Protein tyrosine phosphatase L1 represses endothelial-mesenchymal transition by inhibiting IL-1β/NF-κB/Snail signaling

Abstract

Endothelial–mesenchymal transition (EnMT) plays a pivotal role in various diseases, including pulmonary hypertension (PH), and transcription factors like Snail are key regulators of EnMT. In this study we investigated how these factors were regulated by PH risk factors (e.g. inflammation and hypoxia) in human umbilical vein endothelial cells (HUVECs). We showed that treatment with interleukin 1β (IL‐1β) induced EnMT of HUVECs via activation of NF-κB/Snail pathway, which was further exacerbated by knockdown of protein tyrosine phosphatase L1 (PTPL1). We demonstrated that PTPL1 inhibited NF-κB/Snail through dephosphorylating and stabilizing IκBα. IL‐1β or hypoxia could downregulate PTPL1 expression in HUVECs. The deregulation of PTPL1/NF-κB signaling was validated in a monocrotaline-induced rat PH (MCT-PH) model and clinical PH specimens. Our findings provide novel insights into the regulatory mechanisms of EnMT, and have implications for identifying new therapeutic targets for clinical PH.

Introduction

Endothelial–mesenchymal transition (EnMT) is a cellular process characterized by the loss of endothelial features and the acquisition of mesenchymal, fibroblast, or stem-cell-like characteristics [1]. It is defined by the loss of cellular adhesion and the cytoskeletal reorganization of actin and intermediate filaments that convert apical–basal polarity to front end–back end polarity to form spindle-shaped cells. During this transformation, there is a marked decrease in endothelial biomarkers such as CD144 and CD31, as well as increased expression of mesenchymal biomarkers such as α-SMA and SM-22α [2]. EnMT plays a pivotal role in various pathologic conditions, such as atherosclerosis [3], tumors [4, 5], vein graft failure [6], and fibrosis in key organs [7–9]. A recent study has shown that abnormal differentiation of EnMT‐derived SM‐like cells results in pulmonary hypertension (PH) [10–12]. However, the underlying mechanism remains to be fully elucidated.

Inflammation and hypoxia contribute to the pathogenesis of various types of PH, especially group III PH. Expression of nuclear factor-κB (NF-κB) and the pro-inflammatory cytokine interleukin‐1β (IL‐1β) is elevated in the lung tissue of a rat model of monocrotaline-induced PH (MCT-PH), and NF-κB activation may contribute to the development of this condition [13]. IL-1β could induce EnMT by activating the NF-κB pathway [14]. The NF-κB-regulated gene Snail is a key regulator of EnMT, cell adhesion, and proliferation [15, 16]. These findings highlight the critical role of NF-κB/Snail signaling in EnMT.

Protein tyrosine phosphatase L1 (PTPL1) is a high-molecular-weight (270 kDa) nonreceptor-type phosphatase. It contains multiple domains, providing numerous potential interfaces [17]. Previous studies have shown that PTPL1 plays an important role in the development of tumors [18–20] and idiopathic pulmonary fibrosis [21]. Recent studies have also shown that PTPL1 inhibits EMT in hepatocellular carcinoma [22]. However, the regulatory role and underlying molecular mechanisms of PTPL1 in PH, a disorder attributed necessarily to EnMT of vascular endothelial cells, are not clearly elucidated. Here, we investigated the role of PTPL1 in IL‐1β‐induced EnMT using human umbilical vein endothelial cells (HUVECs). Our findings suggest that PTPL1 represses EnMT by stabilizing IκBα, and therefore inhibiting the IL-1β/NF-κB/Snail pathway. It may help identify new potential drug targets for PH.

Materials and methods

A rat model of MCT-PH

Adult male Sprague-Dawley rats weighing 150–200 g were purchased from Shanghai SLAC Laboratory Animal Co. Ltd (Shanghai, China). All animal studies were approved by the Ethics Committee of Experimental Research at Fudan University Shanghai Medical College and were in accordance with the Institutional Animal Care and Use Committee Guidelines.

