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Am J Physiol Heart Circ Physiol 290: H2393-H2401, 2006. First published January 13, 2006; doi:10.1152/ajpheart.01161.2005
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Caveolin-1-deficient aortic smooth muscle cells show cell autonomous abnormalities in proliferation, migration, and endothelin-based signal transduction

Ghada S. Hassan,1 Terence M. Williams,1 Philippe G. Frank,1,2,3 and Michael P. Lisanti1,3,4

Departments of 1Molecular Pharmacology and Medicine and 2Urology, Albert Einstein College of Medicine, Bronx, New York; 3Department of Cancer Biology, Kimmel Cancer Center, Thomas Jefferson University, Philadelphia, Pennsylvania; and 4Muscular and Neurodegenerative Disease Unit, University of Genoa, and G. Gaslini Pediatric Institute, Genoa, Italy

Submitted 2 November 2005 ; accepted in final form 10 January 2006


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
We previously showed that ablation of caveolin-1 (Cav-1) gene expression in mice promotes neointimal hyperplasia in vivo, a phenomenon normally characterized by smooth muscle cell (SMC) migration and proliferation. Whether these defects are cell autonomous, i.e., due to loss of Cav-1 within SMCs or loss of Cav-1 expression in other adjacent cell types in vivo, remains unknown. Cav-1 has been shown to associate with receptors for many vasoactive factors on the SMC surface. Therefore, Cav-1 might be an important regulator of SMC proliferation, migration, and signal transduction. To mechanistically dissect the role of Cav-1 in SMC signaling, we isolated SMCs from the aortas (AoSMCs) of Cav-1-deficient (Cav-1–/–) mice and characterized these cells with respect to their proliferation, migration, and Ca2+ response to an important vasoactive factor, endothelin-1 (ET-1). 5-Bromo-2'-deoxyuridine incorporation and a wound-healing assay showed an increase in proliferation and migration rates in Cav-1–/– compared with wild-type (Cav-1+/+) AoSMCs. Cav-1–/– AoSMCs demonstrated upregulation of phosphorylated ERK1/2, cyclin D1, and proliferating cell nuclear antigen and reduced expression of the cyclin-dependent kinase inhibitor p27Kip1. The Ca2+ response was examined in the presence of ET-1 and assessed by confocal microscopy with the Ca2+-sensitive fluorescent probe fluo 3. When treated with ET-1, Cav-1–/– AoSMCs exhibited a faster and larger increase in free intracellular Ca2+ than Cav-1+/+ cells. The ET-1-induced response in Cav-1–/– cells was mediated by the ETB receptor, as shown using the ETB receptor antagonist BQ-788 and the ETA receptor antagonist BQ-123. In Cav-1–/– cells, ETA receptor expression was reduced and ETB receptor expression was upregulated. Therefore, Cav-1 ablation increased the ET-1-induced Ca2+ response in SMCs by altering the type and expression level of the ET receptor (i.e., receptor isoform switching). These data suggest a novel regulatory role for Cav-1 in SMCs with respect to their proliferation, migration, and Ca2+-mediated signaling.

vascular disease; calcium response; endothelin receptors; neointimal hyperplasia; caveolae; caveolin

SMOOTH MUSCLE CELLS (SMCs) are used to construct the walls of hollow organs, such as the stomach, intestine, bladder, uterus, and blood vessels (44). Their proliferation, migration, and contraction processes are highly controlled by many factors, ensuring proper functioning of the corresponding organ (31). In the vasculature, abnormalities in these processes are associated with many different classes of vascular disease, such as hypertension, atherosclerosis, and restenosis. However, many of the factors that contribute to the development of vascular disease remain unknown.

Caveolin-1 (Cav-1) has been recently identified as a potential regulator of smooth muscle physiology and pathophysiology (45). Caveolins are the structural units of caveolae organelles. Caveolae microdomains are associated with critical signaling events in various cell types. Multiple studies have shown that caveolins not only form scaffolds for the assembly of signaling molecules, but they also regulate the activation state of these caveolin-associated molecules (41, 42). For example, Cav-1 was shown to negatively regulate the activation state of endothelial nitric oxide synthase, as well as v-Src, H-Ras, and extracellular signal-regulated kinase (ERK1/2) (41–43). Therefore, it was suggested that Cav-1 may possess tumor suppressor capabilities (19).

Cav-1 expression was shown to inversely correlate with the transformed phenotype of NIH/3T3 cells (15, 20, 29). Also, studies have demonstrated that a deficiency of Cav-1 expression induces hypercellularity in lung tissue, hyperproliferation of mouse embryonic fibroblasts in culture (13, 40), hyperplasia of mammary epithelial cells (30), and cardiac hypertrophy (11). In addition, we recently showed that Cav-1 ablation accentuated neointimal formation induced by blood flow interruption, a phenomenon characterized by SMC migration and proliferation (24). On the other hand, Cav-1 was shown to positively regulate the interaction of certain vasoactive factors, such as angiotensin II and endothelin-1 (ET-1), with their corresponding receptors, angiotensin type 1 (AT1) and ET type A and B (ETA and ETB) (8, 25, 26, 46, 51). Therefore, the Ca2+ response of SMCs to certain vasoactive substances may be regulated by Cav-1. Several reports have shown that the presence of intact caveolae is essential for arterial contraction induced by vasoactive substances, such as serotonin and ET-1 (3, 14). However, most of these studies investigated vessel contraction in the presence and absence of caveolae-disrupting drugs (e.g., relatively toxic cholesterol-binding agents, such as nystatin and methyl-beta-cyclodextrin), rather than by evaluation of a direct role for caveolins in Ca2+ signaling events in SMCs.

