β-Catenin is an 88-kDa member of the armadillo family of proteins that is associated with the cadherin-catenin complex in the plasma membrane. This complex interacts dynamically with the actin cytoskeleton to stabilize adherens junctions, which play a central role in force transmission by smooth muscle cells. Therefore, in the present study, we hypothesized a role for β-catenin in the regulation of smooth muscle force production. β-Catenin colocalized with smooth muscle α-actin (sm-α-actin) and N-cadherin in plasma membrane fractions and coimmunoprecipitated with sm-α-actin and N-cadherin in lysates of bovine tracheal smooth muscle (BTSM) strips. Moreover, immunocytochemistry of cultured BTSM cells revealed clear and specific colocalization of sm-α-actin and β-catenin at the sites of cell-cell contact. Treatment of BTSM strips with the pharmacological β-catenin/T cell factor-4 (TCF4) inhibitor PKF115-584 (100 nM) reduced β-catenin expression in BTSM whole tissue lysates and in plasma membrane fractions and reduced maximal KCl- and methacholine-induced force production. These changes in force production were not accompanied by changes in the expression of sm-α-actin or sm-myosin heavy chain (MHC). Likewise, small interfering RNA (siRNA) knockdown of β-catenin in BTSM strips reduced β-catenin expression and attenuated maximal KCl- and methacholine-induced contractions without affecting sm-α-actin or sm-MHC expression. Conversely, pharmacological (SB-216763, LiCl) or insulin-induced inhibition of glycogen synthase kinase-3 (GSK-3) enhanced the expression of β-catenin and augmented maximal KCl- and methacholine-induced contractions. We conclude that β-catenin is a plasma membrane-associated protein in airway smooth muscle that regulates active tension development, presumably by stabilizing cell-cell contacts and thereby supporting force transmission between neighboring cells.
β-catenin is a ubiquitously expressed membrane-associated protein that is localized to the adherens junctions in quiescent cells (26). β-Catenin can play an additional role in gene transcription, as it also functions as a transcriptional coactivator of T cell factor (TCF)/lymphoid enhancer factor (LEF)-responsive genes (10). The most well-characterized role of β-catenin in cellular function is that of the transcriptional coactivator in Wnt signaling. β-Catenin can promote TCF/LEF transcriptional activity when it is stabilized intracellularly and translocated to the nucleus (10). This can occur, for example, due to the presence of growth factors or Wnt ligands that induce the accumulation of cellular and nuclear β-catenin via the inhibition of glycogen synthase kinase-3 (GSK-3)-mediated β-catenin proteasomal degradation and via parallel MAPK/ERK kinase (MEK)-dependent induction of de novo β-catenin protein synthesis. Indeed, we (18, 22, 34) have previously demonstrated a role for GSK-3 and MEK in the accumulation of nuclear β-catenin and subsequent induction of airway smooth muscle cell proliferation. These previous findings indicated the presence of the β-catenin signaling axis in airway smooth muscle and suggest an important role for this pathway in airway smooth muscle phenotype and function. Similar roles for β-catenin have also been described in smooth muscle of different origins and in striated muscles, suggesting that β-catenin regulation of gene transcription is a general mechanism for muscle cell growth (3, 9, 37, 41).
Less well-described is the functional role for β-catenin as a structural protein at the adherens junction. Direct binding of β-catenin to classic type cadherin proteins and clustering with α-catenin, α-actinin, and p120 catenin forms the so-called cadherin-catenin complex that supports homophilic cell-cell contacts at the adherens junction (26, 33). Dynamic coupling of α-actinin to the actin cytoskeleton and the cadherin-catenin complex provides structural support for the adherens junction, suggesting a potentially important role for this complex in the regulation of mechanotransduction (17, 42). The role of this complex in active tension development by smooth muscle is nonetheless not described.
