A novel mechanism by which hydrogen peroxide decreases calcium sensitivity in airway smooth muscle

William J. Perkins1, Robert R. Lorenz1, Michelle Bogoger2, David O. Warner1, Christine R. Cremo2, and Keith A. Jones1

1 Departments of Anesthesiology and Physiology and Biophysics, Mayo Clinic and Mayo Foundation, Rochester, Minnesota 55905; and 2 Department of Biochemistry, University of Nevada, Reno, Reno, Nevada 89509


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The purpose of this study was to test the hypothesis that H2O2 decreases the amount of force produced by a given intracellular Ca2+ concentration (i.e., the Ca2+ sensitivity) in airway smooth muscle (ASM) in part by mechanisms independent of changes in regulatory myosin light chain (rMLC) phosphorylation. A new preparation was developed and validated in which canine ASM strips were first exposed to H2O2 and then permeabilized with 10% Triton X-100 to assess the persistent effects of H2O2 on Ca2+ sensitivity. Experiments in which H2O2 was administered before permeabilization revealed a novel mechanism that contributed to reduced Ca2+ sensitivity independently of changes in rMLC phosphorylation, in addition to an rMLC phosphorylation-dependent mechanism. The mechanism depended on factors not available in the permeabilized ASM strip or in the buffer to which the strip was exposed, since there was no effect when H2O2 was added to permeabilized strips. H2O2 treatment of a maximally thiophosphorylated purified myosin subfragment (heavy meromyosin) significantly reduced actomyosin ATPase activity, suggesting one mechanism by which the phosphorylation-independent reduction in Ca2+ sensitivity may occur.

regulatory myosin light chain phosphorylation; thiophosphorylation; permeabilized smooth muscle; reactive oxidant species; myosin; actomyosin adenosinetriphosphatase


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

REACTIVE OXIDANT SPECIES, such as H2O2, play an important physiological role in both muscle (39) and nonmuscle cells (6) and a pathophysiological role in numerous diseases, including acute lung injury, asthma, pulmonary hypertension, ischemia-reperfusion, and arthritis (6). H2O2 reversibly inhibits receptor agonist-induced contraction of both vascular (18, 19, 32) and airway (11, 12, 15, 38, 45) smooth muscle (ASM). Recently, we reported that H2O2-induced relaxation of intact ASM was caused by a reduction in the amount of force produced by a given intracellular Ca2+ concentration ([Ca2+]i, the "Ca2+ sensitivity"), as shown by the fact that the relaxation of canine ASM produced by H2O2 was accompanied by an increase in [Ca2+]i (30). The mechanism for this effect on Ca2+ sensitivity is not known.

Contraction of ASM is controlled largely by the phosphorylation of the 20-kDa regulatory myosin light chain (rMLC), resulting in the 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; see Ref. 43). The level of rMLC phosphorylation depends on the balance between the activities of myosin light chain kinase (MLCK) and smooth muscle protein phosphatase. MLCK activity is regulated by the binding of Ca2+-calmodulin complexes in response to increased [Ca2+]i produced by receptor stimulation, favoring increased rMLC phosphorylation (26). rMLC phosphorylation can also increase if the activity of smooth muscle protein phosphatase is inhibited. The reduction in Ca2+ sensitivity produced by H2O2 in our prior report of H2O2-induced relaxation of intact ASM (30) was accompanied by a decrease in rMLC phosphorylation, suggesting inhibition of Ca2+ and/or activation of smooth muscle protein phosphatases.

H2O2 may also reduce Ca2+ sensitivity by mechanisms not dependent on rMLC phosphorylation, such as by inhibiting actomyosin ATPase activity. The smooth muscle myosin head contains highly reactive cysteine thiols that result in a reduction in ATPase activity when they are covalently modified (4, 28). Illustrating what effects this might have in an analogous system, the mild oxidant nitric oxide reversibly inhibits skeletal muscle actomyosin ATPase activity, decreases Ca2+ sensitivity, and produces relaxation, effects that are fully reversible by a thiol-reducing agent (37). The possibility of a similar effect with a mild oxidant such as H2O2 in smooth muscle has not been evaluated.

The purpose of this study was to test the hypothesis that H2O2 decreases Ca2+ sensitivity in ASM in part by a previously undescribed mechanism independent of changes in rMLC phosphorylation. To achieve this aim, we developed a novel method in which ASM strips were first exposed to H2O2 and then permeabilized with Triton X-100 to assess the persistent effects of H2O2 exposure on Ca2+ sensitivity. We then used purified protein preparations to examine whether H2O2 has a direct inhibitory effect on the actin-activated myosin ATPase activity in a manner consistent with a phosphorylation-independent reduction in Ca2+ sensitivity.


    MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Tissue preparation. After Institutional Animal Care and Use Committee approval, mongrel dogs of either sex were anesthetized with an intravenous injection of pentobarbital sodium (30 mg/kg) and exsanguinated. The trachea was excised and immersed in chilled physiological salt solution (PSS) of the following composition (in mM): 110 NaCl, 26 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 under microscopic observation to make muscle strips.

