Super‐Enhancer–Driven Syndecan‐4 Regulates Intercellular Communication in Hypoxic Pulmonary Hypertension

Abstract

Background

Unveiling pro‐proliferation genes involved in crosstalk between pulmonary artery endothelial cells and pulmonary artery smooth muscle cells (PASMCs) are important to improving the therapeutic outcome of pulmonary hypertension (PH). Although growing studies have shown that super‐enhancers (SEs) play a pivotal role in pathological and physiological processes, the SE‐associated genes in PH and their impact on PASMC proliferation remain largely unexplored.

Methods and Results

We used serotype 5 adenovirus‐associated virus to interfere with syndecan‐4 and constructed an SU5416 combined with hypoxia–PH model. Chromatin immunoprecipitation sequencing analysis, chromatin immunoprecipitation quantitative polymerase chain reaction, and bioinformatics were used to confirm early growth response 1 was involved in regulating syndecan‐4‐associated SE in PASMCs. The effects of syndecan‐4 and its underlying mechanisms were subsequently elucidated using Western blot, coimmunoprecipitation, and cell coculture assays. Herein, we identified a novel SE‐associated gene, syndecan‐4, in hypoxia‐exposed PASMCs. Syndecan‐4 was transcriptionally driven by early growth response 1 via an SE and was significantly overexpressed in hypoxic PASMCs and plasma from patients with PH. Mechanism studies revealed that syndecan‐4 induces PASMC proliferation by interacting and regulating protein kinase C α ubiquitination. In addition, syndecan‐4 was enriched in exosomes secreted from hypoxic PASMCs, which subsequently transported and led to pulmonary artery endothelial cell dysfunction. Syndecan‐4 inhibition in hypoxia by serotype 5 adenovirus‐associated virus treatment attenuated the pulmonary artery remodeling and development of PH in vivo.

Conclusions

Taken together, our results demonstrate that an SE‐driven syndecan‐4 modulates crosstalk of PASMCs and pulmonary artery endothelial cells and promotes vascular remodeling via the protein kinase C α and exosome pathway, thus providing potential targets for the early diagnosis and treatment of hypoxic PH.

Nonstandard Abbreviations and Acronyms

AAV5

serotype 5 adenovirus‐associated viruses

ChIP

chromatin immunoprecipitation

EdU

ethynyl‐2′‐deoxyuridine

EGR1

early growth response 1

H3K4me1

monomethyl H3K4

HIF

hypoxia‐inducible factor

hPASMCs

human pulmonary artery smooth muscle cells

mPASMCs

mouse pulmonary artery smooth muscle cells

PAECs

pulmonary artery endothelial cells

PASMCs

pulmonary artery smooth muscle cells

PCNA

proliferating cell nuclear antigen

PH

pulmonary hypertension

PRKCA

protein kinase C α

SDC4

syndecan‐4

SE

super‐enhancer

SuHx

SU5416 combined with hypoxia

Clinical Perspective.

What Is New?

• A novel super‐enhancer–driven syndecan‐4 was upregulated in hypoxic pulmonary hypertension.

• Super‐enhancer–driven syndecan‐4 modulates crosstalk of pulmonary artery smooth muscle cells and pulmonary artery endothelial cells and promotes vascular remodeling via the protein kinase C α and exosome pathway.

What Are the Clinical Implications?

• Our results identified a critical function of super‐enhancer–associated syndecan‐4 in the regulation of pulmonary vascular remodeling, providing a new therapeutic target for pharmaceutical intervention of pulmonary hypertension.

Pulmonary hypertension (PH) is a life‐threatening cardiovascular and respiratory system disease characterized by increased vascular remodeling, leading to right heart failure and even death. The structural remodeling process of pulmonary vasculature is mainly due to the dysfunction of pulmonary artery endothelial cells (PAECs) and the increased proliferation/decreased apoptosis of pulmonary artery smooth muscle cells (PASMCs). ¹ , ² Although the mechanism underlying PH is not fully understood, PASMC proliferation plays a center role in the process of vascular remodeling. ³ More importantly, increasing evidence has shown that the intercellular communication between PAECs and PASMCs that is mediated by exosomes plays an essential role in the development and progression of vascular pathology in PH. ⁴ , ⁵ Targeting and characterizing the key molecular pathogenesis involved in the crosstalk of PAECs and PASMCs in mediating vascular remodeling is believed to be a promising direction.

Super‐enhancers (SEs) are large clusters of enhancers with strong transcriptional activity, recruiting core transcription factors, cofactors, and enhancers associated with epigenetic signatures to govern gene expression and regulate pathogenic gene activation. ⁶ , ⁷ Studies have shown that genes driven by SEs are positively correlated with physiological and pathological processes, including acute myeloid leukemia, cardiovascular disease, and especially cancers. For example, SE‐driven TOX2 was reported to mediate tumor formation of natural killer/T cell lymphoma. ⁸ SE‐driven lncRNA Snhg7 was showed to aggravate cardiac hypertrophy via the Tbx5/GLS2/ferroptosis axis. ⁹ Importantly, our previous study demonstrated that SE‐associated circular RNA circKrt4 regulates PAEC dysfunction in hypoxia‐induced PH. ¹⁰ Therefore, it will be of great interest to identify and characterize such SE‐associated genes as biomarkers and therapeutic targets in the pathogenesis of PH.

Exosomes, as extracellular vesicles, serve as crucial carriers of diverse bioactive substances, functioning as pivotal mediators in intercellular communication. These bioactive substances, encompassing proteins, lipids, and RNAs, are internalized by recipient cells, thereby playing integral roles in both physiological and pathological processes. ¹¹ , ¹² Compelling evidence demonstrated that exosomes contribute to the progression of tumor and cardiovascular diseases and could be recognized as potential biomarkers and therapeutic targets, including PH. Aliotta et al showed that exosomes containing microRNAs induces and reverses monocrotaline‐induced PH in mice. ¹³ Our present study showed that hypoxia promoted the enrichment of 15‐lipoxygenase 2 in exosomes in PAECs, and GW4869, the exosomes release inhibitor, restrained the hypoxia‐induced vascular remodeling and PH. ¹⁴ The studies above implied that the modulation of vascular‐derived exosomes has the potential to suppress hypoxia‐induced vascular remodeling. Given the complexity of the biomolecules contained in exosomes, a better exploration of the enrichment of biomolecules in exosomes in PH is required.

Syndecan‐4 is an important member of the transmembrane heparan sulfate proteoglycans. It is widely expressed on a variety of tissues and cell types and acts as a coreceptor to activate multiple signaling pathways by interacting with extracellular matrix proteins, kinases, or other cytokine partners, regulating a variety of cellular processes, such as cell apoptosis, migration, and calcification. ¹⁵ , ¹⁶ , ¹⁷ , ¹⁸ Moreover, studies have shown that syndecan‐4 is a key regulator of inflammation and that it functions via reactive oxygen species. ¹⁹ In hepatocellular carcinoma, syndecan‐4 as a direct anti–hepatocellular carcinoma cellular target of bufalin in inhibiting cell proliferation, invasion, and angiogenesis. ¹⁶ Remarkably, syndecan‐4 was previously shown to regulate the formation of extracellular vesicles and defines extracellular vesicle uptake and oncogenic effects on recipient cell populations in gastric cancer invasion. ²⁰ However, the role of syndecan‐4 in PH occurrence and progression remains unknown.

