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MS 9270 Received 15 February 1999; accepted after revision 31 May 1999.
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ABSTRACT |
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 TopAbstract IntroductionMethodsResultsDiscussionReferences |
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INTRODUCTION |
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Actin is a ubiquitous protein that is required for both non-muscle cell motility and muscle cell contraction. Dynamic actin polymerization to filamentous actin (F-actin) and depolymerization to monomeric actin (G-actin) is a fundamental mechanism of cell motility (Southwick & Stossel, 1983). In contrast, striated muscle contains a stable pool of F-actin, and contraction is mediated by the sliding of actin filaments along myosin. This process is mediated by cyclic attachment and detachment of the myosin head to actin (i.e. cross-bridge cycling), and the hydrolysis of adenosine triphosphate (ATP) by actin-activated, myosin ATPase (actomyosin ATPase). The sliding filament cross-bridge model derived from striated muscle is also thought to be applicable to smooth muscle contraction, although the cytoarchitecture of smooth muscle is not well defined (Somlyo et al. 1983; Somlyo, 1985). Evidence supporting the sliding filament cross-bridge model in smooth muscle includes the characteristic length-isometric force and isometric force-velocity relationships (Murphy, 1989), and the identification of both thick and thin filaments (Somlyo et al. 1983). Additionally, studies have identified dense bodies and membrane-associated dense plaques, which are thought to anchor the actin filaments to other cytoskeleton proteins, thereby coupling actomyosin ATPase activity to myocyte contraction and shortening (Pavalko et al. 1995).
Recent biochemical and mechanical studies of smooth muscle suggest that actin filaments may not be stable as in skeletal muscle, and that dynamic actin reorganization involving actin polymerization and depolymerization may play an important functional role in smooth muscle (Adler et al. 1983; Mauss et al. 1989; Boels & Pfitzer, 1992; Obara & Yabu, 1994). For example, cytochalasins B and D, which inhibit actin polymerization (Cooper, 1987), also reduce isometric force (Adler et al. 1983; Cooper, 1987; Mauss et al. 1989; Obara & Yabu, 1994; Tseng et al. 1997). These findings are consistent with previous studies of non-muscle motile cells, which showed that cytochalasins inhibit cell motility (Southwick & Stossel, 1983) and numerous other cellular processes, such as ion channel activity (Berdiev et al. 1996). Additionally, unlike skeletal muscle, the relationship between smooth muscle cell length and isometric force is not unique, as this relationship varies depending on the prior history of mechanical stretch (Gunst, 1986; Harris & Warshaw, 1991; Gunst et al. 1995).
A fundamental functional characteristic of smooth muscle is that during isometric activation, the rate of ATP hydrolysis by actomyosin ATPase (Kerrick & Hoar, 1994) and maximum unloaded shortening velocity (Vmax) (Dillon et al. 1981; Hai & Murphy, 1988a, b; Jiang & Stephens, 1990), an index of cross-bridge cycling rate, each rapidly reach maximal values but then decline to sustained suprabasal levels even though the increase in isometric force is sustained. Thus, there is a time-dependent decline in tension cost (i.e. ATP hydrolysis rate per unit of isometric force) during steady-state isometric force. Although the underlying mechanism is not fully known, some studies suggest that this decline in tension cost is due to a transition from phosphorylated, rapidly cycling cross-bridges to dephosphorylated, slowly cycling cross-bridges (Hai & Murphy, 1988a, b). It is possible that dynamic actin filament reorganization may also be responsible for the decline in tension, since brevin, a protein that regulates actin gel-sol transformation, increases Vmax and, hence, cross-bridge cycling rate without increasing actomyosin ATPase activity in freshly dissociated, permeabilized smooth muscle cells (Gailly et al. 1990, 1991). These authors postulated that the increase in Vmax was due to a reduction in internal viscous resistance to movement, since brevin also caused a concomitant decrease in cytoplasmic stiffness (Gailly et al. 1991). Accordingly, dynamic actin filament reorganization may account for the time-dependent decrease in tension cost throughout steady-state isometric force in smooth muscle, since cross-bridge cycling rate is load dependent, as in skeletal muscle (Jiang & Stephens, 1990).
The purpose of the current study was to examine the effect of F-actin stabilization on tension cost in airway smooth muscle. We hypothesized that inhibition of F-actin depolymerization by phalloidin increases tension cost during Ca2+-induced contraction of canine tracheal smooth muscle permeabilized with Triton X-100.
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METHODS |
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Experimental techniques
Tissue preparation. All experiments were performed in accordance with the guidelines established by our Institutional Animal Care and Use Committee. Mongrel dogs (15-23 kg) of either sex were anaesthetized with an intravenous injection of pentobarbital (30 mg kg-1) and exsanguinated. A 10-15 cm portion of extrathoracic trachea was excised and immersed in chilled physiological salt solution (PSS) of the following composition (mM): 110·5 NaCl, 25·7 NaHCO3, 5·6 dextrose, 3·4 KCl, 2·4 CaCl2, 1·2 KH2PO4 and 0·8 MgSO4. Fat, connective tissue and the epithelium were removed with tissue forceps and scissors. Thin strips of tracheal smooth muscle (width, 0·1-0·2 mm; length, 6-9 mm; wet weight, 60-135 µg) were teased from the sheet of tissue using a dissecting microscope.
For experiments using rat diaphragm tissue sections, the rats were killed by I.M. injection of 60 mg kg-1 ketamine and 2·5 mg kg-1 xylozine.
