Contribution of transcription factor EB to adipoRon-induced inhibition of arterial smooth muscle cell proliferation and migration

Abstract

Abnormal vascular smooth muscle cell (SMC) dedifferentiation with increased proliferation and migration during pathological vascular remodeling is associated with vascular disorders, such as atherosclerosis and in-stent restenosis. AdipoRon, a selective agonist of adiponectin receptor, has been shown to protect against vascular remodeling by preventing SMC dedifferentiation. However, the molecular mechanisms that mediate adipoRon-induced SMC differentiation are not well understood. The present study aimed to elucidate the role of transcription factor EB (TFEB), a master regulator of autophagy, in mediating adipoRon’s effect on SMCs. In cultured arterial SMCs, adipoRon dose-dependently increased TFEB activation, which is accompanied by upregulated transcription of genes involved in autophagy pathway and enhanced autophagic flux. In parallel, adipoRon suppressed serum-induced cell proliferation and caused cell cycle arrest. Moreover, adipoRon inhibited SMC migration as characterized by wound-healing retardation, F-actin reorganization, and matrix metalloproteinase-9 downregulation. These inhibitory effects of adipoRon on proliferation and migration were attenuated by TFEB gene silencing. Mechanistically, activation of TFEB by adipoRon is dependent on intracellular calcium, but it is not associated with changes in AMPK, ERK1/2, Akt, or molecular target of rapamycin complex 1 activation. Using ex vivo aortic explants, we demonstrated that adipoRon inhibited sprouts that had outgrown from aortic rings, whereas lentiviral TFEB shRNA transduction significantly reversed this effect of adipoRon on aortic rings. Taken together, our results indicate that adipoRon activates TFEB signaling that helps maintain the quiescent and differentiated status of arterial SMCs, preventing abnormal SMC dedifferentiation. This study provides novel mechanistic insights into understanding the therapeutic effects of adipoRon on TFEB signaling and pathological vascular remodeling.

INTRODUCTION

Pathological vascular remodeling is involved in vascular disorders that lead to myocardial infarction, stroke, peripheral vascular diseases, or graft failure in organ transplantation (23, 40, 52). Vascular smooth muscle cell (SMC) is a major cell type of the arterial wall that is normally maintained in homeostatic status and that possesses contractile phenotype with very low synthetic activity. During vascular remodeling, SMCs dedifferentiate into a synthetic state with enhanced proliferative and migratory potential, leading to neointima formation and luminal stenosis (2, 25). Obesity is a major cardiovascular risk factor for the development of vasculopathy (60). Many factors associated with obesity, such as adipokines, can modulate SMC phenotype plasticity with both injurious or beneficial effects (60). There are great clinical implications in understanding the mechanisms underlying the dynamic regulation of SMC phenotypic plasticity by adipokines that helps develop pharmacological interventions.

Adiponectin is the most abundant plasma adipokine that exerts beneficial effects on cardiovascular diseases (13). Hypoadiponectinemia is linked with insulin resistance, hypertension, and coronary artery disease (21, 64). Adiponectin-deficient mice developed insulin resistance and neointima formation after arterial injury, which can be prevented by adenoviral delivery of adiponectin (27, 36). However, adiponectin is relatively large in molecular size and has a short half-life, which limits its clinical use for cardiovascular diseases. AdipoRon has been recently developed as a selective agonist of adiponectin receptors and mimics many beneficial effects of adiponectin on the cardiovascular system. For example, adipoRon was shown to increase postischemic myocardial survival, improve endothelial functions, and cause smooth muscle relaxation (18). Recently, adipoRon was reported to induce SMC differentiation, inhibit SMC proliferation and migration and attenuate neointima formation in the femoral arteries or in aortas with hypertension-induced injury in mice (12, 16). The protective effects of adipoRon on SMCs were attributed to the inhibition of PDGF receptor-dependent signaling or blockade of p38 signaling pathway (12, 16). To date, the downstream mediators for adipoRon effects that interact with SMC dedifferentiation signaling components remain largely unknown.

Transcription factor EB (TFEB) is a member of the microphthalmia family transcription factors and functions as a master controller of autophagy pathway (43, 49, 50, 56, 61, 65, 75). Activation of TFEB leads to its nuclear translocation and binding to coordinated lysosomal expression and regulation (CLEAR) elements of target genes involved in the autophagy pathway, including microtubule-associated protein light chain 3 (LC3), ubiquitin-binding protein p62/sequestosome 1 (p62/SQSTM1), lysosomal-associated membrane protein 1 (LAMP-1), and lysosomal-associated membrane protein 2A (LAMP-2A) (55–59). Consequently, TFEB-mediated transcriptional upregulation of autophagy genes results in autophagosome induction and lysosome biogenesis (46, 56, 61, 62). Recent studies demonstrated that TFEB plays a beneficial role in protecting against cardiovascular diseases. For example, pharmacological activation of TFEB by trehalose induces autophagosome biogenesis and lysosome regeneration in macrophages, and treatment of mice with trehalose also ameliorates atherosclerosis development (9, 11, 54). TFEB gene overexpression can inhibit endothelial inflammation and ameliorate atherosclerosis in mice (34). However, the roles of TFEB in vascular smooth muscle function and in mediating the effects of adipoRon on vascular complications are not investigated.

