Introduction: Cardiovascular disorders are characterized by vascular smooth muscle (VSM) transition from a contractile to proliferative state. Protease-activated receptor 2 (PAR2) involvement in this phenotypic conversion remains unclear. We hypothesized that PAR2 controls VSM cell proliferation in phenotype-dependent manner and through specific protein kinases. Methods: Rat clonal low (PLo; P3–P6) and high passage (PHi; P10–P15) VSM cells were established as respective models of quiescent and proliferative cells, based on reduced PKG-1 and VASP. Western blotting determined expression of cytoskeletal/contractile proteins, PAR2, and select protein kinases. DNA synthesis and cell proliferation were measured 24–72 h following PAR2 agonism (SLIGRL; 100 nM–10 μ<sc>m</sc>) with/without PKA (PKI; 10 μ<sc>m</sc>), MEK1/2 (PD98059; 10 μ<sc>m</sc>), and PI3K (LY294002; 1 μ<sc>m</sc>) blockade. Results: PKG-1, VASP, SM22α, calponin, cofilin, and PAR2 were reduced in PHi versus PLo cells. Following PAR2 agonism, DNA synthesis and cell proliferation increased in PLo cells but decreased in PHi cells. Western analyses showed reduced PKA, MEK1/2, and PI3K in PHi versus PLo cells, and kinase blockade revealed PAR2 controls VSM cell proliferation through PKA/MEK1/2. Discussion: Findings highlight PAR2 and PAR2-driven PKA/MEK1/2 in control of VSM cell growth and provide evidence for continued investigation of PAR2 in VSM pathology.

Cardiovascular disease (CVD) remains the leading cause of morbidity and mortality in the USA [1] and worldwide [2]. Although many basic mechanisms of CVD have been identified over the years, the number of CVD-related deaths including age-adjusted mortality rates continues to rise [1, 2]. Efforts to identify new molecular and/or cellular processes underlying CVD along with potential new pharmacologic targets to combat CVD are clearly needed.

The pathogenesis of CVD including the response to vascular injury is multifactorial and involves aberrant inflammation, uncontrolled vascular smooth muscle (VSM) cell proliferation and migration, and abnormal matrix balance [3]. Mature healthy VSM cells display a differentiated and contractile phenotype with low levels of proliferation to maintain a quiescent steady state. However, during disease or injury, VSM cells dedifferentiate into a synthetic and proliferative phenotype, often with severe clinical consequences such as alterations in regional blood pressure profiles, lumen occlusion, and vessel wall remodeling [3‒5]. Dedifferentiated VSM cells are characterized by loss of cytoskeletal contractile proteins necessary for maintaining quiescence and proper function of the vessel [5, 6]. Thus, approaches for controlling pathologic repercussions of VSM phenotypic switching and growth in the context of CVD are of significant scientific and therapeutic interest.

G protein-coupled receptor (GPCR) signaling is of particular interest in CVD [3, 7]. GPCRs represent the largest group of cell surface receptors in the human genome and constitute ∼34% of all FDA-approved drug targets [7]. Considering their wide-ranging intracellular targets, GPCRs can impact numerous effectors, including members of the equally broad family of protein kinases [3, 4, 8]. Protease-activated receptors (PARs), a GPCR subfamily, have been theorized as important in cardiovascular (patho)physiology [9, 10], and of emerging interest in VSM is PAR2. Activated by extracellular trypsin and mast cell tryptase [11], both liberated during vascular disease or injury, PAR2 has been implicated in cellular changes associated with pathologic VSM [9‒11]; however, the role of PAR2 and its reliance on kinase signals in pathologic VSM phenotypic transition and growth in the setting of CVD is not clear.

Our study aimed to determine impact and signaling mechanisms of PAR2 during VSM cell phenotypic switching and growth. While in vivo examination of CVD is of critical importance, our research strategy was designed a priori to use homogenous clonal primary cell populations (PLo, PHi) to specifically examine PAR2 and its kinase actions on VSM cell growth without associated confounding effects often observed in the heterogenous, multi-cellular in vivo environment. We hypothesized that PAR2 controls VSM cell growth in a phenotype-specific manner, dependent upon specific protein kinases, and that PAR2 and its kinase effectors may represent potential therapeutic targets against aberrant VSM growth. Notable observations from our study include distinction of low passage (PLo) from high passage (PHi) VSM cells through differential expression of cyclic GMP-dependent protein kinase type 1 (PKG-1) and its effector vasodilator-activated serum phosphoprotein (VASP); loss of the cytoskeletal contractile proteins smooth muscle protein 22α (SM22α), calponin 1, and cofilin in PHi cells compared to PLo cells; PAR2 agonism promoting growth in PLo cells yet suppressing growth in PHi cells; reduced expression of mitogen-activated protein kinase kinase 1/2 (MEK1/2), cyclic AMP-dependent protein kinase (PKA), and phosphoinositol-3 kinase (PI3K) in PHi versus PLo cells; and identification of PKA and MEK1/2 as plausible mechanisms of PAR2 control of VSM cell growth. Our findings shed light on the biological and pathophysiological importance of PAR2 and its kinase-dependent mechanisms in VSM growth control in CVD.

This study abided by recommendations of the East Carolina University Animal Care and Use Committee and the Guide for the Care and Use of Laboratory Animals [12]. Animal experiments were designed according to ARRIVE 2.0 guidelines [13]. Only male rats were used in this study to offset potential influence of female hormones on VSM growth as we have documented [14]. Rats were housed with irradiated aspen wood chip bedding and provided food and water ad libitum.

Rat Primary VSM Cell Culture

Thoracic aortae from male CD® (Sprague-Dawley IGS) rats (100–125 g; 4–5 weeks old; strain 001; Charles River) were digested with collagenase/elastase and derived VSM cells were cultured in Dulbecco’s Modified Eagle’s Medium (DMEM; #10-013-CM; Corning) with 10% fetal bovine serum (FBS; #900-108-500; GeminiBio), 2 mml-Glutamine, sodium pyruvate, 5 mm HEPES, and Primocin (100 mg/L; #ant-pm-2; InvivoGen) at 37°C in 95% air/5% CO2 [4, 8]. Cells were serially split (at 80–100% confluence; ∼3 cell doublings per passage [P]) from the same parental clones into two cohorts: PLo cells between P3 and P6, and PHi cells between P10 and P15, recognized passages for distinguishing contractile versus synthetic VSM cells [15, 16].

Pharmacologic PAR2 Modulation

PLo and PHi VSM cells were plated in multi-well plates and incubated (95% air/5% CO2) for 48 h in growth media (10% FBS in DMEM) until adherent. To establish quiescence, cells were switched to low serum media (0.2% FBS in DMEM) for 24 h prior to growth stimulation with/without the addition of vehicle (Veh) or the PAR2 agonist SLIGRL (#1468; Tocris Bioscience, Minneapolis, MN, USA) for varying amounts of time per experiment.

