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Rationale: Airway smooth muscle (SM) of patients with asthma exhibits a greater velocity of shortening (Vmax) than that of normal subjects, and this is thought to contribute to airway hyperresponsiveness. A greater Vmax can result from increased myosin activation. This has been reported in sensitized human airway SM and in models of asthma. A faster Vmax can also result from the expression of specific contractile proteins that promote faster cross-bridge cycling. This possibility has never been addressed in asthma.
Objectives: We tested the hypothesis that the expression of genes coding for SM contractile proteins is altered in asthmatic airways and contributes to their increased Vmax.
Methods: We quantified the expression of several genes that code for SM contractile proteins in mild allergic asthmatic and control human airway endobronchial biopsies. The function of these contractile proteins was tested using the in vitro motility assay.
Measurements and Main Results: We observed an increased expression of the fast myosin heavy chain isoform, transgelin, and myosin light chain kinase in patients with asthma. Immunohistochemistry demonstrated the expression of these genes at the protein level. To address the functional significance of this overexpression, we purified tracheal myosin from the hyperresponsive Fisher rats, which also overexpress the fast myosin heavy chain isoform as compared with the normoresponsive Lewis rats, and found a faster rate of actin filament propulsion. Conversely, transgelin did not alter the rate of actin filament propulsion.
Conclusions: Selective overexpression of airway smooth muscle genes in asthmatic airways leads to increased Vmax, thus contributing to the airway hyperresponsiveness observed in asthma.
An excessive decrease in airway luminal area via bronchoconstriction is one of the final pathways to asthma. However, very little is understood about the molecular mechanics of smooth muscle in airway hyperresponsiveness and asthma.
Selective overexpression of airway smooth muscle genes in asthmatic airways leads to increased Vmax, thus contributing to the airway hyperresponsiveness observed in asthma.
The primary features of asthma are airway inflammation, bronchial hyperresponsiveness, and intermittent airway obstruction. Altered airway smooth muscle (SM) function is considered to be an important contributor to airway hyperresponsiveness and asthma (1). As suggested by mathematical models, airway narrowing results from a balance between the contraction produced by airway SM and the impedance of the surrounding tissues (2, 3). Asthmatic airway SM exhibits enhanced contractility (4, 5). The increased rate and extent of shortening observed in hyperresponsive tissues have traditionally been attributed to increased airway SM mass (6–8), but this has recently been challenged as the main explanation for airway hyperresponsiveness (9, 10). Regardless of possible alterations in airway SM mass, the force normalized to mass generated by asthmatic SM strips is greater than that of nonasthmatic airway SM (10). Another factor that is likely to contribute to power enhancement of hyperresponsive airway SM is the increase in velocity (Vmax) of SM shortening. Indeed, a greater velocity of airway SM shortening has been observed in many animal models of asthma (11–13), in sensitized human bronchi (5), and in single SM cells from asthmatic human airways (14). Two mechanisms have been proposed whereby airway SM that exhibits increased Vmax could lead to excessive bronchoconstriction. The first mechanism involves a greater Vmax during the initial active portion of contraction (15, 16), whereas the second mechanism implicates a greater Vmax after muscle stretching, such as occurs during tidal breathing, thereby counteracting any potential relaxing effects (17, 18).
The contractile apparatus is responsible for muscle force and movement production. Vmax depends on the contractile proteins involved and their level of activation. Myosin is the molecular motor that drives muscle contraction. Alternative splicing of the SM myosin heavy chain (SMMHC) gene generates four isoforms. Splicing in the 5′ region results in the expression of two isoforms that differ by the presence (SM-B) or the absence (SM-A) of a seven-amino-acid insert in the surface loop above the nucleotide binding pocket (19, 20). The SM-B and SM-A isoforms are also referred to as (+) and (−) insert isoforms, respectively. Splicing in the 3′ region leads to the expression of two isoforms that differ by distinct sequences of 43 (SM-1) or 9 (SM-2) amino acids at the carboxy terminal (21, 22). Although no differences in the molecular mechanics of SM-1 and SM-2 have been reported, SM-B propels actin filaments at two times the velocity (νmax) of SM-A in the in vitro motility assay (23–25). The expression and function of these SMMHC isoforms in airway SM hypercontractility has not been thoroughly addressed.
It has been well established that SM contraction is mainly regulated by phosphorylation of the myosin regulatory light chains (LC20) by the 108-kD myosin light chain kinase (MLCK) (26). Increased expression of MLCK has been described in models of asthma and in human asthmatic airway SM (16, 27, 28). Actin-binding proteins are also known to alter SM cross-bridge kinetics. For example, caldesmon decreases ATPase activity and νmax (29, 30), whereas tropomyosin increases only νmax (30). However, the expression of actin-binding proteins has not been addressed in airway hyperresponsiveness and asthma.
