF1 act as potent suppressors of E-cadherin transcription in NMuMG cells. δEF1 Family Proteins Bind to E-box1 and E-box2 in Mouse E-Cadherin Promoter The E-cadherin promoter contains two E-boxes (E-box1 and E-box3) in humans and three E-boxes (E-box1, E-box2, and E-box3) in mice. To determine whether SIP1 and δEF1 affect transcription of the mouse E-cadherin promoter through the E-boxes, we transfected SIP1 expression plasmid with a reporter plasmid driven by mouse E-cadherin core promoter (−178 to + 92) in mouse NMuMG cells. SIP1 and δEF1 induced an ~60% decrease in mouse E-cadherin promoter activity (, B and C). To determine which E-boxes are involved in the repression induced by SIP1 and δEF1, we mutated the E-boxes in the mouse E-cadherin promoter (A). SIP1 and δEF1 repressed the E-cadherin promoter activity of the single E-box point mutants (E1, E2, and E3) to extents similar to those of the wild-type E-cadherin promoter (, B and C). Among three double E-box mutants of the mouse E-cadherin promoter, the E-box1/E-box3 mutant (E13) and E-box2/E-box3 mutant (E23) exhibited activity similar to the wild-type E-cadherin promoter. In contrast, SIP1 and δEF1 failed to repress transcription of the E-box2/E-box1 double mutant (E21), similar to the triple mutant of mouse E-cadherin promoter (E213, , B and C). To confirm these findings, electrophoretic mobility shift assay (EMSA) against radioisotope-labeled mouse E-cadherin probes was performed using nuclear extracts of NMuMG cells overexpressing SIP1 (D). SIP1 bound to the wild-type E-box2/1 probe and gave rise to a specific band that was efficiently competed by the unlabeled wild-type probe but only weakly competed by excess (×50) unlabeled mutant probe. However, no binding of SIP1 protein could be detected in EMSA using double E-box2/1 mutant as a labeled probe. The specificity of binding of Flag-SIP1 to the E-box2/1 probe was confirmed by supershift assay using anti-Flag M2 antibody. Addition of anti-Flag mAb led to the disappearance of the SIP1-specific band and the appearance of a slowly migrating supershift complex (D). Similar results were obtained with overexpression of Flag-δEF1 (data not shown). These findings indicate that SIP1 and δEF1 directly repress mouse E-cadherin promoter activity through interaction with E-box2 and E-box1 elements of the mouse E-cadherin promoter in NMuMG cells. Double Knockdown of SIP1 and δEF1 Blocks TGF-β–induced E-Cadherin Repression and Cell Migration To determine whether SIP1 and δEF1 are required for TGF-β–mediated repression of E-cadherin promoter activity, we used siRNAs directed against SIP1 and δEF1 to reduce the expression of endogenous proteins. SIP1 or δEF1 siRNAs were transfected into NMuMG cells, followed by stimulation of the cells with TGF-β. SIP1 and δEF1 siRNAs each successfully knocked down the expression of corresponding endogenous mRNAs (A). Down-regulation of δEF1 protein expression was also confirmed by immunoblotting (see B). In cells transfected with control siRNA, TGF-β induced about twofold expression of SIP1 and δEF1 mRNAs at 24 h after stimulation and repressed the expression of E-cadherin within 24 h after stimulation (A). In cells transfected with either SIP1 siRNA or δEF1 siRNA alone, TGF-β–mediated E-cadherin repression was not or was only partially blocked. Interestingly, transfection of both SIP1 and δEF1 siRNAs completely abolished TGF-β–induced E-cadherin repression at the mRNA level (A). Immunostaining of E-cadherin in NMuMG cells also revealed that the level of E-cadherin protein was not suppressed by TGF-β in cells transfected with both SIP1 and δEF1 siRNAs (B). Next, we investigated whether functions of SIP1 and δEF1 are required for TGF-β–induced migration and invasion by cell motility assay. In cells transfected with control siRNA, TGF-β accelerated wound closure (C) and invasion (data not shown), possibly through E-cadherin–regulated cell–cell adhesion. These responses were partially impaired in cells transfected with both SIP1 and δEF1 siRNAs (C). These findings indicate that TGF-β requires both SIP1 and δEF1 to repress E-cadherin expression and migratory behavior of NMuMG cells. SIP1 and δEF1 Are Not Involved in the Up-Regulation of Mesenchymal Markers by TGF-β Because transfection with both SIP1 and δEF1 siRNAs completely abolished down-regulation of E-cadherin in response to TGF-β, we examined whether other