Western Blotting

The Pierce Coomassie (Bradford) Protein Assay Kit (#23200; Thermo Fisher) was used to determine protein quantification of cell homogenates per manufacturer’s instructions. Lysates were subjected to sodium dodecyl sulfate-polyacrylamide gel electrophoresis using stain-free gels (#5678123; Bio-Rad) [4, 8]. Separated protein was transferred to polyvinylidene fluoride membranes using the TransBlot Turbo Transfer System (Bio-Rad) at a mixed MW setting (7 min, 2.5 A up to 25 V). Membranes were blocked (2½ h at RT°, 5% milk solution with 0.1% Tris-buffered saline+Tween-20 (TBST)) and incubated at 4°C in 5% BSA/0.1% TBST overnight using 1:1000 dilution for primary antibodies directed against PAR2 (#sc-13504; Santa Cruz Biotechnology), PKG-1 (#3248), phosphorylated (Ser239; #3114) and total VASP (#3112), SM22α (#52011), calponin 1 (#17819), phosphorylated (Ser3; 1:500; #3311) and total cofilin (#3318), phosphorylated (Thr197; #4718) and total catalytic PKA-Cα (#4782), regulatory p85 (#4292) and catalytic p110 (#4225) PI3K, phosphorylated (Ser217/Ser221; #9121) and total MEK1/2 MAPK (#9122), and GAPDH (#2118S; all from Cell Signaling Technology). Membranes were washed in TBST and incubated for 1 h at RT° in 5% dry milk plus TBST using a 1:10,000 HRP-conjugated anti-rabbit or anti-mouse secondary antibody (#AP308P; #12–348; Millipore Sigma) as appropriate. Membranes were developed using Super Signal West Pico or Super Signal West Femto Substrate (#34580; #34095; Thermo Fisher). Chemiluminescent signals were analyzed using a Bio-Rad ChemiDoc MP Imaging System, and for each blot, untransformed (identical) automatic optimal high and low exposure settings including brightness/contrast and gain were used. Densitometry was performed to quantify target protein expression using Bio-Rad Image Lab Touch Software (v.2.2.0.08). Following current best practices for normalization of loading variations in Western blotting [17], target protein values were normalized to corresponding total protein abundance values (using inclusive proteins from the top to the bottom of each gel for each sample) from the same gel, and data are represented as target protein normalized to total protein. Normalization of target protein to total protein eliminates differences in target protein expression due to discrepancies in sample loading and, in turn, addresses biological variabilities inherent within samples across a single cohort [17]. Expression of GAPDH was probed in PLo and PHi cells to ensure that differences in protein expression were not the result of global protein loss due to serial passages; however, GAPDH was not used as a normalization marker as its abundance can change under varying cardiovascular conditions [18] and its molecular weight is identical to that of other target proteins analyzed in this study (which could skew densitometry measurements for proteins close in molecular weight, even following stripping/re-probing).

VSM Cell Growth and Viability

In PLo and PHi VSM cells, with/without SLIGRL and with/without specific protein kinase blockade, complementary approaches were used to determine the impact of PAR2 on cell growth and viability: (1) BrdU incorporation to estimate DNA replication; (2) automated cell number quantification for cell proliferation; and (3) trypan blue exclusion staining and neutral red/MTT assays for cell viability.

BrdU Incorporation

PLo and PHi VSM cells were seeded in 96-well plates (10,000 cells/well) and incubated with SLIGRL (100nM–10 µm), with/without 30-min pretreatment with the following inhibitors: PKI for PKA (10 μm; #BML-P203-0500; Enzo, Farmingdale, NY, USA); LY294002 (LY) for PI3K (1 μm; #440202; Sigma Aldrich, St. Louis, MO); PD98059 (PD) for MEK1/2 (10 μm; #1213; Tocris); or vehicle (appropriate for each inhibitor) as detailed [4] for 24 h, after which media was removed and a colorimetric BrdU cell proliferation ELISA (#2752; Millipore Sigma) was performed per manufacturer’s guidelines.

Automated Cell Number Quantification with Trypan Blue Exclusion Staining

Following PAR-specific interventions, cells were trypsinized and cell numbers were determined through automated quantification with trypan blue exclusion (ViCell; Beckman Coulter) to provide objective measures of viability and total and viable cell numbers as detailed [4, 8].

Neutral Red and MTT Assays

VSM cells were seeded in 96-well plates (10,000 cells/well) and incubated with SLIGRL (100 nm–10 μm), with/without 30-min pretreatment with kinase inhibitors or vehicle for 24 h. A neutral red assay (#ab234039; Abcam) was performed per manufacturer’s instructions with absorbance read at 540 nm, with values corrected for background and normalized to cell density. In separate experiments, media was removed and a colorimetric MTT assay (#30-1010K; ATCC, Manassas, VA, USA) was performed with absorbance read at 590 nm with a reference of 620 nm [4].

Protein Kinase Blockade

VSM cells were pretreated with the following inhibitors for 30 min prior to the addition of SLIGRL: PKI for PKA, LY for PI3K, PD for MEK1/2 or vehicle (appropriate for each inhibitor) as detailed [4].

Statistical Analyses

Statistical significance between cohorts within an experiment was determined using a one-way analysis of variance (ANOVA) or a repeated measures two-way ANOVA for assessment of interaction effects. When the F ratio indicated a significant difference, a multiple replicate Tukey’s (recommended) post hoc test was performed to identify individual paired differences. Two group comparisons were analyzed using an unpaired Student’s t test. Results are reported as mean ± standard error of the mean. Sample sizes per experiment, degrees of freedom (DF), and statistical indicators are included in the figure legends, and statistical significance was determined by a p value <0.05.

Phenotypic Changes in PLo versus PHi VSM Cells

Following confirmed approaches for distinguishing differentiated, contractile VSM cells from dedifferentiated, synthetic cells [15, 16], rat primary PLo (P3–P6) and PHi (P10–P15) VSM cell lysates were generated and probed for the phenotypic markers PKG-1 and the PKG-1 target phosphorylated (Ser239) and total VASP. PHi cells showed significantly reduced (p < 0.001) PKG-1 expression compared to PLo cells (shown in Fig 1a, b; see also online suppl. material; for all online suppl. material, see https://doi.org//10.1159/000532032). Similarly, PHi cells showed significantly reduced expression of both phosphorylated VASP (p = 0.0089; shown in Fig. 1c, d) and total VASP (p < 0.0001; shown in Fig. 1e, f) compared to PLo cells. Considering the relative reduction in total VASP exceeded that observed for phosphorylated VASP in PHi cells, the phosphorylated to total VASP ratio was significantly higher (p = 0.0015) in PHi cells than in PLo cells (shown in Fig. 1g). We have previously documented the biological significance of alterations in solitary phosphorylated VASP and in solitary total VASP as well as the utility of using a phosphorylated to total protein ratio as a relative indicator of kinase activity [4, 8, 19, 20]. For each blot, target proteins were normalized to total protein per sample from the same gel to ensure differences in expression were not the result of variations in sample loading [17].

Fig. 1.

Expression of PKG-1 and VASP in PHi versus PLo VSM cells. a Representative Western blot for PKG-1 expression in PLo and PHi VSM cell lysates along with the total protein blot from the same gel. b Densitometry reveals significantly reduced (p < 0.001; DF = 10) PKG-1 expression (normalized to total protein) in PHi cells compared to PLo cells; n = 6/group. Similarly, PHi cells showed significantly lower expression of both phosphorylated (Ser239) VASP (p = 0.0089; DF = 21; c, d) and total VASP (p < 0.0001; DF = 21; e, f) compared to PLo cells. The ratio of phosphorylated to total VASP was significantly higher (p < 0.0015; DF = 16) in PHi versus PLo cells (g). Each target protein was normalized to total protein within each blot [17], and all PLo and PHi comparisons were performed on the same blot but were separated in the figures for clarity. For c-g, n = 6–12/group. A Student’s t test was performed on data in b, d, f, g; **p < 0.01; ***p < 0.001; ****p < 0.0001 as shown.

Fig. 1.

Expression of PKG-1 and VASP in PHi versus PLo VSM cells. a Representative Western blot for PKG-1 expression in PLo and PHi VSM cell lysates along with the total protein blot from the same gel. b Densitometry reveals significantly reduced (p < 0.001; DF = 10) PKG-1 expression (normalized to total protein) in PHi cells compared to PLo cells; n = 6/group. Similarly, PHi cells showed significantly lower expression of both phosphorylated (Ser239) VASP (p = 0.0089; DF = 21; c, d) and total VASP (p < 0.0001; DF = 21; e, f) compared to PLo cells. The ratio of phosphorylated to total VASP was significantly higher (p < 0.0015; DF = 16) in PHi versus PLo cells (g). Each target protein was normalized to total protein within each blot [17], and all PLo and PHi comparisons were performed on the same blot but were separated in the figures for clarity. For c-g, n = 6–12/group. A Student’s t test was performed on data in b, d, f, g; **p < 0.01; ***p < 0.001; ****p < 0.0001 as shown.

