Atherosclerosis is a primary cause of death globally and is strongly associated with all cardiovascular-related deaths that occur as a result of complications such as stroke, hypertension, and heart attack (1). Atherosclerotic plaques are complex lesions that most likely form as a result of damage to the inner layer of an artery. Aortic smooth muscle cells (ASMCs) are the predominant cell type within the arterial wall. The proliferation and migration of ASMCs play a crucial role in the development of atherosclerotic lesions, where they are responsible for calcium deposition and connective tissue formation (1, 2). In mature blood vessels, ASMCs are found in a differentiated (contractile) phenotypic state, exhibiting a low proliferation rate and a low level of synthetic activity (3). A major focus in the prevention of atherosclerosis is to discover thera-peutics that can effectively inhibit the proliferation, migration, and phenotypic switching of ASMCs.
Vascular injury increases the production of growth factors like platelet-derived growth factor-BB (PDGF-BB). PDGF-BB initiates downstream signaling pathways to modulate the phenotypic switch of ASMCs to a de-differentiated state (3). PDGF-BB is a natural ligand that binds to PDGF receptor-β (PDGFRβ) (4). PDGF-BB suppresses the expression of differentiation markers such as α-smooth muscle actin (α-SMA), smooth muscle 22α (SM22α, also known as transgelin or TAGLN), smooth musclemyosin heavy chain (SM-MHC), and calponin (5). Inflammatory responses play a significant role in various stages of atherosclerosis and contribute to its pathophysiological progression (6, 7). The PDGF-BB stimulated proliferation and migration of vascular smooth muscle cells (VSMCs) is accompanied by an increase in the secretion of inflammatory factors, including interleukin (IL)-1β, IL-8, and tumor necrosis factor (TNF)-α (8). Focal adhesion kinase (FAK), a tyrosine kinase, interacts with multiple signaling molecules to promote cell survival (9). FAK degradation has been shown to promote the expression of cyclin-dependent kinase (CDK) inhibitors, p21Cip1 and p27Kip1, to prevent cell proliferation: p21Cip1 and p27Kip1 bind to cyclin-CDK complexes, inhibiting their activity and inducing cell cycle arrest, thereby reducing the proliferation of HASMCs (10). Furthermore, nuclear FAK acts as a scaffolding protein, facilitating cell survival through enhanced degradation of the tumor suppressor protein p53 during cellular stress (11). FAK inactivation results in p53- and p21-dependent mesodermal cell growth arrest (11). PDGFRβ-mediated signaling pathways, including FAK (12), mitogen-activated protein kinases (MAPKs) (13), serine/threonine kinase AKT-nuclear factor-κB (AKT-NF-κB) (14), and signal transducer and activator of transcription 3 (STAT3) (15), have been suggested to affect cell proliferation and migration.
Hyaluronic acid and proteoglycan link protein 1 (HAPLN1) was first discovered in the proteoglycan component extracted from bovine articular cartilage, and is a component of the extracellular matrix (ECM). HAPLN1 assists in maintaining the stability of two ECM molecules, HA and proteoglycan (16). HAPLN1 mediates myocardial regeneration and upregulates the expression of α-SMA (17, 18). HAPLN1 also plays an important role in maintaining endothelial permeability, and loss of HAPLN1 in the aged microenvironment promotes visceral metastasis (19). HAPLN1-treatment inhibited the migration of fibroblast-like synoviocytes (16). Furthermore, treatment of ASMCs with PDGF-BB has recently been demonstrated to result in decreased expression of HAPLN1 (20). However, the specific role and mechanisms of full-length recombinant human HAPLN1 (rhHAPLN1) in proliferation, migration, and phenotypic switching of SMCs remain unknown. In this study, we aimed to investigate the effects and mechanisms of rhHAPLN1 on PDGF-BB-induced proliferation, migration, and de-differentiation of HASMCs, to ultimately assist in the development of a new therapeutic approach for the treatment of atherosclerosis.