Smooth muscle mechanics. Isometric force, stiffness and the ATP hydrolysis rate were measured simultaneously in muscle strips using a commercially available mechanics and photometric system (Scientific Instruments, Heidelberg, Germany; model PH2) (Kerrick et al. 1990). Maximum unloaded shortening velocity (Vmax) was also measured using this system but in a separate set of tissues obtained from different dogs.
The strips were mounted in a 10 µl quartz cuvette and continuously superfused at 2 ml min-1 with PSS (37°C) aerated with 96 % O2 and 4 % CO2. One end of the strips was anchored via stainless steel microforceps to a calibrated force transducer (resolution, 0·01 mN) and the other end via stainless steel microforceps to a servo-controlled stepper motor (resolution, 0·001 mm). The attachments to the force transducer and stepper motor were not compliant. During a 2 h equilibration period, the length of the muscle strips was incrementally increased after repeated isometric contractions (of 2-3 min duration) induced by 1 µM acetylcholine (ACh) until maximal isometric force (optimal length, Lo) was obtained. The strips were maintained at Lo for the remainder of the experiment. The strips were then cooled to 25°C for 30 min and permeabilized with Triton X-100 (see 'Permeabilization procedure and solutions'). We have previously determined that the contractions of permeabilized canine tracheal smooth muscle induced by 10 µM free Ca2+ are more stable at 25°C than at 37°C. The cross-sectional area was calculated for each strip as the product of the tissue length and weight (g). Isometric force was expressed as newtons per cross-sectional area of tissue (N cm-2).
Tissue stiffness was measured by imposing sinusoidal length oscillations of 0·5 % Lo amplitude at a frequency of 50 Hz. Preliminary studies established that the relationship between the frequency of length oscillation and stiffness was sigmoidal with maximal stiffness at 50 Hz and a phase angle between isometric force and length of less than 10 %. Tissue stiffness was normalized to cross-sectional area and expressed as Young's modulus (N cm-2 Lo-1).
Isotonic shortening velocities under various loads were determined by the quick-release, afterload-clamp method (Jiang & Stephens, 1990). A series of afterloads from 2 to 50 % of maximal isometric force was rapidly (within 2 ms) imposed on the muscle strips. Isotonic shortening velocities (expressed as muscle lengths per second (Lo s-1)) were determined from the length change during a 100-150 ms period beginning 50 ms after release to avoid effects of series elastic recoil. To calculate Vmax, the force-velocity relationship was fitted by a non-linear, least squares method to the Hill equation, and velocity was extrapolated to zero load.
ATP hydrolysis rate measurements. The rate of ATP hydrolysis by permeabilized strips was measured using an enzyme-coupled reduced 
-nicotinamide adenine dinucleotide (NADH) fluorometric technique in which the regeneration of ATP from ADP and phosphoenol pyruvate was catalysed by pyruvate kinase (Kerrick et al. 1990). This reaction was coupled to the oxidation of NADH to NAD+ and to the reduction of pyruvate to lactate; these reactions were catalysed by lactate dehydrogenase. For each mole of ADP produced, 1 mole of NADH, a fluorescent compound, was oxidized to NAD+, a non-fluorescent compound. Thus, the rate of decrease in NADH fluorescence was proportional to the rate of ATP hydrolysis by the tissue.
Light from a xenon lamp was monochromatically filtered to restrict excitation light to 340 nm wavelength and focused by a high-numerical aperture objective onto the quartz cuvette. Fluorescence emitted by the solution in the cuvette was filtered at 500 ± 5 nm and detected by a photomultiplier assembly. Illumination intensity of the excitation light was detected by an absorbance monitor to correct for fluctuations in excitation light intensity.
The cuvette was rapidly flushed for 100 ms every 5 s with fresh solution (
70 µl) containing the constituents necessary to couple ATP hydrolysis to NADH oxidation. Flushing the cuvette with fresh solution caused an abrupt increase in NADH fluorescence (Fig. 1). The rate of decline in NADH fluorescence during a 4·9 s period was measured. NADH fluorescence was determined for known concentrations of NADH (90 and 180 µM) immediately prior to each experiment, so that the amount of NADH oxidized during the 4·9 s period could be calculated and used to quantify the rate of ATP hydrolysis. As shown in Fig. 1, the rate of decline in NADH fluorescence increased when the strip was activated with 10 µM free Ca2+, indicating an increase in ATP hydrolysis rate. ATP hydrolysis rates were normalized for tissue volume and expressed as nanomoles per centimetre cubed per second.
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The rate of decline in NADH fluorescence during a 4·9 s period was measured and used to quantify the ATP hydrolysis rate based on a prior calibration of the system with known NADH concentrations. ATP hydrolysis rates were normalized for tissue volume and expressed as nanomoles per centimetre cubed per second. | ||
Permeabilization procedure and solutions. Muscle strips were either superfused (isometric force, stiffness, ATP hydrolysis and Vmax measurements) or incubated (fluorescence microscopy and regulatory myosin light chain (rMLC) phosphorylation measurements) for 20 min with relaxing solution containing 10 % (v/v) Triton X-100 (25°C). The composition of the relaxing solution was as follows (mM): 85 KCl, 2·1 disodium ATP (Na2ATP), 4 EGTA, 20 imidazole and 1 dithiothreitol (DTT). After permeabilization, the strips were washed with relaxing solution for 15 min to remove excess Triton X-100. Solutions of varying free ion concentrations were prepared using a previously described computer-generated algorithm (Donaldson & Kerrick, 1975) and also contained 1 µM calmodulin. The pH of all solutions was buffered to 7·1 with proprionic acid and the ionic strength was kept constant at 0·150 M by adjusting the concentration of potassium proprionate. In addition to these constituents, solutions for ATP hydrolysis rate measurements also contained 5 mM phosphoenol pyruvate, 0·18 mM NADH, 140 U ml-1 lactate dehydrogenase and 100 U ml-1 pyruvate kinase.