Close modal

Contractile and Cytoskeletal Proteins in PLo versus PHi VSM Cells

To further establish our PLo and PHi VSM cells as respective models of differentiated, contractile and dedifferentiated, synthetic cells, key smooth muscle contractile and cytoskeletal markers were assessed. PHi cells expressed significantly lower SM22α (p = 0.0016) compared to PLo cells (shown in Fig. 2a, b; see also online suppl. material). Calponin 1 also had significantly reduced expression (p < 0.0001) in PHi cells versus PLo cells (shown in Fig. 2c, d). Lastly, both phosphorylated (Ser3) and total cofilin had significantly reduced expression (p = 0.0014) in PHi cells compared to PLo cells (shown in Fig. 2e–h). The phosphorylated to total cofilin ratio was nonsignificantly increased (p = 0.1563) in PHi cells compared to PLo cells (shown in Fig. 2i), likely due to high sample variability within the PHi cohort. There were no significant changes in GAPDH expression between PLo and PHi cells (shown in Fig. 2j, k), measured as confirmation that serial passages do not contribute to global loss of proteins in PHi cells.

Fig. 2.

Expression of cytoskeletal contractile proteins in PLo versus PHi VSM cells. a Representative Western blot for SM22α expression in PLo and PHi VSM cell lysates along with the total protein blot from the same gel. b Densitometry shows that PHi VSM cells have significantly reduced SM22α expression (p = 0.0016; DF = 8) compared to PLo cells. Similarly, calponin 1 expression was significantly reduced (p < 0.001; DF = 8) in PHi cells versus PLo cells (c, d). Both phosphorylated (Ser3) and total cofilin had significantly lower expression levels (p = 0.0067, DF = 30; p = 0.0014, DF = 8, respectively) in PHi cells compared to PLo cells (e-h), with an unchanged (p = 0.1563; DF = 8) phosphorylated to total cofilin ratio (i). Expression of housekeeper GAPDH showed no significant changes in PLo versus PHi cells (j, k). PLo and PHi comparisons were performed on the same blots but were separated in the figures for clarity. Each target protein was normalized to total protein within each blot [17]. n = 5–16/group, and a Student’s t test was used; **p < 0.01; ***p < 0.001; ****p < 0.0001 as shown.

Fig. 2.

Expression of cytoskeletal contractile proteins in PLo versus PHi VSM cells. a Representative Western blot for SM22α expression in PLo and PHi VSM cell lysates along with the total protein blot from the same gel. b Densitometry shows that PHi VSM cells have significantly reduced SM22α expression (p = 0.0016; DF = 8) compared to PLo cells. Similarly, calponin 1 expression was significantly reduced (p < 0.001; DF = 8) in PHi cells versus PLo cells (c, d). Both phosphorylated (Ser3) and total cofilin had significantly lower expression levels (p = 0.0067, DF = 30; p = 0.0014, DF = 8, respectively) in PHi cells compared to PLo cells (e-h), with an unchanged (p = 0.1563; DF = 8) phosphorylated to total cofilin ratio (i). Expression of housekeeper GAPDH showed no significant changes in PLo versus PHi cells (j, k). PLo and PHi comparisons were performed on the same blots but were separated in the figures for clarity. Each target protein was normalized to total protein within each blot [17]. n = 5–16/group, and a Student’s t test was used; **p < 0.01; ***p < 0.001; ****p < 0.0001 as shown.

Close modal

PAR2 and VSM Cell Growth

Following objective characterization of PLo versus PHi VSM cells, the influence of cell passage on PAR2 expression and VSM growth was examined. Expression of PAR2 was significantly reduced (p = 0.0157) in PHi cells compared to PLo cells (shown in Fig. 3a, b; see also online suppl. material). Using BrdU incorporation to estimate DNA replication, SLIGRL-treated PLo cells showed significantly elevated DNA synthesis compared to vehicle controls after 24 h (overall p < 0.0001; post hoc p < 0.0001 for 100 nm; shown in Fig. 3c). In comparison, PHi cells treated with SLIGRL had significantly reduced DNA replication compared to controls after 24 h (overall p = 0.0428; post hoc p = 0.0410 for 1 μm; shown in Fig. 3d). Automated cell counting with trypan blue exclusion staining showed a significant increase in PLo viable cell numbers (p = 0.0021 at 10 μm) and a significant decrease in PHi viable cell numbers (p = 0.0430 at 10 μm) with PAR2 agonism compared to vehicle controls through 48 h (shown in Fig. 3e, f). A repeated measures two-way ANOVA revealed a significant (p = 0.0011) interaction effect between time and SLIGRL concentration in PLo cells and a marked, nonsignificant (p = 0.0527) trend for an interaction between time and SLIGRL in PHi cells. Interestingly, even in the absence of SLIGRL PAR2 agonism, PHi cells showed increased BrdU absorbance and viable cell numbers after 48 h compared to PLo cells. No significant differences in viability were detected between PLo and PHi cells at all times and SLIGRL concentrations used per trypan blue exclusion staining and neutral red/MTT assays (data not shown).

Fig. 3.

PAR2 agonism increases growth in PLo VSM cells but decreases growth in PHi cells. a Representative Western blot for PAR2 expression in PLo and PHi VSM cell lysates along with the total protein blot from the same gel. b Densitometry shows that PAR2 expression is significantly reduced (p = 0.0157; DF = 14) in PHi cells compared to PLo cells. Using BrdU incorporation to estimate DNA synthesis, in (c), PLo cells showed significantly increased DNA synthesis with the PAR2 agonist SLIGRL (overall p < 0.0001; post hoc p < 0.0001 for 100 nm; DF = 3, 56) compared to vehicle controls after 24 h. In PHi cells (d), DNA synthesis was significantly decreased in SLIGRL-treated cells versus controls after 24 h (overall p = 0.0428; post hoc p = 0.0410 for 1 μm; DF = 3, 92). Correspondingly, viable cell numbers were significantly increased (p = 0.0021) in PLo cells (e) but significantly decreased (p = 0.0430) in PHi cells (f) with PAR2 agonism through 48 h. There was a significant (p = 0.0011; DF = 6, 59) trend for interaction of incubation time and SLIGRL concentration in PLo cells and a non-significant (p = 0.0527; DF = 6, 130) trend for interaction in PHi cells. For data in (a) and (b), a Student’s t test was performed. For data in (c, d), a one-way ANOVA was performed, and for data in (e, f), a repeated measures two-way ANOVA was performed including analysis of interaction effects. For data in (c-f), individual comparisons were analyzed using Tukey’s post hoc test. a, bn = 8/group. c, dn = 9–18/group. en = 5–6/group. fn = 11–12/group. *p < 0.05; ****p < 0.0001 as shown.

Fig. 3.

PAR2 agonism increases growth in PLo VSM cells but decreases growth in PHi cells. a Representative Western blot for PAR2 expression in PLo and PHi VSM cell lysates along with the total protein blot from the same gel. b Densitometry shows that PAR2 expression is significantly reduced (p = 0.0157; DF = 14) in PHi cells compared to PLo cells. Using BrdU incorporation to estimate DNA synthesis, in (c), PLo cells showed significantly increased DNA synthesis with the PAR2 agonist SLIGRL (overall p < 0.0001; post hoc p < 0.0001 for 100 nm; DF = 3, 56) compared to vehicle controls after 24 h. In PHi cells (d), DNA synthesis was significantly decreased in SLIGRL-treated cells versus controls after 24 h (overall p = 0.0428; post hoc p = 0.0410 for 1 μm; DF = 3, 92). Correspondingly, viable cell numbers were significantly increased (p = 0.0021) in PLo cells (e) but significantly decreased (p = 0.0430) in PHi cells (f) with PAR2 agonism through 48 h. There was a significant (p = 0.0011; DF = 6, 59) trend for interaction of incubation time and SLIGRL concentration in PLo cells and a non-significant (p = 0.0527; DF = 6, 130) trend for interaction in PHi cells. For data in (a) and (b), a Student’s t test was performed. For data in (c, d), a one-way ANOVA was performed, and for data in (e, f), a repeated measures two-way ANOVA was performed including analysis of interaction effects. For data in (c-f), individual comparisons were analyzed using Tukey’s post hoc test. a, bn = 8/group. c, dn = 9–18/group. en = 5–6/group. fn = 11–12/group. *p < 0.05; ****p < 0.0001 as shown.

