Vascular injury and chronic arterial diseases result in exposure of VSMCs (vascular smooth muscle cells) to increased concentrations of growth factors. The mechanisms by which growth factors trigger VSMC phenotype transitions remain unclear. Because cellular reprogramming initiated by growth factors requires not only the induction of genes involved in cell proliferation, but also the removal of contractile proteins, we hypothesized that autophagy is an essential modulator of VSMC phenotype. Treatment of VSMCs with PDGF (platelet-derived growth factor)-BB resulted in decreased expression of the contractile phenotype markers calponin and α-smooth muscle actin and up-regulation of the synthetic phenotype markers osteopontin and vimentin. Autophagy, as assessed by LC3 (microtubule-associated protein light chain 3 α; also known as MAP1LC3A)-II abundance, LC3 puncta formation and electron microscopy, was activated by PDGF exposure. Inhibition of autophagy with 3-methyladenine, spautin-1 or bafilomycin stabilized the contractile phenotype. In particular, spautin-1 stabilized α-smooth muscle cell actin and calponin in PDGF-treated cells and prevented actin filament disorganization, diminished production of extracellular matrix, and abrogated VSMC hyperproliferation and migration. Treatment of cells with PDGF prevented protein damage and cell death caused by exposure to the lipid peroxidation product 4-hydroxynonenal. The results of the present study demonstrate a distinct form of autophagy induced by PDGF that is essential for attaining the synthetic phenotype and for survival under the conditions of high oxidative stress found to occur in vascular lesions.
VSMCs (vascular smooth muscle cells) are essential regulators of vascular function. In healthy arteries, VSMCs are located in the medial vascular layer where they express contractile proteins that help to regulate vessel tone and blood flow . During atherogenesis and arterial restenosis, VSMCs change from a contractile phenotype to a synthetic phenotype. This promotes their migration to the intima, increases their proliferative capacity and promotes the synthesis of extracellular matrix proteins . These changes in VSMC phenotype appear to be fundamental in regulating the composition and stability of vascular lesions .
Although several factors mediate VSMC phenotype transitions, PDGF (platelet-derived growth factor) is among the most robust of the phenotype-modulating agents and has been shown to be a primary regulator of smooth muscle cell growth and proliferation. Antibodies against PDGF [4–6] or PDGF receptors [7,8], antisense oligonucleotides to PDGF receptors [9,10], or PDGF aptamers  inhibit smooth muscle accumulation in the intima after balloon injury, and VSMCs lacking PDGFR-β (PDGF receptor β) show strikingly diminished neointimal accumulation after carotid artery ligation . Moreover, pharmacological inhibition of PDGF signalling reduces VSMC proliferation, migration and occupancy in the neointima [13–15]. Hence, understanding how PDGF regulates VSMC phenotype may be essential for developing better and more targeted strategies for preventing or ameliorating vascular disease.
The switch from the contractile to the synthetic VSMC phenotype caused by PDGF is initiated upon PDGF binding to surface receptors, which activate several intracellular signalling pathways that ultimately regulate gene expression and cellular function. These signalling pathways have been shown to result in decreased abundance of contractile proteins and increased expression of synthetic proteins such as osteopontin [1,16] and vimentin [17,18]. Such structural changes have been shown to be prerequisites for the enhanced cellular proliferation of VSMCs in culture . Although multiple studies have focused on identifying the molecular mechanisms involved in phenotype switching [1,2], it remains unclear how VSMCs can rapidly change from the contractile to the synthetic phenotype. Previous work from our laboratory has shown that autophagy is activated in VSMCs to remove oxidatively damaged proteins , which can form large aggregates that can be particularly difficult to proteolyse. This suggested to us that the contractile apparatus, which must be removed during cellular transition to the synthetic phenotype, may also be degraded by autophagy and that autophagy might be an essential regulator of VSMC viability in diseased vessels. Therefore we examined whether the PDGF-induced transition from the contractile to the synthetic phenotype is accompanied by an increase in autophagy and whether stimulation of autophagy is required for phenotype switching. The results of the present study support the notion that autophagy is essential for the conversion of VSMCs from a contractile into a synthetic phenotype and that this increase in autophagy in synthetic cells could also function to prevent cell death owing to oxidative stress.