Here, we have systematically evaluated the functional role of Cav-1 in vascular smooth muscle by characterizing SMCs isolated from the aortas (AoSMCs) of Cav-1-deficient (Cav–/–) and wild-type (WT, Cav+/+) mice with respect to their proliferation and migration properties. We also investigated the Ca2+ response of these SMCs after treatment with an important vasoactive factor, ET-1, by assessing Ca2+ influx. For this purpose, we used mice lacking the Ink4a locus (47), a tumor suppressor gene. Ink4a-deficient (Ink4a–/–) mice were previously shown to yield cells that could divide indefinitely in culture (immortalized) and are nontransformed (47). Therefore, we isolated SMCs from the aortas of Ink4a–/– mice (Ink4a–/–/Cav-1+/+). To obtain AoSMCs that lack Cav-1 expression, we interbred Cav-1–/– mice with Ink4a–/– mice to generate double-knockout mice and isolated SMCs from their aortas (Ink4a–/–/Cav-1–/–) as well.

Peterson et al. (37) showed that Cav-1 protein expression is decreased 1) in arterial SMCs during neointimal hyperplasia (in a rabbit artery injury model) and 2) upon platelet-derived growth factor (PDGF)-induced proliferation of WT vascular SMCs in culture. Interestingly, transient overexpression of Cav-1 in these WT cells inhibited their growth response (37). However, these growth-inhibitory effects may be simply due to the toxic effects of Cav-1 transient overexpression. Thus the growth-inhibitory role of Cav-1 in vascular SMCs remains controversial.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Materials

Rabbit polyclonal antibodies directed against total ERK1/2 and activated phosphorylated ERK1/2 were obtained from Cell Signaling (Beverly, MA); rabbit polyclonal antibodies to Cav-1 (N-20) and murine p21Cip1 and p27Kip1 from Santa Cruz Biotechnology (Santa Cruz, CA); anti-cyclin D1 rabbit polyclonal antibody from NeoMarkers (Fremont, CA); anti-proliferating cell nuclear antigen (PCNA) monoclonal antibody and anti-ETA rabbit polyclonal antibody from BD Pharmingen (San Diego, CA); anti-ETB polyclonal antibody from Alamone Laboratories (Jerusalem, Israel); anti-{alpha}-smooth muscle actin ({alpha}-SM actin) from Sigma-Aldrich (St. Louis, MO); anti-GAPDH from Research Diagnostics; and ET-1, the ETA receptor antagonist BQ-123, and the ETB receptor antagonist BQ-788 from American Peptide (Sunnyvale, CA).

Animal Studies

All animals used for isolation of cultured SMCs were from the C57Bl/6 background. Mice were housed and maintained in a barrier facility at the Institute for Animal Studies, Albert Einstein College of Medicine, which approved the animal protocols. Mice were kept on a 12:12-h light-dark cycle with ad libitum access to food and water. Cav-1–/– mice were generated as we previously described (40). Ink4a–/– mice were generated by Dr. Ron DePinho (Harvard Medical School). Genotyping of Cav-1/Ink4a double-knockout mice was performed by PCR as previously described (27, 40, 47).

AoSMC Isolation and Cell Culture

Vascular SMCs were isolated from the aortas of male mice at 1–2 mo of age using a modification of previously described techniques (23, 39). Briefly, animals were killed by CO2 asphyxiation. The aorta attached to the spinal cord was dissected from the ilial bifurcation but left attached to the left ventricle. A 1-ml syringe with a 32-gauge needle was used to puncture the left ventricle and flush the aorta with sterile PBS. The aorta was removed with sterile microdissecting scissors and directly placed in cold Ca2+- and Mg2+-free Hanks' balanced salt solution (HBSS; GIBCO, Rockville, MD) supplemented with penicillin (100 U/ml) and streptomycin (100 µg/ml) and then transferred under aseptic conditions (using a tissue culture hood). With the help of surgical loupes (x2.5 magnification capacity), fat and the adventitial layer were dissected away in a petri dish containing ice-cold HBSS and discarded. The remaining tissue was placed in a fresh solution of HBSS and cut into small (~1- to 2-mm2) pieces with use of a no. 10 scalpel. These pieces were then placed onto a 300-µl drop of enzyme digestion solution consisting of 0.1% (wt/vol) collagenase (type II), 0.037% (wt/vol) soybean trypsin inhibitor, and 0.2% (wt/vol) crystallized BSA in medium 199 (M199; GIBCO). The sample was transferred to a small tissue culture tube and incubated in a standard culture incubator at 37°C for 12–16 h. After the digestion period, 3 ml of M199 were added to the tube, and the sample was transferred to a 15-ml conical tube and centrifuged. The pellet was resuspended in 5 ml of fresh medium and centrifuged. The cells were then resuspended in 200 µl of smooth muscle growth medium (SMGM; Cambrex BioScience, Rockland, IL), transferred to a single well of a 24-well plate, and incubated at 37°C for 5–7 days. Fresh medium was then added. At confluence, cells (primary passage 1) were split and plated into larger wells. This procedure was repeated until the cells reached confluence in a 12.5-cm2 flask (usually passage 2 or 3). At this point, an aliquot of cells was frozen and passaged again and/or used for the experiments described below. All cells were cultured in SMGM supplemented with 5% fetal bovine serum, penicillin (100 U/ml), and streptomycin (100 µg/ml). Only early-passage (passage 3 or 4) cells were used for the experiments described below.