Close modal

Candidate GPCR-/PAR2-Driven Protein Kinases

Considering the impact of protein kinases on VSM growth control [3, 4, 7‒10], we evaluated candidate protein kinases in PLo and PHi VSM cells under quiescent and growth conditions. No differences in phosphorylated (Ser217/Ser221) MEK1/2 MAPK were observed between PLo and PHi cells (shown in Fig. 4a, b; see also online suppl. material), but PHi cells showed significantly (p = 0.0028) reduced expression of total MEK1/2 compared to PLo cells (shown in Fig. 4c, d). Notably, the phosphorylated to total MEK1/2 ratio (shown in Fig. 4e) was significantly higher (p = 0.0336) in PHi cells compared to PLo cells. Further, PHi cells showed significantly reduced (p = 0.0004) levels of both phosphorylated (Thr197) catalytic PKA-Cα (shown in Fig. 4f, g) and total PKA-Cα (p < 0.0001; shown in Fig. 4h, i) compared to PLo cells, with an unchanged phosphorylated to total PKA-Cα ratio (shown in Fig. 4j). Again, discrete changes in phosphorylated proteins and in total proteins cannot be disregarded (especially in lieu of a phospho/total protein ratio) for their potential biological importance [4, 8, 19, 20]. Lastly, PHi cells exhibited significantly (p = 0.0031) lower expression of both PI3K regulatory p85 (shown in Fig. 4k, l) and PI3K catalytic p110 (p < 0.0001; shown in Fig. 4m, n) compared to PLo cells.

Fig. 4.

Expression of MEK1/2, PKA-Cα, and PI3K in PLo and PHi VSM cells. a Representative Western blot for phosphorylated (Ser217/Ser221) MEK1/2 expression in PLo and PHi VSM cell lysates along with the total protein blot from the same gel. b Densitometry reveals that phosphorylated (Ser217/Ser221) MEK1/2 levels were comparable between PLo and PHi cells; however, PHi cells showed significantly lower (p = 0.0028; DF = 22) levels of total MEK1/2 compared to PLo cells (c, d). The ratio of phosphorylated to total MEK1/2 was significantly increased (p = 0.0336; DF = 22) in PHi versus PLo cells (e). PHi cells showed significantly decreased levels of both phosphorylated (Thr197) PKA-Cα (p = 0.0004; DF = 19; f, g) and total PKA-Cα (p < 0.0001; DF = 21; h, i) compared to PLo cells, with comparable phosphorylated to total PKA-Cα ratios (j). PHi cells also exhibited significantly reduced levels of both regulatory PI3K p85 (p = 0.0031; DF = 22; k, l) and catalytic p110 subunits (p < 0.0001; DF = 17; m, n) compared to PLo cells. Target protein expression was normalized to total protein per sample within each blot for each experiment [17]. For data shown in b, d, e, g, i, j and l, n = 9–12/group, and for N, n = 7–12/group, and a Student’s t test was used for analysis. *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001 as shown.

Fig. 4.

Expression of MEK1/2, PKA-Cα, and PI3K in PLo and PHi VSM cells. a Representative Western blot for phosphorylated (Ser217/Ser221) MEK1/2 expression in PLo and PHi VSM cell lysates along with the total protein blot from the same gel. b Densitometry reveals that phosphorylated (Ser217/Ser221) MEK1/2 levels were comparable between PLo and PHi cells; however, PHi cells showed significantly lower (p = 0.0028; DF = 22) levels of total MEK1/2 compared to PLo cells (c, d). The ratio of phosphorylated to total MEK1/2 was significantly increased (p = 0.0336; DF = 22) in PHi versus PLo cells (e). PHi cells showed significantly decreased levels of both phosphorylated (Thr197) PKA-Cα (p = 0.0004; DF = 19; f, g) and total PKA-Cα (p < 0.0001; DF = 21; h, i) compared to PLo cells, with comparable phosphorylated to total PKA-Cα ratios (j). PHi cells also exhibited significantly reduced levels of both regulatory PI3K p85 (p = 0.0031; DF = 22; k, l) and catalytic p110 subunits (p < 0.0001; DF = 17; m, n) compared to PLo cells. Target protein expression was normalized to total protein per sample within each blot for each experiment [17]. For data shown in b, d, e, g, i, j and l, n = 9–12/group, and for N, n = 7–12/group, and a Student’s t test was used for analysis. *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001 as shown.

Close modal

Protein Kinase Control of PAR2-Mediated VSM Cell Growth

Based on the differential expression between PLo and PHi VSM cells for PAR2-driven kinases and considering that PAR2 agonism promoted growth in PLo cells, kinase-specific blockade followed by SLIGRL treatment and evaluation of cell growth in PLo cells was performed. Cells were first treated with antagonists against MEK1/2 (PD; 10 μm), PKA (PKI; 10 μm), PI3K (LY; 1 μm), or respective vehicles for 24 h (in the absence of SLIGRL) to establish baseline effects of these kinase inhibitors on DNA synthesis (shown in Fig. 5a). PD or PKI alone had no significant effects on DNA synthesis compared to respective vehicle controls; however, LY significantly decreased DNA synthesis compared to its vehicle (p < 0.0001). Next, PLo cells were pretreated with PD, PKI, LY, or vehicle for 30 min prior to treatment with SLIGRL (100 nm) for 24 h. As shown in Figure 5b, cells incubated with PKI or LY concomitant with SLIGRL showed significantly (p < 0.0001 each) decreased DNA synthesis compared to cells treated with SLIGRL alone. PD with SLIGRL markedly (p = 0.0707) decreased DNA synthesis compared to SLIGRL alone. Using automated cell counting with trypan blue exclusion staining after 72 h (shown in Fig. 5c), significant reductions in viable numbers of cells treated with SLIGRL and PD (p = 0.0028) or SLIGRL and LY (p < 0.0001), along with a decreased trend in SLIGRL and PKI cells (p = 0.0769), compared to cells treated with SLIGRL alone, were observed. In experiments using trypan blue exclusion staining and neutral red and MTT assays, none of the individual kinase inhibitors alone, SLIGRL alone, or SLIGRL with any inhibitor caused significant alterations in cell viability (at all concentrations and times used) compared to respective vehicle controls (data not shown).

Fig. 5.

Evaluation of PAR2-stimulated VSM cell growth following specific kinase blockade. In a, cells incubated with the MEK1/2 inhibitor PD or the PKA inhibitor PKI had no significant changes in DNA synthesis, but cells incubated with the PI3K inhibitor PKI had significantly decreased (p < 0.0001; DF = 28) DNA synthesis, compared to respective vehicle controls after 24 h. In b, cells incubated with the PKA inhibitor PKI or the PI3K inhibitor LY concomitant with SLIGRL showed significantly decreased (p < 0.0001; DF = 3, 56) DNA synthesis compared to cells treated with SLIGRL alone after 24 h. PD with SLIGRL nonsignificantly reduced DNA synthesis (p = 0.0707; DF = 3, 56) compared to SLIGRL treatment alone. In c, viable cell numbers were significantly reduced in cells treated with PD and SLIGRL (p = 0.0028; DF = 3, 16) or LY and SLIGRL (p < 0.0001; DF = 3, 16) compared to cells treated with SLIGRL alone. PKI with SLIRGRL nonsignificantly (p = 0.0769; DF = 3, 16) reduced viable cell numbers versus SLIGRL alone. n = 5–6 in triplicate per group, and a one-way ANOVA was performed followed by Tukey’s post hoc tests for multiple comparisons. **p < 0.01; ****p < 0.0001 as shown.

