TGF-β regulates isoform switching of FGF receptors and epithelial-mesenchymal transition
2011; Springer Nature; Volume: 30; Issue: 4 Linguagem: Inglês
10.1038/emboj.2010.351
ISSN1460-2075
AutoresTakuya Shirakihara, Kana Horiguchi, Keiji Miyazawa, Shogo Ehata, Tatsuhiro Shibata, Ikuo Morita, Kohei Miyazono, Masao Saitoh,
Tópico(s)Cancer Cells and Metastasis
ResumoArticle11 January 2011Open Access TGF-β regulates isoform switching of FGF receptors and epithelial–mesenchymal transition Takuya Shirakihara Takuya Shirakihara Department of Molecular Pathology, Graduate School of Medicine, University of Tokyo, Tokyo, Japan Division of Cancer Genomics, National Cancer Center Research Institute, Tokyo, Japan Search for more papers by this author Kana Horiguchi Kana Horiguchi Department of Molecular Pathology, Graduate School of Medicine, University of Tokyo, Tokyo, Japan Search for more papers by this author Keiji Miyazawa Keiji Miyazawa Department of Biochemistry, Interdisciplinary Graduate School of Medicine and Engineering, University of Yamanashi, Yamanashi, Japan Search for more papers by this author Shogo Ehata Shogo Ehata Department of Molecular Pathology, Graduate School of Medicine, University of Tokyo, Tokyo, Japan Search for more papers by this author Tatsuhiro Shibata Tatsuhiro Shibata Division of Cancer Genomics, National Cancer Center Research Institute, Tokyo, Japan Search for more papers by this author Ikuo Morita Ikuo Morita Department of Cellular Physiological Chemistry and Global Center of Excellence (GCOE) Program, International Research Center for Molecular Science in Tooth and Bone Diseases, Graduate School, Tokyo Medical and Dental University, Tokyo, Japan Search for more papers by this author Kohei Miyazono Corresponding Author Kohei Miyazono Department of Molecular Pathology, Graduate School of Medicine, University of Tokyo, Tokyo, Japan Search for more papers by this author Masao Saitoh Corresponding Author Masao Saitoh Department of Molecular Pathology, Graduate School of Medicine, University of Tokyo, Tokyo, Japan Department of Biochemistry, Interdisciplinary Graduate School of Medicine and Engineering, University of Yamanashi, Yamanashi, Japan Department of Cellular Physiological Chemistry and Global Center of Excellence (GCOE) Program, International Research Center for Molecular Science in Tooth and Bone Diseases, Graduate School, Tokyo Medical and Dental University, Tokyo, Japan Search for more papers by this author Takuya Shirakihara Takuya Shirakihara Department of Molecular Pathology, Graduate School of Medicine, University of Tokyo, Tokyo, Japan Division of Cancer Genomics, National Cancer Center Research Institute, Tokyo, Japan Search for more papers by this author Kana Horiguchi Kana Horiguchi Department of Molecular Pathology, Graduate School of Medicine, University of Tokyo, Tokyo, Japan Search for more papers by this author Keiji Miyazawa Keiji Miyazawa Department of Biochemistry, Interdisciplinary Graduate School of Medicine and Engineering, University of Yamanashi, Yamanashi, Japan Search for more papers by this author Shogo Ehata Shogo Ehata Department of Molecular Pathology, Graduate School of Medicine, University of Tokyo, Tokyo, Japan Search for more papers by this author Tatsuhiro Shibata Tatsuhiro Shibata Division of Cancer Genomics, National Cancer Center Research Institute, Tokyo, Japan Search for more papers by this author Ikuo Morita Ikuo Morita Department of Cellular Physiological Chemistry and Global Center of Excellence (GCOE) Program, International Research Center for Molecular Science in Tooth and Bone Diseases, Graduate School, Tokyo Medical and Dental University, Tokyo, Japan Search for more papers by this author Kohei Miyazono Corresponding Author Kohei Miyazono Department of Molecular Pathology, Graduate School of Medicine, University of Tokyo, Tokyo, Japan Search for more papers by this author Masao Saitoh Corresponding Author Masao Saitoh Department of Molecular Pathology, Graduate School of Medicine, University of Tokyo, Tokyo, Japan Department of Biochemistry, Interdisciplinary Graduate School of Medicine and Engineering, University of Yamanashi, Yamanashi, Japan Department of Cellular Physiological Chemistry and Global Center of Excellence (GCOE) Program, International Research Center for Molecular Science in Tooth and Bone Diseases, Graduate School, Tokyo Medical and Dental University, Tokyo, Japan Search for more papers by this author Author Information Takuya Shirakihara1,2, Kana Horiguchi1, Keiji Miyazawa3, Shogo Ehata1, Tatsuhiro Shibata2, Ikuo Morita4, Kohei Miyazono 1 and Masao Saitoh 1,3,4 1Department of Molecular Pathology, Graduate School of Medicine, University of Tokyo, Tokyo, Japan 2Division of Cancer Genomics, National Cancer Center Research Institute, Tokyo, Japan 3Department of Biochemistry, Interdisciplinary Graduate School of Medicine and Engineering, University of Yamanashi, Yamanashi, Japan 4Department of Cellular Physiological Chemistry and Global Center of Excellence (GCOE) Program, International Research Center for Molecular Science in Tooth and Bone Diseases, Graduate School, Tokyo Medical and Dental University, Tokyo, Japan *Corresponding authors: Department of Molecular Pathology, Graduate School of Medicine, University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo 113-0033, Japan. Tel.: +81 35841 3345; Fax: +81 35841 3354; E-mail: [email protected] of Biochemistry, Interdisciplinary Graduate School of Medicine and Engineering, University of Yamanashi, 1110 Shimokato, Chuo, Yamanashi 409-3899, Japan. Tel.: +81 55273 9496; Fax: +81 55273 6784; E-mail: [email protected] The EMBO Journal (2011)30:783-795https://doi.org/10.1038/emboj.2010.351 PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info The epithelial–mesenchymal transition (EMT) is a crucial event in wound healing, tissue repair, and cancer progression in adult tissues. Here, we demonstrate that transforming growth factor (TGF)-β induced EMT and that long-term exposure to TGF-β elicited the epithelial–myofibroblastic transition (EMyoT) by inactivating the MEK-Erk pathway. During the EMT process, TGF-β induced isoform switching of fibroblast growth factor (FGF) receptors, causing the cells to become sensitive to FGF-2. Addition of FGF-2 to TGF-β-treated cells perturbed EMyoT by reactivating the MEK-Erk pathway and subsequently enhanced EMT through the formation of MEK-Erk-dependent complexes of the transcription factor δEF1/ZEB1 with the transcriptional corepressor CtBP1. Consequently, normal epithelial cells that have undergone EMT as a result of combined TGF-β and FGF-2 stimulation promoted the invasion of cancer cells. Thus, TGF-β and FGF-2 may cooperate with each other and may regulate EMT of various kinds of cells in cancer microenvironment during cancer progression. Introduction The epithelial–mesenchymal transition (EMT) is a differentiation switch that directs polarized epithelial cells to differentiate into mesenchymal cells. Recent studies have proposed that EMT can be classified into three subtypes based on the biological context (Thiery and Sleeman, 2006; Kalluri and Weinberg, 2009; Kalluri, 2009; Zeisberg and Neilson, 2009). Type 1 EMT involves the transition of primitive epithelial cells to motile mesenchymal cells during gastrulation and the generation of migrating-neural crest cells from primitive neuroepithelial cells. This process can generate mesenchymal cells that have the potential to subsequently undergo the mesenchymal–epithelial transition, leading to the generation of secondary epithelia. Type 2 EMT involves the transition of secondary epithelial or endothelial cells to resident tissue fibroblasts and is associated with wound healing, tissue regeneration, and tissue fibrosis. Type 2 EMT begins as a part of a repair-associated event that normally generates fibroblasts for tissue reconstruction after trauma and inflammatory injury. Type 3 EMT occurs in epithelial carcinoma cells in primary nodules that are transitioning to metastatic tumour cells. This process can affect both oncogenes and anti-oncogenes, and carcinoma cells that undergo type 3 EMT can invade and metastasize, resulting in cancer progression. Thus far, nearly all cases of EMT in adult tissues have been shown to be regulated by extracellular matrix components and soluble growth factors or cytokines, including epidermal growth factor, hepatocyte growth factor, fibroblast growth factors (FGFs), and transforming growth factor (TGF)-βs (Thiery and Sleeman, 