Mechanical measurements. For studies of permeabilized muscle strips (5 mm long × 0.3 mm diameter) and simultaneous measurement of force and [Ca2+]i in intact (nonpermeabilized) tissue, strips were mounted in 0.1-ml cuvettes and continuously superfused with PSS (37°C) aerated with 94% O2-6% CO2. One end of the strips was anchored via stainless steel microforceps attached to a micrometer, and the other end was attached via stainless steel microforceps to a force transducer (model KG4; Scientific Instruments, Heidelberg, Germany). For rMLC phosphorylation measurements in intact tissue, muscle strips were suspended in 25-ml water-jacketed tissue baths that were filled with PSS (37°C) aerated with 94% O2-6% CO2. One end of the strips was anchored to a metal hook at the bottom of the tissue bath; the other end was attached to a calibrated force transducer (model FT03D; Grass Instrument, Quincy, MA).

In both cases, during a 3-h equilibration period, the strips were repeatedly contracted isometrically for 2 min every 10 min by maximal stimulation with 1 µM ACh. The length of the strips was increased after each stimulation until active force was maximal (optimal length, Lo). Each strip was maintained at this Lo for the remainder of the experiment.

Isometric force and fura 2 fluorescence measurements. Muscle strips were incubated with PSS (25°C) containing 5 µM fura 2-AM and aerated with 96% O2-4% CO2 for 3 h (23). Fura 2-AM was dissolved in DMSO and 0.02% cremophor. After fura 2 loading, the strips were washed with PSS (37°C) for 30-50 min to remove extracellular fura 2-AM and DMSO and to allow de-esterification of any remaining cytosolic fura 2-AM.

Fura 2 fluorescence intensity was measured by a photometric system that measures optical and mechanical parameters of isolated tissue simultaneously (16). Light from a mercury high-pressure lamp was passed through rotating bandpass filters to restrict excitation light to 340 and 380 nm. Fluorescence emitted from the strips was filtered at 450 ± 5 nm and detected by a photomultiplier (Scientific Instruments). The emission fluorescence intensities due to excitation at 340-nm (F340) and 380-nm (F380) wavelengths were measured, and the F340-to-F380 ratio was used as an index of [Ca2+]i. Absolute values of [Ca2+]i were not calculated since the dissociation constant of fura 2 for Ca2+ within the smooth muscle cytosol cannot be determined (29). In preliminary work, we found that H2O2 alone did not affect F340 and F380 fluorescence intensities when added to solutions containing Ca2+ and fura 2 (data not shown).

Permeabilization of muscle strips. The strips were either superfused (in studies of force) or incubated (in studies of rMLC phosphorylation and thiophosphorylation) for 20 min with relaxing solution containing 10% Triton X-100 (at 25°C; see Ref. 24). The composition of the relaxing solution was as follows (in mM): 85 K+, 2.1 disodium ATP (Na2ATP), 4 EGTA, and 20 imidazole. After permeabilization, the strips were washed with relaxing solution for 5 min to remove excess Triton X-100. Solutions of varying free Ca2+ concentration were prepared using a previously described algorithm (8). Rigor solutions contained (in mM) 85 K+, 0 Na2ATP, 0 free Mg2+, 4 EGTA, 20 imidazole, 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 adenosine 5'-O-(3-thio)triphosphate (ATPgamma S) for selected experiments in which the rMLC was thiophosphorylated (22). The pH of all solutions was buffered to 7.0 with propionic acid, and the ionic strength was kept constant at 0.2 M by adjusting the concentration of potassium propionate.

Determination of extent of myosin regulatory light chain phosphorylation in muscle strips. After experimental interventions, muscle strips were flash-frozen with dry ice-cooled acetone containing 10% (wt/vol) TCA and 10 mM dithiothreitol (DTT). For intact muscle, strips were frozen immediately after removal from organ baths while maintaining Lo, because we found in preliminary work that length affects rMLC phosphorylation in intact strips. Permeabilized strips for rMLC phosphorylation measurements were prepared separately according to the same procedures as for force measurements but were incubated in wells (without determination of Lo) instead of being superfused. The strips were pinned at both ends to maintain isometric conditions. After permeabilization, all conditions were identical to those present in the strips used to measure isometric force responses. In preliminary data obtained for prior studies (3, 17, 25, 48), we have shown that muscle length does not affect rMLC phosphorylation in permeabilized ASM in which [Ca2+]i is controlled.

The frozen strips were then allowed to warm to room temperature in the same solution. After TCA was washed out with acetone containing 10 mM DTT, strips were allowed to dry. rMLC was extracted as described by Gunst et al. (14), and phosphorylation was determined by glycerol-urea gel electrophoresis followed by Western blotting with a polyclonal affinity-purified rabbit anti-20-kDa rMLC antibody, as previously described (3). Unphosphorylated and phosphorylated bands of rMLC were visualized by PhosphorImage analysis (Cyclone storage phosphor system; Packard Instrument, Downers Grove, IL) using 125I-labeled protein A (New England Nuclear) to bind to the rMLC antibody. Fractional phosphorylation was calculated as the density ratio of the sum of mono- and diphosphorylated rMLC to total rMLC using OptiQuant software (version 3.0; Packard Instrument).