Fig. 5.

Evaluation of PAR2-stimulated VSM cell growth following specific kinase blockade. In a, cells incubated with the MEK1/2 inhibitor PD or the PKA inhibitor PKI had no significant changes in DNA synthesis, but cells incubated with the PI3K inhibitor PKI had significantly decreased (p < 0.0001; DF = 28) DNA synthesis, compared to respective vehicle controls after 24 h. In b, cells incubated with the PKA inhibitor PKI or the PI3K inhibitor LY concomitant with SLIGRL showed significantly decreased (p < 0.0001; DF = 3, 56) DNA synthesis compared to cells treated with SLIGRL alone after 24 h. PD with SLIGRL nonsignificantly reduced DNA synthesis (p = 0.0707; DF = 3, 56) compared to SLIGRL treatment alone. In c, viable cell numbers were significantly reduced in cells treated with PD and SLIGRL (p = 0.0028; DF = 3, 16) or LY and SLIGRL (p < 0.0001; DF = 3, 16) compared to cells treated with SLIGRL alone. PKI with SLIRGRL nonsignificantly (p = 0.0769; DF = 3, 16) reduced viable cell numbers versus SLIGRL alone. n = 5–6 in triplicate per group, and a one-way ANOVA was performed followed by Tukey’s post hoc tests for multiple comparisons. **p < 0.01; ****p < 0.0001 as shown.

Close modal

Protease-activated receptors represent a GPCR subfamily that shows promise for clinical utility in cardiovascular tissue. Of the four identified PARs, PAR1 was the first discovered for its control of platelet function [10], and PAR1 with PAR2 can help control inflammation and proliferation in human colon cancer and embryonic kidney cell lines [21, 22]. In both experimental [23, 24] and clinical [25] settings of vascular disease and injury, PAR2 is proposed to contribute to macrophage inflammation and lipid uptake, fibroblast proliferation, and development of occlusive atherosclerotic plaque; however, PAR2 itself has yet to be defined as an effective regulator of VSM growth, a key component in the response to vascular disease and injury in CVD [3, 4, 8, 19, 20]. Diverse intracellular G protein signaling exists for the PARs [3, 26], with precision of effects relying largely on discrete protein kinases [3, 10, 26, 27], yet the specific kinases involved in PAR2 actions in VSM including those instrumental to CVD and vascular disease and injury are unclear.

Our study aimed to determine regulatory impact and kinase-specific mechanisms of PAR2 on VSM cell growth. We hypothesized that PAR2 controls proliferation of VSM cells in a phenotype-specific manner and reliant upon discrete protein kinases, and we propose that PAR2 and its kinase effectors may represent therapeutic targets against aberrant VSM growth elemental in CVD.

Initial experiments aimed to establish phenotypically distinct VSM cell populations reflective of quiescent and proliferative states. Based on established criteria for distinguishing contractile versus synthetic VSM cell phenotypes [15, 16], we generated PLo VSM cells (P3–P6) with a differentiated, contractile phenotype and PHi VSM cells (P10–P15) with a dedifferentiated, proliferative phenotype typical of an injured or diseased state [28, 29]. To establish objective criteria for defining PLo and PHi cells, we probed cell lysates for documented (PKG-1) [28, 29] and novel (VASP) markers of phenotypic switching. Expression of PKG-1, an established readout of phenotypically altered VSM cells [28, 29], was significantly reduced in PHi cells compared to PLo cells (shown in Fig. 1a, b), supporting phenotypic distinction of our PHi cells from PLo cells. Since PKG phosphorylates VASP at Ser239 [3, 8], we examined total and phosphorylated (Ser239) VASP as a possible marker of phenotypic alteration. We observed significant reductions in both phosphorylated and total VASP in PHi cells compared to PLo cell with an increase in the phosphorylated to total VASP ratio (shown in Fig. 1c–g). To our knowledge, this is the first report showing VASP as a plausible marker of phenotypic switching in VSM cells. These differential expression profiles for PKG-1 and VASP helped establish objective criteria for defining phenotypically unique PLo and PHi VSM cells in our study.

To help validate our use of PLo and PHi cells as respective models of quiescent and proliferative VSM cells, we examined expression of key contractile and cytoskeletal proteins. SM22α and calponin 1, actin-binding proteins required for smooth muscle contractility [30, 31], were both significantly decreased in PHi cells compared to PLo cells (shown in Fig. 2a–d). Expression of both phosphorylated (Ser3) and total cofilin, an actin-binding protein essential for F-actin dynamics [32], was significantly decreased in PHi cells versus PLo cells (shown in Fig. 2e–h). Along with losses of PKG-1 and VASP, reduction in these smooth muscle contractile and differentiation mediators in our PHi cells compared to PLo cells help define them as respective models of proliferative and quiescent VSM cells. Given their similar molecular signatures, our PLo and PHi cell models could also reflect aspects of in vitro vascular aging and cell senescence [33] in addition to phenotypic switching.

To discern impact of PAR2 on PLo versus PHi VSM cell growth, we assessed PAR2 expression and utilized SLIGRL-mediated PAR2 agonism [34] followed by evaluation of growth in PLo and PHi cells. PAR2 expression was significantly decreased in proliferating PHi cells compared to quiescent PLo cells (shown in Fig. 3a, b), suggesting that expression of PAR2 is phenotype-specific and that it operates as a growth inhibitor under basal (unstimulated) conditions. Next, SLIGRL-mediated PAR stimulation increased DNA synthesis in PLo cells but decreased DNA synthesis in PHi cells compared to controls after 24 h (shown in Fig. 3c, d). Control PHi cells showed twice the levels of BrdU absorbance compared to control PLo cells, further supporting the proliferative nature of PHi cells versus the quiescent nature of the PLo cells. Likewise, SLIGRL increased PLo cell numbers but decreased PHi cell numbers compared to controls after 48 h (shown in Fig. 3e, f). These differential growth responses of distinct cell populations to PAR2 agonism support a phenotype-specific manner by which PAR2 controls growth in VSM. Notably, BrdU absorbance and cell numbers after 48 h are higher in the PHi cells compared to PLo cells at baseline conditions without PAR2 agonism. This supports our use of PLo and PHi cells as models of contractile, quiescent and synthetic, proliferative VSM cells, respectively.

Reports linking PAR2 and VSM growth are limited, yet our findings showing PAR2 agonism promotes growth in PLo VSM cells agree somewhat with several other studies [35‒37]. Investigators observed that the PAR2-tethered ligand agonist SLIGKV, differing in its mechanism of PAR activation from that of SLIGRL, induced DNA synthesis in commercial human coronary VSM cells [35]. Key differences in experimental design between our studies, including modes of PAR activation, species, cell types (primary vs. commercial), and assessment of proliferation (time points; sole reliance on DNA synthesis vs. multiple readouts), are notable. Another study [36] transfected PAR2 into commercial human VSM cells and observed that PAR2 increased cyclin E1 and CDK2 and decreased p27, suggesting a pro-growth effect of PAR2. Again, comparable experimental differences between this study and ours warrant caution in comparison. Further, in a murine model of diabetes-induced colon dysmobility [37], PAR2 increased colon length and weight and NIH/3T3 cell proliferation, as measured by immunofluorescence. Once more, important differences in experimental approaches between our studies warrant prudence in comparison.

