Influence of calcium on myosin thick filament formation in intact airway smooth muscle

Ana M. Herrera1,3, Kuo-Hsing Kuo1,3, and Chun Y. Seow1,2,3

1 Department of Anatomy, 2 Department of Pathology and Laboratory Medicine, and 3 The University of British Columbia McDonald Research Laboratories/The iCapture Center, St. Paul's Hospital/Providence Health Care, University of British Columbia, Vancouver, British Columbia, Canada V6T 1Z3


    ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Myosin thick filaments have been shown to be structurally labile in intact smooth muscles. Although the mechanism of thick filament assembly/disassembly for purified myosins in solution has been well described, regulation of thick filament formation in intact muscle is still poorly understood. The present study investigates the effect of resting calcium level on thick filament maintenance in intact airway smooth muscle and on thick filament formation during activation. Cross-sectional density of the thick filaments measured electron microscopically showed that the density increased substantially (144%) when the muscle was activated. The abundance of filamentous myosins in relaxed muscle was calcium sensitive; in the absence of calcium (with EGTA), the filament density deceased by 35%. Length oscillation imposed on the muscle under zero-calcium conditions produced no further reduction in the density. Isometric force and filament density recovered fully after reincubation of the muscle in normal physiological saline. The results suggest that in airway smooth muscle, filamentous myosins exist in equilibrium with monomeric myosins; muscle activation favors filament formation, and the resting calcium level is crucial for preservation of the filaments in the relaxed state.

electron microscopy; muscle contraction; ethylene glycol-bis(beta -aminoethyl ether)-N,N,N',N'-tetraacetic acid; muscle plasticity


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

IT HAS BEEN SUGGESTED that myosin thick filaments in relaxed smooth muscle are partially dissolved and reaggregate upon activation (2, 3, 14, 15). Functional studies of airway smooth muscle have shown that thick filament lengthening could account for the increase in isometric force and the decrease in shortening velocity during the sustained phase of contraction (9). It was observed in airway smooth muscle that within at least a threefold length range and when the muscle was allowed to adapt to the lengths at which it was studied, isometric force was independent of muscle length, and shortening velocity and power output were positively correlated to muscle length (7). The observations suggest that the number of in-series contractile units (of which myosin thick filaments are an essential component) could be variable. We observed in a recent study that the thick filaments in intact airway smooth muscle dissolved partially when subjected to mechanical perturbation (5). There are, therefore, many lines of evidence supporting a notion that myosin thick filaments in smooth muscle are labile; partial disassembly/reassembly of the thick filaments could allow the muscle to quickly adapt to externally imposed mechanical strains that reshape the cell. As a first step toward understanding what regulates cellular plasticity in terms of thick filament formation, the present study was carried out to examine myosin evanescence during activation and to determine the stability of the thick filaments in relaxed airway smooth muscle depleted of calcium and subjected to mechanical perturbation.


    METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Tissue Preparation

Porcine tracheal smooth muscle obtained from a local abattoir was used for the experiments. The tracheas were placed in physiological saline solution at 4°C immediately after their removal from the animals. Smooth muscle strips (~6 × 1.5 × 0.3 mm in dimension) were dissected from the tracheas. The strips were attached to aluminum foil clips at both ends; one end was placed over a stationary hook inside the muscle bath, and the other end was connected to the lever arm of a servo-controlled force/length transducer. Two different solutions were used. Physiological saline solution (PSS) with pH 7.4 at 37°C, bubbled with a gas mixture (5% CO2-95% O2), had a composition (in mM) of 118 NaCl, 5 KCl, 1.2 NaH2PO4, 22.5 NaHCO3, 2 MgSO4, 2 CaCl2, and 2g/l dextrose. Zero-calcium saline (same composition as PSS, but with no added calcium) contained 1 mM EGTA and was bubbled with the same gas mixture to maintain the pH at 7.4.

Apparatus

The servo-controlled force/length lever system had a force resolution of 10 µN and a length resolution of 1 µm. The analog signals were converted to digital signals via a National Instrument analog-to-digital converter and were recorded by a computer. The computer also controlled the sequence of events during the experiments. Electrical field stimulation (EFS) of the muscle was provided by a 60-Hz alternating current stimulator with platinum electrodes. Sinusoidal length oscillations applied to the muscle were provided by the lever system, as described previously (13).

