Heterogeneous nuclear ribonucleoprotein U-actin complex derived from extracellular vesicles facilitates proliferation and migration of human coronary artery endothelial cells by promoting RNA polymerase II transcription.

Coronary artery disease (CAD) represents a fatal public threat. The involvement of extracellular vesicles (EVs) in CAD has been documented. This study explored the regulation of embryonic stem cells (ESCs)-derived EVs-hnRNPU-actin complex in human coronary artery endothelial cell (HCAEC) growth. Firstly, in vitro HCAEC hypoxia models were established. EVs were extracted from ESCs by ultracentrifugation. HCAECs were treated with EVs and si-VEGF for 24 h under hypoxia, followed by assessment of cell proliferation, apoptosis, migration, and tube formation. Uptake of EVs by HCAECs was testified. Additionally, hnRNPU, VEGF, and RNA Pol II levels were determined using Western blotting and CHIP assays. Interaction between hnRNPU and actin was evaluated by Co-immunoprecipitation assay. HCAEC viability and proliferation were lowered, apoptosis was enhanced, wound fusion was decreased, and the number of tubular capillary structures was reduced under hypoxia, whereas ESC-EVs treatment counteracted these effects. Moreover, EVs transferred hnRNPU into HCAECs. EVs-hnRNPU-actin complex increased RNA Pol II level on the VEGF gene promoter and promoted VEGF expression in HCAECs. Inhibition of hnRNPU or VEGF both annulled the promotion of EVs on HCAEC growth. Collectively, ESC-EVs-hnRNPU-actin increased RNA Pol II phosphorylation and VEGF expression, thus promoting HCAEC growth.

Highlights

ESC-EVs promote HCAEC proliferation, migration and angiogenesis.

EVs carry hnRNPU into HCAECs.

Inhibition of hnRNPU partially annuls the promotion of EVs on HCAEC growth.

EVs-hnRNPU-actin complex increases RNA Pol II level on the VEGF gene promoter.

EVs-hnRNPU elevates VEGF levels under hypoxia.

Introduction

Coronary artery disease (CAD) remains a prime contributor to global morbidity and mortality, which arises due to chronic inflammation and cholesterol accumulation on coronary arterial walls, causing atherosclerotic plaque deposits and narrowing of coronary arteries, ultimately resulting in the reduction of the oxygen-rich blood [1-3]. CAD accounts for approximately 9 million global deaths in 2017 [4] and contributes to one-third or more of all deaths in individuals over 35 [5]. Despite developments in prevention strategies and medical treatments, the prevalence in developing countries is still increasing [6]. Vascular endothelial activation caused by cardiovascular risk factors results in the generation of cytokines, chemokines, and adhesion molecules, which interact with leukocytes, ultimately leading to inflammation [7,8]. Endothelial dysfunction, injury, and repair are pivotal in cardiovascular diseases (CVs) [9,10]. The limited EC proliferation and migration and elevated EC apoptosis are implicated in the initiation of atherosclerosis [11]. Herein, this study probed into the molecular mechanism of human coronary artery endothelial cell (HCAEC) growth to exert a reference for CAD therapy.

Recently, extracellular vesicles (EVs) have been perceived as pivotal players in CVs [12,13]. EVs transfer bioactive molecules, including peptides, proteins, nucleic acids, and lipids, which can be carried into recipient cells, thereby affecting cell biological behaviors [14,15]. EVs isolated from diverse stem cells are applied for cardiac repair, including embryonic stem cells (ESCs), mesenchymal stem cells, cardiac progenitor cells, and induced pluripotent stem cells [16-19]. Previous research has evidenced that ESCs-derived exosomes promote endogenous repair and enhance cardiac function after myocardial infarction [17]and also mitigate heart failure caused by transverse aortic constriction by facilitating angiogenesis [20].

On a separate note, existing evidence has unveiled that heterogeneous nuclear ribonucleoprotein U (hnRNPU) is abundantly present in EVs derived from ECs [21]. hnRNPs comprise RNA-binding proteins that are vital for nucleic acid metabolism and primarily expressed in the nucleus [22]. hnRNPU, also acknowledged as scaffold attachment factor A, is the largest component of this complex and is expressed in the fetal brain, adult heart, kidney, liver, cerebrum, and cerebellum, which has crucial functions in mammalian development [21,23,24]. Compelling evidence supports that hnRNPU is necessitated for postnatal heart development and functions, and mice lacking hnRNPU exhibit lethal dilated cardiomyopathy, disorganized cardiomyocytes, aberrant excitation-contraction coupling activity, and impaired contractility [25]. Based on aforesaid findings, we reasonably speculated that EVs presumably influence the proliferation and migration of HCAECs by carrying hnRNPU.

hnRNPU can bind to miRNAs to affect EC proliferation, promote the expression levels of angiogenic genes [26] and bind to different transcription factors to regulate the transcription process [27]. Additionally, hnRNPU binding to actin facilitates the transcription of RNA polymerase II (RNA Pol II) [28]. Actin, as an imperative regulator in RNA Pol II transcription, acts to recruit RNA Pol II activators during elongation of nascent transcripts when in complex with specific hnRNPs, and hnRNPU-actin complex cooperates with PCAF to regulate transcription elongation of RNA Pol II [29]. Gene transcription by RNA Pol II is the first step of gene expression in eukaryotes and the focus of regulation during cell development, differentiation, and response to the environment [30]. Pol II transcribes all protein-coding genes and many non-coding RNAs in eukaryotic genomes [31]. Furthermore, RNA Pol III subunit A is discordantly expressed in CAD individuals [32]. However, the mechanism of the hnRNPU-actin complex in regulating RNA Pol II transcription is rarely studied in HCAEC growth.