Antibodies against α-SMA (α smooth muscle cell actin), calponin and α-tubulin were purchased from Sigma–Aldrich. The anti-osteopontin antibody was purchased from Santa Cruz Biotechnology. Recombinant FGF (fibroblast growth factor)-1 was obtained from eBiosciences. Antibodies against GAPDH (glyceraldehyde-3-phosphate dehydrogenase), ubiquitin, p- (phospho-) Akt/Akt, p-p70S6K [70 kDa ribosomal protein S6 kinase 2; also known as RPS6KB2 (ribosomal protein S6 kinase, 70kDa, polypeptide 2)]/p70S6K, p-AMPK (AMP-activated protein kinase)/AMPK and p-mTOR (mammalian target of rapamycin)/mTOR were purchased from Cell Signaling Technology. The anti-(collagen I) antibody was from Abcam. Recombinant rat PDGF was obtained from R&D Biosystems. Polyclonal antibodies against KLH (keyhole-limpet haemocyanin)–4-HNE (4-hydroxynonenal) were raised and tested as described previously . HRP (horseradish peroxidase)-conjugated anti-(rabbit IgG) and anti-(mouse IgG) secondary antibodies were obtained from Cell Signaling Technology. Goat anti-(mouse IgG)–Alexa Fluor® 555 was obtained from Invitrogen. The 3-MA (3-methyladenine), epoxomicin and bafilomycin A1 were obtained from Sigma–Aldrich. Spautin-1 was obtained from Cellagen Technology. All primers used for real-time PCR were designed using the Primer Express software from Applied Biosystems and then ordered from Integrated DNA Technologies. DAPI (4′,6-diamidino-2-phenylindole) stain was obtained from Invitrogen. Electrophoresis supplies were purchased from Bio-Rad Laboratories. ECL® reagents were purchased from GE Healthcare.
All animal procedures were performed in compliance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals and were approved by the University of Louisville Institutional Animal Care and Use Committee. Rat aortic smooth muscle cells (VSMCs) were isolated from the aortas of 6-week-old male Sprague–Dawley rats (Jackson Laboratories, Bar Harbor, ME, U.S.A.) as described previously . Briefly, rat aortas were cleaned of fat and adventitial tissue and then minced in sterile culture dishes. The minced aortas were then digested at 37°C in 5% CO2 for 1 h in an enzyme solution containing 0.1% collagenase type 1A, 0.05% elastase type III, 2 mg/ml BSA, 2 mM calcium chloride and 1% P/S (penicillin/streptomycin; Life Technologies, Invitrogen). After digestion, the supernatant was carefully removed and discarded. The undigested tissue was further digested for up to 3 h with fresh enzyme solution. Growth medium [DMEM (Dulbecco's modified Eagle's medium; Life Technologies, Invitrogen) containing 10% (v/v) FBS (fetal bovine serum; Atlanta Biologicals) and 1% (v/v) P/S] was added immediately after all of the tissue was digested, and the solution was centrifuged at 500 g for 5 min at 4°C to pellet the cells. The cells were resuspended in growth medium, counted and plated in a final volume of 2.5 ml at a density of 0.4–0.6×106 cells per 25 mm2 flask.
The purity of VSMCs was verified by flow cytometry using anticalponin and anti-α-SMA antibody staining (Supplementary Figure S1 at http://www.biochemj.org/bj/451/bj4510375add.htm). To ensure maintenance of the contractile phenotype, only cells between passages 2–7 were used. Cells were maintained at 37°C in a humidified atmosphere containing 5% CO2. At ~70% confluency, VSMCs were serum-starved in DMEM containing 0.1% FBS for 24 h. After the indicated treatments, the cells were rinsed twice with PBS and then lysed in a protein lysis buffer containing 25 mM Hepes, 1 mM EDTA, 1 mM EGTA, 0.1% SDS, 1% NP-40 (Nonidet P40) and 1× protease and phosphatase inhibitors. The Lowry DC assay (Bio-Rad Laboratories) was used for measuring the protein concentration of the crude cell extracts.
mRNA isolation and real-time PCR
mRNA was isolated from VSMCs using the TRIzol® reagent (Invitrogen) and the concentration was determined by measuring absorbance at 260 nm using a Nanodrop spectrophotometer (Thermo Scientific). A 20 μl reverse transcription reaction mixture containing 1 μg of mRNA, 10 units AMV (avian myeloblastosis virus) reverse transcriptase, 0.4 μM poly-T primer (dT18), 0.2 mM dNTP and 20 units RNasin® (Promega) was subjected to cDNA synthesis in a thermal cycler (Bio-Rad Laboratories). cDNA (2 μl) was then used for the amplification of the gene of interest by real-time PCR using SYBR Green (VWR).
