Articles in PresS. Am J Physiol Cell Physiol (April 16, 2003). 10.1152/ajpcell.00095.2003
Structure-function correlation in airway smooth muscle adapted to different lengths (Running Title: Smooth muscle plasticity)
Kuo-Hsing Kuo1,7, Ana M. Herrera3,7, Lu Wang2,7, Peter D. Paré2,7, Lincoln E. Ford5, Newman L. Stephens6, and Chun Y. Seow1,3,4,7
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Department of Anatomy, 2Department of Medicine, 3Department of Pathology and Laboratory
Medicine, 4Department of Pharmacology and Therapeutics, University of British Columbia, Vancouver, BC V6T 1Z3 5
Krannert Institute of Cardiology, Indiana University, Indianapolis, IN 46220
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Department of Physiology, University of Manitoba, Winnipeg, MB R3E 3J7
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The UBC McDonald Research Laboratories / The iCAPTURE Centre, St. Paul's Hospital /
Providence Health Care, Vancouver, BC, V6Z 1Y6
Corresponding Author: Dr. Chun Y. Seow, Department of Pharmacology & Therapeutics, University of British Columbia, 2176 Health Sciences Mall, Vancouver, BC, Canada, V6T 1Z3 Tel: (604) 822-8006, or (604) 806-9268; Fax: (604) 822-6012, or (604) 806-9274 Email:
[email protected]
Copyright (c) 2003 by the American Physiological Society.
ABSTRACT Airway smooth muscle is able to adapt and maintain a nearly constant maximal force generation over a large length range. This implies that a fixed filament lattice such as that found in striated muscle may not exist in this tissue, and that plastic remodeling of its contractile and cytoskeletal filaments may be involved in the process of length adaptation that optimizes contractile filament overlap. Here we show that isometric force produced by airway smooth muscle was independent of muscle length over a 2-fold length change; cell cross-sectional area was inversely proportional to cell length, implying that the cell volume was conserved at different lengths; shortening velocity and myosin filament density varied similarly to length change: increased by 69.4%r5.7(SE) and 76.0%r9.8 respectively for a 100% increase in cell length. Muscle power output, ATPase rate and myosin filament density also have the same dependence on muscle cell length: increased by 35.4%r6.7, 34.6%r3.4, and 35.6%r10.6 respectively for a 50% increase in cell length. The data can be explained by a model where additional contractile units containing myosin filaments are formed and placed in series with existing contractile units when the muscle is adapted at a longer length.
Key words: muscle contraction, myosin filaments, ATPase activity, electronmicroscopy.
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INTRODUCTION Recent studies have demonstrated that the cycle of contraction and relaxation in airway smooth muscle involves polymerization and depolymerization of the actin (9, 13) and myosin (6) filaments. This lability of contractile filaments during muscle activation resembles that of nonmuscle motile cells that rely on impromptu assembly of contractile apparatus for motility. Oscillatory strains applied to single airway smooth muscle cells have revealed plastic deformation similar to that observed in soft glassy materials (3). Functional studies of airway smooth muscle indicate that the cells maintain maximal isometric force production (optimal contractile filament overlap) over a 3-fold length change by varying the number of contractile units in series (15). This newly emerged “plasticity” model of smooth muscle contraction is incompatible with our current concept of contraction based on the relatively restrictive model of striated muscle that possesses filament arrays in fixed lattice, which cannot accommodate large changes in cell length without compromising force production (4). The feasibility of the plasticity model can be tested experimentally based on the model predictions: the recruitment of additional contractile units in muscles adapted to longer lengths should result in an increase in thick filament content (or mass) in individual muscle cells, and as a consequence, the metabolic rate should increase with muscle length. In the present study, we tested the plasticity hypothesis through quantitative analysis of mechanical properties and ultrastructure, and measurement of the rate of ATP consumption in airway smooth muscles adapted to different lengths.
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MATERIALS AND METHODS
Muscle preparation Muscle tissue was obtained from animals sacrificed for other purposes. Pig tracheas were obtained from a local abattoir while dog tracheas were kindly given to us by colleagues who used other parts of the animals for their experiments. Muscle bundles were dissected from a trachea such that all preparations from one animal had the same dimensions. They were studied in a chamber equipped with a force transducer as previously described (12, 19).