The current study explored the function and implication of syndecan‐4 driven by SE and EGR1 (early growth response 1) is enriched in the hypoxic PASMCs and exosomes. Knockdown of syndecan‐4 attenuated hypoxia‐induced intercellular communication between PAECs and PASMCs in pulmonary vascular remodeling via PRKCA (protein kinase C α). Our data identified a critical function of SE‐associated syndecan‐4 in the regulation of pulmonary vascular remodeling, providing new therapeutic target for pharmaceutical intervention of PH.

Methods

The data supporting this study's findings are available from the corresponding author upon reasonable request.

Study Approval

These experimental procedures were approved by the Ethics Committees of Harbin Medical University (HMUDQ20240513001).

Human Plasma Samples

A total of 20 (control=10, PH=10) plasma samples from controls and patients with PH were obtained at the First Affiliated Hospital of Chongqing Medical University. Male and female patients who had PH between 2018 and 2022 and who met the following criteria were considered to be potential participants: age between 20 and 80 years and mean pulmonary artery pressure ≥20 mm Hg. Patients using medication for PH‐related symptoms were excluded. In addition, the control group must have the same criteria as the patients except mean pulmonary artery pressure ≥20 mm Hg. The clinical information pertaining to the patients included in the plasma sample cohort is detailed in Table S1. Before participation, written informed consent was obtained from all individuals enrolled in this study. Whole blood samples (5 mL) were centrifuged at 1500g for 15 minutes at 4 °C to separate plasma from cellular components. These plasma samples were promptly transferred to RNase/DNase‐free tubes and stored at −80 °C until further analysis was conducted.

Quantitative Reverse Transcription Polymerase Chain Reaction

Total RNA was extracted from cells and lung tissues with TRIzol reagent (Invitrogen, Carlsbad, CA) according to the manufacturer's protocol. cDNA was synthesized from 2 μg of RNA using the Superscript First‐Strand Complementary DNA Synthesis Kit (HaiGene, Harbin, China). Quantitative real‐time polymerase chain reaction (PCR) was performed using SYBR Green real‐time PCR (Toyobo, Japan) in a Roche LightCycler 480II System. The relative expression of target mRNAs was determined using ΔCT (Cycle threshold) against endogenous β‐actin control, and the data were analyzed using the 2^−ΔΔCT method. The nucleotide sequences of the primers used are listed in Table S2.

Serotype 5 Adenovirus‐Associated Virus SU5416 Combined With Hypoxia PH Model

Healthy 6‐week‐old male C57BL/6J mice (weighing 20–25 g each) were obtained from the Laboratory Animal Center of the Second Affiliated Hospital of Harbin Medical University. All experimental procedures were performed following the Guide for the Care and Use of Laboratory Animals published by the US National Institutes of Health. The corresponding syndecan‐4 target RNA cloning construction and serotype 5 adenovirus‐associated virus (AAV5) were packaged by Genechem (Shanghai, China). 10¹¹ genome equivalents were prepared in 20 μL of HBSS, and mice anesthetized with isoflurane were infected with AAV5‐syndecan‐4 and AAV5 negative control via nasal drops. After 7 to 14 days, the infected mice were randomly divided into normoxic (Fi, O₂ 0.21) and hypoxic (Fi, O₂ 0.10) groups as described in a previous study. ¹⁰ , ²¹ , ²² SU5416 (S8442; Sigma Aldrich, Darmstadt, Germany) was injected subcutaneously at a dose of 20 mg/kg, and control mice were injected with the same volume of vehicle alone for the next 3 weeks in 10% oxygen, followed by preexposure to normoxia for 2 weeks.

Echocardiography and Right Ventricular Systolic Pressure Measurements

All mice were anesthetized through an intraperitoneal injection of avertin (200 mg/kg IP, Sigma Aldrich, St. Louis, MO). Mice were subjected to echocardiography using a Vevo2100 imaging system (VisualSonics, Toronto, Ontario, Canada). The pulmonary artery velocity time integral, pulmonary artery acceleration time, left ventricular ejection fraction and heart rate were obtained from stable images. The right ventricular (RV) systolic pressure was measured with PowerLab monitoring equipment (AD Instruments, Colorado Springs, CO). A 1.2 French pressure catheter (Scisense Inc, London, Ontario, Canada) was inserted into the superior vena cava and ultimately into the RV vein, enabling continuous recording of RV systolic pressure for a minimum duration of 5 minutes. Subsequently, the thoracic cavity of the mice was rapidly excised using surgical scissors, and the heart and lung tissues were excised. The RV hypertrophy index was calculated by determining the ratio of the RV free wall weight to the combined weight of the septum and left ventricular free wall. To ensure consistency and accuracy, all measurements were conducted by a single experienced investigator. Offline data analysis was performed using LabChart 8 Reader and Vevo2100 software in a blinded fashion by an investigator, ensuring objectivity and precision in the interpretation of the results.

Morphometric Analysis

In brief, lung tissues were immersed in 4% paraformaldehyde and embedded in paraffin, the blocks were cut into 5‐μm‐thick sections for hematoxylin and eosin staining. Hematoxylin and eosin staining was performed following the manufacturer's instructions. Images were taken using Aperio VERSA 8 Scanning System (Leica, Teaneck, NJ). Notably, all analyses were conducted by an independent investigator blinded to the experimental protocol, ensuring objective and unbiased assessment of the histological data.

Cell Culture

All cells were purchased from Procell Life Science and Technology (Wuhan, China). Human pulmonary artery smooth muscle cells (hPASMCs) and mouse pulmonary artery smooth muscle cells (mPASMCs) were cultured in smooth muscle cell medium (No. 1101, ScienCell Research Laboratories, Carlsbad, CA), which contained 1% smooth muscle cell growth supplement, 15% fetal bovine serum, and 1% penicillin and streptomycin. Human PAECs and mouse PAECs were cultured in endothelial cell medium (No. 1001, ScienCell Research Laboratories), which contained 1% endothelial cell growth supplement, 15% fetal bovine serum, and 1% penicillin and streptomycin. Cells were cultured at 37 °C with 5% CO₂ in humidified conditions. All the experiments were performed with 6 different origins of cells that were less than passage 6. For the hypoxia exposure experiments, cells were incubated in a Tri‐Gas Incubator (Thermo, Waltham, MA) in a water‐saturated atmosphere containing 3% O₂ and 5% CO₂ for 24 hours.