The present study aimed to test the hypothesis that TFEB mediates adipoRon-induced inhibitory effects on SMC dedifferentiation. Our results demonstrated that adipoRon inhibits SMC dedifferentiation, as shown by the reduced proliferative and migratory capacity of SMCs, and this effect of adipoRon is attributed, at least in part, to TFEB activation. Our findings provide novel mechanistic insights into understanding the therapeutic effects of adipoRon on vascular complications.

MATERIALS AND METHODS

Antibodies and reagents.

The primary antibodies used are as follows: TFEB (A303-673A; Bethyl Laboratories), LC3 [Cell Signaling Technology (CST) 12741S], β-actin (CST 3700S), p62/SQSTM1 (ab109012; Abcam), LAMP-1 [553792; Becton Dickinson (BD)], Ki67 (ab16667; Abcam), cyclin D1 (554181; BD), cyclin-dependent kinase 4 (CDK4; 559677; BD), GAPDH (sc47724; Santa Cruz Biotechnology), phospho-ERK1/2 (4370; Cell Signaling), ERK1/2 (4695; Cell Signaling), phospho- Akt (4060; Cell Signaling), Akt (9272; Cell Signaling), phospho-AMPKα (Thr-172) (2535; Cell Signaling), AMPKα (2532; Cell Signaling), phospho- molecular target of rapamycin (mTOR) (Ser2448) (5536; Cell Signaling), and mTOR (2983; Cell Signaling). Secondary antibody for Western blot: IRDye 800CW donkey anti-mouse IgG (H⁺L) (926-32212; LICOR), IRDye 800CW donkey anti-rabbit IgG (H⁺L) (926-32213; LICOR), donkey anti-mouse IgG (H⁺L), HRP (A16011; Thermo Fisher), stabilized peroxidase-conjugated goat anti-rabbit (H⁺L) (32460; Invitrogen), goat anti-rat IgG-HRP (629520; Thermo Fisher). Secondary antibody for immunofluorescence: donkey anti-mouse IgG (H⁺L), Alexa Fluor 488 conjugate (A-21202; Thermo Fisher), donkey Alexa Fluor 488 conjugate anti-rabbit IgG (H⁺L) (Thermo Fisher; A21206), donkey Alexa Fluor 555 conjugate anti-mouse IgG (H⁺L) (Thermo Fisher; A-31570), donkey Alexa Fluor 555 conjugate anti-rabbit IgG (H⁺L) (A-31572; Thermo Fisher).

The following reagents were used: adipoRon (ab144867; Abcam), Aurum total RNA mini kits (732-6820; Bio-Rad), iScript reverse transcription supermix for quantitative RT-PCR (1708841; Bio-Rad), iTaq Universal SYBR Green supermix (1725121; Bio-Rad), and rhodamine phalloidin (R415; Fisher).

Mice.

All experimental protocols were reviewed and approved by the Animal Care Committee of University of Houston. C57BL/6J male mice were used in all experiments. All animals were provided standard rodent chow, water ad libitum, and 12:12-h dark-light cycles in a temperature-controlled room.

Primary culture of arterial SMCs from mice.

SMCs were isolated from mice as we previously described (73, 76). Mice at 6 wk of age were deeply anesthetized with an intraperitoneal injection of pentobarbital sodium (25 mg/kg). Then, the mouse hearts were excised with an intact aortic arch and immediately immersed in a petri dish filled with ice-cold Krebs-Henseleit (KH) solution. While the whole heart was maintained in the ice-cold KH solution, a needle (25 gauge), filled with Hanks’ buffered saline solution (HBSS), was inserted through the aortic lumen opening deep into the heart close to the aortic valve. Then, the needle was tied when the needle tip reached the base of the heart. An infusion pump was started with a 20-mL syringe containing warm HBSS through an intravenous extension set at a rate of 0.1 mL/min for 15 min. The HBSS was replaced with a warm enzyme solution containing 1 mg/mL collagenase type 1, 0.5 mg/mL soybean trypsin inhibitor, 3% BSA, and 2% antibiotics. The enzyme solution was flushed through the heart at a rate of 0.1 mL/min. The outflow perfusion fluid was collected at 30-, 60-, and 90-min intervals. After collecting all outflow fluid at 90 min, the heart was cut to open the apex to flush out the cells inside the ventricle. The flushed cells were centrifuged at 1,000 rpm for 10 min, and the pellets were resuspended in advanced Dulbecco’s modified Eagle’s medium (DMEM) with 10% fetal bovine serum (FBS), 10% mouse serum, and 2% antibiotics. The isolated cells were plated on 2% gelatin-coated six-well plates and incubated in 5% CO₂ at 37°C. These isolated cells were confirmed as SMCs originating mainly from coronary arteries by positive staining with α-SMA antibodies and the SMC morphology. The culture medium was replaced 3 days after cell isolation and then twice each week until the cells grew to confluence. All studies were performed with cells at passages 5–8. In this study, SMCs were cultured under a dedifferentiation condition in full-serum medium (DMEM with 10% FBS), if not particularly mentioned.

Immunoblotting.

Cells lysates were prepared in Laemmli sample buffer (161-0737; Bio-Rad) containing β-mercaptoethanol (M3148; Sigma Aldrich), heated for 10 min at 95°C, and then placed in ice-cold water ultrasonic bath. Twenty micrograms of proteins were separated by 8–12% SDS-PAGE. The proteins on the gel were electrophoretically (35 V) transferred onto a PVDF membrane (Bio-Rad) at 4°C overnight. The membrane was blocked with 5% nonfat milk in TBS. After washing, the membrane was probed with primary antibodies as indicated, according to the manufacturer’s instructions. The membrane was washed again and incubated with corresponding secondary antibodies. The protein bands were visualized and analyzed by LI-COR Odyssey Fc System, as we have described previously (76).

