Airway smooth muscle (ASM) cells may contribute to asthma pathogenesis through their capacity to switch between a synthetic/proliferative and a contractile phenotype. The multimeric dystrophin-glycoprotein complex (DGC) spans the sarcolemma, linking the actin cytoskeleton and extracellular matrix. The DGC is expressed in smooth muscle tissue, but its functional role is not fully established. We tested whether contractile phenotype maturation of human ASM is associated with accumulation of DGC proteins. We compared subconfluent, serum-fed cultures and confluent cultures subjected to serum deprivation, which express a contractile phenotype. Western blotting confirmed that β-dystroglycan, β-, δ-, and ε-sarcoglycan, and dystrophin abundance increased six- to eightfold in association with smooth muscle myosin heavy chain (smMHC) and calponin accumulation during 4-day serum deprivation. Immunocytochemistry showed that the accumulation of DGC subunits was specifically localized to a subset of cells that exhibit robust staining for smMHC. Laminin competing peptide (YIGSR, 1 μM) and phosphatidylinositol 3-kinase (PI3K) inhibitors (20 μM LY-294002 or 100 nM wortmannin) abrogated the accumulation of smMHC, calponin, and DGC proteins. These studies demonstrate that the accumulation of DGC is an integral feature for phenotype maturation of human ASM cells. This provides a strong rationale for future studies investigating the role of the DGC in ASM smooth muscle physiology in health and disease.
- phosphatidylinositol 3-kinase
mature smooth muscle tissues are chiefly designed to stiffen, shorten, or relax, thereby regulating the diameter of the hollow organs they encircle (30). The contractile activity of airway smooth muscle (ASM) is a critical determinant of airway resistance in health and in diseases such as asthma and chronic obstructive pulmonary disease (COPD). Beyond their contractile activity airway myocytes can also be induced to migrate, proliferate, and secrete extracellular matrix (ECM), growth factors, cytokines, and chemokines; thus they contribute to fibroproliferative remodeling of the airways during the pathogenesis of asthma and COPD (29, 31). The multifunctional behavior of mature ASM cells stems from a capacity for reversible phenotype switching between contractile and synthetic/proliferative states (29, 51). A number of recognized molecular markers are abundant in contractile phenotype smooth muscle cells, in particular intracellular contractile apparatus- and cytoskeleton-associated proteins such as smooth muscle myosin heavy chain (smMHC), SM22, calponin, smooth muscle α-actin, and desmin (28, 29, 51). Expression of these proteins can be used as a marker for smooth muscle function in tissue biopsies taken from the airways, vasculature, or other hollow organs (7).
The transmembrane dystrophin-glycoprotein complex (DGC) is expressed when skeletal and cardiac muscle undergo differentiation (37); in mature myotubes it anchors intracellular actin networks to extracellular laminin (17, 36). The DGC is critical for providing mechanical reinforcement to the sarcolemma by damping strain arising from repeated contractions (6, 17). The DGC may also participate in intracellular mechanotransduction and with Ca2+ homeostasis (33, 35). The DGC consists of multiple subunits, including β-dystroglycan (β-DG, the core transmembrane subunit), α-dystroglycan (α-DG, an extracellular subunit that serves as a laminin receptor), dystrophin (a rodlike linker between actin and β-DG), a tetrameric transmembrane sarcoglycan complex (SGC) consisting of α-, β-, δ-, and γ-sarcoglycan (SG), and the SGC-stabilizing protein sarcospan (5, 25, 42, 65, 69). The absence of dystrophin leads to DGC disruption, rendering the sarcolemma susceptible to damage during muscle contraction, which ultimately leads to muscle wasting that characterizes Duchenne muscular dystrophy (46). Similarly, mutations in sarcoglycans are associated with limb-girdle muscular dystrophy (9, 10), and their absence in genetically altered mice can lead to severe myopathies (13, 52, 59). Thus DGC components are essential to support mechanical activity of differentiated striated muscle.
Despite its recognized role in striated muscle, there is only marginal appreciation of the expression profile and role of DGC subunits in smooth muscle. Vascular smooth muscles reportedly express α- and β-DG, dystrophin, and a complement of sarcoglycans (β-, δ-, ε-, and γ- or ζ-subtypes) that is unique from that of striated muscle (2, 5, 65, 69). Recent studies suggest this profile may be consistent in most smooth muscle tissues (2). Mardini et al. (45) reported that age-related loss of ASM mass occurs to a greater extent in dystrophic hamsters, and this correlates with reduced contractile agonist-induced force generation. In contractile smooth muscles dystrophin compartmentalizes to caveolae-rich linear arrays that associate with Ca2+ handling microdomains in the plasmalemma (14, 20, 30, 50). These data suggest an important role for the DGC in the function of contractile smooth muscle cells; however, to date there has been little focus on the expression or role of this protein complex in ASM.