Experimental Procedures

All muscle preparations were equilibrated in the PSS at 37°C and stimulated (EFS) once (for 12 s) every 5 min until the isometric force production reached a steady state. This process took about 1 h. During equilibration, a reference length of the muscle (Lref) was established by stretching the muscle to approximately the in situ length, at which the resting tension was 1-2% of the maximal isometric force.

Determination of thick filament density change due to activation. Two muscle strips from each trachea were used in the paired experiments. One strip was fixed (for electron microscopy analysis) in the relaxed state; the other strip was fixed at the plateau of contraction elicited with 10-4 M acetylcholine. Four such paired experiments were carried out with four tracheas.

Determination of thick filament density change in the relaxed state in the presence and absence of calcium and mechanical perturbation. Tracheas from four animals were used for the experiments. Four strips of muscle per trachea were dissected and fixed under four different conditions, as illustrated in Fig. 1. A muscle strip fixed at time 1 was used as the control. At time 2, a strip of muscle was fixed after five stimulations over a period of 25 min in zero-calcium PSS. Isometric force diminished during the time course of stimulation and was not detectable after the third stimulation. At time 3, a 5-min period of length oscillation was applied to the muscle while it was still in zero-calcium PSS. The amplitude of stretch associated with the oscillation was 30% Lref. At time 4, the muscle was allowed to recover after the oscillation in normal PSS. During the period of recovery (25 min), the muscle was stimulated electrically once every 5 min. Full recovery of isometric force occurred after the first couple of stimulations. The muscle strip was fixed after relaxation from the fifth contraction. Note that all preparations were fixed in the relaxed state in this group of experiments. The same protocol was repeated four times for four groups of muscle preparations from four tracheas. Results from each of the four strips under the same condition from four different animals were averaged to obtain the final data.


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Fig. 1.   Schematics illustrate time points (times 1-4) during the course of experiment when the muscle samples were fixed for electron microscopic examination. Contraction of muscle was elicited by a 12-s electrical field stimulation (EFS). Note that the samples were all fixed in resting state.

Electron Microscopy

Muscle preparations were fixed for 15 min while they were still attached to the experimental apparatus. Care was taken not to disrupt the muscle mechanically during the fixation. The fixing solution contained 1.5% glutaraldehyde, 1.5% paraformaldehyde, and 2% tanic acid in 0.1 M sodium cacodylate buffer that was prewarmed to the same temperature as the bathing solution (37°C). After the initial fixation, the strip was removed from the apparatus, cut into small blocks ~1 × 0.5 × 0.3 mm in dimension, and put in the fixing solution for 2 h at 4°C on a shaker. The blocks were then washed three times in 0.1 M sodium cacodylate (30 min). In the process of secondary fixation, the blocks were put in 1% OsO4-0.1 M sodium cacodylate buffer for 2 h, followed by three washes with distilled water (30 min). The blocks were then further treated with 1% uranyl acetate for 1 h (en bloc staining), followed by washes with distilled water. Increasing concentrations of ethanol (50, 70, 80, 90, and 95%) were used (10 min each) in the process of dehydration. Ethanol (100%) and propylene oxide were used (three 10-min washes each) for the final process of dehydration. The blocks were left overnight in the resin (TAAB 812 mix, medium hardness) and then embedded in molds and placed in an oven at 60°C for 8-10 h. The embedded blocks were sectioned on a microtome with a diamond knife and placed on 400-mesh cooper grids. The section thickness was ~100 nm. The sections were then stained with 1% uranyl acetate and Reynolds lead citrate for 4 and 3 min, respectively. The images of the cross sections of the muscle cells were obtained with a Phillips 300 electron microscope.

Morphometric Analysis

Analysis of thick filament density change due to activation. A total of 40 electron micrographs of muscle cell cross sections were analyzed in this group of experiments. Five pictures per strip of muscle were randomly selected. The numbers of thick filaments in five cells from each of the eight muscle strips (4 pairs) were determined by counting the filaments in multiple small squares. A square equivalent to 1 µm2 was placed over as many organelle-free areas as could be found in each cell, and the numbers of filaments in the square were counted. The filament density assessed in this way was multiplied by the total organelle-free area of the cell to obtain the total number of filaments in the cross section.

