RESEARCH ARTICLE VASCULAR BIOLOGY

Fibroblast growth factor receptor 1 is a key inhibitor of TGFb signaling in the endothelium Pei-Yu Chen,1*† Lingfeng Qin,2* George Tellides,2 Michael Simons1,3† Abnormal vascular homeostasis can lead to increased proliferation of smooth muscle cells and deposition of extracellular matrix, resulting in neointima formation, which contributes to vascular lumen narrowing, a pathology that underlies diseases including transplant vasculopathy, the recurrence of stenosis, and atherosclerosis. Growth of neointima is in part due to endothelial-to-mesenchymal transition (EndMT), a transforming growth factor–b (TGFb)–driven process, which leads to increased numbers of smooth muscle cells and fibroblasts and deposition of extracellular matrix. We reported that endothelial cell–specific knockout of fibroblast growth factor receptor 1 (FGFR1) led to activation of TGFb signaling and development of EndMT in vitro and in vivo. Furthermore, EndMT in human diseased vasculature correlated with decreased abundance of FGFR1. These findings identify FGFR1 as the key regulator of TGFb signaling and EndMT development. INTRODUCTION

Fibroblast growth factors (FGFs) are involved in various cellular functions including proliferation, differentiation, and migration, and FGF signaling is critically important in early development (1, 2). FGF signaling has also been implicated in preserving normal vascular homeostasis (3). This includes maintaining the abundance of vascular endothelial growth factor receptor–2 (VEGFR2) necessary for endothelial cell survival (4), preserving endothelial barrier function by suppressing vascular endothelial cell (VE)– cadherin phosphorylation (5), and preventing endothelial-to-mesenchymal transition (EndMT) through regulation of let-7/TGFb (transforming growth factor b) axis (6). However, the FGFs and FGF receptors (FGFRs) that achieve these effects remain unknown. There are 22 members of the FGF family and 4 high-affinity tyrosine kinase receptors involved in FGF signaling (7, 8). The FGFs mediate their biological effect by binding to one of four FGFRs. FGF1 can activate all four FGFRs and their splice variants, whereas other FGF family members exhibit more selective interactions. Binding of FGFs to their receptors results in phosphorylation of specific intracellular tyrosine residues as well as of an adaptor protein, FGF receptor substrate 2a (FRS2a), that plays a central role in FGF-dependent activation of the MEK [mitogen-activated protein kinase (MAPK) kinase]–ERK1/2 (extracellular signal–regulated kinase 1/2) signaling pathway (9–11), which is critically involved in all three homeostasis maintenance activities of FGF signaling (12). FGFR signaling is complicated by the ability of FGFR family members to form both homo- or heterodimers and the effect of the abundance of various FGFR splice variants on FGF ligand specificity, sensitivity, cellular signaling, and biological functions (13). Despite major progress in the characterization of the FGFR family in different tissues, little is known regarding the role of individual FGFRs in the vasculature. The paucity of data regarding FGFR function in this setting is largely related to very early embryonic lethality in mice with global FGFR1 and FGFR2 deletions and the absence of any vascular phenotype in FGFR3 and FGFR4 knockouts 1 Yale Cardiovascular Research Center, Section of Cardiovascular Medicine, Department of Internal Medicine, Yale University School of Medicine, New Haven, CT 06520, USA. 2Department of Surgery, Yale University School of Medicine, New Haven, CT 06520, USA. 3Department of Cell Biology, Yale University School of Medicine, New Haven, CT 06520, USA. *These authors contributed equally to this work. †Corresponding author. E-mail: [email protected] (M.S.); pei-yu. [email protected] (P.-Y.C.)

(14–18). In addition, endothelial cell–specific deletion of Fgfr1 and Fgfr2 (Fgfr1–/–/Fgfr2–/–) does not affect embryonic vascular development or postnatal vascular growth (19). Mice with endothelial cell–specific deletion of Frs2a show development of EndMT characterized by a profound shape change of normal endothelial cells that express smooth muscle cell markers and deposit collagen-rich extracellular matrix (6). The current study was designed to identify the FGFR in endothelial cells that is responsible for EndMT suppression. We found that FGFR1 was the FGFR responsible for suppressing the activation of TGFb signaling and preventing EndMT in vitro and in vivo in mice. In particular, endothelial FGFR1 deletion led to enhanced neointima formation in graft arteriosclerosis and vein graft adaptation models. Furthermore, we observed the loss of FGFR1 and increased phosphorylation of Smad2 in blood vessels of human coronary arteries from chronic transplant rejection cardiac allografts. Together, these findings demonstrate that FGFR1 is the key FGFR regulating endothelial TGFb signaling that plays an important pathophysiological role in some of the most common cardiovascular diseases.

RESULTS

FGFR1 is responsible for FGF-induced signaling in primary endothelial cells

We first examined the basal expression of mRNAs encoding FGFR family members (FGFR1 to FGFR4) to define their expression profile in endothelial cells. In HUAECs (human umbilical artery endothelial cells), HUVECs (human umbilical vein endothelial cells), and HDLECs (human dermal lymphatic endothelial cells), the mRNA for FGFR1 was the most abundant FGFR-encoding mRNA (Fig. 1A). The mRNAs for FGFR3, FGFR4, and FGF co-receptor bKlotho were also expressed, albeit to a lesser extent. Because the abundance of FGFR2 mRNA was very low, it was not considered further. The abundance of the mRNA encoding FRS2a, the key adaptor molecule involved in FGFR signaling, was similar to that of FGFR1 (Fig. 1A). We next sought to identify the FGFRs that were responsible for FGFmediated signaling in endothelial cells. To this end, FGFR1, FGFR3, and FGFR4 were knocked down using short hairpin RNAs (shRNAs) in HUVECs. Each shRNA was effective in suppressing its target expression at the mRNA and protein levels without affecting the abundance of other FGFRs (Fig. 1B and fig. S1, A, B, D, and E). Knockdown of FGFR1, but