Purified protein preparations. Smooth muscle myosin was prepared from frozen chicken gizzards (20) obtained from Pel-Freeze (Rogers, AR). Heavy meromyosin (HMM) was obtained from a Staphylococcus aureus protease digestion of 200-400 mg of smooth muscle myosin, as described previously (21). SDS gel analysis showed that >95% pure HMM was obtained. HMM concentration was determined using an extinction coefficient (epsilon 1% at 280 nm) of 6.5. Ca2+ was prepared from frozen turkey gizzards by the method of Adelstein and Klee (1). Actin was prepared from frozen leg and back muscles of rabbits by the method of Spudich and Watt (44).

Experimental protocols. Each experimental protocol was conducted using muscle strips obtained from a different set of animals. Previous studies, including those from this laboratory, have shown that smooth muscle relaxation induced by H2O2 is partly mediated by cyclooxygenase products, which activate adenylate cyclase, and that this mechanism is completely inhibited by indomethacin (11, 12, 38). Thus, for each protocol, all strips were incubated with 10 µM indomethacin to prevent the formation of prostanoids and the increase in intracellular cAMP levels induced by H2O2 (11, 12, 30).

We found in preliminary studies, and confirmed as described below, that H2O2 did not affect Ca2+ sensitivity when added to permeabilized smooth muscle. Thus, to examine the proposed hypothesis, we developed an experimental protocol in which intact nonpermeabilized strips were briefly exposed to H2O2. H2O2 was then washed from the strips and subsequently permeabilized with Triton X-100 to allow for control of [Ca2+]i. This experimental approach was first validated by three protocols to demonstrate that the effect of H2O2 on Ca2+ sensitivity persisted after complete washout from the intact tissue and after subsequent permeabilization. Next, two protocols were conducted to determine whether the persistent effect of H2O2 on Ca2+ sensitivity after permeabilization was because of inhibition of proteins that regulated rMLC phosphorylation levels, regulated force independent of rMLC phosphorylation levels, or a combination of both processes.

Validation protocols. The first protocol (Fig. 1) determined whether H2O2 inhibited Ca2+ sensitivity when applied to quiescent (unstimulated), intact muscle strips. After strips were loaded with fura 2 and set at Lo for force development, they were maximally activated with 40 mM isotonic KCl for 10 min, inducing stable increases in both F340/F380 and force. After complete washout of isotonic KCl for ~20 min and recovery of F340/F380 and isometric force to the unstimulated baseline values, the strips were exposed to 1 mM H2O2 for 10 min. The increase in F340/F380 and isometric force values induced by H2O2 was expressed as a percentage of the steady-state values measured during maximal activation with 40 mM isotonic KCl. In a separate set of three strips mounted in organ baths, rMLC phosphorylation levels were determined during basal conditions or after stimulation with either 40 mM isotonic KCl or 1 mM H2O2 for 10 min.


View larger version (18K):
[in this window]
[in a new window]
 
Fig. 1.   Effects of H2O2 treatment on Ca2+ concentration and isometric force in intact quiescent muscle strips. Top: representative isometric force and tracings of the ratio of fluorescence at 340 nm (F340) to that at 380 nm (F380) demonstrating the experimental protocol (see Experimental protocols in MATERIALS AND METHODS for details). Tracings show the effect of H2O2 on an intact (nonpermeabilized), quiescent strip loaded with fura 2. Bottom: mean data for F340/F380 and isometric force response to 1 mM H2O2, expressed as a percentage of responses to 40 mM isotonic KCl (left), and for regulatory myosin light chain (rMLC) phosphorylation (right). Values are means ± SE, n = 6 experiments. * Significant difference from 100% by paired t-test (bottom left) or significant difference from baseline by repeated-measures ANOVA (bottom right).

The second protocol (Fig. 2) determined whether the effect of H2O2 on Ca2+ sensitivity persisted in intact tissue after complete H2O2 washout. Fura 2-loaded strips were first activated with 25 mM isotonic KCl for 15 min, which produced ~50% of maximal force. After steady-state increases in F340/F380 and isometric force were achieved, the strips were exposed to 1 mM H2O2 for 10 min. Next, H2O2 was washed from the strips during continued stimulation with 25 mM isotonic KCl. Changes in F340/F380 and isometric force were expressed as a percent change from the steady-state isotonic KCl-induced responses before exposure to H2O2 (defined as the initial value). In a separate study using tissue obtained from a different set of animals, a set of three strips set at Lo for force development was treated using the same protocol. One unstimulated strip was frozen to obtain baseline rMLC phosphorylation values, and the second strip was frozen 10 min after H2O2 washout during continuous stimulation with isotonic KCl. A third strip was stimulated with 25 mM isotonic KCl for 25 min and was not exposed to H2O2 to serve as a control for any time-dependent changes in rMLC phosphorylation during activation.