To explore the potential role of HAPLN1 in the differentiation of HASMCs, we first examined the concomitant expression of HAPLN1 and smooth muscle‑specific markers, including SM22α, α-SMA, calponin, and SM-MHC, during differentiation of HASMCs. The expression of both HAPLN1 and the markers were increased in cells growing in SMC differentiation at 6 days. However, these increases were significantly attenuated in cells treated with PDGF-BB compared to those growing in PDGF-BB-free media (Fig. 1A), indicating that HAPLN1 plays a role in the differentiation process. The PDGF-BB signaling-induced phenotypic switching of HASMCs from a contractile to a synthetic state is recognized as a crucial step in the proliferation and migration of HASMCs during the development and progression of atherosclerosis (21). Therefore, we aimed to investigate the regulatory effect of exogenous treatment with rhHAPLN1 on phenotypic switching in the presence or absence of PDGF-BB. Cells were grown in SMC differentiation media containing rhHAPLN1 for 6 days. HASMCs subjected to rhHAPLN1-treatment showed a significant increase in mRNA levels of the markers at day 6 compared to day 0 (Fig. 1B). In addition, while protein levels of the markers decreased in cells treated with PDGF-BB, this effect was reversed in cells cultured in rhHAPLN1-containing basal media supplemented with 5% fetal bovine serum (FBS), in a dose-dependent manner (Fig. 1C). Taken together, these results suggest that rhHAPLN1 could attenuate PDGF-BB-induced de-differentiation of HASMCs, even in the absence of SMC differentiation media, and therefore play an important role in the phenotypic switching of HASMCs to a contractile state.
The de-differentiated phenotype of HASMCs induced by PDGF-BB has been reported to result in cell proliferation (22). We therefore examined whether treatment with rhHAPLN1 could inhibit the proliferation of HASMCs. HASMCs proliferation was analyzed using fluorescent staining with Ki-67, a marker of proliferation, and the expression of cell cycle regulatory proteins was determined by western blotting. We observed a significant increase in the number of Ki-67-positive HASMCs in PDGF-BB-containing media, but not in media containing both PDGF-BB and rhHAPLN1 (Fig. 2A), indicating that rhHAPLN1 inhibits the PDGF-BB-induced proliferation (Supplementary Fig. 1A, B). This finding prompted us to examine an effect of rhHAPLN1 on FAK, which is known to be a scaffold protein sequestering p53 (11), as well as a key regulator of p21Cip1 and p27Kip1, cell cycle inhibitors that inhibit cyclin/CDK complexes (10). Our results indicate that rhHAPLN1 downregulated the levels of FAK in a dose-dependent manner (Fig. 2B). Consistent with these results, the levels of p53, p21Cip1, and p27Kip1 increased (Fig. 2B). Additionally, the components of the cyclin/CDK complexes showed a dose-dependent decrease upon treatment with rhHAPLN1 (Fig. 2C). Together, these results indicate that rhHAPLN1 treatment attenuated the proliferation of HASMCs by upregulating the protein levels of p53, p21Cip1, and p27Kip1, and downregulating FAK and cyclin/CDK proteins. Consistent with previous studies highlighting the importance of the cell cycle dynamics in regulating cell proliferation (23), our findings demonstrate that rhHAPLN1 inhibits the proliferation of HASMCs through upregulation of p53 and CDK inhibitors like p21Cip1 and p27Kip1, which may be regulated by FAK.
Matrix metalloproteinase-9 (MMP-9) is a gelatinase that degrades type IV collagen, leading to the migration and invasion of VSMCs, which results in the vascular plaque instability characteristic of vascular diseases such as atherosclerosis (24). An in vitro wound healing assay was performed to determine the effects of rhHAPLN1 on the PDGF-BB-induced migration of cells. Our data showed that the migration of HASMCs increased in PDGF-BB-containing media, but not in media containing PDGF-BB and rhHAPLN1 (Fig. 3A). This implies that rhHAPLN1 inhibits the PDGF-BB-mediated migration of HASMCs. We found that PDGF-BB caused an increase in the levels of MMP-9, however, this was reversed by rhHAPLN1 treatment in a dose-dependent manner (Fig. 3B), suggesting that the MMP-9 may be involved in cellular migration. To determine the mechanism by which rhHAPLN1 reduces MMP-9 levels, as reported previously (25, 26), we analyzed several cytokines such as TNF-α, IL-1β, and IL-8. Our results showed that PDGF-BB treatment enhanced the expression of these inflammatory factors, however, this effect was reversed by rhHAPLN1 treatment (Fig. 3C). In addition, the levels of p-AKT/AKT, p-NF-κB/NF-κB were decreased by rhHAPLN1 in a dose-dependent manner (Fig. 3D). Our findings suggest that rhHAPLN1 exerts a suppressive effect on the PDGF-BB-induced increase of MMP-9. This effect is likely mediated through the inhibitory effect of rhHAPLN1 on the expression of inflammatory cytokines.