Actin filaments link to the adherens junctions in smooth muscle, which are the sites of force transmission between the contractile machinery and the extracellular matrix (38). Indeed, recent findings from Gunst and Zhang (24, 43) indicate that interactions between the actin filaments and membrane-bound integrins are important for airway smooth muscle active tension development. The actin binding protein α-actinin is essential for this interaction as it binds to β1-integrins and is recruited to the adherens junction during active tension development (43). Moreover, integrin-linked kinase (ILK), which is also recruited to β1-integrins during contractile stimulation, is involved in active tension development in airway smooth muscle by supporting N-WASp-mediated actin polymerization (44, 45). During this process of actin polymerization, a submembranous actin network is formed, enhancing membrane rigidity and supporting the interactions with the actin filaments that form the contractile apparatus (24).
These findings support a role for α-actinin and the adherens junction in smooth muscle active tension development by supporting force transmission between cells and the associated extracellular matrix. In addition to cell-matrix interactions, adherens junctions are also sites for homophilic cell-cell interactions, via the cadherin-catenin complex, but the role of this complex in smooth muscle contraction has not yet been described. In the present study, we investigated the role of β-catenin as part of the cadherin-catenin complex at the plasma membrane in supporting active tension development in bovine tracheal smooth muscle (BTSM).
MATERIALS AND METHODS
Tissue preparation and organ culture procedure.
Bovine tracheae were obtained from local slaughterhouses and rapidly transported to the laboratory in Krebs-Henseleit (KH) buffer of the following composition: 117.5 mM NaCl, 5.60 mM KCl, 1.18 mM MgSO4, 2.50 mM CaCl2, 1.28 mM NaH2PO4, 25.00 mM NaHCO3, and 5.50 mM glucose, pregassed with 5% CO2-95% O2, pH 7.4. After dissection of the smooth muscle layer and careful removal of mucosa and connective tissue, tracheal smooth muscle strips were prepared while incubated in gassed KH buffer at room temperature. Care was taken to cut tissue strips at macroscopically identical length (1 cm) and width (2 mm). Tissue strips were washed once in sterile DMEM supplemented with NaHCO3 (7 mM), sodium pyruvate (1 mM), nonessential amino acid mixture (1:100), gentamicin (45 μg/ml), penicillin (100 U/ml), streptomycin (100 μg/ml), and amphotericin B (1.5 μg/ml). Next, tissue strips were transferred into suspension culture flasks, and a volume of 7.5-ml medium was added per tissue strip. Strips were maintained in culture in an incubator shaker (37°C, 55 rpm, to prevent tissue attachment and cellular outgrowth) for 3 days, as described previously (21). No load was applied during the organ culture period. Load may promote the expression of contractile proteins and maintain force production of smooth muscle in culture (1). However, using this organ culture approach, we (21) previously demonstrated force production of the BTSM strips to be maintained over an 8-day period.
Isometric tension measurements.
Tissue strips, collected from suspension culture flasks, were washed with several volumes of KH buffer pregassed with 5% CO2-95% O2, pH 7.4, at 37°C. Subsequently, strips were mounted for isometric recording (Grass force displacement transducer FT03) in 20-ml water-jacked organ baths, containing KH buffer at 37°C, continuously gassed with 5% CO2-95% O2, pH 7.4. During a 90-min equilibration period, with washouts every 30 min, resting tension was gradually adjusted to 3 g. Subsequently, muscle strips were precontracted with 20 and 40 mM isotonic KCl solutions. Following two washouts, basal smooth muscle tone was established, and passive tension was readjusted to 3 g, immediately followed by two changes of fresh KH buffer. After another equilibration period of 30 min, cumulative concentration-response curves (CRCs) were constructed to stepwise increasing concentrations of isotonic KCl (5.6–50 mM) or methacholine (1 nM to 100 μM). When maximal KCl- or methacholine-induced tension was obtained, the strips were washed several times, and residual tension was relaxed using isoprenaline (10 μM).
Alamar blue viability assay.
Tissue strips were washed with HBSS in 24-well cluster plates and incubated with HBSS containing 10% vol/vol Alamar blue solution (Biosource, Camarillo, CA). Conversion of Alamar blue into its reduced form by mitochondrial cytochromes was then assayed by fluorescence spectrophotometry and normalized to tissue wet weight.
Isolation of BTSM cells.