Immunofluorescence Techniques

All steps for immunofluorescence were carried out at room temperature. AoSMCs were grown on coverslips, washed with PBS, and fixed for 30 min in PBS containing 2% paraformaldehyde. After fixation, the cells were briefly rinsed with PBS and permeabilized with 0.1% Triton X-100 for 10 min. The cells were then incubated for 10 min with a 50 mM NH4Cl-PBS solution to quench free aldehyde groups. Next cells were incubated with primary antibodies for 60 min in PBS containing 0.2% BSA and then washed three times with PBS (10 min each) and incubated for 30 min with an FITC-conjugated goat anti-mouse antibody (5 µg/ml) and/or a lissamine rhodamine B sulfonyl chloride-conjugated donkey anti-rabbit antibody (5 µg/ml) in PBS containing 0.2% BSA. The cells were washed three times with PBS (10 min each) and mounted on slides with antifade reagent (Slow-Fade, Molecular Probes, Eugene, OR).

Proliferation Assays: 5-Bromo-2'-Deoxyuridine Incorporation

The cells were incubated with 50 mM 5-bromo-2'-deoxyuridine (BrdU; Sigma-Aldrich, St. Louis, MO) in smooth muscle basal medium (SMBM) supplemented with 0.1% BSA and penicillin-streptomycin for 4 h at 37°C. During this period, BrdU was incorporated into the DNA of dividing cells. The cells were then immunostained using a modification of the immunofluorescent technique described above. Before incubation with the primary antibody monoclonal anti-BrdU IgG (Oncogene, Boston, MA), the cells were incubated in 2% HCl supplemented with 0.5% Triton X-100 for 10 min to denature their DNA. After they were incubated for 30 min with an FITC-conjugated anti-mouse antibody (5 µg/ml) and counterstained with Hoechst nuclear dye (10 µg/ml; Molecular Probes), the cells were mounted. Statistical significance was determined using Student's t-test. P < 0.05 was considered significant.

Migration Assays

For measuring the migration rate of AoSMCs, cells from passage 3–4 were seeded in 60-mm sterile culture dishes. Confluent layers were scraped with a 200-µl pipette tip, and the cells were washed with PBS and kept in SMBM supplemented with 0.1% BSA and penicillin-streptomycin. Images were acquired immediately and 24 and 48 h after wound induction. Migration rates were calculated for cells moving from the wound edge to the cell-free space. Statistical significance was determined using Student's t-test. P < 0.05 was considered significant.

Immunoblot Analysis

To generate cell lysates, AoSMCs were plated at a density of ~1–2 x 106 cells in complete medium and cultured for 18–24 h. Subconfluent cells were collected into an appropriate volume of lysis buffer (10 mM Tris, pH 7.5, 150 mM NaCl, 1% Triton X-100, and 60 mM octylglucoside) containing protease inhibitors (Roche Applied Science). For phosphospecific immunoblotting, the cells were scraped into boiling sample buffer to denature endogenous phosphatases. Cell lysates were centrifuged at 12,000 g for 10 min to remove insoluble debris. Protein concentrations were analyzed using the bicinchoninic acid reagent (Pierce, Rockford, IL), and the volume required for 20 µg of protein was determined. Samples were then separated by SDS-PAGE (10 or 12% acrylamide) and transferred to nitrocellulose membranes (0.2 µm), which were stained with Ponceau S (for visualization of protein bands) and subjected to immunoblot analysis. All subsequent wash buffers contained 10 mM Tris, pH 8.0, 150 mM NaCl, and 0.05% Tween 20 and were supplemented with 4% nonfat dry milk (Carnation) for the blocking solution and 1% BSA for the antibody diluent. Primary antibodies were used at 1:500 to 1:1,000 dilution. Horseradish peroxidase-conjugated secondary antibodies [anti-mouse at 1:6,000 dilution (Pierce) and anti-rabbit at 1:5,000 dilution (BD Pharmingen, San Diego, CA)] were used to visualize bound primary antibodies with the Supersignal chemiluminescence substrate (Pierce, Rockford, IL).

