Abstract
Pulmonary arterial hypertension (PAH) is characterized by elevated pulmonary arterial pressure and carries a very poor prognosis. Understanding of PAH pathogenesis is needed to support the development of new therapeutic strategies. Transforming growth factor β (TGF-β) drives vascular remodeling and increases vascular resistance by regulating differentiation and proliferation of smooth muscle cells (SMCs). Also, sphingosine-1-phosphate (S1P) has been implicated in PAH, but the relation between these two signaling mechanisms is not well understood. Here, we characterize the signaling networks downstream of TGF-β in human pulmonary arterial smooth muscle cells (HPASMCs), which involves mothers against decapentaplegic homolog (SMAD) signaling as well as Rho GTPases. Activation of Rho GTPases regulates myocardin-related transcription factor (MRTF) and serum response factor (SRF) transcription activity and results in upregulation of contractile gene expression. Our genetic and pharmacologic data show that in HPASMCs upregulation of α smooth muscle actin (αSMA) and calponin by TGF-β is dependent on both SMAD and Rho/MRTF-A/SRF transcriptional mechanisms.
The kinetics of TGF-β–induced myosin light chain (MLC) 2 phosphorylation, a measure of RhoA activation, are slow, as is regulation of the Rho/MRTF/SRF-induced αSMA expression. These results suggest that TGF-β1 activates Rho/phosphorylated MLC2 through an indirect mechanism, which was confirmed by sensitivity to cycloheximide treatment. As a potential mechanism for this indirect action, TGF-β1 upregulates mRNA for sphingosine kinase (SphK1), the enzyme that produces S1P, an upstream Rho activator, as well as mRNA levels of the S1P receptor (S1PR) 3. SphK1 inhibitor and S1PR3 inhibitors (PF543 and TY52156/VPC23019) reduce TGF-β1–induced αSMA upregulation. Overall, we propose a model in which TGF-β1 activates Rho/MRTF-A/SRF by potentiating an autocrine/paracrine S1P signaling mechanism through SphK1 and S1PR3.
SIGNIFICANCE STATEMENT In human pulmonary arterial smooth muscle cells, transforming growth factor β depends on sphingosine-1-phosphate signaling to bridge the interaction between mothers against decapentaplegic homolog and Rho/myocardin-related transcription factor (MRTF) signaling in regulating α smooth muscle actin (αSMA) expression. The Rho/MRTF pathway is a signaling node in the αSMA regulatory network and is a potential therapeutic target for the treatment of pulmonary arterial hypertension.
Introduction
Pulmonary arterial hypertension (PAH) is characterized by elevated mean pulmonary arterial blood pressure and right heart failure, which often leads to death. In the United States, PAH has a prevalence of 12.4 cases per million people (Burger et al., 2016). Patients with PAH are diagnosed at a mean age of 50 ± 14, and their three-year survival rate is 63% (McGoon and Miller, 2012; Leopold and Maron, 2016). The standard of care for PAH is focused on alleviating symptoms but fails to stop disease progression. To develop therapeutic approaches that halt PAH progression, it is critical to better understand the cellular and molecular mechanisms underlying disease progression.
Both vasoconstriction and vascular remodeling contribute to increased vascular resistance in the pulmonary circulation, ultimately leading to elevated blood pressure (Vaillancourt et al., 2015). Vasoconstriction results from the contraction of smooth muscle cells (SMCs) (Vaillancourt et al., 2015). Dysregulated proliferation, migration, and hypertrophy of SMCs contribute to vascular remodeling (Vaillancourt et al., 2015), which involves transforming growth factor β (TGF-β) signaling in Pulmonary Artery Smooth Muscle Cells (PASMCs) (Morrell et al., 2001). Also, TGF-β is elevated in the serum of patients with PAH (Yan et al., 2016). TGF-β signaling is one of the main drivers of aberrant SMC behavior in PAH (Rol et al., 2018).
Generally, smooth muscle cells are categorized into two mutually exclusive phenotypes. The contractile phenotype has high expression of contractile proteins, has a lower proliferation rate, and is less migratory (Gomez and Owens, 2012). The proliferative phenotype has low expression of contractile proteins and an elevated rate of proliferation and migration (Gomez and Owens, 2012). In PAH, PASMCs have both increased proliferation and elevated levels of contractile proteins, such as α smooth muscle actin (αSMA) (Zabini et al., 2018). Interestingly, TGF-β promotes differentiation of PASMCs isolated from nondiseased lungs but paradoxically drives proliferation of PASMCs isolated from patients with PAH (Morrell et al., 2001). Further investigation of how TGF-β regulates the proliferation and differentiation of PASMCs is important for understanding the mechanism of this dual proliferative and contractile SMC phenotype in PAH pathogenesis.