Isolation and Identification of Exosomes

To collect exosomes derived from PASMCs, cells were maintained in a serum‐free medium (UR51102, Umibio, Shanghai, China) for exosomes. Exosome isolation was carried out by Total Exosome Isolation Kit (UR52121, Umibio, Shanghai, China) according to the manufacturer's protocol. Subsequently, the medium was collected and centrifuged at 3000g for 10 minutes at 4 °C to separate cellular debris and other impurities. The supernatant was then mixed with an exosome concentration solution and incubated for 2 hours at 4 °C to enrich the exosome fraction. Following this, the mixture was centrifuged at 10 000g for 60 minutes at 4 °C to pellet the exosomes. Finally, the exosome pellet was resuspended in PBS for further analysis or downstream applications. The morphology of exosomes is scanned by transmission electron microscope, and their diameters are estimated by Nanosight instrument (Ribobio, Guangzhou, China). Exosomes were labeled with PKH26 (UR52302, Umibio) according to the manufacturer's protocol. After PKH26 staining, the exosomes were collected by Total Exosome Isolation Kit. Finally, PKH26‐labeled exosomes were resuspended in PBS.

Western Blot Analysis

Proteins were extracted using radio‐immune precipitation assay (P0013B; Beyotime Biotechnology, Shanghai, China) buffer supplemented with phenylmethylsulfonyl fluoride (ST506; Beyotime Biotechnology). Protein samples were fractionated by 10% or 12% SDS‐PAGE and were then transferred to nitrocellulose filter membranes. Membranes were blocked with 5% nonfat milk at room temperature for 1 hour. Anti–proliferating cell nuclear antigen (PCNA) antibody (1:1000, No. 2586, Cell Signaling Technology, Danvers, MA), anti–cyclin A antibody (1:1000, BM4674, Boster, Wuhan, China), anti–cyclin D antibody (1:500, A11022, ABclonal, Wuhan, China), anti‐PRKCA antibody (1:1000, 21991‐1‐AP, Proteintech, IL, USA), anti‐syndecan‐4 antibody (1:500, AB‐07‐3823, Erpantech Laboratory, Shanghai, China), anti‐EGR1 antibody (1:1000, #4153S, Cell Signaling Technology), anti–fibroblast growth factor 2 antibody (1:500, A0235, ABclonal, Wuhan, China), anti‐H3K27ac antibody (1:500, A7253, ABclonal), anti‐H3K4me1 antibody (1:500, A2355, ABclonal, Wuhan, China), anti–PDZ domain–containing protein 1 antibody (1:500, A10550, ABclonal), anti–hypoxia‐inducible factor (HIF)‐1α (1:1000, ab179483, Abcam, Cambridge, UK), anti‐HIF‐2α (1:1000, ab207607, Abcam), anti‐calnexin (1:500, A03372, Boster), and anti‐CD63 antibody (1:1000, ab315108, Abcam) were used as primary antibodies and were incubated overnight at 4 °C. The membranes were subsequently incubated with the appropriate horseradish peroxidase–conjugated secondary antibodies at room temperature for a duration of 1 hour. Subsequently, the membranes were treated with enhanced chemiluminescence reagent to facilitate the visualization of the antibody–antigen complexes. Following this, densitometric quantification was performed using Quantity One software (Bio‐Rad Laboratories, Hercules, CA) to accurately measure the intensity of the signals, enabling quantitative analysis of the immunoblotting results.

ELISA Assay

The levels of syndecan‐4 released in the plasms from human were measured with Human Syndecan‐4 ELISA Kit (EK1356, Boster) according to the manufacturer's protocol.

Cell‐Counting Kit‐8 Assay

Approximately 5000 cells/well were cultured in 96‐well plates according to the different experimental groups for 24 hours. Subsequently, 10 μL of Cell‐Counting Kit‐8 reagent was added to each well, and the plates were incubated at 37 °C for 2 to 4 hours; the absorbance at 450 nm was measured using a spectrophotometric microplate reader.

Plasmid and Small Interfering RNA Construction and Transfection

An overexpression vector for PRKCA was constructed using the vector GV486, and an empty vector alone was used as a negative control (Genechem, Shanghai, China). Small interfering RNA (siRNA) against syndecan‐4, and nontargeted control siRNA were designed and synthesized by GenePharma (Shanghai, China). The sequences are listed in Table S2. Cells were cultured to 70% to 80% confluence, and 2 μg siRNA or 3 μg plasmids and 10 μL transfection reagent (X‐tremeGene siRNA or Lipofectamine 2000) were diluted in 2 mL of serum‐free medium. Next, we incubated the mixture for 20 minutes and then added it directly to the cells. The transfection mixture was removed after 4 to 6 hours, and the cells were incubated for 24 hours and then used as required.

5‐Ethynyl‐2′‐Deoxyuridine Staining

Ethynyl‐2′‐deoxyuridine EdU staining was performed using a BeyoClick EdU Cell Proliferation Kit with Alexa Fluor 555 (C0075S; Beyotime, Shanghai, China) according to the manufacturer's protocol. In brief, cells in different groups were treated with 50 μmol/L of EdU and incubated for 2 hours at 37°C before fixation with 4% formaldehyde. Then the cells were exposed to 0.3% Triton X‐100 for 10 minutes. Nuclei were counterstained with 4′,6‐diamidino‐2‐phenylindole (5 μg/mL) for 10 minutes, after which the cells were observed under a live cell workstation (AF6000; Leica).

Cell Cycle and DNA Analysis

Cells were treated as indicated and then harvested and fixed using 70% ethanol at 4 °C for 24 hours. Following fixation, the ethanol solution was carefully removed, and the cells were stained using a Cell Cycle and Apoptosis Analysis Kit (C1052; Beyotime), strictly adhering to the manufacturer's instructions. Finally, the stained cells were analyzed using a FACSCalibur flow cytometer (BD Biosciences, Bedford, MA) to quantitatively assess the distribution of cells across different cell cycle stages.

Cell Coculture Assay

A noncontact transwell coculture model was used to evaluate the effect of PASMCs on PAECs. Well inserts (Costar, Corning, NY, USA) for 24‐ or 6‐well plates with a 0.4 μm/8 μm pore‐sized filter were used for the EdU and cell migration assays, respectively. Before starting the coculture experiments, PASMCs in 24‐ or 6‐well plates were transfected with negative control or syndecan‐4 siRNA, and PAECs were seeded into the well inserts. Then the inserts were subsequently placed in a 24‐ or 6‐well plate containing PASMCs to establish coculture conditions for the next 24 hours. A 1:1 mixture of endothelial cell medium : smooth muscle cell was used during all the coulture experiments.

Migration Assay

Cells were resuspended in serum‐free medium and cultured in the upper chamber of a transwell, which was inserted into 24‐well plates. Migration was measured after incubation in 4% paraformaldehyde for 10 minutes and then stained for 30 minutes with 0.4% crystal violet at room temperature. The number of migrated PAECs was counted under an inverted microscope (Nikon, Tokyo, Japan).