Human RNA Pol II seventh subunit can induce vascular endothelial growth factor (VEGF) promoter transactivation, mRNA expression, and VEGF protein secretion [33]. VEGF is a mitogen that promotes the proliferation and neovascularization of vascular ECs, and also a glycoprotein secreted by ECs and smooth muscle cells in vessel walls [34]. In particular, upregulation of VEGF-A expression promotes the proliferation of islet microvascular endothelial cells and suppresses apoptosis and oxidative stress [35]. Interestingly, miR-934 deficiency prominently reduces VEGF expression in SW480 and HCT116 cells and represses the ability of colorectal cancer cells to facilitate tube formation in vascular endothelial cells [36]. VEGF gene expression is prominently downregulated in CAD individuals [37,38]. Therefore, we hypothesized that the hnRNPU-actin complex has underlying roles in HCAEC proliferation and migration by facilitating RNA Pol II transcription and VEGF expression.

Based on the aforementioned evidence, we speculated that EVs may carry hnRNPU to form a complex with actin, promote RNA Pol II transcription, and augment the enrichment of RNA Pol II in the VEGF promoter region, thereby promoting the transcription of the VEGF gene and facilitating HCAEC proliferation and migration. In the present study, we established in vitro HCAEC hypoxia models and treated HCAECs with ESC-EVs to investigate the molecular mechanism by which the EVs-derived hnRNPU-actin (EVs-hnRNPU-actin) complex promotes HCAEC proliferation and migration, with the expectation to offer some novel targets for treating CAD.

Materials and methods

Cell culture

HCAECs and ESCs were provided by ATCC (Manassas, VA, USA). HCAECs were cultured in Dulbecco’s modified Eagle medium (DMEM) encompassing 10% fetal bovine serum (FBS), 100 μg/mL penicillin, and 100 U/mL streptomycin. ESCs were cultured in the DMEM (Hyclone, Logan, UT, USA) containing 15% FBS (Hyclone), 1% L-Glutamine (Corning, NY, USA), 1% non-essential amino acid (Gibco, Grand Island, NY, USA), 1% penicillin and streptomycin (Gibco), 1% β-mercaptoethanol (Sigma-Aldrich, St Louis, MO, USA), and 1000 U/mL leukemia inhibitory factor (Millipore, Billerica, MA, USA) at 37°C in an incubator with 5% CO2. Cells of the third to sixth passages were used for subsequent experiments [39-41].

Identification of ESCs

ESCs over 85% confluence were passaged as clumps by utilizing Dissociation Buffer (Cat# RP01007; Nuwacell, Hefei, Anhui, China). The relevant markers, including stage-specific embryonic antigen-4 (SSEA4), TRA-1-60, and TRA-1-81, were detected by flow cytometry. The cells were filtered through a 40-µm cell strainer and centrifuged at 500 g for 5 min at 4°C, with the supernatant removed. Later, cells were resuspended in fluorescence-activated cell sorting (FACS) buffer (cat. no. 00–4222; eBioscience, San Diego, CA, USA) at 2 × 106/mL to prepare single-cell suspension. The cells were incubated with antibodies anti-SSEA4 (1:1000, ab16287, Abcam, Cambridge, UK), anti-TRA-1-60 (10 µg/mL, MAB4360, Merck, Kenilworth, NJ, USA), and anti-TRA-1-81 (0.5 µg/mL, gtx48034, GeneTex, Irvine, CA, USA) on ice for 30 min, respectively. Subsequently, cells were reacted with secondary antibody goat anti-mouse immunoglobulin G (IgG) Alexa Fluor® 488 (1:2000, ab150113, Abcam) for 30 min. The mouse IgG (1 µg/mL, ab170190, Abcam) served as the isotype control. After two washes, cells were resuspended in phosphate-buffered saline (PBS) with 2% FBS and cells with the positive expression of SSEA4, TRA-1-60 and TRA-1-81 were analyzed by a flow cytometer (FACSAria III; BD Biosciences, San Jose, CA, USA). The results were analyzed using FlowJo (version 10.0.7r2; FlowJo LLC, Ashland, OR, USA). [42].

Extraction and identification of EVs

Upon reaching 80% cell confluence, the medium was replaced with FBS-free DMEM for further 48-h culture. The collected supernatant was centrifuged at 300 g for 10 min to remove cell debris and then centrifuged at 12,000 g for 30 min at 4°C. Thereafter, the separated supernatant was filtered with a 0.22-µm filter and centrifuged at 12,000 g for 70 min at 4°C through an Optima XPN-100 ultracentrifuge (Beckman Coulter, Fullerton, CA, USA), and then the supernatant was removed to collect the EVs, which were resuspended with sterile PBS and stored in a freezer at 80°C.