Animals were randomly divided into two groups (n = 8/group). The control group rats received phosphate-buffered saline (PBS). The MCT group rats received MCT (60 mg/kg; Sigma, St. Louis, MO, USA). All animals were raised under a 12 h:12 h light–dark cycle and were freely supplied with food and water. The room temperature was maintained at 25 °C, and the bedding was changed once per week.

Hemodynamic analysis and tissue preparation

In the rat model of MCT-PH, rat pulmonary vascular and hemodynamic alterations and RV remodeling occurred 28 days after MCT injection. At that point, the right jugular vein was carefully isolated. A specially shaped catheter linked to the PowerLab system (AD Instruments, Bella Vista, NSW, Australia) was inserted into the right ventricle (RV) via this vein, and right ventricular systolic pressure (RVSP) was recorded. Then, sternotomy was performed, and rats were perfused with paraformaldehyde. Both lungs and the heart were harvested. The RV and the left ventricle plus septum (LV + S) were weighed, and the RV/LV + S weight ratio was determined as an indication of right ventricular hypertrophy. Next, the lower lobe of the right lung was sectioned into 4-mm-thick slices and soaked in a 10% formalin solution (pH 7.4). The other parts were kept at −80 °C for further experiments.

Hematoxylin and eosin staining

The fixed lungs were sliced in the mid-sagittal plane, embedded in paraffin, and cut into ~5-μm-thick sections with a microtome. Then, the sections were placed on glass slides, stained with hematoxylin and eosin (HE) for morphological analysis, and visualized under an Olympus BX41 microscope (Tokyo, Japan).

Cell culture and induction of EnMT

HUVECs (passages 3–5; ScienCell Research Laboratories, California, USA) were cultured in endothelial cell medium (ECM). To induce EnMT, recombinant IL-1β (10 ng/mL; Peprotech) in 0.5% fetal bovine serum medium was used to treat the cells for 48 h. Human Pulmonary Artery Endothelial Cells (HPAECs) (passages 3–5; ScienCell Research Laboratories, California, USA) were cultured in ECM. Cells were maintained in a humidified atmosphere at 37 °C in 5% CO₂.

Immunofluorescence staining

Cells were fixed with 4% paraformaldehyde for 30 min at room temperature. The cells were then incubated with primary antibodies against α-SMA and CD31 (Abcam, Cambridge, MA, USA) overnight at 4 °C and subsequently with Alexa Fluor‐conjugated secondary antibodies (Invitrogen, Carlsbad, CA) for 2 h at 37 °C. After nucleus staining with 4ʹ,6-diamidino-2-phenylindole (DAPI) dye, cells were examined under a fluorescence microscope. A minimum of three replicates were performed for each examination.

Lentiviral construction and transduction

The lentiviral shuttle vector for the PTPL1-targeted short hairpin RNA, sh-PTPL1, and the packaging vectors, including pMD2.0G and psPAX, were purchased from Addgene (Cambridge, MA, USA). They were transfected into HEK293T cells for packaging and amplification of lentivirus. For transient transduction, cells at approximately 80% confluence were transduced with the lentiviral vectors at a multiplicity of infection of 60 for 4 h. Afterward, the media were changed to fresh ECM, and the transfected cells were cultured for an additional 72 h.

Quantitative RT-PCR

Total RNA was extracted from cultured cells using TRIzol reagent (Invitrogen, Life Technologies, Carlsbad, CA, USA). Reverse transcription was performed using a first-strand cDNA reverse transcription kit (Takara, Otsu, Japan), and real‐time PCR was performed using a SYBR Green/ROX qPCR Master Mix kit according to the manufacturer’s instructions (Takara, Otsu, Japan). The relative quantification was determined using the ΔΔCt method with β-actin as a reference gene. All primer sequences are shown in Table 1.

Table 1.