After the removal of epithelium, mucosa, and connective tissue, tracheal smooth muscle was chopped using a McIlwain tissue chopper three times at a setting of 500 μm and three times at a setting of 100 μm. Tissue particles were washed two times with supplemented DMEM with 0.5% FBS. Enzymatic digestion was performed in the same medium, supplemented with collagenase P (0.75 mg/ml), papain (1 mg/ml), and soybean trypsin inhibitor (1 mg/ml). During digestion, the suspension was incubated in an incubator shaker (Innova 4000) at 37°C, 55 rpm, for 20 min, followed by a 10-min period of shaking at 70 rpm. After filtration of the obtained suspension over 50-μm gauze, cells were washed three times in medium supplemented with 10% FBS. Cells were then plated in culture flasks in supplemented DMEM with 10% FBS. Cell cultures were maintained at 37°C in a humidified 5% CO2 incubator. DMEM was replaced every 2–3 days, and cells were used for experiments in passages 1–2.
siRNA preparation and treatment.
A small interfering RNA (siRNA) generation kit (Gene Therapy Systems, San Diego, CA) was used to prepare dicer-generated siRNA against the bovine β-catenin transcript. To generate bovine β-catenin siRNA, RNA was extracted from BTSM, which was reverse-transcribed to cDNA. Next, a fragment of the bovine β-catenin transcript was amplified using two specific primer sets that amplified β-catenin: 1) forward 5′-GCC GGC TAT TGT AGA AGC TG-3′, reverse 5′-GAC GCT GGG TAT CCT GAT GT-3′, yielding a 587-bp amplicon; and 2) forward 5′-CCC TGA GAC GCT AGA TGA GG-3′, reverse 5′-CCA CCA CTA GCC AGG ATG AT-3′, yielding a 663-bp amplicon. Primer sequences also included the T7 promoter sequence linker (5′-GCG TAA TAC GAC TCA CTA TAG GGA GA-target DNA-3′), which were incorporated into the DNA template PCR product to allow for in vitro transcription with the TurboScript T7 Transcription Kit (Gene Therapy Systems). Following cleanup of the PCR product (QIAquick PCR Purification Kit; Qiagen, Venlo, The Netherlands), double-stranded RNA (dsRNA) was generated using the TurboScript T7 RNA Transcription Kit and then diced into 21-bp fragments using recombinant human dicer enzyme following the manufacturer's instructions (Gene Therapy Systems). Transfection of siRNA (1 μg/ml) was performed with Lipofectamine 2000 (Invitrogen, Breda, The Netherlands). For BTSM cells and tissue, siRNA transfections occurred in DMEM without supplements for 6 h, after which, media were replaced with serum-free DMEM supplemented with antibiotics and nutrients as described above. Control transfections were performed using a nonsilencing control siRNA (Qiagen).
Isolation of membrane fractions.
BTSM strips were pulverized in liquid N2 and then lysed for 10 min on ice in homogenization buffer [50 mM Tris (pH 7.4) supplemented with 1 mM Na3VO4, 1 mM NaF, 10 μg/ml aprotinin, 10 μg/ml leupeptin, 7 μg/ml pepstatin A]. After 20 strokes in a Potter homogenizer, the homogenate was centrifuged for 5 min at 500 g. The supernatant obtained was transferred to a new tube and centrifuged for 30 min at 16,100 g. The membrane pellet was resuspended in 200-μl RIPA buffer (composition: 40 mM Tris, 150 mM NaCl, 1% vol/vol Igepal CA-630, 1% wt/vol deoxycholic acid, 1 mM NaF, 1 mM Na3VO4, 10 μg/ml aprotinin, 10 μg/ml leupeptin, 7 μg/ml pepstatin A, pH 7.4) and sonicated, and protein concentration was determined according to Bradford. Samples were then stored at −20°C until further use.
Primary antibodies directed against β-catenin, smooth muscle α-actin (sm-α-actin), and N-cadherin were conjugated to protein A-Sepharose beads immediately before use by incubating for 2 h (4°C) in 30 μl of a 50% Sepharose slurry. Beads were then washed three times with RIPA buffer, blocked with 10% BSA, and transferred to 500 μg of homogenate. Immunoprecipitates, obtained after overnight incubation at 4°C, were then washed four times with Tris-buffered saline (TBS) supplemented with 0.1% Tween 20 and stored at −20°C until further use.