Analysis of thick filament density change in the relaxed muscle in the presence and absence of calcium and mechanical perturbation. A total of 240 images of cross sections of the muscle cells were analyzed in this group of experiments. The process of sampling and analysis of the images were "blind," and the codes were revealed only after the analysis of all groups was completed. Each image contained a whole cell cross section. Fifteen pictures per strip of muscle (60 images/animal) were selected. Five of these fifteen pictures contained cross sections of cells with nuclei, five contained central clusters of mitochondria, and five contained some or no organelles. This protocol of sampling of cross sections was used to ensure that the ultrastructure in different regions of a cell was examined. Myosin thick filaments were identified by eye and manually counted in the whole cell cross section. A potential inaccuracy in identifying the thick filaments was due to the variable size of the thick filament cross sections. The thick filaments, on average, have a diameter of about 15-20 nm. However, the filaments have tapered ends, and the diameter at both ends is therefore smaller than that in the middle. The thick filaments, however, can be identified by the "rosette" of thin filaments surrounding them under most circumstances, regardless of their size. It is still possible though that the number of thick filaments was underestimated with our counting method, but because the same criteria were used in counting all groups, the underestimation should be the same in all groups. The cell cross-sectional areas were measured with a morphometric digital device made by Carl Zeiss. The thick filament density was obtained by dividing the number of thick filaments by the total area of the cell cross section minus areas occupied by nuclei, mitochondria, and other organelles.

Statistics

The statistical analysis was performed using Microcal Origin 5.0 computer software. The values shown are means ± SE. For the one-way ANOVA, significant difference was associated with a P value <0.05.


    RESULTS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Effects of Activation on Thick Filament Density

Figure 2 shows two examples of airway smooth muscle cell cross sections fixed in the relaxed state (Fig. 2A) and at the plateau of acetylcholine-elicited contraction (Fig. 2B). The average isometric force produced by the contracted muscles was 161.4 ± 11.3 kPa. The thick filament density in the relaxed state was 60.8 ± 3.42 filaments/µm2. The density in the contracted state was 148.4 ± 7.49 filaments/µm2. The cross-sectional areas (total area) of the cells in the relaxed and contracted states were 12.7 ± 0.96 and 14.5 ± 1.07 µm2, respectively, and they were not different statistically.


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Fig. 2.   Examples of electron micrographs of muscle cell cross sections. A: muscle sample fixed in the relaxed state. B: muscle sample fixed in the contracted state (10-4 M Ach). Arrows point to myosin thick filaments surrounded by actin thin filaments. Calibration bar, 1 µm.

Effects of Calcium and Mechanical Perturbation on Thick Filament Density

Figure 3 shows two examples of airway smooth muscle cell cross sections fixed at time 4 (Fig. 3A) and time 2 (Fig. 3B). As shown in Fig. 1, time 4 indicates the time point at which the muscle preparation had recovered from being in zero-calcium PSS and subjected to length oscillation. The isometric force, thick filament density, and other morphometric appearances of the cross sections at time 4 were not different from those at time 1, indicating that the experimental protocol did not cause permanent cell damage. Time 2 indicates the time point at which the muscle preparation produced no isometric force due to the lack of calcium (Fig. 1).


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Fig. 3.   Examples of electron micrographs of muscle cell cross sections. A: muscle sample fixed at time 4 (indicated in Fig. 1). B: muscle sample fixed at time 2. Arrows point to myosin thick filaments surrounded by actin thin filaments. Insets: higher magnification of indicated areas. Calibration bar, 1 µm.

Figure 4 shows the averaged (n = 4) thick filament densities for the samples fixed at the four time points shown in Fig. 1. One-way ANOVA showed that the variation in thick filament density is significant (P < 0.05). However, there was no difference (P = 0.67) between the groups at times 1 and 4. Also, no significant difference (P = 0.59) was found between the groups at times 2 and 3. Isometric force of the muscle was totally abolished in zero-calcium PSS. The force, however, recovered fully after reincubation of the muscle in normal PSS; isometric force produced by the muscle at times 1 and was not different (P = 0.9) (Fig. 4).


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Fig. 4.   Thick filament density and isometric force associated with the 4 conditions under which the muscles were fixed. The 4 conditions are indicated by the 4 time points illustrated in Fig. 1. Isometric tetanic force was elicited by 12-s EFS. At times 2 and 3, force was zero. Values are means ± SE.

The number of thick filaments per cell cross section and the cross-sectional cytosolic areas (the total cell cross-sectional area minus the areas occupied by the cellular organelles) of the muscle cells fixed in the four conditions (or time points in Fig. 1) are listed in Table 1. One-way ANOVA showed that there was no significant difference (P = 0.53) among the cross-sectional areas.