In the present study we characterized the profile of DGC subunits expressed in intact human ASM tissue and cultured myocytes. We also tested whether, in a manner akin to striated muscle differentiation, the expression of DGC proteins is associated with, and dependent upon, phenotype switching between contractile and proliferative states in human ASM cells. We investigated bronchial smooth muscle tissue obtained from human subjects, human bronchial smooth muscle cell lines, and primary cultured human tracheal smooth muscle cells obtained from healthy transplant donors. To characterize DGC subunit expression we used reverse transcriptase (RT)-PCR, Western blotting, and fluorescent microscopy to assess cell and tissue distribution. To assess the relationship between DGC expression and myocyte phenotype we used established prolonged serum-free culture protocols to induce ASM phenotype maturation (27, 67). To directly assess whether DGC expression occurs as a consequence of myocyte maturation we compared DGC expression in response to phosphatidylinositol 3-kinase (PI3K) inhibition or blockade of the binding between laminin and integrins, two mechanisms that are required for myocyte maturation (27, 67). Our studies provide the first assessment of the DGC subunit profile in human ASM and the mechanism regulating their expression in association with phenotype plasticity.
MATERIALS AND METHODS
Reagents and antibodies.
Horseradish peroxidase (HRP)-conjugated goat anti-mouse IgG, HRP-conjugated goat anti-rabbit IgG, and mouse monoclonal antibodies for dystrophin, smMHC, and calponin were obtained from Sigma (St. Louis, MO). Mouse anti-β-DG, -β-SG, and -δ-SG antibodies were obtained from Novocastra Laboratories (Newcastle, UK). Goat anti-ε-SG and rabbit anti-caveolin-1 antibody were obtained from Santa Cruz Biotechnology (Santa Cruz, CA). FITC-, Cy3-, and Texas red-conjugated secondary antibodies were obtained from Jackson ImmunoResearch (West Grove, PA). Rabbit anti-phospho-Akt1 (Thr308) and anti-Akt1 antibody were obtained from Cell Signaling Technology (Beverly, MA). All other chemicals were of analytical grade.
Immortalized human airway smooth muscle cell culture.
For all studies at least four senescence-resistant human ASM cell lines were used; these cell lines were prepared by using Moloney murine leukemia virus (MMLV) retroviral vectors to facilitate stable integration of the human telomerase reverse transcriptase gene (hTERT) as we described previously (21). hTERT-expressing human ASM cell lines retain expression of contractile phenotype markers, including smMHC, calponin, smooth muscle α-actin, and desmin to passage 10 and higher (21). For all experiments, passage 10–17 hTERT ASM cultures were used. The primary cultured human ASM cells used to generate each cell line were prepared as we described previously (49, 54) from macroscopically healthy segments of second- to fourth-generation main bronchus obtained after lung resection surgery from patients with a diagnosis of adenocarcinoma. For some experiments we used primary cultured human tracheal smooth muscle cells that were prepared from healthy transplant donors; after microdissection to isolate the trachealis muscle, myocytes were isolated with procedures mimicking those used to prepare bronchial ASM primary cultures (49, 54). All procedures were approved by the Human Research Ethics Board (University of Manitoba) and all donors gave informed consent. All cells were cultured in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum (FBS); medium was changed every 48 h unless otherwise specified. To induce acquisition of a contractile phenotype, confluent cultures were maintained in DMEM supplemented with insulin-transferrin-selenium (ITS; 5 μg/ml insulin, 5 μg/ml transferrin, 5 ng/ml selenium) for up to 4 days as we described previously (26).
Preparation of protein lysates from human ASM tissue and cells.
Intact ASM tissue was isolated from human bronchial specimens by microdissection at 4°C. Thereafter, tissues were homogenized in ice-cold RIPA buffer (composition: 40 mM Tris, 150 mM NaCl, 1% Igepal CA-630, 1% deoxycholic acid, 1 mM NaF, 5 mM β-glycerophosphate, 1 mM Na3VO4, 10 μg/ml aprotinin, 10 μg/ml leupeptin, 7 μg/ml pepstatin A, 1 mM PMSF, pH 8.0) with a Polytron. The lysate was transferred to a 1.5-ml plastic tube and centrifuged (760 g, 5 min), and the supernatant was stored at −20°C for subsequent protein assay and immunoblot analyses.
