Departments of Anesthesiology and Physiology and Biophysics, Mayo Clinic and Mayo Foundation, Rochester, Minnesota 55905
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ABSTRACT |
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This study
determined whether the time-dependent decline in the rate of ATP
hydrolysis by actomyosin ATPase during sustained isometric force can
occur in the absence of a time-dependent decline in regulatory myosin
light chain (rMLC) phosphorylation in Triton X-100-permeabilized canine
tracheal smooth muscle. Maximal activation with 10 µM
Ca2+ induced sustained increases
in isometric force, stiffness, and rMLC phosphorylation; however, the
increase in the ATP hydrolysis rate was initially high but then
declined to a steady-state level above that of the unstimulated muscle
(basal 31.8 ± 5.8 nmol · cm3 · s
1;
peak 81.4 ± 11.3 nmol · cm
3 · s
1; steady-state
62.2 ± 9.1 nmol · cm
3 · s
1).
Activation of strips in which the rMLC was irreversibly and maximally
thiophosphorylated with adenosine
5'-O-(3-thiotriphosphate) also
induced sustained increases in isometric force and stiffness but a
nonsustained increase in ATP hydrolysis rate. There was no significant
difference in the peak or steady-state isometric force, stiffness, or
ATP hydrolysis rate or in the steady-state maximum unloaded shortening
velocity between strips activated by 10 µM
Ca2+ or rMLC thiophosphorylation
(0.058 ± 0.016 and 0.047 ± 0.011 muscle lengths/s,
respectively). Mechanisms other than changes in rMLC phosphorylation
contribute to the time-dependent decline in actomyosin ATPase activity
during sustained activation of canine tracheal smooth muscle.
adenosine 5'-triphosphate; cross bridges; actomyosin adenosine 5'-triphosphatase activity; maximum unloaded shortening velocity; latch bridges
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INTRODUCTION |
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CONTRACTION OF SMOOTH MUSCLE requires cyclic attachment and detachment of the myosin head to actin (i.e., cross-bridge cycling) and the hydrolysis of ATP by actin-activated myosin ATPase (actomyosin ATPase). Models describing the regulation of cross-bridge cycling kinetics in smooth muscle originate from observations that although the increase in isometric force is sustained after activation, cytosolic Ca2+ concentration ([Ca2+]i) (31), phosphorylation of the 20-kDa regulatory myosin light chain (rMLC), and maximum unloaded shortening velocity (Vmax) each rapidly reach maximal values but then decline to sustained suprabasal levels (6, 17, 18). This slowing of Vmax has been attributed to the transformation of rapidly cycling cross bridges to slowly cycling cross bridges. It has been postulated that in addition to regulating cross-bridge recruitment, the level of rMLC phosphorylation is the primary determinant of the cross-bridge cycling rate and Vmax during smooth muscle contraction (45, 48). In a four-state kinetic model of cross-bridge regulation in smooth muscle (17, 18), it was proposed that rMLC phosphorylation by myosin light chain kinase (MLCK) induces the formation of rapidly cycling, force-generating cross bridges. The subsequent dephosphorylation of rMLC by smooth muscle protein phosphatases as [Ca2+]i declines slows the cross-bridge cycling rate and hence Vmax, producing the "latch" state (i.e., attached, dephosphorylated cross bridges) such that isometric force is maintained at a lower tension cost.
Consistent with this model, in many types of smooth muscle, the rate of ATP hydrolysis is initially high during isometric force development (reflecting rapidly cycling cross bridges) and then decreases throughout sustained isometric force (reflecting slowly cycling, dephosphorylated cross bridges) (2, 26, 27, 41, 48). These changes in the rate of ATP hydrolysis correspond to changes in [Ca2+]i and rMLC phosphorylation, which also decline to sustained suprabasal levels after initial peaks. However, a similar profile of the ATP hydrolysis rate during sustained isometric force has been observed in permeabilized smooth muscle preparations in which [Ca2+]i and rMLC phosphorylation are constant after the initiation of contraction, a finding not predicted by the model (27, 49). Other studies (30, 41) have also suggested that the cross-bridge cycling rate can be modulated without concomitant changes in [Ca2+]i and rMLC phosphorylation, including studies (13, 15, 28) of intact airway smooth muscle. However, the ATP hydrolysis rate has not been directly measured in airway smooth muscle so that the predictions of the model have not been directly tested in this tissue.