View larger version (18K):
[in this window]
[in a new window]
 
Fig. 2.   Effects of H2O2 treatment during continuous activation with 25 mM isotonic KCl on F340/F380 and isometric force in intact muscle strips. Top and middle: representative isometric force and F340/F380 tracings demonstrating the experimental protocol (see Experimental protocols for details). Tracings show responses of isometric force and F340/F380 in a fura 2-loaded intact muscle strip exposed to 1 mM H2O2 for 10 min during continuous activation with 25 mM isotonic KCl followed by washout of H2O2. Bottom: values for parameters at times A, B, and C indicated in middle force trace. Changes in F340/F380 and isometric force were expressed as percentage changes from the steady-state isotonic KCl-induced responses before exposure to H2O2 (defined as the initial value). Values are means ± SE, n = 5. * Significant difference from initial (100%) value, P < 0.05.

A final validation protocol (Fig. 3) determined whether the effect of H2O2 on Ca2+ sensitivity in intact tissue persisted after complete washout and subsequent permeabilization. After contraction at Lo with 1 µM ACh to determine maximal isometric force, unstimulated intact muscle strips were exposed to 0 (control) or 1 mM H2O2 for 10 min, followed by complete H2O2 washout with PSS for 10 min. The strips were then permeabilized (requiring 25 min) and maximally activated with 10 µM free Ca2+ (see Fig. 3). The isometric force induced by 10 µM free Ca2+ was expressed as a percentage of the maximal isometric force induced by 1 µM ACh in the intact tissue.


View larger version (14K):
[in this window]
[in a new window]
 
Fig. 3.   Effects of H2O2 treatment on Ca2+-activated isometric force of permeabilized muscle strips that were treated with H2O2 before permeabilization. Representative isometric force tracings demonstrating the experimental protocol (see Experimental protocols for details). After initial contraction with 1 µM ACh, a muscle strip was exposed to 0 (control, solid line) or 1 (dashed line) mM H2O2 for 10 min. PSS, physiological salt solution. Inset: responses to 10 µM free Ca2+ as a percentage of the initial response to 1 µM ACh; values are means ± SE, n = 6. * Significant difference from control by paired t-test.

Dependence of H2O2-induced effects in permeabilized tissue on rMLC phosphorylation. Two experimental protocols were performed to determine if the effects of H2O2 observed in the permeabilized preparation depended on changes in rMLC phosphorylation. In the first protocol, strips were pinned in wells to maintain isometric conditions for rMLC phosphorylation measurements and then treated using a similar protocol to that represented in Fig. 3. After stimulation with 1 µM ACh, strips were exposed to 0 (control), 1, or 10 mM H2O2 for 10 min and then permeabilized (requiring 25 min) after complete H2O2 washout. After permeabilization, rMLC phosphorylation was measured 10 min after stimulation with either 0.9 or 10 µM free Ca2+ (see data for Fig. 4).


View larger version (19K):
[in this window]
[in a new window]
 
Fig. 4.   rMLC phosphorylation in permeabilized smooth muscle strips exposed before permeabilization to 1 (A) or 10 (B) mM H2O2 according to the protocol depicted in Fig. 3 (see text for details). Controls did not receive H2O2. Values are means ± SE, n = 7 for 1 mM H2O2 and n = 5 for 10 mM H2O2. * Significant difference from control by paired t-test.

In the second protocol, strips were set at Lo for isometric force measurements and treated using a similar protocol to that represented in Fig. 5. After maximal activation with 1 µM ACh, the strips were exposed to either 0 (control) or 1 mM H2O2 for 10 min and then permeabilized (requiring 25 min) after complete H2O2 washout. The strips were then superfused with low-Ca2+ rigor solution for 5 min, followed by high-Ca2+ rigor solution for 15 min to produce maximal rMLC thiophosphorylation. Thiophosphorylation produces actomyosin cross bridges that are resistant to dephosphorylation by smooth muscle protein phosphatases. Finally, the strips were superfused for 10 min with relaxing solution containing 1 nM free Ca2+ and 2.1 mM Na2ATP to induce actomyosin cross-bridge cycling and Ca2+- and rMLC phosphorylation-independent isometric force. The isometric force induced by 2.1 mM Na2ATP was expressed as a percentage of the maximal isometric force induced by 1 µM ACh in the intact tissue. A separate set of strips was pinned in wells to maintain isometric conditions to measure the level of rMLC thiophosphorylation achieved during exposure to the high-Ca2+ rigor solution using the same protocol.