Western blot analysis.
BTSM strips were pulverized in liquid N2, sonicated, and lysed in 200-μl RIPA buffer on ice. Homogenates were then cleared by centrifugation for 5 min at 16,100 g. Protein content in cleared homogenates was determined according to Bradford. Equal amounts of protein from total protein lysates were subjected to electrophoresis, transferred to nitrocellulose membranes, and analyzed for the proteins of interest using specific primary and horseradish peroxidase (HRP)-conjugated secondary antibodies. Bands were subsequently visualized on film using enhanced chemiluminescence reagents and were quantified by densitometry using TotalLab software (Nonlinear Dynamics, Newcastle, United Kingdom).
BTSM cells were plated onto Lab-Tek Chamber Slides (Nunc, Roskilde, Denmark) and grown until ∼80% confluence in supplemented DMEM with 10% FBS. Before the experiments, cells were deprived for 24 h in DMEM with 5 μg/ml insulin, 5 μg/ml transferrin, and 5 ng/ml selenium (ITS). Cells were fixed for 15 min at 4°C in cytoskeletal buffer (CB; 10 mM MES, 150 mM NaCl, 5 mM EGTA, 5 mM MgCl2, and 5 mM glucose at pH 6.1) containing 3% paraformaldehyde (PFA). Cells were then permeabilized by incubation for 5 min at 4°C in CB containing 3% PFA and 0.3% Triton X-100. For immunofluorescence microscopy, fixed cells were first blocked for 2 h at room temperature in Cyto-TBS buffer (20 mM Tris base, 154 mM NaCl, 20 mM EGTA, 20 mM MgCl2, pH 7.2) containing 1% BSA and 2% normal donkey serum. Incubation with primary antibodies occurred overnight at 4°C in Cyto-TBS containing 0.1% Tween 20 (Cyto-TBST). Incubation with FITC- or Cy5-conjugated secondary antibodies was for 2 h at room temperature in Cyto-TBST.
All data represent means ± SE from n separate experiments. The statistical significance of differences between data was determined by a two-tailed Student's t-test or one-way ANOVA, where appropriate. Differences were considered to be statistically significant when P < 0.05.
β-Catenin associates with N-cadherin and sm-α-actin at the plasma membrane.
In airway smooth muscle, the existence of the cadherin-catenin complex has not yet been described. Therefore, we first aimed to determine the expression of the mesenchymal classic N-cadherin subtype and its colocalization with β-catenin and sm-α-actin. To this aim, whole cell lysates and membrane fractions of fresh BTSM strips were prepared and analyzed for the expression of these proteins. Both in whole cell lysates and in membrane fractions, a clear signal for β-catenin, N-cadherin, and sm-α-actin could be demonstrated (Fig. 1A). Moreover, both N-cadherin and sm-α-actin associated with β-catenin, as immunoprecipitates for β-catenin, N-cadherin, and sm-α-actin contained clear β-catenin immunoreactivity, which was absent when either homogenate or primary antibody was omitted during the immunoprecipitation step (Fig. 1B). Clustering of β-catenin, N-cadherin, and sm-α-actin at the adherens junction could also be demonstrated using immunocytochemistry. Double-labeling of BTSM cells for β-catenin and sm-α-actin or for N-cadherin and sm-α-actin showed a clear overlapping pattern, which was most intense at the plasma membrane sites of cell-cell contact and absent at plasma membrane sites that were only in contact with the substrate and not with neighboring cells (Fig. 1C). Collectively, these data indicate the clustering of β-catenin at sites of cell-cell contact, where it associates with N-cadherin and sm-α-actin.
β-Catenin is required for active tension development.