To elucidate the inhibitory mechanisms of rhHAPLN1 on cell proliferation and migration, we investigated the effect of rhHAPLN1 on PDGF-BB-induced PDGFRβ signaling. rhHAPLN1 treatment dramatically reduced PDGF-BB-induced phosphorylation of PDGFRβ (p-PDGFRβ) and FAK (Y397, p-FAK) (Fig. 4A, B). In contrast, rhHAPLN1 upregulated the protein level of PDGFRβ in a dose-dependent manner (Fig. 4A). Furthermore, rhHAPLN1 significantly inhibited the phosphorylation of STAT3 (p-STAT3), and the phosphorylation of MAPKs (p-JNK, p-p38, p-ERK) by PDGF-BB in a dose-dependent manner (Fig. 4C). Similarly, rhHAPLN1 significantly downregulated Ras levels, and thereby inhibited the phosphorylation of c-Raf (p-c-Raf) induced by PDGF-BB in a dose-dependent manner (Fig. 4D). Taken together, our results reveal that rhHAPLN1 can attenuate multiple phosphorylation pathways downstream of PDGF-BB-induced PDGFRβ phosphorylation signaling, suggesting a mechanism by which rhHAPLN1 can inhibit PDGF-BB-mediated proliferation and migration of HASMCs (Supplementary Fig. 2).
The excessive proliferation and migration of abnormal HASMCs are major causes of the development and progression of cardiovascular diseases, such as atherosclerosis (27). Vascular injury or treatment with growth factors like PDGF-BB can induce phenotypic modulation of HASMCs, resulting in a decrease in the expression of contractile markers, including SM22α, α-SMA, calponin, and SM-MHC (28). Activation of PDGF receptor signaling can modulate the phenotype of HASMCs. HASMCs undergo a phenotypic switch from a contractile to a synthetic state after PDGFRβ binding to PDGF-BB (29). In healthy vessels, the differentiated HASMCs exhibit a contractile phenotype with high expression of these contractile markers.
In this study, we provide the first evidence that the expression of HAPLN1 is upregulated in conjunction with contractile markers in HASMCs grown in SMC differentiation media, but significantly downregulated in the presence of PDGF-BB (Fig. 1A). The treatment of cells with exogenous rhHAPLN1 significantly reversed the decreased gene expression and protein levels of such contractile markers elicited by PDGF-BB (Fig. 1A, C). Furthermore, rhHAPLN1 inhibited the PDGF-BB-induced proliferation (Fig. 2A, Supplementary Fig. 1A, B) and migration (Fig. 3A, Supplementary Fig. 1C) of HASMCs. In addition, our data show that such inhibitory effects of rhHAPLN1 on the proliferation and migration are likely to result from its crucial roles in FAK-dependent cell cycle arrest and the inflammatory cytokines-dependent production of MMP-9. In the context of proliferation, it is of note silencing of FAK has been shown to increase the expression levels of p53, p21Cip1, and p27Kip1 (9, 11). On the other hand, the expression of MMP-9 has been shown to be induced via ROS-dependent activation of the ERK1/2 and NF-κB pathways in VSMCs (30). MMP-9 plays an important role in HASMCs migration (25). Treatment with rhHAPLN1 effectively inhibited the expression of MMP-9 via inhibition of both p-NF-κB (Fig. 3D) and p-ERK (Fig. 4C). Furthermore, activation of PDGFRβ by PDGF-BB is one of the key factors involved in the dysregulation of proliferation and migration of HASMCs (4, 12). Here, we demonstrate that rhHAPLN1 blocked PDGF-BB-induced activation of PDGFRβ signaling (Fig. 4A, Supplementary Fig. 2), and subsequently attenuated downstream signaling pathways: FAK, MAPK, STAT3 and Ras/c-Raf (Fig. 4B-D), and/or Ras-AKT-NF-κB (Fig. 3D). These findings strongly suggest that treatment with rhHAPLN1 may suppress the development and progression of vascular aging-related diseases such as atherosclerosis and restenosis by regulating the phenotypic switching of HASMCs.
Our results indicated that rhHAPLN1 inhibited proliferation, migration, and phenotypic switching induced by treatment of HASMCs with PDGF-BB, thus providing a new therapeutic approach for the treatment of atherosclerosis.