Coimmunoprecipitation

Cells were lysed in lysis buffer (Tris 50 mmol, pH 7.4; NaCl 150 mmol; Triton X‐100 1%; EDTA 1 mmol; and phenylmethylsulfonyl fluoride 2 mmol) and then centrifuged at 15 000 rpm for 30 minutes at 4 °C to collect the supernatant. Afterward, the supernatant incubated with 5 μg of the target antibody or IgG. Protein A+G agarose beads were added overnight at 4 °C. Antibody–protein complexes were washed 3 times with PBS, and the pellet was resuspended in protein loading buffer (2×) and subjected to Western blot.

Cell and Tissue Immunofluorescence Staining Analysis

Cells were cultured on coverslips in 12‐well plates and treated with different reagents. Twenty‐four hours after transfection, cells were fixed with 4% paraformaldehyde for 10 minutes at room temperature, permeabilized with 0.5% Triton X‐100 for 10 minutes at room temperature and blocked with 5% normal bovine serum for 15 minutes at room temperature. Cells were incubated with anti‐syndecan‐4 antibody (1:100, sc‐12766; Santa Cruz Biotechnology, Santa Cruz, CA), anti‐PRKCA (1:100, 21991‐1‐AP, Proteintech, Rosemont, IL), anti‐CD63 antibody (1:100, ab315108, Abcam), and anti‐Ki67 antibody (1:100, M00254‐3; Boster) at 4 °C overnight. The next day, cells were washed with PBS and incubated with Cy3‐conjugated goat anti‐rabbit antibody (1:50, A0516; Beyotime Biotechnology) or FITC‐conjugated goat anti‐mouse antibody (1:50, A0568; Beyotime Biotechnology) followed by 4′,6‐diamidino‐2‐phenylindole in the dark. Images were captured by live cell workstation. Frozen sectioning of mouse lung tissues was performed in the same manner. The sections were incubated with anti‐PCNA antibody (1:100, #2586; Cell Signaling Technology), anti–α‐smooth muscle actin antibody (1:100, #19245; Cell Signaling Technology), anti‐CD31 (1:200, 28083‐1‐AP; Proteintech) and anti‐PRKCA (1:100, 21991‐1‐AP; Proteintech). PCNA⁺ PASMCs were counted using the Image‐Pro Plus 6.0 software and were expressed as a percentage of the total number of α‐smooth muscle actin–positive PASMCs. For quantification of PRKCA, ImageJ was used to detect mean gray value, the mean gray value (mean)=integrated density (IntDen)/area. Approximately 5 arteries per animal were randomly examined under microscope at magnification ×200, and an average was calculated.

Chromatin Immunoprecipitation Sequencing and Chromatin Immunoprecipitation Quantitative PCR

Chromatin immunoprecipitation (ChIP) sequencing was performed by Guangzhou Epibiotek Co., Ltd. The bigwig files of H3K27ac were visualized via Integrative Genomics Viewer and WashU Epigenome Browser. ChIP assays were performed using ChIP Assay Kits (P2078, Beyotime Biotechnology) according to the manufacturer's instructions. Briefly, cells were cross‐linked with 1% formaldehyde and quenched by 0.125 M glycine, followed by ultrasonic shear to obtain chromatin fragments (100–500 bp). The chromatin fragments were incubated with magnetic beads combined with anti‐H3K27ac antibody (1:100, A7253; ABclonal), anti‐H3K4me1 antibody (1:100, A2355; ABclonal) and anti‐EGR1 antibody (1:1000, #4153S; Cell Signaling Technology) at 4 °C overnight. Finally, the immunoprecipitated DNA was eluted, purified, and detected by quantitative reverse transcription PCR. The primers sequences used are listed in Table S2.

Statistical Analysis

Statistical analysis was performed using Prism 8.0 (GraphPad Software Inc., La Jolla, CA). All data collection and processing in this study are expressed as mean ±SEM. Before statistical testing, the normality of the data was rigorously evaluated using the Shapiro–Wilk test, and the equality of group variance was assessed through the Brown–Forsythe test. Student's t test was used to compare the data between 2 groups with equal variance, and the Welch correction test was used for 2‐group analysis with unequal variance. One‐way or 2‐way ANOVA with Tukey's post hoc test was used to compare multiple groups with equal variance, and Brown–Forsythe and Welch ANOVA with Tamhane T2 post hoc test was used to compare multiple groups with unequal variance. For nonnormally distributed data, we performed nonparametric analyses such as the Mann–Whitney U test for 2 groups or the Kruskal–Wallis test followed by the Dunn posttest for multiple groups. A 2‐tailed P<0.05 was considered to indicate statistical significance.

Results

Identification of Syndecan‐4 as a Novel SE‐Driven Gene in PH

To characterize SE‐associated genes in PH, we first performed ChIP sequencing with antibody against H3K27ac in hypoxic hPASMCs and found that the syndecan‐4 transcript was marked by the epigenetic signature of active enhancers H3K27ac signaling peaks (Figure 1A). To determine whether SE activated the expression of syndecan‐4, we treated hPASMCs with BRD4 (master regulator of SEs) inhibitor, JQ‐1, and found that JQ‐1 specifically inhibited the transcription of SE‐associated syndecan‐4 compared with hypoxia (Figure 1B). Moreover, the specifical inhibition of JQ1 on the expression of BRD4 was determined in hPASMCs and the expression of HIF‐1α was detected as a quality control for hypoxia (Figure S1A). We detected the expression of syndecan‐4, HIF‐1α, and HIF‐2α (a quality control for hypoxia endothelial cells) using Western blot analysis, the results demonstrated that the expression of syndecan‐4 were increased in hPASMCs under hypoxic conditions but not in human hPAECs (Figure 1C and 1D). Notably, due to syndecan‐4 was soluble‐related protein, ²³ we measured syndecan‐4 expression by using ELISA, and found that the syndecan‐4 concentration was higher in plasma of patients with PH compared with healthy donors (Figure 1E). Importantly, syndecan‐4 has high homology in humans and mice, and syndecan‐4 was also upregulated in hypoxic lung tissues and mPASMCs (Figure S1B, Figure 1F and 1G). Quantitative reverse transcription PCR result showed that JQ‐1 specifically inhibited the expression of syndecan‐4 in hypoxic mPASMCs (Figure 1H). The above results demonstrate that the upregulated syndecan‐4 in hypoxia is a novel SE‐driven gene and may be a key regulator involved in hypoxic PH.

Figure 1. Syndecan‐4 was an SE‐driven gene in hypoxia.