We next investigated whether β-catenin was involved in active tension development. BTSM strips were cultured in the presence of PKF115-584 (10 and 100 nM), an inhibitor of β-catenin/TCF4 interactions that downregulates β-catenin expression (6, 30, 32, 40). Although at 10 nM no effects of the compound on β-catenin were observed (optical density = 86 ± 12% of control; P = 0.32), pretreatment of BTSM strips for 3 days with 100 nM PKF115-584 significantly decreased the expression of β-catenin in these strips, both in whole cell lysates (optical density = 50 ± 8% of control; P = 0.003) and in membrane fractions (optical density = 55 ± 5% of control; P = 0.003; Fig. 2A). As a result, the association of N-cadherin with sm-α-actin was significantly impaired in BTSM strips treated with PKF115-584, as immunoprecipitates for sm-α-actin contained significantly less N-cadherin after PKF115-584 treatment (Fig. 2B). Viability of these strips was not affected by the treatment, which was assayed using an Alamar blue mitochondrial conversion assay. Alamar blue conversion was corrected for muscle wet weight and was found to be similar for all three treatment protocols (viability was 116 ± 10 and 110 ± 6% of control for 10 and 100 nM PKF115-584 treatment, respectively; not significant).
Downregulation of β-catenin protein by PKF115-584 had significant effects on active tension development of BTSM strips (Fig. 2, C and D). Cumulative dose-response relationships to both KCl and methacholine were constructed using PKF115-584-pretreated BTSM strips, representing both a receptor-independent and a receptor-dependent mechanism for Ca2+ generation and contraction. Maximal responses to both agonists were significantly and similarly reduced by PKF115-584 pretreatment, although only at a concentration of 100 nM [maximal active tension development (Fmax) KCl = 56 ± 9% of control, P = 0.016; Fmax methacholine = 54 ± 9% of control, P = 0.012]. Treatment with 10 nM was ineffective, which correlates well with the observed effects on β-catenin protein regulation. A higher concentration of PKF115-584 (1.0 μM) was also tested (data not shown), which reduced β-catenin protein expression in whole tissue lysates even further (16 ± 3% of control; P < 0.001) and inhibited KCl (14 ± 6% of control; P < 0.001)- and methacholine (5 ± 3% of control; P < 0.001)-induced maximal contractions almost completely. However, at this concentration, also a significant reduction in viability of the strips was measured (42 ± 13% of control; P < 0.001).
To further confirm the role of β-catenin in regulating active tension development, a second strategy was used to downregulate β-catenin protein in BTSM strips. For these experiments, we used an siRNA approach to specifically reduce β-catenin expression. Since siRNA against the bovine β-catenin transcript is not commercially available, this was custom-generated using a dicer siRNA generation kit. For this, first the β-catenin transcript was amplified by PCR, for which two separate primer pairs were evaluated. Both primer pairs successfully yielded their respective 587- and 663-bp PCR products, and after transcription to mRNA and digestion of the dsRNA product by recombinant dicer enzyme into siRNA, both procedures successfully reduced β-catenin protein expression in BTSM cells, which was maximal 3 days after transfection (Fig. 3A). Since the relative reduction in β-catenin expression using the 663-bp amplicon was slightly better, this strategy was used for further experiments. Transfection of BTSM strips in organ culture using this β-catenin siRNA significantly reduced β-catenin protein expression at 3 days after transfection (optical density = 61 ± 8% of control; P = 0.019) and attenuated maximal KCl (77 ± 4% of control; P = 0.015)- and methacholine-induced contraction (73 ± 3% of control; P = 0.008; Fig. 3, B–D). Collectively, these data indicate that β-catenin expression is required for active tension development in BTSM.
β-Catenin downregulation does not affect contractile protein expression.
β-Catenin also regulates gene expression when it is translocated to the nucleus by acting as a transcriptional coactivator of TCF/LEF-mediated gene transcription (10). Also, in smooth muscle, we (18, 34) and others (37, 41) have previously demonstrated a role for β-catenin in regulating smooth muscle cell responses including cell growth. Therefore, we aimed to study the effects of β-catenin knockdown in the smooth muscle strips on contractile protein expression to verify that the depressed maximal responses to KCl and methacholine were not primarily due to changes in contractile protein abundance. Irrespective of the treatment used (PKF115-584 or β-catenin siRNA), the reductions in β-catenin expression and force production were not correlated to changes in sm-α-actin or sm-myosin heavy chain (MHC) expression in the smooth muscle strips (Fig. 4). This indicates that the changes in active tension development observed are not due to a reduction in smooth muscle-specific protein expression.