View larger version (14K):
[in this window]
[in a new window]
 
Fig. 5.   Representative isometric force tracings demonstrating the experimental protocol used to produce thiophosphorylation (see text for details). After initial contraction with 1 µM ACh, a muscle strip was exposed to 0 (control, solid line) or 1 (dashed line) mM H2O2 for 10 min. Next, the strips were washed with PSS for 10 min to remove H2O2 and permeabilized with 10% Triton X-100 for 25 min. Finally, after superfusion for 5 min with low-Ca2+ rigor (LCR) solution (see text for composition) followed by 10 min with high-Ca2+ rigor (HCR) solution (see text for composition), the strips were activated with 2.1 mM ATP to initiate actomyosin cross-bridge cycling and isometric force development.

Treatment of thiophosphorylated HMM with peroxide and ATPase measurements in the presence of actin. To further characterize rMLC-independent H2O2 effects on this system, we directly measured actomyosin ATPase activity. We studied this using purified HMM, which is a soluble proteolytic fragment of myosin containing two myosin heads adjoined by the hinge region. The regulatory light chains of HMM were thiophosphorylated by incubation in 10 mM MOPS (pH 7.0), 0.1 mM EGTA, 50 mM NaCl, 2.5 mM MgCl2, 2.5 mM CaCl2, 4 µg/ml calmodulin, 1 mM ATPgamma S, and 30 µg/ml MLCK at 25°C for 1 h, followed by overnight incubation on ice (7). Thiophosphorylated HMM (HMM-P) was separated from reagents by spinning the sample through a 5-ml buffer exchange column (36) prepared with Sephadex G-50-80 resin in nonreducing buffer [50 mM NaCl and 10 mM MOPS (pH 7.0)]. The extent of thiophosphorylation before and after H2O2 treatment was verified by 8 M urea gel electrophoresis, which can detect 5% or greater unphosphorylated rMLC (7). The ratio of thiophosphate to HMM was at least 1.9:1 (gels not shown) for all samples studied.

HMM-P in nonreducing buffer was treated with either 0 or 1 mM H2O2 for 30 min at 37°C and immediately spun through a buffer exchange column in nonreducing buffer to remove H2O2. ATPase measurements were performed immediately in nonreducing buffer at 25°C. A previously described stopped-flow single-turnover method using the fluorescent ATP analog methylanthraniloyl ATP (Molecular Probes) was used to determine the steady-state ATPase activity of HMM-P (1-2 mg/ml) in the presence of 5 µM F-actin, as previously described (5, 7). Steady-state rates were calculated from the two amplitudes (A1 and A2; expressed as percent of total) and the two rates (r1 and r2) obtained from a double-exponential fit to a biphasic decay using the following equation
steady-state rate (s<SUP>−1</SUP>) = <IT>A</IT><SUB>1</SUB>(r<SUB>1</SUB>) + <IT>A</IT><SUB>2</SUB>(r<SUB>2</SUB>) (1)
The steady-state ATPase activity of samples treated with 0 mM H2O2 in the above protocol did not differ significantly from that obtained for untreated HMM-P and ranged from 1 to 1.2 s-1.

Materials. The polyclonal affinity-purified rabbit anti-20-kDa rMLC antibody was a generous gift of Dr. Susan J. Gunst (Dept. of Physiology and Biophysics, Indiana University School of Medicine, Indianapolis, IN). Na2ATP was purchased from Research Organics (Cleveland, OH). All other drugs and chemicals were purchased from Sigma Chemical (St. Louis, MO).

Statistical analysis. Data are expressed as means ± SE; n represents the number of dogs. Paired t-tests were used to compare two groups. Repeated-measures ANOVA was used to compare multiple groups, and the Dunnett's test was used for post hoc comparisons. A value of P < 0.05 was considered to be statistically significant.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Validation of preparation. Exposure of intact quiescent strips to 1 mM H2O2 produced a greater increase in F340/F380 and a smaller increase in isometric force compared with prior maximal responses to 40 mM isotonic KCl (Fig. 1). This was accompanied by a smaller increase in rMLC phosphorylation than that observed for a maximal KCl response that was not significantly different from baseline unstimulated values (Fig. 1). Thus, when applied to unstimulated strips, H2O2 inhibited Ca2+ sensitivity, in part by inhibiting the increase in rMLC phosphorylation produced for a given [Ca2+]i.

When added to intact strips contracted with 25 mM isotonic KCl, 1 mM H2O2 increased F340/F380 and caused an initial decrease in isometric force (Fig. 2, top). However, at 10 min of exposure to H2O2, force had recovered such that it was not significantly different from that measured before H2O2 exposure. When H2O2 was washed from the muscle strips during continuous stimulation with 25 mM isotonic KCl, the F340/F380 rapidly returned to pre-H2O2 values. However, force decreased to nearly baseline values measured before stimulation with isotonic KCl. At this point (10 min after H2O2 washout), the level of rMLC phosphorylation measured in a separate set of strips was significantly less in strips that had been exposed to H2O2 compared with time control strips that had not been exposed to H2O2 (11.0 ± 5.3 and 23.2 ± 2.4%, respectively, P = 0.03, n = 5). In fact, this level of rMLC phosphorylation 10 min after H2O2 washout was not significantly different from the baseline level measured in the unstimulated strips before stimulation with isotonic KCl (4.1 ± 3.2%, P = 0.16). Thus the effect of H2O2 on Ca2+ sensitivity and rMLC phosphorylation persists in intact strips 10 min after complete H2O2 washout.