TGF-β regulates gene expression through mothers against decapentaplegic homolog (SMAD) 2/3, which is generally considered to be the canonical signaling pathway (Xing et al., 2015). TGF-β also regulates gene expression through SMAD-independent mechanisms, such as Rho/myocardin-related transcription factor (MRTF)/serum response factor (SRF) (Xing et al., 2015). This starts with the activation of RhoA subfamily small GTPases, which, in turn, induces actin polymerization. Actin polymerization drives the transcriptional coactivator MRTF to translocate into the nucleus, where MRTF binds to SRF to regulate gene expression (Guo and Chen, 2012). The suites of genes regulated by SMAD and Rho/MRTF/SRF pathways overlap in the modulation of fibrosis and cellular migration by TGF-β, suggesting that these two transcriptional mechanisms may cooperate to regulate gene transcription (He and Dai, 2015). Both SMAD and Rho/MRTF/SRF upregulate the expression of contractile genes and markers of differentiation, such as αSMA (Guo and Chen, 2012). Elevated expression of contractile genes results in increased SMC contractility (Bai et al., 2020), which could result in excessive vasoconstriction and SMC hypertrophy in PAH. During myofibroblast differentiation in mouse fibroblast 10T1/2 cells, SMAD3 interacts directly with an SRF-associated complex and mediates TGF-β–induced expression of SM22, another contractile protein (Qiu et al., 2003). It is unknown whether a similar mechanism applies to the transcriptional regulation of αSMA and calponin 1 (CNN1) in human pulmonary arterial smooth muscle cells (HPASMCs).
It is also unclear how TGF-β activates Rho/MRTF/SRF. In human-derived fibroblasts, TGF-β increases the level of the Rho activator sphingosine-1-phosphate (S1P) by upregulating its synthetic enzyme sphingosine kinase 1 (SphK1) (Yamanaka et al., 2004). This suggests a model wherein TGF-β activates Rho signaling through S1P. S1P is a bioactive sphingolipid which binds to a GPCR family of S1P receptors, activates RhoA, and stimulates expression of αSMA and CNN1 in SMCs (Wamhoff et al., 2008). SMAD3 activation is responsible for SphK1 upregulation in C2C12 myoblasts (Cencetti et al., 2010). It is possible that TGF-β upregulates SphK1 levels through an SMAD pathway in HPASMCs. This in turn could promote S1P synthesis and activate Rho/MRTF/SRF in an autocrine/paracrine manner. Consistent with this, plasma S1P levels are increased in patients with idiopathic PAH and in a rodent model of PAH (Gairhe et al., 2016). Genetic and pharmacologic inhibition of SphK1 activity is protective in PAH animal models (Chen et al., 2014; Gairhe et al., 2016), highlighting the importance of S1P signaling in PAH. Further clarifying the signaling interaction between the TGF-β and the S1P pathways will increase our understanding of the molecular mechanisms underlying PAH pathogenesis.
In this study we explore crosstalk mechanisms between SMAD signaling and Rho/MRTF/SRF signaling to better understand how TGF-β modulates SMCs. We provide evidence that S1P signaling bridges the SMAD and Rho/MRTF/SRF pathways to coregulate gene expression in HPASMCs. Our data also suggest that this is primarily mediated by S1P receptor (S1PR) 3.
Materials and Methods
Cell Culture
Human pulmonary artery smooth muscle cells (ThermoFisher, Waltham, MA; C0095C) were cultured in Medium 231 (ThermoFisher; M231500) supplemented with smooth muscle growth supplement (SMGS; ThermoFisher; S00725) and 1% antibiotic-antimycotic (ThermoFisher; 15240062). HPASMCs (passage 6-8) were starved in 0.1% SMGS Medium 231 overnight prior to any experiments.