Protein lysates were collected from human ASM cultures at three stages: serum fed, 50–70% confluence (proliferative stage); serum fed, 90–100% confluence (day 0); and 4 days (day 4) after switching the medium of confluent (day 0) cultures to serum-free DMEM supplemented with ITS. To prevent phenotype maturation, in some experiments during serum-free culture cells were treated with PI3K inhibitor (20 μM LY-294002 or 100 nM wortmannin) or with laminin competing peptide YIGSR (1 μM) or an inactive scrambled peptide, GRADSP (1 μM). For protein lysate preparation plates were washed twice with ice-cold PBS, and cells were homogenized by scraping in ice-cold RIPA buffer. Lysates were transferred to 1.5-ml plastic tubes and centrifuged (760 g, 5 min), and the supernatants were stored at −20°C for subsequent protein assay and immunoblotting.
Protein content in supernatant samples was determined with the Bio-Rad protein assay and bovine serum albumin (BSA) as a reference (Bio-Rad, Hercules, CA). Immunoblotting was performed with standard techniques (26). Briefly, after samples were reconstituted in denaturing buffer, 18–25 μg of protein was loaded per lane and size separated electrophoretically under reducing conditions with SDS-polyacrylamide gels. Thereafter, proteins were electroblotted onto nitrocellulose membranes, which were subsequently blocked with 5% (wt/vol) skim milk in Tris-buffered saline (TBS) (composition: 10 mM Tris·HCl, pH 8.0, 150 mM NaCl) with (0.2%) or without Tween 20. Blocked membranes were incubated with primary antibodies diluted in TBS containing 1% (wt/vol) skim milk with (0.2%) or without Tween 20. The membranes were developed by subsequent incubation with HRP-conjugated secondary antibody and then visualized on photographic film with enhanced chemiluminescence reagents (Amersham, Little Chalfont, UK). β-Actin was used to correct for equal loading of all samples. Densitometry and quantification of the relative protein abundance were performed with the Epson Perfection 4180 Station and TotalLab TL100 software (Nonlinear Dynamics, Durham, NC).
RNA isolation and RT-PCR.
Total RNA was extracted with the Qiagen RNeasy Mini Kit in accordance with the manufacturer's recommendations (Qiagen, Mississauga, ON, Canada) from human bronchial tissue enriched in ASM that was microdissected from two different human donors. Total RNA (1 μg) was reverse transcribed with MMLV reverse transcriptase (Promega, Madison, WI) for 2 h at 37°C followed by 5-min incubation at 95°C and then diluted 1:10 with RNase-free water. The RT-PCR reactions for cDNAs of interest were carried out in a thermal cycler (Mastercycler, Eppendorf, Germany) with primer pairs listed in Table 1. The coding regions corresponding to the primers were taken from the National Center for Biotechnology Information (NCBI), and then primers were designed with PRIMER-3 and IDT programs available online. Cycle parameters were denaturation (94°C for 45 s), annealing (60°C for 45 s), and extension (72°C for 45 s). The initial denaturation period was 4 min, and the final extension was 5 min. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was amplified as an internal control. Amplified products were analyzed by DNA gel electrophoresis in 2% agarose and visualized by Gelstar staining under ultraviolet illumination with a gel documentation system (AlphaEaseFC, Alpha Innotech, San Leandro, CA).
Segments of second- to-fourth-generation main stem bronchi from healthy portions of human lung removed during lung resection surgery were frozen fresh in optimum cutting temperature (OCT) embedding medium. Tissue was cryosectioned serially into thin slices (5 μm), transferred onto precleaned microscope slide, and then air dried briefly and equilibrated in cold cytoskeletal buffer (CB) (mM: 10 MES, 150 NaCl, 5 EGTA, 5 MgCl2, and 5 glucose at pH 6.1). Sections were fixed for 15 min in cold 3% paraformaldehyde and then permeabilized with 0.3% Triton X-100 in CB for 40 min. Sections were blocked for 2 h with 3% donkey serum in cyto-TBS (mM: 20 Tris base, 154 NaCl, 20 EGTA, and 20 MgCl2 at pH 7.2) containing 0.1% Tween 20 (cyto-TBST) and then incubated at 4°C overnight in primary antibody diluted in cyto-TBST (0.1% Tween 20): anti-smMHC (1:200), anti-β-DG (1:50), anti-β-SG (1:50), or anti-dystrophin (1:50) antibody. For negative controls, samples were incubated with isotype-matched mouse IgG. Cy3-conjugated secondary antibodies were used to detect primary antibody bound to tissues. Nuclei were stained with Hoechst 33342 (10 μg/ml). After staining coverslips were mounted with ProLong antifade medium (Molecular Probes), and fluorescent imaging was performed with an Olympus BX-51 fluorescent microscope (Olympus America, Melville, NY).