Following overnight fixation in PBS (pH 7.4) containing 2% glutaraldehyde, EVs were rinsed with PBS and then fixed with 1% osmium tetroxide overnight at 4°C. After dehydration in acetone, they were embedded with epoxy resin, sectioned, and then adsorbed onto formvar-coated copper grids, followed by staining with uranyl acetate and lead citrate. Thereafter, EVs were observed under a Tecnai transmission electron microscope (TEM, FEI Company, Hillsboro, OR, USA). The particle size distribution of EVs was detected using a NanoSight nanoparticle tracking analyzer (Malvern Panalytical, Malvern, Worcestershire, UK).

The expression levels of EV surface positive markers CD9 (ab92726, 1:2000, Abcam), CD81 (ab109201, 1:1000, Abcam), tumor susceptibility gene 101 (TSG101, 1:1000, ab133586, Abcam), and negative marker Calnexin (ab92573, 1:20,000, Abcam) were determined using Western blotting (WB). In addition, after ESCs were treated with 20 μM EVs inhibitor GW4869 (Sigma-Aldrich) for 2 h [43], PBS resuspension obtained from the ESCs by conducting the same EV extraction steps was used as the control. The identified EVs were lysed and then the total protein content was determined using the bicinchoninic acid (BCA) protein quantification kit (BL-FZ-1008, Bio-light Biotech, Shanghai, China). The protein content was regarded as the standard of EVs [40,41].

EVs grouping

The EVs were grouped as follows: (1) EVs; (2) GW: after ESCs were treated for 2 h with 10 μM GW4869 (Sigma-Aldrich) [44], the PBS resuspension was obtained by implementing the same EV extraction steps; (3) EVs-si-hnRNPU: EVs were separated and extracted following transfection of si-hnRNPU into ESCs for 24 h; (4) EVs-si-NC: EVs were isolated and extracted after ESCs were delivered with si-negative control (NC) for 24 h; (5) EVs + Prot K: EVs were treated with 20 mg/mL Proteinase K (Cat# 25–530-049) [21]; (6) EVs + Prot K + Triton: EVs were treated with 20 mg/mL Proteinase K and 1% TritonX-100 [21]. Lipofectamine RNAiMAX Transfection reagents (Cat# 13,778,150, Invitrogen, Carlsbad, CA, USA) were used to transfect 10 nmol/L si-hnRNPU or si-NC at final concentration of 3.75 µL/mL [21]. The si-hnRNPU and si-NC and Proteinase K were provided by Thermo Fisher Scientific (Waltham, MA, USA).

HCAECs treatment and grouping

HCAECs were cultured to the 3rd passage, and cells reached 80% confluence. Firstly, different concentrations (5, 10, 20, and 30 μg) of EVs were co-cultured with HCAECs for 12 h, 24 h, 36 h, and 48 h under hypoxia, respectively; next, the CCK-8 assay was utilized to detect the cell proliferation ability, with the optimal treatment concentration determined as 20 μg and the optimal treatment time determined as 24 h. HCAECs were allocated into the following groups: (1) Hypoxia group; (2) Hypoxia + EVs group: treated with 20 μg EVs; (3) Hypoxia + GW group: treated with 20 μg GW4869; (4) Hypoxia + VEGF group: treated with 10 ng/mL VEGF [45] instead of EVs; (5) Hypoxia + EVs-si-NC group: treated with 20 μg EVs-si-NC; (6) Hypoxia + EVs-si-hnRNPU group: treated with 20 μg EVs-si-hnRNPU; (7) Hypoxia + EVs + si-NC group: treated with 20 μg EVs and transfected with si-NC; (8) Hypoxia + EVs + si-VEGF group: treated with 20 μg EVs and transfected with si-VEGF. In addition, HCAECs cultured in a CO2 incubator at 37°C for 24 h under normoxia were used as the normal control (Normoxoia group). si-NC and si-VEGF were purchased from Thermo Fisher Scientific.

Cell counting kit-8 (CCK-8) assay

HCAECs were placed in 96-well plates at 2 × 104/cm2 for 48 h-culture. Cell viability and proliferation were examined using CCK-8 kits (A311-01, Vazyme, Nanjing, Jiangsu, China). Briefly, each well was supplemented with 10 μL CCK-8 solution (diluted in 100 μL culture fluid) and incubated for 1 h at 37°C. Next, the optical density at 450 nm was detected and quantified using an automatic microplate reader (Rayto, Jinan, Shandong, China) [39].

Flow cytometry

The collected cells were analyzed by fluorescein isothiocyanate (FITC) Annexin V apoptosis detection kits 556,547, BD Biosciences. Cells were double-stained with FITC Annexin V and propidium iodide. Cell apoptosis was detected using the flow cytometer (FACScan; BD Biosciences) and analyzed using CellQuest software (BD Biosciences). The percentage of apoptosis was considered as the apoptotic rate. The unstained cells were used as negative controls [46].

Scratch test

The migration of cells was tested using a scratch assay. HCAECs were cultured in 6-well plates and grown to approximately 2 × 106/well, followed by scratches in the center of the slide with 200 μL sterile pipette tips. After 24 h, the remaining cell-free area was measured and compared with the initially scratched area (in percentage). The images were obtained by a Zeiss Axio Observer microscope (Zeiss, Jena, Germany), and wound fusion was analyzed using the IncuCyte software (Essen BioScience, Ann Arbor, MI, USA) to quantify cell migration [26,39,47,48].