Our results conflict with a report [38] showing lack of a growth effect from SLIGRL in bovine coronary VSM cells. In addition to differences in species and tissue origin of derived cells, the reported EC50 for SLIGRL is ∼ 5 μm [39], and these opposing results could be due to off-target effects from the high SLIGRL concentration (100 μm) used in that study [38] compared to those (100 nm–10 μm) used in our study. We were unable to identify reports showing growth inhibition by PAR2 in any cell type, as we show here with our PHi VSM cells. Considering that PAR2 is activated by serine proteases, including extracellular trypsin and mast cell tryptase [3, 11] secreted during inflammation [36], our observations of VSM growth following PAR2 agonism, as well as divergent effects of PAR2 on growth in PLo versus PHi cells, have major clinical implications.

Wide-ranging impacts of protein kinases on VSM growth are known [3, 4, 8, 19, 20], and we proposed that PAR2 elicits growth control in VSM cells through specific kinases. PLo and PHi VSM cells displayed comparable levels of phosphorylated (Ser217/Ser221) MEK1/2 MAPK (Fig. 4a, b), which is required for kinase activation [40], yet PHi cells showed significantly lower total MEK1/2 compared (shown in Fig. 4c, d) and a significantly higher phospho/total MEK1/2 ratio (shown in Fig. 4e) compared to PLo cells. PLo cells also showed significantly higher levels of phosphorylated (Thr197, essential for full PKA activation [41]) and total PKA-Cα (shown in Fig. 4f–i versus PHi cells, with comparable levels of phosphorylated to total PKA-Cα between PLo and PHi cells (shown in Fig. 4j). Lastly, significantly lower levels of regulatory p85 and catalytic p110 subunits of PI3K, both required for PI3K activation [42], were found in PHi versus PLo cells (shown in Fig. 4k–n. These observations correlate with diminished PAR2 expression in PHi cells (shown in Fig. 3a, b) and offer evidence that PKA, MEK1/2, and/or PI3K mediate growth effects of PAR2 stimulation in PLo and PHi VSM cells.

In SLIGRL-treated PLo VSM cells, kinase blockade was then used to determine their involvement in the mitogenic actions of PAR2. If kinases were causative for the increased growth seen in PAR2-activated PLo cells, then kinase blockade would reverse those observations and show reduced proliferation, implying a kinase-dependent mechanism of PAR2 growth control. Baseline effects (lacking SLIGRL) of the kinase inhibitors on DNA synthesis were first determined. While no significant impacts on DNA synthesis with the PKA inhibitor PKI or the MEK1/2 inhibitor PD were observed, the PI3K inhibitor LY significantly decreased DNA synthesis compared to vehicle controls (shown in Fig. 5a), consistent with our earlier observations [43]. Compatible with the MEK1/2 expression data (shown in Fig. 4a–e, no significant effects on DNA synthesis were observed in SLIGRL-treated cells with/without PD; however, a significant decrease in viable cell numbers was observed (shown in Fig. 5b, c). Comparatively, SLIGRL-treated cells with PKI showed significantly reduced DNA synthesis compared to SLIGRL controls (shown in Fig. 5b), while SLIGRL-treated cells with LY showed significantly reduced DNA synthesis and viable cell numbers compared to SLIGRL-treated cells (shown in Fig. 5b, c). An aforementioned study [37] found that PAR2 induced NIH/3T3 cell growth in PI3K-dependent manner which, despite differences in cell types and modes of PAR agonism, support our observations of a PI3K-specific mechanism for PAR2 control of VSM growth. However, we believe the observed effects on DNA synthesis and viable cell numbers following LY treatment are likely due to substantial baseline effects of the inhibitor (shown in Fig. 5a) irrespective of its interaction with SLIGRL, and we cannot conclude that PI3K is involved in PAR2 growth control based on these observations. It is likely that PI3K controls cell proliferation and DNA synthesis independent of PAR2 signaling, as we have previously noted [43]. However, our observations argue for a PKA/MEK1/2 mechanism in PAR2 control of VSM cell growth (shown in Fig. 5b, c). While VSM growth control by PKA and by MEK1/2 has been documented [4, 44, 45], alone as well as collaboratively with associated signals such as Epac-1 and Erk1/2, their link to PAR2 signaling, including serving as mechanisms of PAR2-mediated VSM growth control, is unknown. The results of this study indicate that PAR2 activity may be controlled in a phenotype-dependent manner and that PAR2 agonism in synthetic, proliferative VSM cells could potentially inhibit abnormal VSM growth in vascular disease conditions. Our findings also support the use of PLo and PHi VSM cells as models of contractile, quiescent cells and synthetic, proliferative cells, respectively. PLo and PHi VSM cells could be used in future studies to further understand phenotypic switching seen in altered VSM in CVD.

Several limitations became apparent during our study that warrant discussion. Female rats were not used as sources of primary cells due to potential influence of female hormones such as estrogen on VSM growth [14]; nonetheless, full evaluation of PAR2 and its impact on cell growth mandates consideration of both male and female donors to identify potential sex-dependent effects. Our primary cells were harvested from the thoracic aorta, a large conduit vessel that serves as a suitable source for primary cell culture in basic science [4, 8, 16, 19, 20, 46] and clinical investigation [47, 48]; however, the reader should be aware of important physiological differences in cells derived from small resistance vessels versus those from conduit vessels [49, 50]. Lastly, for complete understanding of the functional capacities of PAR2 (and other GPCRs), one should consider complementary experiments using PAR2 antagonism as well as cooperativity observed for PAR2 with PAR1 during inflammation and proliferation [21, 22] and with its close homolog, PAR4, identified in membrane trafficking and related cellular functions [51].

In this study, we hypothesized that PAR2 controls VSM cell growth in a phenotype-specific manner dependent upon specific protein kinases, and, in turn, PAR2 and its kinase effectors may represent potential therapeutic targets against pathologic VSM growth. Our chief findings support our hypothesis and include objective distinction of PLo from PHi VSM cells through reduced expression of PKG and VASP, confirmed by loss of SM22α, calponin 1, and cofilin, in PHi versus PLo cells; decreased expression of PAR2 in PHi cells versus PLo cells; PAR2 agonism promoting growth in PLo cells yet suppressing growth in PHi cells; reduction of MEK1/2, PKA, and PI3K in PHi cells compared to PLo cells; and pharmacologic blockade of PKA and MEK1/2 reversing growth induction by PAR2 in PLo cells. Given the complexity and context-dependency of these actions of PAR2 in VSM cell growth, prudence in interpretation should be heeded, and additional studies are certainly warranted; nonetheless, our findings shed light on the pathophysiological importance and therapeutic promise of PAR2 and PAR2-driven PKA/MEK1/2 in VSM cell growth control in CVD.

The authors would like to thank the academic and support staff of the East Carolina University Department of Physiology for their assistance in processing this manuscript. The authors would also like to thank the many investigators who have contributed significantly to cardiovascular science research but whose works were not cited in this article due to formatting constraints.

All protocols and procedures employed in this study were reviewed and approved by the East Carolina University Animal Care and Use Committee and conformed to the guidelines described in the Guide for the Care and Use of Laboratory Animals [12]. In vivo studies were designed according to the Animal Research: Reporting In Vivo Experiments (ARRIVE) 2.0 Essential 10 guidelines [13].

The authors declare that this research was conducted in the absence of any commercial or financial relationships that could be construed as potential conflicts of interest.

This work was supported in part by NIH grants R01HL81720 (DAT), R01HL81720-05S1 (DAT), and R15HL135699 (DAT), an ECU Brody School of Medicine Seed/Bridge Grant (DAT), and a Brody Brothers Endowment Fund Award (DAT). This content is solely the responsibility of the authors and does not necessarily represent the official views of the NIH, ECU, or the Brody Brothers Endowment Fund.

