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Am J Physiol Lung Cell Mol Physiol 295: L479-L488, 2008. First published June 27, 2008; doi:10.1152/ajplung.00421.2007
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Airway smooth muscle cell tone amplifies contractile function in the presence of chronic cyclic strain

Nigel J. Fairbank,1,* Sarah C. Connolly,1,* James D. MacKinnon,1 Kathrin Wehry,2 Linhong Deng,3,4 and Geoffrey N. Maksym1

1School of Biomedical Engineering, Dalhousie University, Halifax, Nova Scotia, Canada; 2Department of Medical Engineering, University of Applied Sciences Wilhelmshaven, Wilhelmshaven, Germany; 3National 985 Project, Institute of Biorheology and Gene Regulation, Bioengineering College, Chongqing University, Chongqing, China; and 4Program in Molecular and Integrative Physiological Sciences, Department of Environmental Health, Harvard School of Public Health, Boston, Massachusetts

Submitted 11 October 2007 ; accepted in final form 23 June 2008


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Chronic contractile activation, or tone, in asthma coupled with continuous stretching due to breathing may be involved in altering the contractile function of airway smooth muscle (ASM). Previously, we (11) showed that cytoskeletal remodeling and stiffening responses to acute (2 h) localized stresses were modulated by the level of contractile activation of ASM. Here, we investigated if altered contractility in response to chronic mechanical strain was dependent on repeated modulation of contractile tone. Cultured human ASM cells received 5% cyclic (0.3 Hz), predominantly uniaxial strain for 5 days, with once-daily dosing of either sham, forskolin, carbachol, or histamine to alter tone. Stiffness, contractility (KCl), and "relaxability" (forskolin) were then measured as was cell alignment, myosin light-chain phosphorylation (pMLC), and myosin light-chain kinase (MLCK) content. Cells became aligned and baseline stiffness increased with strain, but repeated lowering of tone inhibited both effects (P < 0.05). Strain also reversed a negative tone-modulation dependence of MLCK, observed in static conditions in agreement with previous reports, with strain and tone together increasing both MLCK and pMLC. Furthermore, contractility increased 176% (SE 59) with repeated tone elevation. These findings indicate that with strain, and not without, repeated tone elevation promoted contractile function through changes in cytoskeletal organization and increased contractile protein. The ability of repeated contractile activation to increase contractility, but only with mechanical stretching, suggests a novel mechanism for increased ASM contractility in asthma and for the role of continuous bronchodilator and corticosteroid therapy in reversing airway hyperresponsiveness.

asthma; mechanical stress; cytoskeletal remodeling; myosin light-chain kinase; optical magnetic twisting cytometry

IN ASTHMA, AIRWAY SMOOTH MUSCLE (ASM) is constantly subjected to external mechanical stress due to breathing as well as internal stress due to chronic and episodic elevations of baseline contraction, or tone. For skeletal muscle, it is known that exercise similarly involves both stretch and muscle activation, which lead to increased muscle mass and altered contractile function. Changes in smooth muscle function have also been reported with obstruction of the bladder or intestine, although the effect of altered tone with stretch is largely unknown (13, 24, 30). Moreover, the effects on ASM of stretch and muscle activation in combination have seldom been examined despite the fact that both are present and likely elevated in asthma.

Elevated ASM tone manifests itself as reversible airway obstruction, which is typically characterized by a bronchodilator-induced increase in forced expiratory volume in 1 s (FEV1). In addition, external ASM stress may be elevated in asthma due to disease-associated wheezing, coughing, etc. In this report, we explore the hypothesis that repeated contractile activation in the presence of continuous cyclic stretching leads to increased contractile function of ASM.

Application of external stress or strain to cultured ASM cells has been shown to elicit a number of procontractile changes in cytoskeletal and contractile function, including increased cytoskeletal remodeling, stiffness, force production, shortening capacity, and contractile enzyme activity and content (12, 43–45, 47). However, little is known about the contribution of tone to strain-induced changes. Cultured ASM cells have an elevated level of contractile tone, evident in a substantial decrease in force production or stiffness to a variety of relaxant agonists (22, 34, 41). Furthermore, tone has been shown to be required for growing and maintaining integrin-dependent cytoskeletal linkages to the ECM (40) as well as to promote and have intrinsic dependence on actin polymerization and remodeling (12, 20, 36). As both internally and externally applied forces are carried by the cytoskeleton and its attachments, tone may be an important determinant of the ASM cell response to strain.

We (11) previously reported that ablation of tone during application of acute (2 h) localized external mechanical stress prevented a twofold increase in cell stiffness and inhibited associated actin remodeling in cultured canine ASM cells. Here, we extended these studies, using human ASM cells to investigate the effects of repeated contractile activation or deactivation on changes in baseline stiffness, contractility, and "relaxability" induced by chronic (5 day) substrate stretch, or strain. Cytoskeletal stiffness was used as a measure of cytoskeletal prestress, which has been shown to be an index of contractile activation in response to acutely delivered muscle agonists or antagonists (22, 34, 55). We also measured the proportion of phosphorylated myosin light-chain (pMLC) to further associate the changes in contractility with the contractile state as well as changes in the expression of rate-limiting contractile enzyme, myosin light-chain kinase (MLCK), which is known to be sensitive to strain. We show that strain-induced increases in contractility were modulated by a history of repeated tone alteration and introduced a positive dependence of contractile function on tone not observed in static culture.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cell culture. Human ASM cells were cultured from macroscopically healthy bronchial tissue explants, each visibly containing one segment of cartilage. The tissue was obtained, by informed consent, from eight patients following lung resection surgery, as approved by the Capital Health Research Ethics Board (Halifax, Nova Scotia, Canada). Each patient was diagnosed with adenocarcinoma but had no history of airway disease. Tissue containing ASM was separated from any surrounding cartilage, gently scraped to remove epithelial cells, and cut into sections approximately 3 x 3 mm. These were placed in a 100-mm plastic tissue-culture Petri dish and compressed slightly using square glass coverslips (VWR; West Chester, PA) anchored in one corner with sterile silicone grease. The tissue sections were incubated for ~2 wk in 10 ml of culture medium (1:1 DMEM/Ham's F-12 mixture supplemented with 10% FBS) with one medium change per week. Resulting adherent cells were then trypsinized and seeded into culture flasks at an average density of 3.2 x 103 cells/cm2. At passage 1, the cells were characterized morphologically by immunostaining for smooth muscle (SM)-{alpha}-actin and by contractility assessment using optical magnetic twisting cytometry (OMTC), described below. Cells were grown to passages 2–3 in culture medium changed every 2–4 days before being used for experimentation.

