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Published ahead of print on February 2, 2006, doi:10.1165/rcmb.2005-0306OC
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American Journal of Respiratory Cell and Molecular Biology. Vol. 34, pp. 695-703, 2006
© 2006 American Thoracic Society
DOI: 10.1165/rcmb.2005-0306OC

Retinoic Acid Inhibits Airway Smooth Muscle Cell Migration

Regina M. Day, Young H. Lee, Ah-Mee Park and Yuichiro J. Suzuki

Department of Pharmacology, Georgetown University Medical Center, Washington, DC; and Department of Pharmacology, Uniformed Services University of the Health Sciences, Bethesda, Maryland

Correspondence and requests for reprints should be addressed to Dr. Yuichiro J. Suzuki, Department of Pharmacology, Georgetown University Medical Center, NW403 Medical-Dental Building, 3900 Reservoir Road NW, Washington, DC 20057. E-mail: ys82{at}georgetown.edu


   Abstract
Top
 Abstract
Introduction
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 References
 
Airway remodeling in chronic asthma is characterized by increased smooth muscle mass that is associated with the reduction of the bronchial lumen as well as airway hyperresponsiveness. The development of agents that inhibit smooth muscle growth is therefore of interest for therapy to prevent asthma-associated airway remodeling. All-trans retinoic acid (ATRA) suppresses growth of vascular smooth muscle cells (SMCs) from the systemic and pulmonary circulation. The present study investigated the effects of ATRA on human bronchial (airway) SMCs. Human bronchial SMCs were found to express mRNAs for retinoic acid receptor (RAR)-{alpha}, -beta, -{gamma}, and retinoid X receptor (RXR)-{alpha}, -beta, but not RXR-{gamma}. Although ATRA was not effective in inhibiting proliferation or in inducing apoptosis in airway SMCs, we found that ATRA (0.2–2 µM) inhibited the SMC migration in response to platelet-derived growth factor (PDGF), as determined in a modified Boyden chamber assay. Both RAR and RXR agonists also blocked PDGF-induced airway SMC migration. ATRA also inhibited PDGF-induced actin reorganization associated with migration. PDGF-induced actin reorganization and migration were blocked by inhibitors of phosphatidylinositol 3 kinase (PI3K) and Akt. However, migration was blocked by inhibitors of the MEK/ERK pathway, with no effect on cytoskeletal reorganization. ATRA suppressed PDGF-induced Akt activation without influencing ERK activation. RAR was found to form protein–protein interactions with the p85 PI3K subunit. These results suggest that retinoic acid inhibits airway SMC migration through the modulation of the PI3K/Akt pathway.

Key Words: airway • migration • retinoic acid • signal transduction • smooth muscle


   Introduction
Top
 Abstract
 Introduction
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 References
 
Asthma is a chronic airway disorder characterized by airflow obstruction, airway inflammation, and persistent airway hyperreactivity (1). Although airway remodeling involves alterations of several of cell types, airway smooth muscle cells (SMCs) appear to play a critical role. The airway SMCs change from a contractile phenotype to a proliferative (or synthetic) phenotype characterized by increased growth, migration, production of extracellular matrix proteins, and secretion of a variety of chemokines, cytokines, and growth factors (2). Not surprisingly, the proliferative/synthetic phenotype also has reduced expression of contractile proteins (2, 3).

The change in airway SMC phenotype is believed to occur after exposure to growth factors, which initiate SMC proliferation, inhibit apoptosis, and cause migration of these cells. Several factors have been identified that may aid the phenotypic switch of airway SMCs, including platelet-derived growth factor (PDGF), epidermal growth factor, and basic fibroblast growth factor (2, 3), and these factors have been shown to be elevated in the plasma of patients with asthma (4, 5). Importantly, growth arrest of the proliferative/synthetic SMCs in culture restores the cells to a contractile phenotype (3). The ability of growth arrest to revert the pathologic phenotype of airway SMCs has led to a search for pharmacologic agents to inhibit airway SMC growth.

All-trans retinoic acid (ATRA) is an active metabolite of vitamin A that has been demonstrated to inhibit the growth of cancer cells (6), some types of epithelial cells (7), and vascular smooth muscles (8–10). ATRA inhibits PDGF-induced proliferation and induces apoptosis in rat and human aortic SMCs (11–13). In cultured pulmonary artery SMCs, ATRA inhibits serotonin-induced proliferation (8). In vivo studies indicate that ATRA reduces systemic and pulmonary vascular smooth muscle remodeling; both in the carotid artery balloon injury model system in rats (9), and in pulmonary hypertension induced by monocrotaline in rats (14), ATRA inhibited remodeling, primarily through the regulation of SMC growth.