Quantitative real-time PCR.

Total RNA was isolated using the Aurum total RNA mini kits (732-6820; Bio-Rad), according to the manufacturer’s instructions. The cDNA was generated from the total RNA using iScript reverse transcription supermix for quantitative RT-PCR (1708841; Bio-Rad). The real-time PCR was performed using the iTaq Universal SYBR Green supermix (1725121; Bio-Rad) on the Bio-Rad CFX Connect real-time system using the following primers: TFEB forward: 5′-CAGCAGGTGGTGAAGCAAGAGT-3′ and reverse: 5′-TCCAGGTGATGGAACGGAGACT-3′; LC3 forward: 5′-CGTCCTGGACAAGACCAAGT-3′ and LC3 reverse: 5′-ATTGCTGTCCCGAATGTCTC-3′; p62/SQSTM1 forward: 5′-AGGGAACACAGCAAGCT-3′ and p62/SQSTM1 reverse: 5′-GCCAAAGTGTCCATGTTTCA-3′; LAMP-2A forward: 5′- CCAAATTGGGATCCTAACCTAA-3′ and LAMP-2A reverse: 5′-TGGTCAAGCAGTGTTTATTAATTCC-3′; LAMP-1 forward: 5′- ACATCAGCCCAAATGACACA-3′ and LAMP-1 reverse: 5′-GGCTAGAGCTGGCATTCATC-3′; β-actin forward: 5′-TCGCTGCGCTGGTCGTC-3′ and β-actin reverse: 5′- GGCCTCGTCACCCACATAGGA-3′. The cycle threshold values were converted to relative gene expression levels using the 2^−ΔΔCt method. The data were normalized to that of internal control β-actin, as we described previously (76).

Immunofluorescence staining.

Approximately 1 × 10⁴ cells were seeded on gelatin-coated coverslips in 24-well plates. Cells were washed two times with PBS, and fixed in 4% paraformaldehyde at room temperature for 15 min. After fixation, cells were washed twice with PBS Tween (PBST; 0.05% Tween20 in PBS) and then permeabilized with 0.3% Triton X-100 in PBST for 15 min. Nonspecific sites were blocked with 5% BSA in PBS at room temperature for 1 h, and then cells were incubated with indicated primary antibodies at 4°C overnight. Cells were then incubated with corresponding secondary antibodies conjugated with Alexa Fluor 488 or Alexa Fluor 555 for 1 h at room temperature. The cell nucleus was stained with DAPI for 15 min at room temperature and then mounted with an antifluorescence quenching agent. For phalloidin staining of F-actin in cultured cells, fixed and permeabilized cells were incubated with Alexa Fluor 568-conjugated phalloidin (1:50) for 30 min, washed with PBS, and mounted with an antifluorescence quenching agent. The cells were visualized using Olympus IX73 imaging system. Pearson’s correlation for colocalization efficient and mean fluorescence density was analyzed using Image-Pro Plus 6.0 software, as described previously (70).

Cell viability assay.

The cell viability was assessed by lactate dehydrogenase (LDH) assay. LDH activity in the cell culture medium was determined by Pierce LDH assay kit (Thermo Fisher Scientific), according to the manufacturer’s protocol. The absorbance was spectrophotometrically quantified at 490 nm using CLARIOstar microplate reader (BMG Labtech). The maximum LDH activity (LDH_max) was obtained from samples treated with lysis buffer. The basal level of LDH activity (LDH₀) due to spontaneous LDH release was obtained from unstimulated control samples. The survival was calculated as the percentage of (LDH_max − LDH_sample) over (LDH_max − LDH₀).

Wound scratch assay of SMC migration.

Cell migration was assessed by a wound scratch assay. Briefly, 90% confluent SMCs were starved in low-serum media (0.1% FBS) overnight. Scratch wounds were created using a 2-mm-wide pipette tip. Cells were cultured in full-serum medium (10% FBS) with indicated treatment. After 24 h, the scratched area of cells was imaged by using Olympus IX73 imaging system. The average wounded area was quantified using Image-Pro Plus 6.0 software.

Electric cell-substrate impedance-sensing assay of SMC migration.

An electric cell-substrate impedance-sensing (ECIS) system (Applied Biophysics) was used to conduct migration assay, as we have described previously (15). SMCs (5 × 10⁴ per well) were cultured in 8W1E+ arrays (Applied Biophysics), and the resistance was subsequently monitored and measured using the ECIS software provided. The cells were incubated until the resistance reached a plateau, suggesting a confluent monolayer had formed over the array electrodes. The cell monolayer was subsequently wounded electrically by applying current pulses of 5,000 μA s at 64,000 Hz for 30 s. This wounding procedure created a simultaneous physical break of equal dimensions in the cell monolayer of each well. After wounding, the resistance increased, as cells migrated back onto the electrode. The slope of changes in resistance after wounding before reaching the plateau was calculated and used to reflect the cell migration rate. The maximal change in resistance was also calculated.

RNA interference by siRNA transfection.