Compounds and Antibodies
Recombinant human TGF-β1 protein was purchased from Research And Diagnostic Systems, Inc. (Minneapolis, MN). Y-27632 (S1049) was purchased from Selleckchem, Houston, TX. SIS3 (15945), JTE-013 (10009458), TY 52156 (19119), VPC23019 (13240), and PF-543 (17034) were purchased from Cayman Chemical (Ann Arbor, MI). All compounds were dissolved in DMSO and frozen at −20°C. Antibodies against MRTF-A (sc21558) and MRTF-B (sc98989) were purchased from Santa Cruz Biotechnology (Dallas, TX). Antibodies against MRTF-A (14760), Smad2/3 (8685), phosphorylated myosin light chain (pMLC) 2 (3674), and myosin light chain (MLC) 2 (#3672) were ordered from Cell Signaling (Danvers, MA). αSMA antibody (7817) and phosphorylated SMAD3 antibody (52903) were purchased from Abcam (Cambridge, MA), and CNN1 antibody (13938-1-AP) was purchased from Proteintech (Rosemont, IL). All secondary antibodies [Donkey anti-Mouse680 (C31216-02), Donkey anti-Mouse800 (C90507-03), Donkey anti-Goat680 (C41105-05), Donkey anti-Rabbit680 (C40130-02), and Donkey anti-Rabbit800 (C90129-05)] were all purchased from LI-COR (Lincoln, NE). The MRTF/SRF pathway inhibitor CCG-222740 (Hutchings et al., 2017) was obtained from the laboratory of Dr. Scott Larsen at the University of Michigan.
Small Interfering RNA Transfection
ON-TARGETplus small interfering RNA (siRNA) for MRTF-A (Dharmacon L-015434-00-0010; Lafayette, CO), MRTF-B siRNA (Dharmacon L-019279-00-0010), and nontargeting pool control (Dharmacon D-001810-10-05) were used based on the manufacturer’s protocol. siRNAs were diluted in OptiMEM, mixed with DharmaFECT (Dharmacon T-2001-01), and then mixed with fresh medium 231 with 5% SMGS at a final concentration of 25 nM. Cells were seeded at a density of ∼80% confluence and were transfected overnight. The next day, the cells were serum-starved for 16-20 hours prior to the treatment with TGF-β.
Reverse transcription-quantative polymerase chain reaction (RT-qPCR)
HPASMCs were resuspended in complete medium, and 180,000 cells were seeded in each well of a 6-well plate. The cells were allowed to reach confluence (approximately 4 days) before being serum-starved in 0.1% SMGS Medium 231 overnight. Cells were treated as described in the figure legends, and total cellular RNA was collected using the RNeasy kit (Qiagen, Hilden, Germany; 74104) according to the manufacturer’s protocol. The High-Capacity cDNA RT kit (ThermoFisher; 4368814) was used to reverse transcribe the RNA into cDNA following the manufacturer’s protocol. SYBR Green PCR Master Mix (ThermoFisher; 4309155) was used to perform quantitative polymerase chain reaction following the manufacturer’s protocol on the Stratagene Mx3000P machine. Fold change of gene expression was normalized to GAPDH and analyzed by the ΔΔCT method. Primer sequences are listed in Supplemental Table 1.
Immunoblotting
HPASMCs were cultured and treated as described in the figure legends. Total cellular protein was collected in 2× Laemmli Sample Buffer (Biorad, Hercules, CA; 1610737). After heating the samples at 100°C for 10 minutes, protein samples were resolved on 10% (MRTF) or 12% (pMLC2/MLC2) polyacrylamide gels and transferred to PVDF membranes (Millipore Sigma, Burlington, MA; IPFL00010). Blots were blocked in Odyssey Blocking buffer in PBS (LI-COR; 927-40000) at room temperature for 1 hour and then incubated with primary antibody at room temperature for 1 hour or overnight at 4°C. Blots were washed three times for 5 minutes each with Tris-buffered saline with 0.1% Tween-20 (TBST) and then incubated with the appropriate secondary antibodies diluted 1:10,000 in blocking buffer at room temperature for 1 hour. After 3 washes for 5 minutes each with TBST, blots were imaged using a LI-COR Odyssey FC instrument and analyzed using Image Studio Lite software v.4.0.
Cell Proliferation
HPASMCs were cultured and were seeded in 96-well plates at a density of 10,000 cells per well. The next day, the cells were starved in 1% SMGS Medium 231 for ∼14–18 hours. Then the cells were cultured in 1% SMGS Medium 231 with or without 10 ng/mL TGF-β1 for 2 days. Cells were then fixed with 3.7% formaldehyde for 10 minutes. After three washes with PBS for 5 minutes, cells were stained with 500 ng/mL 4',6-diamidino-2-phenylindole. Images were captured at the center of each well using a Cytation 3 automated microscope (Biotek). All images were blinded by an automated R script before quantifying the cell numbers with Image J.