All authors made substantial, direct, and intellectual contributions to the work and have approved it for publication, and all authors are eligible for authorship on this manuscript. Specifically, Madison D. Williams, Michael T. Bullock, Sean C. Johnson, Nathan A. Holland, Danielle M. Vuncannon, Joani Zary Oswald, Shaquria P. Adderley, and David A. Tulis designed and performed experiments; analyzed and interpreted data; performed statistical analyses; prepared figures; approved the final manuscript; and hold accountability for accuracy and integrity of the work. Additionally, Madison D. Williams, Nathan A. Holland, and David A. Tulis drafted the manuscript and reviewed and revised the manuscript to its final form.

1.
Tsao
CW
,
Aday
AW
,
Almarzooq
ZI
,
Anderson
CAM
,
Arora
P
,
Avery
CL
.
Heart disease and stroke statistics: 2023 update. A report from the American heart association
.
Circulation
.
2023
;
147
(
8
):
e93
621
.
2.
World Health Organization [Internet]
. World Health Statistics 2021: Monitoring Health for the SDGs 2021 [cited 2022 March 15]. Available from: https://www.who.int/publications/i/item/9789240027053.
3.
Holland
N
,
Francisco
J
,
Johnson
S
,
Morgan
JS
,
Dennis
TJ
,
Gadireddy
NR
.
Cyclic nucleotide-directed protein kinases in cardiovascular inflammation and growth
.
J Cardiovasc Dev Dis
.
2018
;
5
(
1
):
6
.
4.
Adderley
SP
,
Joshi
CN
,
Martin
DN
,
Mooney
S
,
Tulis
DA
.
Multiple kinase involvement in the regulation of vascular growth
. In:
Da Silva Xavier
G
, editor.
Advances in protein kinases
InTech Open Access Publishers
2012
. p.
131
50
.
5.
Beamish
JA
,
He
P
,
Kottke-Marchant
K
,
Marchant
RE
.
Molecular regulation of contractile smooth muscle cell phenotype: implications for vascular tissue engineering
.
Tissue Eng Part B Rev
.
2010
;
16
(
5
):
467
91
.
6.
Thyberg
J
,
Hedin
U
,
Sjölund
M
,
Palmberg
L
,
Bottger
BA
.
Regulation of differentiated properties and proliferation of arterial smooth muscle cells
.
Arteriosclerosis
.
1990
;
10
(
6
):
966
90
.
7.
Hauser
AS
,
Attwood
MM
,
Rask-Andersen
M
,
Schioth
HB
,
Gloriam
DE
.
Trends in GPCR drug discovery: new agents, targets, and indications
.
Nat Rev. Disc
.
2017
;
16
(
12
):
829
42
.
8.
Holt
AW
,
Martin
DN
,
Shaver
PR
,
Adderley
SP
,
Stone
JD
,
Joshi
CN
.
Soluble guanylyl cyclase-activated cyclic GMP-dependent protein kinase inhibits arterial smooth muscle cell migration independent of VASP-Serine 239 phosphorylation
.
Cell Signal
.
2016
;
28
(
9
):
1364
79
.
9.
Leger
AJ
,
Covic
L
,
Kuliopulos
A
.
Protease-activated receptors in cardiovascular diseases
.
Circulation
.
2006
;
114
(
10
):
1070
7
.
10.
Hirano
K
.
The roles of proteinase-activated receptors in the vascular physiology and pathophysiology
.
Arterioscler Thromb Vasc Biol
.
2007
;
27
(
1
):
27
36
.
11.
Coelho
AM
,
Ossovskaya
V
,
Bunnett
NW
.
Proteinase-activated receptor-2: physiological and pathophysiological roles
.
Curr Med Chem Cardiovasc Hematol Agents
.
2003
;
1
:
61
72
.
12.
National Research Council (NRC)
Guide for the Care and use of laboratory Animals
8th Ed
Washington, DC
The National Academies Press
2011
.
13.
Percie du Sert
N
,
Ahluwalia
A
,
Alam
S
,
Avey
MT
,
Baker
M
,
Browne
WJ
.
Reporting animal research: explanation and elaboration for the ARRIVE guidelines 2.0
.
PLoS Biol
.
2020
;
18
(
7
):
e3000411
.
14.
Yuan
Y
,
Liao
L
,
Tulis
DA
,
Xu
J
.
Steroid receptor coactivator-3 is required for inhibition of neointima formation by estrogen
.
Circulation
.
2002
;
105
(
22
):
2653
9
.
15.
Nakano-Kurimoto
R
,
Ikeda
K
,
Uraoka
M
,
Nakagawa
Y
,
Yutaka
K
,
Koide
M
.
Replicative senescence of vascular smooth muscle cells enhances the calcification through initiating the osteoblastic transition
.
Am J Physiol Heart Circ Physiol
.
2009
297
5
H1673
84
.
16.
Chang
S
,
Song
S
,
Lee
J
,
Yoon
J
,
Park
J
,
Choi
S
.
Phenotypic modulation of primary vascular smooth muscle cells by short-term culture on micropatterned substrate
.
PLoS One
.
2014
;
9
(
2
):
e88089
.
17.
Brooks
HL
,
Lindsey
ML
.
Guidelines for authors and reviewers on antibody use in physiology studies
.
Am J Physiol Heart Circ Physiol
.
2018
314
4
H724
32
.
18.
Lindsey
ML
,
Iyer
RP
,
Zamilpa
R
,
Yabluchanskiy
A
,
DeLeon-Pennell
KY
,
Hall
ME
.
A novel collagen matricryptin reduces left ventricular dilation post-myocardial infarction by promoting scar formation and angiogenesis
.
J Am Coll Cardiol
.
2015
;
66
(
12
):
1364
74
.
19.
Mendelev
NN
,
Williams
VS
,
Tulis
DA
.
Antigrowth properties of BAY 41-2,272 in vascular smooth muscle cells
.
J Cardiovasc Pharmacol
.
2009
;
53
(
2
):
121
31
.
20.
Joshi
CN
,
Martin
DN
,
Fox
JC
,
Mendelev
NN
,
Brown
TA
,
Tulis
DA
.
The soluble guanylate cyclase stimulator BAY 41-2272 inhibits vascular smooth muscle growth through the cAMP-dependent protein kinase and cGMP-dependent protein kinase pathways
.
J Pharmacol Exp Ther
.
2011
;
339
(
2
):
394
402
.
21.
Hu
L
,
Xia
L
,
Zhou
H
,
Wu
B
,
Mu
Y
,
Wu
Y
.
TF/FVIIa/PAR2 promotes cell proliferation and migration via PKCα and ERK-dependent c-Jun/AP-1 pathway in colon cancer cell line SW620
.
Tumour Biol
.
2013
;
34
(
5
):
2573
81
.
22.
Fan
L
,
Yotov
WV
,
Zhu
T
,
Esmailzadeh
L
,
Joyal
JS
,
Sennlaub
F
.
Tissue factor enhances protease-activated receptor-2-mediated factor VIIa cell proliferative properties
.
J Thromb Haemost
.
2005
;
3
(
5
):
1056
63
.
23.
D’Andrea
M
,
de Garavilla
L
,
Cheung
WM
,
Andrade-Gordon
P
,
Damiano
B
.
Increased expression of protease activated receptor-2 (PAR-2) in balloon-injured rat carotid artery
.
Thromb Haemost
.
1999
;
81
(
05
):
808
14
.
24.
Hara
T
,
Phuong
PT
,
Fukuda
D
,
Yamaguchi
K
,
Murata
C
,
Nishimoto
S
.
Protease-activated receptor-2 plays a critical role in vascular inflammation and atherosclerosis in apolipoprotein E-deficient mice