Ca2+ Influx Experiments Using Confocal Laser Scanning Microscopy

Loading of fluo 3-AM for confocal microscopy. The cells were cultured on 35-mm glass-bottom dishes (Plastek Cultureware, MatTek, Ashland, MA) and then loaded with the Ca2+-sensitive fluorescent dye fluo 3-AM (Molecular Probes) at a final concentration of 13.5 µM in Tyrode-BSA solution for 45 min, as described previously (6). The loaded cells were left at 28°C to ensure complete hydrolysis of the acetoxymethyl ester groups before initiation of the experiments.

Confocal microscopy. The cells were examined with a confocal laser scanning microscope (Radiance 2000, Bio-Rad Laboratories) equipped with epifluorescence x10, x20, x40, and x60 oil objectives. The 488-nm laser line was directed to the sample via a 510-nm primary dichromic filter and attenuated with a neutral-density filter to reduce photobleaching. Laser line intensity, photometric gain, and filter attenuation were kept constant throughout the experiments.

Ca2+ measurements. Cells were laser scanned before (basal level) and 1, 5, 10, 20, 30, and 40 min after addition of 10–6 M ET-1. Intracellular Ca2+ fluorescence was measured from three-dimensional reconstructions of the cells. In all experiments, the cells were bathed with Tyrode balanced salt solution containing 5 mM HEPES, 136 mM NaCl, 2.7 mM KCl, 1 mM MgCl2, 1.9 mM CaCl2, and 5.6 mM glucose, buffered to pH 7.4 with Tris base. The osmolarity of the buffer solution was adjusted with sucrose to 310 mosmol/l.

Sustained intracellular Ca2+ levels were expressed as the mean absolute Ca2+-fluo 3 fluorescence intensity levels. Values are means ± SE for n different cells. Statistical significance was determined using Student's t-test. P < 0.05 was considered significant.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Cav-1 Expression and Localization in WT and Cav-1–/– AoSMCs

Cells isolated from the aortas of Ink4a–/–/Cav-1+/+ and Ink4a–/–/Cav-1–/– double-knockout mice were identified as SMCs by staining for {alpha}-SM actin. For simplicity, these cells were designated Cav-1+/+ and Cav-1–/– AoSMCs.

Greater than 80–90% of these Cav-1+/+ and Cav-1–/– cells exhibited positive {alpha}-SM actin staining, which was localized to actin filaments (Fig. 1). Cav-1 was localized at the level of the plasma membrane in Cav-1+/+ AoSMCs. On the other hand, Cav-1–/– cells expressed no Cav-1, as expected.


Figure 1
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  		Fig. 1. Immunolocalization of {alpha}-smooth muscle ({alpha}-SM) actin and caveolin-1 (Cav-1) in Cav-1+/+ and Cav-1–/– aortic smooth muscle cells (AoSMCs). Smooth muscle cells (SMCs) were isolated from aortas of Ink4a–/–/Cav-1+/+ and Ink4a–/–/Cav-1–/– double-knockout mice and immunostained for {alpha}-SM actin and Cav-1. Representative images are shown. Note characteristic filamentous pattern of {alpha}-SM actin staining in Cav-1+/+ AoSMCs in A and B. In C, Cav-1 was localized at the plasma membrane in Cav-1+/+ cells. In D and E, Cav-1–/– AoSMCs also showed positive {alpha}-SM actin staining. As expected, Cav-1–/– AoSMCs in F showed no Cav-1 immunostaining. Arrows in B, C, E, and F point at the plasma membrane. N, nucleus.

 
Cav-1–/– SMCs Show Increases in Cell Proliferation as Assessed via BrdU Incorporation

Previous studies have suggested that Cav-1 may function as a negative regulator of proliferation in many cell types (11, 13, 15, 20, 24, 29, 30, 40, 41). However, the role of Cav-1 in SMC proliferation remains unknown. Therefore, we assessed the proliferation rates of AoSMCs in the presence and absence of Cav-1. Cav-1+/+ and Cav-1–/– AoSMCs were maintained in SMGM supplemented with 5% FBS. After the dividing cells were incubated with BrdU, they were immunostained using a specific BrdU antibody-FITC complex and counterstained with Hoechst nuclear stain to label all nucleated cells. Significantly more BrdU-positive-stained cells were observed in fields of Cav-1–/– than Cav-1+/+ AoSMCs (Fig. 2). BrdU-positive cells were then scored and compared with the whole population of cells labeled with Hoechst nuclear stain. Quantitation revealed 15.5 ± 2.4% and 27.2 ± 3.5% BrdU-positive cells among Cav-1+/+ and Cav-1–/– cells, respectively, approaching a nearly twofold increase in proliferating cell number. These data clearly show a higher proliferation rate for Cav-1–/– AoSMCs than for Cav-1+/+ cells.