Experimental protocol. Cells were plated in six-well format culture plates with well bottoms of collagen I-coated silastic membrane (BioFlex culture plate; Flexcell International, Hillsborough, NC) and grown to ~70% confluence. They were then left unstrained for 5 days (static controls) or subjected to 5% linear sinusoidal (0.3 Hz), predominantly uniaxial strain for 5 days in an incubator. Strain was applied with a device (Flexcell Tension Plus System, Flexcell International) that employed computer-controlled vacuum pressure beneath the membranes of the culture plate, causing cyclic downward deflection.

During the period of strain application, the culture medium was refreshed once daily. One day before the cessation of strain, the serum concentration of the medium was reduced to 1% and supplemented with 5.8 µg/ml insulin and 1 µg/ml transferrin. Unstrained control cells were grown on the same membranes and treated in the same manner as the strained cells.

Using changes in cell stiffness measured by OMTC and pMLC (methods described below) to detect changes in contractile activation, we conducted a search from a variety of pharmaceutical agents for those able to alter contractile activation periodically over a 5-day period, for several hours at a time, without exhibiting tachyphylaxis. For instance, contraction-inhibiting agents evaluated included long- and short-acting β-agonists, MLCK inhibitors, phosphodiesterase inhibitors, cAMP analogs, and adenylyl cyclase activators. Consistent behavior was found using forskolin (5 µM) to lower contraction and carbachol (100 µM) or histamine (5 µM) to raise contraction. During the 5-day period of strain application, these agonists and a sham control (identical volume of culture medium used to dilute the agonists) were separately added once daily, following each medium change, to both strained and unstrained cells. Cells from the eight human donors were subjected to the eight resulting experimental conditions (4 agonist treatments including sham, with and without strain). The figure legends indicate the number of donors and minimum beads or repeats where appropriate per assay.

To measure the effects of the above treatments on cytoskeletal stiffness and acute forskolin and KCl responses, pie-shaped portions of membranes were placed, strain-free, in 35-mm dishes and covered with medium from the culture plates, similar to Smith et al. (42). Ferrimagnetic microbeads were bound to the cells for 40 min in an incubator, with two washes, to remove beads not bound to cells, and a replacement of medium with insulin- and transferrin-containing DMEM/F-12 (free of contraction-altering agonists) after 20 min. OMTC was subsequently used to measure baseline cytoskeletal stiffness (details in OMTC) and then the acute (<10 min) change in stiffness due to the addition of either contractile agonist, KCl (80 mM, isoosmotic), or relaxant agonist, forskolin (10 µM). For baseline stiffness, results were obtained from an average of 800 beads and 6 repeated treatments (wells), whereas results for KCl and forskolin responses were from 400 beads and 3 wells. Wherever the cells were aligned as a result of strain, measurements were taken in a direction parallel to the long axes of the cells as described further in DISCUSSION.

To assess the speed of contraction in strained cells, the time constant of KCl-induced stiffness increase was calculated by fitting stiffness vs. time to a step-response function as follows:

Formula 1(1)
where G' is stiffness calculated at an individual bead, as described in OMTC, GFormula 1 is the stiffness before drug addition, GFormula 1 is the plateau stiffness following drug addition, and {tau} is the time constant of rising to the plateau, in seconds.

For measurements of cellular alignment, multiple images per membrane (1–2 membranes per donor) were taken of adherent cells using Hoffman modulation contrast at x20 magnification. These images were analyzed using custom software (details in Measuring cell alignment) to determine the extent of alignment.

For measurements of the effects of strain and alteration of contraction on the abundance of contractile protein, MLCK, membrane-adhered cells from all experimental conditions were fixed and stained and then an in-cell Western was done on MLCK as outlined in Infrared staining and in-cell Western.

To measure myosin activation potential, downstream of MLCK, pMLC was quantified by Western blot following the addition of KCl (80 mM, isoosmotic) to induce acute contraction, as outlined in Western blotting.