A, Gene tracks depicting the SE region of syndecan‐4 in hPASMCs by performing ChIP‐seq with antibody against H3K27ac. B, JQ1 specifically diminished the expression of syndecan‐4 in hypoxic hPASMCs (n=5). C, Western blot analysis was used to verify the expression of syndecan‐4 and HIF‐1α in hypoxic hPASMCs (n=6). D, Western blot analysis was used to verify the expression of syndecan‐4 and HIF‐2α in hypoxic human PAECs (n=6). E, ELISA assay was used to detect the expression of syndecan‐4 in plasma of patients with PH (n=10). (F) Western blot analysis was used to verify the expression of syndecan‐4 and HIF‐1α in mouse lung tissues (n=6). (G) Western blot analysis was used to verify the expression of syndecan‐4 and HIF‐1α in hypoxic mPASMCs (n=6). (H) JQ‐1 specifically diminished the expression of syndecan‐4 in hypoxic mPASMCs (n=5). All values are presented as the mean±SEM. Statistical analysis was performed with one‐way ANOVA or Student's t‐test. *P<0.05, **P<0.01, ***P<0.001. ChIP‐seq indicates chromatin immunoprecipitation sequencing; hPAECs,  human pulmonary artery endothelial cells; hPASMCs, human pulmonary artery smooth muscle cells; HYP, hypoxic; mPASMCs, mouse pulmonary artery smooth muscle cells; NOR, normoxia; and SDC4, syndecan‐4.

SE Regions of Syndecan‐4 Is Occupied by Transcription Factor EGR1 in Hypoxia

Consistent with the ChIP sequencing results, we found that the syndecan‐4 transcript was marked by the monomethyl H3K4 (H3K4me1) and H3K27ac signaling peaks in the SE region of human lung tissue and primary lung cells through analysis of the WashU Epigenome Browser databases. We further divided the SE region of syndecan‐4 into 4 constituents (E1–E4) according to joint analysis of the ChIP sequencing and WashU Epigenome Browser databases (Figure 2A). As SEs stimulate transcription across large genomic distances by binding transcription factors and coactivators, we identified candidate transcription factors that could bind to both of SE region and promoter region of syndecan‐4 by using SE databases (SEanalysis and SEdb) and promoter databases (JASPAR and AnimalTFDB). The transcription factors EGR1 and interferon regulatory factor 1 were noted, and with more binding sites of EGR1 occupied in syndecan‐4 promoter (Figure 2B and 2C). Western blot results showed that expression level of EGR1 was upregulated in hypoxic hPASMCs (Figure 2D). The coimmunoprecipitation assay results verified the interaction of EGR1 with H3K27ac and H3K4me1 (Figure S1C).

Figure 2. SE‐associated syndecan‐4 was transcriptionally activated by EGR1 in hPASMCs.

A, Four constituents (E1–E4) of SE region of syndecan‐4. B, EGR1 and IRF1 were identified as the candidate transcription factors. C, Representative motif of EGR1 and IRF1 binding sites. D, Western blot analysis was used to verify the expression of EGR1 and HIF‐1α in hypoxic hPASMCs (n=6). E and F, hPASMCs were subjected to ChIP analysis using antibodies against H3K27ac, H3K4me1, and EGR1. The association with the SE region (E1–E4) of syndecan‐4 was quantified by quantitative reverse transcription PCR (n=3). G and H, hPASMCs were subjected to ChIP analysis using antibodies against H3K27ac, H3K4me1 and EGR1. The association with the promoter region (P1–P3) of syndecan‐4 was quantified by quantitative reverse transcription PCR (n=3). (I) Silencing of EGR1 significantly decreased the expression of EGR1 and syndecan‐4 in hypoxic hPASMCs (n=6). J and K, hPASMCs were treated with EGR1 small interfering RNA and subjected to ChIP analysis using antibodies against H3K27ac. The association with the E2 SE (J) and P2 promoter (K) regions of syndecan‐4 was quantified by quantitative reverse transcription PCR (n=3–4). (L) Schematic diagram of transcribing syndecan‐4 in hPASMCs. All values are presented as the mean±SEM. Statistical analysis was performed with 1‐way ANOVA or Student's t test. *P<0.05, **P<0.01, ***P<0.001. ChIP indicates chromatin immunoprecipitation; EGR1, early growth response 1; hPASMCs, human pulmonary artery smooth muscle cells; HYP, hypoxic; IRF1, interferon regulatory factor 1; NC, negative control; NOR, normoxia; and SDC4, syndecan‐4.

To clarify the direct regulatory effects of H3K27ac, H3K4me1, and EGR1 on syndecan‐4 at the transcriptional level through the SE, ChIP–quantivative PCR assays revealed that the binding of the H3K27ac to the individual constituent E1 to E4 of syndecan‐4 and H3K4me1 was enriched in E2 and E3 of syndecan‐4 in hPASMCs (Figure 2E). In addition, hypoxia promoted the association between EGR1 and E2 of syndecan‐4 in hypoxia (Figure 2F). The above results indicating E2 as a regulation region of the syndecan‐4 SE in hypoxia.

To further validate the binding of H3K27ac, H3K4me1 and EGR1 on syndecan‐4 promoter, we divided the syndecan‐4 promoter into 2 equal segments (P1–P2). We performed ChIP with an antibody against H3K27ac and H3K4me1 followed by quantitative PCR and observed that H3K27ac and H3K4me1 enrichment at the promoter regions of syndecan‐4 in hPASMCs (Figure 2G). Furthermore, hypoxia increased EGR1 enrichment at the P2 promoter regions of syndecan‐4 in hPASMCs (Figure 2H). Meanwhile, hPASMCs treatment with EGR1 siRNA (Figure S1D) counteracted hypoxia‐mediated upregulation of EGR1 and syndecan‐4 (Figure 2I). Similar results were observed in cells that EGR1 loss decreased H3K27ac enrichment at the P2 and E2 regions of syndecan‐4 (Figure 2J and 2K). Taken together, these data demonstrate that EGR1 is recruited by the SE upstream E2 region and acts on the promoter P2 region of syndecan‐4, forming a chromatin loop to enhance its transcription in hypoxia hPASMCs (Figure 2L).

Moreover, syndecan‐4 transcript was marked by H3K27ac and H3K4me1 signaling peaks in the SE regions of mouse lung tissue via WashU Epigenome Browser databases analysis (Figure S2A). We further confirmed the specific interaction of H3K27ac, H3K4me1, and EGR1 on SE and promoter regions of syndecan‐4 by ChIP–quantitative PCR assays. The results demonstrate that H3K27ac, H3K4me1, and EGR1 interact with both the E1 and P1 regions of syndecan‐4 to enhance its transcription in hypoxic mPASMCs (Figure S2B through S2F). Collectively, these findings identify SE‐associated syndecan‐4 as a downstream target regulated by EGR1, which are recurrently coamplified in hypoxia‐induced PH.