The retinoic acid receptors (RAR) and retinoid X receptors (RXR) mediate the biological effects of ATRA. These receptors are members of the superfamily of steroid hormone ligand–activated transcription factors (15, 16). RAR bind ATRA as well as 9-cis retinoic acid, a naturally occurring isomer, whereas the RXR bind only 9-cis retinoic acid. When bound to their ligand, RAR–RXR heterodimers activate gene transcription by binding to specific promoter elements (16), and also affect the activities of other transcription factors, such as activator protein (AP)-1 (17). ATRA has additionally been demonstrated to directly interfere with the activation of signal transduction proteins, including extracellular signal–regulated kinases p44/p42 (ERK1/2) (18), as well as phosphatidylinositol 3 kinase (PI3K) and Akt (19). Thus, ATRA regulation of cell activities potentially occurs through both nuclear and cytoplasmic mechanisms; studies suggest that the operative mechanism in any case is cell-type–specific.

The present study examined effects of ATRA on airway SMC growth and migration. Although ATRA has little or no effect on airway smooth muscle proliferation and apoptosis, we found that ATRA is an effective inhibitor of airway SMC migration induced by PDGF. The mechanisms of ATRA actions involve its ability to inhibit PI3K/Akt-dependent reorganization of actin cytoskeleton.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 References
 
Cell Culture
Human bronchial SMCs and human pulmonary artery SMCs were purchased from Cell Applications (San Diego, CA) and maintained in SMC Growth Medium (Cell Applications) or Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% FBS, 1% penicillin/streptomycin, and 0.5% fungizone. Bovine pulmonary artery SMCs were isolated from adult bovine pulmonary artery and cultured in RPMI-1640 medium supplemented with 10% FBS, 1% penicillin/streptomycin, and 0.5% fungizone, as previously described (20). Cells at passages 2–6 were used for experiments. ATRA, 9-cis retinoic acid, 13-cis retinoic acid (Sigma-Aldrich, St. Louis, MO), 4-[E-2-(5,6,7,8-tetrahydro-5,5,8,8-tetramethyl-2-naphthalenyl)-1-propenyl]benzoic acid (TTNPB) and methoprene acid (BIOMOL, Plymouth Meeting, PA) were dissolved in DMSO for stock solutions. For working solutions, a further dilution was made using cell culture medium with no serum, so that the final concentration of DMSO did not exceed 0.02%.

Methylthiazolyldiphenyl-Tetrazolium Bromide Assay
Human bronchial SMCs were cultured in 96-well plates for 24 h in DMEM containing 10% FBS followed by 72 h of growth arrest in DMEM containing 0.1% FBS. Human bronchial SMCs were then treated with PDGF (10 ng/ml) with or without 30-min ATRA (2 µM) pretreatment, or ATRA alone, for 4 d. Medium was aspirated, and 100 µl/well of methylthiazolyldiphenyl-tetrazolium bromide (MTT, Sigma-Aldrich) solution was added (0.5 mg/ml MTT in serum free DMEM). Cells were incubated at 37°C, 5% CO2, for 4 h. MTT stain was aspirated, and 150 µl/well of DMSO was added; the plate was then agitated for 5 min before reading at 570 nm, with 595-nm reference, in a SpectraMax 340PC Microplate spectrophotometer (Molecular Devices, Sunnyvale, CA).

Measurements of Apoptosis
The neutral comet assay was used to measure double-stranded DNA breaks as an indication of apoptosis, as previously described (21). Cells were treated with apoptotic stimuli, washed in PBS, embedded in 1% agarose, and placed on a comet slide (Trevigen, Gaithersburg, MD). Cells were then placed in lysis solution (2.5 M NaCl, 1% Na-lauryl sarcosinate, 100 mM EDTA, 10 mM Tris base, 0.01% Triton X-100) for 30 min. The nuclei were subsequently electrophoresed for 20 min at 1 V/cm in 1x Tris/borate/EDTA buffer (TBE; 5x TBE stock has 250 mM Tris, 250 mM Boric acid, and 5 mM EDTA), fixed in ethanol, followed by staining with Sybr Green (Molecular Probes, Eugene, OR) and visualization with a fluorescence microscope at 478 nm excitation and 507 nm emission wavelengths. Between 100 and 150 comets were scored per experiment and assigned into type-A, -B, or -C categories based on their tail moments. Type-C comets were defined as apoptotic cells, as described by Krown and colleagues (22).