In vitro tube formation assay

After 24-well plates containing 1 mL culture medium (including 200 μL pre-gelled Matrigel from BD Biosciences) were placed in an incubator with 5% CO2 at 37°C for 30 min, HCAECs were seeded into the Matrigel at 5 × 104 cells/well and incubated in a CO2 incubator at 37°C. After 24 h, tube formation of HCAECs was photographed by a phase-contrast microscope and quantified by counting all the tubular structures in the gel [48,49].

Uptake of EVs

EVs were labeled using the PHK26 (Sigma-Aldrich). Briefly, 20 μg EVs were mixed with 1 mL PHK26 staining solution for 20 min, rinsed with PBS, and then centrifuged at 1,000 g for 70 min. Following the co-culture of PHK26-labeled EVs with HCAECs for 24 h, the uptake of EVs by HCAECs was observed under a confocal fluorescence microscope (Zeiss) [50].

WB assay

Cells or EVs were collected and then washed with PBS. Next, proteins were extracted from cells, EVs, or tissue homogenate using radio-immunoprecipitation assay (RIPA) lysis buffer (Solarbio, Beijing, China), followed by quantification with BCA protein assay kits. Total proteins were separated by sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis and then transferred to polyvinylidene fluoride membranes (Millipore, Bedford, MA, USA). After blocking with 5% skim milk for 2 h at room temperature, membranes were incubated overnight at 4°C with primary antibodies anti-hnRNPU (1–5 μg/mL, SAB1402227, Sigma-Aldrich), anti-VEGF (1 μg/mL, ab155944, Abcam), anti-RNA Pol II S2 (1:5000, ab193468, Abcam), and anti-actin (1:40, ab179467, Abcam). The next day, membranes were rinsed 3 times and incubated with secondary antibody goat anti-mouse anti-IgG (1:2000, ab97023, Abcam) for 1 h at room temperature. After that, membranes were washed 3 times, added with luminescent liquid, and imaged on the machine. The gray values of images were analyzed to assess the protein levels of hnRNPU, VEGF, and RNA Pol II (S2) using the Image J software (NIH, Bethesda, MD, USA), with anti-glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (1:10,000, ab8245, Abcam) as the internal control [51].

Co-immunoprecipitation (Co-IP) assay

Cells were collected and washed with PBS. Thereafter, proteins were extracted using RIPA lysis buffer containing protein inhibitor (Solarbio). Afterward, 300 μL extracts were added with 50% Protein A/G beads (Upstate Biotechnology, Lake Placid, NY, USA) for 1 h-incubation at 4°C for pre-clearing. After centrifugation to remove the beads, the supernatant was obtained and incubated overnight with antibody: anti-hnRNPU (1 μg/mL, A300-690A, Thermo Fisher Scientific) or normal rabbit IgG (1:1000, ab172730, Abcam) (negative control) at 4°C, followed by 2 h-incubation with 50% Protein A/G beads at 4°C. Later, the protein-antibody complexes were rinsed with RIPA buffer for 5 min × 5 times, followed by resuspension of protein-antibody complex beads in buffer containing SDS. Immunoprecipitated proteins were boiled at 95–100°C for 3–5 min and the supernatant was collected by centrifugation. The hnRNPU and actin protein levels were then analyzed by WB assay [52,53].

Chromatin immunoprecipitation (ChIP) analysis

Cells were resuspended with PBS and sonicated by adding ChIP nuclear lysis buffer to obtain the ideal chromatin fragments with a length of 150–1000 bp for IP reactions. The fragments were treated overnight with magnetic beads and antibody (anti-RNA Pol II, 5:25, ab26721, Abcam) followed by de-crosslinking. Purified DNA was obtained using a DNA purification kit (TIANGEN, DP214-02, Biolink Biotechnology, Beijing, China). Subsequently, quantitative polymerase-chain reaction (PCR) was performed to assess the enrichment level of RNA Pol II at the binding sites of the VEGF promoter, with total chromatin as standard changes (input). Rabbit IgG antibody (ab172730, Abcam) was used as the negative reference, and primers for the VEGF promoter region were designed: VEGF-F 5’-3’: GACGTTCCTTAGTGCTGGCGGGTAGGTTTGA; VEGF-R 5’-3’: GGCACCAAGTTTGTGGAGCTGAGAAC. Reaction conditions were as follows: pre-denaturation at 95°C for 3 min, 40 cycles of denaturation at 95°C for 10s, annealing at 55°C for 30s, and extension at 62°C for 20s. The PCR products were subjected to electrophoresis on 1% Tris-acetate-ethylenediamine tetraacetic acid agarose gels in triplicate [53,54].

Statistical analysis

All data were statistically analyzed and graphed with GraphPad Prism 8.0.1 software (GraphPad Software, San Diego, CA, USA). Measurement data were described as mean ± standard deviation (SD). The t test was used for comparisons between two groups and one-way analysis of variance (ANOVA) was employed for comparisons between multiple groups. Tukey’s multiple comparisons test was implemented for the post hoc analysis. The p < 0.05 indicated statistically significant.