TFEB siRNA (sc-38510; Santa Cruz Biotechnology) or scramble control siRNA (sc-37007; Santa Cruz Biotechnology) were commercially available. SMCs with 70% confluency were transfected with siRNA using SiLentFect Lipid Reagent (170-3361; Bio-Rad), according to the manufacturer’s instructions. The transfection efficiency of TFEB siRNA was examined by quantitative RT-PCR of mRNA level of TFEB, LC3, p62/SQSTM1, LAMP-1 and LAMP-2A, and Western blot analysis of TFEB and LC3.

Ex vivo aortic ring sprouting assay.

The mouse aortic ring sprouting assay was performed to examine the migration and proliferation of SMCs in an ex vivo condition, as described previously (66, 76). In brief, fresh thoracic aorta was harvested from 12-wk old mice and placed in sterile DMEM buffer. The endothelial layer was removed by the injection of air bubbles. The periadventitial fat and connective tissues were carefully removed. Then the aorta was sliced into ring segments ~0.5 mm in thickness. The aortic rings were rinsed with DMEM medium and then embedded in 24-well plates precoated with 200 µL of synthetic basement membrane (Corning Matrigel) per well on ice. Approximately 10–15 rings per group were used for each experiment. The plate with aortic rings was first placed at room temperature for 15 min, and then it was incubated in a humidified incubator (37°C, 5% CO₂) for 1 h followed by addition of 500 μl prewarmed DMEM containing 10% FBS into each well. The explants were maintained in a humidified incubator (37°C, 5% CO₂) with medium replaced every other day. For the treatment group, the culture medium was replaced by fresh DMEM containing 10% FBS and adipoRon at the indicated concentration (0–100 μM). For shRNA treatment groups, aortic rings were preincubated with control or TFEB shRNA lentiviral particles (sc-38510-V; Santa Cruz) for 24 h in the presence of polybrene (final concentration was 5 μg/mL), and then treated with vehicle or 50 μΜ adipoRon. After 7 days, the outgrowth of cells was observed and imaged using an inverted microscope (Olympus IX73). Digital images of aortic ring sections were analyzed using Image-Pro Plus software. Each outgrowth emerging directly from the ring was identified (number of sprouts) and traced, and branch points were marked. The ImageJ segmented line tool was then used to measure the migrated area of all branches for each outgrowth that reflects migration distance.

Statistics analysis.

Data are presented as means ± SD. All experiments were analyzed by Student’s t test or one/two-way ANOVA with treatments as category factors, followed by Bonferroni’s multiple comparison test, if applicable. Student’s t test was used to detect significant difference between the two groups. The statistical analysis was performed by GraphPad Prism 6.0 software (GraphPad Software). P < 0.05 was considered statistically significant.

RESULTS

AdipoRon activates TFEB and autophagy signaling in SMCs.

Previous studies demonstrated that adiponectin receptor-mediated signaling protects against the proliferation and neointima formation (12). Here, we explored the possible role of TFEB in the protective effects on SMCs exerted by adipoRon, a selective adiponectin receptor agonist. Our immunofluorescent studies showed that adipoRon significantly increased the nuclear translocation of TFEB from the cytosol, a key event in activating transcription factor TFEB (Fig. 1A). AdipoRon also increased the expression of TFEB and TFEB-controlled autophagy genes, including LC3, p62, LAMP-1, and LAMP-2A (Fig. 1, B and C). Moreover, we demonstrated that adipoRon further increased LC3 expression when autophagic flux was blocked by chloroquine or bafilomycin (Fig. 1D). This suggests that adipoRon-induced increases in LC3 and p62 were likely due to the induction of gene expression rather than inhibition of autophagic flux. We also directly monitored the autophagolysosome formation by detecting increased colocalization of LC3 with lysosome protein LAMP-1 (Fig. 1E).

Fig. 1.

AdipoRon activates transcription factor EB (TFEB) and induces autophagy in smooth muscle cells (SMCs). SMCs cultured in full-serum medium (10% FBS) were treated with adipoRon (AdR; 0–100 μM) for 6 or 24 h. A: immunofluorescence images and quantification data show the translocation of TFEB in the nucleus (green) upon adipoRon treatment for 6 or 24 h (n = 4). Nuclei were stained with DAPI. B: immunoblotting analysis and quantification data show the protein expression of LC3 and p62/SQSTM1 in SMCs upon adipoRon treatment for 24 h (n = 4). C: real-time RT-PCR analysis of mRNA levels of TFEB, LC3, p62/SQSTM1, LAMP-1, and LAMP-2A (n = 4–6). To arrest autophagic flux, cells were treated with adipoRon for 20 h and then incubated with or without lysosome function inhibitors chloroquine (CQ, 100 μM) or bafilomycin (Baf; 50 nM) for another 4 h. D: immunoblotting analysis shows the protein expression of LC3 and p62/SQSTM1 (n = 4). E: immunofluorescence images and quantification data show the colocalization of LC3 and LAMP-1 (n = 4). Scale bar = 20 μm. *P < 0.05 vs. vehicle control or as indicated.

AdipoRon inhibits the proliferation of SMCs.

As shown in Fig. 2A, adipoRon significantly arrested cell growth of SMCs cultured in full-serum medium in a dose-dependent manner. We then determined the protein expression of cell cycle-related proteins cyclin D1 and CDK4. As shown in Fig. 2B, adipoRon markedly decreased cyclin D1 and CDK4 expression. The antiproliferative effect of adipoRon was also confirmed by the decreased expression of proliferation marker Ki67 (Fig. 2C). This inhibitory effect of adipoRon on cell growth was not associated with induction of cell death, as confirmed by LDH cytotoxicity analysis (Fig. 2D) and flow cytometric analysis of PI/Annexin V staining (Fig. 2E).