rMLC phosphorylation measurements. rMLC phosphorylation was measured in separate muscle strips, prepared from a different set of dogs, by gel electrophoresis followed by Western blot analysis (Gunst et al. 1994). After an equilibration period of 15 min in aerated PSS at 25°C, the strips were incubated in nominally Ca2+-free PSS containing 2 mM EGTA for 15 min. Then intracellular Ca2+ stores were depleted by adding 10 µM ACh to this solution for 10 min. ACh was removed by exchanging solutions repeatedly with nominally Ca2+-free PSS over 15 min before tissues were permeabilized with Triton X-100. After experimental interventions, the permeabilized strips (width, 0·1-0·2 mm; length, 1·0-1·5 cm; wet weight, 85-150 µg) were flash-frozen by rapid immersion in dry ice-acetone slurry containing 10 % (w/v) trichloroacetic acid and 10 mM DTT (-80°C). Then the frozen strips were thawed to 25°C, washed in acetone containing 10 mM DTT to remove the trichloroacetic acid, and transferred to 200 µl Eppendorf tubes containing 65 µl of extraction buffer (7·3 M urea, 20 mM Tris, 21 mM glycine and 10 mM DTT). Proteins were separated by glycerol-urea polyacrylamide gel (10 % (w/v) acrylamide, 0·5 % (w/v) bisacrylamide, 40 % (v/v) glycerol, 20 mM Tris and 21 mM glycine) electrophoresis. The electrophoresis buffer contained 20 mM Tris, 21 mM glycine, 1 mM DTT and 1 mM sodium thioglycolate. The gels were subjected to pre-electrophoresis for 1 h at 400 V (10°C) to remove urea, and to allow DTT and thioglycolate to enter the gels. Then 50 µl of sample was injected into the wells, and initially subjected to electrophoresis for 1 h at 100 V and then for 17 h at 400 V (10°C).
For Western blot analysis, the proteins were transferred to nitrocellulose sheets (0·22 µm) for 4 h at 1·6 A (15°C) in 25 mM Na2HPO4 (pH 7·6). The nitrocellulose sheets were washed twice with 10 mM Tris-buffered saline (TBS) containing 5 % (w/v) bovine serum albumin for 1 h (25°C) before labelling with polyclonal, affinity-purified, rabbit anti-rMLC antibody (Gunst et al. 1994). The anti-rMLC antibody was detected with 125I-labelled protein A (Du Pont, Boston, MA, USA).
The unphosphorylated and phosphorylated bands of rMLC were visualized by phosphoimage analysis (PhosphoImager, Molecular Dynamics, Inc., Sunnyvale, CA, USA) and quantified by ImageQuaNT software (Molecular Dynamics, Inc.). After local background subtraction, rMLC phosphorylation was calculated by integrating the bands corresponding to the mono- and diphosphorylated rMLC as a fraction of the total integration of both the phosphorylated and unphosphorylated rMLC.
Fluorescence microscopy. Strips were fixed with 4 % (w/v) paraformaldehyde in 10 mM TBS for 20 min and then washed three times with TBS to remove excess fixative. After the strips had been embedded, 10 µm-thick transverse sections were prepared, mounted on glass slides and permeabilized with 1 % (v/v) Triton X-100 in TBS for 20 min. The sections were first treated with TBS containing 5 % (w/v) bovine serum albumin (blocking buffer) for 20 min, and then co-labelled (25°C) with 3 µg ml-1 tetramethylrhodamine-conjugated DNase I (rhodamine-DNase I) and 5 U ml-1 Alexa-488-conjugated phalloidin (Alexa-488-phalloidin) in blocking buffer for 20 min. After washing the sections three times with TBS, a coverslip was mounted on the slide using ProLong antifade reagent (Molecular Probes) to reduce photobleaching.
Localization of G- and F-actin was determined using a Zeiss LSM510 confocal fluorescence microscope (Carl Zeiss, Inc., Oberkochen, Germany). Fluorescence from Alexa-488-phalloidin and rhodamine-DNase I was measured simultaneously. The excitation and emission maxima for Alexa-488-phalloidin were 492 and 518 nm, respectively, and for rhodamine-DNase I were 555 and 580 nm, respectively. Fluorescence from each fluorophore was distinguished by both the laser excitation line and the emission optical filters. An argon-krypton laser produced excitation lines of 488 and 568 nm. The images were acquired using a × 100 oil immersion lens with a numerical aperture of 1·4. The emission fluorescence wavelengths were restricted by a 505-550 nm bandpass filter and a 585 nm longpass filter. Prior to each experiment, the excitation line intensity and photomultiplier gain were adjusted to eliminate possible contamination of the rhodamine-DNase I fluorescence by the fluorescence of Alexa-488-phalloidin (and vice versa).