EMT markers, including fibronectin and N-cadherin, are affected by these siRNAs in NMuMG cells. Cells were transfected with both SIP1 and δEF1 siRNAs. Nontransfected cells and those transfected with a control siRNA were used as controls. In the control cells, TGF-β down-regulated the expression of E-cadherin and up-regulated those of fibronectin and N-cadherin at both the mRNA and protein levels (, A and B). In contrast to the effect of knockdown of both SIP1 and δEF1 on TGF-β–induced E-cadherin repression, cells transfected with SIP1 and δEF1 siRNAs exhibited expression profiles of fibronectin and N-cadherin similar to those of the control cells (, A and B). Moreover, examination of actin reorganization by TRITC-phalloidin staining in the control cells revealed dramatic actin fiber formation by TGF-β, which was also observed in cells transfected with both SIP1 and δEF1 siRNAs (C). These findings strongly suggest that SIP1 and δEF1 are not involved in regulation of the expression of mesenchymal markers by TGF-β. Activation of Smad Signaling and Subsequent De Novo Protein Synthesis Are Required for TGF-β–induced SIP1 Expression To elucidate whether de novo protein synthesis is required for TGF-β–induced SIP1 and δEF1 expression, RNAs were extracted from cells stimulated with TGF-β in the presence or absence of cycloheximide, an inhibitor of protein synthesis, and analyzed by semiquantitative RT-PCR. Because rapid increase in Smad7 mRNA by TGF-β has been reported as a direct effect of Smad signaling activated by TGF-β ( Akiyoshi et al., 2001 ), Smad7 was used as a negative control to evaluate the effect of cycloheximide. Snail mRNA was up-regulated until 12 h by TGF-β ( and see E). Similar to the effect of cycloheximide on TGF-β–induced increase in Smad7, induction of Snail was not affected by cycloheximide, indicating that Snail is a direct target of the TGF-β-Smad pathway. In contrast, although TGF-β induced expression of SIP1 mRNA in the absence of cycloheximide, pretreatment of cells with cycloheximide blocked the TGF-β–mediated induction of SIP1 () and δEF1 (data not shown). The up-regulation of SIP1 and δEF1 mRNAs may thus be an indirect transcriptional response and require de novo protein synthesis by TGF-β. Increase in Ets1 by TGF-β Is Involved in Expression of SIP1 and δEF1 To examine the molecular mechanisms by which δEF1 and SIP1 are transcriptionally activated through de novo–synthesized proteins in response to TGF-β, we investigated TGF-β–regulated genes from our microarray data for human HaCaT keratinocyte cells ( Akiyoshi et al., 2001 ) and from published data of NMuMG cells ( Xie et al., 2003 ). Expression of several representative genes was up-regulated by TGF-β in HaCaT cells; among such genes, we found that Ets family genes were induced by TGF-β, even in the presence of cycloheximide in HaCaT cells ( Akiyoshi et al., 2001 ). Expression of Ets family genes was also up-regulated in NMuMG cells ( Xie et al., 2003 ). Semiquantitative RT-PCR analyses using RNAs from NMuMG cells treated with TGF-β revealed that, among several Ets family proteins, including Ets1, Ets2, and ESE-1, only Ets1 was increased after TGF-β stimulation (A), and that this increase was unaffected by cycloheximide pretreatment (data not shown). To examine the roles of Ets1 in TGF-β–induced EMT, we determined the effect of Ets1 on E-cadherin promoter activity. Transfection of Ets1 in NMuMG cells resulted in dose-dependent repression of E-cadherin promoter activity (B), suggesting that repression of E-cadherin by Ets1 may be mediated by SIP1 and/or δEF1. To explore this possibility, δEF1 promoter was cloned from genomic DNA of HaCaT cells. Computer analyses showed that sequence similarity between the mouse and human δEF1 promoter regions is ~65–70% in the −620 to +1 region of the mouse promoter (corresponding to −810 to +1 of the human promoter) and that these regions contain two potential binding sites for Ets1 and three binding sites for bHLH E47 proteins. Transient transfection of Ets1 into NMuMG cells induced a threefold increase in δEF1 promoter activity (C). siRNA directed against Ets1 successfully and specifically knocked down the expression of endogenous Ets1 without off-target effects on other Ets family genes (D). In cells transfected with the Ets1 siRNA, TGF-β–induced expression of SIP1 and δEF1 was inhibited compared with that in cells