Depending on the target protein, 0.5–25 μg of crude cell protein was applied to each lane of a 10.5–14% Bis-Tris/HCl gel and electroblotted on to a PVDF membrane. The membrane was then incubated overnight at 4°C using the appropriate dilutions of primary antibodies. PVDF membranes were then incubated at room temperature (22°C) with HRP-conjugated secondary antibodies. Immunoreactive bands were detected using a Typhoon scanner (SA Biosciences) after exposure to the ECL® detection reagent. Band intensity was quantified by using the TotalLab TL120 software.
Measurement of protein-bound 4-HNE
VSMCs were serum-starved for 24 h and then treated without or with PDGF (20 ng/ml) for 48 h. After PDGF treatment, cells were exposed to 50 μM 4-HNE in HBSS (Hanks balanced salt solution) for 30 min and then the medium was replaced with DMEM containing 10% FBS. Cells were then incubated in the 4-HNE-free medium for 3.5 h. Cells were lysed using lysis buffer and 25 μg was used for Western blotting to quantify protein–4-HNE adducts using an anti-protein–4-HNE antibody .
LDH (lactate dehydrogenase) activity assay
VSMCs were serum-starved for 24 h and then treated without or with PDGF (20 ng/ml) for 48 h. After PDGF treatment, cells were exposed to 50 μM 4-HNE in HBSS for 30 min which was then replaced with DMEM containing 10% FBS. After 16 h, a LDH assay was performed as described previously .
Adenoviral gene transfection
The GFP (green fluorescent protein)–LC3 (microtubule-associated protein light chain 3 α; also known as MAP1LC3A) plasmid was a gift from Professor Roberta Gottlieb (San Diego State University, San Diego, CA, U.S.A.). The plasmid was amplified in Escherichia coli cells, and the GFP–LC3 fragment was excised from the plasmid backbone. The GFP–LC3 adenovirus was then developed at Vector Biolabs. Briefly, the GFP–LC3 construct was cloned into a pAd5-dE1/E3 vector for viral packaging and amplification in HEK (human embryonic kidney)-293 cells. AdGFP without LC3 was used as a control viral vector. Adenoviruses were used at a MOI (multiplicity of infection) of 100 and GFP expression was used to ascertain transduction efficiency. AdGFP or AdGFP–LC3-transduced cells were then stimulated with PDGF at the indicated times. Before imaging, cells were incubated with DAPI for nuclear visualization.
Immunofluorescence staining and confocal imaging
VSMCs were cultured on glass chamber slides or glass-bottomed culture dishes and after the appropriate treatments were washed with PBS and then fixed in cold (−20°C) methanol for 5 min. The cells were then incubated for 30 min at room temperature in a blocking solution (5% BSA in PBS) followed by incubation for 1 h with a 1:500 dilution of anti-α-SMA primary antibody at room temperature. A 1:1000-diluted solution of goat anti-(mouse IgG)–Alexa Fluor® 555 secondary antibody was used for incubation after treatment with the primary antibody. After three washes with PBS, the cells were stained with DAPI for 10 min at room temperature. The α-SMA staining and GFP–LC3 puncta were visualized using a Nikon TE-2000E2 microscope interfaced with a Nikon A1 confocal system. DAPI was used to visualize nuclei. Prior to imaging the live cells, the cells were placed in Phenol Red-free DMEM supplemented with 25 mmol/l Hepes (pH 7.4). The individual fluorophores were illuminated as indicated below. All images were acquired through a 60× oil-immersion DIC (differential interference contrast) objective [NA (numerical aperture)=1.4], natively averaged at 2× magnification, scan direction set to one-way, image size was 1024×1024 and channel series was ON. Laser lines were modulated and integrated in the A1 system's AOTF (acousto-optic tuneable filter). For DAPI/GFP imaging, a 405 nm excitation line [Power=1.1, PMT HV (photomultiplier tube high voltage)=80 and Offset=0] with emission collected through a 450/50 nm bandpass filter and a 488 nm excitation line (Power=1.2, PMT HV=90 and Offset=0) with emission collected through a 525/50 nm bandpass filter were used with a pinhole size of 36.69 μm, the scanner zoom was set at 1.205 with a scan speed of 1. For DAPI/Alexa Fluor® 555 (α-SMA), a 405 nm excitation line (Power=2.0, PMT HV=75 and Offset=0) with emission collected through a 450/50 nm bandpass filter and a 561 nm excitation line (Power=1.1, PMT HV=90 and Offset=0) with emission collected through a 595/50 nm bandpass filter were used with a pinhole size of 36.07 μm, the scanner zoom was set at 1.000 with a scan speed of 1/8.