For terminal deoxynucleotidyl transferase–mediated deoxyuridine triphosphate nick-end labeling (TUNEL) assay, cells were grown on 12-mm-diameter glass coverslips in 24-well dishes for 24 h. Cells were then treated with or without testing agents for 24 h. TUNEL stain was performed by using the TACS-XL In Situ Apoptosis Detection Kit (Trevigen) on 4% paraformaldehyde–fixed cells.

To measure mitochondrial membrane potential disruption, cells were grown in 24-well dishes for 24 h, followed by treatment with or without testing reagents for 24 h. DePsipher Kit (Trevigen) was used to detect changes in mitochondrial membrane potential with a cationic dye, 5,5'6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolylcarbocyanine iodide. In accordance with the manufacturer's instruction, cells were incubated for 20 min in 1x reaction buffer, stabilizing solution, and DePsipher solution, at 37°C. The cells were examined under a fluorescence microscope (Olympus, Melville, NY) with a red/green dual filter cube.

Migration Assay
Human bronchial SMCs were trypsinized, washed, and plated at a density of 4 x 105 cells/well in 12 well/plate Transwell (Corning, Corning, NY) dishes (12-µm pore size). Cells were allowed to attach for 3 h, then treated in both upper and lower chambers pretreated with or without ATRA (2 µM) for 20 min before the addition of PDGF (1 µg/ml), or ATRA alone. After 16 h, cells attached to the Transwell insert were fixed for 15 min in 100% methanol and stained for 1 h in 0.94% crystal violet (Fisher, Fairlawn, NJ) in 20% methanol. The insert was rinsed in a large volume of H2O, and the upper side of the membrane was cleaned using a cotton swab. Remaining migrated cells were counted by microscopy.

Immunohistochemistry
For fluorescent labeling of the actin cytoskeleton, cells were grown in 35-mm dishes on glass coverslips coated with 0.015–0.15% gelatin, for 8–16 h. Cells were grown to 80% confluence and treated as described. Cells were then washed once with 1 ml warm PBS for 3 min before fixing for 5 min in 1 ml formalin (4% formaldehyde in PBS). Cells were then gently washed twice with 1 ml PBS for 3 min. Cells were permeabilized for 5 min with Triton buffer (0.1% Triton X-100, 50 mM PIPES, pH 7.0, 90 mM HEPES, pH 7.0, 0.5 mM MgCl2, 75 mM KCl, 0.5 mM EGTA) and washed three times in PBS. Rhodamine-labeled phalloidin (Molecular Probes) (1:50 in 1% BSA in PBS) was added drop-wise onto the center of the cover slip and incubated for 1 h at ambient temperature. Finally, cells were washed twice with 1 ml PBS at ambient temperature for 3 min. Cover slips were mounted in 9:1 glycerol PBS. Fluorescent proteins were visualized on a Zeiss fluorescence microscope (Carl Zeiss, Thornwood, NY) at x40 magnification.

Western Blotting
To prepare whole-cell lysates, the cells were washed in PBS and solubilized with 50 mM Hepes solution (pH 7.4) containing 1% (vol/vol) Triton X-100, 4 mM EDTA, 1 mM sodium fluoride, 0.1 mM sodium orthovanadate, 1 mM tetrasodium pyrophosphate, 2 mM PMSF, 10 µg/ml leupeptin, and 10 µg/ml aprotinin. To prepare nuclear extracts, cells were washed in PBS and incubated for 15 min at 4°C in 10 mM Hepes (pH 7.8), 10 mM KCl, 2 mM MgCl2, 4 mM EDTA, 0.1 mM PMSF, 5 µg/ml leupeptin, 5 µg/ml aprotinin, 95 mM sodium fluoride, 2.7 mM sodium orthovanadate, and 10 mM tetrasodium pyrophosphate. Nonidet P-40 was then be added at a final concentration of 0.6%, mixed vigorously, and centrifuged. Pelleted nuclei were resuspended in 50 mM Hepes (pH 7.8), 50 mM KCl, 300 mM NaCl, 0.1 mM EDTA, 0.1 mM PMSF and 10% (vol/vol) glycerol, then mixed for 30 min at 4°C, centrifuged, and the supernatant harvested.