Human ASM cells were plated onto precleared glass coverslips in six-well culture dishes. Cells were fixed for 15 min at 4°C in CB 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 containing 1% BSA and 2% normal donkey serum. Incubation with primary antibodies occurred overnight at 4°C in cyto-TBST with anti-smMHC (1:200), anti-β-DG (1:50), anti-β-SG (1:50), or anti-dystrophin antibody (1:50). For negative controls, samples were incubated with either isotype-matched mouse IgG or rabbit antiserum. Incubation with FITC- or Texas red-conjugated secondary antibodies was for 2 h at room temperature in cyto-TBST. Coverslips were mounted with ProLong antifade medium (Molecular Probes). Fluorescent imaging was performed by capturing a midcell section of 0.3-μm focal depth with an Olympus LX-70 FluoView confocal laser scanning microscope (Olympus America) equipped with a ×40 objective.
Values reported for all data represent means ± SE. For all studies, replicate data from three or four different cell lines were obtained. The statistical significance of differences between two means was determined by an unpaired two-tailed Student's t-test or, when appropriate, by one-way ANOVA with Bonferroni's multiple comparison test for comparison between treatments. Differences were considered to be statistically significant when P < 0.05.
Characterization of dystrophin-glycoprotein complex subunits in human ASM tissue.
We first addressed the question of whether DGC subunits are present in human ASM tissue by assessing protein abundance with Western blotting. Protein lysates were collected from human bronchial smooth muscle tissue dissected from intact bronchial specimens. All lysates were characterized by abundant smMHC, a stringent marker for contractile smooth muscle tissue (Fig. 1A). In addition, samples were probed for several DGC subunits, including β-DG, β-SG, δ-SG, ε-SG, and dystrophin; each of the DGC protein subunits was easily detected in all human ASM tissues analyzed (Fig. 1A). Because reliable antibodies were not available for all DGC subunits, we complemented our immunoblot studies with RT-PCR to profile mRNA for DGC subunits in human ASM tissue. Consistent with our immunoblot analyses, RT-PCR confirmed the presence of transcripts for β-DG, β-SG, δ-SG, ε-SG, and dystrophin but also demonstrated the presence of mRNA for α-DG, α-SG, γ-SG, ζ-SG, sarcospan, and utrophin (Fig. 1B). RT-PCR also confirmed the presence of mRNA for calponin, a contractile apparatus-associated smooth muscle marker, and caveolin-1, which we showed previously is abundant in contractile ASM cells (21), where it is thought to be associated with the DGC complex and involved in excitation-contraction coupling (30, 32). To further characterize DGC expression in human ASM tissue, we performed immunohistochemistry of bronchi obtained from human donors (Fig. 2). Consistent with our immunoblot and RT-PCR analyses, marked labeling of β-DG, β-SG, and dystrophin was seen in the ASM tissue layer, which was clearly identifiable from smMHC staining. Collectively, our experiments demonstrate that DGC subunits are abundant in human ASM tissue.
Phenotype-dependent expression of dystrophin-glycoprotein complex in cultured human ASM cells.
Smooth muscle cells retain the capacity for reversible phenotype switching, which enables myocytes to exhibit contractile and synthetic/proliferative phenotypes in vitro and in vivo. Because our initial experiments exploring DGC expression were conducted in intact, contractile ASM tissues, we next used primary cultured and immortalized human ASM cells to investigate whether the expression of DGC subunits is phenotype dependent. Myocytes were analyzed under conditions that promoted a proliferative phenotype (serum-fed, subconfluent cultures), or a contractile phenotype (4-day serum-free, confluent cultures) (Fig. 3). Western blot analysis showed that DGC protein subunits, including β-DG, β-SG, δ-SG, ε-SG, and dystrophin, were markedly increased after 4-day culture in serum-free conditions, which promotes phenotype maturation, as indicated by the concomitant accumulation of calponin, smMHC, and caveolin-1. Notably, densitometry confirmed that contractile phenotype marker proteins (calponin, caveolin-1, and smMHC) increased approximately two to three times in serum-deficient conditions, and we observed concomitant multifold accumulation of DGC subunits (Fig. 3B). For example, we observed more than a sevenfold increase in β-DG (P < 0.01) after 4-day serum deprivation compared with serum-fed proliferative phenotype cultures. The magnitude of this increase was mimicked by δ-SG (P < 0.05), ε-SG (P < 0.05), and dystrophin (P < 0.01), and the level of β-SG was also increased by more than fivefold (P < 0.05) in cultures induced to a contractile phenotype. These results indicate that the abundance of DGC components is variable and correlates with the dynamics of ASM cell phenotype expression in vitro.