Results

We hypothesized that ESC-EVs carried hnRNPU to form a complex with actin, promoted RNA Pol II transcription, and elevated the enrichment of RNA Pol II in the VEGF promoter region, thus facilitating the transcription of the VEGF gene and accelerating HCAEC proliferation and migration. The in vitro hypoxia models of HCAECs were established and EVs were extracted from ESCs. The proliferation and migration abilities of HCAECs were detected by CCK-8 kits, flow cytometry, scratch assay, and in vitro tube formation assay. The interaction between hnRNPU and actin was verified by the Co-IP assay, and the level of RNA Pol II on the VEGF gene promoter was determined using CHIP assay. It was found that ESC-EVs promoted HCAEC proliferation and migration under hypoxia. After treatment with ESC-EVs, hnRNPU level was raised in HCAECs and EVs could carry hnRNPU into HCAECs. The inhibition of hnRNPU partially counteracted the promotion of EVs on proliferation, migration, and angiogenesis of HCAECs. Moreover, EVs-hnRNPU-actin complex potentiated RNA Pol II transcription in HCAECs, increased RNA Pol II level on the VEGF gene promoter, and enhanced VEGF expression.

Identification of ESCs and EVs

ESCs were subcultured to the 3rd passage. The flow cytometer unveiled that cells with the positive expression of ESC surface markers SSEA-4, TRA1-60, and TRA1-81 were more than 99% (Figure 1(a)), indicating the good purity of ESCs. Subsequently, we adopted the ultracentrifugation method to separate and extract EVs from the supernatant of the 4th passage medium. TEM observed that isolated EVs exhibited a typical cup-shaped morphology (Figure 1(b)). Nanoparticle tracking analysis (NTA) unraveled that the particle size of EVs ranged from 30 nm to 150 nm, with a concentration of 2.19 × 109/mL (Figure 1(c)). In addition, WB assay showed that the expressions of EV surface markers CD9, CD81, and TSG101 were positive and Calnexin expression was negative compared with the GW group (Figure 1(d)). The aforesaid results elicited the successful isolation of ESC-EVs.

Figure 1.

Identification of ESCs and EVs. (a) Cells with the positive expression of ESC surface markers SSEA-4, TRA1-60, and TRA1-81 detected using flow cytometer; (b) Morphology of EVs observed using TEM; (c) Size distribution of EVs analyzed by NTA; (d) Expression levels of EV surface positive markers CD9, CD81, TSG101 and negative marker Calnexin measured using WB. EVs: EVs were extracted from ESCs by ultracentrifugation; GW: after treatment with GW4869 for 2 h, the PBS resuspension was obtained by implementing the same EV extraction steps.

ESC-EVs promoted HCAEC proliferation and migration under hypoxia

It is noteworthy that ESC-EVs can enhance post-myocardial infarction angiogenesis, promote myocardial repair, and potentiate cardiac function [17]. To investigate the role of ESC-EVs on the proliferation and migration of HCAECs under hypoxia, HCAECs were, respectively, cultured under normoxia and hypoxia. Next, CCK-8 kits suggested that the Hypoxia group possessed noticeably lower cell viability and proliferation than the Normoxoia group (Figure 2(a), p < 0.01). Flow cytometry revealed a significantly elevated apoptosis in the Hypoxia group (Figure 2(b), p < 0.01). The scratch test demonstrated that the wound fusion was lowered under hypoxia relative to normoxia (Figure 2(c), p < 0.01). Tube formation assay indicated the obvious formation of tubular capillary structural networks under normoxia (Figure 2(d), p < 0.01). The above results indicated the impairment of hypoxia on the proliferation, migration, and angiogenesis of HCAECs. Afterward, HCAECs were treated with different concentrations (5, 10, 20, and 30 μg) of EVs or 10 ng/mL VEGF [55] and cultured under hypoxia for 12 h, 24 h, 36 h, and 48 h. EVs were found to be internalized by HCAECs via the uptake assay (Figure 2(e)). CCK-8 assay indicated that EVs significantly elevated cell proliferation (p < 0.05). Meanwhile, we found that the cell viability was slowly and stably changed following 24 h co-culture with the EVs. The change in cell viability was minimal when the EVs concentration was 20 μg, which was close to the effect of VEGF treatment and showed no statistical difference with the treatment of 30 μg EVs (p > 0.05) (Figure 2(a)). It can be concluded that the optimal treatment concentration of EVs was 20 μg and the optimal treatment time was 24 h. In subsequent experiments, HCAECs were treated with 20 μg ESC-EVs or 10 ng/mL VEGF for 24 h under hypoxia, and next we detected the migration and angiogenesis of HCAECs. The results unveiled that 24 h-treatment with EV or VEGF elevated the viability and proliferation of HCAECs, reduced apoptosis, enhanced wound fusion, and increased the number of tubular capillary structures under hypoxia (Figure 2(a-d), p < 0.01). Altogether, ESC-EVs stimulated the proliferation and migration of HCAECs under hypoxia.

Figure 2.

ESC-EVs promoted HCAEC proliferation and migration under hypoxia. (a) CCK-8 kits assessed cell proliferation; (b) Flow cytometry detected cell apoptosis; (c) Scratch test tested wound fusion; (d) Tube formation assay; (e) Uptake of EVs. Cell experiments were replicated 3 times. Data were presented as mean ± SD. One-way ANOVA was employed for comparisons between multiple groups, followed by Tukey’s test. **p < 0.01. Normoxoia: HCAECs were cultured under normoxia for 24 h; Hypoxia: HCAECs were cultured under hypoxia for 24 h; Hypoxia + GW: HCAECs were treated with GW4869, followed by culture under hypoxia for 24 h; Hypoxia + EVs: HCAECs were treated with EVs, followed by culture under hypoxia for 24 h; Hypoxia + VEGF: HCAECs were treated with VEGF instead of EVs, followed by culture under hypoxia for 24 h.