transfected with control siRNA, suggesting that Ets1 may be involved in TGF-β–regulated induction of δEF1 and SIP1 and that it may in turn repress E-cadherin transcription. We and another group have previously reported that, in NMuMG cells, endogenous Id2 acts as a negative regulator of TGF-β–induced EMT and that overexpression of Id2 partially blocks the E-cadherin repression and EMT phenotype evoked by TGF-β ( Kondo et al., 2004 ; Kowanetz et al., 2004 ). To investigate whether Id2 regulates the effect of Ets1 on up-regulation of δEF1 and SIP1, we examined the induction of SIP1/δEF1 by TGF-β in NMuMG cells retrovirally infected with pBabe-mouse Id2. In cells infected with control pBabe vector, endogenous Id2 expression was down-regulated by TGF-β, and δEF1 mRNA was increased by TGF-β (E), similar to the noninfected NMuMG cells (see A). In Id2-infected NMuMG cells, high levels of endogenous Id2 mRNA were maintained and were not affected by TGF-β. TGF-β failed to up-regulate the expression of δEF1 (D) and SIP1 (data not shown) in these cells, despite the finding that TGF-β could increase the expression of Ets1 (data not shown). Supporting these observations, the δEF1 promoter activity induced by Ets1 was enhanced by cotransfection with constitutively active TβR-I (ALK5TD), which was partially inhibited by cotransfection with Id2 (C). Taken together, these findings suggest that Id2 may regulate the function of Ets1 to modulate the transcription of SIP1 and δEF1 without alteration of the transcription of Ets1. Ets1-regulated δEF1/SIP1 transcription may thus be of great importance for TGF-β–induced EMT in NMuMG cells. In the present study, we investigated the roles of transcriptional repressors for E-cadherin in regulation of TGF-β–induced EMT in NMuMG cells and found that the δEF1 family proteins, δEF1 and SIP1, are required for TGF-β–induced repression of E-cadherin but not for up-regulation of mesenchymal markers. We also found that TGF-β–induced expression of Ets1 promoted transcription of SIP1 and δEF1 mRNAs through potential cooperation with E12/E47 bHLH transcription factors. δEF1 Family Proteins Are Essential for Repression of E-Cadherin Expression by TGF-β Various transcription factors have been reported to be involved in EMT ( Barrallo-Gimeno and Nieto, 2005 ). Among such factors, expression of SIP1 and δEF1 mRNAs was gradually increased by TGF-β in NMuMG cells with expression profiles reciprocal to that of E-cadherin (D). Although SIP1 and δEF1 (ZEB1) have been reported to play opposing roles in TGF-β and BMP signaling ( Postigo, 2003 ), we have found that both SIP1 and δEF1 repress the expression of E-cadherin and are essential for TGF-β–induced EMT in NMuMG cells. In agreement with our findings, it has been reported that overexpression of SIP1 in the absence of TGF-β fully suppressed E-cadherin promoter activity in NMe cells, a line of cells derived from NMuMG cells ( Comijn et al., 2001 ) and that overexpression of δEF1 also repressed E-cadherin expression in epithelial cells ( van Grunsven et al., 2003 ). SIP1 was originally identified as a Smad-interacting protein by yeast two-hybrid screening and was shown to contain the SBD at its N-terminus ( Verschueren et al., 1999 ). SIP1 interacts with R-Smads through the SBD in a TGF-β–dependent manner. In contrast, δEF1 interacts with Smads only weakly even in the presence of TGF-β (data not shown and Postigo, 2003 ), possibly due to the low degree of sequence similarity in the SBD between δEF1 and SIP1. Although δEF1 and SIP1 have been reported to regulate the promoter activity of p3TP-lux and p21-lux in TGF-β–dependent manner ( Postigo, 2003 ; Postigo et al., 2003 ) and Comijin et al. (2001) reported that a SIP1 deletion mutant lacking SBD failed to repress the transcription of human E-cadherin in MCF7 cells, the present findings revealed that interaction of Smads with SIP1 is not required for repression of transcription of E-cadherin. These findings indicate that SIP1 and δEF1 regulate mouse E-cadherin transcription independently of interaction with Smads in NMuMG cells. Comijin et al. (2001) also reported that the human E-cadherin promoter contains E-box1 and E-box3, but lacks E-box2, and that SIP1 suppresses transcription of human E-cadherin promoter activity in human MCF7 cells through E-box1 and E-box3. However, in mouse NMuMG cells, E-box1 and E-box2 were sufficient for the