Effects of maturation on human ASM cell morphology and phenotype.
In canine and human ASM cells subjected to prolonged serum starvation, phenotype maturation occurs in a select subset of myocytes that become characteristically elongate, reacquire responsiveness to contractile agonists, and accumulate abundant contractile marker proteins such as smMHC, calponin, and desmin (26, 27, 66). Thus we assessed whether accumulation of DGC subunits induced by serum deprivation was directly associated with the specific subset of human airway myocytes that undergo phenotype maturation. Using fluorescence immunocytochemistry, after 4-day serum deprivation we double-labeled primary cultured human tracheal smooth muscle cells with smMHC and β-DG, β-SG, or dystrophin (Fig. 4). Consistent with previous reports, myocytes exhibited phenotype heterogeneity, with 15–20% of ASM cells acquiring a contractile phenotype, as evidenced by a dramatic accumulation of smMHC. Notably, the maturation of individual human ASM cells to a contractile phenotype was uniquely associated with a dramatic increase in staining for DGC subunits, including β-DG, β-SG, and dystrophin, whereas little or no labeling for DGC proteins was evident in cells devoid of smMHC (Fig. 4, D–F). In contrast to serum-deprived cultures, staining of very low intensity for smMHC, β-DG, β-SG, and dystrophin was observed for all myocytes in serum-fed, subconfluent conditions (not shown). These results demonstrate, at the single-cell level, an association between the acquisition of a contractile phenotype and expression of DGC subunits.
Effect of PI3K inhibitors on human ASM cell maturation and DGC expression.
We next investigated whether accumulation of DGC proteins is merely coincident with ASM phenotype maturation in response to serum deprivation or represents an integral component of the maturation process, making it a reliable marker for active acquisition of the contractile phenotype. Previous studies have demonstrated that signaling through the PI3K pathway, including Akt1, p70S6 kinase, and mammalian target of rapamycin (mTOR), is required for ASM maturation, hypertrophy, and concomitant accumulation of contractile protein markers (27, 71). Thus we performed experiments in which we blocked maturation of human hTERT ASM cells by treating serum-deprived cultures with pharmacological inhibitors of PI3K: LY-294002 and wortmannin. To confirm that the addition of PI3K inhibitors to the culture medium blocked downstream signaling we first assayed phosphorylation of Thr308 on Akt1 before (day 0) and after (day 4) serum deprivation (Fig. 5A). Consistent with previous studies, 4 days of serum deprivation in ITS-containing medium resulted in elevated PI3K signaling, as revealed by a sevenfold increase in phospho-Thr308 Akt-1 (P < 0.001). The increase in Akt1 phosphorylation was reduced by nearly 60% in cultures treated with 20 μM LY-294002 (P < 0.01) and was virtually abolished by treatment with 100 nM wortmannin (P < 0.001) (Fig. 5B). Of note, and consistent with previous studies, accumulation of the contractile phenotype markers calponin and smMHC was prevented by PI3K inhibition with LY-294002 (P < 0.05) or wortmannin (P < 0.001) (Fig. 5, C and D). Having confirmed that PI3K signaling is essential for hTERT ASM cell maturation in vitro, we next assessed the effects of PI3K inhibition on the accumulation of DGC subunits during serum-free culture. Importantly, whereas β-DG was increased by more than fourfold (P < 0.001) after 4-day serum deprivation, this was virtually abolished by LY-294002 (P < 0.001) or wortmannin (P < 0.001) (Fig. 5E). Similarly, the accumulation of β-SG and dystrophin that we saw associated with phenotype maturation of human ASM cell lines (Fig. 3) was also suppressed significantly by inhibition of PI3K (Fig. 5, F and G). Together, these results demonstrate that inhibition of PI3K signaling, which is essential for contractile phenotype expression by human ASM cells, is also required for the accumulation of DGC proteins.