Primers used for qPCR

Gene	Forward 5′–3′	Reverse 5′–3′
CD31 (Human)	ACCAGAGCTATTCCCAAAAGACC	TGCGGCGATTCATCAGGAAAT
CD144 (Human)	AGAGCTCCACTCACGCTCAG	CATCTTCCCAGGAGGAACAG
α-SMA (Human)	AAGCACAGAGCAAAAGAGGAAT	ATGTCGTCCCAGTTGGTGAT
SM-22α (Human)	AGTGCAGTCCAAAATCGAGAAG	CTTGCTCAGAATCACGCCAT
PTPL1 (Human)	TGGCTCTCCAGGCTGAGTATG	AACACCAAACAAAATGGTCCTG
β-actin (Human)	GGACCTGACTGACTACCTCAT	CGTAGCACAGCTTCTCCTTAAT
IL-1β (Rat)	TTCAGCAGTAAAGCGTGTGG	GCGTGTGTGTGTGTGTGTGT
IL-6 (Rat)	ACCACCCACAACAGACCAGT	TTTCACAGCCTACCCACCTC
TNF-α (Rat)	AGAGCCCCCAATCTGTGTC	TTCAGCGTCTCGTGTGTTTC
PTPL1 (Rat)	TGACACACCATGGTCACAGTT	CGACAGAAGTAGCATCGCCG
GAPDH (Rat)	CCATTCTTCCACCTTTGATGCT	TGTTGCTGTAGCCATATTCATTGT

Western blotting

Proteins from cultured cells were extracted with radioimmunoprecipitation assay lysis buffer and quantified with the bicinchoninic acid protein assay kit. Then, the proteins were separated with sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS‐PAGE) and transferred to a polyvinylidene fluoride transfer membrane. The membranes were blocked in 5% nonfat dry milk for 1 h, followed by incubation overnight at 4 °C with primary antibodies. The membranes were then washed and incubated with secondary antibodies, washed with Tris‐buffered saline and Tween 20 (0.1%), and treated with electrochemiluminescence reagents for blot detection. Blots were quantified by densitometry using ImageJ (NIH Image, Bethesda, MD). The primary antibodies used are as follows: CD31 (1:1000), α-SMA (1:100), CD144 (1:1000), SM-22α (1:1000), IκBα (1:1000), phospho-IκBα (Y42, 1:1000), and β-actin (1:1000) were purchased from Abcam (Cambridge, MA, USA); PTPL1 (1:1000) was purchased from Novus Biologicals; and NF-κB (1:1000), phospho-NF-κB (p65, 1:1000), and Snail (1:1000) were purchased from Cell Signaling Technology (Danvers, MA, USA). Secondary horseradish‐peroxidase‐conjugated antibodies were purchased from Jackson ImmunoResearch Laboratories (1:5000, dilution; ImmunoResearch Laboratories, Inc., West Grove, PA, USA).

Immunohistochemical assay

All the specimens were fixed with 4% paraformaldehyde and embedded with paraffin. Four-micrometer-thick sections were cut and transferred to glass slides coated with 100 g/L polylysine. The immunohistochemical (IHC) staining was performed as described [23]. Anti-PTPL1 antibody was purchased from Novus Biologicals. The primary antibodies were diluted in PBS (1:50). IHC was also performed on paraffin-embedded sections. The sections were autoclaved for 10 min at 121 °C for antigen retrieval. An Envision kit (DAKO, Glostrup, Denmark) was used for staining according to the manufacturer’s instructions.

Immunoprecipitation

Cells were lysed in buffer containing 1% (v/v) Nonidet P-40, 0.5 mM EGTA, 5 mM sodium orthovanadate, 10% (v/v) glycerol, 100 μg/mL phenylmethylsulfonyl fluoride, 1 μg/mL leupeptin, 1 μg/mL pepstatin A, 1 μg/mL aprotinin, and 50 mM HEPES (pH 7.5). Thereafter, 1000 μL of diluted lysate (1 μg protein/μL) was incubated overnight with 5 μg of an antibody for immunoprecipitation (IP). Immune complexes were captured by adding 40 μL of a 1:1 (v/v) suspension of protein A:Sepharose 4B beads and rotating the mixture for 2 h at 4 °C. The beads were harvested, and the bound proteins were resolved by SDS-PAGE and analyzed by Western blotting. The primary antibodies used for IP are described in the Western blotting section.