repression of mouse E-cadherin transcription by SIP1 and δEF1, and E-box3 was not required for this repression (). Using chromatin immunoprecipitation assays, we have found that overexpressed δEF1 and SIP1 bound to E-box2/1 elements of mouse E-cadherin promoter (data not shown). Although the reason for this difference is unclear, it is possible that SIP1 and δEF1 act on E-box3 through interaction with Smads and that E-box2, which is located close to E-box1 in the mouse E-cadherin promoter, may compensate for the function of E-box3. Further studies using mouse and human E-cadherin promoters and using chromatin immunoprecipitation assays by endogenous δEF1, SIP1, and Snail are required in the future. Snail Is Not Involved in the Repression of E-Cadherin Expression in NMuMG Cells Snail has been shown to be induced by TGF-β and to play pivotal roles in E-cadherin repression and EMT in various cells in vitro and in vivo ( Barrallo-Gimeno and Nieto, 2005 ). Although Snail mRNA expression was strongly induced by TGF-β in NMuMG cells (E), we were not able to determine Snail protein levels because of the low efficiency of several commercially available antibodies to Snail (our unpublished data). Overexpression of Snail failed to affect the phenotypes of NMuMG cells and to regulate the expression of EMT markers, including E-cadherin, N-cadherin, fibronectin, and vimentin (A and Supplementary Figure 1, D and E). Experiments using siRNA also demonstrated that TGF-β efficiently induced EMT markers in NMuMG cells transfected with Snail siRNA (Supplementary Figure 1, A–C). Snail has recently been reported to be activated during EMT through various mechanisms, including up-regulation by HMGA2, phosphorylation by GSK3β, activation by lysyl oxidase-like 2, and induction of nuclear localization by zinc transporter LIV1 ( Yamashita et al., 2004 ; Zhou et al., 2004 ; Peinado et al., 2005 ; Thuault et al., 2006 ). The present findings, however, suggest that Snail is not the primary mediator of TGF-β–mediated EMT in NMuMG cells and that it may require other molecules to induce EMT in these cells. Ets1 and Id Proteins Act as Upstream Factors for SIP1 and δEF1 Ets1 has been shown to be a direct target of TGF-β-Smad signaling in some cells ( Akiyoshi et al., 2001 ; del Valle-Perez et al., 2004 ). In the present study, we also observed that TGF-β up-regulated the expression of Ets1, which was not affected by cycloheximide treatment (data not shown). Because cycloheximide treatment blocked TGF-β–induced expression of SIP1 and δEF1 mRNAs, Ets1 is a direct target, and SIP1 and δEF1 are indirect targets, of TGF-β-Smads pathways in NMuMG cells. The promoter-reporter assay we performed revealed that Ets1 activated the activity of δEF1 promoter and suppressed the activity of E-cadherin promoter (, B and C). Knock-down of Ets1 decreased the mRNA levels of endogenous δEF1 or SIP1 (D). Interestingly, in Ets1 siRNA-transfected cells, expression of Ets1 mRNA was remarkably suppressed in the presence and absence of TGF-β, whereas mRNA levels of SIP1 and δEF1 in the absence of TGF-β were similar to those in control siRNA-transfected cells (D). This may have been due to redundant effects of other Ets family transcription factors; we observed that expression of Ets2 and ESE1, a breast cancer-specific Ets protein, was up-regulated in NMuMG cells when Ets1 expression was knocked down by Ets1-specific siRNA (our unpublished data). Previous studies reported that the transcription activity of Ets1 is regulated through phosphorylation by certain serine-threonine kinases, including ERK1/2 ( Tootle and Rebay, 2005 ). Recently, importance of the cross-talk between Ras and TGF-β signaling pathways has been reported, although roles of Ras signaling on TGF-β signaling appear to be context-dependent ( Gotzmann et al., 2002 ; Peinado et al., 2003 ). Smad1 is phosphorylated by Ras-activated ERK MAP kinases and degraded by Smurf1 ( Sapkota et al., 2007 ), whereas Smad2/3 are irregularly translocated to the nucleus in certain cancer cells transfected with active Ras ( Yamagata et al., 2005 ; Oft et al., 2002 ). Interestingly, it was reported that TGF-β–induced EMT was found only in the presence of active Ras ( Gotzmann et al., 2002 ), suggesting that Ets1 would be a key molecule for the cross-talk between TGF-β and Ras signaling pathways. In the present study, we demonstrated that, although