OMTC. OMTC is a technique used to measure cytoskeletal stiffness, as described previously (12). Ferrimagnetic microbeads (4.5 µm diameter), kindly provided by Dr. J. J. Fredberg, Harvard School of Public Health, Boston, MA, were coated with a synthetic Arg-Gly-Asp (RGD)-containing peptide (12). These beads bind to surface integrin receptors and the cytoskeleton through the formation of focal contacts. Briefly, OMTC involves magnetizing the beads in the horizontal plane and subsequently applying a vertical, sinusoidally varying (0.5 Hz) external magnetic field that causes the beads to rotate, or twist, back and forth. Cytoskeletal stiffness is measured in hundreds of cells simultaneously by tracking the translational motion of the beads during 30 s of twisting with a CCD camera (SensiCam; Cooke, Auburn Hills, MI). At each bead, a complex modulus is computed in the Fourier domain at the twisting frequency by dividing the known torque applied to the bead, Formula 1, by its displacement, Formula 1:

Formula 2(2)
where G', the real component, is an elastic modulus or stiffness of the cytoskeleton that has units of torque normalized to bead volume, or Pascals per nanometer; G", the imaginary component, is a loss modulus; and i is the unit imaginary number –1.

Infrared staining and in-cell Western. Membranes with adherent cells, which had received tone modulation as well as strain or no strain, were fixed in 3% paraformaldehyde for 15 min at 4°C. They were then permeabilized in 3% paraformaldehyde and 0.3% Triton X-100 for 5 min and stored at 4°C until staining. MLCK was stained by blocking the cells in TBS containing 1% BSA and 2% goat serum for 2 h and then incubating at room temperature for 1 h with MLCK primary antibody (clone K36; Sigma-Aldrich, Oakville, Ontario, Canada) at a 1:100 dilution in TBST (TBS + 0.1% Tween 20). Following washes with TBST, cells were incubated in TBST containing the secondary antibody, IRDye 800CW-conjugated goat anti-mouse IgG (1:1,000; Rockland, Gilbertsville, PA), and TO-PRO-3 iodide (1:1,000; Invitrogen, Burlington, Ontario, Canada), a nuclear dye, for 1 h at room temperature.

Membranes were imaged, from the cell side, on an Odyssey near-infrared imaging system (LI-COR Biosciences, Lincoln, NE) at excitation wavelengths of 800 and 700 nm, constant gain, and resolutions at both wavelengths of 12 bits and 21 µm. This spatial resolution was insufficient to resolve individual cells but more than sufficient to quantify changes in protein quantity across large or small populations of cells. Resulting images demonstrated very low background noise, which is typical of infrared immunofluorescence due to reduced autofluorescence and scattering at these wavelengths compared with visible-light fluorescence. Custom software, written using LabVIEW (National Instruments, Toronto, Ontario, Canada) and MATLAB (The MathWorks, Natick, MA), was used to determine the median intensity of each channel image in concentric rings 1 mm wide, following the removal of pixels with obvious defects (debris or lifting cells due to removal of membranes from 6-well plate and subsequent handling during staining and imaging). MLCK per cell was estimated by dividing the median MLCK intensity by median nuclear intensity, which was proportional to cell number (verified by manual cell counts, not shown) and measured in the same region.

Western blotting. Membranes with adherent cells, from all experimental conditions, were rinsed on ice with 80 mM KCl in TBS, following 10-min contractile stimulation in 80 mM KCl. Cell scrapers were used to transfer the cells into tubes, which were centrifuged for 15 min at 18,000 g. Supernatant was then removed, and the pellets were diluted with 2x Laemmli sample buffer containing 50 mM dithiothreitol, 1% protease inhibitor cocktail, and 1% phosphatase inhibitor cocktail 2 (all from Sigma). Samples were vortexed for 90 min at 4°C followed by 3 min of centrifugation at 10,000 g. Cellular extracts were then heated for 4 min at 97°C.

Proteins were separated by electrophoresis on acrylamide gradient mini-gels (NuPAGE Bis-Tris Gels) in MES running buffer using the XCell SureLock apparatus at 200 V for 50 min (all products from Invitrogen). Following electrophoresis, proteins were transferred overnight to pure nitrocellulose membranes presoaked in 1x transfer buffer (Invitrogen) containing 20% methanol using the XCell blot module (Invitrogen) at 25 V.

Membranes were then blocked in 2% blocking buffer (Bio-Rad, Mississauga, Ontario, Canada) in TBST for 1 h at room temperature and subsequently incubated for 1 h at room temperature with rabbit anti-phospho-MLC (1:1,000; detects phosphorylation of serine 19, the major phosphorylation site and preferred site for MLCK) and mouse anti-β-actin (1:20,000; clone AC-15) primary antibodies (both from Sigma). All antibodies were diluted in 2% blocking buffer in TBST. Following primary incubation, the resulting blots were washed repeatedly in TBST and then labeled by 1-h incubation at room temperature with infrared secondary antibodies IRDye 700DX-conjugated donkey anti-mouse IgG (1:1,000) and IRDye 800-conjugated donkey anti-rabbit IgG (1:20,000), both from Rockland. After repeated washes with TBST, proteins were detected using the Odyssey near-infrared imaging system (LI-COR Biosciences) at excitation wavelengths of 700 nm and 800 nm. Tagged antibodies were then removed by incubation in stripping buffer (62.5 mM Tris·HCl with 2% SDS, pH 6.8) at 50°C for 15 min. Primary and secondary antibody binding and detection was subsequently repeated following the above protocol using mouse anti-MLC primary (1:1,000; clone MY-21 from Sigma) in place of anti-phospho-MLC. Quantification was performed using Quantity-One image software (Bio-Rad).

Measuring cell alignment. Custom software was used to determine the orientation of the long axes of cells from images acquired at known points on silastic membranes. On a given image, the software randomly placed 80 dots, which marked the cells to be measured. For each of these cells, a line was drawn from end to end of the cell, and the angle (0–180°) between this line and a radial vector, which indicated the predominant strain direction, was determined automatically.