Figure 2
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  		Fig. 2. Proliferation and migration rates of Cav-1+/+ and Cav-1–/– AoSMCs. For proliferation studies, AoSMCs were incubated with 5-bromo-2'-deoxyuridine (BrdU), which incorporates into the nuclei of dividing cells (a measure of DNA synthesis). Cells were immunostained with a monoclonal anti-BrdU antibody and counterstained with Hoechst nuclear dye to label all nucleated cells. BrdU positively stained nuclei were scored and compared with the total number of nuclei (labeled with Hoechst nuclear dye). Percentage of BrdU-positive cells was used as a measure of their proliferation rate. Two representative fields of different cell density (low vs. high) are shown for each genotype. A–D: Cav-1+/+ AoSMCs labeled with Hoechst nuclear dye (A and C) exhibited few cells positively stained for BrdU (B and D). KO, Knockout. E–H: significantly more BrdU-positive cells (F and H) were detected in Cav-1–/– cells labeled with Hoechst nuclear dye (E and G). Note BrdU positivity in 15.5% of Cav-1+/+ AoSMCs and 27.2% of Cav-1–/– cells (~1.75-fold increase). *Statistically significant difference. For migration studies, confluent monolayers of Cav-1+/+ and Cav-1–/– AoSMCs were scraped, and images of the denuded area were studied immediately (time 0) and 24 and 48 h after wound induction. Number of cells in the wounded area, indicative of migrating cells, was counted for each group of cells and plotted against time. More Cav-1–/– than Cav-1+/+ AoSMCs were observed in the denuded area 24 and 48 h after wound induction, indicating a higher migration rate in Cav-1–/– cells. *P < 0.05; **P < 0.01.

 
Increases in Cav-1–/– SMC Migration Shown Using a Cell Culture Wound-Healing Assay

Migration was assessed in AoSMCs kept in basal medium supplemented with 0.1% BSA for 24 h. Briefly, the confluent monolayers were scraped with a 200-µl pipette tip to create a wound in cultures of AoSMCs. Images were acquired immediately and 24 and 48 h after wound induction.

Cell migration rates were quantitatively assessed by counting the number of cells in the denuded area at 0, 24, and 48 h after wound induction. At time 0, the scraped area was mostly devoid of cells. At as early as 6 h, the cells appeared to be migrating from the edge of the wound toward the empty space within the wound. At 24 and 48 h after wound induction, there were clearly more cells in the denuded area of Cav-1–/– than Cav-1+/+ AoSMCs: 12.3 ± 1.2 vs. 7.5 ± 0.7 at 24 h and 17.1 ± 1.3 vs. 12.3 ± 1.1 at 48 h (Fig. 2). These findings show a higher (~1.45-fold) migration rate for Cav-1–/– AoSMCs than Cav-1+/+ cells.

p42/44 MAP Kinase (ERK1/2) Is Hyperactivated in Cav-1–/– SMCs

Evidence has accumulated suggesting a regulatory relation between Cav-1 and the p42/44 MAP kinase (ERK1/2) cascade (11, 15, 20, 35). Thus we next examined the activation state of ERK1/2 in Cav-1+/+ and Cav-1–/– AoSMCs by Western blot analysis using phosphospecific antibodies.

As demonstrated in Fig. 3, phosphorylated ERK1/2 levels are dramatically increased (by ~5-fold) in Cav-1–/– AoSMCs compared with Cav-1+/+ cells. Evaluation with a phospho-independent antibody showed equivalent levels of total ERK1/2, which also serves as an internal control for equal protein loading. These results provide a potential mechanism to explain the hyperproliferative phenotype of Cav-1–/– AoSMCs.


Figure 3
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  		Fig. 3. ERK1/2 activation and cell cycle regulators in Cav-1+/+ and Cav-1–/– AoSMCs. Activation state of ERK1/2 was assessed by immunoblotting cell lysates with phosphospecific antibodies directed against activated (phospho)-ERK1/2. Phospho-ERK1/2 was upregulated in Cav-1–/– AoSMCs compared with Cav-1+/+ cells, consistent with a predisposition toward hyperproliferation. Levels of total ERK1/2 remained unchanged. Expression levels of various proliferative signal transducers were assessed by immunblotting cell lysates with specific antibodies. Cav-1–/– AoSMCs showed an upregulation of proliferating cell nuclear antigen (PCNA) and cyclin D1 and reduced levels of p27Kip1, consistent with a predisposition toward hyperproliferation. Antibodies directed against GAPDH were used as a control for equal protein loading.

 
Levels of Important Cell Cycle Regulatory Proteins, Including Cyclin D1, PCNA, and p27Kip1, Are Altered in Cav-1–/– SMCs

To mechanistically dissect the observed increases in proliferation and migration rates of Cav-1–/– AoSMCs, we next performed immunoblotting with antibodies directed against a panel of important regulatory proteins, including cyclin D1, p27Kip1, and PCNA.

Interestingly, Cav-1–/– AoSMCs showed an upregulation of the proliferative proteins cyclin D1 and PCNA compared with Cav-1+/+ cells. Conversely, levels of the cell cycle inhibitor p27Kip1 were dramatically decreased in Cav-1–/– AoSMCs (Fig. 3). Also, we evaluated the expression of GAPDH as a control for equal loading, and its levels were equivalent in Cav-1+/+ and Cav-1–/– AoSMC samples.