In this study, we hypothesized that the expression of genes that code for contractile proteins is altered in asthmatic airway SM, thus contributing to their increased Vmax. We found that the expression of SM-B, transgelin (SM22), and MLCK is increased in endobronchial biopsies from humans with mild asthma. Furthermore, we found that the myosin purified from airways of the Fisher rats, an animal model of innate bronchial hyperresponsiveness that overexpresses SM-B in airway SM, has a greater νmax than myosin from control animals. Conversely, SM22 had no effect on cross-bridge cycling rate. Our combined human and rat data suggest that the selective contractile protein gene expression measured in asthmatic airway SM leads to increased velocity of shortening, as measured in the rats, thus contributing to airway hyperresponsiveness.
Thirteen subjects with asthma, whose diagnoses were made according to the definition of the American Thoracic Society, and 14 healthy subjects without allergy history, asthma, or occupational exposure to sensitizing agents were recruited. The clinical characteristics of the subjects are provided in Table 1. Evaluation included a medical history, physical examination, skin prick tests to common allergens (Omega, Montreal, PQ, Canada), spirometry, and measurements of airway responsiveness to inhaled methacholine according to standardized procedures (31). All subjects were nonsmokers and had not had respiratory infection within the last 2 months. Subjects with asthma were stable and were receiving a treatment of inhaled β2-agonist on demand but no inhaled corticosteroid therapy. Control subjects had no systemic disease, were not receiving medication, and had negative skin prick tests. The study was approved by the Laval Hospital and the Montreal Chest Institute Research Ethics Board of McGill University Health Center, and all subjects provided written informed consent.
Oxygen was administered at 5 liters per minute by a nasal catheter, and vital signs, electrocardiogram, and oxymetry data were monitored during the bronchoscopy. Local anesthesia of the airways was done with 2 or 4% lidocaine up to a total dose of 400 mg. A flexible bronchoscope (Olympus OES 10 fiberscope; Olympus, Markham, ON, Canada) and alligator forceps (Olympus FB-15C-1) were used for the procedure. In five subjects with asthma and five control subjects, six or seven specimens were taken from the origins of subsegmental, segmental, or lobar bronchial carinae and kept in RNA-later solution (Qiagen, Inc., Mississauga, ON, Canada) for mRNA extraction. In one control subject and two subjects with asthma, additional biopsies were taken and fixed in 4% paraformaldehyde for immunohistochemistry. Nine additional control and eight asthmatic specimens, collected as mentioned above, were obtained from The Tissue Bank of the Respiratory Health Network of the FRSQ.
We quantified by real-time PCR the mRNA expression of myosin isoforms (SM-1, SM-2, SM-A, SM-B), actin isoforms (α, γ), SM myosin light chain kinase (MLCK), tropomyosin isoforms (α, β), SM caldesmon, and transgelin (SM22) in the endobronchial biopsies of five subjects with asthma and five healthy subjects. All primer sets spanned at least one intron. The sequence of the primers is shown in Table 2. The primer set called “Total SMMHC” amplifies the four SMMHC isoforms because it targets a region not subject to alternative splicing. The six to seven biopsies from each patient were pooled and homogenized in RNAlater buffer, and total RNA was extracted using a commercial kit (Mini Prep, Qiagen) following the manufacturer's recommendations. RNA (1 μg) from each of these subjects was reverse transcribed simultaneously to minimize variability. PCR reactions were performed in a volume of 20 μl, containing 1 μl cDNA, 10 μL 2× QuantiTect SYBR Green PCR (Qiagen), 7 μl of nuclease-free H2O, and 1 μl of both the forward and reverse primers (final concentration, 0.1 μmol each). The samples were amplified in a LightCycler system (Roche Diagnostics, Laval, PQ, Canada). The real-time PCR conditions consisted of a denaturation step of 15 minutes at 95°C followed by an amplification of 50 cycles (denaturation at 95°C for 15 seconds, annealing at 60°C for 20 seconds, and extension at 72°C for 20 seconds) and one melting curve cycle. PCR reactions were performed in triplicate (i.e., three repeats on the same RNA sample pooled from each subject). Each primer set generated only one PCR product. PCR reaction efficiencies were calculated for each reaction following a previously described method (32, 33) and were used for relative quantification with REST software (v384) (34). REST uses the following equation to calculate the ratio of expression of a target gene between two groups (sample and control) when the target gene is referenced to a housekeeping gene:
where CT is the threshold cycle number (a fixed threshold where the PCR amplification is still in the exponential phase and the reaction components are not limiting gene amplification [32, 33]), and E is the efficiency of the reaction. Thus, results were expressed in fold increase in patients with asthma compared with control subjects using the ribosomal protein S9 as a reference.