Experimental protocols
Four experimental protocols were conducted, each on canine tracheal smooth muscle strips obtained from a different set of dogs. For each protocol, permeabilized strips were superfused (isometric force, stiffness, ATP hydrolysis and Vmax measurements) or incubated (fluorescence microscopy and rMLC phosphorylation measurements) for 1 h either with relaxing solution containing 1 % methanol (control) or with relaxing solution containing 50 µM phalloidin in 1 % methanol. Then all strips were washed with relaxing solution containing 1 µM calmodulin for 10 min to remove the methanol and phalloidin, and allow the tissues to equilibrate with calmodulin. All subsequent solutions also contained 1 µM calmodulin. For those protocols in which ATP hydrolysis rate was measured, all solutions also contained the appropriate enzymes (see 'Permeabilization procedure and solutions').
Effect of phalloidin on the time course of isometric force, stiffness and ATP hydrolysis rate. This experimental protocol determined whether phalloidin altered the time-dependent relationship between isometric force, stiffness and ATP hydrolysis rate, during maximal activation by 10 µM free Ca2+. Isometric force, stiffness and ATP hydrolysis rate were measured simultaneously in individual muscle strips. The strips were superfused with 10 µM free Ca2+ for 30 min and then with relaxing solution for an additional 5 min to allow isometric force, stiffness and the ATP hydrolysis rate to return to unstimulated baseline values.
Effect of phalloidin on the time course of rMLC phosphorylation. To measure the time course of rMLC phosphorylation, two sets of six muscle strips were placed in 5 ml polyethylene vials and permeabilized. One set was incubated with 1 % methanol to provide control rMLC phosphorylation measurements and the second set was incubated with 50 µM phalloidin in 1 % methanol. One strip from each set was flash-frozen prior to activation for baseline rMLC phosphorylation measurements. The remaining five strips from each set were incubated with 10 µM free Ca2+ and then flash-frozen for rMLC phosphorylation measurements at 1, 2, 5, 10 and 30 min following activation.
Effect of phalloidin on Vmax. Permeabilized muscle strips were maximally activated with 10 µM free Ca2+ for 30 min as previously described. Then, simultaneous with a rapid (< 2 ms) length step, afterloads (P) equal to 2, 5, 10, 20 or 50 % of maximal isometric force (Po) were imposed on each muscle strip in random order. These quick-release measurements were performed during a single contraction, since preliminary studies demonstrated that repeated isometric contractions induced by 10 µM free Ca2+ were not reproducible. Preliminary studies also demonstrated that the multiple quick-release measurements of P/Po of 0·05 performed during a single contraction yielded reproducible shortening velocity measurements.
Localization of G- and F-actin. In the first set of experiments we determined the specificity of rhodamine-DNase I labelling of G-actin by comparing the extent of labelling between tissue sections of rat diaphragm, which does not contain G-actin, and canine tracheal smooth muscle. In a second set of experiments, we determined whether both G- and F-actin are present in the permeabilized canine tracheal smooth muscle. Paired sections of rat diaphragm and intact or permeabilized canine tracheal smooth muscle were mounted on slides. One section of each pair was co-labelled with 3 µg ml-1 rhodamine-DNase I and 5 U ml-1 Alexa- 488-phalloidin. The second section of each pair was not labelled and was used to determine the level of background autofluorescence at a given excitation line intensity and photomultiplier gain.
Materials
The polyclonal affinity-purified rabbit anti-rMLC antibody was a generous gift of Susan J. Gunst, Ph.D., Department of Physiology and Biophysics, Indiana University School of Medicine, Indianapolis, IN, USA. Na2ATP was purchased from Research Organics, Inc. (Cleveland, OH, USA). Protein A was purchased from Du Pont. Alexa-488-phalloidin, rhodamine-DNase I, and the ProLong antifade kits were purchased from Molecular Probes, Inc. Stock solutions of the fluorescent probes were prepared in methanol. Phalloidin was purchased from Calbiochem (La Jolla, CA, USA) and was dissolved in methanol. The final concentration of methanol in all experimental solutions was 1 %; at this concentration, methanol had no effect on the increases in isometric force, stiffness, ATP hydrolysis rate or rMLC phosphorylation induced by free Ca2+ (data not shown). Trichloroacetic acid was purchased from Fisher Scientific (Fair Lawn, NJ, USA). Pyruvate kinase and lactate dehydrogenase were purchased from Boehringer Mannheim (Indianapolis, IN, USA). Calcium oxide and magnesium oxide were purchased from Aldrich Chemical Company, Inc. (Milwaukee, WI, USA). All other drugs and chemicals were purchased from Sigma. Unless otherwise specified, all drugs and chemicals were dissolved in distilled water.
Statistical analysis
Data are expressed as mean values ± standard error of the mean (S.E.M.); n represents the number of dogs. All comparisons were made by Student's unpaired t test. A P value < 0·05 was considered statistically significant.
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RESULTS |
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Effect of phalloidin on the time course of isometric force, stiffness and ATP hydrolysis rate
In permeabilized strips incubated with methanol (control), increasing the free Ca2+ concentration from 1 nM to 10 µM induced sustained increases in isometric force (Fig. 2, upper panel) and stiffness (Fig. 2, middle panel). By contrast, the increase in ATP hydrolysis rate (Fig. 2, lower panel) was initially high, reaching a maximal value at 1-2 min following activation, but thereafter gradually declined to a sustained suprabasal level at 10 min that was sustained throughout activation, as the ATP hydrolysis rates measured at 10 and 25 min were not significantly different (Table 1). Decreasing the free Ca2+ concentration from 10 µM to 1 nM induced complete relaxation, which was preceded by a decline in ATP hydrolysis rate to a level similar to that of the unstimulated strips.