2006; Acloque et al, 2009). Among these factors, TGF-β and FGF are considered key mediators of EMT and are frequently and abundantly expressed in various tumours. TGF-β signalling not only contributes to EMT during embryonic development, but also induces EMT in cancer cells during cancer progression (Moustakas and Heldin, 2007). TGF-β activates Smad proteins and activated Smads transcriptionally regulate several genes including δEF1, Snail, Twist, HMGA2, and Ids, which lead to EMT particularly through the transcriptional repression of E-cadherin (Zavadil and Bottinger, 2005; Horiguchi et al, 2009). The E-cadherin repressors that are regulated by TGF-β appear to function in a cell context-dependent manner, and δEF1 is one of the most well-characterized factors that is involved in TGF-β-induced EMT in mouse epithelial cells (Shirakihara et al, 2007). δEF1 (also known as ZEB1 or TCF-8) and SIP1 (known as ZEB2) are members of the δEF1 family. δEF1 can bind directly to the E-cadherin promoter and repress the E-cadherin expression. In addition, δEF1 also regulates the TGF-β-mediated epithelial–myofibroblastic transition (EMyoT). δEF1 seems to positively regulate the transcription of α-smooth muscle actin (α-SMA), a representative myofibroblast marker, by binding to its promoter in vascular smooth muscle cells (Nishimura et al, 2006). Among various mammalian FGFs, FGF-2 and FGF-4 are key regulators of EMT during development and cancer progression (Strutz et al, 2002; Strutz and Neilson, 2003). FGFs execute diverse functions by binding and activating members of the FGF receptor (FGFR) family. There are four FGFR genes (FGFR1–FGFR4) that encode functional receptors that consist of three extracellular immunoglobulin domains (Ig-I, Ig-II, and Ig-III), a single-transmembrane domain, and a cytoplasmic tyrosine kinase domain (Eswarakumar et al, 2005). FGFRs have several isoforms as exon skipping removes the Ig-I domain. In addition, alternative splicing in the second half of the Ig-III domain in FGFR1–FGFR3 produces the IIIb (FGFR1IIIb–FGFR3IIIb) and IIIc (FGFR1IIIc–FGFR3IIIc) isoforms that have distinct FGF-binding specificities and are predominantly expressed in epithelial and mesenchymal cells, respectively. FGF-2 binds preferentially to the IIIc isoforms, whereas FGF-7 and FGF-10 bind exclusively to the IIIb isoforms. FGF-2 is highly expressed in wounds and tumour tissues. However, it has not been determined how FGF-2 transmits signals to induce EMT and promote cancer progression because epithelial cells typically express the FGFRIIIb isoforms, which do not bind to FGF-2. Recently, it was reported that advanced cancer cells overexpress FGFR1IIIc (Foster et al, 1999; Acevedo et al, 2007), but it is still unknown how epithelial cells and cancer cells upregulate the expression of the FGFRIIIc isoforms. In the present study, we investigated the properties of EMT induced by TGF-β in cooperation with FGFs. TGF-β induced EMT, and prolonged treatment with TGF-β induced EMyoT. During TGF-β-mediated EMT, TGF-β altered the sensitivities of cells from FGF-7 to FGF-2 through FGFR isoform switching. In addition, FGF-2 prevented TGF-β-mediated EMyoT and enhanced EMT with more aggressive characteristics that resemble those of activated fibroblasts. Moreover, the cells generated through EMT mediated by FGF-2 and TGF-β facilitated cancer cell invasion, when the cells undergoing EMT were mixed with cancer cells. Therefore, TGF-β and FGF-2 cooperate with each other and may regulate the EMT of various kinds of cells in cancer microenvironment. Results FGFR isoform switching during TGF-β-induced EMT Although the FGFRIIIc isoforms are not predominantly expressed in epithelial cells, their expression and FGF-2 expression have been correlated with cancer progression. We hypothesized that TGF-β primes FGFR isoform switching in cells undergoing EMT and subsequently alters the sensitivity of cells to FGF ligands. To verify this possibility, we first examined the mRNA profiles of FGFR variants in response to TGF-β by quantitative RT–PCR analyses. FGFR1 mRNA gradually increased to ∼3.5-fold in mouse normal mammary epithelial NMuMG cells 24 h after TGF-β stimulation, whereas FGFR2 mRNA levels decreased over this time period (Figure 1A). Conventional