Protein expression of the SM-B SMMHC isoform, SM22, and MLCK was verified by immunohistochemistry in 10 subjects with asthma and 10 control subjects (some of the subjects were tested at the mRNA and protein levels; see Table 1). The paraformaldehyde-fixed biopsy specimens were dehydrated in alcohol, incubated in xylene, and embedded in blocks of fresh molten paraffin wax. Sections (5 μm) were made with the cutting angle in the axis of the bronchial mucosa. For general assessment of the tissue, three slides were deparaffinized, rehydrated, and stained with hematoxylin-eosin. Only biopsies with smooth muscle bundles and preserved morphology were used. A peroxidase protocol for paraffin-embedded sections was used for immunohistochemistry. For each target protein, all procedures were performed simultaneously and in the same conditions for subjects with asthma and normal subjects. Slides were deparaffinized in xylene and in decreasing concentrations of alcohol. Tissues were permeabilized with Triton 0.2% and incubated with 5% peroxide except for MLCK, where they were initially incubated in citrate buffer (10 mM citric acid [pH 6.0]). After blocking with universal blocking solution for 30 minutes (Dako, Carpinteria, CA), slides were incubated overnight with the following antihuman primary antibodies against the SM-B SMMHC isoform (Genscript Corp., Piscataway, NJ), SM22 (ab10135; ABCam), and MLCK (M7905; Sigma, St. Louis, MO). Subsequently, the slides were incubated with biotinylated secondary antibody (E0453; Dako) and with ABC complex (Dako) and stained with DAB (diaminobenzidine)-chromogen (Dako) and counterstained with hematoxylin.
To control for nonspecific binding, a competitive preadsorption test was performed. A synthetic peptide of the seven-amino-acid insert sequence (QGPSFAY), purified smooth muscle MLCK (generous gift from Dr. J. Haeberle), and recombinant SM-22 (see below) were incubated for 2 hours at room temperature with their respective antibodies at a molar ratio of 1:100. Incubation of slides with these antigen–antibody complexes were then performed in the same conditions as described above.
We used the in vitro motility assay to quantify the rate of actin filament movement (νmax) when propelled by myosin molecules. The in vitro motility assay was described before (24, 35). The buffers were as follows: myosin buffer (300 mM KCl, 25 mM imidazole, 1 mM EGTA, 4 mM MgCl2, 25 mM DTT [pH 7.4]), actin buffer (25 mM KCl, 25 mM imidazole, 1 mM EGTA, 4 mM MgCl2, 25 mM DTT [pH 7.4]), and motility buffer (0.5% methylcellulose, 2 mM MgATP, 25 mM KCl, 25 mM imidazole, 1 mM EGTA, 4 mM MgCl2, 25 mM DTT [pH 7.4]) combined with an oxygen scavenger (0.1 mg/ml glucose oxidase, 0.018 mg/ml catalase, 2.3 mg/ml glucose). Myosin regulatory light chains were thiophosphorylated with 0.6 mM ATPγS, 7.5 μM calmodulin, 5 μM MLCK, 0.2 mM CaCl2, and 2 mM MgCl2. Inactive myosin molecules were removed by ultracentrifugation (24). Myosin (0.25 mg/ml) was then perfused into a flow-through chamber (36) and allowed to attach randomly onto the nitrocellulose-coated glass. Inactive myosin molecules were further inhibited by the addition of monomeric unlabeled actin dissolved in actin buffer. Actin fluorescently labeled with tetramethylrhodamine isothiocyanate phalloidin (TRITC P1951; Sigma) in actin buffer was then perfused into the chamber, followed by motility buffer. Actin movement was recorded by a SIT camera, digitized, and analyzed (Scion-Image software). Measurements were made at 30°C. νmax was reported in μm/s ± SD. νmax was measured for three separate purifications from each rat strain. For each in vitro motility assay, the movement of a minimum of 90 filaments was analyzed.