Solutions Intact muscle experiments were done at 37 qC, pH 7.4 in physiological saline equilibrated with 95% O2, 5% CO2 and containing, in mM, NaCl 118, NaHCO3 22.5, KCl 5, NaH2PO4 1.2, MgSO4 2, CaCl2 2 and glucose 2g/liter. Permeabilized muscle was studied at room temperature and pH 7.0 in solutions containing EGTA 5, K-acetate 85, imidazole 20, MgCl2 6.1, Na2ATP 5.6, dithiothritol 1, NaN3 1, phosphoenol pyruvate 5, P1, P5-di(adenosine-5') pentaphosphate 0.2, NADH 0.6, lactic dehydrogenase 70U/ml, and pyruvate kinase 50 U/ml. Activating solution contained sufficient calcium to buffer the free [Ca++] to 25 PM while relaxing solution had no added calcium. A rinse solution containing 0.1 mM EGTA was used to lower intracellular EGTA just prior to activation.
Muscle force, velocity, and power measurements For the group of experiments examining changes in mechanical properties associated with a 1.5fold change in length, canine trachealis preparations were used (in order to match the ATPase
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data); whereas in the group examining changes associated with a 2-fold change in length swine trachealis preparations were used. The reason for using different animal tissues is because this study is a combination of three independent studies carried out in three laboratories (Seow, Ford, and Stephens). Methods for measuring force-velocity parameters have been described in detail previously by us (15, 19). Briefly, aluminum clips were attached to both ends of a muscle preparation; one end of the muscle was attached to a stationary hook at the bottom of a vertical muscle bath, the other end was attached to the force/length transducer (lever system) via a lowcompliance surgical silk suture about 10 cm in length. Muscle preparations were equilibrated/adapted for about hour at 37 qC in physiological saline bubbled with a 95% O2-5% CO2 mixture until the isometric tension produced by the muscle reached a maximal, steady state. During the equilibration period the muscle was stimulated electrically to produce a brief tetanus (12 s) once every 5 min. During the period a reference length (Lref) for the muscle was determined. The length where rest tension was just detectable, approximately the in situ length, defines Lref. After equilibration, isotonic shortening velocity was measured by quick-releasing the muscle from plateau of an isometric contraction to a pre-set isotonic load. Velocity of shortening was determined by measuring the slope of the shortening trace 100 ms after the quick release when the servo-controlled shortening reached a steady state. 6-10 such releases were performed on a muscle preparation to obtain force-velocity data used in the curve fitting to generate a hyperbolic (Hill’s) force-velocity curve (7) of the form: V = b(Fmax – F)/(F + a), where V is shortening velocity, Fmax is the maximal isometric force, F is the isotonic load, and a and b are Hill’s constants. Velocity change in a muscle adapted to different lengths was
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determined by scaling velocity values of the force-velocity curve obtained at Lref by a variable until the best fit was obtained for the force-velocity data obtained at the test length (.5 or 2 Lref). The same procedure was used in our previous studies (15). Maximal muscle power output was determined from the force-velocity curve as described previously (19). Briefly, the power output (P) of the muscle as a function of load (F) was obtained by multiplying V and F: P = FV = Fb(Fmax – F)/(F + a). The maximal power (Pmax) was calculated by differentiating the forcepower function (with respect to F) to obtain the force associated with Pmax: F’ = a[(Fmax/a + 1)-1/2 – 1], and substitute F’ for F in the force-power function. The number of animals used for the .5-Lref group of experiments was 9; for the 2-Lref group, the number was 6.
ATPase measurements Muscles (canine trachealis) were first adapted to the reference length as described above. A pair of muscle preparations from one trachea was studied in a single experiment; three such experiments were carried out. One member of the pair was adapted to Lref and the other to 1.5 Lref. Muscles were permeabilized by 2 hrs immersion in a relaxing solution containing 1% (v/v) Triton X-100 and studied in an apparatus designed by Güth and Wojciechowski (5). Both muscles were studied at 2 lengths, first the length where it was adapted and then at the other length. Structural constraints of the apparatus limited muscle length changes to 1.5-fold. One end of the muscle was tethered to a stationary hook; the other end was attached to a force transducer. A quartz cuvette containing solution was slipped over the muscle, which was illuminated with a 340 nm light source. ATP hydrolysis was measured by the rate of decline in fluorescence of NADH, a reduced form of nicotinamide adenine dinucleotide used to regenerate ATP in conjunction with the phosphoenol pyruvate/pyruvate kinase intermediate. Fresh NADH
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was injected into the cuvette at 25 s intervals to maintain a high level. Fluorescence at 450 nm (wavelength) was measured over a 1 mm length of muscle, and total muscle ATPase rate was estimated by multiplying the rate of fluorescence decline by the length of the muscle. The fluoresced light passed through a rectangular aperture in the photomultiplier, 1 mm wide in the direction parallel to the muscle and 2 mm in the other dimension. Because the cross-sectional area of the cuvette was at least 20 times larger than that of the muscle, and because diffusion of ADP and NADH from the 0.2 mm wide muscle is very rapid, while diffusion over the >5 mm muscle length is comparatively very slow, it is likely that the optical signal derived almost entirely form the NADH in the extracellular space, and was not influenced by either the optical properties of the tissue or by diffusion to and from ends of the muscle.