RT–PCR analyses using isoform-specific primers revealed that FGFR1 upregulated by TGF-β was FGFR1IIIc (the mesenchymal isoform), whereas the FGFR2 downregulated by TGF-β was FGFR2IIIb (the epithelial isoform) (Figure 1B). FGFR1IIIb, FGFR2IIIc, FGFR3, and FGFR4 were not clearly detected in NMuMG cells. The isoform switching of FGFRs was also detected in mouse mammary gland epithelial Eph4 cells that underwent EMT by TGF-β. Interestingly, the addition of FGF-2 to TGF-β-treated cells sustained expression of FGFR1 and FGFR2 at higher and lower levels, respectively, for 72 h (Figure 1C). Consistent with the changes in the mRNA expression profiles of the FGFR isoforms, Erk phosphorylation was observed in NMuMG cells in response to FGF-7, but not FGF-2, whereas Erk phosphorylation was observed in TGF-β-pretreated cells in response to FGF-2, but not FGF-7 (Figure 1D). These findings indicate that TGF-β primes isoform switching of FGFRs during EMT and thereby changes the sensitivities of cells from FGF-7 to FGF-2. Figure 1.Isoform switching of FGFRs during TGF-β-induced EMT. (A) The kinetics of FGFR1 and FGFR2 expression were examined in NMuMG cells treated with 1 ng/ml TGF-β for the indicated periods by quantitative RT–PCR analysis. The ratio of the mRNA levels in TGF-β-treated cells compared with non-treated cells is shown. Each value represents the mean±s.d. of duplicate determinations from a representative experiment. Similar results were obtained in at least three independent experiments. (B) Expression of the alternatively spliced forms of FGFR1 and FGFR2 was examined by RT–PCR using RNA samples from NMuMG cells treated with TGF-β for 36 h. A schematic illustration of the primers is shown at the top. The primers for the IIIb and IIIc isoforms were the sp1–ap1 pair and sp2–ap1 pair, respectively. Ig, extracellular immunoglobulin-like domain; KD, kinase domain. FGFR3 and FGFR4 were not detected in NMuMG cells by quantitative RT–PCR. (C) Regulation of FGFR1 expression (left) and FGFR2 expression (right) by TGF-β and/or FGF-2 were examined by quantitative RT–PCR analysis in NMuMG cells. Values were normalized to the housekeeping gene TBP and are indicated as fold differences compared with the non-treated cells. Each value represents the mean±s.d. of duplicate determinations from a representative experiment. Similar results were obtained in three independent experiments. (D) Erk1/2 phosphorylation (p-Erk1/2) by FGF-2 or FGF-7 was examined by immunoblot analysis. NMuMG cells were preincubated with or without TGF-β for 2 days and then stimulated with 30 ng/ml FGF-2 or 30 ng/ml FGF-7 for 15 min. F2, FGF-2; F7, FGF-7. Download figure Download PowerPoint Activated phenotype of cells generated by FGF-2 and TGF-β treatment Next, we determined whether FGF-2 enhanced TGF-β-induced EMT after TGF-β-induced FGFR isoform switching, because previous studies have shown that TGF-β-induced EMT can be enhanced with oncogenic signals or growth factors (Horiguchi et al, 2009). To examine this possibility, NMuMG cells were continuously exposed to TGF-β, FGF-2, or both for several days. As we previously reported (Shirakihara et al, 2007), the morphology of NMuMG cells clearly changed from a cobblestone-like shape to a fibroblastic spindle shape upon prolonged TGF-β treatment, whereas cells treated with FGF-2 alone did not exhibit morphological changes (Figure 2A). Compared with cells treated with TGF-β alone, the addition of FGF-2 to TGF-β-treated cells led to drastic changes in cell morphology and actin fibre formation that are typical of fibroblastic differentiation (Figure 2A and B). Figure 2.Activated phenotype of cells generated by treating with FGF-2 and TGF-β. (A) NMuMG cells were incubated for 4 days in the absence or presence of 1 ng/ml TGF-β alone, 30 ng/ml FGF-2 alone, or both TGF-β and FGF-2, and visualized by phase-contrast microscopy. Scale bar indicates 100 μm. (B) Actin cytoskeleton reorganization was visualized by TRITC-phalloidin staining 4 days after TGF-β or TGF-β+FGF-2 treatment. Scale bar indicates 100 μm. (C) NMuMG cells were cultured for 4 days in the absence or presence of TGF-β, FGF-2, or both, and then the monolayers were wounded. After 12 h, the migratory behaviours of the cells were determined