Fig 2: Decreased ALK1 expression is associated with reduced NOTCH activity and NOTCH target gene expression in the colonic epithelial cells of CD patients. (A) Representative blot (left) and the difference of JAG1 and NOTCH intracellular domain (NICD) protein expression between NIBD and CD patients (right). (B) BMP9 concentration in the serum of NIBD controls (N = 17) and CD patients (N = 23). (C) NOTCH target gene expression in colonic epithelial cells from CD patients (N = 15) and NIBD controls (N = 12). (D) NOTCH target gene expression in NIBD patient-derived colonic epithelial cell monolayers. Expanded cells were cultured in expansion media in the presence or absence of BMP9 and ALK1–Fc chimera protein. N = 6 per group. Each gene expression was normalized to (C) GAPDH or (D) RPLP0. ∗P < .05, ∗∗P < .01, and ∗∗∗P < .001. P values were determined by the (A–C) Mann–Whitney test or the (D) Friedman test followed by the Dunn multiple comparison test.

Fig 3: BMP9 protein is present in growth plate cartilage. Immunofluorescence analysis of WT, Tgfbr1Col2, and Bmp9−/− mice to detect the presence of BMP9 in cartilage. WT and Tgfbr1Col2 sections are from E18.5 embryos, and Bmp9−/− sections are from P0 newborn pups. α-BMP9 staining is red. Nuclei are stained with DAPI (blue). n = 3 mice. (A and B) In WT and Tgfbr1Col2 mice, BMP9 protein is localized in bones as expected. However, BMP9 also presents in the cartilage in these 2 mouse lines. Arrow points to a blood vessel. C, cartilage; B, bone; M, muscle. (C) The specificity of anti-BMP9 antibody was validated by using Bmp9−/− mice. No positive signal was detected in Bmp9−/− mice. Arrow points to a blood vessel. C, cartilage; B, bone; M, muscle.

Fig 4: BMP9–ALK1 signaling restricts the stemness of human colonic IECs. (A) EdU assay in NIBD patient-derived colonic epithelial cell monolayers (N = 4–8 per group). Expanded cells were cultured in EM in the presence or absence of BMP9 and ALK1–Fc chimera protein. Red, EdU; blue, Hoechst 33342. (B) Proliferation- and stemness-related gene expression in NIBD patient-derived colonic epithelial cell monolayers (N = 6 per group). (C) Proliferation- and stemness-related gene expression in colonic epithelial cells isolated from CD patients (N = 15) and NIBD controls (N = 12). (D) Representative immunohistochemical images of OLFM4 expression in the colonic crypts of NIBD controls (left) and CD patients (right). The percentage of OLFM4 staining area in colonic crypts was compared between CD patients and NIBD controls (N = 4 per group). Each gene expression was normalized to (B) RPLP0 or (C) GAPDH. ∗P < .05, ∗∗P < .01, and ∗∗∗P < .001. P values were determined by the (A) Kruskal–Wallis test or the (B) Friedman test followed by the Dunn multiple comparisons test, or the (C and D) Mann–Whitney test.

Fig 5: Endothelial PHD2 deficiency suppressed BMP signaling in lung endothelial cells. (a) RNA‐seq analysis demonstrated suppression of BMP signaling by PHD2 deficiency in HIF‐2α dependent manner in PH mice. Egln1 f/f (WT), Egln1 Tie2Cre (CKO), and Egln1/Hif2a Tie2Cre (EH2) mice. (b) mRNA expression of Bmpr2, Bmpr1b, Acrvl1, and Grem1 were dysregulated in the lung of CKO mice and normalized in EH2 mice. (c) Downregulated protein expression of BMPR2 and upregulation of GREM1 in the CKO lungs were normalized in EH2 lungs. (d) Decreased BMPR2 and increased GREM1 in the lung ECs of CKO lungs via immunostaining. V, vessel. (e) RNA‐sequencing analysis showed that PHD2 deficiency reduced the expression of BMP signaling molecules. HLMVECs were transfected with control siRNA (siCtl) or PHD2 siRNA for 48 h. Three replicates were pooled in an equal amount for RNA‐seq analysis. (f) qRT‐PCR analysis confirmed the reduction of BMP signaling, including upregulation of GREM1 and downregulation of BMPR1A and ID1 in PHD2‐deficient HLMVECs. N = 3. (g) BMPR2 expression and activities were reduced by PHD2 knockdown, but not affected by HIF‐2α knockdown in IPAH patient‐derived PAECs. IPAH PAECs were transfected with siCtl or PHD2 or HIF‐2α siRNA for 48 h, followed by treatment of BMP9 (5 ng/ml) for 16 h. (h) GREM1 is upregulated by HIF‐2α secondary to PHD2 deficiency. HLMVECs were transfected with siCtl or PHD2 siRNA or PHD2 plus HIF2A siRNA for 48 h. N = 3. (i) A diagram showing three putative hypoxia responsible element (HRE) sites in the promoter region of the human GREM1 promoter. Red highlighted texts indicate mutated HRE sequences. (j) Luciferase assay demonstrates that human GREM1 promoter activities were induced by PHD2 deficiency in an HIF‐2α‐dependent manner. HLMVECs were cotransfected with human GREM1 promoter firefly luciferase plasmids, control Renilla luciferase plasmids, and siCtl or PHD2 siRNA for 48 h, followed by treatment with dimethyl sulfoxide or HIF‐2α translational inhibitor C76 (20 μm) for 16 h. N = 3. (k) Mutagenesis studies and luciferase assays demonstrated that HRE sites in the GREM1 promoter mediate GREM1 activation in PHD2‐deficient HLMVECs. N = 3 to 4. (l) FK506 treatment reduced right ventricular (RV) systolic pressure in CKO mice. (m) RV hypertrophy was inhibited in FK506‐treated CKO mice compared to PBS. One‐way analysis of variance with Tukey posthoc analysis for multiple group comparisons (b, h, j, k). Student t test (f, l, m). CKO, conditional knockout; DAPI, 4′,6‐diamidino‐2‐phenylindole; HIF, hypoxia‐inducible factor; HLMVEC, human lung microvascular endothelial cell; IPAH, idiopathic pulmonary arterial hypertension; mRNA, messenger RNA; PAEC, pulmonary arterial endothelial cell; PBS, phosphate‐buffered saline; PH, pulmonary hypertension; PHD2, prolyl hydroxylase 2; qRT‐PCR, quantitative reverse‐transcription polymerase chain reaction; RVSP, right ventricular systolic pressure; siRNA, small interfering RNA; WT, wild‐type. *p < 0.05; **p < 0.01, and ***p < 0.001