Migration of VSMCs was measured using the scratch wound assay [23,24]. Briefly, VSMCs were plated in 96-well cell culture plates and allowed to grow to a 70% confluency. The cells were then serum-starved for 24 h in DMEM containing 0.1% FBS and then stimulated with PDGF (20 ng/ml) or vehicle in the absence or presence of spautin-1 (10 μM) for 24 h. A p200 pipette tip was used to create a scratch on the bottom of each well and the cells were incubated further for 24 h. The distance between the edges of the scratch was measured immediately after scratching and again after 24 h. Migration was quantified and expressed as follows: percentage closure=[(distance immediately after scratching−distance 24 h after scratching)/distance immediately after scratching ×100.
Transmission electron microscopy
VSMCs were grown on cover slips, serum-starved for 24 h and then treated without or with PDGF (20 ng/ml) for 48 h. The cells were fixed and imaged as described previously .
Results are means±S.E.M. Multiple groups were compared using one-way ANOVA, followed by Bonferroni post-hoc tests. Unpaired Student's t test was used for two-group comparisons. P<0.05 was considered significant.
Platelet-derived growth factor induces phenotype switching in VSMCs
To examine the mechanisms regulating the phenotypic transition of VSMCs, we first measured changes in molecular markers of contractile and synthetic VSMC phenotypes. For this, VSMCs were deprived of serum for 24 h to induce cell-cycle arrest. The cells were then incubated with vehicle or PDGF-BB (20 ng/ml) for 24 and 48 h. Relative mRNA and protein abundance was measured by real-time PCR and Western blotting respectively. As shown in Figure 1(A), PDGF treatment for 24 h resulted in a ~50% decrease in the mRNA levels of α-SMA and calponin and a robust up-regulation of osteopontin (12-fold) and vimentin (2-fold). After 48 h, the levels of α-SMA protein were decreased by 25%, calponin abundance was decreased by 75% and osteopontin protein was elevated more than 2-fold (Figures 1B and 1C). As observed by immunofluorescence imaging, the α-SMA filaments showed marked disorganization in the PDGF-treated cells compared with the control cells, and this was accompanied by marked changes in cell morphology (Figure 1D). These results confirm previous findings showing that PDGF promotes the phenotypic transition of VSMCs [1,19,25].
PDGF induces a contractile-to-synthetic phenotype switch in VSMCs
PDGF induces autophagy in VSMCs
The relatively rapid loss of contractile proteins suggested to us that proteolytic mechanisms may function to remove the contractile apparatus to aid in the transition from the contractile to the synthetic phenotype. Indeed, previous studies have shown that proteasome and calpain inhibitors affect VSMC phenotype [26–28]; however, these studies showed that proteasome inhibition, counterintuitively, decreases markers of the contractile phenotype , resulting in reduced cell contractility . Therefore we hypothesized that autophagy may be important for the removal of contractile proteins and protein complexes. To test this hypothesis, we first assessed the effects of PDGF on autophagy in VSMCs by examining LC3-II formation, an indicator of autophagy [29,30]. As shown in Figures 2(A–C), PDGF stimulation caused a time-dependent increase in LC3-II formation. A significant increase in LC3-II was observed 12 h after PDGF stimulation and maximal increases in LC3-II were obtained 48 h after PDGF treatment, suggesting that PDGF promotes autophagy.
PDGF-BB induces autophagy in VSMCs
Because the formation of LC3-II is transient and the protein can be rapidly degraded in the lysosome, an increase in LC3-II could also paradoxically indicate a decrease in autophagy. Hence, to more accurately assess autophagic flux, we inhibited the lysosomal degradation of LC3-II by treating cells with bafilomycin A1, an inhibitor of the vacuolar-type H+-ATPase [30,31]. As shown in Figures 2(D–F), treatment of the control cells with bafilomycin A1 increased LC3-II abundance, which was further increased in PDGF-stimulated cells that were treated with the inhibitor. These data indicate that autophagic flux is higher in PDGF-treated cells. These effects appear to be specific to PDGF because treatment with FGF-1, a known VSMC mitogen , did not affect the abundance of contractile proteins or increase LC3-II formation, despite increasing cell proliferation to levels similar to that of PDGF (Figures 2G–2J). To obtain further evidence for the induction of autophagy by PDGF, we transduced VSMCs with a GFP–LC3 adenovirus and assessed the formation of punctate LC3, which indicates autophagosomal localization . As shown in Figures 3(A) and 3(B), there was a 2-fold increase in fluorescent puncta in PDGF-treated cells, and this was further increased by bafilomycin A1 treatment.