When added to strips before permeabilization, H2O2 induced a small increase in isometric force (Fig. 3), as in the first experiment (Fig. 1). This exposure decreased the force response to 10 µM free Ca2+ after permeabilization by 44% compared with control strips that were not exposed to H2O2. When added to strips that were stimulated with 10 µM free Ca2+ after permeabilization (i.e., the control strips for this experiment), 1 mM H2O2 did not affect isometric force (Fig. 6). Thus the effect of H2O2 on Ca2+ sensitivity that occurred in the intact tissue (Fig. 1) and persisted after H2O2 washout for 10 min (Fig. 2) also persisted 25 min after permeabilization. By contrast, H2O2 did not affect Ca2+ sensitivity when added after the muscle strips had been permeabilized.


View larger version (9K):
[in this window]
[in a new window]
 
Fig. 6.   Representative tracing showing the lack of effect of 1 mM H2O2 on isometric force in a Triton X-100-permeabilized smooth muscle strip activated with 10 µM free Ca2+.

Dependence of H2O2 effects on rMLC phosphorylation in permeabilized muscle. rMLC phosphorylation was measured in permeabilized strips after exposing the intact tissue to 0 (control), 1, or 10 mM H2O2 before permeabilization. H2O2 did not affect rMLC phosphorylation under conditions of low free Ca2+ concentration (Fig. 4). H2O2 (10 mM but not 1 mM) significantly inhibited the increase in rMLC phosphorylation induced by either 0.9 or 10 µM free Ca2+. Thus the persistent effect of 1 mM H2O2 on Ca2+ sensitivity in permeabilized strips observed in the previous experiment (Fig. 3; as assessed by isometric force responses) was produced by mechanisms independent of changes in rMLC phosphorylation. For 10 mM H2O2, inhibition of the Ca2+-dependent increase in rMLC phosphorylation was also present.

To confirm the existence of H2O2 effects on Ca2+ sensitivity that are independent of changes in rMLC phosphorylation, ATPgamma S was used to produce stable, maximal rMLC thiophosphorylation in the permeabilized preparation (Fig. 5). The isometric force induced by 2.1 mM ATP in muscle strips in which the rMLC was irreversibly, maximally thiophosphorylated (Fig. 7) was similar to that induced by maximal activation with 10 µM free Ca2+ in the prior experiment (57 ± 4 and 48 ± 6% of maximal isometric force for ATP- and Ca2+-stimulated strips, respectively). Exposure to 1 mM H2O2 before permeabilization significantly decreased the isometric force produced in response to ATP (Figs. 5 and 7). Measurements confirmed that the thiophosphorylation treatment produced near-maximal levels of rMLC thiophosphorylation that were not affected by H2O2 exposure before permeabilization (Fig. 7). These findings confirm the results from Fig. 4 that the effects of 1 mM H2O2 on Ca2+ sensitivity that persist after permeabilization were produced by mechanisms independent of changes in rMLC phosphorylation.


View larger version (17K):
[in this window]
[in a new window]
 
Fig. 7.   Isometric force (left) and rMLC phosphorylation (right) in permeabilized strips after maximal, irreversible rMLC thiophosphorylation according to the protocol depicted in Fig. 6 (see text for details). Before permeabilization, muscle strips were exposed to 0 (control) for 1 mM H2O2 for 10 min. Values are means ± SE, n = 8. * Significant difference from control by paired t-test.

Treatment of purified HMM-P for 30 min with 1 mM H2O2 at 37°C resulted in a 48% reduction in the steady-state ATPase activity measured in the presence of 5 µM F-actin (Fig. 8). Treatment with H2O2 did not alter the extent to which HMM was thiophosphorylated (data not shown).


View larger version (9K):
[in this window]
[in a new window]
 
Fig. 8.   Steady-state actin-activated ATPase activity of smooth muscle heavy meromyosin (HMM) that has been maximally thiophosphorylated (HMM-P). After 30 min of treatment with 0 (control) or 1 mM H2O2 for 30 min, HMM-P is purified away from the excess H2O2, and the ATPase activity is determined using a single turnover method. Values are means of 8 experiments for the control group and 4 experiments for H2O2-treated HMM-P. * Significant difference from control by paired t-test.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The findings of this study confirm prior observations that H2O2 relaxes intact ASM by decreasing Ca2+ sensitivity, in part by decreasing rMLC phosphorylation maintained for a given [Ca2+]i (Fig. 2). However, results in the permeabilized smooth muscle preparation used in these studies also demonstrate for the first time that H2O2 also inhibits Ca2+ sensitivity by mechanisms independent of changes in rMLC phosphorylation. Although we were unable to demonstrate inhibition of actomyosin ATPase using an in situ ATPase assay, we demonstrate that a purified subfragment of smooth muscle myosin is modified by H2O2 treatment and that this modification results in inhibition of its ATPase activity in the presence of F-actin. These results suggest that an oxidant species may decrease force at a given [Ca2+]i in part by a direct effect on a key contractile protein, specifically myosin.