EVs carried hnRNPU into HCAECs

Existing studies have found that hnRNPU can affect EC proliferation and facilitate angiogenic gene expressions by binding to miRNAs [21,26]. To explore whether EVs transfer hnRNPU into HCAECs, we further employed WB assay to detect hnRNPU levels in the cells, which illustrated that hnRNPU level in HCAECs was prominently diminished under hypoxia, while raised after treatment with EVs (Figure 3(a), p < 0.01). Moreover, hnRNPU was remarkably elevated in EVs. Subsequently, EVs were treated with Proteinase K and we found no obvious changes in hnRNPU levels in EVs, while simultaneous treatment of Proteinase K and Triton X-100 decreased hnRNPU levels in EVs (Figure 3(b)), indicating the presence of hnRNPU in EVs. Thereafter, ESCs were transfected with si-hnRNPU, followed by isolation and extraction of EVs. The WB assay showed decreased hnRNPU levels in EVs (Figure 3(c), p < 0.01). Finally, EVs-si-hnRNPU was co-cultured with HCAECs under hypoxia for 24 h, and then WB detection unveiled a markedly diminished hnRNPU level in the Hypoxia + EVs-si-hnRNPU group compared with the Hypoxia + EVs-si-NC group (Figure 3(a), p < 0.01). Overall, EVs could transport hnRNPU into HCAECs.

Figure 3.

EVs carried hnRNPU into HCAECs. HCAECs were treated with different EVs. (a) hnRNPU levels detected using WB; (b) The presence of hnRNPU in EVs verified by WB; (c) hnRNPU levels in EVs determined via WB. Cell experiments were repeated 3 times. Data were exhibited as mean ± SD. One-way ANOVA was adopted for comparisons between multiple groups, followed by Tukey’s test. **p < 0.01. Normoxoia: HCAECs were cultured under normoxia for 24 h; Hypoxia: HCAECs were cultured under hypoxia for 24 h; Hypoxia + GW: HCAECs were treated with GW4869, followed by culture under hypoxia for 24 h; Hypoxia + EVs: HCAECs were treated with EVs, followed by culture under hypoxia for 24 h; Hypoxia + EVs-si-NC: HCAECs were treated with EVs-si-NC, followed by culture under hypoxia for 24 h; Hypoxia + EVs-si-hnRNPU: HCAECs were treated with EVs-si-hnRNPU, followed by culture under hypoxia for 24 h; EVs: EVs were extracted from ESCs by ultracentrifugation; GW: after treatment with GW4869 (Sigma-Aldrich) for 2 h, the PBS resuspension was obtained by implementing the same EV extraction steps; EVs-si-NC: after ESCs were transfected with si-NC for 24 h, EVs were isolated and extracted; EVs-si-hnRNPU: after ESCs were transfected with si-hnRNPU for 24h, EVs were isolated and extracted.

Inhibition of hnRNPU partially annulled the promotion of EVs on HCAEC proliferation and migration

To further investigate whether EVs promote the proliferation and migration of HCAECs under hypoxia through hnRNPU transmission, HCAECs were co-cultured with EVs-si-hnRNPU for 24 h under hypoxia, and then we detected the proliferation and migration of HCAECs. The results elicited that compared with the Hypoxia + EVs-si-NC group, the cell viability and proliferation were predominantly lowered (Figure 4(a), p < 0.01), cell apoptotic rate was increased (Figure 4(b), p < 0.01), cell wound fusion was decreased (Figure 4(c), p < 0.01), and the number of tubular capillary structures was reduced (Figure 4(d), p < 0.01) in the Hypoxia + EVs-si-hnRNPU group. Briefly, inhibition of hnRNPU partially abrogated the promoting effects of EVs on the proliferation, migration, and angiogenesis of HCAECs.

Figure 4.

Inhibition of hnRNPU partially annulled the promotion of EVs on HCAEC proliferation and migration. HCAECs were co-cultured with EVs-si-hnRNPU under hypoxia for 24 h. (a) CCK-8 kits detected cell proliferation; (b) Flow cytometry assessed cell apoptosis; (c) Scratch test; (d) Tube formation assay. Cell experiments were duplicated 3 times. Data were described as mean ± SD. The t test was used for comparisons between two groups. **p < 0.01. Hypoxia + EVs-si-NC: HCAECs were treated with EVs-si-NC, followed by culture under hypoxia for 24 h; Hypoxia + EVs-si-hnRNPU: HCAECs were treated with EVs-si-hnRNPU, followed by culture under hypoxia for 24 h.

EVs-hnRNPU-actin complex facilitated RNA Pol II transcription, increased RNA Pol II level on the VEGF gene promoter, and promoted VEGF expression in HCAECs

Subsequently, we further examined the molecular mechanism of EVs in HCAECs. The Co-IP assay unraveled that the interaction between hnRNPU and actin was significantly attenuated in HCAECs under hypoxia, enhanced by EV treatment, but weakened upon inhibition of hnRNPU (Figure 5(a)). After that, WB assay revealed that the trend of RNA Pol II phosphorylation level paralleled the interaction of hnRNPU with actin in HCAECs (Figure 5(b), p < 0.05). In addition, CHIP assay suggested that RNA Pol II level on the VEGF gene promoter was decreased in HCAECs under hypoxia, elevated by EV treatment, while reduced after the suppression of hnRNPU (Figure 5(c), p < 0.05). Afterward, WB detection demonstrated diminished VEGF under hypoxia, increased VEGF after EV treatment, and decreased VEGF after hnRNPU inhibition (Figure 5(d), p < 0.05). To sum up, EVs-hnRNPU-actin complex promoted RNA Pol II transcription, elevated RNA Pol II level on the VEGF gene promoter, and facilitated VEGF expression in HCAECs.