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Fig. 1. FGFR1 relays FGF signaling in primary endothelial cells. (A) Quantitative real-time polymerase chain reaction (qRT-PCR) analysis of FGFRs, FRS2a, and Klotho family gene expression in HUAECs, HUVECs, and HDLECs. Data are presented as means ± SD (*P < 0.001, comparing FGFR2, FGFR3, or FGFR4 to FGFR1). b-Actin was used for sample loading normalization. Histogram of qRT-PCR results is representative of four independent experiments. (B) qRT-PCR analysis of FGFR gene expression in control HUVECs and those with knockdown of FGFRs. Data are presented as means ± SD (*P < 0.001, compared to control). b-Actin was used for sample loading normalization. Histogram of qRT-PCR results is representative of three independent ex-

periments. (C) Phase-contrast and immunofluorescence images of HUVECs with knockdown of FGFRs. DAPI, 4′,6-diamidino-2-phenylindole. Scale bar, 12 mm. Images are representative of three independent experiments. (D) Control or specific shFGFR-treated HUVECs were treated with FGF1 (50 ng/ml) and heparin (100 mg/ml) for the indicated times, and downstream signaling was analyzed by immunoblotting. p-ERK, phosphorylated ERK. Blots are representative of four independent experiments. (E) Cell proliferation was analyzed in control, FGFR1, and FRS2a knockdown HUVECs that were serumstarved overnight and then treated with FGF2 (50 ng/ml). Cell proliferation curves are representative of four independent experiments.

not of FGFR3 or FGFR4, resulted in altered endothelial cell morphology (Fig. 1C and fig. S1, C and F) and was associated with a decrease in FGF1-induced phosphorylation of ERK (Fig. 1D). In response to FGF2 stimulation, the tyrosine phosphorylation of the adaptor protein FRS2a is increased, and its electrophoretic mobility on SDS–polyacrylamide gel electrophoresis gels changes, which has been attributed to serine-threonine phosphorylation (9, 20). This mobility shift is caused by ERK-mediated phosphorylation and can be blocked by MAPK inhibitor U0126 (21). In agreement with this, FGF-induced phosphorylation of FRS2a in FGFR1 knockdown HUVECs was reduced, as demonstrated by a reduction in the mobility shift of FRS2a (Fig. 1D). To verify that knockdown of FGFR1 inhibited FGF signaling and its proangiogenic effects, HUVECs and HDLECs treated with FGFR1 shRNA were stimulated with FGF2. In both cell types, knockdown of FGFR1 inhibited FGF2-induced ERK signaling (fig. S2A), suppressed in vitro cord formation in Matrigel (fig. S2B), and reduced FGF2-driven proliferation of HUVECs (Fig. 1E). These effects were similar in magnitude to the effect of FRS2a knockdown on these processes (Fig. 1E and fig. S2, A and B). The proliferation defects in HUVECs with FGFR1 and FRS2 knockdown may be due to G1 cell cycle arrest and the increased expression of the mRNAs for the cell cycle inhibitors p21 and p27 (fig. S2, C and D).

TGFb2, and TGFbR1, leading to activation of TGFb signaling as demonstrated by increased phosphorylation of Smad2 (Fig. 2, A and B, and fig. S3, A to E). Real-time PCR analysis confirmed FGFR1 knockdown–dependent induction of EndMT in HUVECs characterized by increased abundance of mRNAs encoding smooth muscle markers (SM a-actin, SM22a, SM-calponin, and Notch3) and mesenchymal markers [COL1A1 (collagen 1A1), COL3A1 (collagen 3A1), fibronectin, and vimentin]. In addition, the amount of mRNAs encoding transcription factors Snail, Slug, and Kruppel-like factor 4 (KLF4), which are downstream targets of TGFb and markers for EndMT, was significantly induced by FGFR1 knockdown in HUVECs (Fig. 2C). We previously reported that FGF inhibits TGFb signaling in endothelial cells by increasing the abundance of the microRNA (miRNA) let-7, which targets the mRNA encoding TGFbR1 (6). To investigate whether EndMT in HUVECs induced by FGFR1 knockdown was due to a decrease in let-7 abundance (fig. S3F) and a subsequent increase in TGFbR1 signaling, two different approaches were used. Treatment of HUVECs with FGFR1 knockdown with the TGFbR1 kinase inhibitor SB431542 significantly decreased the abundance of mRNAs encoding SM-calponin, SM a-actin, SM22a, fibronectin, and COL1A1 (Fig. 2D). Overexpression of let-7c in these cells resulted in reduced TGFb signaling and decreased expression of smooth muscle and mesenchymal markers (Fig. 2E).

FGFR1 knockdown increases TGFb-dependent signaling and induces EndMT

Endothelial cells in human patients with chronic cardiac graft rejection lose FGFR1 and activate Smad2

We next examined whether endothelial FGFR1 knockdown was sufficient to increase TGFb signaling and induce EndMT. In HUVECs, knockdown of FGFR1, but not of FGFR3 or FGFR4, led to increased expression of TGFb1,

To study the role of FGFR1 in the regulation of EndMT in disease settings, we next evaluated the abundance of FGFR1 and the phosphorylation of Smad2 by immunocytochemistry in samples of coronary arteries from