Fig. 2.

AdipoRon inhibits the proliferation of smooth muscle cells (SMCs) induced by serum. SMCs cultured in full-serum medium (10% FBS) were treated with adipoRon (AdR; 0–100 μM) for 24 h. A: cell proliferation was analyzed by counting the cell numbers (n = 4). B: immunoblotting analysis shows the effects of adipoRon on the cell cycle protein cyclin D1 and CDK4 (n = 4). C: immunofluorescence images and summarized data show the effects of adipoRon on proliferative cell marker Ki67 (green). Nuclei were stained with DAPI. Quantified data show the percentage of Ki67-positive cells (n = 4). Scale bar = 20 μm. AOI, area of interest. D: lactate dehydrogenase (LDH) assay for the cytotoxicity of adipoRon (n = 4). E: flow cytometric analysis of apoptotic cells by FITC-annexin V and propodium iodide (PI) staining (n = 4). *P < 0.05 vs. vehicle control.

AdipoRon inhibits migration of SMCs.

As shown in Fig. 3A, adipoRon significantly decreased the migration of SMCs, as assessed by the scratch assay. To exclude the effects of cell proliferation on the wound-healing process in scratch assay, we used ECIS method to monitor the migration process in the wounded area under full-serum (10% FBS) or low-serum (0.1% FBS) conditions. The recovery of resistance was considered to correlate with cell migration, but not proliferation, particularly for cells under the low-serum condition. As shown in Fig. 3B, in both full-serum and low-serum cultured SMCs, adipoRon significantly retarded the recovery of resistance, as shown by the reduced slope in the cell resistance trace curve and the attenuated maximal increase in cell resistance. These data further suggest that adipoRon had inhibitory effects on the migratory ability of SMCs. The remodeling of F-actin, a filamentous actin in the cytoskeleton, is a marker event associated with cell migration (24). SMCs within full serum media exhibited migratory phenotype with disassembled distribution and aggregation around the perinuclear region of actin filaments without clear filamentous organization (Fig. 3C). Conversely, SMCs treated with adipoRon showed a spindle-like shape and organization of the actin filaments (Fig. 3C). Matrix metalloproteinases (MMP) are peptidase enzymes involved in extracellular matrix degradation contributing to cell migration (37). AdipoRon decreased the mRNA level of MMP9 (Fig. 3D).

Fig. 3.

AdipoRon (AdR) inhibits the migration of smooth muscle cells (SMCs) induced by serum. SMCs cultured in full-serum medium (10% FBS) were treated with adipoRon (0–50 μM) for 24 h. A: cell migration was analyzed by scratch assay (n = 4). Scale bar = 100 μm. B: electric cell-substrate impedance sensing (ECIS) analysis of wound healing in SMCs. SMCs were cultured on chamber slides with electrodes until confluency that transmonolayer electrical resistance reaches a plateau. Cells were then incubated with or without adipoRon (50 μM) for 30 min before the introduction of electrical pulse for cell wounding. Traces show cell migration as monitored by determining the resistance of the wounded area using ECIS. Quantified data show the slope of the recovery resistance curve after wounding, which reflects the rate of unwounded cell migration toward the wounded area (n = 4). C: representative immunofluorescence images show the arrangement of F-actin filaments by phalloidin staining (red) from four independent experiments. Scale bar = 20 μm. D: real-time RT-PCR analysis of MMP-9 mRNA levels (n = 4). *P < 0.05 vs. vehicle control.

TFEB gene silencing attenuates the effects of adipoRon on SMC proliferation and migration.

TFEB gene silencing in SMCs effectively decreased expression of TFEB and LC3 (Fig. 4A) and downregulated the mRNA levels of TFEB itself and TFEB-target genes (LC3, p62, LAMP-1, and LAMP-2A) (Fig. 4B). More importantly, adipoRon-induced inhibition of proliferation was significantly attenuated when TFEB gene is silenced (Fig. 4C). Similarly, adipoRon-induced downregulation of SMC migration was reversed (Fig. 4D). Together, these data suggest that agonism of adipoRon promotes SMC differentiation, leading to reduced proliferative and migratory capacity through a TFEB-dependent manner.

Fig. 4.

Transcription factor EB (TFEB) gene silencing ameliorates adipoRon-induced differentiation in smooth muscle cells (SMCs). SMCs were cultured in full-serum medium (10% FBS) and transfected with control scramble siRNA (si-ctrl) or TFEB siRNA (si-TFEB) for 24 h. Then, cells were treated with vehicle or adipoRon (50 μM) for another 24 h. A: immunoblot analysis shows the efficiency of TFEB siRNA transfection on downregulating TFEB and LC3 (n = 3). B: real-time RT-PCR analysis of mRNA levels of TFEB, LC3, p62/SQSTM1, LAMP-1, and LAMP-2A (n = 4). C: cell number counting of siRNA-transfected SMCs treated with vehicle or 25 μM adipoRon (n = 4). D: scratch assay and summarized data of transfected SMCs treated with vehicle or 50 μM adipoRon (AdR; n = 4). Scale bar = 100 μm *P < 0.05 vs. si-ctrl.