Migration assays

Cell migration was assessed using transwell chambers (24 wells, 8 μm pore size; BD Biosciences, San Diego, CA, USA). Samples containing 1 × 10⁵ cells were resuspended in serum‐free ECM and loaded into the upper chamber. The chambers were incubated for 24 h, with complete culture medium added to the lower chamber. Nonmobile cells were removed, and the chambers were stained with crystal violet. Five randomly selected fields were counted under an inverted light microscope.

Tube formation assays

HUVECs were harvested and suspended in ECM. Forty-eight-well plates were coated with Matrigel (BD Biosciences), which was allowed to polymerize at 37 °C in a 5% CO₂ chamber for 30 min. HUVECs (1 × 10⁴ cells/mL) were added to each chamber, treated as indicated, and incubated for 8 h at 37 °C in 5% CO₂, followed by observation of capillary-like tube formation using phase-contrast microscopy.

RNA-Seq analysis

We analyzed the change in the mRNA profiles between HPAECs that were treated with and without hypoxia for 6 h. Total RNA was isolated using TRIzol reagent (Invitrogen, Carlsbad, CA, USA) according to the manufacturer’s instructions. cDNA library preparation was performed using the RNA-Seq Sample Prep Kit (Illumina, NEB, Ipswich, MA, USA) according to the manufacturer’s instructions. For the QC step, an Agilent 2100 Bioanalyzer and an ABI StepOnePlus Real-Time PCR System were used to qualify and quantify the sample library. Each cDNA library was amplified once before sequencing. Sequencing was performed on an Illumina HiSeq X Ten at Biotecan Co., Ltd sequencer (Shanghai, China). The RNA-Seq data were analyzed as previously described [24, 25]. Differentially expressed genes (with a fold change over 1.5 and P < 0.05) were selected.

Statistical analysis

Data are presented as the means ± SEMs. All experiments were repeated at least three times. Statistical analysis was performed with GraphPad Prism 7.0 software. Comparisons among groups were made using one‐way analysis of variance or Student’s t-test. P values <0.05 were considered significant.

Results

IL-1β induces EnMT in HUVECs by activating the NF-κB/Snail pathway

Exposure of HUVECs to IL-1β (10 ng/mL) for 48 h caused an obvious alteration in cellular morphology from a polygonal, cobblestone‐like shape to a more spindle and fibroblast‐like shape (Fig. 1a). Quantitative RT-PCR (qRT‐PCR) showed that IL‐1β treatment reduced the mRNA levels of endothelial markers (CD31 and CD144 were greatly reduced) and increased those of the mesenchymal markers α-SMA and SM-22α (Fig. 1b). To further confirm that EnMT was induced by IL-1β, the protein levels of the endothelial or mesenchymal markers were examined by Western blot assay and immunofluorescent staining. As a result, CD31 and CD144 were significantly downregulated, and α-SMA and SM-22α were remarkably upregulated by IL‐1β treatment (Fig. 1c, d). In addition, HUVEC tube formation was significantly inhibited by IL‐1β (Fig. 1e). Consistent with the transition to a mesenchymal phenotype, HUVECs treated with IL-1β exhibited enhanced migration capabilities when compared with untreated cells (Fig. 1f). Exposure of HUVECs to IL-1β activated NF-κB signaling and increased the level of Snail (Fig. 1g). These results suggest that IL-1β likely elicits EnMT of HUVECs via NF-κB/Snail signaling.

Fig. 1.