The extent of alignment was quantified by angular dispersion, where high alignment corresponds to low dispersion. Dispersion was calculated as the mean angular distance, or absolute difference in degrees, from the mean cell angle.

Statistics. As stiffness measurements were distributed lognormally, the effects of mechanical strain and tone history on baseline cell stiffness, as well as acute changes in stiffness due to KCl and forskolin, were examined by logarithmic transformation followed by two-way ANOVA. Post hoc comparisons were done by t-test with Bonferroni correction. The effect of strain and tone history on pMLC was examined in the same way, without transformation. Kruskal-Wallis nonparametric one-way ANOVA was used, with Bonferroni correction, for time constants of contraction, MLCK, pMLC (yielding the same results as the above test), and cell angle dispersion. This method of examining dispersion was suggested by Wallraff (52).

Reagents. All chemicals were purchased from Sigma-Aldrich unless otherwise stated. Tissue culture reagents, including DMEM/F-12 medium and trypsin, were purchased from Invitrogen.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Strain amplified the range of baseline stiffness induced by contractile modulation. We found that baseline stiffness following 5 days of repeated application of forskolin, carbachol, or histamine was rank-ordered according to the acute change in stiffness that could be induced with these agents. Cells that had been treated with forskolin demonstrated the lowest stiffness, and those that had been treated with histamine, the highest stiffness. This rank-ordering was observed in both unstrained and strained cells but was amplified in strained cells (P < 0.01; Fig. 1). In all but forskolin-treated cells, strain also increased baseline stiffness, raising it by 0.30 (SE 0.08), 0.25 (SE 0.06), and 0.38 (SE 0.08) Pa/nm in sham-, carbachol-, and histamine-treated cells, respectively, compared with unstrained cells with the same treatment. A significant interaction effect between strain and history of contractile activation on stiffness was found by two-way ANOVA (P < 0.001), meaning strain had a significantly greater effect on stiffness in the presence of repeated contractile activation. All of the above measurements, and those presented below, were made after the final day of strain application and concurrent contractile modulation, following washout of contraction-altering agents as described in MATERIALS AND METHODS.


Figure 1
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  		Fig. 1. Baseline cell stiffness and contraction/relaxation responses following 5 days with strain (Strain) or without (Static) in medium containing either forskolin (F; 5 µM), sham (S), carbachol (C; 100 µM), or histamine (H; 5 µM). Following strain, tone-altering agonists were removed, baseline stiffness was measured, and then the change in stiffness due to acute KCl (80 mM, isoosmotic)-induced contraction or acute forskolin (10 µM)-induced relaxation was measured. See RESULTS for description of significant differences. Contractile (Cont.) scope is the range in stiffness from plateau relaxation to plateau contraction. Values are medians ± SE (donors = 3, beads > 300) normalized to the baseline of sham-treated, strained cells.

 
Strain-enhanced contractility was modified by contractile activation history. The plateau stiffness following acute KCl-induced contraction, or the contractile ceiling, was in all cases raised by strain (P < 0.001; Fig. 1). Furthermore, strain introduced a positive dependence on the history of contractile activation that was not observed in unstrained (static) cells. For instance, whereas plateau stiffness increased by 101.6% (SE 17.9) in cells that had received histamine pretreatment, it increased by only 29.6% (SE 8.4) in those that received forskolin pretreatment (Fig. 1). Strain also introduced in all groups a rank-ordering of the contractile floor with history of contractile activation, where the contractile floor is the plateau stiffness following acute addition of forskolin.

The contractile scope in response to acute changes in contractile activation was considered to be the range from the contractile floor to the contractile ceiling, as described by An et al. (3). Strain increased the contractile scope in rank-order with history of contractile activation (Fig. 1; P < 0.001). In the presence of strain, contractile scope was also highly correlated with baseline stiffness (R2 = 0.98), whereas there was no correlation in unstrained cells.

Like the contractile scope, KCl responses were uncorrelated with history of contractile activation in static cells (Fig. 2). Strain introduced a positive correlation between KCl contractile response and baseline stiffness (R2 = 0.90), with the greatest increase, 176.0% (SE 58.6; P < 0.001), occurring in cells that had been incubated with histamine. Strain also led to a greater KCl response in cells that had been incubated with forskolin, but the increase was smaller at 43.7% (SE 19.9; P < 0.05). As with baseline stiffness, a significant interaction effect between strain and history of contractile activation on KCl responses was found (P < 0.05).


Figure 2
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  		Fig. 2. Increase in stiffness due to acute KCl (80 mM, isoosmotic)-induced contraction following strain or no strain for 5 days. Letters indicate tone-modulating agents as in Fig. 1. In static cells, baseline stiffness was greater (rightward shift) when tone had been elevated. However, contractility was not correlated with tone. In strained cells, there was a strong dependence of both baseline stiffness and contractility on chronic tone modulation, with contractility increasing (shifting upward) substantially in high-tone cells (P < 0.001). Values are medians ± SE. *P < 0.01 compared with static sham, $P < 0.05 compared with strain sham.

 
In addition, the speed of contraction, i.e., KCl-induced increase in stiffness to its plateau value, was greater in cells cultured with a history of increased contractile activation (Fig. 3) than those cultured with repeated relaxation, as indicated by a lesser time constant (P < 0.01). This suggests an enhanced contractile efficiency in the presence of strain and elevated contraction.


Figure 3
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  		Fig. 3. Time constant of acute KCl-induced contraction in strained cells. Cells with tone contracted significantly faster, indicated by a lower time constant, than those with tone reduced by forskolin during strain (P < 0.01). Inset: a representative graph of cell stiffness vs. time for cells that had previously undergone chronic incubation with forskolin or histamine; KCl was added 60 s after stiffness measurement began. Values are medians ± SE. *P < 0.01 compared with sham.