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Fig. 2. FGFR1 suppresses TGFbR1-induced mesenchymal marker gene expression. (A) qRT-PCR analysis of TGFb target gene expression in control and FGFR1 knockdown HUVECs (*P < 0.05; **P < 0.01, compared to control). b-Actin was used for sample loading normalization. Histogram of qRT-PCR results is representative of three independent experiments. (B) Immunoblot analysis of TGFb signaling in control and FGFR1 knockdown HUVECs. Blots are representative of three independent experiments. p-Smad2, phosphorylated Smad2. (C) qRT-PCR analysis of gene expression for smooth muscle cell (SMC) markers, mesenchymal markers, and transcription factors in control and FGFR1 knockdown HUVECs (*P < 0.05; **P < 0.01; ***P < 0.001, compared to control). b-Actin was used for sample loading normalization. Histograms of qRT-PCR results are representative of three independent experiments. COL1A1, collagen 1A1; COL3A1, col-

lagen 3A1. (D) Left: Immunoblots of TGFbR1 and Smad2 in control and FGFR1 knockdown HUVECs with or without SB431542 (SB) treatment. Blots are representative of three independent experiments. Right: qRT-PCR analysis of gene expression of smooth muscle cell and mesenchymal markers (*P < 0.05; **P < 0.01, compared to control). b-Actin was used for sample loading normalization. Histogram of qRT-PCR results is representative of three independent experiments. (E) Left: Immunoblots of TGFbR1 and Smad2 in control and FGFR1 knockdown HUVECs with or without let-7c overexpression. Blots are representative of three independent experiments. Right: qRT-PCR analysis of gene expression of smooth muscle cell and mesenchymal markers (*P < 0.05; **P < 0.01, compared to control). b-Actin was used for sample loading normalization. Histogram of qRT-PCR results are representative of three independent experiments.

healthy individuals or from explanted rejected cardiac allografts (fig. S4, A and B). Whereas healthy human coronaries demonstrated extensive FGFR1 staining in the endothelium (Fig. 3A), FGFR1 staining was lost in ~80 to 90% of luminal and neointima endothelial cells in arteries from patients with chronic cardiac graft rejection (Fig. 3, B and C). This loss of FGFR1 staining was also associated with increased phosphorylation of Smad2 (Fig. 3, D to F). To further study the effect of allograft rejection on FGFR1 abundance, we used a human-mouse chimeric model (22), in which segments of normal human coronary arteries from fresh surgical specimens were implanted into the infrarenal aortae of severe combined immunodeficient (SCID)/beige mice, which were then sensitized a week later by an injection of human allogeneic peripheral blood mononuclear cells (PBMCs). Intimal expansion was greater in PBMC-injected grafts than in uninjected grafts (fig. S4, C and D). Quantitative analysis demonstrated that although FGFR1 was present in most endothelial cells in uninjected human grafts (Fig. 4, A and B), FGFR1 was present in only 5% of luminal and 10% of neointimal endothelial cells after PBMC injection and the onset of rejection (Fig. 4, C to E). In addition, the phosphorylation of Smad2 in

CD31+ endothelial cells was significantly higher in PBMC-injected human vessels than in uninjected vessels (Fig. 4, F to J).

Endothelial cell–specific deletion of Fgfr1 enhances EndMT in vivo To further corroborate that the loss of endothelial FGFR1 leads to the onset of EndMT in vivo, we generated mice with an endothelial cell–specific Fgfr1 deficiency (Fgfr1ECKO) using the Cdh5-CreERT2 line (fig. S5A). Postnatal Cre activation starting at P1 resulted in highly efficient deletion of the target allele (fig. S5, B and C). In agreement with a previous study (19), Fgfr1ECKO mice appeared normal. In particular, there was no delay in the formation of retinal vasculature in P6 pups and adults (fig. S5, D to G) or the density or appearance of the blood vasculature in various organs (fig. S5H). Interposition of a segment of a male donor (BALBc/J strain) thoracic aorta into the abdominal aorta of a male recipient (C57BL/6 strain) led to the development of extensive neointima formation in the graft after 2 weeks (fig. S6, A and B). To study the contribution of EndMT to this process, we used endothelial fate–mapped mice that were generated by crossing Cdh5-CreER2;Fgfr1flox/flox and mT/mG mouse lines (fig. S5A). mT/mG is a double-fluorescent reporter

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RESEARCH ARTICLE Fig. 3. FGFR1 protein abundance is deA creased and phosphorylation of Smad2 is increased in endothelial cells in coronary arteries from chronically rejecting human heart allografts. (A) Representative images of immunofluorescence staining for CD31 (green) and FGFR1 (red) in healthy human coronary arteries. L, lumen. Nuclei were stained with DAPI (blue). Scale bar, 62 mm. a, high-power images of endothelial cells in the lumen of a healthy human coronary arC tery. Scale bar, 16 mm. Images are representative of three normal human coronary artery samples. (B) Representative images of immunofluorescence staining for CD31 (green) and FGFR1 (red) in a coronary artery from a chronically rejecting human heart allograft. White triangle, internal elasD tic lamina (IEL). Nuclei were stained with DAPI (blue). Scale bar, 62 mm. b and c, high-power images of endothelial cells in the lumen (b) and neointima (c) of a coronary artery from a chronically rejecting allograft artery. White arrows, endothelial cells without FGFR1 staining. Yellow arrowhead, endothelial cell with FGFR1 staining. Scale bar, 16 mm. Images are representative of eight coronary arteries from patients with chronically rejecting heart allografts for luF minal endothelial cells (ECs), six of which had neointima ECs that were positive for the human endothelial cell marker CD31. Loss of EC markers may occur during disease. (C) Percentage of FGFR1+CD31+ cells in the lumen or in neointima (***P < 0.001, compared to normal artery; Ø, not detected). Three normal human coronary arteries and eight coronary arteries from patients with chronically rejecting heart allografts were evaluated. (D) Representative images of immunofluorescence staining for CD31 (green) and p-Smad2 (red) in healthy human coronary arteries. Nuclei were stained with DAPI (blue). Scale bar, 62 mm. d, high-power images of endothelial cells in the lumen of a healthy human coronary artery. Scale bar, 16 mm. Images are representative of three normal human coronary artery samples. (E) Representative images of immunofluorescence staining for CD31 (green) and phosphorylated Smad2 (red) in a coronary artery from a chronically rejecting human allograft. White triangle, internal elastic lamina. Nuclei were stained with DAPI (blue). Scale bar, 62 mm. e and f,

high-power images of endothelial cells in the lumen (e) or neointima (f) of a chronically rejecting human coronary artery. White arrows, endothelial cells with phosphorylated Smad2. Scale bar, 16 mm. Images are representative of eight human coronary arteries from patients with chronically rejecting heart allograft samples for luminal ECs, six of which had neointima ECs that were positive for the human endothelial cell marker CD31. Loss of EC markers may occur during disease. (F) Percentage of p-Smad2+CD31+ cells in the lumen or neointima (***P < 0.001, compared to normal artery; Ø, not detected). Three normal human coronary arteries and eight human coronary arteries from patients with chronically rejecting heart allografts were evaluated.