overexpression of Ets1 accelerated δEF1 promoter activity, coexpression with constitutively active TβR-I (ALK5TD) further enhanced the δEF1 promoter activity induced by Ets1 (C). These findings suggest that Ets1 is activated through posttranslational modifications, including phosphorylation, by TGF-β in NMuMG cells. We and another group have previously reported that HLH transcriptional inhibitor Id proteins inhibit TGF-β–induced E-cadherin repression through constitutive association with E12/E47 bHLH transcriptional factors in NMuMG cells ( Kondo et al., 2004 ; Kowanetz et al., 2004 ). Thus, down-regulation of Id proteins by TGF-β abolishes this inhibition, leading to EMT in NMuMG cells. Consistent with these findings, when Id2 is overexpressed in NMuMG cells, TGF-β–induced E-cadherin repression and δEF1 induction were partially impaired (E and Kondo et al., 2004 ). Previous studies reported that Id proteins bind directly to Ets1 through the conserved HLH domain in vitro ( Yates et al., 1999 ), and we have shown here that Ets1-induced activation of the δEF1 promoter was partially suppressed by overexpression of Id2 (C). These findings suggest inhibitory effects of Id proteins on Ets1 in TGF-β–induced EMT. In addition, Ets1 activity on δEF1 promoter was partially repressed by transfection of E12/E47 siRNA (Supplementary Figure 2, A and B), indicating that Ets1 may act in cooperative manner with E47 to regulate expression of δEF1. However, it is currently unknown whether Ets1 directly interacts with E47 or binds with it indirectly through other molecule(s) in TGF-β–induced EMT ( Dang et al., 1998 ). Taken together, these findings suggest that collaboration of Ets1 with E47 may regulate the expression of SIP1 and δEF1, which may occur in a cell- and promoter-context manner and that down-regulation of Id protein expression by TGF-β may thus enhance this cooperation in inducing expression of δEF1 family proteins, leading to EMT. SIP1 and δEF1 Regulate the Expression of E-Cadherin, But Not That of Mesenchymal Markers The findings of the present study suggest that transcriptional regulation of δEF1 family proteins is required for regulation of E-cadherin during TGF-β–induced EMT and that Ets1 may act as an inducer of δEF1 family genes in collaboration with E47. However, the molecular mechanisms by which Id proteins inhibit the effects of Ets1 on induction of ZEB and SIP1 in NMuMG cells remain to be elucidated. In the process of EMT, the transcriptional program of epithelial cells shifts toward that of mesenchymal cells. Various transcription factors have been suggested to play key roles in this process. In TGF-β–induced EMT, Smad signaling has been reported to be essential for EMT ( Piek et al., 1999 ), whereas certain signals involved in EMT are transmitted via non-Smad pathways. In the present study, we showed that, in addition to SIP1 ( Comijn et al., 2001 ), δEF1 plays a key role in TGF-β–induced EMT in NMuMG cells. Moreover, several non-Smad pathways, including the RhoA ( Bhowmick et al., 2001 ) and p160 ROCK pathways ( Ozdamar et al., 2005 ), have been reported to transduce signals for TGF-β–induced EMT, and expression of fibronectin occurs independently of Smad activation ( Hocevar et al., 1999 ). Although SIP1 and δEF1 are transcription factors of critical importance to the induction of EMT, they appear to regulate only certain subsets of EMT markers. EMT may thus be induced by an array of signals activated by TGF-β as well as by other cytokines. The mechanism of induction of mesenchymal markers during TGF-β–induced EMT will require further determination in the future. We thank Ms. E. Ohara for technical assistance, Drs. N. Kobayashi, K. Tobiume, M. Taki, and N. Kamata for their advice, and all the members of the Molecular Pathology Laboratory and the Biochemistry Laboratory of the Cancer Institute of the Japanese Foundation for Cancer Research, Tokyo, for critical comments. This work was supported by KAKENHI (Grants-in-Aid for Scientific Research) from the Ministry of Education, Culture, Sports, Science and Technology, Japan and a grant from the Uehara Memorial Foundation. | | TGF-β | transforming growth factor-β | | | HLH | helix-loop-helix | | | EMT | epithelial-mesenchymal transition | | | SIP1 | Smad-interacting protein1 | | | δEF1 (ZEB1) | delta-crystallin/E2-box factor 1. |
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