Tracheal SM myosin was purified from eight Fisher and eight Lewis rats, following our previously described protocol for small samples (35). All experiments were conducted in compliance with the animal ethics committee of McGill University. Chicken gizzard SM myosin was purified following a standard protocol (37). Skeletal myosin was a generous gift from Dr. P. VanBuren (University of Vermont, Burlington, VT). Unregulated actin was purified from chicken pectoralis acetone powder following the Pardee and Spudich protocol (38). Recombinant SM22 was generated in Escherichia coli as previously described (39). Briefly, cDNA containing the full-length SM22 coding region was ligated in frame into BamH I– and Hind III–digested pQE-30 bacterial expression vector (Qiagen) containing an NH2-terminal MRGHHHHHHGS “His” tag. Recombinant His-tagged SM22 was expressed in Escherichia coli and batch purified under native conditions using nickel (Ni-nitrilotriacetic acid [NTA]) beads (Qiagen). Transformed E. coli were grown to optical density of 0.6 to 1.0 at 600 nm, and then recombinant protein expression was induced for 6 hours with 1 mM isopropyl-β-D-thiogalactopyranoside. Bacteria were lysed on ice for 30 minutes in buffer containing 50 mM sodium phosphate (pH 8.0), 300 mM NaCl, 10 mM imidiazole, and 1 mg/ml lysozyme followed by sonication on ice using a microtip. Lysate was centrifuged to remove cell debris, and then Ni-NTA beads were added and allowed to bind at 4°C for 30 minutes. Bound proteins and beads were washed four times with 50 mM sodium phosphate (pH 8.0), 300 mM NaCl, and 50 mM imidiazole. The retained proteins were eluted and stored in buffer containing 50 mM sodium phosphate (pH 8.0), 300 mM NaCl, 250 mM imidazole, and protease (serine, cysteine, calpain, and metalloprotease) inhibitors (Roche). Two purifications of SM22 recombinant protein were done and tested in the in vitro motility assay.
SM22 was bound to unregulated actin following a previously described protocol (39) with slight modifications. Briefly, recombinant SM22 (3 μM) was incubated in presence of unregulated monomeric actin (10 μM) and tetramethylrhodamine isothiocyanate phalloidin (10 μM) (Sigma) for 12 hours at 4°C in a buffer containing 12 mM KPi (pH = 6.8), 20 mM imidazole, 24 mM NaCl, 2 mM MgCl2, 1 mM ATP, and 1 mM EGTA. To ascertain that only SM22-bound actin was used in the in vitro motility assay, cosedimentation of actin with SM22 was performed, and the pellet was washed twice and resuspended in actin buffer before use in the in vitro motility assay. For cosedimentation, the actin-SM22 solution was spun at 100,000 × g for 21 minutes. We further verified that the pellet contained mostly actin-bound SM22 using immunoprecipitation. The acto-SM22 mixture was incubated with an actin-specific antibody (#4700; Sigma) at 4°C. Protein-A sepharose beads (Amersham, Baie D'Urfe, PQ, Canada) were then incubated with the acto-SM22 solution for 1 hour before being spun down at 15,000 rpm for 2 minutes. The supernatant and the pellet were then boiled in Leamli buffer and loaded onto a gel for SDS-PAGE, followed by Western blotting (described below), with a primary antibody that targeted SM22 (ab10135; ABCam, Cambridge, MA). To ascertain that the sepharose beads were not nonspecifically pulling down actin or SM22, control experiments were performed using different primary and secondary antibodies.
The relative content of the SM-B myosin isoform with respect to total myosin in the Fisher and Lewis rat trachealis purifications was determined by Western blot analysis. Proteins were separated by SDS-PAGE using a 7% acrylamide gel. Equal volumes of myosin solutions were loaded onto a single gel. Protein concentration was estimated by a standard Bradford assay, and 100 μg of proteins were loaded in each well. Proteins were transferred electrophoretically onto nitrocellulose membranes (Bio-Rad, Hercules, CA). Membranes were blocked with 5% nonfat milk and probed with the polyclonal SM-B6 antibody that specifically recognizes the seven-amino-acid insert QGPSFAY (20, 40) or with the polyclonal BT-562 antibody (Biomedical Technologies Inc., Stroughton, MA) that recognizes all SMMHC isoforms. Antibody detection was done by enhanced chemiluminescence (Amersham), and quantification was performed with a Fluorchem 8500 imaging system using AlphaEase software (Alpha Innotech, San Leandro, CA). Quantification was performed for three separate purifications and loaded in duplicate for each rat strain on six nitrocellulose membranes. The results are reported as mean ± SE.
Differences in mRNA expression between biopsies from subjects with asthma and control subjects were tested using Pair Wise Fixed Reallocation Randomization Test in REST software (v 2). Differences in the SM-B tracheal protein content and in νmax between Fisher and Lewis rats and between actin bound SM22 and no SM22 were tested using Student's t tests. Significance was considered for values of P ≤ 0.05.