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Permeabilized strips were superfused with relaxing solution (see Methods for composition) containing 1 % methanol or 50 µM phalloidin in 1 % methanol for 1 h prior to activation. In preliminary studies, 1 % methanol had no effect on the increases in isometric force (upper panel), stiffness (middle panel) or ATP hydrolysis rate (lower panel) induced by 10 µM free Ca2+. Values are means ± S.E.M. using data obtained from 7 animals. | ||
Incubating the permeabilized strips with phalloidin had no significant effect on baseline isometric force, stiffness or ATP hydrolysis rate (Table 1). Likewise, there was no significant difference in initial isometric force or stiffness development (measured at peak ATP hydrolysis rate) between strips incubated with or without phalloidin. However, in contrast to tissues not incubated with phalloidin, the increases in isometric force and stiffness induced by 10 µM free Ca2+ were not sustained (Fig. 2, upper and middle panel, respectively). Isometric force and stiffness reached maximal levels at 2-3 min following activation, but then gradually declined throughout the remaining 27-28 min of activation (Table 1). Phalloidin had no effect on the relationship between isometric force and stiffness (Fig. 3).
Table 1. Effect of phalloidin on isometric force, stiffness, ATP hydrolysis rate and tension cost
| Isometric force (N cm-2) |
Stiffness (N cm-2 Lo-1) |
ATP hydrolysis rate (nmol cm-3 s-1) |
Tension cost (nmol N-1 cm-1 s-1) |
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| Methanol | Phalloidin | Methanol | Phalloidin | Methanol | Phalloidin | Methanol | Phalloidin | |
| Baseline | 3·2 ± 0·8 | 3·1 ± 0·4 | 94 ± 29 | 91 ± 18 | 18·2 ± 1·7 | 23·2 ± 2·9 | 6·8 ± 1·1 | 7·8 ± 1·1 |
Peak  |
14·2 ± 2·3 | 14·5 ± 0·8 | 879 ± 157 | 873 ± 74 | 47·7 ± 4·3 | 71·4 ± 9·4 * | 3·3 ± 0·4 | 4·9 ± 0·6 * |
| 10 min | 17·0 ± 2·1 | 12·6 ± 0·6 * | 1115 ± 156 | 726 ± 67 * | 30·3 ± 3·7 | 54·1 ± 6·8 * | 1·9 ± 0·2 | 4·3 ± 0·5 * |
| 25 min | 16·4 ± 2·0 | 10·2 ± 0·5 * |
1050 ± 138 | 553 ± 49 * |
28·8 ± 2·8 | 51·5 ± 7·0 * | 1·8 ± 0·2 | 5·0 ± 0·6 * |
Values measured at the time of peak ATP hydrolysis rate. * Significantly different from peak values measured in permeabilized strips incubated with methanol alone (control). Significantly different from values measured 10 min after activation with 10 µM free Ca2+.
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Permeabilized strips were superfused with relaxing solution (see Methods for composition) containing 1 % methanol or 50 µM phalloidin in 1 % methanol for 1 h prior to activation. These relationships were constructed from the data shown in Fig. 2, upper and middle panels. Values are means using data obtained from 7 animals. Error bars have been omitted for the purpose of clarity. | ||
Incubation of permeabilized strips with phalloidin had no effect on the time course of the increase in ATP hydrolysis rate induced by 10 µM free Ca2+. As in the control tissues, the suprabasal ATP hydrolysis rate was not sustained, reaching peak values within 1-2 min, but then declining to steady-state, suprabasal levels by 10 min following activation (Fig. 2, lower panel). There was no significant difference in the ATP hydrolysis rate measured at 10 and 25 min. The magnitudes of both the peak and steady-state ATP hydrolysis rates were significantly greater in strips incubated with phalloidin compared with control strips. However, the magnitudes of the decline in ATP hydrolysis rate from peak to steady-state levels measured at 25 min following activation were similar (Table 1).
The time course for changes in tension cost (ATP hydrolysis rate divided by isometric force) is shown in Fig. 4. In permeabilized strips incubated with methanol only, tension cost declined throughout activation with 10 µM free Ca2+. By contrast, in strips incubated with phalloidin, activation caused an initial decrease in tension cost that reached a nadir by 7-10 min, but then gradually increased throughout activation; the tension cost measured at 25 min was significantly greater than that measured at 10 min following activation. Thus, after an initial decline, tension cost increased throughout activation. Tension cost was significantly greater in permeabilized strips incubated with phalloidin throughout activation compared with control strips (Table 1).
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Permeabilized strips were superfused with relaxing solution (see Methods for composition) containing 1 % methanol or 50 µM phalloidin in 1 % methanol for 1 h prior to activation. Tension cost was calculated as ATP hydrolysis rate (nmol cm-3 s-1) divided by isometric force (N cm-2). Values are means ± S.E.M. using data obtained from 7 animals | ||
Effect of phalloidin on the time course for rMLC phosphorylation
Similar to the pattern observed in measurements of isometric force and stiffness, 10 µM free Ca2+ induced a sustained increase in rMLC phosphorylation (Fig. 5). There was no significant difference in baseline rMLC phosphorylation between permeabilized strips incubated with methanol or phalloidin in methanol. Likewise, incubation with phalloidin did not affect the time course or magnitude of the increase in rMLC phosphorylation induced by 10 µM free Ca2+ (Fig. 5).