Cell lysates or nuclear extracts (10 µg protein/sample) were electrophoresed through a reducing SDS polyacrylamide gel (10%) and electroblotted onto a nitrocellulose membrane. The membrane was blocked and incubated with the polyclonal IgG for phosphospecific ERK1/2 or phosphospecific Akt (Cell Signaling Technology, Beverly, MA) or ERK1/2 (Santa Cruz Biotechnology, Santa Cruz, CA). The levels of proteins and phosphoproteins were detected with horseradish peroxidase–linked secondary antibodies and the ECL System (Amersham Life Science, Arlington Heights, IL).

RT-PCR
PCR primer sequences for human RARs have previously been described (8). Denaturing was performed at 94°C for 45 s, annealing for 45 s at 55°C, and polymerase reactions for 2 min at 72°C. Reactions were performed using PCR SuperMix (Invitrogen, Carlsbad, CA), according to the manufacturer's instructions.

Electrophoretic Mobility Shift Assays
Cells were washed in PBS and incubated in 10 mM Hepes (pH 7.8), 10 mM KCl, 2 mM MgCl2, 0.1 mM EDTA, 0.1 mM PMSF, 5.0 µg/ml leupeptin, 5.0 µg/ml aprotinin, 1.0 mM sodium fluoride, 0.1 mM sodium orthovanadate, and 1.0 mM tetrasodium pyrophosphate for 15 min at 4°C. Nonidet P-40 was then be added at a final concentration of 10%, mixed vigorously, and centrifuged. Pelleted nuclei were resuspended in 50 mM Hepes (pH 7.8), 50 mM KCl, 300 mM NaCl, 0.1 mM EDTA, 0.1 mM PMSF and 10% (vol/vol) glycerol, then mixed for 30 min at 4°C, centrifuged, and the supernatant harvested.

For electrophoretic mobility shift assay (EMSA), the binding reactions were performed for 20 min in 5 mM Tris-HCl (pH 7.5), 37.5 mM KCl, 4% (wt/vol) Ficoll 400, 0.2 mM EDTA, 0.5 mM DTT, 1 µg poly (dI-dC)·poly(dI-dC), 0.25 ng (> 20,000 cpm) 32P-labeled double-stranded oligonucleotide, and 2 µg protein of nuclear extract. Electrophoresis of samples through a native 6% polyacrylamide gel run in 0.25x TBE buffer was followed by autoradiography. The double-stranded oligonucleotide probe containing RAR response element and RXR response element consensus elements have the sequence 5'-TCG AGG GTA GGG TTC ACC GAA AGT TCA CTC G-3' and 5'-AGC TTC AGG TCA GAG GTC AGA GAG CT-3' (Santa Cruz Biotechnology), respectively.

Adenoviral Infection
The adenovirus-directed gene transfer was implemented by adding the gene-carrying replication-deficient adenovirus (50 pfu/cell). Adenoviruses expressing the dominant negative mutant Akt (T308A,S473A) and the constitutively active mutant Akt (with c-src myristoylation sequence fused to the N-terminus) were kind gifts from Dr. Kenneth Walsh (Boston University).

Statistical Analysis
Means ± SE were calculated, and statistically significant differences between two groups were determined by the Student's t test at P < 0.05.


   RESULTS
Top
 Abstract
 Introduction
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 References
 
Effects of ATRA on Airway SMC Proliferation and Apoptosis
ATRA typically inhibits 50–90% of aortic or pulmonary SMC proliferation induced by growth factors (8, 10, 11, 13, 23). We tested the hypothesis that ATRA may also inhibit the proliferation of airway SMCs. To induce proliferation, human bronchial SMCs were treated with PDGF with or without pretreatment with ATRA. In contrast with findings from vascular SMCs, we found that ATRA reduced PDGF-induced proliferation of airway SMCs by only 5–10% (Figure 1A).


Figure 1
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  		Figure 1. Retinoic acid receptors (RAR) expression in airway SMCs and the effect of all-trans retinoic acid (ATRA) on airway smooth muscle cell (SMC) proliferation and apoptosis. (A) Human bronchial SMCs (HBSMCs) were treated with ATRA (2 µM) and then with platelet-derived growth factor (PDGF; 10 ng/ml). The number of viable cells was determined by the methylthiazolyldiphenyl-tetrazolium bromide (MTT) assay. (B) Cultured bovine (BPASMCs) or human pulmonary artery SMCs (HPASMCs) or (C) HBSMCs were treated with ATRA (2 µM) or daunorubicin (DNR; 2 µM) for 24 h. Incidence of apoptosis was determined by the neutral comet assay. The values in the bar graph represent means ± SEM of percent apoptotic cells (n = 4). * Value is significantly different from the untreated control value at P < 0.05. (D) HBSMCs were treated with ATRA or DNR, and TUNEL assay was used to measure apoptotic cells. The bar graph represents means ± SEM. (E) HBSMCs were treated with ATRA or hydrogen peroxide; mitochondrial membrane potential was monitored by DePsipher. In healthy mitochondria, the DePsipher dye aggregates and emits red fluorescence. When the membrane potential is disrupted during the early stage of apoptosis, the dye cannot cross the mitochondrial membrane and is visualized as a monomeric form, with green fluorescence in the cytosol.