Statistical Analysis
Data were analyzed through either paired t test or one-way repeated-measures ANOVA followed by Dunnett’s post-test using GraphPad Prism 7. Data are presented as the mean ± S.D., and P < 0.05 is considered statistically significant.
Results
Both SMAD and Rho/MRTF/SRF Pathways Are Necessary for TGF-β–Induced Contractile Gene Expression
To identify the mechanism by which TGF-β1 regulates expression of the contractile proteins αSMA and CNN1 in HPASMCs, we first inhibited phosphorylation of SMAD3 using SIS3. TGF-β1–induced αSMA and CNN1 expression was reduced to control levels by 10 μM SIS3 (Fig. 1A). This suggests that phosphorylation of SMAD3 is important for TGF-β1–induced contractile gene expression. To test the role of Rho/MRTF/SRF in regulating contractile gene expression, we used the ROCK inhibitor Y27632 and the MRTF/SRF pathway inhibitor CCG-222740 (Hutchings et al., 2017). Y27632 and CCG-222740 reduced TGF-β1–induced expression of contractile genes by approximately 60% and 100% respectively (Fig. 1, A and B). Finally, siRNA-mediated silencing of MRTF-A reduced TGF-β1–induced protein levels of αSMA; however, there was only a minor reduction upon MRTF-B silencing. (Fig. 1C). Taken together, these data show that both SMAD and the Rho/MRTF-A/SRF pathways are required for TGF-β1–induced contractile gene expression.
TGF-β1–induced αSMA and CNN1 expression is dependent on both the SMAD pathway and the Rho/MRTF/SRF pathway. HPASMCs were seeded into 6-well plates (for the small interfering RNA MRTF protein experiment, cells were seeded into 60-mm petri dish) and allowed to reach confluence. Then the cells were serum-deprived in 0.1% SMGS Medium 231 overnight before being treated the next day. (A) Cells were pretreated with 10 μM SMAD3 inhibitor SIS3 for 1 hour and then along with 10 ng/ml TGF-β1 for another 12 hours. Cells were also concurrently treated with 10 μM ROCK inhibitor Y-27632 and 10 ng/ml TGF-β1 for 24 hours. DMSO was used as a vehicle control. After treatment, total cellular RNA was extracted, and RT-qPCR was used to measure αSMA and CNN1 expression. (B) HPASMCs were cotreated with 10 ng/ml TGF-β1 and 10 μM CCG-222740 or vehicle. Both mRNA and protein were collected to measure the expression levels of αSMA and CNN1. (C) HPASMCs were transfected with 5 μM siMRTF-A or siMRTF-B as described in Materials and Methods. The cells were then starved as described above before being treated with 10 ng/ml TGF-β1 for 24 hours. Protein (N = 3 Santa Cruz anti-MRTF-A, N = 1 Cell Signaling anti-MRTF-A) levels of αSMA, MRTFs, and GAPDH were examined (N ≥ 3. Mean with S.D. Paired t test * P < 0.05, ** P < 0.01).
TGF-β Indirectly Activates Rho Signaling
To better understand how TGF-β1 activates Rho signaling in HPASMCs, we first measured the kinetics of TGF-β1–induced phosphorylation of MLC2, a commonly used readout of Rho activation (Yu et al., 2017). To assess the kinetics of TGF-β1–induced MLC2 phosphorylation, HPASMCs were treated with TGF-β1 for 1–9 hours; pMLC2 was increased only after 6-9 hours (Fig. 2B). In contrast, S1P-induced MLC2 phosphorylation was maximal after 0.5–1 hours of treatment (Fig. 2A). S1P is a GPCR agonist that signals through G12/13, which can rapidly activate RhoA and induce MLC2 phosphorylation (Ambesi and McKeown-Longo, 2009). The delayed kinetics of TGF-β1–induced MLC2 phosphorylation suggest that the action of TGF-β1 is through an indirect and perhaps transcriptional/translational signaling mechanism. Consistent with this hypothesis, cycloheximide blocked TGF-β1–induced MLC2 phosphorylation but did not suppress and even slightly enhanced S1P-induced MLC2 phosphorylation (Fig. 2). Thus, TGF-β1–induced Rho signaling activation, as indirectly detected by pMLC2 levels, requires the translation of new proteins (Fig. 2B). In contrast, TGF-β1–induced SMAD3 phosphorylation peaked at 1 hour and was not blocked by cycloheximide, consistent with the expected direct activation of SMAD3 phosphorylation by the TGF-β receptor (Supplemental Fig. 1).