Laminin competing peptide inhibits human ASM cell maturation and DGC expression.
We recently showed (66, 67), using the laminin competing peptide YIGSR, that binding of endogenously expressed laminin to α7-integrin subunits is required for maturation of primary cultured and human ASM cell lines. For these studies we used peptides based on the integrin-binding motif of laminin β1-chains to effectively inhibit phenotypic maturation of ASM cells in vitro (67). In the present study we determined whether blocking laminin-mediated myocyte maturation with YIGSR also affected DGC protein accumulation during 4 days of serum-free culture. Consistent with our previous findings, treatment of hTERT ASM cells with YIGSR (1 μM) prevented the accumulation of contractile phenotype marker proteins calponin (P < 0.01) and smMHC (P < 0.001) (Fig. 6, A–C). Similarly, YIGSR treatment prevented the accumulation of β-DG, β-SG, and dystrophin (Fig. 6, D–F). As a negative control some cultures were treated with the inactive peptide GRADSP, which was without effect on induction of smMHC, calponin, β-DG, β-SG, and dystrophin (Fig. 6). These results demonstrate that the accumulation of DGC subunits in human ASM cells is directly associated with phenotype maturation. Moreover, expression of DGC proteins appears to be regulated by mechanisms that are essential determinants of the contractile phenotype.
This study was undertaken to profile the proteins that comprise the DGC in human ASM cells, and to determine whether their expression correlates with phenotype switching. Our focus on the DGC stems from reports showing that expression of its subunits is developmentally regulated and required for skeletal muscle maturation and maintenance (1), and that animal models lacking sarcoglycan subunits exhibit altered arterial function that may be associated with changes in Ca2+ homeostasis (12, 43). Our study is the first to provide systematic profiling of the DGC subunits expressed in human ASM tissue and cultured cells and to show that the composition of the DGC is consistent with reports for other smooth muscles, because it includes α- and β-DG, dystrophin, sarcospan, and a sarcoglycan complex that includes β-, δ-, ε-, γ-, ζ-, and, perhaps, α-SG. Our studies using cultured human ASM reveal that the expression of DGC protein subunits is dynamic, being lost or markedly reduced on modulation to a proliferative phenotype, while they accumulate when individual cells reacquire a contractile phenotype. The binding of laminin-2 to α7-integrin is required for ASM phenotype maturation (66, 67); our study revealed that blocking laminin-integrin binding prevented both phenotype maturation and the accumulation of DGC proteins, confirming that they are reliable markers for acquisition of the contractile phenotype in ASM cells. We also show for the first time that PI3K activity, which is critical for ASM maturation (27), is required for DGC protein accumulation. Collectively our results suggest that DGC protein expression is dynamically regulated by mechanisms that control ASM maturation. Moreover, we show that the DGC is abundant in contractile ASM, which suggests it could be associated with functional aspects of contraction in a manner similar to that seen for skeletal muscle.
DGC protein expression has been studied in vascular and visceral smooth muscle tissues, where the complex appears to include dystrophin, α- and β-DG, sarcospan, and sarcoglycans, including β-, δ-, and ε-SG (2, 5, 65, 69). There has been some debate about whether γ- or ζ-SG is the final member of the tetrameric sarcoglycan complex; however, a recent report indicates that they are functionally indistinguishable, indicating that the distinction may be moot (62). Our studies suggest a DGC composition for human ASM that is similar to other smooth muscles, with a core consisting of an α- and β-DG that is associated with β-, δ-, and ε-SG. We demonstrate that mRNA for both γ- and ζ-SG is present in ASM, as well as for α-SG, a subunit that has been generally accepted to be skeletal muscle specific (65). However, our detection of α-SG is consistent with some recent immunohistochemical surveys of normal smooth muscle tissues, including ASM, that do suggest the presence of low levels of α-SG protein (2, 3). We also observed mRNA for sarcospan, which is thought to stabilize sarcoglycan tetramers by forming a sarcospan-sarcoglycan complex within the DGC. Our RT-PCR analyses also indicate that utrophin, the autosomal homolog of dystrophin thought to support cellular architecture rather than a stress-bearing role like dystrophin, is expressed abundantly in ASM tissue. This is consistent with other reports indicating that both utrophin and dystrophin are expressed in vascular smooth muscle (55). A number of other proteins, such as the dystrobrevins, syntrophins, and α-actinin-associated LIM protein, are linked with the DGC and its function in skeletal muscle (36, 42, 53); our studies did not evaluate the expression of these ancillary subunits in ASM, although future studies may be warranted. Collectively our study provides the first systematic assessment of DGC protein expression in human ASM and supports a model in which α- and β-DG may be linked to the actin cytoskeleton by dystrophin or utrophin and are associated with a sarcospan-stabilized, tetrameric sarcoglycan subcomplex that may exist in multiple heteromeric forms.