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Permeabilized strips were incubated in relaxing solution (see Methods for composition) containing 1 % methanol or 50 µM phalloidin in 1 % methanol for 1 h prior to activation. Values are means and S.E.M. using data obtained from 5 animals. | ||
Effect of phalloidin on Vmax
There was no significant difference in Vmax (Fig. 6), measured 30 min after activation with 10 µM free Ca2+, between permeabilized strips incubated with methanol or phalloidin in methanol (0·07 ± 0·01 Lo s-1 and 0·05 ± 0·01 Lo s-1, respectively).
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Vmax in Triton X-100-permeabilized canine tracheal smooth muscle strips maximally activated by 10 µM free Ca2+ Permeabilized strips were incubated in relaxing solution (see Methods for composition) containing 1 % methanol (upper panel) or 50 µM phalloidin in 1 % methanol (lower panel) for 1 h prior to activation. The data were fitted with the hyperbolic Hill equation (P + a)(V + b) = (Po + a)b, where a and b are the asymptotes of the hyperbola. Values are means ± S.E.M. using data obtained from 5 animals. | ||
Localization of F- and G-actin
Figure 7 shows rat diaphragm (upper panels) and canine tracheal smooth muscle (lower panels) co-labelled with rhodamine-DNase I and Alexa-488-phalloidin. Whereas F-actin was detected in both tissue types, G-actin was detected only in the canine tracheal smooth muscle. These data demonstrate that the rhodamine-DNase I labelling was specific for G-actin. Figure 8 is an example of fluorescence images of the G- and F-actin in intact canine tracheal smooth muscle strips incubated in PSS and permeabilized strips incubated in relaxing solution. The major finding was that appreciable G-actin was detected in both the intact and the permeabilized canine tracheal smooth muscle strips.
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The G-actin was labelled with rhodamine-DNase I and imaged using confocal fluorescence microscopy. The image intensity gain was determined using an unlabelled tissue section to eliminate tissue autofluorescence and was kept constant throughout the experiment. | ||
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The G- and F-actin were labelled with rhodamine-DNase I and Alexa-488-phalloidin, respectively, and visualized using confocal fluorescence microscopy. The image intensity gain was determined for each channel using an unlabelled tissue section to eliminate tissue autofluorescence and was kept constant throughout the experiment. | ||
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DISCUSSION |
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The major findings of this study were that incubation of permeabilized canine tracheal smooth muscle with phalloidin, an agent known to stabilize F-actin filaments, resulted in decreased isometric force and stiffness, increased actomyosin ATPase activity, and no significant change in rMLC phosphorylation during maximal Ca2+ activation. Thus, incubation with phalloidin increased tension cost. These results are the first to demonstrate that dynamic actin filament reorganization may be necessary for optimal energy utilization and the persistent decline in tension cost observed in smooth muscle during Ca2+ activation.
Actin is a cellular protein that is essential for muscle contraction and the motility of non-muscle cells. In non-muscle cells, actin polymerization and depolymerization play a central role in cell motility (Southwick & Stossel, 1983). These cells contain significant amounts of both G-actin and F-actin, and the rate of protrusion or propulsion is inversely proportional to the half-life of F-actin (Theriot, 1994). Thus, cell motility is predicated on rapid dynamic actin reorganization. Smooth muscle, while similar to striated muscle in that contraction requires actin- myosin interaction, does not contain a morphologically distinct sarcomeric structure (Somlyo et al. 1983). Additionally, whereas skeletal muscle contains stable actin filaments and little G-actin, 25 % or more of the actin in unactivated smooth muscle is in the monomeric form (Tseng et al. 1997). A recent study of cultured human airway smooth muscle cells demonstrated that activation with carbachol induced a marked quantitative transition from G- to F-actin and an increase in the number of stress fibres, indicating that actin filaments can be dynamically regulated by a contractile agonist (Togashi et al. 1998). In another study, the ratio of F-actin to G-actin in vascular smooth muscle cells under relaxing conditions was significantly decreased by incubation with cytochalasin B, a compound that inhibits actin polymerization, suggesting that transitions from F-actin to G-actin take place even in resting smooth muscle (Wang et al. 1996). Smooth muscle contractions are also inhibited by cytochalasins (Adler et al. 1983; Obara & Yabu, 1994), suggesting that dynamic actin reorganization is involved in force development (Cooper, 1987; Tseng et al. 1997). ADP-ribosylation of monomeric actin, which decreases actin polymerization induced by botulinum C2 toxin (Aktories et al. 1986), inhibited electromechanical coupling in longitudinal guinea-pig ileum smooth muscle (Mauss et al. 1989). Taken together, these results provide evidence that dynamic actin reorganization occurs in smooth muscle, that interference with this process can inhibit smooth muscle contraction, and that dynamic actin reorganization may be modulated by contractile agonists.
Another molecular probe that has been used to investigate the role of dynamic actin reorganization in motile and smooth muscle cell function is the bicyclic heptapeptide phalloidin. Phalloidin binds specifically to F-actin (Faulstich et al. 1988) and both increases the stability of F-actin and markedly decreases the G-actin concentration at which polymerization to F-actin occurs, thereby reducing the level of G-actin (Dancker et al. 1975; Wehland et al. 1977; Estes et al. 1981). Stabilization of F-actin and a reduction in guanidine hydrochloride-extractable G-actin has been reported in Triton X-100-permeabilized vascular smooth muscle incubated with phalloidin (Boels & Pfitzer, 1992). Unlike the cytochalasins, phalloidin has not been widely used as a functional probe for dynamic actin reorganization in tissue, because of inefficient uptake by cells (Cooper, 1987). However, microinjection of phalloidin into motile, non-muscle cells induced actin polymerization and interfered with cell locomotion, which suggested that motility in some cells is contingent upon both polymerization and depolymerization of actin (Wehland et al. 1977).