IL-1β induces EnMT in HUVECs by activating the NF-κB/Snail pathway. a Microscopy of HUVECs treated with or without IL-1β (10 ng/mL) for 48 h. IL-1β‐treated HUVECs had a fibroblast‐like phenotype, whereas untreated control cells had an endothelial cell phenotype. b The relative mRNA levels of the endothelial markers CD31 and CD144 and the mesenchymal markers α-SMA and SM-22-α in control and IL-1β-treated HUVECs were assessed by qRT‐PCR (each bar represents the mean ± SEM. **P < 0.01, ***P < 0.001 vs. the control group). c The protein levels of CD31, CD144, α-SMA, and SM-22-α in control and IL-1β-treated HUVECs were determined by Western blot analysis. (each bar represents the mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001 vs. the control group). d Immunofluorescent staining of CD31 (green) and α-SMA (red) expression in control and IL-1β-treated HUVECs. e Tube formation assays for unstimulated and IL-1β-exposed HUVECs (each bar represents the mean ± SEM. *P < 0.05 vs. the control group). f Transwell assay to assess the migration of unstimulated and IL-1β-exposed HUVECs (wach bar represents the mean ± SEM. ***P < 0.001 vs. the control group). g The protein levels of p-NF-κB (p65), NF-κB, and Snail were determined by Western blot analysis using lysates from unstimulated and IL-1β-exposed HUVECs (each bar represents the mean ± SEM. *P < 0.05, ***P < 0.001 vs. the control group)

PTPL1 impairs IL-1β-induced EnMT by inactivating NF-κB

We next examined the regulatory role of PTPL1 in the IL‐1β‐induced EnMT of HUVECs. PTPL1 was knocked down in HUVECs via lentiviral vector-based sh-RNAs (Fig. 2a–c). Consequently, PTPL1 knockdown increased the mRNA and protein expression of α-SMA and SM-22α and decreased CD31 and CD144 expression (Fig. 2d, e). The HUVEC tube formation capacity was significantly inhibited after PTPL1 silencing (Fig. 2f). In accordance with the transition to a mesenchymal phenotype, PTPL1-knockdown HUVECs displayed enhanced migration abilities compared with the control cells (Fig. 2g). Control and PTPL1-knockdown HUVECs were further subjected to treatment with IL-1β (10 ng/mL) for 48 h. We found that knockdown of PTPL1 synergized with IL‐1β to upregulate the mesenchymal proteins α‐SMA and SM-22α and impair the expression of the endothelial biomarkers CD31 and CD144 (Fig. 2h). Similarly, PTPL1-knockdown HUVECs showed much higher NF-κB activity and Snail levels than control cells in response to treatment with IL‐1β (Fig. 2i). These data indicate that PTPL1 represses the IL‐1β-induced EnMT of HUVECs by counteracting NF-κB/Snail signaling.

Fig. 2.

PTPL1 impairs IL-1β-induced EnMT by inactivating NF-κB. a Fluorescent microscopy of HUVECs infected with recombinant lentiviruses expressing control or PTPL1-targeted sh-RNA for 72 h. b–e The PTPL1 levels in cells in a were measured by qRT‐PCR (b, d) and Western blot (c, e) (each bar represents the mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001 vs. the sh-mock group). f Tube formation assay for cells in a (each bar represents the mean ± SEM. *P < 0.05 vs. the sh-mock group). g Cell migration was assessed by transwell assays (each bar represents the mean ± SEM. ****P < 0.0001 vs. the sh-mock group). h, i The cells in a were treated with IL-1β (10 ng/mL) and were subjected to Western blot analysis (each bar represents the mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001 vs. the sh-mock group; ^#P < 0.05, ^##P < 0.01 vs. the sh-PTPL1 group)

PTPL1 inhibits the NF-κB signaling pathway by stabilizing IκBα

PTPL1 has been predicted to dephosphorylate various proteins, including IκBα [26]. We thus investigated whether PTPL1 inhibits NF-κB activity and impedes the EnMT of HUVECs by targeting IκBα. Indeed, PTPL1 was coimmunoprecipitated with IκBα from HUVEC lysates (Fig. 3a). While IL‐1β induced the phosphorylation of IκBα at tyrosine 42 and triggered its degradation in HUVECs, PTPL1 knockdown further promoted the phosphorylation and downregulation of the IκBα protein (Fig. 3b). Therefore, PTPL1 may attenuate NF-κB signaling by dephosphorylating IκBα and increasing its half-life in vascular endothelial cells.