There are numerous reports linking expression of the DGC with skeletal muscle differentiation and myotube survival (11, 47, 68). Nonetheless, before our present study, the association of the DGC with human ASM phenotype expression had not been investigated. We show here, and have described previously, that cell culture provides a useful tool for assessing mechanisms regulating phenotype plasticity of ASM cells, because they spontaneously modulate to a proliferative state in subconfluent, serum-fed conditions but can be induced to a mature, fully contractile phenotype by prolonged culture in insulin-supplemented serum-deficient conditions (26, 27, 61). Importantly, on the basis of our Western blot analyses of serum-deprived human ASM cells, it appears that contractile ASM cells express a repertoire of DGC proteins that mirrors ASM tissue. Using fluorescence immunocytochemistry, we saw that β-DG, β-SG, and dystrophin accumulated exclusively in myocytes from serum-deprived cultures that acquired an elongate morphology marked by abundant smMHC. These observations provide first-time, compelling evidence on a cell-by-cell basis for a direct association between the expression of DGC proteins and the acquisition of a contractile phenotype by human ASM cells.
We showed previously (66, 67) that α7-integrin, which like α-DG is a receptor for extracellular laminin, is preferentially expressed by contractile phenotype ASM cells. This parallels studies in skeletal muscle showing that expression of both α7-integrin and DGC proteins correlates with myogenesis (11). The association of laminin-binding receptors with ASM maturation is fully consistent with previous work showing that endogenously expressed extracellular laminin promotes both the differentiation of ASM in the developing lung and the expression of a functional contractile phenotype by mature myocytes (57, 67). In the present study we confirmed that blockade of ASM phenotype maturation with a laminin competing peptide, YIGSR, was sufficient to prohibit DGC protein accumulation. The YIGSR peptide corresponds to amino acids 929-933 of the β1 laminin chain (23), and it has been used frequently as an inhibitor of laminin binding in vitro and in vivo (24, 39, 60, 67). Laminin is a trimer of α-, β-, and γ-polypeptide chains that possess a number of binding sites for integrins (19, 63, 70) and non-integrin receptor subtypes, including α-DG (15, 17, 56). Because YIGSR mimics a motif in laminin β1 that selectively binds to integrins, our studies do not address the extent to which laminin binding to α-DG might directly regulate phenotype maturation of ASM cells. Interestingly, cross talk and linkages between integrins and the DGC have been reported, and these appear to modulate downstream signaling (18). There is also an emerging body of evidence that the DGC, in particular the cytoplasmic tail of β-DG, acts as a scaffold that contributes to signal transduction (42, 64). This suggests that α-DG and the DGC could be a functional determinant in ASM phenotype switching. Furthermore, our findings suggest that the DGC has potential to be functionally important during lung development and in other scenarios involving ASM phenotype maturation and growth, such as in asthma, which is marked by the accumulation of contractile phenotype ASM. Future studies that investigate these possibilities will be enlightening.
We (27) and others (71) have reported that PI3K-mediated signal transduction is critical for contractile phenotype maturation and for myocyte elongation and hypertrophy of ASM cells. Similarly, in skeletal muscle, myotube hypertrophy and the accumulation of contractile proteins requires PI3K/Akt1/mTOR and PI3K/Akt1/GSK3 signal transduction pathways (8, 58). Notably, PI3K signaling has been linked with laminin binding to the DGC and integrin subunits in skeletal and smooth muscle, respectively (34, 41). Because our experiments with YIGSR peptide showed that laminin is required for myocyte maturation and DGC expression, we used two different chemical inhibitors of PI3K activity, LY-294002 and wortmannin, to test whether PI3K activity also modulates DGC expression. We observed that PI3K inhibition concomitantly prevented ASM contractile phenotype maturation and the accumulation of the DGC proteins dystrophin, β-DG, and β-SG. To the best of our knowledge, these observations are the first to show that PI3K signaling regulates expression of DGC proteins. PI3K signaling cascades are complex and involve multiple downstream effectors that regulate proliferation, protein synthesis, cell survival, differentiation, and gene transcription (27, 34, 58). Although in the context of our present studies it is not possible to discern whether PI3K mediates its effects directly or indirectly on DGC expression through transcription or protein translation-associated mechanisms, our studies do indicate that careful dissection of these mechanisms is warranted.