Inhibition of GSK-3 induces β-catenin expression and active tension development.
A major protein kinase known to regulate the expression of β-catenin is GSK-3, a serine/threonine kinase that in its active, nonphosphorylated form induces β-catenin phosphorylation, priming β-catenin for ubiquitination and proteasomal degradation (8). Thus inhibition of GSK-3 is well-known to induce the expression of β-catenin in many experimental settings, including airway smooth muscle (8, 18, 34). We aimed to perform gain of function experiments by inducing β-catenin protein expression and studying the subsequent regulation of smooth muscle force production. To this aim, three different strategies were used: we employed pharmacological GSK-3 inhibition using two GSK-3 inhibitors, SB-216763 and LiCl, that are distinct in their structure, specificity profiles and mechanism of GSK-3 inhibition (5). These inhibitors are known to induce β-catenin protein expression in airway smooth muscle (18). Also, we pretreated smooth muscle strips with insulin, a hormone known to inhibit GSK-3 by Ser9/21 phosphorylation (13). All of these treatments induced significantly the expression of β-catenin protein in BTSM strips after 3 days of treatment (Fig. 5A). The induction of β-catenin protein was most profound for insulin. As expected, insulin also induced the most profound Ser9/21 GSK-3 phosphorylation, as SB-216763 and LiCl inhibit GSK-3 by direct pharmacological inhibition. Nonetheless, LiCl did have a small effect on the abundance of this phosphoprotein.
Importantly, all three pretreatment protocols, particularly the pretreatment with insulin and SB-216763, affected smooth muscle force production (Fig. 5, B and C). LiCl had only small effects, as it augmented the KCl-induced contraction to minor, yet significant extent (Fmax = 110 ± 4% of control; P = 0.032), whereas methacholine-induced contraction was not affected significantly (Fmax= 110 ± 6% of control; P = 0.122). SB-216763 pretreatment, on the other hand, significantly induced the maximal contractile responses to both agonists (Fmax = 118 ± 7% of control, P = 0.016 for KCl; Fmax = 137 ± 9% of control, P = 0.001 for methacholine). The most profound effects, however, were observed with insulin pretreatment (Fmax = 142 ± 13% of control, P = 0.008 for KCl; Fmax = 137 ± 9% of control, P = 0.002 for methacholine).
These data indicate that loss of β-catenin protein expression reduces maximal contraction to KCl and methacholine, whereas gain of β-catenin protein expression induces maximal contraction to these agonists. Indeed, when combining all data points for all treatment protocols in this study, a strong relationship, fitted as a linear equation, existed between β-catenin abundance and maximal contraction (r2 = 0.85 for KCl and 0.85 for methacholine; Fig. 6). Collectively, these data support our hypothesis that β-catenin, as part of the cadherin-catenin complex at the plasma membrane, supports active tension development in BTSM.
Regulation of smooth muscle contraction is a key determinant of organ physiology and plays a central role in the pathophysiology of several human diseases. Excessive airway smooth muscle contraction contributes to airway narrowing in obstructive airways diseases such as asthma and COPD (2). Also, in other organ systems, for example the vasculature, smooth muscle plays a key role in determining blood pressure and in the pathophysiology of hypertension. Therefore, it is of importance to understand in detail the physiological mechanisms of smooth muscle contraction and their regulation. In the present study, we describe a novel mechanism that supports active tension development during smooth muscle contraction, involving β-catenin, as part of the cadherin-catenin complex. These findings provide new insight into the regulation of smooth muscle contraction and suggest the presence of a novel regulatory mechanism in smooth muscle that can be modulated pharmacologically.