 
MLCK decreased with strain by 22.4% (SE 3.9; P < 0.05) in forskolin-treated cells, relative to forskolin-treated unstrained cells (data not shown). MLCK was statistically unchanged by strain with sham treatment [although increasing numerically by 12.3% (SE 2.0)], and increased significantly with strain with carbachol and histamine treatment. The greatest strain-induced increase in MLCK, 55.4% (SE 13.7; P < 0.01), occurred with histamine treatment, demonstrating a positive dependence of MLCK on strain, history of contractile activation, and baseline stiffness as further described in DISCUSSION. Furthermore, whereas MLCK in unstrained cells was inversely dependent on history of contractile activation, decreasing by 18.4% (SE 3.8; P < 0.05) with histamine treatment and increasing by 24.2% (SE 4.7; P < 0.05) with forskolin treatment relative to sham (see DISCUSSION), MLCK in strained cells showed a strong positive correlation with baseline stiffness (Fig. 4).


Figure 4
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  		Fig. 4. Myosin light-chain kinase (MLCK) content measured in strained cells showing that cells that were subjected to chronic tone elevation contained more MLCK than those subjected to tone reduction with forskolin (MLCK content in unstrained cells can be found in Fig. 7). Inset: a representative false-gray image of infrared-labeled MLCK in cells adhered to a membrane portion (bar = 1 mm; mean cell length, ~140 µm). Values are means ± SE (donors = 4) normalized to sham-treated cells. *P < 0.05 compared with forskolin.

 
The increased MLCK, relative to unstrained controls, observed with strain and increased contractile activation corresponded to increased myosin activity following acute KCl (80 mM)-induced contraction (P < 0.05), as demonstrated by the level of pMLC (Fig. 5). In cells pretreated with histamine, strain induced a 103.6% (SE 23.3) increase in KCl-stimulated pMLC, above that in unstrained cells, whereas this increase was completely absent in cells pretreated with forskolin. A significant interaction effect between strain and history of contractile activation on pMLC was found by two-way ANOVA (P < 0.01).


Figure 5
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  		Fig. 5. A: cellular content of phosphorylated/total MLC vs. baseline stiffness, following chronic incubation with forskolin ({circ}, bullet), sham ({square}, {blacksquare}), carbachol ({triangleup}, {blacktriangleup}), and histamine ({lozenge}, {blacklozenge}) and then agonist washout and acute contraction with KCl (80 mM). Unstrained cells are indicated by open symbols and strained cells by closed symbols. B: sample Western blot of phosphorylated (Ser19) MLC and total MLC for all treatments (F, H, C, and S, as in Fig. 1) in unstrained and strained cells. Values are means ± SE (donors = 5, lanes = 2) normalized to strain sham. *P < 0.05 compared with static sham, $P < 0.05 compared with strain sham. Inset: strain-induced increases (Inc.) in MLCK and MLC phosphorylation (pMLC) were approximately proportional (R2 = 0.98).

 
Repeated reduction of contractile activation decreased strain-induced cell alignment. With strain, all groups became more aligned perpendicular to the principal direction of applied strain (Fig. 6; P < 0.001). Unstrained (static) cells had no preferred orientation, as indicated by visual inspection as well as a measured angular dispersion matching that predicted by a uniformly random distribution of angles. However, whereas all strained cells became aligned, cells pretreated with forskolin aligned significantly less than all other groups (P < 0.001).


Figure 6
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  		Fig. 6. Extent of cell alignment, quantified as dispersion of angle, {theta}, illustrated in inset. Static cells of all treatments (F, S, C, and H, as in Fig. 1) had no preferred orientation, indicated by a high dispersion, equal to that of a uniform distribution (Dist'n). However, all strained cells tended to align perpendicular to the direction of strain, with mean {theta} close to 90°. Alignment of cells that had tone reduced (F) was less than that of cells that had unaltered or elevated tone (P < 0.001), with the latter cells demonstrating strong alignment and correspondingly low angular dispersion. Values are mean angular distance from mean {theta} ± SD (donors = 5, measured angles > 400). *P < 0.001 compared with strain sham.

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
We found that chronic uniaxial strain was a potent stimulus for increased ASM stiffness, cellular organization, and contractility. Furthermore, these properties exhibited dependence on repeated modulation of tone by pharmacological stimulation, revealing that repeated contraction amplified the maladaptive effects of strain. In the following, we compare our principal data with previous findings and then discuss the physiological significance.

Magnetic microbeads have been used extensively to probe cytoskeletal mechanics and changes in cellular contractility (10, 12, 34), with results in close agreement with those obtained by other methods, such as atomic force microscopy (1, 12, 15, 38) and traction microscopy (55). Stiffness data acquired by OMTC are highly heterogeneous, but it has been shown that statistically reliable values and significant differences can be obtained relatively easily with a small number of experimental repetitions due to the large number of cells probed simultaneously in each measurement (15).

The strain-induced increases in stiffness and contractility we observed (Figs. 1–3) are consistent with previous studies that also reported increases in stiffness and contractile function due to long-term application of cyclic stretch (42, 44, 46, 47). However, in the absence of repeated contractile activation, we did not observe a significant increase in MLCK as had been reported earlier (44). A number of factors could account for the difference. We employed a shorter period of strain, i.e., 5 vs. 14 days, and we also, importantly, employed human ASM cells rather than canine as well as a smaller, more physiological strain amplitude of 5% vs. 10% (2). Here, we report the above findings in human ASM cells for the first time and investigate the effects of repeated alteration of contractile tone.