 
ATRA has also been shown to induce apoptosis of vascular SMCs (12). The neutral comet assay, which determines the incidence of double-stranded DNA breaks, was used to demonstrate that ATRA treatment induced apoptosis of bovine and human pulmonary artery SMC (Figure 1B). In contrast, apoptotic cell death was not produced in response to ATRA in human bronchial airway SMC, as monitored by comet assay (Figure 1C). Similarly, TUNEL assay (Figure 1D) and the measurement of mitochondrial membrane potential (Figure 1E) failed to demonstrate significant apoptosis by ATRA in airway SMCs. Positive controls, daunorubicin (DNR) and hydrogen peroxide, effectively elicited apoptosis in airway SMC.

Effects of ATRA on Airway SMC Morphology, Actin Polymerization, and Migration
Treatment of airway SMCs with PDGF induces an alteration of airway SMC shape (Figure 2A). Cells appear to be more elongated, with multiple processes extending from the cell body. These changes in morphologic characteristics in response to PDGF are not accompanied by indications of cell death. ATRA pretreatment attenuated PDGF-induced processes from forming on the cells.


Figure 2
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  		Figure 2. ATRA inhibits PDGF-induced airway SMC migration. (A) HBSMCs were pretreated with ATRA (2 µM), then treated with PDGF (10 ng/ml), and morphologic changes were observed by phase-contrast microscopy. (B) Cells were pretreated with retinoids (2 µM) for 30 min, then treated with PDGF. Cells were stained for filamentous actin. (C) HBSMCs were split into Boyden chambers and allowed to attach for at least 2 h. Attached cells were treated with ATRA (2 µM) for 30 min before treatment with PDGF (10 ng/ml). After 18 h, cells were fixed and stained, and migrated cells were counted. Values represent means ± SEM. * Value is significantly different from the untreated control at P < 0.05. (D) Dose–response. Values are represented in percent of PDGF-induced migration without ATRA treatment. (E) Cells in Boyden chambers were pretreated with ATRA and treated with PDGF for 4 h, and migrated cells were counted. Values represent means ± SEM. (F) Cells in Boyden chambers were pretreated with actinomycin D (5 µg/ml) for 30 min before treating with ATRA and then with PDGF for 4 h. Migrated cells were counted, and percent inhibition of PDGF- induced migration by ATRA in the absence and the presence of actinomycin D (Act D) using the equation: ([–Act D] – [+Act D])/ (–Act D).

 
The actin cytoskeleton of airway SMCs is also altered in response to PDGF treatment (Figure 2B). Untreated airway SMCs contain multiple actin stress fibers, mostly oriented along the length of the cell body. With PDGF treatment, the cells appear to lose most of the actin stress fibers. These changes are consistent with cytoskeletal rearrangements that occur during migratory responses (24). ATRA treatment alone did not appear to alter the actin fibers. However, pretreatment with ATRA inhibited most of the actin rearrangement of PDGF.

Because ATRA inhibited both morphologic and actin cytoskeletal changes induced by PDGF, we characterized the effects of ATRA on PDGF-induced airway SMC migration using a modified Boyden chamber assay, which allows for determination of motility in random directions. As shown in Figure 2C, PDGF caused a 4-fold increase in migration of airway SMCs after 24 h, and ATRA blocked these events. ATRA by itself had no effect. While the therapeutic level of ATRA in human plasma could reach 1–2 µM (25), the effects on airway SMC migration were observed with ATRA concentrations as low as 0.2 µM (Figure 2D). DMSO, which is used as vehicle for ATRA and other retinoids, has no effect on PDGF-induced airway SMC migration. This does not appear to be due to the effects of ATRA on cell proliferation, as MTT assay showed that ATRA is not effective in inhibiting PDGF-induced cell proliferation; additionally, migration assay with 4 h of PDGF treatment also exhibits the ability of ATRA to inhibit migratory responses, as monitored using a modified Boyden chamber assay (Figure 2E). Thus, although ATRA is ineffective in inhibiting proliferation and inducing apoptosis of airway SMCs, ATRA is an efficient inhibitor of airway SMC migration. Furthermore, our results using actinomycin D, a general inhibitor of gene transcription (Figure 2F), showed that ATRA inhibition of SMC migration does not mediate gene transcriptional events.