The transcriptional role of β-catenin in cell physiology, including smooth muscle cell physiology, is well-described. β-Catenin is part of the cadherin-catenin complex at the plasma membrane and plays an important role in smooth muscle remodeling by regulating TCF/LEF-dependent gene transcription when targeted to the nucleus (8, 10, 22). Nuclear targeting of β-catenin can be regulated by its liberation from cell-cell contacts, as described for vascular smooth muscle, in response to mitogenic stimulation or in response to matrix metalloproteinase-dependent proteolytic cleavage of R- and N-cadherin (31, 35, 37, 41). This results in dissociation of the cadherin-catenin complex and subsequent induction of β-catenin-dependent gene transcription. In airway smooth muscle, dissociation of β-catenin from the plasma membrane is not induced in response to mitogen stimulation. Rather, nuclear accumulation of β-catenin in these cells appears to be regulated by de novo β-catenin protein synthesis via H-Ras and MEK, which, in parallel with a reduced GSK-3-mediated β-catenin degradation, results in the accumulation of cellular and nuclear β-catenin protein (18, 34). Accumulation of nuclear β-catenin and subsequent induction of TCF/LEF-mediated gene transcription is associated with smooth muscle cell proliferation (12, 18, 34, 37, 41) and VEGF-A secretion (11). Indeed, increased β-catenin expression by smooth muscle cells is a feature of proliferative phenotype myocytes in atherosclerotic lesions (39). Although these published findings support the functional role of β-catenin as a transcriptional coactivator in smooth muscle, the stabilizing role of β-catenin at the plasma membrane in the cadherin-catenin complex is still largely unknown. Here, we demonstrate that β-catenin is of significant importance in the regulation of active tension development during smooth muscle contraction, which reveals that β-catenin as part of the cadherin-catenin complex also plays an important physiological role in smooth muscle cell structure and function that is distinct from its transcriptional role in the nucleus. This contention is supported by our observations that smooth muscle-specific protein expression was not affected in our protocols that were aimed at reducing β-catenin protein expression using PKF115-584 and β-catenin siRNA.
The role of β-catenin in supporting smooth muscle contraction is likely explained by its stabilizing effect on the attachment of actin filaments to the adherens junctions. β-Catenin binding to N-cadherin and the association of p120 catenin, α-catenin, and α-actinin forms the so-called cadherin-catenin complex that interacts dynamically with the actin cytoskeleton and supports its association with adherens junctions (26, 33). This complex is already present in smooth muscle in the relaxed state, as all experiments shown in Fig. 1 were performed in unstimulated cells and tissues. Also, no recruitment of β-catenin to the plasma membrane could be observed after contractile stimulation with methacholine (data not shown). Since homophilic N-cadherin binding between neighboring cells provides structural support, a reduction in β-catenin content in the plasma membrane could thus limit the structural support that is necessary for tension development in the smooth muscle tissue. This contention is supported by the observation that N-cadherin, sm-α-actin, and β-catenin colocalized at the plasma membrane, coimmunoprecipitated in whole cell lysates, and colocalized at the sites of cell-cell contact. Interestingly, immunocytochemistry revealed that N-cadherin, sm-α-actin, and β-catenin also colocalized at the nucleus. A functional cadherin-catenin complex in the nuclear membrane could also contribute to the effects of β-catenin on force transmission, as actin filament binding to the nuclear envelope is required for force transmission in airway smooth muscle tissue (29).
β-Catenin binding to sm-α-actin was recently also proposed to regulate portal hypertension during the development of liver cirrhosis, suggesting a similar structurally supportive role for β-catenin in liver cells (28). Moreover, a similar role for the adherens junction was recently proposed by Gunst and Zhang (24, 43), who reported that the dynamic association of actin binding proteins to integrins at adherens junctions provides structural support and supports active tension development by providing a structural link between the actin cytoskeleton and the extracellular matrix. Collectively, it appears that actin filaments can bind to the adherens junction via multiple mechanisms (integrins, cadherin-catenin complex) and that this binding provides structural support to both the extracellular matrix and to neighboring cells that is critical during active tension development.