We quantified, by real-time PCR, the expression of SM-specific contractile protein genes in endobronchial biopsies from subjects with asthma and control subjects (Figure 1). The housekeeping gene ribosomal S9 unit was very stable and had similar levels in asthmatic and control biopsies (asthmatic/control: +1.03; P = 0.78). Thus, ribosomal S9 was an adequate housekeeping gene. There was no significant difference in mRNA expression between patients with asthma and control subjects in the SMMHC SM-1 isoform, tropomyosin β, caldesmon, and α-actin. The expression of tropomyosin α (asthmatic/control: 0.78 ± 0.16 SE; P < 0.03) and γ-actin (asthmatic/control: 0.53 ± 0.15 SE; P < 0.01) was significantly lower in patients with asthma compared with control subjects. Conversely, the mRNA expression of total SMMHC, SM-2, SM-A, and SM-B isoforms was significantly up-regulated in subjects with asthma, along with significant up-regulation of SM22 and MLCK. The increase in SM-A expression (asthmatic/control: 1.63 ± 0.40 SE; P < 0.002) and SM-2 (asthmatic/control: 1.53 ± 1.03 SE; P < 0.04) was of similar magnitude as that of total SMMHC (asthmatic/control: 1.69 ± 0.28 SE; P < 0.001). SM-B (asthmatic/control: 2.41 ± 0.80 SE; P < 0.006) showed the greatest up-regulation among SMMHC isoforms and was the only myosin isoform with a significant up-regulation in patients with asthma relative to total SMMHC. SM22 (asthmatic/control: 2.18 ± 1.14 SE; P < 0.01) was also highly up-regulated, whereas the up-regulation of MLCK (asthmatic/control: 1.21 ± 0.24 SE; P < 0.05) was modest. Thus, these results show that the mRNA encoding for the SM-B SMMHC isoform, SM22, and MLCK is up-regulated in mild asthmatic endobronchial SM. When normalized to α-actin, the mRNA encoding for SM-B (asthmatic/control: 2.21 ± 0.74; P = 0.03), SM22 (asthmatic/control: 1.91 ± 0.62; P = 0.04), total SMMHC (asthmatic/control: 1.55 ± 0.25; P = 0.05), and SM-A (asthmatic/control: 1.50 ± 0.37; P = 0.05) was still seen to be up-regulated in patients with asthma compared with control subjects, whereas that of MLCK and SM-2 was not.
To confirm that SM-B, SM22, and MLCK are expressed at the protein level in human airways SM and to visualize their localization, we performed immunohistochemistry on the endobronchial biopsies. The positive staining for the SM-B SMMHC isoform had a uniform pattern (Figures 2A and 2B). At high magnification, the staining was intracellular and had an elongated spindle shape when the muscle bundle was cut longitudinally (Figures 2A and 2B). The SM-B–positive cells were usually dispersed homogeneously in the SM tissue, although they were seldom grouped (Figures 2A and 2B). The positive staining for SM22 was uniform and intracellular (Figures 3A and 3B). The SM22-positive cells were dispersed homogeneously in the SM tissue (Figures 3A and 3B). The staining for MLCK confirmed its expression in the airway SM cells at the protein level (Figures 4A and 4B). The absence of staining after competitive preadsorption (Figures 2B, ,3B,3B, and and4B)4B) confirmed that the signal was specific in all cases.
To determine if overexpression of the SM-B isoform contributes to muscle hypercontractility, we measured the rate of actin filament propulsion (νmax) in the in vitro motility assay of purified Fisher and Lewis rat trachealis myosin. νmax was significantly faster for the myosin purified from Fisher rat tracheae (0.61 ± 0.01 μm/s) than from Lewis rat tracheae (0.47 ± 0.01 μm/s; P < 0.001) (Figure 5A). To verify that myosin purification did not alter the previously reported SM-B content in Fisher and Lewis rat trachealis (41), we performed Western blot analysis. The hyperresponsive Fisher rat trachealis had 1.8 ± 0.2 fold more of the SM-B isoform (P < 0.001) than the hyporesponsive Lewis rat (Figure 5B). The difference between Fisher and Lewis rats is of the same magnitude as what we observed between the asthmatic and control biopsies (see Figure 1). Altogether, these data suggest that the overexpression of the SM-B isoform results in a faster myosin cycling velocity and thus a faster velocity of SM contraction.
To determine if the overexpression of SM22 observed in the subjects with asthma contributes to muscle hypercontractility, the molecular mechanics of SM22-bound actin filaments, when propelled by myosin molecules, was assessed in the in vitro motility assay. No difference in νmax was observed when SM22 was absent (0.79 ± 0.01 μm/s) or when it was bound to actin (0.79 ± 0.02 μm/s) (Figure 6). The binding of SM22 to actin was verified by immunoprecipitating the co-sedimented actin-SM22 with an actin-specific antibody and analyzing by Western blotting with an SM22-specific antibody (Figure 7A). SM22 was mostly found in the pellet obtained after immunoprecipitation of actin, showing that actin was effectively bound to SM22 (Figure 7A). Specificity of the immunoprecipitation procedure was verified by controlled experiments with antibodies against P44 MAP kinase and the chemokine CCR5 (Figure 7B). These data show that, at the molecular level, SM22-bound actin does not directly alter cross-bridge cycling rate.