We repeatedly altered the contractile state of the ASM cell using forskolin to decrease activation, or carbachol or histamine to increase activation, relative to sham controls. However, these agonists have a number of nonspecific effects, via a variety of cell signaling pathways, on a variety of cell functions. For instance, forskolin and downstream cAMP have been shown to inhibit proliferation (26), alter histamine H1 and β2-adrenergic receptor function (32, 37), and induce an array of changes in gene expression, largely mediated by the transcription factor, cAMP response element binding protein (58). Carbachol and histamine have been shown in ASM to potentiate mitogen-induced proliferation (27) as well as alter adenylyl cyclase activity and cAMP formation (7, 14), whereas carbachol incubation has been observed to increase inflammatory gene transcription and augment transcription induced by length oscillation (25). Thus some of the altered contractility observed here may be due to complex indirect effects other than induced changes in the contractile state. Although we do not know the precise mechanisms that led to altered contractile function, each of the agonists employed is known to directly modulate force development and alter cytoskeletal tension as we observed here, via changes in cell stiffness.

Indeed, stiffness is directly dependent on the state of contractile activation and is a measure of cytoskeletal tension, which is often termed prestress (49, 55). Prestress has been shown to be linearly related to both stiffness, as measured by OMTC, and the traction force applied by the cell to its substrate (55). Moreover, stiffness and contractile force, in turn, are established to be directly modulated in concert with changes in contractile activation (49). We show here that repeated stimulation to activate or deactivate the contractile apparatus, in the presence and absence of strain, led to changes in baseline stiffness and likely prestress, which was maximal with strain and contractile activation (Figs. 1 and 2). Furthermore, strain and repeated contractile stimulation altered KCl-induced contraction, measured by pMLC and induced increase in cytoskeletal stiffness, that were matched by changes in baseline stiffness. This implies that changes in contractility were coupled with changes in baseline prestress, indicating change in the overall contractile ability of the cell in the 5-day strain period. Although the changes in contractility could have arisen from direct or indirect effects of agonist stimulation, it is nevertheless likely that they reflect an alteration of contractile phenotype. This is supported by the fact that a change in contractile phenotype can occur with strain alone (44). To explore the possibly that the changes in contractility observed with stretch and tone were accompanied by a change in contractile phenotype, we determined whether MLCK, a marker of contractile phenotype and function, was altered similarly to contractility.

Negative modulation of MLCK by tone in static culture. We found that MLCK content was decreased in static culture with increases in tone (Fig. 7), but this was dramatically reversed with the application of mechanical strain, as we discuss further below. A negative regulation of contractility with sustained contraction is consistent with previous reports in static culture. For instance, using organ-cultured bovine tracheal smooth muscle strips, Gosens et al. (18) reported that 8-day incubation with methacholine caused a concentration-dependent reduction in contractile responses to receptor-dependent and -independent agonists as well as concentration-dependent decreases in SM-{alpha}-actin and SM-myosin heavy-chain protein expression. Additionally, the magnitudes of the changes in contractile proteins they found are comparable and in agreement with the MLCK decrease in static culture we report here.


Figure 7
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  		Fig. 7. Cellular content of MLCK and degree of strain-induced alignment (quantified as angular dispersion) vs. baseline stiffness, following chronic incubation with forskolin ({circ}, bullet), sham ({square}, {blacksquare}), carbachol ({triangleup}, {blacktriangleup}), and histamine ({lozenge}, {blacklozenge}). Unstrained cells are indicated by open symbols, strained cells by closed symbols, and alignment data by an adjacent letter "a". MLCK in static cells exhibited a negative correlation with baseline stiffness, indicated by linear fit (dotted line), whereas in strained cells, MLCK and alignment exhibited a positive correlation with baseline stiffness (dashed and solid lines, respectively). Values are means ± SE normalized to strain sham, and statistical differences are provided in RESULTS. Mean angular dispersion of static cells (data not shown) was 3.91 (SE 0.07).

 
Such decreases in contractility were shown by Gosens et al. (18) to be dependent on chronically elevated intracellular calcium, as had been demonstrated previously in organ-cultured rat tail artery (29) and guinea pig ileum (17). Conversely, increases in contractility have been shown in rat tail arterial rings cultured with reduced-calcium media (28). This also is in agreement with the increase in MLCK we found in static culture following incubation with forskolin, which induces large increases in cAMP and, likely, cAMP-dependent decreases in intracellular calcium (4).

However, all of the above studies were conducted in the absence of physiological cyclic strain, which we have found to significantly alter contractile behavior and its history of contractile activation dependence, as discussed below.

Positive modulation of MLCK by tone in mechanically strained culture. The observed reversal of MLCK dependence on history of tone modulation in the presence of strain is novel (Figs. 4 and 7) but is supported by Wahl et al. (51), who observed that a cyclic stretch-dependent decrease in SM-{alpha}-actin mRNA in bovine tracheal strips could be prevented when the strips were activated with carbachol or histamine. Such a strain-induced reversal in MLCK tone dependence may be important due to the central role MLCK plays in smooth muscle contraction, as well as some evidence that it may be altered in asthma, as we discuss further below. Increases in MLCK in cultured ASM cells, such as we observed with strain and repeated contractile activation, have also been shown to directly contribute to increases in velocity and extent of shortening, calcium sensitivity, and force production (44–47).