HASMCs (Cell Applications, CA, USA) were maintained in SMC growth medium (Cell Applications, CA, USA) at 37°C in 5% CO2 incubator. Media were changed every 2 days. HASMCs were seeded in growth medium for 24 h. The cells were changed to basal medium (Cell Applications) supplemented with 5% FBS containing the indicated concentration of rhHAPLN1 (WUXI, Shanghai, China) or PDGF-BB (R&D Systems, MN, USA). For SMC differentiation, HASMCs were incubated in SMC differentiation medium (Cell Applications) for 6 days, changing the medium every 2 days, after which cells were characterized. Cells at passage 4-9 were used in subsequent experiments.
Total protein extractions were performed using the RIPA buffer (BioWorld, Ohio, USA) supplemented with protease and phosphatase inhibitors (Roche, Mannheim, Germany). Protein concentrations were determined using a BCA assay (Thermo Fisher Scientific, MA, USA). From each extract, 15 μg of protein was subjected to SDS-PAGE on an 8-12% gel and transferred onto a PVDF membrane (Bio-Rad, CA, USA). Membranes were blocked for 1 h using 5% skim milk or 5% bovine serum albumin (BSA) and incubated at 4°C overnight with the following primary antibodies: SM22α, α-SMA, SM-MHC (Abcam, Cambridge, UK); Calponin (Sigma, MO, USA); Cyclin D, p-PDGFRβ, PDGFRβ, p-FAK, FAK, p-STAT3, STAT3, p-ERK, ERK, JNK, p-JNK, p38, p-p38, AKT, p-AKT, p-NF-κB, NF-κB, Ras, c-Raf, p-c-Raf (Cell Signaling, MA, USA); MMP-9, GAPDH, Cyclin E, CDK2, CDK4, p53, p27Kip1, p21Cip1 (Santa Cruz, TX, USA). After washing three times with TBST (TBS with 0.1% Tween-20), membranes were incubated with anti-rabbit or anti-mouse secondary antibodies (Cell Signaling) for 1 h at room temperature. Bands were visualized with the ECL reagent (Gen-DEPOT, TX, USA). Quantitative analysis was performed with scanning densitometry using ImageJ (NIH, MD, USA).
Total RNA was extracted from cells using an RNeasy mini kit (Qiagen, CA, USA). For each sample, 2 μg of total RNA was used for cDNA synthesis using the PrimeScript RT Reagent kit (TaKaRa, Shiga, Japan) according to the manufacturer’s instructions. qRT-PCR was performed using the SYBR Green Supermix and CFX-Connect (Bio-Rad, CA, USA). Primer sequences are listed in Supplementary Table 1. Data were analyzed using the 2-DDCt method and GAPDH was used as an internal control.
HASMCs were seeded in 6-well plates. The medium was replaced with fresh medium containing 20 ng/ml rhHAPLN1 an incubated for 24 h. The next day, cells were treated with 10 ng/ml PDGF-BB in the presence or absence of 20 ng/ml rhHAPLN1 and incubated for 24 h. After incubation, a single wound was created in the cell monolayer by scratching the surface with an SPLScar Scratcher (SPL Life Science, Pocheon, Korea). Images were captured by inverted microscopy (Nikon ECLIPSE Ts2, Tokyo, Japan) at 0 and 8 h after wounding, and wound areas were determined using ImageJ software.
For immunofluorescence staining of Ki-67 (Abcam, Cambridge, UK), cells were fixed with 4% paraformaldehyde and then washed three times using PBS. Cells were permeabilized with 0.03% Triton X-100 at room temperature for 5 min and subsequently blocked with 3% BSA for 1 h at room temperature. Ki-67 antibody staining was performed overnight at 4°C in 3% BSA, after which cells were incubated with the Alexa Fluor 488-conjugated secondary antibody (Invitrogen, CA, USA), for 1 h at room temperature. Cell nuclei were stained using DAPI (Thermo Fisher Scientific) for 15 min at room temperature. After staining, images were captured using an inverted microscope (Nikon ECLIPSE Ts2). Ratios of the number of Ki-67 positive cells to total cells were determined for each substrate on three views per sample.
All statistical analyses were performed using GraphPad Prism 9 software (GraphPad Software, CA, USA). All results are presented as the means ± standard error of the mean (SEM) of at least three replicates. Student’s t-test was used to compare the results between two groups. Results with a P-value < 0.05 were considered statistically significant.
D.Z., Z.F., J.M.J., G.Y. and I.C.S. are employees of HaplnScience Inc. J.M.J. and D.K.K. are shareholders of HaplnScience Inc. The remaining authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.