One experimental technique used to examine the role of attached,
dephosphorylated cross bridges employs thiophosphorylation of rMLC by
adenosine
5'-O-(3-thiotriphosphate)
(ATPS) (24). Thiophosphorylation produces cross bridges that are
resistant to dephosphorylation by smooth muscle protein phosphatases.
Thus any experimental behavior observed after thiophosphorylation
cannot be attributed to attached, dephosphorylated cross bridges. These data are also not available for airway smooth muscle.
The purpose of the present study was to test the applicability of the
four-state kinetic model of cross-bridge cycling to Triton
X-100-permeabilized canine tracheal smooth muscle. According to the
predictions of this model, we hypothesized that activation with a
constant concentration of free
Ca2+ would produce a sustained
increase in rMLC phosphorylation, isometric force, and ATP hydrolysis
rate. We further hypothesized that for a given level of isometric
force, the presence of dephosphorylated, slowly cycling cross bridges
formed during activation with a constant concentration of free
Ca2+ would result in a lower
Vmax compared
with that measured in muscle strips activated by irreversible rMLC
thiophosphorylation produced by ATPS, which produces only
phosphorylated, rapidly cycling cross bridges.
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METHODS AND MATERIALS |
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Experimental Techniques
Tissue preparation. Mongrel dogs (15-23 kg) of either sex were anesthetized with an intravenous injection of pentobarbital sodium (30 mg/kg) and exsanguinated. A 10- to 15-cm portion of the extrathoracic trachea was excised and immersed in chilled physiological salt solution (PSS) of the following composition (in 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 5-7 mm, wet weight 65-145 µg) were cut from the sheet of tissue with a dissecting microscope.Smooth muscle mechanics. Isometric force, stiffness, and ATP hydrolysis rate were measured simultaneously in muscle strips with a commercially available system (model PH2, Scientific Instruments, Heidelberg, Germany) (24). Vmax was also measured with this system 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 with PSS (37°C) aerated with 94% O2-6% CO2. One end of the strips was anchored via stainless steel microforceps to a calibrated force transducer and the other end via stainless steel microforceps to a servo-controlled stepper motor. The force transducer and stepper motor attachments were not compliant. The resolutions of the force and length transducers were 0.01 mN and 0.001 mm, respectively. During a 2-h equilibration period, the length of the muscle strips was incrementally increased after repeated isometric contractions (of 2- to 3-min duration) induced by 1 µM acetylcholine (ACh) until isometric force was maximal [optimal length (Lo)]. The strips were then cooled to 25°C and maintained at Lo for the remainder of the experiment. Cross-sectional area of the muscle strip was calculated based on measurement of Lo and tissue weight.
Tissue stiffness was measured by imposing sinusoidal length oscillations of 0.5% Lo at a frequency of 50 Hz. Preliminary studies determined that the relationship between the frequency of length oscillation (at 0.5% Lo) and stiffness was sigmoidal, with maximal stiffness at 50 Hz and a phase angle between force and length < 10%. Tissue stiffness was normalized to cross-sectional area and is expressed as Young's modulus.
Isotonic shortening velocities under various loads were determined by the quick-release, afterload-clamp method (23). 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 Lo /s) were determined from the length change during a period of 100-150 ms after release to avoid effects of series elastic recoil. To calculate Vmax, the force-velocity relationship was fit by a nonlinear, least-squares method to the Hill equation.