AdipoRon increases ERK1/2, Akt, and AMPK activation but does not affect mTOR in SMCs.

In non-SMCs, phosphorylation of TFEB by protein kinases, such as mTOR, promotes its degradation in the cytoplasm and inhibits TFEB activity (35, 51, 59), whereas calcium-dependent phosphatase can enhance TFEB activity by dephosphorylation (38). Protein kinases, such as ERK1/2, Akt, and AMPK are upstream regulators of mTOR activity in SMCs. Here, we found that adipoRon, particularly at a concentration of 50 μM, increased phosphorylation of ERK1/2 (Fig. 5A), Akt (Fig. 5B), and AMPK (Fig. 5C) in arterial SMCs. However, adipoRon did not affect phosphorylation of mTOR (Fig. 5D).

Fig. 5.

Effects of adipoRon (AdR) on activation of ERK1/2, Akt, and AMPK-molecular target of rapamycin (mTOR) complex 1 axis in SMCs. SMCs were cultured in full-serum medium (10% FBS) and treated with adipoRon (0–50 μM) for 24 h. Representative immunoblots and quantified data show the effects of adipoRon on the phosphorylation of ERK1/2 (A), Akt (B), AMPK (C), or mTOR (D) (n = 4). *P < 0.05 vs. vehicle control.

AdipoRon-induced TFEB activation depends on intracellular Ca²⁺.

Calcium-dependent phosphatases were shown to dephosphorylate TFEB and, thereby, promote TFEB stability and activity in an mTOR-independent manner (38). As shown in Fig. 6A, the treatment of SMCs with cell-permeable Ca²⁺ blocker BAPTA attenuated adipoRon-induced TFEB nuclear translocation. In contrast, extracellular Ca²⁺ chelator EGTA had no effects. Thapsigargin, an inhibitor of the sarcoendoplasmic reticulum Ca²⁺ ATPase, can cause Ca²⁺ leakage from sarco/endoplasmic reticulum and, thereby, increase intracellular Ca²⁺ concentration in endothelial cells and SMCs (10, 74). Our data demonstrated that thapsigargin could activate TFEB nuclear translocation. These data further implicate a role of intracellular Ca²⁺ in TFEB activation. The inhibition of TFEB activity was also confirmed by the finding that BAPTA reduced the adipoRon-induced protein expression of LC3 and p62/SQSTM1 (Fig. 6B) and mRNA transcription of TFEB-controlled autophagy pathway genes (Fig. 6C). It should be noted that BAPTA alone significantly decreased the mRNA levels of TFEB, and its targeted genes without affecting the basal TFEB activity. These data may suggest that BAPTA affects the autophagy clearance pathway, or TFEB is dispensable for basal autophagy.

Fig. 6.

Activation of transcription factor EB (TFEB) by adipoRon (AdR) requires intracellular Ca²⁺. Smooth muscle cells (SMCs) were cultured in full-serum medium (10% FBS) and treated with adipoRon (50 μM) for 24 h in the absence or presence of BAPTA-AM (5 μM) or EGTA (1 mM), or with thapsigargin (TG; 10 μM). A: immunofluorescence images and quantification data show the translocation of TFEB (green) in the nucleus in SMCs (n = 4). Nuclei were stained with DAPI. Scale bar = 20 μm. B: representative immunoblotting analysis and summarized data show the protein expression level of LC3 and p62/SQSTM1 (n = 4). C: real-time RT-PCR analysis of mRNA levels of TFEB, LC3, p62/SQSTM1, LAMP-1, and LAMP-2A (n = 4). *P < 0.05.

AdipoRon inhibits the formation of aortic ring sprouts in a TFEB-dependent manner.

The role of the adipoRon-TFEB pathway in regulating SMC proliferation and migration was further examined in aortic explants from mice. The endothelium-denuded aortic rings were cultured ex vivo in a Matrigel-precoated 24-well plate. The number of sprouts that had outgrown from aortic rings was quantified by analyzing cells stained with DAPI. The cell migration distance was measured for DAPI⁺ cells that emigrate from aortic explants. As shown in Fig. 7A, adipoRon-treated rings showed a significantly decreased number of sprouts and cell migration distance compared with aortic rings cultured under control condition. As a negative control, aortic rings under serum starvation condition showed almost no cell migration (Fig. 7A). To suppress TFEB function ex vivo, aortic rings were transduced with lentiviral particles containing TFEB shRNA plasmids. This TFEB gene silencing significantly attenuated the inhibitory effects of adipoRon on aortic ring sprouts (Fig. 7B).

Fig. 7.

AdipoRon (AdR) prevents the formation of aortic ring sprouts ex vivo. A: aortic rings were isolated and treated with adipoRon (0–100 μM) for 7 days. Aortic rings were also cultured under starvation (Stav) condition in low serum medium (0.1% FBS) as a negative control. Representative bright-field and immunofluorescent images show the aortic rings stained with DAPI (blue). The summarized data show the number of sprouts that emerge from aortic rings and the migrated area of all branches for each outgrowth that reflects migration distance (n = 10–15 rings). B: aortic rings were transfected with TFEB shRNA (shTFEB) or scramble control shRNA (Scr) for 24 h and then treated with or without adipoRon (50 μM) for 7 days. Immunofluorescent analysis of the number of sprouts that emerge from aortic rings and the migrated area of all branches for each outgrowth that reflects migration distance (n = 10–15 rings). Scale bar = 200 μm. *P < 0.05; #P < 0.05 vs. vehicle.