mouse that expresses membrane-targeted tandem dimer Tomato (mT) before Cre-mediated excision and membrane-targeted green fluorescent protein (GFP) (mG) after excision (23). Thus, Cdh5-GFP–positive cells after Cre excision serve as an indicator not only of endothelial cells but also of FGFR1 deletion. Transplantation of an aortic segment into Fgfr1ECKO mice resulted in a marked increase in the number of luminal and neointimal Cdh5-GFP– positive cells with smooth muscle cell markers compared to the control (Fig. 5, A to D) and a significant increase in artery graft neointima thickness and collagen 1 deposition (Fig. 5E). These Cdh5-GFP–positive endothelial cells were negative for MYH11 (a marker of mature smooth

muscle cells), FSP1 (a fibroblast marker), and NG2 (a pericyte marker) (fig. S6C). A different set of control and Fgfr1ECKO mice was used for studies using the vein graft adaptation model, in which a segment of the intrathoracic inferior vena cava from C57BL/6 male mouse was interposed into the infrarenal aorta of a genetically matched wild-type or Fgfr1ECKO male mouse (fig. S6D). Similar to arterial grafting, vein grafting in an arterial position also led to neointima formation (fig. S6E). The loss of FGFR1 in endothelial cells led to a significant increase in the neointima thickness of vein grafts, collagen 1 deposition, and the numbers of luminal and neointimal Cdh5-GFP–positive cells with smooth muscle cell markers compared to the control (Fig. 5, F to J).

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Fig. 4. FGFR1 protein abundance is decreased and phosphorylation of Smad2 is increased in a human-mouse chimera artery transplant model. (A) Representative images of immunofluorescence staining for CD31 (green) and FGFR1 (red) in a human coronary artery that was not injected with human PBMCs before transplantation into SCID/beige mouse (control). L, lumen. Nuclei were stained with DAPI (blue). Scale bar, 62 mm. (B) High-power images of the area indicated by the white box in (A). Scale bar, 16 mm. Images are representative of five injected human grafts. (C) Representative images of immunofluorescence staining for CD31 (green) and FGFR1 (red) in a human coronary artery injected with human PBMCs before transplantation into SCID/beige mouse. Nuclei were stained with DAPI (blue). Scale bar, 62 mm. Images are representative of 10 human grafts injected with PBMCs. (D) High-power images of the area indicated by the white box in (C). Note the loss of FGFR1 staining. White triangle, internal elastic lamina (IEL); white arrows, endothelial cells without FGFR1. Scale bar, 16 mm. (E) Percentage of CD31+FGFR1+endothelial cells in the lumen (left) and neointima (right) in PBMC-injected and uninjected mice (Ø, not detected; ***P < 0.001, compared to no PBMCs). Five uninjected human grafts and 10 human grafts injected with PBMCs were evaluated. (F)

Representative images of immunofluorescence staining for CD31 (green) and phosphorylated Smad2 (red) in a human coronary artery transplanted into SCID/beige mouse not injected with human PBMCs. Nuclei were stained with DAPI (blue). Scale bar, 62 mm. (G) High-power images of the area indicated by the white square in (F). White arrows, endothelial cells expressing phosphorylated Smad2. Scale bar, 16 mm. Images are representative of five uninjected human grafts. (H) Representative images of immunofluorescence staining for CD31 (green) and phosphorylated Smad2 (red) in a human coronary artery transplanted into SCID/beige mouse injected with human PBMCs. Nuclei were stained with DAPI (blue). Scale bar, 62 mm. (I) High-power images of the area indicated by the white box in (H). Note increased phosphorylated Smad2 staining. White triangle, internal elastic lamina; arrows, endothelial cells with phosphorylated Smad2. Yellow arrowhead, endothelial cell negative for phosphorylated Smad2. Scale bar, 16 mm. Images are representative of 10 human grafts injected with PBMCs. (J) Percentage of CD31+, p-Smad2+ endothelial cells in lumen (left) and neointima (right) (**P < 0.01, compared to no PBMC; Ø, not detected). Five uninjected human grafts and 10 human grafts injected with PBMCs were evaluated.

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Fig. 5. Fgfr1ECKO knockout mice: EndMT in arteriosclerosis and vein graft adaptation models. (A) Histological analysis of artery with anti–a-SMA (a–smooth muscle actin), anti-Notch3, and anti–collagen 1 antibodies. Colocalization (yellow) of Cdh5-GFP+ (green) with Notch3 (red) or a-SMA (red) staining indicates EndMT. Nuclei were counterstained with DAPI (blue). L, lumen; N, neointima; white triangle, endothelial cells; yellow triangle, internal elastic lamina (IEL); white arrows, endothelial cells expressing smooth muscle cell markers. Scale bar, 10 mm (Notch3 and a-SMA); 63 mm (collagen 1). (B) Percentage of Notch3-positive cells with Cdh5-GFP fluorescence (left) or a-SMA staining (right) cells in the neointima (*P < 0.05, compared to control). (C) Percentage of Cdh5-GFP–positive cells with Notch3 (left) or a-SMA (right) staining in the neointima. (D) Percentage of Cdh5-GFP–positive cells with Notch3 (left) or a-SMA (right) staining in lumen (*P < 0.05, compared to control). (E) Morphometric assessment of artery graft neointima thickness and collagen-1 deposition performed by

computer-assisted microscopy (*P < 0.05, compared to control). n = 4 mice per group were evaluated in (A) to (E). (F) Histological analysis with anti–a-SMA, anti-Notch3, and anti–collagen 1 of vein grafts transplanted into an arterial position. Colocalization (yellow) of Cdh5-GFP (green) with Notch3 (red) or a-SMA (red) staining indicates EndMT. Nuclei were counterstained with DAPI (blue). Symbols are the same as in (A). Scale bar, 10 mm (Notch3 and a-SMA); 63 mm (collagen 1). (G) Percentage of Notch3-positive (left) or a-SMA–positive (right) cells that have Cdh5-GFP fluorescence in neointima (*P < 0.05, compared to control). (H) Percentage of Cdh5-GFP– positive cells that have Notch3 (left) or a-SMA (right) staining in neointima. (I) Percentage of Cdh5-GFP–positive cells that have Notch3 (left) or a-SMA (right) staining in the lumen (*P < 0.05, compared to control). (J) Morphometric assessment of artery graft neointima thickness and collagen 1 deposition performed by computer-assisted microscopy (*P < 0.05; **P < 0.01, compared to control). n = 6 mice per group were evaluated in (F) to (J).