ATP hydrolysis rate measurements. The
rate of ATP hydrolysis was measured with an enzymatic-coupled -NADH
fluorometric technique in which the regeneration of ATP from ADP and
phospho(enol)pyruvate is catalyzed
by pyruvate kinase (24, 40, 49). This reaction is coupled to the
oxidation of NADH to NAD+ and to
the reduction of pyruvate to lactate; these reactions are catalyzed by
lactate dehydrogenase. For each mole of ADP produced, 1 mol of NADH, a
fluorescent compound, is oxidized to
NAD+, a nonfluorescent compound.
Thus the rate of decrease in NADH fluorescence is proportional to the
rate of ATP hydrolysis by the tissue.
Light from a xenon lamp was monochromatically filtered to restrict excitation light to a 340-nm wavelength and focused with a high-aperture objective onto the quartz cuvette. Fluorescence emitted by the solution in the cuvette was filtered at 500 ± 5 nm and detected with a photomultiplier assembly. Illumination intensity of the excitation light was detected with an absorbance monitor and used to correct for fluctuations in excitation light intensity.
The cuvette was rapidly flushed for 100 ms every 4.9 s with fresh
solution (~50 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 the 4.9-s period was measured. NADH fluorescence
was determined for known concentrations of NADH (90 and 180 µM)
immediately before 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 tissue was activated with 10 µM free
Ca2+, indicating an increase in
ATP hydrolysis rate. ATP hydrolysis rates are expressed as nanomoles
per second and were normalized for tissue volume.
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Permeabilization procedure and
solutions. Muscle strips were either superfused (in
studies of muscle mechanics and ATP hydrolysis rate) or incubated (in
studies of rMLC phosphorylation) for 20 min with a relaxing solution
containing 10% Triton X-100 (25°C). The composition of the
relaxing solution was (in mM) 85 K+, 2.1 Mg2ATP, 4 EGTA, 20 imidizole, and
1 dithiothreitol (DTT). After permeabilization, the strips were washed
with the relaxing solution for 15 min to remove excess Triton X-100.
Solutions of varying free ion concentrations were prepared with a
previously described computer-generated algorithm (7) and also
contained 1 µM calmodulin. Rigor solutions contained (in mM) 85 K+, 0 Mg2ATP, 0 free
Mg2+, 4 EGTA, 20 imidizole, 1 DTT,
and either 1 nM (low-Ca2+ rigor
solution) or 10 µM (high-Ca2+
rigor solution) free Ca2+. The
rigor solutions also contained 2.1 mM magnesium adenosine 5'-O-(3-thiotriphosphate)
(Mg2ATPS) for selected
experiments in which the rMLC was thiophosphorylated. The pH of all
solutions was buffered to 7.1 with proprionic acid, and the ionic
strength was kept constant at 0.15 M by adjusting the concentration of potassium proprionate. In addition to these constituents, solutions for
ATP hydrolysis rate measurements also contained 5 mM
phospho(enol)pyruvate, 0.18 mM NADH,
140 U/ml of lactate dehydrogenase, and 100 U/ml of 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 (4, 15). 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
the tissues were permeabilized with Triton X-100. After experimental
interventions, the permeabilized strips were flash-frozen by rapid
immersion in a dry ice-acetone slurry containing 10% (wt /vol)
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). The proteins were separated by
glycerol-urea polyacrylamide gel [10% (wt /vol) acrylamide,
0.5% (wt /vol) bis-acrylamide, 40% (vol/vol) 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
thioglycate. The gels were subjected to preelectrophoresis 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 were injected into the wells and
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 a buffer of 25 mM Na2HPO4 (pH 7.6). The nitrocellulose sheets were washed twice with 10 mM Tris-buffered saline containing 5% (wt /vol) bovine serum albumin for 1 h (25°C) before being labeled with polyclonal, affinity-purified rabbit anti-rMLC antibody (4, 15). The anti-rMLC antibody was detected with 125I-labeled protein A (DuPont, Boston, MA).