DISCUSSION

The aim of the present study was to determine the role of TFEB in mediating the protective effects of adipoRon on SMCs. Our study demonstrated that adipoRon activates the TFEB-autophagy pathway that contributes to inhibited proliferation and migration in SMCs. Such adipoRon-induced TFEB activation was dependent on intracellular calcium but independent of Akt, ERK1/2, AMPK, and mTOR.

AdipoRon is a widely used small-molecule agonist of adiponectin receptors (47). However, the role of the TFEB-autophagy pathway in the adiponectin receptor signaling pathway has not been elucidated. Autophagy is an evolutionarily conserved, nonstop, reparative, and life-sustaining catabolic process to maintain normal cellular homeostasis. In this process, double-membraned autophagosomes engulf unhealthy organelles and long-lived proteins and degrade them into small molecules via autophagic flux. In autophagic flux, autophagosomes fuse with acidic lysosomes to form autophagolysosomes, in which autophagic substrates are degraded by lysosome proteinases and hydrolases (22, 30, 33, 71). A recent study demonstrated that adipoRon enhanced autophagosome formation, lysosome protein LAMP2 expression, and autophagosome clearance in adult cardiomyocytes from mice (67). In the present study, we demonstrated that adipoRon treatment increased the TFEB activation, as indicated by nuclear translocation of TFEB and subsequent protein expression of autophagosome marker LC3 and autophagic substrate p62/SQSTM1 in SMCs. Such TFEB activation was also accompanied by transcriptional upregulation of TFEB-targeted genes, including TFEB itself and autophagy genes LC3, p62/SQSTM1, LAMP-1, and LAMP-2A. Moreover, we demonstrated that arresting autophagic flux enhanced adipoRon-induced LC3 expression, indicating that adipoRon induces autophagosome biogenesis rather than inhibits autophagic flux. By observing the colocalization of LC3 and LAMP-1, we further demonstrated that adipoRon increases the formation of autophagolysosomes. Collectively, our data demonstrated an inducing role of adipoRon in autophagosome formation and its flux in SMCs through activating TFEB. To our knowledge, this is the first study that links adipoRon action to the TFEB-autophagy pathway.

Serum is well known to promote SMC dedifferentiation leading to synthetic phenotypes. In the present study, we demonstrated that activation of TFEB by adipoRon markedly inhibited the proliferation of SMCs under serum stimulation. Our findings are consistent with previous studies showing that adipoRon exerts antiproliferative effects on SMC proliferation induced by PDGF-BB or ANG II (12, 16). The checkpoints for the G1/S and G2/M phases are critical for cell progression. Because adipoRon downregulated the regulatory proteins cyclin D1 and CDK4 for G1 phase, it is plausible that TFEB activation by adipoRon prevents cells from entering G1 phase. Moreover, in this study, we revealed that activating TFEB by adipoRon leads to inhibition of SMC migration, as characterized by F-actin reorganization and MMP-9 downregulation. Our data provided the first evidence that agonism of the adiponectin receptor exerts its inhibitory effects on SMC proliferation and migration in a TFEB-dependent manner. Autophagy plays a critical role in regulating the phenotypic plasticity of SMCs (28, 63). Enhanced, but not excessive, induction of autophagy by either rapamycin or statins contributes to maintaining the quiescent differentiated phenotype of vascular smooth muscle (29). This prevents SMC proliferation and migration induced by different atherogenic factors, including oxidized low-density lipoproteins, advanced glycation end products (AGEs), and thrombin (8, 19, 69). In another aspect, impaired autophagy induction contributes to cell growth or migration, which promotes neointimal lesions and atherosclerosis (14, 42, 48). Alternatively, the autophagy pathway can be impaired by lysosome dysfunction (33, 77). LAMP2 is a lysosome-associated membrane protein that is critical for lysosome function. A recent study reported that aberrant autophagy was found in SMCs of LAMP2-deficient mice, which was linked to luminal stenosis and medial thickening (45). Our recent studies also demonstrated that genetic deletion of acid sphingomyelinase or CD38 results in lysosome dysfunction and impairs autophagy maturation and autophagolysosome formation, which contributes to imbalanced SMC homeostasis (33, 72, 76, 77). Therefore, these previous findings support the view that adipoRon inhibits SMC dedifferentiation through enhancing the TFEB-dependent autophagy pathway.