and quantified (right). Bars represent the median for each category from a representative experiment. Similar results were obtained in two independent experiments. The median migration distance of the cells was significantly different for non-treated versus TGF-β+FGF-2, TGF-β versus TGF-β+FGF-2, and FGF-2 versus TGF-β+FGF-2 (P<0.001; non-parametric Mann–Whitney U test). Scale bar indicates 100 μm. (D) NMuMG cells were cultured for 4 days in the absence or presence of TGF-β or both TGF-β and FGF-2, and then seeded into cell culture inserts. After 12 h, the cells that had migrated to the opposite side were stained and quantified. Each value represents the mean±s.d. of triplicate determinations from a representative experiment. Similar results were obtained in two independent experiments. (E) NMuMG cells were preincubated with or without the indicated combinations of TGF-β, FGF-2, and 10 μM GM6001 for 5 days, and then mixed in collagen matrices. After polymerization, the mixtures were released from the culture dishes and incubated for an additional 2 days. Three independent experiments were performed in duplicate, and representative results are shown (#1 in blue and #2 in red). (F) The conditioned media from NMuMG cells treated with TGF-β or both TGF-β and FGF-2 were collected and applied to 10% polyacrylamide gel impregnated with 1 mg/ml gelatin. After removing SDS, the gels were incubated overnight and stained with CBB. Download figure Download PowerPoint Next, cell motility was examined using a wound closure assay with NMuMG cells. After treating with TGF-β, FGF-2, or both for 4 days, the cells were wounded by scratching and were then analysed after 12 h. Figure 2C shows that NMuMG cells treated with TGF-β alone had slightly enhanced cell motility, compared with non-treated cells or FGF-2-treated cells. On the other hand, treating with TGF-β and FGF-2 strongly promoted the motility of NMuMG cells within only 12 h. Similar to their ability to enhance migration, combined TGF-β and FGF-2 treatment remarkably promoted the invasion of NMuMG cells in an in vitro invasion assay (Figure 2D). As one of the most characteristic features of fibroblasts is their ability to degrade extracellular matrix, this property was also determined by a collagen gel degradation/contraction assay (Mikko et al, 2008). Cells were pretreated with TGF-β, FGF-2, or both and then suspended in a collagen type I gel. After the collagen had solidified, the gel was detached from the sides and bottoms of the dishes and floated in media containing ligands for 48 h. There was no significant degradation of the collagen gel in either the control or FGF-2-treated cells, but the volume of the collagen gel was reduced by ∼60% in cells treated with TGF-β (Figure 2E). More importantly, the cells treated with TGF-β and FGF-2 drastically decreased the volume of collagen gel by ∼30%, and these changes were inhibited by the matrix metalloprotease inhibitor, GM6001. Gelatin zymography further showed that MMP9 activity was enhanced by TGF-β and FGF-2, compared with that by TGF-β alone (Figure 2F). These findings suggest that NMuMG cells treated with TGF-β alone reveal incomplete EMT, and that the cells exposed to TGF-β and FGF-2 exhibit EMT and more aggressive characteristics that resemble those of activated fibroblasts. Myofibroblastic differentiation by prolonged TGF-β treatment To biochemically discriminate between cells that were continuously exposed to TGF-β alone and those exposed to TGF-β and FGF-2, we examined the expression of representative TGF-β-target genes based on our previous study (Kondo et al, 2004). The phosphorylation levels of Smad2 or Smad1/5/8 were not different, and expression levels of some TGF-β-target genes, including Smad7 and fibronectin, were not affected by the addition of FGF-2, indicating that FGF-2 did not affect general TGF-β-Smad signalling (Supplementary Figure S1A and B). Interestingly, the mRNA expression levels of well-known myofibroblast markers, α-SMA and calponin, were increased in TGF-β-treated cells, and α-SMA expression was confirmed in the cells by immunohistochemical analyses (Figure 3A and B), suggesting that TGF-β induced EMyoT. Conversely, the addition of FGF-2 to TGF-β-treated cells markedly decreased