PDGF-BB increases autophagosome formation and promotes extensive vacuolization
PDGF induces ultrastructural changes and extensive vacuolization in VSMCs
Although LC3-II formation is a validated marker of autophagy, changes in the ultrastructure showing the formation of autophagosomes is considered the ‘gold-standard’ for documenting autophagy. Hence, to examine ultrastructural changes, VSMCs were treated with PDGF for 48 h and then visualized by transmission electron microscopy. Transmission electron micrographs showed that treatment with PDGF (20 ng/ml) for 48 h caused a ~3-fold increase in single-membrane autophagic vacuoles (some containing electron-dense material) as well as early double-membrane vacuoles (Figures 3C–3F), similar to those shown previously [20,33,34]. In addition, PDGF treatment resulted in the consistent formation of a singular large double-membrane phagosome per cell (labelled in Figures 3D and 3E), which may serve as a designated compartment for the digestion of intracellular materials. Collectively, these structural changes are in concordance with our immunological and fluorescence imaging data and suggest that PDGF is a robust inducer of the autophagic programme.
PDGF-induced autophagy occurs via an AMPK- and mTOR independent mechanism
Activation of autophagy in mammalian cells usually involves the inhibition of the mTOR, an important signalling molecule necessary for the growth and proliferation of cells. During autophagy activation mTOR is a downstream target of multiple signalling pathways, such as the Akt signalling pathway and the AMPK pathway, which inhibit and activate autophagy respectively. We found that both Akt and mTOR, as well as the substrate of mTOR p70S6K/p85S6K, were activated despite the induction of autophagy (Figure 4). This suggested to us that PDGF-induced autophagy does not occur via inhibition of mTOR. Because AMPK activation was inhibited after PDGF stimulation, PDGF-induced autophagy is probably independent of AMPK activity as well. We also examined whether autophagy induced by PDGF could be owing to nutrient starvation. For this we incubated the cells with PDGF in the absence or presence of excess essential amino acids or pyruvate. As shown in Supplementary Figure S2 (at http://www.biochemj.org/bj/451/bj4510375add.htm), increasing amino acid and pyruvate availability did not affect LC3 expression in PDGF-treated cells, suggesting that this form of autophagy could not be attributed to well-described starvation pathways.
PDGF-induced autophagy is AMPK and mTOR independent
PDGF-induced autophagy is required for phenotype switching
To examine whether autophagy plays a role in VSMC phenotype switching, we pre-treated VSMCs with the pharmacological inhibitor of autophagy, 3-MA , prior to stimulating the cells with PDGF. As shown in Figures 5(A) and 5(B), 3-MA decreased GFP–LC3 puncta formation by more than 50%. This inhibition of autophagy was associated with partial stabilization of the contractile markers calponin and α-SMA (Figures 5C–5E) and inhibition of smooth muscle cell proliferation (Figure 5F). We further confirmed the role of autophagy in protein degradation during phenotype switching by using the downstream inhibitor of autophagy bafilomycin A1. As shown in Supplementary Figure S3 (at http://www.biochemj.org/bj/451/bj4510375add.htm), bafilomycin A1, which inhibits the acidification of autophagosomes and the proteolytic degradation of their contents, also prevented the degradation of calponin in PDGF-treated cells.