                              
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Table 1.   Number of myosin thick filaments per cell cross section and cell cross-sectional cytosolic areas


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Lability of myosin thick filaments has been demonstrated in some intact smooth muscle preparations (2, 3, 14, 15), including that of airway smooth muscle (5). The labile nature can be seen in the variation of the thick filament density in cross sections of cells fixed under different experimental conditions (2, 5, 15). The ability of smooth muscle thick filaments to depolymerize and repolymerize may facilitate the muscle in adapting to large changes in cell length (7).

The 144% increase in myosin filament density in contracted airway smooth muscle (compared with the density in the relaxed state) observed in the present study represents the largest density change due to activation found in any smooth muscles so far. The density change due to activation in rat anococcygeus was about 23%, whereas in guinea pig taenia coli there was no significant increase (15). The difference in the observed myosin evanescence was attributed to the difference in the "active state" in resting muscles (2, 15). Taenia coli normally does not exist in a truly relaxed state because of its ability to spontaneously contract. The thick filaments may not have a chance to disassemble in such a muscle. Anococcygeus muscle, on the other hand, has a long quiescent period in between contractions; this may allow the thick filaments to partially depolymerize. The airway smooth muscle preparation that we used has an extremely low resting activity, judging from the absence of active tone and zero myosin light chain (MLC) phosphorylation found in this preparation (6). This may explain the large difference in the thick filament density between the relaxed and contracted states found in this study. Other explanations, however, cannot be excluded; the variation in myosin evanescence may be due to differences in properties of myosin molecules and other cellular properties associated with different types of smooth muscle.

It has been reported in rat anococcygeus that the cell cross-sectional area decreases when the muscle contracts (2, 15). The cell shrinkage was not observed in guinea pig taenia coli (3), at least not consistently (15). No significant change in the cell cross-sectional area due to activation was found in this study, and therefore no correction for the area variation was carried out.

The underlying mechanism for the thick filament lability in intact smooth muscle is not clear. Studies of isolated myosins in solution have suggested that calcium-dependent phosphorylation of the regulatory MLC was essential in the formation of the thick filaments (4, 11). The present finding of a substantial increase in myosin filament density upon muscle activation supports the notion that MLC phosphorylation could enhance thick filament formation. However, this does not mean that dephosphorylation of the light chain will cause total dissolution of the thick filaments. Thick filaments have been found in relaxed, dephosphorylated muscles (10, 12). Results from the present study also indicate that thick filaments exist in relaxed airway smooth muscle, even after removal of extracellular calcium and in the presence of the calcium-chelating agent EGTA. Calcium removal, however, does lower the thick filament density significantly (35%; Fig. 4). An intriguing question is whether this decrease in the filament density is due to a decrease in the level of MLC phosphorylation below its normal resting level. This is not likely considering that the normal resting level of phosphorylation in our preparation is virtually zero (6). Also, in a recent study (Qi D, Mitchell RW, Burdyga T, Ford LE, Kuo K, and Seow CY, unpublished observations) we used wortmannin (a potent inhibitor of MLC kinase) to examine the effect of MLC phosphorylation on thick filament formation; we found that inhibition of MLC phosphorylation by wortmannin in the presence of calcium did not cause a reduction in the thick filament density from its normal resting level (although length oscillation was able to reduce the filament density under this condition, and recovery of the density required removal of wortmannin). The reduction in thick filament density (without mechanical agitation) associated with calcium removal therefore is likely due to disruption of some pathways that rely on a normal resting calcium level. The condition of zero extracellular calcium with EGTA likely creates a nonphysiological state for the muscle. The experiment, however, demonstrated that a normal resting level of calcium was important for preservation of filamentous myosins in relaxed airway smooth muscle. It is interesting to observe that spontaneous calcium release (that does not contribute significantly to the global intracellular calcium concentration) appears to be essential for myofibrillogenesis in striated muscle. Blockade of the transient calcium release disrupts myosin thick filament assembly in the developing Xenopus myocytes (1). It is not known whether the occasional calcium "sparks" observed in resting smooth muscle have the same function in maintaining integrity of the thick filaments in the relaxed state.