Silencing of Syndecan‐4 Inhibits Hypoxia‐Induced PASMC Proliferation

To explore the function of syndecan‐4 in hypoxic PH, we transfected syndecan‐4 siRNA into hPASMCs and the Western blot results showed that siRNA‐3 effectively inhibited syndecan‐4 expression (Figure S3A). EdU, Ki67, and Cell‐Counting Kit‐8 assays were performed and showed that syndecan‐4 silencing retarded hypoxia‐induced proliferation of hPASMCs (Figure 3A and 3B). The effects of syndecan‐4 on cell cycle progression were verified by flow cytometry, in which knockdown of syndecan‐4 reversed the increasing percentage of cells in S+G2/M phase promoted by hypoxia (Figure 3C). To determine whether there were the off‐target effects, we have examined the expression of syndecan‐4 using Western blot. The results showed the success of syndecan‐4 knockdown under hypoxic conditions (Figure 3D). In addition, Western blot analysis showed that silencing of syndecan‐4 abolished hypoxia‐upregulated expressions of PCNA and cell cycle–related proteins (cyclin A and cyclin D) (Figure 3D). Similarly, hypoxia‐induced cell proliferation in mPASMCs were abrogated by syndecan‐4 knockdown (Figure S3B, Figure 3E through 3G). Taken together, these findings imply that SE droven‐syndecan‐4 play a key role in controlling hypoxia‐induced PASMC proliferation.

Figure 3. Silencing of syndecan‐4 inhibited hypoxia‐induced PASMC proliferation.

A, Silencing of syndecan‐4 blocked the effects of hypoxia on EdU incorporation and Ki67 expression in hPASMCs, EdU (red), Ki67 (green), 4′,6‐diamidino‐2‐phenylindole (blue). Scale bar, 50 μm. B, hPASMCs viability was determined by Cell‐Counting Kit‐8 assay (n=6). C, Cell cycle analysis by flow cytometry indicated that hypoxia stimulated cell progression into G2/M+S phase, and this effect was inhibited by syndecan‐4 small interfering RNA. D, Western blot analysis of syndecan‐4, HIF‐1α, PCNA, cyclin A, and cyclin D in hPASMCs (n=6). E, mPASMC viability was determined by Cell‐Counting Kit‐8 assay (n=6). F, Silencing of syndecan‐4 blocked the effects of hypoxia on EdU incorporation in mPASMCs. EdU (red), 4′,6‐diamidino‐2‐phenylindole (blue). Scale bar, 50 μm (n=6). G, Western blot analysis of syndecan‐4, HIF‐1α, PCNA, cyclin A, and cyclin D in mPASMCs (n=6). All values are presented as the mean±SEM. Statistical analysis was performed with 1‐way ANOVA. *P<0.05, **P<0.01, ***P<0.001. EdU indicates ethynyl‐2′‐deoxyuridine; hPASMCs, human pulmonary artery smooth muscle cells; HYP, hypoxic; mPASMCs, mouse pulmonary artery smooth muscle cells; NC, negative control; NOR, normoxia; PASMC, pulmonary artery smooth muscle cell; PCNA, proliferating cell nuclear antigen; and SDC4, syndecan‐4.

Syndecan‐4 Regulates PASMC Proliferation Via PRKCA

To gain a better understanding of the mechanisms underlying syndecan‐4 mediated hypoxia‐induced PASMC proliferation, we screened and predicted target proteins (fibroblast growth factor 2, PRKCA and PDZ domain–containing protein 1) interacted with syndecan‐4 on the basis of bioinformatics (STRING, GENEMANIA, and inBio Discover) (Figure 4A). Then, Western blot experiments were performed and showed that only the expression of PRKCA was increased in hypoxia, which was decreased by sisyndecan‐4 in hPASMCs (Figure 4B), and the same results were found in mPASMCs (Figure S4A). To confirm the interaction between syndecan‐4 and PRKCA, coimmunoprecipitation experiments were performed and revealed that syndecan‐4 directly bound to PRKCA (Figure 4C, Figure S4B). Consistent with these results, immunofluorescence assay showed that syndecan‐4 colocalized with PRKCA both in hPASMCs and mPASMCs (Figure 4D, Figure S4C). Finally, we predicted and visualized the 3‐dimensional structural docking of syndecan‐4–PRKCA, the result demonstrated that syndecan‐4 and PRKCA have multiple binding sites (Figure 4E). These data suggest that PRKCA is likely involved in the regulation of syndecan‐4 in PASMCs.

Figure 4. PRKCA is involved in syndecan‐4‐mediated hPASMC proliferation.

A, FGF2, PRKCA, and GIPC1 were identified as the candidate target proteins. B, Cells were transfected with syndecan‐4 siRNA, and the protein expression of syndecan‐4, HIF‐1α, FGF2, PRKCA, and GIPC1 was estimated (n=6). C, Coimmunoprecipitation assay verified the interaction of syndecan‐4 and PRKCA. D, Syndecan‐4 and PRKCA were colocalized in the cytoplasm. Scale bar, 50 μm. E, Schrodinger2019.01 software predicted and visualized the 3‐dimensional structural docking of syndecan‐4 (magenta) and PRKCA (cyan). Yellow dash represents hydrogen bond and salt bridge interaction. F, Online websites were used to predict PRKCA binding to E3 ubiquitin ligases (http://ubibrowser.ncpsb.org/). G, Effect of syndecan‐4 small interfering RNA on PRKCA ubiquitination level. H, Cells were treated with syndecan‐4 siRNA and proteasome inhibitor MG132, and the expression levels of syndecan‐4, HIF‐1α and PRKCA were examined (n=5). I and J, Cell‐Counting Kit‐8 and EdU incorporation assays showed that overexpression of PRKCA abolished the effect of syndecan‐4 deficiency on cell viability and proliferation in hypoxic hPASMCs. EdU (red), 4′,6‐diamidino‐2‐phenylindole (blue). Scale bar, 50 μm (n=6). K, Western blot analysis showing that overexpression of PRKCA attenuated the effect of syndecan‐4 siRNA on protein expression of syndecan‐4 and PCNA (n=5). (L) Cell cycle analysis by flow cytometry indicated that overexpression of PRKCA attenuated the effect of syndecan‐4 siRNA on cell cycle. All values are presented as the mean±SEM. Statistical analysis was performed with 1‐way ANOVA. *P<0.05, **P<0.01, ***P<0.001. FGF2 indicates fibroblast growth factor 2; GIPC1, PDZ domain–containing protein 1; HYP indicates hypoxic; NC, negative control; NOR, normoxia; PRKCA, protein kinase C α; and SDC4, syndecan‐4.

The significantly downregulated protein expression of PRKCA in PASMCs with syndecan‐4 knockdown suggested that syndecan‐4 might be involved in PRKCA protein degradation. We found that PRKCA can serve as substrate for a variety of E3 ubiquitin ligases through UbiBrowser (http://ubibrowser.ncpsb.org/) prediction (Figure 4F). Subsequently, we performed ubiquitination analysis and the coimmunoprecipitation result demonstrated that hypoxia significantly decreased ubiquitination level of PRKCA, knockdown of syndecan‐4 reversed this effect (Figure 4G). Furthermore, Western blot showed that silencing of syndecan‐4 suppressed the protein levels of PRKCA induced by hypoxia, whereas the ubiquitin–protease system inhibitor MG‐132 reversed these trends (Figure 4H). These results suggest that syndecan‐4 interacted with the PRKCA protein and inhibited the degradation of PRKCA through the ubiquitin–proteasome pathway.