Inhibition of autophagy prevents PDGF-induced phenotype switching
Because of potential off-target effects of 3-MA [36–38] and the relatively high concentrations of the compound needed to inhibit the autophagic machinery, we tested whether spautin-1, a structurally dissimilar inhibitor of autophagy, would affect the VSMC phenotype. Spautin-1 has been recently reported to prevent autophagy by inhibiting the deubiquitinases USP10 (ubiquitin-specific peptidase 10) and USP13 (ubiquitin-specific peptidase 13), which leads to beclin 1 and Vps34 [vacuolar protein sorting-associated protein 34; also known as PI3K (phosphoinositide 3-kinase) III degradation . Treatment of VSMCs with spautin-1 (10 μM) prevented PDGF-induced increases in LC3 abundance and prevented losses of calponin and α-SMA after PDGF exposure (Figures 6A–6D). Spautin-1 also prevented the PDGF-induced disorganization of actin filaments (Figure 6E). Interestingly, we did not find changes in the levels of beclin 1 and Vps34 in spautin-1-treated cells (Supplementary Figure S4 at http://www.biochemj.org/bj/451/bj4510375add.htm), suggesting that its mechanism of inhibition of autophagy in VSMCs may be independent of its previously described effect on deubiquitinase activity.
Spautin-1, an inhibitor of autophagy, prevents PDGF-induced phenotype switching
To determine whether the inhibition of autophagy prevents the functional changes associated with the synthetic VSMC phenotype, we measured cell proliferation, migration and the extracellular matrix component collagen I after PDGF treatment in the absence or presence of spautin-1. Similar to that shown with 3-MA (Figure 5F), spautin-1 prevented PDGF-induced cell proliferation (Figure 7A). In addition, PDGF-induced migration, as measured by the scratch wound assay [23,24], was inhibited in spautin-1-treated cells (Figures 7B and 7C), as was collagen I synthesis (Figures 7D and 7E). Collectively, these results suggest that autophagy is a major regulator of VSMC phenotype and that spautin-1 is particularly effective in preventing conversion into the synthetic phenotype.
Spautin-1 prevents the functional characteristics of the synthetic phenotype typically induced by PDGF
To determine if the proteasome plays a role in regulating VSMC phenotype switching, we treated cells with the proteasome inhibitor epoxomicin  prior to PDGF treatment and examined contractile protein expression. Although epoxomicin treatment inhibited proteasomal activity, as shown by the increased abundance of ubiquitinated proteins (Figures 8A and 8B), the loss of contractile proteins owing to PDGF treatment remained largely unaffected (Figures 8C–8E). Collectively, these results suggest that autophagy, and not proteasomal activity, is required for phenotypic changes in PDGF-stimulated VSMCs.
Inhibition of the proteasomal pathway does not affect contractile protein abundance in PDGF-treated cells
Synthetic VSMCs are resistant to aldehyde-induced cell death
Lesions in atherosclerotic and restenotic vessels are characterized by increased amounts of proteins modified by lipid peroxidation products such as 4-HNE [41,42]. Because previous findings suggest that autophagy prevents toxicity owing to 4-HNE , we hypothesized that synthetic VSMCs, having a higher autophagic flux, might be protected from 4-HNE-induced cell death. This could be important because synthetic VSMCs are the abundant phenotype found in vascular lesions; hence, their survival may be impacted by their ability to handle oxidative insults. Indeed, as shown in Figure 9(A), the viability of synthetic VSMCs challenged with 4-HNE was significantly enhanced compared with 4-HNE-challenged contractile VSMCs. Consistent with the role for autophagy in clearing oxidatively damaged proteins [20,43], synthetic VSMCs resisted accumulation of protein–4-HNE adducts owing to exogenous 4-HNE exposure (Figures 9B and 9C). Inhibiting autophagy with spautin-1 promoted accumulation of 4-HNE-damaged proteins (Figures 9D and 9E). These results suggest that PDGF-induced autophagy may be important for promoting the survival of VSMCs found in diseased vessels.
Synthetic VSMCs are resistant to 4-HNE-induced toxicity
The present study demonstrates a novel role for autophagy in regulating VSMC phenotype. We found that PDGF, which promotes the development of the synthetic VSMC phenotype, is a robust inducer of autophagy and that pharmacological inhibition of autophagy by three structurally unrelated inhibitors blocked the degradation of contractile proteins. Spautin-1 was particularly robust in preventing the PDGF-induced synthetic phenotype. This inhibitor of autophagy not only prevented degradation of calponin and α-SMA, but it also prevented proliferation, migration and synthesis of collagen I after exposure of VSMCs to PDGF. We also found that the synthetic phenotype was more resistant to the electrophilic stress commonly encountered in diseased vessels, and this was associated with an increased ability to remove proteins damaged or modified by electrophiles. These findings uncover PDGF-mediated autophagy as an important regulator of VSMC phenotype and survival.