The unphosphorylated and phosphorylated bands of rMLC were visualized by phosphorimage analysis (PhosphorImager, Molecular Dynamics, Sunnyvale, CA) and quantified by ImageQuant software (Molecular Dynamics). After local background subtraction, rMLC phosphorylation was calculated by integrating the bands corresponding to the mono- and diphosphorylated rMLCs as a fraction of the total integration of both the phosphorylated and unphosphorylated rMLCs.
Experimental Protocols
Effect of wortmannin on isometric force and ATP hydrolysis rate. This experimental protocol (Fig. 2) determined whether the basal ATP hydrolysis rate is modulated by free Ca2+ and whether the suprabasal ATP hydrolysis rate requires rMLC phosphorylation by MLCK and actomyosin cross-bridge formation. After permeabilization, muscle strips were superfused for 15 min with the low-Ca2+ rigor solution either with or without (control) 10 µM wortmannin, an inhibitor of MLCK (33). Then the muscle strips were superfused with relaxing solution for 10 min to remove excess wortmannin and to equilibrate the strips with Mg2ATP, calmodulin, and the enzymes required for ATP hydrolysis rate measurements. The muscle strips were sequentially activated with 0.7, 1, and 10 µM free Ca2+ for 10 min at each concentration and then deactivated with the relaxing solution to allow the ATP hydrolysis rate and isometric force to return to baseline values (Fig. 2).
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Time course for isometric force, stiffness, ATP hydrolysis rate, and rMLC phosphorylation. This experimental protocol determined the time course of the relationship among isometric force, stiffness, ATP hydrolysis rate, and rMLC phosphorylation during maximal activation by 10 µM free Ca2+. Isometric force, stiffness, and ATP hydrolysis rate were measured simultaneously in individual muscle strips, and rMLC phosphorylation was measured in separate muscle strips prepared from different dogs. After permeabilization and equilibration in the relaxing solution, the strips were activated with 10 µM free Ca2+ for ~10 min and then deactivated with the relaxing solution for an additional 5 min to allow isometric force, stiffness, and the ATP hydrolysis rate to return to baseline values. All solutions contained 1 µM calmodulin and the enzymes required for ATP hydrolysis rate measurements.
To measure the time course for rMLC phosphorylation, seven muscle strips were placed in 5-ml polyethylene vials and permeabilized. Then, after all strips were equilibrated in the relaxing solution for 15 min, one strip was flash-frozen in the relaxing solution for baseline rMLC phosphorylation measurements. Five strips were incubated with 10 µM free Ca2+ and then flash-frozen for rMLC phosphorylation measurements at 0.5, 1, 2, 5, and 10 min after activation. The seventh strip was first incubated with 10 µM free Ca2+ for 10 min, then incubated in the relaxing solution for an additional 5 min, and finally flash-frozen for rMLC phosphorylation measurements. All solutions contained 1 µM calmodulin.
Time course for isometric force, stiffness, and ATP
hydrolysis rate during rMLC thiophosphorylation. This
experimental protocol determined the time course for the relationship
among isometric force, stiffness, and ATP hydrolysis rate during
maximal activation by irreversible rMLC thiophosphorylation with
ATPS. Permeabilized muscle strips were superfused with the
low-Ca2+ rigor solution for 20 min
to remove Mg2ATP and then with the high-Ca2+ rigor solution
containing 2.1 mM ATP
S for 10 min to thiophosphorylate the rMLC.
Preliminary studies determined that superfusion with this solution for
10 min produced >95% rMLC thiophosphorylation, which was constant
throughout the 10-min time course of the protocol. Then the strips were
superfused with the low-Ca2+ rigor
solution containing 10 µM wortmannin for 10 min to ensure complete
inhibition of MLCK and remove ATP
S. Finally, the strips were
superfused with the relaxing solution containing 2.1 mM
Mg2ATP. All solutions contained 1 µM calmodulin and the enzymes required for ATP hydrolysis rate measurements.