Because our data show that DGC proteins are abundant in intact ASM tissue and individual cultured myocytes of a contractile phenotype, a functional and/or structural role in differentiated myocytes is implicated. In patients with muscular dystrophy, defects in gastrointestinal smooth muscle function have been postulated to underpin frequent dysphasia, vomiting, chronic constipation, and acute digestive dilatations (4, 40). In dystrophic hamsters, DGC deficiency correlates with more pronounced loss of ASM mass and contractile responses with aging (45), an effect that could contribute to suppressed airway responsiveness in vivo. Although the functional role of the DGC in contractile smooth muscle cells has not been established, some studies provide clues in this area. North et al. (50) reported that dystrophin is highly organized in contractile smooth muscle cells, segregating into longitudinal linear arrays in association with caveolae-rich plasma membrane domains. Interestingly, this more or less mimics skeletal muscle, in which the DGC is sequestered to costamers and provides a mechanical link between the ECM and the Z disk (42). In skeletal muscle the DGC appears to act as a molecular shock absorber during contraction and relaxation (17). To date no studies have directly assessed the role of the DGC in mechanical load bearing in smooth muscle cells, although a recent report from Dye and colleagues (16) reveals that carotid arteries from mdx and δ-SG knockout mice exhibit decreased pressure-induced distensibility and increased circumferential and axial stress. Most reports have suggested that disruption of the DGC in smooth muscle may be linked to changes in Ca2+ homeostasis. The absence of dystrophin in portal veins from mdx mice leads to reduced intercellular myocyte communication associated with stretch-induced myogenic contractile responses (44). Notably, ectopic smooth muscle-specific expression of dystrophin can improve aberrant vasoregulation in mdx mice (38). Morel et al. (48) reported that decreased mechanical activity of duodenal smooth muscle in mdx mice is due to reduced type 2 ryanodine receptor expression that compromises sarcoplasmic reticulum Ca2+ release. Interestingly, Cohn et al. (12) showed that cardiac myopathy associated with coronary artery vasospasm in sarcoglycan knockout mouse models can be prevented by verapamil, a vasodilatory Ca2+ channel blocker. The precise mechanism linking the DGC with Ca2+ handling in smooth muscle cells is not clear; however, its association with caveolae (50) provides an interesting lead. In smooth muscle, including ASM, caveolae are sites where ion channels and Ca2+-binding proteins are sequestered, and caveolae are thought to be spatially aligned with the sarcoplasmic reticulum to facilitate receptor-mediated Ca2+ release (14, 30). In fact, we recently confirmed (22) that caveolae facilitate G protein-coupled receptor-mediated contraction and Ca2+ release in ASM muscle. Future studies are needed to better ascertain the precise role of DGC in excitation-contraction coupling and mechanical force transmission in contractile smooth muscle cells and tissue.
In summary, our study characterized DGC subunit expression in human ASM tissue and demonstrates that the expression of DGC proteins is associated with, and dependent upon, phenotype switching between contractile and proliferative states in human ASM cells. Notably, we also demonstrate that DGC expression is subject to regulation by mechanisms involving laminin-integrin binding and the induction of PI3K signaling that are essential for ASM cell maturation. Our studies provide an important new platform for future studies investigating the direct role of the DGC, in particular laminin-2 binding to α-DG, in generating intracellular signaling cascades that support myocyte maturation and/or maintenance of a contractile phenotype. In addition, our studies provide compelling evidence to support future studies investigating the functional role of the DGC in contractile ASM, in particular in relation to mechanical load bearing and Ca2+ homeostasis.
This project is supported by a grant from the Canadian Institute of Health Research (CIHR) and is supported in part by funding through the Canada Research Chair program, Canada Foundation for Innovation, and the Manitoba Institute of Child Health (MICH). P. Sharma holds a graduate studentship from MICH and the CIHR National Training Program in Allergy and Asthma. T. Tran was supported by a CIHR/Canadian Lung Association/GlaxoSmithKline Fellowship and the CIHR National Training Program in Allergy and Asthma. R. Gosens received support from the National Training Program in Allergy and Asthma (NTPAA) and is the recipient of a Marie Curie Outgoing International Fellowship from the European Community (008823). A. J. Halayko holds a Canada Research Chair in Airway Cell and Molecular Biology.
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.
- Copyright © 2008 the American Physiological Society