In a prior report, we found that H2O2-induced relaxation of canine ASM contracted by muscarinic stimulation or membrane depolarization was caused entirely by decreases in Ca2+ sensitivity, since isometric force decreased while [Ca2+]i increased. These effects on isometric force, Ca2+ sensitivity, and [Ca2+]i were spontaneously reversible after H2O2 washout, demonstrating that these observations do not reflect permanent damage to smooth muscle myocytes in ASM (30). In the current study, we also found a dramatic, reversible increase in [Ca2+]i when H2O2 was added to quiescent, unstimulated strips. This is consistent with prior reports in other cell types (18, 31, 40, 41). Proposed mechanisms include lipid peroxidation (31), stimulation of sarcoplasmic reticulum Ca2+ channels (9), direct activation of intracellular kinases (47), and activation of membrane Ca2+ channels (40). These increases in [Ca2+]i probably account for prior reports that H2O2 increases tone in resting ASM (2, 12, 15, 34, 38, 45). However, the increase in isometric force associated with the increase in [Ca2+]i induced by H2O2 is much less than that expected from the force and F340/F380 responses induced by isotonic KCl. These data indicate that H2O2 decreases Ca2+ sensitivity when applied to quiescent, unstimulated tissue.

We found that H2O2 had no effect on Ca2+ sensitivity when added to strips that were already permeabilized. This finding suggests that components of the solutions bathing the permeabilized muscle inhibited the ability of H2O2 to oxidize intracellular protein targets that regulate isometric force or that a factor important for H2O2 actions was lost during permeabilization. A number of factors, including the chelation of divalent metal ions that may catalyze peroxide-mediated thiol oxidation (35, 46) and the presence of the buffer imidazole, which inefficiently reacts with peroxide (42, 49), may also contribute to this lack of peroxide effect in the permeabilized preparation. This finding has implications regarding the mechanism by which peroxide mediates its effects in this smooth muscle tissue that deserve further exploration.

The results of the experiment in which H2O2 was administered and then removed from isotonic KCl-depolarized strips showed that increases in [Ca2+]i produced by H2O2 can offset H2O2-induced decreases in Ca2+ sensitivity such that at 10 min of exposure there was no net change in isometric force (Fig. 2). However, complete H2O2 washout during continuous activation with isotonic KCl (Fig. 3) uncovered an underlying, persistent effect on Ca2+ sensitivity that was at least partially mediated by decreases in rMLC phosphorylation. This decrease in rMLC phosphorylation is consistent with that observed in the presence of H2O2 found in our previous study (30). The persistence of this effect on Ca2+ sensitivity made it possible to pursue a strategy of permeabilization after treatment of intact strips with H2O2, an exposure that occurs under conditions allowing relevant chemical intermediates to be generated.

At least one component of H2O2-induced inhibition of Ca2+ sensitivity survived permeabilization, as indicated by decreases in the isometric force induced by 10 µM free Ca2+ in permeabilized strips treated before permeabilization with H2O2 (Fig. 3). However, with 1 mM H2O2, this effect was not associated with a decreased level of rMLC phosphorylation (Fig. 4). Thus the effect of this concentration of H2O2 on rMLC phosphorylation was either reversed by permeabilization or had dissipated over the 30-40 min required by the protocol. After exposure to 10 mM H2O2, the effect on rMLC phosphorylation did persist. Thus two mechanisms were responsible for the decrease in Ca2+ sensitivity induced by H2O2 in the intact strips: 1) a component dependent on a decrease in rMLC phosphorylation, which was present after permeabilization only after exposure to a high concentration of H2O2 (10 mM) and 2) a component that did not depend on changes in rMLC phosphorylation, which persisted after permeabilization observed after exposure to 1 mM H2O2. The presence of the latter mechanism was confirmed by the thiophosphorylation experiments (Figs. 5 and 7), which support the study hypothesis.

The mechanisms of H2O2 effects on Ca2+ sensitivity remain speculative. There are several examples of H2O2-induced oxidation of intracellular proteins, with resultant changes in the activity of the protein, including creatine kinase (13, 27) and the ryanodine receptor Ca2+ release channel (9). Many proteins are susceptible to oxidative stress by selective modification of sulfur-containing amino acid residues, such as cysteine (9) and methionine (10), which can impact enzyme and cellular function. Cysteine thiols can be oxidized by H2O2 to several states, depending on the duration of exposure to H2O2, H2O2 concentration, and the reactivity of the thiol (9). Potential targets responsible for the effects dependent on changes in rMLC phosphorylation include calmodulin, smooth muscle protein phosphatase, and Ca2+. The latter is known to contain reactive thiols that are potential targets of H2O2 (33).