To determine the role of FGFR1 in vascular regeneration, we used a mouse hindlimb ischemia model (24). FGFR1 abundance in the vasculature of ischemic limbs was not altered by induction of ischemia as determined by immunohistochemistry with anti-FGFR1 and anti-CD31 antibodies (Fig. 6, A and B). When compared to control mice, mice carrying endothelial Fgfr1 deletion demonstrated a significant but only moderately reduced blood flow restoration in the ischemic limb (Fig. 6, C and D) and decreased capillary density (Fig. 6, E and F).

ture. Loss of FGFR1 expression in vitro or in vivo is associated with increased TGFb signaling, activation of Smad2, and the appearance of various mesenchymal markers. In vivo, the loss of FGFR1 is associated with the presence of inflammation. This is demonstrated by a nearly complete loss of the receptor in a mouse strain-mismatched vascular transplant model and in human vascular tissues from rejecting cardiac allografts, settings associated with intense inflammation. At the same time, FGFR1 abundance was not altered in the hindlimb ischemia model that is characterized by ischemia but not an intense inflammatory response. As a consequence, there was only a mild impairment of perfusion recovery. These findings are consistent with our previous report of control of FGFR1 expression by inflammatory and immune response mediators such as interferon-g, tumor necrosis factor a, and interleukin-1b (6). This important

DISCUSSION

The evidence presented in this study points to the endothelial FGFR1 as the principal FGFR that confers protection from EndMT to the vascula-

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Fig. 6. Fgfr1ECKO knockout mice: mouse hindlimb ischemic model. (A) Representative images of H&E (hematoxylin and eosin) and FGFR1 (red) and CD31 (green) immunofluorescence staining in fresh-frozen sections of nonischemic and ischemic lower limb at days 0, 3, and 7 after ligation. Images are representative of three mice per time point. (B) Left: Percentage of CD31-positive cells that also have FGFR1 staining in nonischemic and ischemic muscles at days 0, 3, and 7. Right: Quantification of capillary density (number/mm2 muscle area) in (A). NS, not significant compared to nonischemic muscle. Three mice per time point were evaluated. (C) Laser Doppler images showing blood flow before and after the induction of ische-

mia in control and Fgfr1ECKO mice. Images are representative of three mice per time point. (D) Laser Doppler analysis of blood flow recovery in the left foot, calculated as the ratio of blood flow in left to right foot (L/R). *P < 0.05, control compared to Fgfr1ECKO. Three mice per time point were evaluated. (E) Representative sections from nonischemic and ischemic groups of control and Fgfr1ECKO on day 14 after ischemia. Images are representative of three mice at day 14. (F) Quantification of vascular density (number/mm2 muscle area). Data are means ± SD from 10 fields per section (3 sections per mouse). *P < 0.05, control compared to Fgfr1ECKO. Three mice per group were evaluated.

biological role of FGFR1 is further supported by the enhancement of EndMT and neointima formation in mouse graft arteriosclerosis and vein-to-artery grafting models by endothelial cell–specific deletion of FGFR1. FGFs comprise a large family of 22 proteins that are involved in the regulation of various biological processes including cell proliferation, migration, survival, and differentiation (1, 25). These effects are mediated at the cellular level by four FGF receptor tyrosine kinases and several nontyrosine kinase receptors including Klotho and syndecan family members (26–28). FGFRs are widely distributed in many tissues and organs, and variations in their distribution or abundance may lead to functional differences essential for the coordinate regulation of tissue homeostasis and orchestration of complex processes such as angiogenesis or tissue repair. Aberrant FGFR signaling, whether enhanced or reduced, has been implicated in human pathologies including developmental and genetic diseases, metabolic disorders, and cancer (29). Little is known about the abundance and biological and pathological functions of various FGFRs in the vasculature. Several studies have demonstrated that a disruption of FGF signaling input to the vasculature, achieved either by expression of a soluble pan-FGF trap or by endothelial expression of a dominant-negative construct capable of blocking all FGFR signaling, results in a profound decline in VEGFR2 expression, increased permeability, and, eventually, loss of vascular integrity (3, 4). We have reported that a disruption of FGF signaling in blood endothelium achieved by a knockout of a pan-FGFR adaptor, Frs2a, results in a steep decline in let-7 miRNA expression, which in turn induces TGFb signaling and the development of EndMT (6).