Next, we attempted to gain insight into the functional recovery experiments to determine whether PRKCA is involved in syndecan‐4–mediated hPASMC proliferation. PRKCA protein expression was significantly upregulated after transfection of the PRKCA overexpression plasmid into hPASMCs (Figure S3C). Cell‐Counting Kit‐8 and EdU assays showed that silencing of syndecan‐4 partially suppressed hypoxia‐induced proliferation of hPASMCs, whereas PRKCA overexpression reversed this effect (Figure 4I and 4J). Additionally, hPASMC transfection with si‐syndecan‐4 decreased the expression of the PCNA protein and cell cycle progression under hypoxia, an effect that was reversed by overexpression with PRKCA (Figure 4K and 4L). The above results suggest that PRKCA, a downstream target of syndecan‐4, is involved in syndecan‐4–mediated PASMC proliferation.

PASMC‐Derived Syndecan‐4 Promotes PAEC Dysfunction

Syndecan‐4 has been reported to be associated with exosomes. ²⁴ To explore the transport capacity of syndecan‐4 in PH, we isolated extracellular vesicles and transmission electron microscope revealed the “cup‐shaped” vesicles of exosomes with a median diameter of 70 nm, indicating that most of the isolated vesicles comprise exosomes (Figure 5A). The expression of syndecan‐4, CD63 (a positive marker of exosomes), and CANX (a negative marker of exosomes) were observed using Western blot, the results confirmed the syndecan‐4 in exosomes secreted by PASMCs (Figure 5B). Immunofluorescence staining revealed the colocalizations of syndecan‐4 and CD63 in PASMCs (Figure 5C). These results showed that syndecan‐4 was increased in exosomes under hypoxia. Knockdown of the syndecan‐4 led to a decrease in the level of syndecan‐4 in the exosomes secreted by hPASMCs under hypoxic conditions (Figure S5A). We further examined the intercellular transport of exosomes, and as shown in Figure 5D, PAECs exhibited efficient uptake of the exosomes derived from PASMCs.

Figure 5. PASMC‐derived syndecan‐4 promotes PAEC dysfunction.

A, Exosomes extracted from hPASMCs were prepared for electron microscopy assay, and the particle size of the vesicles was measured by NanoSight analysis. Scale bar, 100 nm. B, The expression levels of CD63, syndecan‐4, CANX and β‐Actin in the cell lyses and exosomes. C, syndecan‐4 and CD63 were colocalized. Scale bar, 50 μm. D, Exosomes from PASMCs were labeled with PKH26 and then incubated PAECs. Scale bar, 50 μm. E, Schematic of the transwell coculture model. F, The expression levels of PRKCA in transwell coculture model (n=4). G, EdU incorporation in transwell coculture model (n=6). Scale bar, 50 μm. H, Cell migration assay in transwell coculture model (n=6). Scale bar, 100 μm. All values are presented as the mean±SEM. Statistical analysis was performed with 2‐way ANOVA. *P<0.05, **P<0.01, ***P<0.001. EdU indicates ethynyl‐2′‐deoxyuridine; Exo, exosomes; hPASMCs, human pulmonary artery smooth muscle cells; HYP, hypoxic; NC, negative control; NC, negative control; NOR, normoxia; PAEC, pulmonary artery endothelial cell; PASMC, pulmonary artery smooth muscle cell; and SDC4, syndecan‐4.

Moreover, we constructed a noncontact transwell co‐culture system (Figure 5E). The expression of syndecan‐4 and PRKCA were increased in PAECs when PASMCs were cocultured with PAECs (Figure S5B and S5C). Western blot experiments were performed and showed that knockdown of syndecan‐4 in PASMCs decrease the protein levels of PRKCA when PASMCs were cocultured with PAECs under either normoxia or hypoxia (Figure 5F). We further studied whether PASMC‐secreted syndecan‐4 plays a role in PAEC proliferation and migration. EdU staining showed that PASMC‐derived exosomes induced PAEC proliferation, but this effect was inhibited when syndecan‐4 was silenced or by GW4869 (exosome inhibitor) administration in PASMCs (Figure 5G). In addition, migration assay suggested that knockdown of syndecan‐4 or GW4869 inhibited PAEC migration when PASMCs were cocultured with PAECs under either normoxia or hypoxia (Figure 5H). These results suggest that PASMC‐derived syndecan‐4 mediates endothelial function through a mechanism dependent on paracrine associated with exosomes.

Inhibition of Syndecan‐4 Reverses SU5416 Combined With Hypoxia–Induced PH

To determine the role of syndecan‐4 in hypoxic PH in vivo, we constructed a SU5416 combined with hypoxia (SuHx)–PH model and used AAV5 to interfere with syndecan‐4 (Figure 6A). The infection efficiency of AAV5 within the lungs is shown in the Figure S6A. Quantitative reverse transcription PCR and Western blot analysis confirmed that syndecan‐4 was effective knockdown in lung tissues infected with AAV5 (Figure S6B and S6C). To assess the effect of syndecan‐4 silencing on SuHx‐induced PH, we evaluated right ventricular systolic pressure and right ventricle‐to‐left ventricle plus septum ratio. The results showed that AVV5 loaded syndecan‐4 knockdown inhibited the above PH index induced by SuHx (Figure 6B and 6C). Correspondingly, echocardiography showed that inhibiting of syndecan‐4 significantly improved the impaired pulmonary artery acceleration time and pulmonary artery velocity time integral induced by SuHx treatment with no differences in left ventricular ejection fraction and heart rate (Figure 6D). Furthermore, morphological and immunofluorescence staining for PCNA assays demonstrated that knockdown of syndecan‐4 reversed SuHx‐induced distal pulmonary vascular remodeling and proliferation (Figure 6E and 6F; Figure S6D and S6E). Besides, knockdown of syndecan‐4 prevented increases in PRKCA expression in SuHx‐treated mice (Figure 6G and 6H; Figure S6F). The above results suggested that syndecan‐4 plays an important role in contributing to the development of PH associated with hypoxia.

Figure 6. Inhibition of syndecan‐4 reverses SuHx‐induced PH progression.

A, Schematic illustration showing the construction of mice AAV5 and the treatment protocol. B and C, Right ventricular systolic pressure and right ventricular/left ventricular+septum weight ratio in the SuHx‐induced PH mice models (n=7–8). D, Pulmonary artery velocity time integral, pulmonary artery acceleration time, left ventricular ejection fraction and heart rate of the SuHx‐induced PH mice models infected with AAV5‐NC and AAV5‐syndecan‐4 (n=7–8). E, Immunofluorescence of PCNA and α‐smooth muscle actin in mouse lung sections. Scale bars, 50 μm. F, Quantification of PCNA‐positive α‐smooth muscle actin–positive cells in (E) (n=7–8). G, Immunofluorescence of PRKCA and α‐smooth muscle actin in mouse lung sections. Scale bars, 50 μm. H, Quantification of PRKCA expression in (G) (n=7–8). All values are presented as the mean±SEM. Statistical analysis was performed with one‐way ANOVA. ***P<0.001. AAV5 indicates serotype 5 adenovirus‐associated virus; HYP, hypoxic; NC, serotype 5 adenovirus‐associated virus–negative control; NOR, normoxia; PCNA, proliferating cell nuclear antigen; PH, pulmonary hypertension; PRKCA, protein kinase C α; and SuHx, SU5416 combined with hypoxia.