To further investigate the relevance of the expression of these contractile protein genes to the clinical status of the subjects, we compared the PC20 values with the mRNA data. The S9 normalized target gene abundance was calculated as:
A linear relationship was seen between the PC20 and total SMMHC expression normalized to S9 (Figure 8; R2 = 0.6247). No clear relationship was seen between PC20 and SM-B, SM22, or MLCK (data not shown).
In this study, we found that the expression of specific genes encoding for SM contractile proteins is up-regulated in asthmatic bronchial biopsies. We showed from a rat model of airway hyperresponsiveness that the up-regulation of the SM-B myosin isoform leads to a faster rate of νmax in the in vitro motility assay. Because νmax correlates with unloaded shortening velocity (42, 43), a greater expression of the SM-B isoform contributes to the faster rate of airway SM shortening. We also found that SM22 is overexpressed in asthmatic bronchial biopsies. However, we did not elucidate its function at the molecular level.
Two mechanisms have been suggested to explain how an increased Vmax could potentially lead to airway hyperresponsiveness. The first theory emerged from the measurements performed by Jiang and coworkers, who showed in their allergic dog model that the development of Vmax occurs during the first 2 seconds of a 10-second contraction (15, 16). Their results also suggested that the remaining contraction is handled by more slowly cycling cross-bridges, so-called “latch-bridges.” Similarly, they showed in human bronchial SM cells that 90% of the shortening occurs during the first 1.5 seconds of the contraction (14). These observations led to the conclusion that a faster contractile machinery would produce a greater extent of shortening during the first few seconds of active contraction, thus a greater airway narrowing (15, 16). The second theory suggested to explain how a greater Vmax could enhance airway responsiveness emerged from the more recent advances on the effects of deep inspiration. When subjects without asthma are prevented from taking deep breaths, they experience airway hyperresponsiveness similar to subjects with asthma (44). Thus, airway SM stretching by tidal inflation or deep breaths decreases airway resistance in normal lungs but not in patients with asthma (44, 45). Modern imaging techniques have demonstrated that airways of patients with asthma also dilate upon stretching but that this dilation is transient because asthmatic airways quickly narrow back to their initial diameter (46). These results suggest that asthmatic airway SM can shorten faster than normal SM after a stretch. This rapid reconstriction might maintain asthmatic airway SM in a more constricted state because it would have time to shorten significantly between each breath, which would counteract the relaxing effect of tidal breathing (17, 18). Indeed, it is thought that the degree of airway narrowing is an equilibrium state that depends on the rate of acto-myosin cross-bridge interaction versus the rate of stretch imposed to airway SM by tidal breathing (47). Therefore, a greater rate of SM shortening is more likely to compensate for the disruptions of the cross-bridges induced by oscillatory stretches (17, 18, 47, 48). Our results of alterations in expression of mRNA from the contractile machinery of patients with asthma, along with our in vitro motility data from the Fisher-Lewis rat model of airway hyperresponsiveness, offer molecular mechanisms to explain the increased rate of airway SM shortening observed in asthma.
The rate of shortening of airway SM is increased in human asthmatic SM cells (14), human sensitized airways (5), and animal models of bronchial hyperresponsiveness (11–13, 49). In the current study, we showed that the mRNA expression of SM-B doubles in airway SM from patients with mild asthma compared with normal subjects. Furthermore, we demonstrated that an increased SM-B isoform content in airway SM has a functional impact by measuring νmax for myosin purified from Fisher and Lewis rat trachealis muscle. We chose this animal model because fresh human airway SM is not readily available and because the Fisher rats are hyperresponsive (41, 50) and express more SM-B than the Lewis rats (41) (see Figure 5B). The difference in SM-B isoform expression between Fisher and Lewis rats is of the same magnitude as what we observed between asthmatic and control human biopsies. One limitation of our study is that molecular or muscle strip level measurements do not take into account any potential differences in the load on the muscle between subjects with asthma and control subjects. Nonetheless, the data in the literature supporting such differences in load are still very limited.