EndMT has emerged as a critical pathological process that occurs in various disease settings including myocardial ischemia, cancer, and liver injury. Moreover, it has been causally implicated in the development of cardiac fibrosis (30), pulmonary hypertension (31), neointima formation (6, 32), and cavernous cerebral malformations (33). Here, we showed that FGFR1 is the key FGFR that prevents EndMT and maintains vascular normalcy. FGFR1 was highly abundant in the luminal endothelium of nondiseased human coronary arteries and in the mouse vasculature, whereas its abundance was decreased in the endothelium of coronary arteries from rejecting human hearts or from human arteries implanted into mice. Morphologically, vasculature in rejecting organ grafts had increased phosphorylation of Smad2, consistent with activated TGFb signaling, and extensive neointima formation and fibrosis, all of which are hallmarks of EndMT. This adverse role of TGFb signaling in the vasculature is consistent with its role in preventing endothelial cell differentiation and interfering with VEGFR2 signaling (34). Furthermore, we and others have demonstrated that suppression of TGFb signaling suppresses neointima formation in vein graft adaptation and tissue-engineered vascular graft models (35, 36). The loss of FGFR1 and the development of EndMT demonstrate an important regulatory role for FGFR1 in these settings. The direct evidence of the FGFR1 loss–dependent induction of EndMT comes from studies in Fgfr1ECKO mice, using two different EndMT models: graft arteriosclerosis and vein-to-artery grafting models. Histologic examination of vascular grafts of mice in these models revealed that deletion of

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RESEARCH ARTICLE the Fgfr1 resulted in a significant increase in EndMT, collagen deposition, and neointima formation, compared to control mice. In summary, this study demonstrates that FGFR1 is the key FGFR in endothelial cells that regulates TGFb signaling. Decreased abundance of FGFR1 leads to EndMT and the development of extensive vascular pathology. In human tissues, loss of endothelial FGFR1 is also associated with enhanced TGFb signaling and EndMT. These findings identify the FGFR1TGFb pathway as an important contributor to vascular pathologies and as a potential target for therapeutic interventions.

MATERIALS AND METHODS

Growth factors and chemicals Recombinant human FGF1 (100-17A, PeproTech), FGF2 (233-FB-001MG/ CF, R&D), and TGFb1 (580702, BioLegend) were reconstituted in 0.1% bovine serum albumin (BSA)/phosphate-buffered saline (PBS). The TGFbR1 kinase inhibitor SB431542 (S4317, Sigma) was reconstituted in dimethyl sulfoxide (D2650, Sigma) and used at a final concentration of 10 mM in cell culture.

Antibodies used for immunodetection of proteins We used the following antibodies for immunoblotting (IB), immunofluorescence (IF), or immunohistochemistry (IHC): CD31 (sc-1506, Santa Cruz; IB), CD31 (sc-1506, Santa Cruz; IHC), CD31 (M0823, Dako; IHC), CD31 (553370, BD Pharmingen; primary mouse endothelial cell isolation), collagen 1 (NB600-408, Novus Biologicals; IHC), phosphorylated ERK (M8159, Sigma; IB), FGFR1 (#2144-1, Epitomics; IB), FGFR1 (FB817; IHC for human paraffin samples and for mouse fresh muscle tissue samples) (37), FGFR1 [sc-121, Santa Cruz; IHC for mouse 4% paraformaldehyde (PFA)–fixed optimum cutting temperature compound (OCT) samples], FGFR1 (5G11; IB) (37), FGFR3 (sc-9007, Santa Cruz; IB), FGFR4 (#8562, Cell Signaling; IB), FRS2a (sc-8318, Santa Cruz; IB), FSP1 (ab27957, Abcam; IHC), GAPDH (glyceraldehyde phosphate dehydrogenase) (#2118, Cell Signaling; IB), HMGA2 (#59170AP, BioCheck; IB), p44/p42 MAP Kinase (#9102, Cell Signaling; IB), MYH11 (M7786, Sigma; IHC), NG2 (AB5320, Millipore; IHC), Notch3 (Ab23426, Abcam; IHC), p21 (#2947, Cell Signaling; IB), p27 (#3688, Cell Signaling; IB), Smad2 (#3122, Cell Signaling; IB), phosphorylated Smad2 (Ser465/467) (#3108, Cell Signaling; IB), phosphorylated Smad2 (Ser465/467) (#3101, Cell Signaling; IHC for human paraffin samples), smooth muscle a-actin–APC (allophycocyanin) (IC1420A, R&D; IHC), TGFbR1 (sc-398, Santa Cruz; IB), b-tubulin (T7816, Sigma; IB), VE-cadherin (C-19, Santa Cruz; IB, IF), and VEGFR2 (#2479, Cell Signaling; IB).

Cell culture and reagents Human 293T T17 cells (human embryonic kidney cells, ATCC CRL11268) were maintained in Dulbecco’s modified Eagle’s medium (Gibco) with 10% fetal bovine serum (FBS) (Invitrogen) and penicillin-streptomycin (Gibco). HUAECs (passages 5 to 10, Lonza), HUVECs (passages 5 to 10, Lonza), and HDLECs (passages 5 to 10, Lonza) were cultured in endothelial basal medium–2 (EBM-2) supplemented with EGM-2MV BulletKit (CC-3202, Lonza). Culture vessels were coated with 0.1% gelatin (G6144, Sigma) for 30 min at 37°C immediately before cell seeding. Primary mouse endothelial cells were isolated from the lung, using rat anti-mouse CD31 antibody (clone MEC13.3, Pharmingen, #553370) and Dynabeads (cat. no. 110.35, Invitrogen) as previously described (6).

Generation of lentivirus constructs

Bam H1–cleaved complementary DNA (cDNA) fragments encoding Xenopus kinase-dead (KD) FGFR1 were inserted into the lentivirus pLVX-IRES-Puro

vector. The constructs were verified by sequence analysis (Yale Keck DNA sequencing core facility), and protein expression was confirmed by immunoblot analysis.

Generation of lentiviruses FGFR1, FGFR3, and FGFR4 shRNA lentiviral constructs were purchased from Sigma, and FRS2a shRNA lentiviral construct was purchased from Open Biosystems. For the production of shRNA lentivirus, 3.7 mg of D8.2, 0.2 mg of vesicular stomatitis virus glycoprotein, and 2.1 mg of pLKO.1 carrying the control, FGFR1, FGFR3, FGFR4, or FRS2a shRNA were cotransfected into 293T cells, using FuGENE 6. Forty-eight hours later, the medium was harvested, cleared by 0.45-mm filter, mixed with polybrene (5 mg/ml) (H9268, Sigma), and applied to the cells. After a 6-hour incubation, the virus-containing medium was replaced by fresh medium. For the production of FGFR1 KD or miRNA lentivirus, 10 mg of pLVX-IRES carrying the FGFR1 KD or 10 mg of pMIRNA1 carrying the let-7 miRNA expression cassette (System Biosciences), 5 mg of pMDLg/PRRE, 2.5 mg of RSV-REV (Rous sarcoma virus– regulator of expression of virion proteins), and 3 mg of pMD.2G were cotransfected into 293T cells, using FuGENE 6. Forty-eight hours later, the medium was harvested, cleared by 0.45-mm filter, mixed with polybrene (5 mg/ml) (H9268, Sigma), and applied to the cells. After a 6-hour incubation, the virus-containing medium was replaced by fresh medium.