Fig. 3.

PTPL1 inhibits the NF-κB signaling pathway by stabilizing IκBα. a Immunoprecipitation of PTPL1 and IκBα using lysates from HUVECs. b HUVECs infected with sh-mock or sh-PTPL1 lentiviruses were further treated with IL-1β (10 ng/mL) and were subjected to Western blot analysis (each bar represents the mean ± SEM. *P < 0.05, **P < 0.01 vs. the sh-mock group; ^#P < 0.05, ^##P < 0.01 vs. the sh-PTPL1 group)

PTPL1 is downregulated by IL-1β and hypoxia in human vascular endothelial cells

We next asked how the expression of PTPL1 was regulated in vascular endothelial cells. We found that treatment of HUVECs with IL-1β significantly reduced the levels of PTPL1 (Fig. 4a, b). Tissue hypoxia is a risk factor for diverse clinical disorders, e.g., group 3 PH, that pathologically involves EMT [27, 28] or EnMT [29–31]. We thus probed whether hypoxia plays a regulatory role in PTPL1 expression in vascular endothelial cells. Our RNA-Seq results showed that hypoxia treatment of HPAECs induced robust changes in the expression profile of genes, including PTPL1 (Fig. 4c). The downregulation of PTPL1 by hypoxia in HUVECs was further verified by Western blot (Fig. 4d). These results show that PTPL1 downregulation by inflammatory factors or hypoxia contributes to the EnMT of vascular endothelial cells.

Fig. 4.

PTPL1 is downregulated by IL-1β and hypoxia in human vascular endothelial cells. a, b HUVECs were treated with IL-1β (10 ng/mL) for 48 h and subjected to qRT-PCR (a) and Western blot analysis (b) (each bar represents the mean ± SEM. *P < 0.05, **P < 0.01 vs. the control group). c Heatmap of RNA-Seq data in HPAECs after hypoxia treatment (T: hypoxia; C: normoxia). d Western blot analysis of HUVECs exposed to hypoxia (1% O₂, 94% N₂, and 5% CO₂) for 48 h (each bar represents the mean ± SEM. *P < 0.05 vs. the normoxia group)

PTPL1/NF-κB signaling participates in PH pathogenesis

We evaluated the involvement of PTPL1 and the NF-κB pathway in the pathogenesis of PH. A rat model of MCT-induced PH (MCT-PH) was established via exposure to MCT for 4 weeks, as verified by significantly increased RVSP (Fig. 5a) and mean pulmonary arterial pressure (mPAP) (Fig. 5b) and a significantly increased ratio of right ventricle to left ventricle plus septum (RV/LV + S) weight (Fig. 5c). To validate NF-κB activation in vivo, we generated a rat model of MCT-PH characterized by abnormal media thickening (Fig. 5d). Quantitative RT-PCR showed that the expression of pro-inflammatory cytokines, such as tumor necrosis factor‐α (TNF‐α), interleukin (IL)‐6 and interleukin (IL)‐1β, was increased, whereas the level of PTPL1 was reduced in the lung tissue of MCT-PH rats compared to control rats (Fig. 5e, f). We also found that the level of phosphorylated NF-κB protein was significantly elevated in the lung tissue of MCT-PH rats compared to control rats (Fig. 5g). In addition, we collected lung tissue samples from five patients with PH and examined the expression of PTPL1 in the lung tissues of different patients by immunohistochemical staining. The results showed that PTPL1 levels were lower in the pulmonary arterial endothelial cells of PH patients than in control lung cancer patients (Fig. 5h). Therefore, PTPL1 might suppress the development of PH via inhibition of NF-κB activity.

Fig. 5.