These data suggest that activation of cyclin D1 and inhibition of p27Kip1 may be additional mechanisms that explain the hyperproliferative phenotype of Cav-1–/– AoSMCs.

Ca2+ Response to ET-1 Is Abnormal in Cav-1–/– SMCs

In this series of experiments, we examined the effects of 10–6 M ET-1 on intracellular Ca2+ levels in AoSMCs isolated from Cav-1+/+ and Cav-1–/– mice. For this purpose, we employed the Ca2+-sensitive fluorescent probe fluo 3.

Intracellular Ca2+ levels were increased by 20 min after addition of ET-1 in Cav-1+/+ AoSMCs, and this increase was sustained for 30 and 40 min (Fig. 4; 40 min response not shown). A faster and more pronounced response to ET-1 developed in Cav-1–/– AoSMCs. Intracellular Ca2+ was elevated in Cav-1–/– cells as early as 1 min after ET-1 addition, and the response progressed to a highly significant increase at 20 and 30 min after addition of ET-1.


Figure 4
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  		Fig. 4. Ca2+ fluorescence intensity levels in Cav-1+/+ and Cav-1–/– AoSMCs before and after addition of ET-1. Representative images of Cav-1+/+ and Cav-1–/– AoSMCs before and after ET-1 addition are shown. Note scale of fluorescence intensity used to visualize intensity levels. Cells were labeled with the fluorescent Ca2+ probe fluo 3 and kept in physiological buffer (Tyrode balanced salt solution). After confocal laser scanning microscopic imaging of labeled cells at basal levels, 10–6 M ET-1 was added to the bathing buffer, and images were taken 1 and 30 min after addition of ET-1. Settings, such as laser intensity and the photon multiplier, were kept constant during all experiments. Fluorescence intensity was measured using Image J software and plotted at time 0 (control) and 1–30 min after ET-1 addition. Cav-1+/+ AoSMCs showed a significant increase in Ca2+-fluo 3 complex at 20 min. This increase was sustained at 30 and 40 min (40-min data not shown). Cav-1–/– AoSMCs showed a significant increase in Ca2+-fluo 3 fluorescence intensity as early as 1 min. This Ca2+ increase was progressive, until it reached a plateau at 20 min. The sustained Ca2+ fluorescence increase was greater in Cav-1–/– than in Cav-1+/+ AoSMCs. *P < 0.05; **P < 0.01; ***P < 0.001.

 
To identify the ET receptor subtype(s) implicated in these responses, we next applied ET antagonists to the cells: the ETA antagonist BQ-123 or the ETB antagonist BQ-788, both in the presence of 10–6 M ET-1. The Ca2+ increase induced by ET-1 in Cav-1+/+ AoSMCs was reversed by BQ-123, as expected. However, the Ca2+ response to ET-1 in Cav-1–/– cells was not altered by BQ-123 (Fig. 5). On the other hand, the ETB antagonist BQ-788 reversed the ET-1-induced Ca2+ increase in Cav-1–/– AoSMCs to control levels but did not affect the response observed in Cav-1+/+ cells (Fig. 5). These results suggest that different ET receptor subtypes are mediating ET-1 effects in Cav-1–/– vs. Cav-1+/+ cells.


Figure 5
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  		Fig. 5. Expression levels of ETA and ETB receptors in Cav-1+/+ and Cav-1–/– AoSMCs. Left: receptor antagonist studies. ETA receptor antagonist BQ-123 (10–4 M) or ETB receptor antagonist BQ-788 (10–4 M) was added to bathing buffer. Images were taken at control levels, at different times after the addition of ET-1, and after addition of receptor antagonists. Fluorescence intensity was measured using Image J software and plotted. In Cav-1+/+ AoSMCs, increase in Ca2+-fluo 3 fluorescence intensity in response to ET-1 was restored to control levels by BQ-123 but was not affected by BQ-788. Ca2+ response upon addition of ET-1 in Cav-1–/– AoSMCs was unaltered by BQ-123 but was completely abolished by BQ-788, suggesting that different receptor subtypes mediate responses to ET-1 in AoSMCs derived from Cav-1+/+ and Cav-1–/– mice. **P < 0.01; ***P < 0.001. Right: immunoblot analysis. Cell lysates were immunoblotted with specific antibodies directed against ETA or ETB receptors. Cav-1–/– AoSMCs exhibited a downregulation of the ETA receptor and increased ETB receptor expression compared with Cav-1+/+ cells. Antibodies directed against GAPDH were used as control for equal protein loading.

 
Expression of ETA and ETB Receptors in Cav+/+ and Cav-1–/– SMCs

To examine the protein expression of the ETA and ETB receptors in Cav-1+/+ and Cav-1–/– AoSMCs, we performed immunoblotting of cell lysates using specific antibodies directed against each of the receptors.

Our results show that protein levels of the ETA receptor are downregulated and those of the ETB receptor are increased in Cav-1–/– AoSMCs compared with Cav-1+/+ cells (Fig. 5). These data could explain the differential sensitivity of Cav-1+/+ and Cav-1–/– AoSMCs to distinct ET receptor antagonists (ETA vs. ETB) in our Ca2+-response experiments.