Western blot analysis Cells were lysed with T-PER (Thermo Scientific) containing complete mini EDTA-free protease inhibitors (#11836170001, Roche) and phosphatase inhibitors (#04906837001, Roche). Twenty micrograms of total protein from each sample was resolved on a 4 to 12% bis-tris gel (Bio-Rad) with MOPs Running Buffer (Bio-Rad) and transferred onto nitrocellulose membranes (#162-0094, Bio-Rad). The blots were then probed with various antibodies. Chemiluminescence measurements were performed using SuperSignal West Femto Maximum Sensitivity Substrate (Thermo Fisher Scientific).

RNA isolation and qRT-PCR Cells were suspended in TRIzol reagent (#15596018, Invitrogen), and total RNA (#74134, Qiagen) and miRNA-enriched fraction (#74204, Qiagen) were isolated according to the manufacturer’s instructions. Reverse transcriptions were performed by using iScriptcDNA synthesis kit (170-8891, Bio-Rad) for mRNA or RT2 miRNA First Strand Kit (331401, Qiagen) for miRNA. qRT-PCR was performed using Bio-Rad CFX94 (Bio-Rad) by mixing equal amounts of cDNAs, iQ SYBR Green Supermix (170-8882, Bio-Rad), and gene-specific primers (Qiagen). All reactions were done in a 25-ml reaction volume in duplicate. Individual mRNA or miRNA expression was normalized in relation to expression of endogenous b-actin or small nuclear SNORD47, respectively. PCR amplification consisted of 10 min of an initial denaturation step at 95°C, followed by 46 cycles of PCR at 95°C for 15 s and 60°C for 30 s (for mRNA cDNA) and 10 min of an initial denaturation step at 95°C, followed by 46 cycles of PCR at 95°C for 15 s, 55°C for 30 s, and 70°C for 30 s (for mRNA cDNA).

Immunofluorescence staining

Cultured primary endothelial cells were grown on fibronectin (10 mg/ml, F2006, Sigma)–coated glass-bottomed dishes (P35G-1.5-20-C, MatTek Corporation) 24 hours before fixation. The next day, cells were first fixed with 2% PFA (18814, Polysciences Inc.) in PBS for 20 min at 37°C, then permeabilized with 0.1% Triton X-100 in PBS containing 2% PFA at room temperature for 5 min, and blocked with 3% BSA (001-000-162, Jackson ImmunoResearch Laboratories Inc.) at room temperature for 60 min. Cells were washed with PBS and incubated with VE-cadherin antibody (1:100 in 1% BSA) at 4°C overnight, washed three times with

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RESEARCH ARTICLE PBS, and incubated with diluted Alexa Fluor–conjugated secondary antibody (1:500) (Invitrogen) for 1 hour at room temperature. The dishes were then washed three times with PBS and mounted using ProLong Gold antifade reagent with DAPI (P36935, Life Technologies).

Matrigel cord formation assay Twenty-four–well plates were coated with growth factor–reduced Matrigel (354230, BD Biosciences) and allowed to solidify for 30 min at 37°C. Cells were trypsinized, seeded on Matrigel-coated plates at a density of 6 × 104 cells per well in EBM-2 medium with 0.5% FBS with and without FGF2 (50 ng/ml), and incubated for 6 hours at 37°C. Photomicrographs were taken at ×100 magnification.

of 10- to 12-week-old male recipient Fgfr1ECKO mice, using an end-to-end microsurgical anastomotic technique.

Hindlimb ischemia model and laser Doppler blood flow analysis Mouse hindlimb ischemia was induced as previously described (24). For the laser Doppler blood flow analysis, hindlimb blood flow was measured on preoperative days and postoperative days 0, 3, 7, and 14 using a laser Doppler blood flow analyzer (moorLDI, Moor Instruments). Blood flow was quantitatively assessed as changes in the laser images, using different color pixels. To avoid mouse-to-mouse and experiment-to-experiment variations, hindlimb blood flow was expressed as the ratio of left (ischemic) to right (nonischemic) laser Doppler signal.

Cell cycle analysis Cell cycle analysis was performed using propidium iodide (PI) staining and flow cytometry. Cells were trypsinized, washed twice in PBS, and fixed in 70% ethanol at –20°C overnight. After being washed twice in PBS, the cells were treated with RNase A (100 mg/ml) (R5125, Sigma) at 37°C for 30 min and stained in PI solution (50 mg/ml) (P4170, Sigma). Then, the cells were transferred to flow cytometry tubes with filters (#352235, BD) for cell cycle analysis. For each sample, 10,000 events were collected. The data were collected and analyzed with ModFit software.

Morphometric analysis To harvest transplanted grafts, animals were anesthetized, and tissues were perfused with normal saline and 4% PFA. For cryosection preparation, grafts were isolated from anesthetized mice, fixed in 4% PFA overnight at 4°C, cryoprotected for 4 hours in 15% sucrose at 4°C, and embedded in OCT (Tissue-Tek). Graft area measurements of the intima (between the endothelium and internal elastic lamina) were calculated from five serial cross sections, 150 mm apart for each graft, using computer-assisted image analysis and National Institutes of Health (NIH) Image 1.60.