Electronmicroscopy Samples (swine trachealis) were prepared for electron microscopy as previously described (6, 12, 16). Briefly, 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% tannic acid in 0.1 M sodium cacodylate buffer that was pre-warmed to the same temperature as the bathing solution (37 ºC). After the initial fixation, the strip was removed from the apparatus and cut into small blocks, approximately 1 x 0.5 x 0.2 mm in dimension, and put in the fixing solution for 2 hrs at 4 ºC on a shaker. The blocks were then washed three times in 0.1 M sodium cacodylate (3x10 min). In the process of secondary fixation, the blocks were put in 1% OsO4, 0.1 M sodium cacodylate buffer for 2 hours followed by three washes with distilled water (3x10 min). The blocks were then further treated with 1% uranyl acetate for 1 hour (en bloc staining)
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followed by washes with distilled water. Increasing concentrations of ethanol (50%, 70%, 80%, 90% and 95%) were used (ten minutes each) in the process of dehydration. 100% ethanol 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 ºC for 8-10 hrs. The embedded blocks were sectioned on a microtome using a diamond knife, and place 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. Images of the cross-sections of cells were made with a Phillips 300 electron microscope at various magnifications.
Morphometric Analysis The myosin thick filament had an amorphous cross-sectional profile with an average diameter of about 15 nm. The actin thin filament had a circular cross-section with a diameter of 6 nm. The number of thick filaments in 5 cells from each muscle preparation (n=4 for the 2xLref group and n=5 for the .5xLref group, where n is the number of pairs of muscle strips from one trachea and also equals the number of animals used). Examples of electron micrographs are given in Fig. 1. The filament density was determined by dividing the total filament number in a cross-section by the cross-sectional cytoplasmic area, which was the cell’s cross-sectional area minus the areas occupied by organelles such as nucleus, mitochondria, sarcoplasmic reticulum, and caveolae. The same method of quantifying myosin filament density was used in our previous studies (6, 12, 16). Morphometric measurements were carried out “blind” – the tissue samples were relabeled by a person not involved in the experiments and the true identity of the samples was
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revealed only after all samples were analyzed. Identification of the filaments was done manually and quantification was done with the assistance of the software Image Pro Plus.
RESULTS Length-dependence of force-velocity parameters The same measurements have been done in our previous studies (15). The reason for repeating the measurements was to match animal species and to make sure that the same muscle behavior was observed. As shown in Fig. 2 and 3, and also Table 1, isometric force was not significantly different over a 2-fold length change. Note that the ratios of isometric forces at different muscle lengths were pooled together from different groups of experiments; the ratios from each group showed no significant difference from unity. Shortening velocity and power output, in contrast to isometric force, increased significantly at both 1.5 and 2.0 Lref compared to the corresponding values at Lref (Table 1). The value of maximal shortening velocity listed in Table 1 has been normalized by Lref. The un-normalized value was 1.65±0.37 (SEM) mm/s.
Length-dependence of muscle metabolic rate ATPase activities were measured fluorimetrically in canine trachealis muscles permeabilized by partial dissolution of the plasma membranes and stimulated to contract by the addition of calcium. Two lengths were studied, Lref and 1.5 Lref. At these two lengths, force was not significantly different and ATPase activity was 34.6%±3.4(SE) greater for the 1.5-times longer muscle (Fig. 2, Table 1). The ATPase activity was normalized by isometric force to reduce inter-preparation difference in the rates. Without normalization, the ATPase activity increase was 32±8% for a 50% increase in muscle length. This is not statistically different from
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the normalized value; the standard error, however, is larger without normalization. The finding that inter-preparation variation in ATPase rate could be reduced when the developed force was taken into account is consistent with the finding in skinned gizzard muscle that ATPase activity was directly proportional to the isometric force that a muscle has developed (10).