TGF-β1 indirectly activates the Rho signaling. HPASMCs were seeded into 6-well plates and allowed to reach confluence before being serum-deprived in 0.1% SMGS Medium 231 overnight. The next day the cells were treated with 10 ng/mL cycloheximide or vehicle and were stimulated with: A)1 μM S1P for 0.5, 1, 3, or 6 hours or B) 10 ng/ml TGF-β1 for 1, 3, 6, or 9 hours. Protein was extracted, and the levels of pMLC, total MLC, and GAPDH were measured. The ratio of pMLC/MLC is compared (N = 5. Data are represented as mean with S.D. Paired t test * P < 0.05).
TGF-β Induces mRNA Expression of SphK1
Given our hypothesis that TGF-β1 induces Rho activation indirectly by upregulating Rho activators, we next wanted to identify which factors may be mediating this process. We found that TGF-β1 increases the mRNA level of endothelin-1, connective tissue growth factor, and SphK1. However, in our preliminary experiments (unpublished data), only S1P signaling inhibitors reduced TGF-β1–induced αSMA expression, so we focused the remainder of our studies on S1P signaling. We compared the kinetics by which TGF-β1 upregulates SphK1 with those of αSMA and CNN1 upregulation (Fig. 3A). HPASMCs were treated with TGF-β1 for 1, 3, 6, 12, or 24 hours. αSMA mRNA was upregulated 3.8-fold by TGF-β1 at 12 hours, whereas SphK1 mRNA was increased 4- or 7-fold at 3 or 6 hours, respectively. The peak of SphK1 mRNA level at 6 hours was approximately the same time when αSMA mRNA started to increase, supporting the idea that increased S1P activated αSMA expression. This is similar to results observed in fibroblasts in which TGF-β1 activated Rho signaling through S1P (Cencetti et al., 2010). The time course of the increase in CNN1 mRNA (Fig. 3A) was more similar to that for SphK1 than for αSMA, but CNN1 mRNA had a delayed peak at 12 hours (8-fold) as compared with SphK1. CNN1 mRNA increased by 2.6-fold at 3 hours, which was earlier than TGF-β1–induced phosphorylation of MLC2 (Fig. 2B). This suggests that TGF-β1 regulation of CNN1 expression may have a distinct mechanism from αSMA.
Kinetics of TGF-β1–induced contractile genes and TGF-β1 modulation of S1P receptors. HPASMCs were seeded into 6-well plates and allowed to reach confluence before being serum-deprived in 0.1% SMGS Medium 231 overnight. (A) Cells were then treated with 10 ng/ml TGF-β1 for 1, 3, 6, 12, and 24 hours in low serum medium. Afterward, mRNA was extracted, and the levels of αSMA, CNN1, and SphK1 were measured by RT-qPCR. (B) The mRNA levels of S1PR1, S1PR2, and S1PR3 were measured by RT-qPCR. (C) HPASMCs were pretreated with 10 μM SIS3 for 1 hour and then treated with 10 ng/ml TGF-β1 for 12 hours in low serum medium. RT-qPCR was used to assess the mRNA level of S1PR3 (N = 5. Data are represented as mean with S.D. For (A), one-way repeated-measures ANOVA followed by Dunnett’s post-test was performed. For (B), paired t test was conducted. * P < 0.05, ** P < 0.01).
TGF-β Modulates the Expression of S1P Receptor 3
In HPASMCs, 10 µM S1P induces approximately a 1.4-fold increase in αSMA protein levels (Supplemental Fig. 2). Surprisingly, with a lower concentration of S1P (1 µM), we did not observe induction of αSMA expression, which is inconsistent with the observation that S1P induces αSMA expression and differentiation in smooth muscle cells (Lockman et al., 2004). The blunted response of HPASMCs to S1P may be a result of different cell types having a different response to S1P, or it could be because primary cells may respond differently to S1P than cultured cell lines. In addition to the levels of S1P, the amount and composition of its receptors could also determine cellular responses to S1P. Consequently, we tested whether TGF-β1 also modulates the expression of S1P receptors in addition to SphK1. Based on the literature, HPASMCs express three subtypes of S1P receptors. Both S1PR2 and S1PR3 are coupled to G12/13 which, in turn, result in Rho activation (Wamhoff et al., 2008). We found that S1PR3 mRNA was elevated after 3 hours of TGF-β1 treatment and was further increased after 12 hours. S1PR1 and S1PR2 mRNA were unaffected by TGF-β1 (Fig. 3B). The TGF-β1–upregulated S1PR3 mRNA levels were reduced by the SMAD3 phosphorylation inhibitor SIS3 from 2.7- to 1.6- fold over control (95% confidence interval of the difference was −2.5 to 0.3, Fig. 3C). Although this effect was not statistically significant because of variability, treatment with the SMAD inhibitor SIS3 reduced the stimulation of S1PR3 expression by about 65%.