Figure 5.

EVs-hnRNPU-actin complex facilitated RNA Pol II transcription, increased RNA Pol II level on the VEGF gene promoter, and promoted VEGF expression in HCAECs. HCAECs were co-cultured with EVs-si-hnRNPU under hypoxia for 24 h. (a) Interaction between hnRNPU and actin assessed using Co-IP assay; (b) Phosphorylation of RNA Pol II detected by WB; (c) RNA Pol II level on the VEGF gene promoter determined through CHIP assay; (d) Phosphorylation level of VEGF evaluated using WB. Cell experiments were repeated 3 times. Data were presented as mean ± SD. One-way ANOVA was used for comparisons between multiple groups, followed by Tukey’s test. *p < 0.05, **p < 0.01. Normoxoia: HCAECs were cultured under normoxia for 24 h; Hypoxia: HCAECs were cultured under hypoxia for 24 h; Hypoxia + GW: HCAECs were treated with GW4869, followed by culture under hypoxia for 24 h; Hypoxia + EVs: HCAECs were treated with EVs, followed by culture under hypoxia for 24 h; Hypoxia + EVs-si-NC: HCAECs were treated with EVs-si-NC, followed by culture under hypoxia for 24 h; Hypoxia + EVs-si-hnRNPU: HCAECs were treated with EVs-si-hnRNPU, followed by culture under hypoxia for 24 h.

VEGF inhibition partially reversed the promotion of EVs on HCAEC proliferation and migration

HCAECs were treated with EVs and si-VEGF under hypoxia to further validate the role of VEGF on the proliferation and migration of HCAECs under hypoxia. After 24 h, the proliferation, migration, and angiogenesis of HCAECs were detected by CCK-8 kits, flow cytometry, scratch test, and tube formation assay. The results illustrated that the Hypoxia + EVs + si-VEGF group exhibited prominently lowered cell viability and proliferation (Figure 6(a), p < 0.01), elevated apoptotic rates (Figure 6(b), p < 0.01), decreased wound fusion (Figure 6(c), p < 0.01), and diminished number of tubular capillary structures (Figure 6(d), p < 0.01) compared with the Hypoxia + EVs + si-NC group. In a word, repression of VEGF partially nullified the promotion of EVs on the proliferation, migration, and angiogenesis of HCAECs.

Figure 6.

VEGF inhibition partially reversed the promotion of EVs on HCAEC proliferation and migration. HCAECs were treated with EVs and si-VEGF under hypoxia for 24 h. (a) CCK-8 kits detected cell proliferation; (b) Flow cytometry evaluated cell apoptosis; (c) Scratch test; (d) Tube formation assay. Cell experiments were duplicated 3 times. Data were exhibited as mean ± SD. The t test was performed for comparisons between two groups. **p < 0.01. Hypoxia + EVs + si-NC group: HCAECs were treated with EVs and si-NC, followed by culture under hypoxia for 24 h; Hypoxia + EVs + si-VEGF: HCAECs were treated with EVs and si-VEGF, followed by culture under hypoxia for 24 h.

Discussion

CAD has emerged as a predominant threat to public health globally, which is an atherosclerotic disease with the manifestations of unstable angina, stable angina, myocardial infarction, or sudden cardiac death [56,57]. The normal vascular endothelium is considered a gatekeeper of cardiovascular health, whereas endothelial dysfunction is a chief contributor to atherosclerosis and CAD [58-60]. Circulating EVs released from diverse cells can alter CAD pathophysiological processes by transporting plentiful biological information [55]. This study verified the molecular mechanism of EVs in regulating HCAEC proliferation and migration by carrying hnRNPU to bind to actin, affecting Pol II transcription, and mediating the expression of the VEGF gene.

The utilization of ESCs has been documented in cardiac repair in animal studies [61,62]. Additionally, human ESCs can differentiate into contracting cardiomyocytes and human ESC-derived cardiomyocytes can regenerate infarcted myocardium [63]. EVs are biological particles released by various cell types, such as ESCs and considered for therapeutic utilization, including atherosclerosis treatment [64,65]. Furthermore, the regulation of EVs in cardiac regeneration and homeostasis has been alluded [66]. Firstly, after successful isolation and identification of normal ESC-EVs, we observed that ESC-EVs could be internalized by HCAECs. Additionally, hypoxia lowered HCAEC viability, proliferation, wound fusion and the number of tubular capillary structures, and elevated apoptosis, whereas ESC-EVs annulled these effects of hypoxia on HCAECs. Hypoxia/reoxygenation, as a cause of CAD, is predominately induced by HCAEC damage [67]. Alterations of EC functions, such as decreased endothelial nitric oxide availability, raised EC apoptosis, chemokine/adhesion molecule expressions, and pro-thrombotic activation, contribute to the incidence of atherosclerosis and CAD [68]. Hypoxia can induce cell viability decrease and apoptosis in human cardiac microvascular ECs [69]. ESC-derived exosomes facilitate endogenous repair mechanisms and augment cardiac function after myocardial infarction [17]. EVs derived from various stem cells exert different properties, including pro-angiogenesis, anti-fibrosis, and anti-apoptosis [70]. Briefly, ESC-EVs promoted proliferation, migration, and angiogenesis of HCAECs.