Length-dependence of myosin thick filament density Two groups of experiments were carried out to assess changes in myosin filament density at different muscle lengths: .5 Lref vs. Lref and 2.0 Lref vs. Lref. Two adjacent trachealis strips with the same initial length were dissected from each animal, one was set at Lref and the other at a test length (.5 or 2.0 Lref). After at least 1 h of adaptation at a set length, the muscle was stimulated to produce a ~120 s contraction by the addition of 0.1 mM acetylcholine and fixed for electron microscopy in the contracted state. Acetylcholine was used for the final stimulation to ensure continued activation during fixation. Cell cross-sectional areas measured in the electron micrographs were decreased by approximately half in muscles fixed at 2xLref (Fig. 3, Table 1) compared to that at Lref, indicating that cell volume was conserved. The myosin filament density increased by 35.6%r10.6(SE) and 76.0%r9.8 with 50% and 100% increase in muscle length, respectively. These values were obtained from density ratios of paired muscle preparations from the same trachea. The pooled absolute values of filament density are listed in Table 1. The filament density found in this study was much higher than that found in our previous studies (6, 12, 16); this is because the density values listed in our previous publications were those determined in the relaxed muscles, whereas the density reported here was determined in the activated muscles.
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DISCUSSION In our previous study (15) isometric force of canine trachealis was found to be independent of muscle length (after length adaptation) and velocity was shown to be increased at long lengths, and the force-velocity curves obtained at different lengths could be superimposed by plotting force relative to its isometric level and scaling velocities by a constant factor for each length. The scale factor therefore indicated changes in velocity. The scale factor was found to be less than the length change, ~67% when the length was doubled (15). We interpreted the data as suggesting that the number of contractile units (of which the thick filaments are an essential component) in series increased by 67% for length doubling (15). This interpretation is illustrated in Fig. 4. In the highly simplified case, an increase in the number of contractile units from 3 to 5 (67% increase) as a result of length doubling should cause a 67% increase in shortening velocity and power output, but no change in isometric force, because the additional contractile units are in series with the existing ones (Fig. 4). The present findings in terms of length dependence of force-velocity parameters are nearly identical to our previous findings (15); the model depicted in Fig. 4 therefore applies to the present results. The model predicts that the energetic cost of contraction increases with adapted muscle length, and that the increase should match the power increase. This is exactly what we found (Fig. 2). The model also predicts that the content (or mass) of myosin filament increases with adapted muscle length. Currently the only reliable measurement available to us that indirectly estimates myosin filament content is the thick filament density found in a transverse section of a cell. This indirect measurement of myosin filament content also agrees with the model prediction, as discussed below. If the thick filament length is much less than the length of a cell, and also if they are randomly distributed within the cell regardless of the cell length, the thick filament density in a
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cell cross-section should remain constant at different cell lengths provided the filament content (or mass) is length independent (i.e., no filament polymerization or depolymerization with length change). This is analogous to a bag of black and white marbles; as long as the balls are evenly mixed, the number of black marbles per unit area (density) found in any imaginary planes cutting across the bag will be the same whether the bag is stretched or squashed. The present finding that the thick filament density increased in muscles adapted at longer length suggests filamentogenesis. Furthermore, the filament density in a cross-section should reflect filament content of a cell provided that the cell volume is conserved. (Use the same analogy above, the number of black marbles found per unit area in a cross-section is directly correlated to the total number (or content) of black marbles in the bag). The inverse relationship between cell crosssectional area and cell length (Fig. 3, Table 1) indicates that the cell volume was indeed the same within the lengths studied. The 35.6%r10.6 and the 76%r9.8 increase in thick filament density found at 50% and 100% length increase therefore closely fit the model prediction (Fig. 4). The present study therefore provides the first ever evidence that myosin filament content in airway smooth muscle is regulated by muscle length. How is it regulated is unknown. It could however be speculated that cytoskeletal deformation could lead to activation of integrinlinked kinase that in turn promotes myosin light chain phosphorylation (2) and thick filament formation (16). The observation that there is a large increase in the thick filament density (in a matter of seconds) when airway smooth muscle is activated (6) suggests that there is a large pool of monomeric myosins in the trachealis cells. There are alternative explanations of our data. The increase in mechanical power output at longer lengths could be a result of increased energetic efficiency in the muscle, although this is unlikely because the rate of ATP consumption found in the present study was similarly