During acute contraction, actomyosin force generation is dependent on the phosphorylation of MLC, which is largely achieved by calcium-calmodulin-dependent activation of MLCK (48) and opposed by MLC phosphatase (MLCP). Indeed, in the presence of strain, we observed like changes in MLCK and pMLC due to a history of tone modulation (Figs. 4 and 5), and these mirrored contraction-induced changes in stiffness (Fig. 2). Changes in MLCK and pMLC from the unstrained condition were also highly correlated (Fig. 5, inset). Although this points to the fact that changes in MLCK protein levels drove changes in pMLC, it is also possible and perhaps quite likely that other regulatory pathways of MLC phosphorylation contributed to these changes, such as a strain-induced decrease in MLCP activity or increase in MLCK activity, both of which have been observed previously in canine ASM (47). Nonetheless, with strain, MLCK, pMLC, and KCl-induced increase in stiffness changed along with baseline stiffness, or prestress, illustrating a tight coupling of the regulation of the contractile ability of the cell with its baseline tension, as well as indicating a shift to a more contractile phenotype. Furthermore, strain-induced increases in pMLC have previously been shown to be accompanied by increased activation of actomyosin ATPase (47), suggesting high prestress and contractility due to the combination of strain and repeated contractile activation are a result of an increased number of attached actomyosin cross bridges.

It is possible that ECM proteins, both the collagen I coating of the membranes and proteins deposited by the cells themselves, played a role in the strain dependence and tone dependence of contractility. ECM proteins can alter ASM phenotype and affect the responses of ASM to other stimuli, such as the proliferative response to mitogen stimulation (21) in the presence and absence of strain (8). In addition, laminin promotes a more contractile ASM phenotype than collagen and fibronectin (21). Although little is known about how cyclic strain may affect the deposition of ECM proteins by ASM, tenascin and matrix metalloproteinases are known to be increased (19). It has also been shown in vascular smooth muscle cell culture that ECM coating can modulate strain-induced expression of SM-myosin (39). Since stress due to mechanical strain is transmitted to the cytoskeleton via focal adhesions, which attach to specific ECM motifs, it is likely that initial ECM protein content and ASM ECM deposition play a role in the regulation of strain- and tone-dependent changes in ASM contractility.

Cytoskeletal organization. Organization of the ASM cell, which reorients perpendicular to the direction of mechanical strain, has been found to play an important role in strain-induced changes in contractile function (42, 46). Here, we report changes in stiffness and response to KCl in a direction parallel with aligned actin fibers. We (42) have previously shown that stiffness measured by OMTC in the parallel direction is 24% (SE 5) greater than the perpendicular direction. Thus some of the increase in baseline stiffness observed in this study may be due to increased cytoskeletal alignment but likely not all. As strained cells were stiffer than unstrained cells in the absence of changes in contractile stimulation, and pMLC increased collinearly with MLCK and cytoskeletal stiffness, this implies a strong interdependence between changes in contractile protein content and cellular organization, which is also suggested by the interaction effects we found. Indeed, perhaps regulation of MLCK content, contractile scope, and range of MLC phosphorylation may depend on collinear arrangement of the contractile machinery.

Increased baseline stiffness and contractility could also be a result of increased cytoskeletal fiber formation through contraction-dependent actin polymerization. Actin polymerization and contractile activation are highly codependent. Activation by contractile agonist is known to increase actin polymerization and focal adhesion formation, at least acutely (11, 36). Thus a similar mechanism may explain the increased baseline stiffness with repeated contractile activation in either strained or unstrained cells, as contractile activation generally led to an increase in the contractile floor, which was the lowest stiffness achieved with maximal relaxation (Fig. 1). Similarly, repeatedly decreasing contractile activation over 5 days may have led to reduced actin polymerization. Relaxation with a variety of agonists for 2 h is known to inhibit mechanically induced stiffening and associated cytoskeletal and focal adhesion organization (5, 11, 16). Although this is not established in response to chronic strain, periodic relaxation did reduce cytoskeletal alignment relative to all other groups (Fig. 6).

Thus our study revealed a strong relationship between contractility, MLCK, alignment, and prestress (baseline stiffness) in cells subjected to chronic strain. This can be seen in Fig. 7, where the data have been normalized to those from strained cells cultured in medium treated with tone-unaltering sham. What can also be seen in this figure is that changes in cell stiffness, and thus cytoskeletal tension, induced by modulating contractile activation may be analogous to changes in cytoskeletal tension induced by cyclic stretching. Looking at the data from strained cells (solid symbols in Fig. 7), a move from left to right along either the solid or dashed line of best-fit corresponds to a history of greater activation, resulting in increased MLCK, alignment, and cytoskeletal tension. Similarly, a move from unstrained to strained cells with the same history of contractile activation (open to solid symbol of the same shape) corresponds to an increase in MLCK, alignment, and stiffness for all agonist treatments except forskolin. For example, moving from histamine-treated unstrained cells to histamine-treated strained cells demonstrates the greatest strain-induced increase in MLCK of 55.4%, whereas forskolin-treated cells demonstrated a strain-induced decrease in MLCK and no change in baseline stiffness accompanying the addition of strain. This implies that, when cytoskeletal tension changes, the resulting effects on cytoskeletal organization and contractility are similar whether the tension change is due to altered mechanical strain or altered contractile activation history.