An interesting aspect of our studies is that our experiments demonstrate that the expression of β-catenin in smooth muscle tissue can be modulated pharmacologically. PKF115-584, a natural compound isolated from fungal origin that interferes with β-catenin/TCF4 binding, also reduced the expression of β-catenin and the association of N-cadherin with sm-α-actin, which is in accordance with earlier published studies (6, 30, 32, 40). The strong inhibitory effects of this compound on airway smooth muscle contraction suggest that inhibition of β-catenin expression may be a strategy worth pursuing in the identification of new drug targets for chronic obstructive airways diseases. Such drugs could also be effective against the remodeling associated with these diseases, as β-catenin appears to play a role in these processes as well (22). Our studies also suggest that factors that induce GSK-3 inhibition in airway smooth muscle exert the opposite effects and augment airway smooth muscle contraction. For example, our experiments using insulin demonstrate that sustained GSK-3 inhibition induces the expression of β-catenin and augments smooth muscle contraction. These studies follow up on our previous observations indicating that also PDGF, transforming growth factor-β (TGF-β), and acetylcholine modulate the GSK-3/β-catenin signaling axis (18, 20, 34), suggesting that targeting this pathway may provide significant beneficial effects in chronic airways disease. Indeed, enhanced GSK-3 phosphorylation within the airway smooth muscle bundle of allergen-challenged mice has been reported that correlated well with the changes in smooth muscle phenotype and function that were observed in these mice, including increased contractile protein expression and airway smooth muscle cell hyperplasia and hypertrophy (7). Increased GSK-3 phosphorylation could also impact on β-catenin expression, and future investigations in this area would be of interest in view of the role of this protein in the regulation of force production and proliferation of airway smooth muscle.
The role of GSK-3 in the regulation of airway smooth muscle force production can, however, be explained in more than one way. GSK-3 is a multitasking kinase that targets numerous protein kinases, transcription factors, cell cycle regulatory proteins, enzymes, and others (16). GSK-3 induces an inhibitory phosphorylation of eukaryotic initiation factor (eIF) 2B, a guanine nucleotide exchange factor that promotes translation initiation, resulting in inhibition of the translation of smooth muscle-specific proteins. In airway smooth muscle, it has been demonstrated that GSK-3 inhibition using LiCl or SB-216763 induces cell hypertrophy and the accumulation of contractile proteins via this mechanism (15). Furthermore, active GSK-3 inhibits myocardin, a transcriptional coactivator that is critical for smooth muscle-specific protein accumulation in smooth muscle (4, 16). Indeed, in addition to the induction of β-catenin expression, we observed significant increases in the expression of sm-α-actin and sm-MHC (data not shown) in the BTSM strips treated with GSK-3 inhibitors (LiCl, SB-216763) or insulin. Thus prolonged GSK-3 inhibition in smooth muscle appears to support force production via several mechanisms, including enhanced eIF2B-mediated smooth muscle-specific protein translation, myocardin-mediated smooth muscle-specific gene expression, and β-catenin-mediated stabilization of cell-cell contacts.
The induction of a hypercontractile (airway) smooth muscle phenotype by prolonged exposure to insulin is well-documented (14, 16, 23, 27, 36). Insulin activates a series of intracellular signaling cascades that explain its effects on smooth muscle phenotype, of which phosphatidylinositol 3-kinase-dependent signaling to smooth muscle-specific gene expression (via Foxo transcription factors) and protein translation (via Akt/p70S6 kinase) and RhoA-mediated smooth muscle-specific gene expression [via serum response factor (SRF) and myocardin-related transcription factor (MAL)] are the most extensively characterized (25). Insulin signaling results in increased sm-α-actin and sm-MHC expression, increased active tension development, and morphological changes associated with a hypercontractile phenotype (23, 36). Our current findings indicate that insulin also acts via the GSK-3/β-catenin signaling pathway in smooth muscle to increase active tension development.
Collectively, we demonstrate that β-catenin is associated with the cadherin-catenin complex in smooth muscle that associates with sm-α-actin at the adherens junctions. In this capacity, β-catenin regulates active tension development in smooth muscle. Pharmacological modulation of β-catenin expression and function could provide an effective means of reducing smooth muscle contraction in diseases such as asthma, in which airway hyperresponsiveness plays a pathological role.
This work was supported by a Veni grant (916.86.036) from the Dutch Organisation for Scientific Research (NWO).
No conflicts of interest, financial or otherwise, are declared by the author(s).
We thank Dr. Esther Schmitt (Novartis Pharmaceuticals) for the generous gift of PKF115-584.
- Copyright © 2010 the American Physiological Society