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dependent on the muscle length (Fig. 2). In fact, the results suggest that the simplest explanation is that the efficiency of the muscle is unchanged at different lengths. The increased ATPase activity at longer muscle length could also be due to activation of cross bridges previously not activated at short lengths. Measurements of thick filament density however suggest that there is a net increase in thick filament number in the cell as the cell length increases (Fig. 2 & 3). A simpler explanation therefore is that the increase in ATPase rate is due to a matching increase in the thick filament content. Furthermore, the findings that shortening velocity and ATPase activity increase in the same proportion together with the finding that the force-velocity curves obtained at different lengths can be superimposed by velocity scaling imply that cross-bridge function (in terms of cycling rate and force per bridge) is the same at the two lengths. The absence of a change in cross-bridge function would require that the individual filaments slide at the same rate in long and short muscles, so that the amount of increase in velocity and power would require the same amount of increase in filaments in series when length was increased. Taken together, the results are consistent with the interpretation that the increase in the cell content of myosin filaments is the underlying cause for the increases in the other three measures of muscle activity (velocity, power, and ATPase activity). The model shown in Fig. 4 assumes that the length of thick filaments is the same at different cell lengths, and the increase in thick filament content at longer muscle length is due to more thick filaments added in series, and not due to the existing thick filaments getting longer, to account for the present finding that isometric force is length independent. Another model that predicts the same amount of increase in filament content and length-independence of isometric force allows the thick filament length to increase in inverse proportion to the number of thick filaments in parallel (to keep isometric force constant). This model also has to allow the number
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of thick filaments in series to vary with muscle length in order to account for the lengthdependent shortening velocity. Results from the present study do not allow us to favor one model over the other. The model presented in Fig. 4 however is simpler in the sense that it does not rely on a potentially complicated mechanism that adjusts the thick filament length and the number of filaments in parallel precisely according to the cell length. Although there is always a danger in extrapolating interpretation on one smooth muscle type to another, it could however be speculated that the very long functional length range (up to 10-folds change in length) of rabbit urinary bladder muscle (18) and pedal retractor muscle of Mytilus edulis (blue mussel) (8) could be due to a similar plastic mechanism described above. The length dependence of smooth muscle force (20), intracellular calcium concentration and myosin light chain phosphorylation (17), ATPase activity (1), and rate of oxygen consumption (14) have been documented. The present results, however, cannot be compared to those from the early studies because of the length adaptation protocol (cyclic activation of the muscle at a constant length) used in our experiments has largely eliminated the dependence of isometric force on muscle length and altered the ultrastructure of the cell. The need for plastic adaptation is quite obvious in smooth muscles lining the wall of hollow organs that undergo large volume changes. It is not clear why such plasticity exists in airway smooth muscle. It is clear, however, that there are a variety of in vivo conditions under which airway smooth muscle could be adapted to pathologically short lengths, especially when inflammation of the airways is involved: frequent and prolonged stimulation of the muscle, adventitial edema of the airway wall that decreases resting muscle length, reduced elastic tethering of lung parenchyma on the airway wall (due to chronic inflammation) that diminishes resting muscle length, to name just a few (11). Elucidating the mechanism of airway smooth
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muscle plasticity therefore will, in addition to gaining insights into the basic mechanism of contraction, shed light on the pathophysiology of exaggerated airway narrowing seen in diseases such as asthma and emphysema.
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ACKNOWLEDGEMENT
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 would like to thank Cathy Pollock, Inspector in Charge, Canada Food Inspection Agency (Establishment # 362) for her help in obtaining the tracheas. This study was supported by operating grants from Canadian Institutes of Health Research (MT-13271, MT-4725).
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bridge phosphorylation in the swine carotid media. J Physiol 488:729-739, 1995.
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FIGURE LEGENDS
Figure 1: Examples of electron micrographs of cross sections of trachealis fixed in the contracted state at (A) reference length (Lref), and (B) .5 Lref. Arrows point to myosin thick filaments. D, dense body. Inset: Magnified rectangular area shown in A, arrowheads point to myosin thick filaments surrounded by actin thin filaments. Calibration bar indicates 1 Pm. The cross-sectional areas of cells in A and B are 8.27 and 5.02 Pm2 respectively, and their cytoplasmic areas are 7.48 and 3.94 Pm2 respectively. The thick filaments counted in A are 532 and 379 in B, the thick filament densities are therefore 71.1 and 96.2 filaments/Pm2 in A and B respectively.
Figure 2: Isometric force (F), power output (P), ATPase rate (R) and myosin filaments density (D) measured at 1.5xLref normalized by the corresponding values at Lref. Asterisks denote statistical difference from reference (p