Discussion

In this study, we defined that syndecan‐4, driven by SE, is upregulated in plasma of patients with PH as well as lung tissues and cells from hypoxic PH models. Both in vitro and in vivo experiments demonstrated a novel function of syndecan‐4 in regulating of PASMCs and PAECs dysfunction related to hypoxic pulmonary vascular remodeling. As shown in Figure 7. syndecan‐4 combined with PRKCA to protect it from ubiquitination degradation, leading to PASMC proliferation. Furthermore, syndecan‐4 transported by exosomes mediated intercellular communication between PASMCs and PAECs, contributing to proliferation and migration of PAECs in hypoxia. Our data suggest that syndecan‐4 plays a crucial role in PH pathogenesis and serves as a potential target for treatment and diagnosis of PH.

Figure 7. Proposed mechanism for the role of syndecan‐4 in vascular remodeling.

Transcription factor EGR1 directly bound SE and promoter region of syndecan‐4, and activated the transcription of syndecan‐4 in hypoxic PASMCs. On the one hand, syndecan‐4 interacted with protein kinase C alpha to regulate PRKCA ubiquitination, which leading to PASMC proliferation. On the other hand, PASMCs secrete exosomes enriched with syndecan‐4, which are transported to PAECs, inducing PAEC proliferation and migration. Finally, PASMC proliferation and PAEC dysfunction regulated by SE‐driven syndecan‐4 promote vascular remodeling and PH. EGR1 indicates early growth response 1; HYP, hypoxic; PAEC, pulmonary artery endothelial cell; PASMC, pulmonary artery smooth muscle cell; PH, pulmonary hypertension; PRKCA, protein kinase C α; SDC4, syndecan‐4; and Ubb, ubiquitin B.

SEs are important transcriptional regulation elements that can bind to H3K4me1, H3K27ac, and transcription factors to drive gene expression. ²⁵ More important, SE‐associated proteins and noncoding RNAs regulate cardiovascular disease and tumor pathogenesis. ²⁶ , ²⁷ , ²⁸ , ²⁹ Indeed, SE inhibitor JQ‐1 has been proven to suppress proliferation in breast cancer, endometrial cancer, and prostate cancer, which indicates that SEs could be one of the promising therapeutic targets for diseases treatment. ³⁰ , ³¹ , ³² In this study, we first analyzed the profiles of SEs by ChIP sequencing with antibody against H3K27ac in hPASMCs and identified the novel SE‐associated syndecan‐4. Moreover, we revealed that H3K27ac, H3K4me1, and EGR1 occupy the promoter of syndecan‐4 and activate its transcription by ChIP–quantitative PCR experiments. The present study comprehensively screened SE‐associated genes from the point of hypoxic PH and identified a new upstream mechanism by which SE‐driven syndecan‐4 expression in PASMCs. Moreover, we revealed for the first time that syndecan‐4 expression was abnormally higher in the plasma of patients with PH and PASMC‐derived exosomes in hypoxia. Knockdown of syndecan‐4 significantly reversed hypoxia‐induced vascular remodeling by regulating PASMC proliferation and the intercellular communication with PAECs. Thus, our findings suggest that SE‐driven syndecan‐4 is a vital regulator in PH. However, further studies are needed to determine the specific pathway of syndecan‐4 involved in the regulation of proliferation, such as glycolysis, endoplasmic reticulum stress, and mitochondrial dysfunction, which will provide crucial insight into the contribution of the SE‐associated gene in complex pathological process.

More importantly, our results from the ELISA analysis revealed a significant upregulation of syndecan‐4 in the plasma of patients with PH, suggest that syndecan‐4 may be a potential biomarker for hypoxic PH. Undeniably, this possibility still needs to be further verified by a large number of samples from patients with PH, given the multiple subtypes of PH, whether syndecan‐4 play the same roles in all the PH subtypes remains to be elucidated.

It has been reported that exosomes regulate PASMC and PAEC crosstalk in the pulmonary vascular microenvironment. For instance, exosomes secreted from PASMCs mediate the transfer of miR‐143/145 from PASMCs to PAECs in a paracrine manner, inducing migration and angiogenesis in PH. ³³ Exosomes secreted from SOX17‐overexpressing PAECs regulate endothelial function and pulmonary vascular homeostasis by releasing miR‐224‐5p and miR‐361‐3p. ³⁴ The present study revealed that syndecan‐4 was incorporated into exosomes of PASMCs and acts on the PAECs, contributing to proliferation and migration. We provide the first evidence that syndecan‐4 acts as a paracrine signaling mediator and is involved in the regulation of pathogenesis and development of PH. Nevertheless, because the lack of exosome‐deficient or syndecan‐4 knockout mice, we have not been able to clarify the proposed mechanism about syndecan‐4 transported by exosomes and mediated intercellular communication in‐depth. Moreover, another limitation of this study is that the mode of syndecan‐4 loading into exosomes and its receptors or ligands are not clarified, which is worthy of further study.

As an important protein kinase, PRKCA has been shown to regulate multiple signaling pathways, which are abnormally active in cancer, promoting tumor progression via proliferation, migration, and invasion. ³⁵ , ³⁶ In addition, hypoxia can activate PRKCA to participate in autophagy and proliferation in colorectal cancer cells, suggesting a possible relationship between PRKCA and PH. ³⁷ In this study, we identified that syndecan‐4 tends to interact with PRKCA and promotes its expression. PRKCA was predicted to function as substrate for a variety of E3 ubiquitin ligases. Thus, we hypothesized that syndecan‐4 could affect the ubiquitination of PRKCA. According to our results, knockdown of syndecan‐4 increased the ubiquitination level of PRKCA. Our results unraveled that syndecan‐4 interacts with PRKCA, inhibiting the degradation of PRKCA by proteasome. Although the specific protein interaction domain and ubiquitination site of syndecan‐4 and PRKCA are not revealed, this study provides a new direction for posttranslational modification of PRKCA involved in PASMC proliferation.

To summarize, we demonstrate for the first time that SE‐driven syndecan‐4 has a critical regulatory role in PASMC proliferation and intercellular communication with PAECs via the PRKCA and exosome pathway. These findings suggest that syndecan‐4 holds the potential to emerge as a novel therapeutic target and may serve as a biomarker for the diagnosis of PH.

Sources of Funding

This study was supported by the National Natural Science Foundation of China (82170059 and 32400949), Natural Science Foundation of Heilongjiang Province (ZD2023H003), Natural Science Foundation of Chongqing (cstc2021jcyj‐msxmX0474).

Disclosures

None.

Supporting information

Data S1