Vmax during maximal activation by free
Ca2+ or rMLC
thiophosphorylation.
This experimental protocol determined
Vmax 10 min after
maximal activation by 10 µM free
Ca2+ or 10 min after activation
with ATP after rMLC thiophosphorylation with ATPS. Measurements at
several different loads were performed in random order in each strip
during a single contraction because preliminary studies demonstrated
that repeated isometric contractions induced by 10 µM free
Ca2+ were not reproducible. In
support of this procedure, preliminary studies demonstrated that
multiple quick-release measurements performed during sustained
contraction at a single load produced reproducible shortening velocity
measurements. All solutions contained 1 µM calmodulin. As a quality
control measure,
Vmax was also
determined in each muscle strip before permeabilization at 37°C 10 min after stable contractions induced by 10 µM ACh. The
Vmax measured
under these conditions (before permeabilization) did not differ between strips later activated after permeabilization by 10 µM free
Ca2+ or rMLC thiophosphorylation
(0.15 ± 0.04 and 0.17 ± 0.04 Lo/s, respectively).
Chemicals
The polyclonal affinity-purified rabbit anti-rMLC antibody was a generous gift from Dr. Susan J. Gunst (Department of Physiology and Biophysics, Indiana University School of Medicine, Indianapolis, IN). ATP disodium salt was purchased from Research Organics (Cleveland, OH). Protein A was purchased from DuPont. TCA was purchased from Fisher Scientific (Fair Lawn, NJ). Pyruvate kinase and lactate dehydrogenase were purchased from Boehringer Mannheim (Indianapolis, IN). Calcium oxide and magnesium oxide were purchased from Aldrich (Milwaukee, WI). All other drugs and chemicals were purchased from Sigma (St. Louis, MO). All drugs and chemicals were dissolved in distilled water.Statistical Analysis
Data are expressed as means ± SE; n is the number of dogs. All comparisons were made by Student's unpaired t-test. A P value < 0.05 was considered significant. Values of force, stiffness, and ATPase hydrolysis rate were quantified by averaging over 60-s (baseline and steady-state values) or 30-s (peak values) intervals. ![]() |
RESULTS |
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Effect of Wortmannin on Isometric Force and ATP Hydrolysis Rate
The basal ATP hydrolysis rate (at 1 nM free Ca2+) was 26 ± 1.1 nmol · cm
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Time Course for Isometric Force, Stiffness, rMLC Phosphorylation, and ATP Hydrolysis Rate
Increasing the free Ca2+ from 1 nM to 10 µM induced sustained increases in isometric force (Fig. 4, top trace), stiffness (Fig. 4, second trace), and rMLC phosphorylation (Fig. 4, bottom trace). In contrast, the increase in ATP hydrolysis rate produced by Ca2+ activation was not sustained (Fig. 4, third trace). The rate reached a maximal value at ~2 min (baseline and peak values of 31.8 ± 5.8 and 81.4 ± 11.3 nmol · cm
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Time Course for Isometric Force, Stiffness, and ATP Hydrolysis Rate During rMLC Thiophosphorylation
Similar to the pattern observed during activation by 10 µM free Ca2+, activation of muscle strips in which the rMLC was thiophosphorylated by ATP
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Vmax During Maximal Activation by Free Ca2+ or rMLC Thiophosphorylation
There was no significant difference in Vmax measured 8 min after activation between strips activated by 10 µM free Ca2+ or rMLC thiophosphorylation (Fig. 7).
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DISCUSSION |
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The most significant finding of this study is that whereas isometric force induced by either rMLC phosphorylation or thiophosphorylation is sustained during activation of permeabilized airway smooth muscle, actomyosin ATPase activity declines from a peak value. This finding is not predicted by the four-state kinetic model of cross-bridge cycling and suggests that factors other than dephosphorylation of rMLC may contribute to the time-dependent decline in actomyosin ATPase activity during sustained contraction of airway smooth muscle.