Generation of mice

Fgfr1flox/flox mice have been previously described (38). Fgfr1flox/flox mice were crossed to Rosa26 fluorescent reporter mT/mG reporter line (JAX SN:007676) and then bred with mice expressing Cre recombinase under the Cdh5 promoter (39). PCR genotyping analysis was done using the following primers: Fgfr1flox/flox (5′-CTGGTATCCTGTGCCTATC-3′ and 5′-CAATCTGATCCCAAGACCAC-3′), mT/mG (5′-CTCTGCTGCCTCCTGGCTTCT-3′, 5′-CGAGGCGGATCACAAGCAATA-3′, and 5′-TCAATGGGCGGGGGTCGTT-3′), and Cdh5-CreERT2 (5′-GCCTGCATTACCGGTCGATGCAACGA-3′ and 5′-GTGGCAGATGGCGCGGCAACACCATT-3′).

Human tissue De-identified human coronary arteries were obtained from organ donors with normal cardiac function, from end-stage cardiomyopathy patients undergoing heart transplantation, or from cardiac allograft recipients with chronic rejection at autopsy or undergoing retransplantation. Tissue collection and analysis were approved by the review boards of Yale University School of Medicine, the New England Organ Bank, and the University of British Columbia.

Animal models for examining EndMT in vivo All experiments were performed under protocols approved by Yale University Institutional Animal Care and Use and Human Investigation Committees. For the human-to-mouse arterial transplant model, adjacent segments of human coronary artery from surgical specimens or cadaveric organ donors were implanted into the infrarenal aortae of 10-week-old CB.17 SCID/beige mice. Each mouse received 1 × 108 human allogeneic PBMCs, injected intraperitoneally into an artery 1 week after transplantation. Animals were euthanized 4 weeks after coronary artery transplantation, and transplanted arteries were removed and frozen in OCT for further analysis. For the mouse arteriosclerosis model, segments of thoracic aorta from 4- to 5-week-old male BALBc/J mice were interposed into the infrarenal aortae of 10- to 12-week-old male recipient Fgfr1ECKO mice, using an end-to-end microsurgical anastomotic technique. For the mouse vein graft adaptation model, segments of intrathoracic inferior vena cava from 4- to 5-week-old male C57BL/6 mice were interposed into the infrarenal aortae

Immunohistochemical staining Frozen blocks were sectioned at 5-mm intervals using a Microm cryostat. For frozen-tissue sections, slides were fixed in acetone for 10 min at –20°C. For paraffin sections, slides were dewaxed in xylene, boiled for 20 min in citrate buffer (10 mM, pH 6.0) for antigen retrieval, and rehydrated. After being washed three times with PBS, tissue sections were incubated with primary antibodies diluted in blocking solution (10% BSA and horse serum in PBS) overnight at 4°C in a humidified chamber. Sections were washed three times with tris-buffered saline, incubated with appropriate Alexa Fluor 488–, Alexa Fluor 594–, or Alexa Fluor 633–conjugated secondary antibodies diluted 1:500 in blocking solution for 1 hour at room temperature, washed again three times, and mounted on slides with ProLong Gold mounting reagent with DAPI (P36935, Life Technologies). All immunofluorescence micrographs were acquired using a Zeiss microscope. Images were captured using Velocity software, and quantifications were performed using ImageJ (NIH).

Computer-assisted image analysis (quantitative immunofluorescence) Collagen-positive areas were measured using NIH Image 1.60. Neointima thickness was measured from four regions of a section along a cross, and the average was calculated from five serial cross-sections, 150 mm apart, for each graft, using computer-assisted image analysis and NIH Image 1.60. For evaluation of a-SMA+/Cdh5-GFP+ or Notch3+/Cdh5-GFP+ cells, at least 10 sections 150 mm apart per graft were analyzed for colocation of endothelial and smooth muscle markers, using immunofluorescence staining imaged under ×100 magnification.

Statistical analysis Statistical analysis was performed using GraphPad Prism software v.5. All data are presented as means ± SD, and two group comparisons were done with a twotailed Student’s t test. A value of P < 0.05 was taken as statistically significant. SUPPLEMENTARY MATERIALS www.sciencesignaling.org/cgi/content/full/7/344/ra90/DC1 Fig. S1. Efficient FGFR shRNA knockdown in HUAECs and HDLECs. Fig. S2. Effects of FGFR knockdown on FGF2 signaling in vitro.

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RESEARCH ARTICLE Fig. S3. Effects of FGFR knockdown on TGFb1 signaling in vitro. Fig. S4. Increased intimal thickening in human coronary artery from chronically rejecting heart allografts and in human-mouse chimera graft rejection arteries. Fig. S5. Mice with endothelial cell–specific knockout of Fgfr1 have normal baseline vascular density. Fig. S6. Schematic of the mouse arteriosclerosis and mouse vein graft adaptation models.

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Adams, Inducible gene targeting in the neonatal vasculature and analysis of retinal angiogenesis in mice. Nat. Protoc. 5, 1518–1534 (2010). Acknowledgments: We thank R. Friesel (Maine Medical Center Research Institute) for providing FGFR1 constructs and FGFR1 antibodies, R. Adams (Max Planck Institute, Munster) for the Cdh5-CreERT2 mouse, and B. McManus (University of British Columbia) for human coronary artery specimens. We are grateful to R. Webber, N. Copeland, and W. Evangelisti for maintaining the mice used in these studies. Funding: This work was supported by NIH grant R01 HL053793 (to M.S.). Author contributions: P.-Y.C., L.Q., G.T., and M.S. designed the research; P.-Y.C. and L.Q. performed the research; P.-Y.C., L.Q., G.T., and M.S. analyzed the data; and P.-Y.C. and M.S. wrote the manuscript. Competing interests: The authors declare that they have no competing interests. Data and materials availability: The FGFR1 floxed mice require a materials transfer agreement from C.-X. Deng (NIH). Submitted 20 May 2014 Accepted 5 September 2014 Final Publication 23 September 2014 10.1126/scisignal.2005504 Citation: P.-Y. Chen, L. Qin, G. Tellides, M. Simons, Fibroblast growth factor receptor 1 is a key inhibitor of TGFb signaling in the endothelium. Sci. Signal. 7, ra90 (2014).

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