ATRA Inhibits PDGF Activation of Akt, but Not MEK/ERK
To further determine the mechanism of retinoic acid actions, signal transduction pathways were examined. Two pathways, PI3K/Akt and MEK/ERK, have been shown to be involved in activation of migration by a number of growth factors (26–28). Similarly to observations by Goncharova and colleagues (29) in pulmonary vascular SMC, PDGF-induced airway SMC migration appears to be dependent on PI3K/Akt pathway, as a PI3K inhibitor, LY294002 (Figure 3A), inhibited PDGF-induced SMC migration. Similarly, an inhibitor of MEK, U0126, also blocked migration in response to PDGF (Figure 3A). Interestingly, although LY294002 is effective in inhibiting PDGF-induced actin reorganization, U0126 did not inhibit this activity (Figure 3B), suggesting that inhibition of migration by U0126 occurs via a different mechanism.


Figure 3
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  		Figure 3. Effects of PI3K/Akt and MEK/ERK pathway inhibitors on PDGF-induced migration in airway SMCs. (A) HBSMCs were pretreated with the PI3K inhibitor, LY294002 (20 µM), or the MEK inhibitor, U0126 (10 µM), then treated with PDGF (10 ng/ml) for 2 h. Cells were washed, trypsinized, and counted for seeding in migration chambers, where they were allowed to attach for 2 h before treatment with PDGF (10 ng/ml). Cells were fixed and stained, and migrated cells were counted 18 h later. Bar graphs represent means ± SEM. * Significantly different values from control at P < 0.05. (B) Cells were pretreated with LY294002 or U0126, then treated with PDGF for 2 h. Actin organization was determined by immunofluorescence analysis.

 
ATRA has been shown in carcinoma cells to inhibit the PI3K/Akt pathway (30). ATRA has also been shown to inhibit p42/p44 ERK activation in some cell types and activate ERK in others, including aortic SMCs (31, 32). Western blot experiments using phosphospecific antibodies revealed that ATRA inhibits PDGF-induced Akt activation, although cellular levels of Akt were not changed (Figure 4A). ATRA alone does not activate ERK in airway SMC and does not affect either the phosphorylation of ERK by PDGF or cellular levels of total ERK protein (Figure 4B). Smith and colleagues (18) reported that serum-induced nuclear translocation of phospho-ERK is inhibited by ATRA treatment in F9 carcinoma cells. We show that 30 min pretreatment of airway SMCs with ATRA does not inhibit nuclear translocation of PDGF-induced phospho-ERK (Figure 4C).


Figure 4
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  		Figure 4. ATRA inhibits PDGF activation of Akt, but not activation of ERK. Bronchial SMCs were grown to 80% confluence, placed in serum-free medium overnight, and then treated with PDGF (10 ng/ml), with or without pretreatment with ATRA (2 µM) for 30 min. Cell lysates were prepared, and equal amounts of protein (10 µg) were used for Western blot determination of the phosphorylation states and total protein of (A) Akt and (B) ERK. Bar graphs represent means ± SEM of percent control of intensity of the bands, as determined by densitometry (n = 3). (C) Equal amounts of nuclear extract protein (10 µg) were used for Western blots of nuclear phosphorylated ERK.

 
Consistent with our finding that Akt activation is blocked by ATRA, SMC migration was inhibited by adenovirus-mediated gene transfer of dominant negative Akt (dnAkt) (Figure 5A). Moreover, a constitutive active mutant of Akt attenuated the inhibition of migration by ATRA (Figure 5B).


Figure 5
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  		Figure 5. Effects of adenovirus expressing Akt mutants on the ATRA inhibition of PDGF-induced SMC migration. HBSMCs were infected for 24 h with (A) adenovirus expressing dominant negative Akt (dnAkt) or control adenovirus (Control Adv), or (B) with adenovirus expressing constitutively active Akt (caAkt Adv) or control adenovirus. Cells were then split into migration chambers, were allowed to attach to membranes for 2 h, and were then treated with PDGF (10 ng/ml), with or without 30-min pretreatment with ATRA (2 µM). Cell were fixed, stained, and counted 20 h after treatment. Bar graphs represent means ± SEM. * Significantly different values from control at P < 0.05. Western blot results for the expression of Akt mutants are also shown.