Our previous study (5) shows that mechanical oscillation imposed on relaxed airway smooth muscle (in the presence of normal extracellular calcium) resulted in a transient reduction in the thick filament density. The density (and isometric force) recovered fully after a period of 25-30 min during which the muscle was stimulated electrically once every 5 min (5). In the present study, the same oscillation protocol was used but with the muscle bathed in zero-calcium PSS. Figure 4 shows that the amount of decrease in myosin thick filament density after oscillation (Fig. 1, time 3) was the same as that after removal of calcium without oscillation (Fig. 1, time 2). This is somewhat surprising considering that mechanical oscillation has been shown to be able to "break up" the labile (presumably nonphosphorylated) thick filaments in relaxed muscles (5). It has been shown under in vitro conditions that there exists an equilibrium between filamentous and monomeric myosins (4), and once a critical concentration of monomeric myosins is reached, thick filaments will form even if the myosins are not phosphorylated. It is possible that the amount of thick filaments seen under zero-calcium conditions in the present study represents the minimum amount of thick filaments maintained by the equilibrium. Mechanical agitation may be able to cause dissolution of the filaments, but the equilibrium force would quickly reestablish the minimum concentration of thick filaments. It also is possible that the intracellular pathway that conveys the mechanical signal leading to dissolution of the thick filaments is calcium dependent; with removal of calcium, this pathway may not be functional, resulting in the insensitivity of thick filament density to mechanical stretch.

In conclusion, activation of airway smooth muscle resulted in a 144% increase in the amount of myosin filaments seen in muscle cell cross sections; the activation-dependent increase in the filament density may be related to the increase in MLC phosphorylation during muscle activation. Removal of calcium resulted in a 35% decrease in the filament density from its normal resting level; this change in filament density appears to be independent of MLC phosphorylation.


    ACKNOWLEDGEMENTS

We offer special thanks to Pitt Meadows Meats Limited (Pitt Meadows, BC) for the supply of fresh porcine tracheas in kind support for this research project. In particular, we thank Cathy Pollock, Inspector in Charge, Canada Food Inspection Agency (Establishment no. 362) for help in obtaining the tracheas.


    FOOTNOTES

This study is supported by operating grants from Canadian Institutes of Health Research to C. Y. Seow (MT-13271).

Address for reprint requests and other correspondence: C. Y. Seow, Dept. of Anatomy, Univ. of British Columbia, 2177 Wesbrook Mall, Vancouver, BC, Canada V6T 1Z3 (E-mail: cseow{at}interchange.ubc.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.

First published October 24, 2001; 10.1152/ajpcell.00390.2001

Received 10 August 2001; accepted in final form 22 October 2001.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Ferrari, MB, Ribbeck K, Hagler DJ, Jr, and Spitzer NC. A calcium signaling cascade essential for myosin thick filament assembly in Xenopus myocytes. J Cell Biol 141: 1349-1356, 1998[Abstract/Free Full Text].

2.   Gillis, JM, Cao ML, and Godfraind-De Becker A. Density of myosin filaments in the rat anococcygeus muscle, at rest and in contraction. II. J Muscle Res Cell Motil 9: 18-29, 1988[ISI][Medline].

3.   Godfraind-De Becker, A, and Gillis JM. Analysis of the birefringence of the smooth muscle anococcygeus of the rat, at rest and in contraction. I. J Muscle Res Cell Motil 9: 9-17, 1988[ISI][Medline].

4.   Kendrick-Jones, J, Smith RC, Craig R, and Citi S. Polymerization of vertebrate non-muscle and smooth muscle myosins. J Mol Biol 198: 241-252, 1987[ISI][Medline].

5.   Kuo, K, Wang L, Paré PD, Ford LE, and Seow CY. Myosin thick filament lability induced by mechanical strain in airway smooth muscle. J Appl Physiol 90: 1811-1816, 2001[Abstract/Free Full Text].

6.   Mitchell, RW, Seow CY, Burdyga T, Maass-Moreno R, Pratusevich VR, Ragozzino J, and Ford LE. Relationship between myosin phosphorylation and contractile capability of canine airway smooth muscle. J Appl Physiol 90: 2460-2465, 2001[Abstract/Free Full Text].

7.   Pratusevich, VR, Seow CY, and Ford LE. Plasticity in canine airway smooth muscle. J Gen Physiol 105: 73-94, 1995[Abstract].

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10.   Somlyo, AV, Butler TM, Bond M, and Somlyo AP. Myosin filaments have non-phosphorylated light chains in relaxed smooth muscle. Nature 294: 567-569, 1981[ISI][Medline].

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Am J Physiol Cell Physiol 282(2):C310-C316
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