the expression of α-SMA and calponin (Figure 3B and C), although the levels of the representative EMT marker, E-cadherin, were repressed by TGF-β and unaffected by addition of FGF-2 (Figure 3C; Supplementary Figure S1C). Inhibition of FGF-2 signalling by SU5402 or short interfering RNA (siRNA)-mediated knockdown of FGFR1IIIc by its specific siRNAs did not obviously alter the expression of E-cadherin regulated by TGF-β (Figure 3C; Supplementary Figure S1D), suggesting that autonomously secreted ligands to FGFR1IIIc exhibit only limited effects on TGF-β-induced EMT. In addition, these findings were not restricted to NMuMG cells, and were also observed in another epithelial cell line α-TN4 (Supplementary Figure S2A and B). These findings indicate that prolonged TGF-β treatment induces the differentiation of epithelial cells into myofibroblastic cells, and that FGF-2 inhibits the TGF-β-mediated EMyoT. Figure 3.Prevention of EMyoT by FGF-2. (A) Immunohistochemical analysis was performed with anti-α-SMA antibody (green) and propidium iodide to detect nuclei (red) in cells treated with 1 ng/ml TGF-β for 3 days and 13 days. (B) Cells were treated with either 1 ng/ml TGF-β or 30 ng/ml FGF-2 alone, or both and then examined for the induction of α-SMA (left) and calponin (right) at the indicated time points by quantitative RT–PCR analysis. Each value represents the mean±s.d. of duplicate determinations from a representative experiment. Similar results were obtained at least three independent experiments. (C) Protein levels of endogenous E-cadherin and α-SMA were analysed by immunoblotting 4 days after treating with combinations of TGF-β, FGF-2, and 10 μM SU5402. α-Tubulin levels were monitored as a loading control for the whole-cell extracts. (D) Phase-contrast micrographs (upper panels) and confocal images after anti-α-SMA antibody staining (green in lower panels) of NMuMG cells. The ligand and inhibitor concentrations were as follows: 1 ng/ml TGF-β, 30 ng/ml FGF-2, 10 μM SU5402, 1 μM PI-103, 1 μM U73122, and 10 μM U0126. Scale bar indicates 100 μm. (E, F) Immunoblot analyses were performed with lysates from NMuMG cells treated with the indicated combinations of ligands and inhibitors. α-Tubulin was used as a loading control (E). The ligands and inhibitors were incubated in culture media containing 10% FBS for 4 days (E), and 2 days (2d in F) or 4 days (4d in F). The same reagent concentrations were used as described in (D). F, FGF-2; Tβ, TGF-β; SU, SU5402; PI, PI-103; U7, U73122; U0, U0126. Download figure Download PowerPoint FGF stimulation leads to tyrosine phosphorylation of FGFR, the recruitment of multiple complexes with adaptor proteins, and activation of the MEK-Erk, phosphoinositide 3 kinase (PI3K) and phosphatidylinositol-specific phospholipase C (PLC) γ pathways. To determine how FGF-2 inhibits TGF-β-mediated EMyoT, we used several inhibitors, including SU5402 for FGFR1, U0126 for MEK 1 and 2, PI-103 for PI3K, and U73122 for PLC. Among these inhibitors, SU5402 and U0126 clearly inhibited the effects of FGF-2 and restored TGF-β-induced α-SMA expression and cell morphology, while neither PI-103 nor U73122 significantly affected these cells (Figure 3D and E). More importantly, TGF-β reduced the levels of Erk phosphorylation, which was accompanied by a reciprocal induction of α-SMA, whereas the addition of FGF-2 returned the Erk phosphorylation levels to baseline (Figure 3F). The cells expressing α-SMA had weaker Erk phosphorylation compared with α-SMA-negative cells, as determined by immunohistochemical analyses (Supplementary Figure S2C). Thus, these findings suggest that Erk inactivation by TGF-β results in α-SMA induction and that restored Erk activation by FGF-2 is required to prevent the induction of α-SMA by TGF-β. Prevention of α-SMA induction by the CtBP1–δEF1 interaction We further examined how FGF-2-induced Erk activation suppressed the induction of α-SMA by TGF-β. Based on a previous report and our previous study (Nishimura et al, 2006; Shirakihara et al, 2007), we focused on the δEF1 family proteins, δEF1/ZEB1 and SIP1/ZEB2. After confirming that specific siRNAs successfully knocked down δEF1 and SIP1 at both the mRNA (Figure 4A) and protein levels (Shirakihara et al, 2007), we found that