S1PR3 Antagonists Reduce TGF-β–Induced αSMA Expression, but an S1PR2 Antagonist Does Not
To determine which S1P receptors might be functionally relevant for TGF-β1–induced αSMA and CNN1 expression, HPASMCs were treated with TGF-β1 for 24 hours along with the S1PR2 antagonist JTE013, the S1PR3 antagonist TY52156, or the dual S1PR1/3 antagonist VPC23019. Both the S1PR3 and the S1PR1/3 antagonists reduced TGF-β1–induced stimulation of αSMA and CNN1 mRNA levels after 24 hours of cotreatment (Fig. 4A). There was no effect of the S1PR2 antagonist JTE013. αSMA protein levels were also reduced by the two antagonists (TY52156 and VPC23019) targeting the S1PR3 receptor (Fig. 4B). Inhibition of CNN1 protein levels by these two antagonists was only modest and did not achieve statistical significance. The overlapping effect of S1PR3 and S1PR1/3 antagonists leads us to conclude that TGF-β1 regulates αSMA expression through S1P signaling mainly via S1PR3, whereas CNN1 expression may also be regulated by S1PR3-independent signaling mechanisms.
TGF-β1–induced αSMA and CNN1 expression is mediated by S1PR3 but not by S1PR2. HPASMCs were seeded into 6-well plates and were allowed to reach confluence before being serum-deprived in 0.1% SMGS Medium 231 overnight. Cells were treated with 10 ng/ml TGF-β1 for 24 hours and 5 μM JTE013 (S1PR2 antagonist), 1 μM TY52156 (S1PR3 antagonist), 2 μM VPC23019, or vehicle control. The mRNA (A) and protein (B) levels of αSMA and CNN1 were analyzed by RT-qPCR and Western blot, respectively. Both blots were probed on the same membrane, but the bands were cropped to include only relevant lanes. αSMA protein levels in TGF-β1–treated groups with or without TY52156 and VPC23019 show a mean difference of −0.47 with 95% CI of −0.93 to 0.01 and −0.94 with 95% CI of −1.58 to −0.31 (N ≥ 3. Data are represented as mean with S.D. Paired t test. * P < 0.05). CI, confidence interval.
SphK1 Inhibitors Reduce TGF-β–Induced αSMA Expression
To test whether SphK1 regulates TGF-β–induced contractile genes expression, HPASMCs were treated with TGF-β1 for 24 hours with and without an SphK1 inhibitor, PF-543 at 10 µM (Fig. 5). PF-543 decreased both the αSMA and CNN1 mRNA by 80% and 50%, respectively. PF-543 trended toward decreasing αSMA protein from 2.6- to 1.9-fold of control and CNN1 protein from 1.9- to 1.4-fold of control. Although not statistically significant, this finding is similar to that for the S1PR3 receptor antagonists. In addition to SphK1, SphK2, another Sphingosine kinase, is also expressed in HPASMCs. However, the function of SphK2 is less well characterized than SphK1. TGF-β treatment did not affect the mRNA level of SphK2 in HPASMCs (Supplemental Fig. 3). Several studies demonstrated that SphK2 may be able to compensate for the deficiency of SphK1 and, in this case, may maintain the S1P levels when SphK1 is silenced. It required deletion of SphK1 and SphK2 to completely abolish the production of S1P and inhibit the effect of S1P (Mizugishi et al., 2005; Meng et al., 2011; Xiong et al., 2013). This could explain the less significant inhibitory effect of PF-543 on TGF-β–induced αSMA expression, since PF-543 might not fully block the function of SphK2 at 10 µM (Yang et al., 2019). Overall, these findings still suggest that SphK1 is important for regulating TGF-β–induced αSMA expression and that CNN1 expression is not completely dependent on S1P signaling.