 
Roles of RAR
Aortic SMCs (11) and pulmonary artery SMCs (8) express RAR{alpha}, RARbeta, RAR{gamma}, RXR{alpha}, and RXRbeta, but not RXR{gamma}. Given the different response of the airway SMCs to ATRA compared with vascular SMCs, we hypothesized that airway SMCs may express a different subset of ATRA receptors, and RT-PCR was used to determine the receptor expression patterns in airway SMCs. As shown in Figure 6A, 30 cycles of PCR generated products of 201, 752, and 327 bp, which correspond to RAR{alpha}, RARbeta, and RAR{gamma}, respectively. Detection of RXR required 40 cycles of PCR, which generated 93- and 510-bp products, corresponding to RXR{alpha} and RXRbeta, respectively. However, the RXR{gamma} product with the expected size of 532 bp was not generated. We have performed PCR up to 50 cycles to confirm that RXR{gamma} is not detectable. Control experiments with no reverse transcription show that no bands were produced, confirming that our PCR products resulted from cDNA and not from contaminating genomic DNA. These results indicate that, similarly to vascular SMCs, airway SMCs express RAR{alpha}, RARbeta, RAR{gamma}, RXR{alpha}, and RXRbeta, but not RXR{gamma}.


Figure 6
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  		Figure 6. Roles of RAR agonists on PDGF-induced migration. (A) Total RNA was isolated from HBSMCs. RAR-{alpha}, -beta, -{gamma}, RXR-{alpha}, -beta, and -{gamma} expressions were monitored by RT-PCR. (B) Cells were treated with ATRA. Nuclear extracts were subjected to EMSA with RAR or RXR consensus elements. (C) HBSMCs were split into Boyden chambers and allowed to attach for at least 2 h. Attached cells were pretreated with ATRA, 4-[E-2-(5,6,7,8-tetrahydro-5, 5,8,8-tetramethyl-2-naphthalenyl)-1-propenyl]benzoic acid (TTNPB) or methoprene acid (meth acid; 2 µM) and then treated with PDGF (10 ng/ml). (D) Attached cells were pretreated with ATRA, 9-cis-retinoic acid (9cisRA), or 13-cis-retinoic acid (13cisRA) at 2 µM and then treated with PDGF (10 ng/ml). After 16–20 h, cells were fixed and stained, and migrated cells were counted. Values represent means ± SEM. * Value is significantly different from the untreated control at P < 0.05. (E) Cell lysates were immunoprecipitated with a mouse p85 IgG and GammaBind G Sepharose. RAR{alpha}–p85 interactions were assessed by Western blot using the rabbit RAR{alpha} IgG.

 
Treatment of the airway SMCs with ATRA increased DNA binding toward both RAR and RXR response elements, as demonstrated by EMSA (Figure 6B). These results are similar to those we obtained previously using human pulmonary artery SMCs (8). These results confirm that, in human bronchial SMCs, RAR and RXR are functional DNA-binding proteins.

Pharmacologic methods were used to determine the identities of specific RARs and RXRs required for the inhibition of PDGF-induced SMC migration by ATRA. TTNPB (RAR agonist) and methoprene acid (RXR agonist) both mimicked the actions of ATRA in the inhibition of PDGF-induced airway SMC migration (Figure 6C). These results indicate that the activation of either RAR or RXR is sufficient to block the migratory effects of PDGF. This suggests that RAR-RXR heterodimer activation may be involved in the mechanism of ATRA inhibition of PDGF-induced migration. In addition to ATRA, biological retinoic acids include 9-cis and 13-cis retinoic acids. Although ATRA is a natural agonist for RAR, 9-cis retinoic acid can activate both RAR and RXR. Although 13-cis retinoic acid does not directly affect these receptors, it can be isomerized to ATRA or 9-cis retinoic acid. We found that both 9-cis and 13-cis retinoic acid isomers are effective inhibitors of PDGF-induced airway SMC migration (Figure 6D). Inhibitory effects were comparable to that exerted by ATRA.

To provide evidence for the possible roles of RAR in PI3K/Akt pathway inhibition, we explored whether RAR might interact with PI3K. Immunoprecipitation/Western blot experiments revealed that, in human bronchial SMC, RAR{alpha} forms protein–protein interactions with p85 PI3K subunit (Figure 6E), suggesting the RAR-dependent modulation of the PI3K-mediated signal transduction.