   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
We previously showed that ablation of Cav-1 gene expression in mice promotes neointimal hyperplasia in vivo, a phenomenon normally characterized by SMC migration and proliferation (24). However, it remains unknown whether these defects are cell autonomous, i.e., due to loss of Cav-1 within the SMCs themselves or loss of Cav-1 expression in other adjacent cell types in vivo. Thus, to clarify the distinct role of Cav-1 in SMCs, we have characterized the phenotypic behavior of AoSMCs isolated from Cav-1–/– and Cav-1+/+ mice with regard to their proliferation, migration, and Ca2+ response to ET-1 treatment.

First, with use of BrdU incorporation, Cav-1–/– AoSMCs were shown to have a higher proliferation rate than Cav-1+/+ cells. The role of Cav-1 as a potential tumor suppressor has been reported in many cell types (10, 41). For instance, ablation of the Cav-1 gene was shown to increase the incidence, multiplicity, and area of skin tumors induced by topical application of the carcinogen 7,12-dimethylbenzanthracene (7). In addition, a Cav-1 deficiency induced an earlier onset, a higher number, and a larger area of mammary tumors in a tumor-prone transgenic model of breast cancer, namely, MMTV-PyMT mice (48). Moreover, Cav-1 gene deletion enhanced neointimal formation, a phenomenon characterized by vascular SMC migration from the medial to the intimal layer and their proliferation (24). Peterson et al. (37) showed that Cav-1 protein expression is decreased 1) in arterial SMCs during neointimal hyperplasia (in a rabbit artery injury model) and 2) on PDGF-induced proliferation of WT vascular SMCs in culture. Interestingly, transient overexpression of Cav-1 in these WT cells inhibited their growth response to PDGF and serum (37). However, these growth-inhibitory effects may be simply due to the toxic effects of Cav-1 transient overexpression. Thus the growth-inhibitory role of Cav-1 in vascular SMCs was controversial before our present studies.

In addition to proliferation, migration of AoSMCs was assessed using a cell culture wound-healing assay. With use of this approach, Cav-1–/– AoSMCs were shown to have a higher migration potential than Cav-1+/+ cells. We cannot rule out the possibility that the increases in cell migration may be due to the increased cell proliferation. However, this possibility is highly unlikely, inasmuch as we performed our migration studies under serum-free conditions. Our laboratory previously reported such an inhibitory role for Cav-1 in the migration of mammary adenocarcinoma-derived cells. We showed that the transduction of mammary adenocarcinoma cells with a Cav-1 adenoviral vector inhibited their migration and lamellipod extension (52). Moreover, the enhanced neointimal formation observed in Cav-1–/– mice in response to blood flow interruption could be due not only to the increased proliferation of vascular SMCs, but also to the increased migration of Cav-1–/– SMCs from the medial to the intimal layer (24). Therefore, the migration of vascular SMCs is another important cellular process that is negatively regulated by Cav-1.

The contraction of SMCs is a process characterized by an increase in intracellular Ca2+ levels (38). Ca2+ then forms a complex with calmodulin and activates the myosin light-chain kinase. This activated enzyme phosphorylates myosin, which can then interact with actin filaments, thereby initiating contraction (1). In our present study, we assessed the role of a well-known activator of Ca2+-mediated signaling in AoSMCs. One of the important vasoactive factors known to induce an intracellular Ca2+ increase and, therefore, contraction of SMCs is ET-1 (9, 32, 50). Using the Ca2+ fluorescent probe fluo 3 and confocal laser scanning microscopy, we measured the intracellular Ca2+ levels of live cells before (basal levels) and after addition of ET-1. Cav-1+/+ AoSMCs exhibited increased intracellular Ca2+ levels at 20 min after treatment with ET-1. However, Cav-1–/– AoSMCs showed a faster response to ET-1, inasmuch as their Ca2+ level was increased after as little as 1 min after peptide addition. Moreover, the Ca2+ increase, which was sustained at 30 min (and even 40 min; data not shown) of ET-1 treatment in both groups of cells, was even more prominent in Cav-1–/– cells. These results indicate that a different subtype or, possibly, an increased number of ET receptors is mediating the effects of ET-1 in Cav-1–/– AoSMCs. Consistent with previous findings from other laboratories, the Ca2+ increase we observed was preferentially localized at the level of the nucleus, which acts as a buffer for cytosolic Ca2+ in SMCs (4–6).

ET-1 normally induces a biphasic increase in intracellular Ca2+ levels. Initially, on binding to its receptor, ET-1 induces a transient increase of Ca2+ levels due to the release of Ca2+ from intracellular stores. The second response phase is a sustained Ca2+ increase caused by the activation of Ca2+ channels and the influx of Ca2+ from the extracellular milieu (21, 32). However, in our cells, the transient Ca2+ response could not be detected in both groups, and the intracellular Ca2+ increase was gradual and sustained after 20 min, thereby reflecting the influx of Ca2+ via Ca2+ channels, such as voltage-dependent, receptor-operated non-voltage-dependent, and store-operated Ca2+ channels (4, 6, 18, 22).