TGF-β1–induced αSMA and CNN1 expression is reduced by SphK1 inhibitors. HPASMCs were seeded into 6-well plates and were allowed to reach confluence before being serum-deprived in 0.1% SMGS Medium 231 overnight. Cells were treated with 10 ng/ml TGF-β1 for 24 hours and 10 μM PF-543 or vehicle control. The mRNA (A) and protein (B) levels of αSMA and CNN1 were analyzed by RT-qPCR and Western blot, respectively. Both blots were probed on the same membrane, but the bands were cropped to include only relevant lanes. The αSMA protein levels in TGF-β1–treated groups with or without PF-543 show a mean of differences of −0.6779 with 95% CI of −1.646 to 0.2898 (N ≥ 3. Data are represented as mean with S.D. Paired t test * P < 0.05). CI, confidence interval.
Summary overview of the interaction of TGF-β and S1P signaling in regulation of αSMA expression in HPASMCs.
Discussion
TGF-β signaling is enhanced in patients with PAH (Yan et al., 2016), and transgenic mice overexpressing TGF-β1 spontaneously develop PAH (Calvier et al., 2019). The contributions and interactions of SMAD, Rho/MRTF, and other mechanisms downstream of TGF-β that contribute to PASMC activation remain controversial (Tang et al., 2011; Zabini et al., 2018; Calvier et al., 2019). Additionally, S1P plays an important role in PAH (Xing et al., 2015). Here, in HPASMCs, we investigated the interaction between TGF-β1 and S1P signaling to further define the signaling network downstream of TGF-β, with a focus on the SMAD and Rho/MRTF pathways.
SMAD signaling interacts with the Rho/MRTF/SRF pathway in multiple contexts. For example, in Monc-1 neural crest cells, RhoA directly regulates the phosphorylation of SMAD (Chen et al., 2006). In cardiac myoblasts, an MRTF-A/phosphorylated SMAD3 complex serves as a transcriptional regulatory element controlling the expression of αSMA (Parmacek, 2010). In HPASMCs, the Zabini group reported that loss of SMAD3 disinhibits MRTF and drives the αSMA expression in PAH (Zabini et al., 2018). However, the Hansmann group did not observe reduced expression of SMAD3 in the lungs from SUGEN/Hypoxia rats and pulmonary arteries from patients with PAH and claimed that TGF-β1 signaling drives αSMA expression in HPASMCs through canonical SMAD3 activation instead of by SMAD3 downregulation (Calvier et al., 2019). Our results suggest that both SMAD and Rho/MRTF/SRF signaling are involved in the regulation of αSMA and CNN1 in HPASMCs. We also show that MRTF-A but not MRTF-B is required for regulating αSMA expression in HPASMCs. Based on these data, we suggest a model wherein SMAD3 induces the activation of Rho/MRTF/SRF and regulates αSMA expression by potentiating the pathway of Rho activators, such as S1P.
The interaction between TGF-β and S1P signaling has been characterized in fibrosis and cancer. In those contexts, TGF-β increases the expression of SphK1 and in turn the level of S1P in fibroblasts, which contributes to TGF-β–mediated modulation of gene expression. SphK1/S1P has been reported to mediate TGF-β1–induced proliferation in rat PASMCs (Wang et al., 2019); however, we did not observe a significant pro-proliferative effect of TGF-β1 in human PASMCs (Supplemental Fig. 4). This highlights potential differences between the responses of rat and human PASMCs to the same stimuli. In addition to SphK1, TGF-β upregulates S1PR3 through the SMAD3 signaling axis in lung adenocarcinoma cell lines, and in those cells, S1PR3-mediated signaling drives the lung carcinoma cell growth (Zhao et al., 2016). We found a similar interaction between TGF-β and S1P signaling in regulation of αSMA in HPASMCs; TGF-β1 elevates αSMA expression, which is reduced by inhibition of either SphK1 or S1PR3. However, it is important to note that although we saw statistically significant differences at the mRNA level, in some cases the differences at the protein level did not achieve statistical significance. TGF-β1 induced an approximately 2- to 3-fold increase in αSMA protein levels, which gives a narrow window to test the inhibitory effect of antagonist, and maybe contributes to the variability in some experiments. Despite this, we believe that the similar response to several modulators targeting the S1P pathway suggests that SphK1 and S1PR3 are important for regulating TGF-β1–induced αSMA expression.