   DISCUSSION
Top
 Abstract
 Introduction
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
References
 
The major finding of this study is that ATRA and other biologically active retinoids inhibit airway SMC migration in the absence of antiproliferative or apoptotic effects. ATRA inhibits PDGF activation of the PI3K/Akt pathway in airway SMC, but does not block phosphorylation or nuclear localization of p42/p44 ERK. We also show that inhibition of Akt, by the adenoviral expression of a dnAkt, can block PDGF-induced migration, and the expression of constitutively active Akt attenuates the inhibition by ATRA of PDGF-induced migration. Finally, we demonstrate that RAR{alpha} can form protein–protein interactions with the p85 PI3K subunit in human airway SMC.

The scheme in Figure 7 describes a model in which retinoids affect airway SMC migration. PDGF activation of airway SMC migration requires both MEK/ERK and PI3K/Akt pathways. Inhibition of PI3K/Akt by either pharmaceutical inhibitors or dnAkt blocks both migration and actin cytoskeletal rearrangement, but inhibition of ERK blocks migration without inhibiting actin changes. ATRA or its isomers bind a retinoid binding protein and prevent PDGF activation of Akt, possibly by blocking the upstream activation of PI3K. The identity of the ATRA binding protein directly involved in PI3K/Akt regulation in airway SMCs is still under investigation in our laboratories. Another retinoid binding protein, the cellular retinol-binding protein-I, has been shown to act as an inhibitor of PI3K by directly blocking heterodimerization of the p85 and p110 PI3K subunits in breast epithelial cells (30), which might serve as the target of ATRA in airway SMCs. ATRA has also been shown to affect the subcellular localization of PI3K and ERK1/2 in carcinoma cells (18, 33). The mechanisms by which ATRA regulates subcellular localization of signal transduction proteins is also not well defined, although an intact actin cytoskeleton appears to be required for regulation of ERK (18).


Figure 7
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  		Figure 7. Hypothetical model for the mechanisms of retinoid actions to inhibit PDGF-induced airway SMC migration.

 
Interestingly, ATRA does not result in growth arrest or apoptosis of airway SMCs, despite the fact that airway SMCs express the same profile of RARs and RXRs as vascular SMCs, which readily undergo apoptosis with ATRA treatment. Consequently, it appears that the effects of ATRA are specific for SMCs of different tissue origins. The mechanism of ATRA regulation of SMC growth has been shown to involve both nuclear (gene regulatory) and non-nuclear actions (10, 34, 35). Although we have determined that ATRA regulation of migration in airway SMCs involves the PI3K/Akt and not the ERK pathway, the signaling pathway(s), which allow airway SMCs to proliferate in the presence of ATRA, require further investigation.

Recently, work from our group demonstrated that patients with idiopathic pulmonary arterial hypertension have decreased levels of plasma retinoids (8). We also found that ATRA inhibits proliferation of pulmonary artery SMCs. Given the possibility that ATRA may negatively regulate pulmonary artery SMC mass, the decreased ATRA level in the plasma of pulmonary hypertension patients implies a potential role of this specific retinoid in pulmonary vascular remodeling. Decreased levels of plasma retinol, a precursor of ATRA, have been found in patients with chronic obstructive pulmonary disease; this disease, like asthma, often includes increased smooth muscle mass in airway remodeling (36). Further work is required to determine whether plasma levels of ATRA are involved in reduced inhibition of airway SMC migration in patients with asthma. Naturally occurring negative regulatory agents, such as ATRA, may provide novel strategies for prevention of airway remodeling. Furthermore, the results of the present study, together with those of a recent work by Fang and coworkers (37), showing that ATRA inhibited airway inflammation in asthmatic rats, suggest that ATRA might have beneficial effects in patients with asthma through multiple mechanisms.


   Acknowledgments
 
The authors thank Tufani SenGupta for excellent technical assistance.


   Footnotes
 
This work was supported in part by National Institutes of Health grants HL73929 (to R.M.D.), HL67340, and HL72844 (to Y.J.S.).

The observations presented in this manuscript are the opinions of the authors and do not reflect the views of the Uniformed Services University of the Health Sciences, the U.S. Department of Defense, or the federal government.

Originally Published in Press as DOI: 10.1165/rcmb.2005-0306OC on February 2, 2006

Conflict of Interest Statement: None of the authors has a financial relationship with a commercial entity that has an interest in the subject of this manuscript.

Received in original form August 8, 2005

Accepted in final form January 8, 2006


   References
Top
 Abstract
 Introduction
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 References
 

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