PTPL1/NF-κB signaling is involved in in vivo PH pathogenesis. a–c Rats were treated with MCT as described in the Materials and methods. RVSP (a), mPAP (b), and RV/(LV + S) (c) data were obtained from each group and plotted (each bar represents the mean ± SEM. ****P < 0.0001 vs. the control group). d HE staining of paraffin-fixed lung sections for morphological analysis of pulmonary arteries from control or MCT-PH rats. e–g Pulmonary artery samples from control or MCT-PH rats were used for qRT-PCR (e, f) and Western blot (g) analyses. h Immunohistochemical staining of lung tissues from PH patients and control lung cancer patients. Bar, 50 μm (each bar represents the mean ± SEM. **P < 0.01, ****P < 0.0001. vs. the control group)

Discussion

In this study, we demonstrated that PTPL1 suppresses the IL‐1β‐induced EnMT process in HUVECs by impairing NF-κB/Snail signaling. It has been established that EnMT plays critical roles in PH [23, 32, 33]. IL‐1β is one of the most important pro-inflammatory cytokines that elicit EnMT [34–36]. We found that HUVECs exposed to IL-1β could undergo EnMT, as verified by the transition of cells to a spindle-like shape, varied expression of endothelial and fibroblast markers, increased migration ability, and reduced tube formation capacity. Consistent with studies reporting that the NF-κB/Snail pathway plays an important role in EMT [37, 38], we found that IL-1β induced EnMT in HUVECs through activation of NF-κB and upregulation of Snail. The activity of NF-κB is regulated by divergent stimuli dependent upon the intracellular signaling context [39, 40], which prompted us to explore the regulatory mechanisms of NF-κB activation in the inflammatory factor-induced EnMT of vascular endothelial cells.

PTPL1 is a nonreceptor-type protein tyrosine phosphatase involved in various cellular signal pathways. Previous studies have found that PTPL1 acts as both a tumor promoter [41] and a suppressor [42]. A previous study showed that PTPL1 overexpression significantly inhibited the progression of hepatocellular carcinoma cells by inhibiting EMT [22]. In this study, we found that PTPL1 could inhibit IL‐1β‐induced EnMT in HUVECs, and knockdown of PTPL1 could activate NF-κB signaling and increase the level of Snail. Unlike previous studies showing that PTPL1 controls NF-κB activation via p75NTR [43], we found that PTPL1 attenuates NF-κB activation by dephosphorylating IκBα at tyrosine 42 and consequently increasing the half-life of IκBα. These findings are in agreement with a recent report that PTPL1 could target IκBα in ovarian cancer cells [44]. Nonetheless, given the wide crosstalk between intracellular signaling pathways and the relatively low number of protein tyrosine phosphatases compared with protein tyrosine kinases, we cannot rule out that PTPL1 also regulates EnMT by targeting other pathways in vascular endothelial cells.

Both inflammation [45–47] and hypoxia [48, 49] contribute to the pathogenesis of various forms of PH. Our study showed that hypoxia or IL-1β could downregulate PTPL1 in vascular endothelial cells, consistent with a previous finding that PTPL1 expression was impeded upon exposure of prostate cancer cells to hypoxia [50]. We also found that the expression of inflammatory cytokines such as TNF‐α, IL‐6, and IL-1β was increased in the lung tissue of a rat model of MCT-PH, adding weight to the probability that the inflammatory microenvironment and endothelial PTPL1 in the lung form a feedback circuit to control EnMT and the pathogenesis of PH. Together, the present findings demonstrated a suppressive role of PTPL1 in EnMT and the deregulation of PTPL1/NF-κB signaling in the pathogenesis of PH.

Author contributions

SQL designed the experiments and prepared the initial manuscript. XMW and GW conducted the experiments. LD and NZ cultivated the cells. CWL and JWX were responsible for the experimental calculations. YZZ, XJZ, YYZ, and PZ were involved in the experimental analysis.

Competing interests

The authors declare no competing interests.