We demonstrated the essentiality of S1PR3 in the upregulation of αSMA by TGF-β1 in HPASMCs. Meanwhile, PASMCs from patients with PAH showed elevated S1PR2 levels (Chen et al., 2014). Silencing of S1PR2 or pharmacological inhibition of S1PR2 has been shown to ablate S1P-stimulated SMC proliferation (Chen et al., 2014). S1P regulates proliferation and differentiation of SMCs through different S1P receptors, which are coupled to different G proteins (Wamhoff et al., 2008). Thus, TGF-β–induced S1P levels could result in increased differentiation or proliferation of PASMCs based on the composition of S1P receptors that the cells express. Overall, Sphingosine kinase and S1PR3 are two critical components of the S1P signaling pathway, and this highlights the importance of S1P signaling in regulating αSMA expression and further supports their potential as PAH drug targets.
Our results showed that the S1PR3 antagonists markedly but not completely reduced the elevation of αSMA levels. One possibility is that TGF-β1 regulates αSMA expression through the SMAD pathway in parallel. Alternatively, TGF-β1 may also upregulate other Rho activators, such as endothelin-1 and connective tissue growth factor, which in turn contribute to the αSMA elevation. Another observation we had is that CNN1 is regulated in a different manner from αSMA. CNN1 mRNA increased at a similar time course as SphK1 and earlier than TGF-β1–induced phosphorylation of MLC2. However, the upregulation of CNN1 mRNA at 24 hours was substantially suppressed by the ROCK inhibitor Y27632 (Fig. 1A). The S1PR3 and sphingosine kinases inhibitors inhibited the TGF-β1–induced CNN1 expression to a lesser extent. These suggest that the regulation of CNN1 expression relies on a Rho-dependent mechanism at later times but SMAD or other mechanisms early on during stimulation.
Phosphorylation of MLC2 causes contraction of SMCs. Additionally, increased expression of the contractile protein αSMA also contributes to the elevated vasoconstriction and SMC hypertrophy (Bai et al., 2020; Zabini et al., 2018). In this study we demonstrated the role of SphK1 and S1PR3 in TGF-β1–induced αSMA expression. We found that both SMAD3 and Rho/MRTF-A/SRF are important mediators of TGF-β1–induced αSMA expression in HPASMCs. We proposed a model similar to what was demonstrated in fibroblasts, wherein TGF-β1 upregulated αSMA expressions in HPASMCs by potentiating S1P signaling (Figure 6). Our results show that SIS3 tends to inhibit TGF-β1–induced S1PR3 mRNA, although this observation did not reach statistical significance. Although we lack evidence that strongly implicates the SMAD pathway in this model, our data suggest that TGF-β1 potentiates S1P signaling through SMAD pathway, which is activated at an earlier time point. In total, these findings suggest that both S1P and MRTF-A/SRF signaling are potential therapeutic targets to reduce vascular contraction and SMC hypertrophy in PAH.
Authorship Contributions
Participated in research design: Ji, Lisabeth, Neubig.
Conducted experiments: Ji.
Performed data analysis: Ji, Lisabeth, Neubig.
Wrote or contributed to the writing of the manuscript: Ji, Lisabeth, Neubig.
Footnotes
- Received March 24, 2020.
- Accepted April 30, 2021.
This research was supported by American Heart Association predoctoral fellowship 19PRE34450084 and Michigan State University (MSU) Scleroderma/Fibrosis Research Fund.
Dr. Neubig is President and cofounder of FibrosIX Inc., which holds an option on intellectual property for CCG-222740 and related compounds.
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This article has supplemental material available at molpharm.aspetjournals.org.
Abbreviations
- CNN1
- calponin 1
- HPASMC
- human pulmonary arterial SMC
- MLC
- myosin light chain
- MRTF
- myocardin-related transcription factor
- PAH
- pulmonary arterial hypertension
- PASMC
- pulmonary arterial SMC
- pMLC2
- phosphorylated MLC
- ROCK
- Rho-associated protein kinase
- RT-qPCR
- reverse transcription-polymerase chain reaction
- siRNA
- small interfering RNA
- αSMA
- α smooth muscle actin
- SMAD
- mothers against decapentaplegic homolog
- SMC
- smooth muscle cell
- SMGS
- smooth muscle growth supplement
- S1P
- sphingosine-1-phosphate
- SphK1
- sphingosine kinase 1
- S1PR
- S1P receptor
- SRF
- serum response factor
- TGF-β
- transforming growth factor β
- Copyright © 2021 by The American Society for Pharmacology and Experimental Therapeutics