The ET-1 peptide acts on SMCs via two types of receptors: ETA and ETB (12, 21, 32). Although the ETA receptor was considered the main receptor type on SMCs, many studies have reported that ETB receptors are present as well and can mediate ET-1-induced effects in SMCs (12, 33). To identify the subtype(s) of ET receptors implicated in the ET-1-induced response in our AoSMCs, we used specific antagonists of the ETA and ETB receptors and examined their effects on the Ca2+ increase caused by ET-1. The ETA receptor antagonist BQ-123 completely reversed the Ca2+ increase induced by ET-1 in Cav-1+/+ AoSMCs without affecting the response observed in Cav-1–/– cells. On the other hand, the ET-1-induced Ca2+ increase in Cav-1+/+ AoSMCs was not altered by the ETB receptor antagonist BQ-788, which completely abolished the response to ET-1 in Cav-1–/– cells. These results show that the ET-1-induced Ca2+ increase is mediated via the ETA receptor in Cav-1+/+ AoSMCs and via the ETB receptor in Cav-1–/– cells.

Studies have shown that the ETA receptor colocalizes with Cav-1 in caveolae microdomains (8, 34). Moreover, disruption of caveolae (using cholesterol-chelating agents) was shown to inhibit the ET-1 effect mediated via the ETA receptor, suggesting a positive regulatory role of caveolae (and Cav-1) on ETA activation (25, 51). As such, Cav-1 may induce the existence of preformed signaling complexes within caveolae, allowing for a rapid, efficient, and regulated response to ET-1. However, the ETB receptor only colocalizes with caveolae in unstimulated cells. Addition of ET-1 then displaces the ETB receptor from caveolae (49). Thus Cav-1 may only maintain the ETB receptor in caveolae in an inactive state. Therefore, we suggest that Cav-1 may inhibit ET-1 signaling via the ETB receptor but facilitate ET-1 signaling via the ETA receptor. We speculate that in the absence of Cav-1, as is the case of Cav-1–/– AoSMCs, ETA receptors could not be activated by ET-1, and ETB receptors may "take over" mediating the ET-based effects.

Moreover, our results showed that the protein levels of the ETA receptor were downregulated and that ETB receptor levels were increased in Cav-1–/– AoSMCs. This finding is consistent with several in vivo and in vitro studies that demonstrated a shift from ETA to ETB receptors in proliferating SMCs of atherosclerotic vessels of animal, as well as human, origin (2, 28, 36). Taken together, our findings suggest that, in the absence of Cav-1, SMCs depend on the ETB receptor to mediate their responses to ET-1 because of the decreased expression of the ETA receptor and the compensatory upregulation of the ETB receptor (isoform switching).

Interestingly, loss of Cav-1 in whole animal studies is atheroprotective (16). We believe that this atheroprotection is conveyed by Cav-1–/– endothelial cells, rather than Cav-1–/– AoSMCs, via a block in the endothelium-based transcytosis of modified lipoproteins (17). In fact, this is one of the reasons why we chose to isolate AoSMCs from Cav-1–/– mice to study their phenotype in isolation and to establish whether the effects we observed are cell autonomous. In a separate study, we also tested the in vivo behavior of Cav-1–/– SMCs and their role in neointimal hyperplasia (24). Our in vivo and in vitro results are consistent with the idea that loss of Cav-1 causes the hyperproliferation and migration of AoSMCs (for review see Ref. 17).

In conclusion, we have characterized AoSMCs derived from Cav-1–/– mice as well as from their Cav-1 WT counterparts, Cav-1+/+ mice. With respect to their proliferative and migration properties, our data demonstrate that Cav-1–/– AoSMCs exhibited higher proliferation and migration rates. In addition, Cav-1–/– cells show the hyperactivation of ERK1/2, upregulation of cyclin D1 and PCNA, and downregulation of p27Kip1. Moreover, by monitoring increases in intracellular Ca2+ levels, we assessed the ET-1-mediated Ca2+ response of Cav-1–/– AoSMCs. Our results show that Cav-1–/– cells exhibit a faster and more sustained Ca2+ influx in response to the vasoactive factor ET-1, functionally mediated via the ETB receptor.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by grants from the National Institutes of Health and the American Heart Association, as well as the Hirschl/Weil-Caulier Career Scientist Award (all to M. P. Lisanti). G. S. Hassan is a recipient of a postdoctoral fellowship from the Foundation of Health Research (Quebec, Canada). P. G. Frank was supported by a Scientist Development Grant from the American Heart Association and by a grant from the Elsa U. Pardee Foundation.


   FOOTNOTES
 

Address for reprint requests and other correspondence: P. G. Frank or M. P. Lisanti, Dept. of Cancer Biology, Kimmel Cancer Center, Thomas Jefferson Univ., 233 S. 10th St., BLSB (Bluemle Life Sciences Bldg.), Rm. 933, Philadelphia, PA (e-mail: philippe.frank{at}jefferson.edu or michael.lisanti{at}jefferson.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

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