SIP1 siRNA did not affect the induction of α-SMA (Figure 4B). However, in cells transfected with δEF1 siRNAs, TGF-β-mediated α-SMA upregulation was significantly reduced to a level similar to that obtained with the addition of FGF-2 (Figure 4B and C). More interestingly, FGF-2 did not affect the expression of δEF1 at either the mRNA or protein level (Figure 4A and lower panels in Figure 4E), nor did it affect the expression of microRNA-200b that directly targets δEF1 (Supplementary Figure S2D) (Gregory et al, 2008). These findings indicate that δEF1 is required for the TGF-β-mediated induction of α-SMA and that FGF-2 reduces α-SMA expression without affecting the levels of δEF1 that are upregulated by TGF-β. Thus, the function of δEF1 is regulated by mechanisms other than those involved in regulating transcription or protein stability. Figure 4.Interaction of CtBP1 with δEF1 to prevent α-SMA induction. (A, B) NMuMG cells transfected with either δEF1 siRNA or SIP1 siRNA were stimulated with or without 1 ng/ml TGF-β, 30 ng/ml FGF-2, or both for 24 h and then examined by quantitative RT–PCR analysis for the expression levels of δEF1 (A, left), SIP1 (A, right), and α-SMA (B). NC, control siRNA. Each value represents the mean±s.d. of duplicate determinations from a representative experiment. Similar results were obtained at least three independent experiments. (C) After transfection with different types of siRNAs against δEF1 (δEF1-B and δEF1-C), α-SMA induction by TGF-β was evaluated by immunoblot analysis. α-Tubulin was used as a loading control. The ratio of δEF1 or α-SMA to α-tubulin was validated by densitometric analysis and shown at the lower panels. NC, control siRNA; Tβ, TGF-β; F2, FGF-2. (D) NMuMG cells transfected with control siRNA (NC) or two different types of CtBP1 siRNAs (#1 and #2) were stimulated with TGF-β, FGF-2, or both for 60 h as indicated. After measuring the protein concentrations, the cell lysates were examined by immunoblot analysis. α-Tubulin was used as a loading control. (E) After NMuMG cells were incubated in culture media containing 10% FBS for 4 days with the indicated ligands, cell lysates from ∼1 × 106 cells were examined for protein concentrations and then subjected to immunoprecipitation with normal rabbit IgG (NRS) or an anti-δEF1 antibody. Copurified CtBP1 was detected by immunoblotting with an anti-CtBP1 antibody (top panel). The expression levels of δEF1, CtBP1, and α-tubulin in the same lysates were verified by immunoblotting (lower three panels). The ratio of δEF1 to α-tubulin was validated by densitometric analysis and shown at the bottom. Download figure Download PowerPoint The transcriptional function of δEF1 is regulated through interactions with CtBP1, C-terminal binding protein 1 (Chinnadurai, 2009; Vandewalle et al, 2009). We also found that the inhibitory effect of FGF-2 on TGF-β-induced α-SMA expression was abolished by knocking down CtBP1, resulting in restored α-SMA expression, whereas E-cadherin repression by TGF-β was not affected (Figure 4D). Moreover, endogenous CtBP1 coimmunoprecipitated with endogenous δEF1 (Figure 4E). TGF-β treatment reduced the interaction between CtBP1 and δEF1, while the addition of FGF-2 restored this interaction to a level similar to that in untreated cells (Figure 4E). Inhibition of the MEK-Erk pathway with U0126 and PD98059 decreased this interaction compared with that in the cells treated with TGF-β and FGF-2 (Supplementary Figure S3A). To further define this interaction, we transiently transfected HEK-293T cells with expression plasmids encoding Myc-tagged δEF1 and FLAG-tagged CtBP1. The interaction between δEF1 and CtBP1 was confirmed in overexpressing HEK-293T cells (Supplementary Figure S3B). A CtBP1 mutant that harbours mutations in conserved sites putatively phosphorylated by MAP kinases also interacted with δEF1. This interaction was enhanced by FGF-2 stimulation and repressed by U0126 (Supplementary Figure S3B). In addition, FLAG-CtBP1 that was immunoprecipitated from FGF-2-treated cells exhibited an enhanced interaction with Myc-δEF1 compared with that from non-treated cells in an in vitro binding assay (Supplementary Figure S3C). To examine whether the associ
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