A fundamental functional characteristic of smooth muscle is that during activation, tension cost declines during sustained isometric force (Krisanda & Paul, 1988; Kuhn et al. 1990), a process that may be attributable to the development of dephosphorylated, slowly cycling actomyosin cross-bridges (Hai & Murphy, 1988a, b). However, it is possible that dynamic actin filament reorganization also contributes to this decline in tension cost during sustained isometric force. This assertion is supported by studies of permeabilized taenia coli smooth muscle, which was treated with brevin, an endogenous protein that severs actin filaments into shorter ones. During activation with Ca2+, brevin treatment caused an increase in Vmax and, hence, actomyosin cross-bridge cycling rate (Gailly et al. 1990), and a decrease in cytoplasmic stiffness in the absence of actomyosin cross-bridge formation (Gailly et al. 1991). These data suggest that dynamic actin reorganization may be an important determinant of internal resistance to movement, cross-bridge cycling rate and, hence, tension cost.
To examine this possibility, the current study determined the effect of F-actin stabilization with phalloidin on the time-dependent change in tension cost induced by free Ca2+ in Triton X-100-permeabilized canine tracheal smooth muscle. The fluorescence imaging studies (Figs 7 and 8) documented that both G- and F-actin are present in this tissue before and after permeabilization with Triton X-100, thereby establishing this preparation as suitable for examining dynamic actin reorganization. The predominant source of suprabasal ATP hydrolysis during Ca2+ activation in this preparation is the actomyosin ATPase (Kuhn et al. 1990; Kerrick & Hoar, 1994; Zhang et al. 1994). For example, the calmodulin antagonist W7 (Zhang et al. 1994) and the MLC kinase (MLCK) inhibitor wortmannin (Jones et al. 1999) both inhibit the increase in isometric force and ATP hydrolysis rate induced by free Ca2+ in Triton X-100-permeabilized smooth muscle. By contrast, neither basal nor the Ca2+-activated, suprabasal ATP hydrolysis rate are affected by inhibitors of the sarcoplasmic reticulum, mitochondria or plasma membrane Ca2+-ATPases, or the Na+-K+-ATPase (Kuhn et al. 1990; Zhang et al. 1994). Also, activation of permeabilized canine tracheal smooth muscle by irreversible, maximal rMLC thiophosphorylation (which induces isometric force without activating MLCK) results in peak and steady-state suprabasal ATP hydrolysis rates that are not significantly different from that induced by maximal activation with free Ca2+ (which activates both MLCK and actomyosin ATPase). Taken together, these data indicate that the suprabasal ATP hydrolysis rate requires rMLC phosphorylation by MLCK and the formation of force-generating, actomyosin cross-bridges. This preparation also eliminates possible effects of dynamic actin reorganization on intracellular Ca2+ homeostasis and rMLC phosphorylation (Tseng et al. 1997), and allows phalloidin to freely diffuse into the cell, as intact cells are essentially impermeable to phalloidin (Faulstich et al. 1988).
In the current study, free Ca2+ induced sustained increases in isometric force and rMLC phosphorylation, and a non-sustained increase in ATP hydrolysis rate. Thus, tension cost decreased throughout sustained isometric force. These findings are consistent with previous studies of permeabilized smooth muscle (Zhang et al. 1994). By contrast, in those permeabilized strips incubated with phalloidin, the increase in isometric force induced by free Ca2+ was not sustained. The initial isometric force developed by the phalloidin-treated strips was the same as that produced by the untreated control strips, suggesting that dynamic actin reorganization is not involved in the initial development of isometric force but may be important in the maintenance of isometric force. These findings corroborate those of a previous study of permeabilized vascular smooth muscle (Boels & Pfitzer, 1992). These changes in isometric force in phalloidin-treated strips were accompanied by an increase in peak and steady-state ATP hydrolysis rate compared with control strips, such that tension cost initially declined but then gradually increased throughout activation. The net effect of phalloidin treatment was to cause an increase in actomyosin ATPase activity even as isometric force declined. Thus, there was a persistent increase in tension cost throughout activation that was due to both an increase in ATP hydrolysis rate by actomyosin ATPase and a decline in isometric force.
One possible explanation for the increase in ATP hydrolysis rate by phalloidin is that MLCK, which is bound to F-actin, is activated. However, incubation with phalloidin did not increase basal ATP hydrolysis rate or basal rMLC phosphorylation. Additionally, there was no effect of phalloidin treatment on the rate and extent of rMLC phosphorylation following activation with free Ca2+. These data are consistent with the previous finding that phalloidin had no effect on either MLCK or smooth muscle protein phosphatase activities determined using extracts from vascular smooth muscle (Boels & Pfitzer, 1992). Taken together, these data suggest that the increase in ATP hydrolysis rate by phalloidin was not due to increased MLCK activity and rMLC phosphorylation.