To interpret our findings, it was necessary to first determine whether
enzymes other than actomyosin ATPase and MLCK contribute to the
increase in the ATP hydrolysis rate induced by free
Ca2+. A previous study (25) has
demonstrated that the basal ATP hydrolysis rate in permeabilized smooth
muscle is due primarily to ATPases that are tightly bound to the plasma
membrane (e.g., ecto-ATPases) and are markedly inhibited by Triton
X-100. It is possible that changes in tissue stress during contraction
will alter the diffusional access of ATP to ATPases and thus contribute to a time-dependent decline change in the suprabasal ATP hydrolysis rate. To evaluate this possibility, we measured the basal ATP hydrolysis rate in five permeabilized canine tracheal smooth muscle strips that had been passively stretched to 1.4 Lo. This degree of stretch produced a passive force of ~43% of the active isometric force induced by maximal activation with 10 µM free
Ca2+ or rMLC thiophosphorylation.
The basal ATP hydrolysis rate measured in strips stretched to 1.0 and
1.4 Lo were 31.8 ± 5.8 and 32.0 ± 5.0 nmol · cm3 · s
1, respectively.
In the present study, wortmannin, an inhibitor of MLCK, completely blocked both the initial peak and steady-state increase in ATP hydrolysis rate and isometric force during activation with 0.7 and 1 µM free Ca2+. These data suggest that both the initial peak and steady-state increase in ATP hydrolysis rate require rMLC phosphorylation by MLCK and the formation of actomyosin cross bridges. This finding is consistent with that previously reported (49) for Triton X-100-skinned swine carotid artery where the calmodulin antagonist W-7 inhibited Ca2+-stimulated isometric force and Ca2+-activated ATP hydrolysis. Furthermore, in preliminary studies, neither the basal ATP hydrolysis rate nor the increase in ATP hydrolysis rate induced by 1 µM free Ca2+ were affected by thapsigargin (an inhibitor of sarcoplasmic reticulum Ca2+-ATPase), ouabain, (an inhibitor of Na+-K+-ATPase), oligomycin (an inhibitor of mitochondrial Ca2+-ATPase), or quercetin (an inhibitor of plasma membrane Ca2+-ATPase) (data not shown). These observations are consistent with those previously reported for other permeabilized smooth muscle preparations (27, 49). Thus the only apparent sources of the Ca2+-activated suprabasal ATP hydrolysis in this preparation are MLCK and actomyosin ATPase.
The four-state kinetic model of cross-bridge regulation postulates that the initial phosphorylation of rMLC by MLCK induces the formation of rapidly cycling, force-generating cross bridges (17, 18). As [Ca2+]i declines during activation of intact smooth muscle, MLCK activity decreases and rMLC is dephosphorylated by smooth muscle protein phosphatases. Dephosphorylation of attached cross bridges decreases the cycling rate (reflected by a decrease in Vmax) so that isometric force is maintained at a lower tension cost. The four-state model implies that 1) there is a time-dependent correlation among [Ca2+]i, the level of rMLC phosphorylation, and actomyosin ATPase activity; 2) ATP hydrolysis by MLCK represents a significant fraction of the suprabasal increase in the ATP hydrolysis rate during contraction (19); and 3) the formation of slowly cycling cross bridges requires rMLC dephosphorylation and thus requires smooth muscle protein phosphatase activity. Although much experimental data have accumulated in support of this theory (for a review, see Ref. 32), other studies in intact smooth muscle have demonstrated that the cross-bridge cycling rate can be modulated without changes in rMLC phosphorylation (30), including studies of intact airway smooth muscle (10, 11, 15, 28).