.
Circulation
.
2018
;
138
(
16
):
1706
19
.
25.
Napoli
C
,
de Nigris
F
,
Wallace
JL
,
Hollenberg
MD
,
Tajana
G
,
De Rosa
G
.
Evidence that protease activated receptor 2 expression is enhanced in human coronary atherosclerotic lesions
.
J Clin Pathol
.
2004
;
57
(
5
):
513
6
.
26.
Sanderlin
EJ
,
Justus
CR
,
Krewson
EK
,
Li
VY
.
Emerging roles for the pH-sensing G protein-coupled receptors in response to acidotic stress
.
Cell Health Cytoskelet
.
2015
;
7
:
99
109
.
27.
Sheng
J
,
Deng
X
,
Zhang
Q
,
Liu
H
,
Wang
N
,
Liu
Z
.
PAR-2 promotes invasion and migration of esophageal cancer cells by activating MEK/ERK and PI3K/Akt signaling pathway
.
Int J Clin Exp Pathol
.
2019
;
12
(
3
):
787
97
.
28.
Boerth
NJ
,
Dey
NB
,
Cornwell
TL
,
Lincoln
TM
.
Cyclic GMP-dependent protein kinase regulates vascular smooth muscle cell phenotype
.
J Vasc Res
.
1997
;
34
(
4
):
245
59
.
29.
Anderson
PG
,
Boerth
NJ
,
Liu
M
,
McNamara
DB
,
Cornwell
TL
,
Lincoln
TM
.
Cyclic GMP-dependent protein kinase expression in coronary arterial smooth muscle in response to balloon catheter injury
.
Arterioscler Thromb Vasc Biol
.
2000
;
20
(
10
):
2192
7
.
30.
Han
M
,
Dong
LH
,
Zheng
B
,
Shi
JH
,
Wen
JK
,
Cheng
Y
.
Smooth muscle 22 alpha maintains the differentiated phenotype of vascular smooth muscle cells by inducing filamentous actin bundling
.
Life Sci
.
2009
84
13–14
394
401
.
31.
Gimona
M
,
Herzog
M
,
Vandekerckhove
J
,
Small
JV
.
Smooth muscle specific expression of calponin
.
FEBS Lett
.
1990
274
1–2
159
62
.
32.
Albinsson
S
,
Nordström
I
,
Hellstrand
P
.
Stretch of the vascular wall induces smooth muscle differentiation by promoting actin polymerization
.
J Biol Chem
.
2004
;
279
(
33
):
34849
55
.
33.
Uryga
AK
,
Bennett
MR
.
Ageing induced vascular smooth muscle cell senescence in atherosclerosis
.
J Physiol
.
2016
;
594
(
8
):
2115
24
.
34.
Villari
A
,
Giurdanella
G
,
Bucolo
C
,
Drago
F
,
Salomone
S
.
Apixaban enhances vasodilatation mediated by protease-activated receptor 2 in isolated rat arteries
.
Front Pharmacol
.
2017
;
8
:
480
.
35.
Indrakusuma
I
,
Romacho
T
,
Eckel
J
.
Protease-activated receptor 2 promotes pro-atherogenic effects through transactivation of the VEGF receptor 2 in human vascular smooth muscle cells
.
Front Pharmacol
.
2016
;
7
:
497
.
36.
Wei
M
,
Liu
Y
,
Zheng
M
,
Wang
L
,
Ma
F
,
Qi
Y
.
Upregulation of protease-activated receptor 2 promotes proliferation and migration of human vascular smooth muscle cells (VSMCs)
.
Med Sci Monit
.
2019
;
25
:
8854
62
.
37.
Li
YJ
,
Ao
JP
,
Huang
X
,
Lu
HL
,
Fu
HY
,
Song
NN
.
Involvement of PAR2 in platelet-derived growth factor receptor-α-positive cell proliferation in the colon of diabetic mice
.
Physiol Rep
.
2021
;
9
(
21
):
e15099
.
38.
Bretschneider
E
,
Kaufmann
R
,
Braun
M
,
Wittpoth
M
,
Glusa
E
,
Nowak
G
.
Evidence for proteinase-activated receptor-2 (PAR-2)-mediated mitogenesis in coronary artery smooth muscle cells
.
Br J Pharmacol
.
1999
;
126
(
8
):
1735
40
.
39.
Nystedt
S
,
Emilsson
K
,
Wahlestedt
C
,
Sundelin
J
.
Molecular cloning of a potential proteinase activated receptor
.
Proc Natl Acad Sci U S A
.
1994
;
91
(
20
):
9208
12
.
40.
Cowley
S
,
Paterson
H
,
Kemp
P
,
Marshall
CJ
.
Activation of MAP kinase kinase is necessary and sufficient for PC12 differentiation and for transformation of NIH 3T3 cells
.
Cell
.
1994
;
77
(
6
):
841
52
.
41.
Moore
MJ
,
Kanter
JR
,
Jones
KC
,
Taylor
SS
.
Phosphorylation of the catalytic subunit of protein kinase A. Autophosphorylation versus phosphorylation by phosphoinositide-dependent kinase-1
.
J Biol Chem
.
2002
;
277
(
49
):
47878
84
.
42.
Neri
LM
,
Borgatti
P
,
Capitani
S
,
Martelli
AM
.
The nuclear phosphoinositide 3-kinase/AKT pathway: a new second messenger system
.
Biochim Biophys Acta
.
2002
1584
2–3
73
80
.
43.
Adderley
SP
,
Martin
DN
,
Tulis
DA
.
Exchange protein activated by cAMP (EPAC) controls migration of vascular smooth muscle cells in concentration and time-dependent manner
.
Arch Physiol
.
2015
;
2
(
1
):
2
.
44.
Jiang
S
,
Tang
DD
.
Plk1 regulates MEK1/2 and proliferation in airway smooth muscle cells
.
Respir Res
.
2015
;
16
(
1
):
93
.
45.
Hewer
RC
,
Sala-Newby
GB
,
Wu
Y-J
,
Newby
AC
,
Bond
M
.
PKA and Epac synergistically inhibit smooth muscle cell proliferation
.
J Mol Cell Cardiol
.
2011
;
50
(
1
):
87
98
.
46.
Xu
S
,
Fu
J
,
Chen
J
,
Xiao
P
,
Lan
T
,
Le
K
.
Development of an optimized protocol for primary culture of smooth muscle cells from rat thoracic aortas
.
Cytotechnology
.
2009
61
1–2
65
72
.
47.
Bogunovic
N
,
Meekel
JP
,
Majolee
J
,
Hekhuis
M
,
Pyszkowski
J
,
Jockenhövel
S
.
Patient-specific 3-dimensional model of smooth muscle cell and extracellular matrix dysfunction for the study of aortic aneurysms
.
J Endovasc Ther
.
2021
;
28
(
4
):
604
13
.
48.
Mizrak
D
,
Feng
H
,
Yang
B
.
Dissecting the heterogeneity of human thoracic aortic aneurysms using single-cell transcriptomics
.
Arterioscler Thromb Vasc Biol
.
2022
;
42
(
8
):
919
30
.
49.
Matsuki
T
,
Hynes
MR
,
Duling
BR
.
Comparison of conduit vessel and resistance vessel reactivity: influence of intimal permeability
.
Am J Physiol
.
1993
264
4 Pt 2
H1251
8
.
50.
Archer
SL
,
Huang
JMC
,
Reeve
HL
,
Hampl
V
,
Tolarová
S
,
Michelakis
E
.
Differential distribution of electrophysiologically distinct myocytes in conduit and resistance arteries determines their response to nitric oxide and hypoxia
.
Circ Res
.
1996
;
78
(
3
):
431
42
.
51.
Cunningham
MR
,
McIntosh
KA
,
Pediani
JD
,
Robben
J
,
Cooke
AE
,
Nilsson
M
.
Novel role for proteinase-activated receptor 2 (PAR2) in membrane trafficking of proteinase-activated receptor 4 (PAR4)
.
J Biol Chem
.
2012
;
287
(
20
):
16656
69
.