Previous studies have identified numerous factors that regulate phenotype plasticity in VSMCs. For example, contractile agonists, reactive oxygen species and extracellular matrix components regulate the expression of contractile genes [44–50]. Conversely, growth factors, such as PDGF, extracellular matrix components and oxidized phospholipids, are known to favour the synthetic phenotype [25,51–53]. The signalling pathways known to trigger the synthetic phenotype culminate in the displacement of myocardin, a co-activator of serum-response factor, from the consensus CArG box upstream of smooth muscle cell marker genes . PDGF is known to act through transcription factors such as KLF4 (Krupple-like factor 4) and Elk-1 as well as microRNAs (such as miR221 and miR222), leading to the displacement of myocardin and down-regulation of VSMC contractile genes [55–58]. Hence, most studies to date examined transcriptional and translational changes regulating phenotypic transition.
We posited that one limiting event to transition to the synthetic phenotype would be removal of the contractile apparatus. Degradation of contractile proteins would likely involve the proteasome, autophagy or both. Interestingly, the proteasome has been shown to play a role in VSMC hyperplasia, but not phenotype switching [59,60]. Paradoxically, inhibitors of the proteasome decrease markers of the contractile phenotype  resulting in reduced cell contractility . In the present study we found that inhibition of the proteasome with epoxomicin had little effect on contractile protein abundance, suggesting that autophagy could be the major proteolytic device employed by the cell to remove contractile proteins.
We found that PDGF promoted a robust form of autophagy characterized by increased LC3-II abundance, augmented autophagic flux, and the formation of autophagosomes and autophagic vacuoles. The induction of autophagy was specific to PDGF; the mitogen FGF, while increasing proliferation to a similar extent as PDGF, did not induce autophagy. Whereas factors such as TNFα (tumour necrosis factor α) and osteopontin have been shown to promote autophagy in VSMCs [61,62], there are surprisingly few reports of induction of autophagy by growth factors. In fibroblasts and cancer cells, connective tissue growth factor activates autophagy and regulates proliferation , and transforming growth factor β activates autophagy in hepatocellular carcinoma cells . In these cases, the induction of autophagy is usually associated with diminished proliferation; however, we found that autophagy may be important for PDGF-mediated hyperproliferation, as inhibition of autophagy with 3-MA or spautin-1 prevented the cellular proliferation induced by PDGF. This is consistent with a recent study showing that 3-MA prevents arterial restenosis . The discrepancy in proliferation between the different inhibitors is likely to be related to their distinct molecular targets. Although 3-MA inhibits autophagy via inhibiting the PI3K–Akt pathway , which is also important in regulating cell proliferation, spautin-1 has no effect on the Akt pathway (Supplementary Figure S5 at http://www.biochemj.org/bj/451/bj4510375add.htm) and inhibits autophagy through a distinct mechanism. It is also possible that PDGF-mediated proliferation may not be directly related to the induction of autophagy. PDGF-mediated proliferation in SMCs may depend on FGF release and FGFR (FGF receptor) activation , and we show that FGF-1 alone can promote proliferation despite having no effects on autophagy or phenotype switching. Hence, it is possible that the PDGF-mediated FGF autocrine effects are retarded when autophagy is inhibited. Additionally, it may be that the decreased generation of metabolic building blocks, e.g. free amino acids, a natural consequence of enhanced autophagic activity, could be used to supply the energy requirements of increased proliferation.
The induction of autophagy by PDGF does not appear to be mediated by AMPK or mTOR. Exposure of cells to PDGF resulted in a decrease in the phosphorylation of AMPK and increased phosphorylation of mTOR and its downstream substrates p85S6k/p70S6k (Figure 4). Because activation of autophagy triggered by nutrient deprivation is typically linked to enhanced AMPK phosphorylation or inhibition of mTOR, it is unlikely that these pathways regulate the autophagic programme initiated by PDGF. It remains unclear why rapamycin, which is an inhibitor of mTOR and activator of autophagy, promotes the contractile phenotype in VSMCs [68,69], whereas PDGF-mediated autophagy induces the synthetic phenotype. It is probable that the form of autophagy induced by PDGF is uniquely poised to degrade contractile proteins and is tailored to regulate the phenotype switch.