Recent studies demonstrated that protein kinases, including mTOR, ERK2, and Akt, can phosphorylate TFEB, promote its cytosolic degradation, and thereby inhibit its activity (35, 51, 59). In contrast, calcium-dependent phosphatases dephosphorylate TFEB, enhances TFEB stability, and increases its activity (38). In the present study, we explored whether adipoRon-induced TFEB activation is associated with inhibition of kinases. Unexpectedly, adipoRon had no obvious effect on mTOR activity or even increased the activities of AMPK, ERK1/2, and Akt (Fig. 5). AMPK inhibits mTOR and increases autophagy in mammalian cells, including SMCs (53). In contrast, ERK1/2 and Akt are well-known activators of mTOR in SMCs. These previous findings reveal a complex temporal and spatial regulation of mTOR by its kinase activators and inhibitors. In our experiments, adipoRon did not affect mTOR activity, but it increased activities of both activators (ERK1/2 and Akt) and inhibitors (AMPK) of mTOR; therefore, the effects of these regulators on mTOR activity seem to counteract each other. A recent study reported that adipoRon inhibits PDGF-induced mTOR activity in murine aortic SMCs in an AMPK-independent manner, and this inhibitory effect is mediated through suppressing PDGF-receptor tyrosine phosphorylation (12). It should be noted that in this study, the SMC dedifferentiation is induced by serum, which contains multiple growth factors and nutrients that may increase mTOR activity independent of the PDGF-receptor pathway. Nonetheless, our data suggest that adipoRon-induced TFEB activation is not associated with inhibition of kinases. In contrast, our data reveal a role of intracellular Ca²⁺ in adipoRon-induced TFEB activation. AdipoRon increases ceramidase activity of adiponectin receptors and the production of a sphingolipid known as sphingosine-1-phosphate (S1P) (5, 17). S1P binds to membrane G protein-coupled receptors leading to elevated intracellular Ca²⁺ concentration via inositol trisphosphate IP₃-mediated Ca²⁺ mobilization (7, 32). AdipoRon may increase activities of calcium-dependent phosphatases such as calcineurin or protein phosphatase 2A, which, in turn, may dephosphorylate TFEB and enhance its nuclear translocation (3, 38). The detailed mechanisms of TFEB activation in SMCs deserve further elucidation.

The present study did not identify the downstream mediator(s) of the adipoRon-TFEB-autophagy axis that acts on proliferation and migration of SMCs. Recent studies suggest that the autophagic substrate p62/SQSTM1 cross talks with several signaling pathways that may interact with SMC dedifferentiation, such as nuclear factor erythroid 2-related factor 2 (Nrf2) signaling (41, 44) and DNA damage repair (68). Nrf2 is a critical transcriptional regulator of antioxidant responses to oxidative damages. Increased reactive oxygen species (ROS) causes oxidation of Kelch-like ECH-associated protein 1 (Keap1), the repressor of Nrf2, resulting in Nrf2 activation. Alternatively, accumulated p62/SQSTM1 can interact with Keap1 and, thereby, can cause a ROS-independent activation of Nrf2 (26). Nrf2 pathway is involved in the regulation of vascular smooth muscle homeostasis. For example, Levonen et al. (31) demonstrated that Nrf2 gene overexpression suppressed human and rabbit aortic SMC proliferation in vitro. In other studies, the Nrf2-mediated NADPH quinone dehydrogenase 1 contributes to the inhibition of vascular damage and SMCs proliferation and migration (20). Conversely, Nrf2 depletion enhances PDGF-stimulated SMC migration through increased ROS, Rac1, and sustained ERK1/2 signaling (1). TFEB-p62-Nrf2 axis may account for adipoRon’s effect on SMCs dedifferentiation. In addition to Nrf2, p62/SQSTM1 may also cross-talk with DNA double-strand repair machinery. Wang et al. (68) demonstrated that in cancer cells, defective autophagy leads to accumulation of nuclear p62, which, in turn, binds to E3 ligase ring finger protein 168 (RFN168) and inhibits RNF168-mediated recruitment of homologous recombination-related DNA repair proteins to double-strand breaks, resulting in decreased homologous recombination-mediated repair. Increased DNA damage activates the p53 pathway, which causes cell cycle arrest, apoptosis, or senescence in SMCs (4, 39). Therefore, it is plausible that the cross-talk between p62 and DNA damage response contributes to the adipoRon effect on SMC homeostasis.

Pathological vascular remodeling is involved in the pathogenesis of various vascular disorders that lead to myocardial infarction, stroke, peripheral vascular diseases, or graft failure in organ transplantation. In response to injury, SMCs dedifferentiate into synthetic phenotypes, leading to vascular SMC proliferation and migration, which are essential to the development of neointima. AdipoRon was previously reported to attenuate neointima formation in the femoral arteries in mice (12, 16). In the present study, we also confirmed that adipoRon inhibits SMC proliferation and migration using an ex vivo aortic ring sprouting assay. Importantly, aortic rings transduced with TFEB shRNA reversed the inhibitory effects of adipoRon. Recent studies demonstrated that in the vasculature, enhanced autophagy prevents progressive atherosclerosis or restenosis, whereas defective or excessive autophagy promotes these disease states (6). Thus, our findings support the view that instigation of TFEB autophagy by adipoRon maintains SMCs in a differentiated quiescent status with reduced proliferative and migratory potential, which contributes to the prevention of intimal hyperplasia and neointima development in the arterial wall.

In summary, our results show that adipoRon activates the TFEB-autophagy pathway that helps maintain the quiescent status of arterial SMCs and, thereby, prevents SMC dedifferentiation. Our findings provide novel insights into understanding the therapeutic effects of adipoRon on pathological vascular remodeling associated with metabolic disorders.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

Y.-T.W., Y.C., and Y.Z. conceived and designed research; Y.-T.W., J.C., and X.L. performed experiments; Y.-T.W., J.C., Y.C., and Y.Z. analyzed data; Y.-T.W. and Y.Z. interpreted results of experiments; Y.-T.W. and Y.Z. prepared figures; Y.-T.W., X.L., M.U., Y.C., and Y.Z. drafted manuscript; Y.-T.W., X.L., M.U., Y.C., P.-L.L., and Y.Z. edited and revised manuscript; Y.-T.W., J.C., X.L., M.U., Y.C., P.-L.L., and Y.Z. approved final version of manuscript.