EVs, which can transfer miRNAs, DNA, proteins, and lipids, are effective in regenerative and preventive medicine and in protecting the heart from myocardial infarction and arrhythmias [71]. hnRNPs are expressed in separated EVs, including hnRNPU, hnRNPK, and hnRNPA2B1 [72]. In the present study, we constructed the in vitro hypoxia models of HCAECs, and additionally, HCAECs were cultured with ESC-EVs for 24 h under hypoxia. Our results elicited that the protein level of hnRNPU in HCAECs was decreased from 1.81-fold to 0.36-fold of GAPDH under hypoxia, a decrease of 80.43%. However, after treatment of EVs, hnRNPU expression was increased to 1.59-fold, an upregulation of 341.67%, and all the protein expression changes of hnRNPU were statistically significant. In addition, hnRNPU was encapsulated in EVs and EVs could transfer hnRNPU into HCAECs. Subsequently, we explored the role of hnRNPU in cellular functions and found that inhibition of hnRNPU annulled the promotion of EVs on HCAEC proliferation, migration, and angiogenesis. hnRNPU is required for cardiac pre-mRNA splicing and postnatal heart progression [25]. The studies about the regulation of hnRNPU on CAD and HCAECs are scarce. But, there are some relevant studies about other hnRNPs in that field. For example, silencing of hnRNPE1 represses the aggregation of endogenous LC3 and diminishes the protein level of LC3-II, thus promoting vascular EC apoptosis and inhibiting autophagy [73]. Intriguingly, the substantial roles of hnRNPA1 in human atherosclerotic lesions, damage-induced vessel remodeling, and vascular smooth muscle cell function are documented [74]. Reduced expression of hnRNPA1 damages neurological property, learning and memory abilities, which also aggravates degeneration and apoptosis of neuronal cells [75]. To sum up, EVs exerted a function on HCAEC growth by delivering hnRNPU.

The complex of hnRNPs and actin has been unraveled to regulate RNA Pol II transcription [76]. Blocking the binding between RNA Pol II and VEGF promoter inhibits VEGF expression [33,77]. Moreover, VEGF expression is tightly related to the proliferation and migration of vascular endothelial cells [35]. Next, we investigated the specific mechanism of EVs-transported hnRNPU in HCAECs. Our subsequent results revealed that EV treatment could abate the function of hypoxia on weakening hnRNPU–actin interaction, reducing RNA Pol II phosphorylation, and diminishing RNA Pol II level on the VEGF gene promoter and VEGF expression, while hnRNPU inhibition further nullified the effects of EVs treatment. Consistently, hnRNPU can promote transcription of RNA Pol II by binding to actin [28,29]. Similarly, hnRNPG interacts with RNA Pol II to mediate expression and alternative splicing of target mRNAs [74]. hnRNPK binds to the 3'UTR of mRNAs to post-transcriptionally regulate its levels by modulating its stability and involves in regulating angiogenesis pathway by activating VEGF transcription [72]. Moreover, in hypoxic breast cancer cells, RAB11B-AS1 upregulates levels of angiogenic factors, such as VEGFA and ANGPTL4 by elevating the recruitment of RNA Pol II [78]. Notably, our results discovered that VEGF inhibition counteracted the promotion of EVs on HCAEC proliferation, migration, and angiogenesis, which tied well with the existing finding that VEGF can trigger aortic and coronary EC proliferation [79] and inhibit EC apoptosis [80] and that VEGF stimulation varies EC gene transcription to support vessel formation [81]. Conjointly, EVs-hnRNPU-actin complex facilitated RNA Pol II transcription, increased VEGF expression, thus promoting HCAEC growth. The cell cycle regulates the expressions of numerous proteins [82-84] and consequently affects various cellular processes, including glycolysis and oxidative phosphorylation [85]. The link between cell cycle progression and apoptosis has been proposed many years ago and the regulation of the cell cycle may prevent or induce apoptotic responses [86]. Long-term time-lapse experiments have already provided insight into the cell cycle dependency of intrinsic apoptosis [87-92]. In the present study, we investigated the proliferation-promoting and apoptosis-inhibiting effects of ESC-EVs-hnRNPU on HCAECs, but its role in the cell cycle remains unclear, which is a limitation of this study.

Conclusion

To conclude, the present study highlighted that ESC-EVs elevated RNA Pol II transcription and VEGF expression by carrying hnRNPU to form a complex with actin, thereby facilitating proliferation and migration of HCAECs. Furthermore, ESC-EVs could provide a novel cell-free system to exploit the paramount regenerative capacity of ESCs and avoid the risk of transplantation and teratomas induced by direct ESCs or ESC-derived cells. A major source of limitation is due to the intricate effects of EVs-hnRNPU-actin complex on RNA Pol II transcription. To remedy the deficiency, future research shall target the hnRNPU-actin complex to deeply explore its effect on RNA Pol II transcription and HCAEC proliferation and migration at the gene level, providing more theoretical options for the treatment of CVs.

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