In the present study, a sustained increase in free Ca2+ concentration induced sustained increases in rMLC phosphorylation and isometric force. These findings are consistent with previous studies of permeabilized smooth muscle (24, 49) and are consistent with the predictions of the four-state model in that rMLC phosphorylation remains constant under these conditions of a constant free Ca2+ concentration. However, the increase in ATP hydrolysis rate was initially high but then gradually declined to a steady-state level that was greater than that of the unstimulated muscle (i.e., was not sustained). Thus the sustained increase in rMLC phosphorylation induced by a constant free Ca2+ concentration did not correlate with the time-dependent decline in ATP hydrolysis rate after the initial peak, a finding that is not predicted by the four-state model. Similar results have also been reported in permeabilized vascular (49) and gut (27) smooth muscles. We also found no significant difference in the magnitude or time course of the suprabasal ATP hydrolysis rate between tissues activated by free Ca2+ and tissues activated by rMLC thiophosphorylation. In addition, there was no significant difference in steady-state isometric force or stiffness between tissues activated by these two modes, thus permitting comparisons at similar numbers of attached cross bridges.
Three main conclusions can be drawn from our observations. First, the fact that the profile of ATP hydrolysis rate was similar during activation by Ca2+ and rMLC thiophosphorylation suggests that the predominant source of ATP hydrolysis in this preparation is actomyosin ATPase. If ATP hydrolysis by MLCK was a significant fraction of the total ATP hydrolysis during Ca2+ activation, ATP hydrolysis should have been significantly lower with rMLC thiophosphorylation. Indeed, this tendency was present but is not significant. Thus although we cannot exclude that some portion of the total suprabasal ATP hydrolysis rate during Ca2+ activation was due to MLCK, the majority was due to actomyosin ATPase. This finding is consistent with prior work in skinned vascular (49) and gut (20, 24) smooth muscle. Second, the ATP hydrolysis rate decreased with time after an initial peak with both modes of activation. The decline observed during rMLC thiophosphorylation, a condition not examined in prior studies, confirms that such decreases cannot be associated with smooth muscle protein phosphatase activity and the dephosphorylation of attached cross bridges. Finally, the fact that isometric force, stiffness, and Vmax were not different in tissues activated by these two means suggests that the number of attached cross bridges and their cycling rates were not different. If attached, dephosphorylated cross bridges were present during Ca2+ activation, Vmax should have been less than during rMLC thiophosphorylation, a condition that does not permit cross-bridge dephosphorylation. Similar findings have been reported in skinned guinea pig taenia coli (20).
The mechanism(s) responsible for the nonsustained increase in actomyosin ATPase activity under these experimental conditions is not known. However, several models have been proposed. First, there is evidence that actomyosin ATPase activity and the cross-bridge cycling rate may be under the biochemical regulation of another Ca2+- and phosphorylation-dependent system (16, 22, 38). Biochemical studies, including in vitro motility assays, have suggested that the actin-associated proteins caldesmon and calponin may regulate actomyosin ATPase activity (34, 39, 47). For example, whereas unphosphorylated caldesmon inhibited actomyosin ATPase activity in vitro, no inhibition was observed with phosphorylated caldesmon (34). It has also been shown that unphosphorylated caldesmon (1, 35, 36) or calponin (21, 44) inhibit isometric force in skinned smooth muscle preparations and that both proteins are phosphorylated during smooth muscle contraction (14, 29, 37).
However, it is likely that phosphorylation of caldesmon and calponin
was constant during activation of the thiophosphorylated strips with
ATP in the absence of Ca2+ because
phosphorylation of both proteins is regulated largely by
Ca2+ (12, 16, 22, 38).
Furthermore, it is likely that ATPS also irreversibly
thiophosphorylated caldesmon and calponin during incubation of the
strips in the high-Ca2+ rigor
solution because both proteins are readily thiophosphorylated in vitro
(1, 35, 36). Thus the presence of a nonsustained increase in ATP
hydrolysis during activation of thiophosphorylated strips with ATP
suggests that the similar decrease in the ATP hydrolysis rate during
Ca2+ activation is not dependent
on progressive caldesmon or calponin phosphorylation.