Myosin is a mechano-enzyme: The hydrolysis of MgATP leads to actin binding and a conformational change, which results in the translational movement of actin. Structural differences between the SM-B and SM-A SMMHC isoforms explain the differences in their contractile kinetics. The seven-amino-acid insert is strategically located to alter the in and out movement of the nucleotide (20, 23, 51). Indeed, it has been shown that the presence of the insert increases the flexibility of the loop above the nucleotide binding pocket and as a consequence facilitates the binding of MgATP and the release of the hydrolysis product, MgADP (52). This was confirmed in our previous study (24) in which we used a laser trap to demonstrate that the time of attachment of SM-B to actin is half that of SM-A, thereby explaining the twofold faster νmax for SM-B measured in the in vitro motility assay (23, 25). We also showed that it is the proportion of the SM-B to SM-A isoform that determines νmax (24).
The presence of the myosin SM-B isoform is also known to affect SM kinetics at the whole organ level. We previously purified myosin from multiple rat organs and showed a rank correlation in νmax from the slowest contracting tonic organ to the fastest contracting phasic organ (35). Furthermore, by measuring the pulmonary mechanics of the SM-B isoform knockout mouse (53), we showed the impact of the seven-amino-acid insert at the whole animal level (54). We observed an 18% increase in the time to peak airway resistance in the knockout compared with the wild-type mice (54). It is also noteworthy that the expression of the SM-B isoform has been shown to be altered in several models of disease. For example, bladder or intestinal obstruction causes a reduction in mRNA encoding for SM-B, with concurrent reductions in maximal shortening velocity (55–57). Hormones (58) and stages of development also alter the expression of the SM-B myosin isoform (51).
Immunohistochemistry showed intercellular heterogeneity in the distribution of SM-B in biopsies from normal subjects and subjects with asthma. Positive SM-B cells were not usually observed in clusters but individually dispersed. Similar results have been reported in swine stomach SM and rat lung airway SM (40, 59). Parisi and Eddinger (59) have suggested that such heterogeneity might be critical in conferring a wider range of shortening velocity to SM. For example, they suggested that cells expressing high levels of the SM-B isoform could, by contracting rapidly, stretch and activate SM-B–deprived neighborhood cells via mechano-transduction. Such a mechanism could explain how heterogeneity in cell phenotype would lead to amplification of the rate of SM cell activation, provided that the SM-B–deprived cells do not get stretched beyond their optimal length.
It is widely accepted that airway SM mass is increased in asthma (2, 6, 28, 60, 61). Our results of a 1.7-fold up-regulation of SMMHC mRNA (see Figure 1) in the asthmatic biopsies may indirectly reflect an increase in airway SM mass. However, the mRNA data were normalized to a ubiquitously expressed ribosomal S9 gene, so our results do not allow for distinction between a greater number of SM cells expressing SMMHC (hyperplasia) and a similar number of SM cells, each expressing more SMMCH (hypertrophy). Nevertheless, the fact that the expression of the α-actin was not greater in the subjects with asthma compared with control subjects and that the γ-actin in subjects with asthma was even lower than that of control subjects strongly suggests that the airway smooth muscle mass was not significantly greater in the subjects with asthma. These results may seem at odds with other studies that have reported a greater muscle mass (i.e., greater α-actin expression ) in subjects with asthma compared with control subjects, but this is most likely a reflection of technical differences. Our biopsies are probably taken deeper, leading to a greater amount of muscle collected in the control subjects than reported in other studies (alligator jaw forceps [FB-15C-1; Olympus, Centre Valley, PA] of volume 2.89 mm3 were used). Our biopsies have been analyzed in a separate study for the SM area normalized to the total surface of the biopsy (62). No statistical differences were observed between subjects with mild to moderate asthma and control subjects. Nevertheless, not having differences in amount of SM between our subjects with asthma and control subjects would have only contributed to reducing the differences that we observed in gene expression.
MLCK is the key activator of SM contraction by phosphorylation of the LC20, thereby initiating the cross-bridge cycling (63). The maximal velocity of SM contraction is tightly dependent on the acto-myosin cross-bridge cycling rate (42, 43). Many studies have suggested that the cross-bridge cycling rate is proportional to the amount of MLCK and the resulting LC20 phosphorylation levels (64, 65). However, this direct link remains controversial (66–68). Nonetheless, several studies have reported significant increases of MLCK expression in asthma and in models of asthma. MLCK protein content was found to be increased in human bronchial SM sensitized ex vivo (27) and in a dog model of allergic airway hyperresponsiveness (16, 69). Our data show an up-regulation of MLCK mRNA in patients with asthma compared with control subjects. Our data agree with those of Ma and coworkers (14), who studied whole tissue biopsies from a similar subject population but used conventional RT-PCR. Others used laser capture to dissect out the SM portion of biopsies, along with mRNA amplification techniques, and did not find significant changes in MLCK expression between asthmatic and control bronchial biopsies (60). At the protein level, a morphometric analysis in patients with severe and moderate asthma showed a significant increase in MLCK in the patients with severe asthma (28). Discrepancies between studies may be due to differences in patient populations and quantification techniques.