How do mechanical strain and changes in contractile activation signal changes in cell structure and function? Both strain and contractile activation involve the application of force through the cytoskeleton. Balaban et al. (5) and others (16) have suggested the cytoskeleton of the living cell has a stress "set-point" that is maintained by cytoskeletal adaptation. They report that with added internal force (tone) or added external force (strain) the cytoskeleton increases focal adhesion size and attachments to maintain a constant stress, or force per unit of cross-sectional area through which the force is transmitted. Conversely, when tone or strain decreases, the cell responds by decreasing focal adhesion size and cytoskeletal attachments. This is reflected in our data with either reduced strain or reduced tone by a decrease in cell stiffness and MLCK, of which the latter may occur with decreases in actin-myosin bundling.

However, although regulation of cytoskeletal attachments has been demonstrated in response to acute stress, it has not yet, to our knowledge, been demonstrated in response to chronic strain applied to the entire cell. Furthermore, in contrast to the increased stiffness and contractility due to continuous cyclic stretch, the acute ASM response to a single, or a few, large stretches is now established to be a transient and large decrease in stiffness, interpreted as "fluidization" of the cytoskeleton (50). We do not know the mechanisms for the difference in response to acute vs. long-term stretching. Although we did not measure acute responses to substrate stretch, our data suggest that over longer durations of stretch–of days rather than seconds, i.e., on the time scale of altered protein expression–there may be something of a set-point phenomenon similar to that described for local perturbation of the cytoskeleton. This model appears to break down in the absence of mechanical strain, where the dependence of MLCK on the history of contractile activation, or at least the specific agonist effects on tone, is reversed (Fig. 7, dotted line). It may be, for instance, that the increase in MLCK that resulted from forskolin administration and reduced stretch (static culture) was an attempt by the cell to increase cytoskeletal stress to the set-point through increasing MLCK-dependent contractile force. In this case, we may speculate that the associated decrease in stiffness indicates a failure to achieve normal physiological cytoskeletal stress.

Strain profile. In contrast with the uniaxial strain applied in this study, equibiaxial strain does not induce orientation in fibroblasts or ASM cells (42, 53, 54) or promote procontractile changes (54). Approximately equibiaxial strain is thought to be the predominant form of strain in the healthy lung as airways both dilate and lengthen by similar amounts during lung inflation (23). Furthermore, its potentially anticontractile influence is supported in vivo by a decrease in acetylcholine response of ex vivo tissue from the lungs of rabbits that had been ventilated with continuous positive airway pressure for 4 days (57). However, in the presence of tone or heterogeneous airway remodeling such as occurs in asthma, the airways are not as likely to experience equibiaxial strain but rather more uniaxial or anisotropic strain. We (33) have suggested, and others have observed (9), that with airway constriction due to tone or narrowing in asthma, airway dilation is reduced relative to lengthening, indicating strain anisotropy. Since ASM is predominantly circumferentially oriented around the airways, stretch due to breathing under a condition of elevated tone would become more uniaxial and would more closely resemble strain oriented transverse to the cell long axes, as employed in this study. If this is true, it implies that biaxial stretch may be protective, in agreement with the aforementioned biaxial studies in vitro and in vivo, whereas uniaxial stretching promotes enhanced contractility and airway hyperresponsiveness.

Physiological significance. Our results suggest that stretch and tone may work together to promote increased contractility through cytoskeletal reorganization and increases in MLCK. Although we have no data on cytoskeletal organization in asthma, currently there are conflicting findings on changes in MLCK with asthma. MLCK is present in the lung in both a low and a high molecular-weight isoform; one study reported no difference in the low molecular-weight isoform between asthmatic and healthy tissues (35). However, they did not report on the other isoform. Another in vivo study reported that total MLCK protein is increased in asthma (6), whereas data on MLCK mRNA from bronchial biopsies are equivocal, with reports of either an increase (31) or no change (56). In any case, our data indicate that increases in MLCK depend on the history of contractile stimulation, which suggests increases in MLCK might occur in vivo in asthma when therapy has been ineffective at reducing ASM activation.

We report an ability of ASM to respond to repeated increases in contractile activation or tone with increased stiffness and contractility. Furthermore, this ability was highly dependent on mechanical strain. If present together in vivo, strain and elevated tone could promote airway hyperresponsiveness, airway elasticity, and impairment of dilation, which are present in asthma. This report also suggests a novel mechanism for the benefits of current therapy on ASM remodeling. Long-acting bronchodilators and corticosteroids act to directly and indirectly decrease ASM tone, which, even in the presence of stretch due to breathing, may reduce procontractile phenotypic modulation of the ASM, leading to reduced airway hyperresponsiveness.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This study was supported by the Canadian Institutes of Health Research. N. J. Fairbank is supported by a Canada Graduate Scholarship from the Natural Sciences and Engineering Research Council of Canada. N. J. Fairbank and S. C. Connolly are supported by the Canadian Institutes of Health Research Strategic Training Fellowship STP-53877. S. C. Connolly is supported by the Nova Scotia Health Research Foundation.


   ACKNOWLEDGMENTS
 
We thank Drs. Drew Bethune and Zhaolin Xu of The Queen Elizabeth II Health Sciences Centre, Halifax, Nova Scotia, Canada, for kindly providing tissue; Dr. Jeffrey Fredberg of the Harvard School of Public Health, Boston, MA, for supplying the ferrimagnetic beads and for helpful comments; Drs. Reinoud Gosens and Per Hellstrand for their helpful comments; and Darren Cole for help with cell preparation.


   FOOTNOTES
 

Address for reprint requests and other correspondence: G. N. Maksym, School of Biomedical Engineering, Dalhousie Univ., 5981 Univ. Ave., Halifax, Nova Scotia B3H 1W2, Canada (e-mail: gmaksym{at}dal.ca)

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.

* N. J. Fairbank and S. C. Connolly contributed equally to this work. Back


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