The observation of the present study that three structurally distinct inhibitors of autophagy stabilized the contractile phenotype supports the view that autophagy is required for PDGF-induced phenotype conversion. Spautin-1 was most robust in stabilizing the abundance of contractile proteins, and it completely prevented hyperproliferation, migration and synthesis of collagen I after PDGF exposure. The inhibition of autophagy with spautin-1 did not affect PDGF signalling through Akt and Erk (extracellular-signal-regulated kinase; Supplementary Figure S5), suggesting further that the inhibition of phenotype switching was due to stabilization of the contractile apparatus. Interestingly, spautin-1 appeared to inhibit autophagy in a manner different from what has been published previously ; spautin-1 treatment did not decrease the levels of Vps34 or beclin 1 in VSMCs (Supplementary Figure S4), despite completely inhibiting PDGF-induced autophagy and preventing phenotype transition. Therefore the mechanism by which spautin inhibits autophagy in VSMCs remains unclear.
It should be noted that the inhibitors of autophagy used in the present study could have autophagy-independent effects that might regulate the VSMC phenotype. For example, the effects of bafilomycin A1 on vesicular trafficking have been shown to inhibit canonical Wnt signalling , which is an important regulator of VSMC proliferation, migration and development . Similarly it is possible that unidentified off-target effects of spautin-1 could affect the VSMC phenotype. Nevertheless, the finding that all three inhibitors of autophagy, i.e. spautin-1, 3-MA and bafilomycin A1, prevented PDGF-induced losses of contractile proteins supports the concept that autophagy regulates the contractile-to-synthetic VSMC phenotype transition. Moreover, the remarkable efficacy of spautin-1 for inhibiting PDGF-induced autophagy and preventing phenotype switching in vitro suggests that it might be a useful therapeutic agent for preventing phenotype switching and proliferation in certain circumstances of vascular injury, such as restenosis.
In addition to contractile proteins, autophagy has been shown to be important in preventing the accumulation of damaged or aggregated proteins [20,43,72]. We found that the increase in autophagic flux in PDGF-treated VSMCs was sufficient to decrease cell death and the abundance of protein–4-HNE adducts after aldehyde challenge, suggesting that PDGF-induced autophagy primes synthetic VSMCs to better tolerate the high levels of 4-HNE [41,73,74] and oxidant stress prevalent in vascular lesions. This tolerance may be important for maintaining plaque stability, which would be consistent with findings showing that induction of autophagy in macrophages prevents the accumulation of damaged proteins and stabilizes atherosclerotic plaques . Thus, overall, the promotion of autophagy, cell survival and increased formation of extracellular matrix by PDGF might safeguard plaque-resident VSMCs against oxidative injury while also stabilizing lipid-laden atherosclerotic plaques. Interestingly, the form of autophagy induced by PDGF appears to be functionally different from what has been described to occur with other stimuli. For example, TNFα and IGF-1 (insulin growth factor 1) were shown to increase autophagy in VSMCs, yet promote an autophagic form of cell death . Further studies are required to elucidate how apparently divergent forms of autophagy regulate atherosclerosis progression and plaque stability.
In summary, the present study identifies a new mechanism by which autophagy is activated and shows that autophagy is critical for attaining a synthetic VSMC phenotype and for increasing resistance to oxidative stress. Our observations also suggest that therapies targeting autophagy might be useful in preventing aberrant smooth muscle cell growth, but that such interventions may have deleterious effects on synthetic smooth muscle cell survival under conditions of oxidative stress. Future studies are required to identify the detailed molecular mechanism(s) by which PDGF activates the autophagic programme and to test how modulating PDGF-mediated autophagy affects the progression of vascular disease.
AMP-activated protein kinase
Dulbecco's modified Eagle's medium
fetal bovine serum
fibroblast growth factor
green fluorescent protein
Hanks balanced salt solution
microtubule-associated protein light chain 3 α
mammalian target of rapamycin
platelet-derived growth factor
HV, photomultiplier tube high voltage
70 kDa ribosomal protein S6 kinase
α smooth muscle actin
tumour necrosis factor α
vacuolar protein sorting-associated protein 34
vascular smooth muscle cell
Joshua Salabei designed and performed the experiments, analysed data and helped write the paper; Timothy Cummins helped perform experiments and analyse data; Mahavir Singh helped standardize the experiments using the GFP–LC3 vector; Steven Jones provided reagents, materials and guidance for confocal imaging experiments as well as helping to write the paper; Aruni Bhatnagar helped to design experiments, analyse data and write the paper; and Bradford Hill designed experiments, analysed data and wrote the paper.
This work was supported by the National Institutes of Health [grant numbers GM103492, HL083320, HL094419, HL055477 and HL078825].