SM22 is highly expressed in SM, but it has also been reported in fibroblasts (70). Because our analysis was performed in whole biopsy samples, we cannot be sure of the cellular source of the SM22 overexpression observed in our patients with asthma. Moreover, although SM22 was first purified from SM more than a decade ago (71), its function remains unknown. SM22 binds to filaments of actin in SM cells in vitro and in vivo (39, 71, 72) and is not essential for life and SM development (72). We hypothesized that SM22 would behave as several other actin-binding proteins and directly alter the cross-bridge cycling rate and/or ATPase activity (73). Our results demonstrate no difference in νmax when SM22 is bound to actin. It is possible that SM22 has a role in actin filament remodeling. In this sense, actin elongation has been suggested to play a role in bronchial hyperresponsiveness (74). It is also possible that the effect of SM22 involves multiple actin filaments, as suggested by the SM22 jellifying action on actin filaments, observed in specific ionic strength conditions (75). This would be missed when performing measurements at the molecular level but should be investigated at the cellular level.
Although several markers of SM have been studied in asthma, myosin, being the molecular motor that drives contraction, is the marker of choice. The linear relationship between PC20 and the mRNA expression level of total SMMHC supports the notion that airway SM plays a critical role in the response to MCh. To our knowledge, these relationships have not been addressed before. Further studies are necessary to fully understand the role of myosin proteins in airway hyperresponsiveness and asthma.
In summary, we have shown that the mRNA levels of SM-B, SM22, and MLCK are increased in mild asthmatic bronchial biopsies. Furthermore, we showed that up-regulation of rat airway SM-B increases νmax. This specific contractile protein gene up-regulation most likely leads to increased SM velocity of shortening, thus contributing to the airway hyperresponsiveness as observed in asthma. Further quantitative analysis is needed to confirm the up-regulation of these genes at the protein level.
The authors thank C. Racine, S. Al-Mot, Dr. S. Lajoie-Kaldoch, and Dr. S. Létuvé for technical assistance. The authors also thank the Tissue Bank of the Respiratory Health Network of the FRSQ for providing human specimens and Marvid Poultry for chicken gizzards and pectoralis.
Supported by Canadian Institute of Health Research grants MGC-42667 and MOP-79545; Réseau en Santé Respiratoire du Québec, Natural Sciences and Engineering Research Council grants RGPIN 217457-00 and EQPEQ 229606-00; and by National Institutes of Health grant R01 HL79398. The Meakins-Christie Laboratories are supported in part by a Center grant from Le Fonds de la Recherche en Santé du Québec.
Originally Published in Press as DOI: 10.1164/rccm.200609-1367OC on November 14, 2008
Conflict of Interest Statement: R.L. does not have a financial relationship with a commercial entity that has an interest in the subject of this manuscript. M.L. participated or participates in clinical trials for Altana, AstraZeneca, Boehringer-Ingelhiem, GlaxoSmithKline, Bayer, Merck, Asthmatx, and Topingen. For the Asthmatx trials, M.L.'s institution, the Laval Hospital, received $182,000 in 2004, $115,000 in 2005, $151,000 in 2006, and $169,000 in 2007 for reimbursement for expenses related to clinical trials of bronchial thermoplasty. M.L. gave lectures sponsored by Altana, AstraZeneca, 3M, GlaxoSmithKline, Merck, and Asthmatz. M.L. received a medical research grant ($50,000) from Merck-Frosst in 2004. C.B. does not have a financial relationship with a commercial entity that has an interest in the subject of this manuscript. N.Z. does not have a financial relationship with a commercial entity that has an interest in the subject of this manuscript. P.K. does not have a financial relationship with a commercial entity that has an interest in the subject of this manuscript. J.S. has received or will receive consultant fees of $1,000 in 2005 from Critical Therapeutics, $1,000 in 2006 from Tanox, $1,600 in 2006 from Merck, $2,700 in 2007 from AstraZeneca, $3,500 in 2007 from Genentech, $6,000 in 2008 from Cytokinetics, and $2,500 in 2008 from Sepracor. J.S. has served as principal investigator on a grant of $125,031 from AstraZeneca in 2008. L.K. does not have a financial relationship with a commercial entity that has an interest in the subject of this manuscript. Q.M. does not have a financial relationship with a commercial entity that has an interest in the subject of this manuscript. A-M.L. does not have a financial relationship with a commercial entity that has an interest in the subject of this manuscript.