A second alternative model is the intriguing possibility that during smooth muscle contraction, changes in load on the cross bridges may account for the time-dependent change in actomyosin ATPase activity because the cross-bridge cycling rate is load dependent (23). The change in load on the cross bridges could be due to 1) active shortening of the myocytes within the compliant extracellular connective tissue matrix of the multicellular preparation (i.e., the myocytes are not themselves isometrically activated) or 2) actin (3, 43) or myosin (9) reorganization because both proteins are known to undergo dynamic change between filamentous and monomeric conformations. For example, biochemical studies have demonstrated that there is more myosin incorporated into thick filaments during activation than in unstimulated tissue (46) and that this process is augmented by rMLC phosphorylation (5, 42). Based on these observations, it has been proposed that a series-to-parallel transition in the arrangement of the actin-myosin contractile units could account for the time-dependent decline in shortening velocity and, hence, actomyosin ATPase activity during sustained isometric force (9). It is possible that the load on the cross bridges varies after activation such that the initial peak increase in actomyosin ATPase activity observed during isometric force development (i.e., within the first 2 min after Ca2+ activation) may correspond to a period of relative low load on the cross bridges. Conversely, the sustained phase of isometric force may correspond to a period of increasing load on the cross bridges and, hence, a decline in the cross-bridge cycling rate and ATP hydrolysis rate. In support of this possibility, Sieck et al. (40) recently demonstrated in detergent-skinned porcine tracheal smooth muscle that the ATP hydrolysis rate increases with power output during isovelocity contractions induced by free Ca2+. The ATP hydrolysis rate increased by approximately sixfold at peak power output, which was achieved at a load corresponding to ~33% of maximal isometric force. Thus energy utilization of smooth muscle increases in proportion to work as originally demonstrated for striated muscle (8).
It should be emphasized that the results of this study do not preclude the possibility that attached, dephosphorylated cross bridges can form under conditions of changing intracellular Ca2+ that exist in intact airway smooth muscle stimulated with receptor agonists such as ACh. Furthermore, the applicability of results in permeabilized preparations to intact muscles has been questioned. In particular, it has been suggested that the reduced temperatures employed in permeabilized preparations to minimize experimental deterioration may preferentially reduce smooth muscle protein phosphatase activity relative to MLCK activity (32). This scenario could discourage formation of attached, dephosphorylated cross bridges. However, we note that rMLC phosphorylation values measured during maximal Ca2+ activation in our preparation at 25°C are similar to those previously reported in maximally stimulated, intact airway smooth muscle studied at 37°C (15), suggesting a similar balance of smooth muscle protein phosphatase and MLCK activities in the two conditions. In any event, it is apparent from our data that actomyosin ATPase activity declines during sustained contraction of airway smooth muscle even in the absence of any decreases in rMLC phosphorylation.
In summary, this study demonstrates that during activation of Triton X-100-permeabilized canine tracheal smooth muscle by sustained rMLC phosphorylation or thiophosphorylation, actomyosin ATPase activity declines from an initial peak value, whereas isometric force remains constant. This finding is not predicted by the four-state kinetic model of cross-bridge regulation. Changes in load on the cross bridges during activation may be a plausible alternative mechanism to explain this time-dependent decline in actomyosin ATPase activity.
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ACKNOWLEDGEMENTS |
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Our thanks to K. Street for expert technical assistance.
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FOOTNOTES |
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This study was supported in part by National Heart, Lung, and Blood Institute Grants HL-54757, HL-45532, and HL-33009; by a grant from Abbott Laboratories (Abbott Park, IL; to Y. S. Prakash); and the Mayo Foundation (Rochester, MN).
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. §1734 solely to indicate this fact.
Address for reprint requests and other correspondence: K. A. Jones, Mayo Clinic, 200 First St. SW, Rochester, MN 55905.
Received 8 October 1998; accepted in final form 29 March 1999.
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