A second possibility is that phalloidin binds at or near the myosin-binding domain at the actin surface, thereby directly perturbing cross-bridge attachment and cycling. There are conflicting results concerning this possibility. Some studies of striated muscle have shown that phalloidin has no effect on actomyosin ATPase activity measured in vitro (Dancker et al. 1975). It has also been reported that phalloidin has no effect on isometric force induced by Ca2+ in permeabilized skeletal muscle fibres (Prochniewicz-Nakayama et al. 1983). However, other studies suggest that phalloidin may increase both isometric force and actomyosin ATPase activity in permeabilized skeletal (Bukatina et al. 1996) and cardiac (Bukatina & Fuchs, 1994; Bukatina et al. 1995) muscle fibres. Whereas the isometric force and actomyosin ATPase activity measurements were not concurrent in these studies (Bukatina & Fuchs, 1994; Bukatina et al. 1995, 1996), the data suggest that tension cost was either decreased or unchanged by phalloidin. Given that in these tissues, actin is regulated by Ca2+-modulated, actin-associated proteins and that phalloidin has no effect on unregulated actin (Dancker et al. 1975), it is possible that the reported effects in striated muscle are due to an interaction between phalloidin and actin-associated proteins (e.g. troponin C), rather than an effect of phalloidin on the myosin binding domain of actin. In addition, the reported effects of phalloidin on striated muscle are the opposite of those reported in the current study of permeabilized airway smooth muscle. Thus, it is unlikely that phalloidin directly affects the actomyosin cross-bridges in smooth muscle, since the primary mechanism of Ca2+ regulation of smooth muscle contraction is thick filament phosphorylation, which was not affected in the current study.
There are additional observations that argue against a direct effect of phalloidin on the actomyosin cross-bridge in smooth muscle. First, the initial isometric force developed in phalloidin-treated and control strips was similar, a finding consistent with that reported for permeabilized vascular smooth muscle (Boels & Pfitzer, 1992). Second, phalloidin treatment does not affect rigor force in permeabilized vascular smooth muscle (Boels & Pfitzer, 1992). It is also unlikely that the increase in ATP hydrolysis rate was caused by activation of the ATPase associated with dynamic actin polymerization and depolymerization, since this ATPase is completely inhibited by phalloidin (Dancker et al. 1975).
The mechanism for the decline in isometric force during constant activation with free Ca2+ in phalloidin-treated strips is not clear. According to the Brenner analytic model of muscle contraction (Brenner, 1985), increased actomyosin ATPase activity reflects an increase in either the number of force-producing cross-bridges (cross-bridge recruitment) or the cross-bridge cycling rate. It is unlikely that the increase in ATP hydrolysis rate was due to the former, since treatment with phalloidin resulted in a decrease in isometric force and had no effect on the relationship between isometric force and stiffness. Consequently, the increase in ATP hydrolysis rate must be due to an increase in actomyosin cross-bridge cycling rate, which should be reflected in an increase in Vmax. However, there was no significant difference in Vmax between control strips and those incubated with phalloidin.
Previous studies of smooth muscle mechanics have shown that the relationship between smooth muscle cell length and isometric force is not unique, as this relationship varies depending on the prior history of mechanical stretch (Gunst, 1986; Harris & Warshaw, 1991; Gunst et al. 1995). There is recent evidence to suggest that this plasticity of mechanical responses may result from dynamic cytoarchitectural reorganization (Harris & Warshaw, 1991; Gunst et al. 1995). It has been proposed that this reorganization process could be mediated by a dynamic interaction between F-actin and cytoskeleton-associated proteins (Gunst et al. 1995; Pavalko et al. 1995). While there is very little direct structural evidence for dynamic cytoskeleton reorganization in smooth muscle (Togashi et al. 1998), actin filaments are known to be attached to dense bodies and dense plaques (Somlyo et al. 1983; Pavalko et al. 1995). Such attachments have been described as conferring sarcomere-like structure on actin and myosin in smooth muscle (Somlyo et al. 1983) and play a critical role in transducing cross-bridge cycling into isometric force development and shortening of the myocyte. By preventing dynamic actin reorganization, phalloidin may have caused the formation of actomyosin cross-bridges that were 'uncoupled' from the cytoskeleton and, thus, less able to transduce actomyosin ATPase activity and cross-bridge cycling into isometric force. Myosin associated with F-actin that is less structurally able to couple actomyosin ATPase activity with the cytoskeleton may 'unload' the actomyosin cross-bridges, and result in an increase in actomyosin ATPase activity (cross-bridge cycling rate) that is not reflected by a change in Vmax. This possibility is supported by the findings of the current study, which showed that phalloidin treatment reduced the ability of the tissue to maintain isometric force and stiffness, and increased the actomyosin ATPase activity without affecting Vmax. Reconciling the phalloidin-stabilizing effect on F-actin with these findings could suggest that during activation of airway smooth muscle, F-actin either detaches or is initially detached from cytoskeletal proteins and cannot form a new attachment to the cytoskeleton, possibly due to inhibition of F-actin depolymerization. The finding of the current study that phalloidin treatment had no effect on the initial isometric force lends support to this conjecture. The finding that the actomyosin ATPase activity was already significantly greater in phalloidin-treated airway smooth muscle at peak isometric force is, however, difficult to reconcile with this model.
In conclusion, phalloidin, an agent that both stabilizes F-actin and decreases G-actin concentration, results in increased tension cost, thereby implicating dynamic actin reorganization in the time-dependent decrease in tension cost normally observed during activation of airway smooth muscle.
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This study was supported in part by research RO1 grants HL-54757 and HL-45532, and PO1 grant HL33009 from the National Institutes of Health. We thank Mrs K. Street for her expert technical assistance.
Corresponding author
K. A. Jones: Mayo Clinic, 200 First Street SW, Rochester, MN 55905, USA.
Email: kjones{at}mayo.edu
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