Effects independent of changes in rMLC phosphorylation could be related to effects on smooth muscle actomyosin ATPase. The myosin head contains reactive cysteine thiols, which when oxidized or covalently modified inhibit actomyosin ATPase activity (4, 28). Such a modification would decrease isometric force independent of [Ca2+]i or rMLC phosphorylation, an effect noted for myosin oxidation produced by sodium nitroprusside in skeletal muscle (37). Using a purified protein preparation, we have demonstrated that 1 mM H2O2, under conditions similar to those used in intact tracheal smooth muscle experiments (30), inhibited smooth muscle actin-activated myosin ATPase activity by ~50% (Fig. 8). The studies were performed on maximally thiophosphorylated HMM, indicating that the observed reduction in actomyosin ATPase activity occurred independently of rMLC phosphorylation. Although allowing for the fact that H2O2 treatment of intact tracheal smooth muscle occurs in a different microenvironment, we find it plausible that the smooth muscle myosin itself is an H2O2-sensitive target. Inhibition of actomyosin ATPase activity on this basis would account for the phosphorylation-independent reduction in Ca2+ sensitivity. Other possible explanations for this novel observation, including H2O2 oxidation of actin, actin-associated proteins, or of enzymes regulating actin-associated proteins, have not yet been addressed.

In summary, we confirm that H2O2 decreases Ca2+ sensitivity in ASM in part by decreasing the rMLC phosphorylation produced by a given Ca2+ concentration. This effect depends on factors not present or active under the study conditions used in permeabilized ASM. However, experiments in which H2O2 is administered before permeabilization have made it possible for the first time to observe a distinct mechanism that is independent of changes in rMLC phosphorylation that also contributes to H2O2-induced decreases in Ca2+ sensitivity. We provide evidence that oxidation of myosin may provide an explanation for this mechanism of H2O2-mediated reduction in Ca2+ sensitivity.


    ACKNOWLEDGEMENTS

This work was supported by National Heart, Lung, and Blood Institute Grants HL-45532 and HL-54757 and by funds from Mayo Foundation.


    FOOTNOTES

Address for reprint requests and other correspondence: K. A. Jones, Mayo Clinic, 200 First St. SW, Rochester, MN 55905 (E-mail: jones.keith{at}mayo.edu).

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 18, 2002;10.1152/ajplung.00159.2002

Received 22 May 2002; accepted in final form 2 October 2002.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Adelstein, RS, and Klee CB. Purification and characterization of smooth muscle myosin light chain kinase. J Biol Chem 256: 7501-7509, 1981[Abstract/Free Full Text].

2.   Bauer, V, Oike M, Tanaka H, Inoue R, and Ito Y. Hydrogen peroxide induced responses of cat tracheal smooth muscle cells. Br J Pharmacol 121: 867-874, 1997[Abstract].

3.   Bremerich, DH, Kai T, Warner DO, and Jones KA. Effect of phorbol esters on Ca2+ sensitivity and myosin light-chain phosphorylation in airway smooth muscle. Am J Physiol Cell Physiol 274: C1253-C1260, 1998[Abstract/Free Full Text].

4.   Chandra, TS, Nath N, Suzuki H, and Seidel JC. Modification of thiols of gizzard myosin alters ATPase activity, stability of myosin filaments, and the 6-10 S conformational transition. J Biol Chem 260: 202-207, 1985[Abstract/Free Full Text].

5.   Cremo, CR, Wang F, Facemyer K, and Sellers JR. Phosphorylation-dependent regulation is absent in a nonmuscle heavy meromyosin construct with one complete head and one head lacking the motor domain. J Biol Chem 276: 41465-41472, 2001[Abstract/Free Full Text].

6.   Droge, W. Free radicals in the physiological control of cell function. Physiol Rev 82: 47-95, 2002[Abstract/Free Full Text].

7.   Ellison, PA, Sellers JR, and Cremo CR. Kinetics of smooth muscle heavy meromyosin with one thiophosphorylated head. J Biol Chem 275: 15142-15151, 2000[Abstract/Free Full Text].

8.   Fabiato, A, and Fabiato F. Calculator programs for computing the composition of the solutions containing multiple metals and ligands used for experiments in skinned muscle cells. J Physiol 75: 463-505, 1979.

9.   Favero, TG, Zable AC, and Abramson JJ. Hydrogen peroxide stimulates the Ca2+ release channel from skeletal muscle sarcoplasmic reticulum. J Biol Chem 270: 25557-25563, 1995[Abstract/Free Full Text].

10.   Gao, J, Yin DH, Yao Y, Sun H, Qin Z, Schoneich C, Williams TD, and Squier TC. Loss of conformational stability in calmodulin upon methionine oxidation. Biophys J 74: 1115-1134, 1998[Abstract/Free Full Text].

11.   Gao, Y, and Vanhoutte PM. Effects of hydrogen peroxide on the responsiveness of isolated canine bronchi: role of prostaglandin E2 and I2. Am J Physiol Lung Cell Mol Physiol 263: L402-L408, 1992[Abstract/Free Full Text].

12.   Gao, Y, and Vanhoutte PM. Products of cyclooxygenase mediate the responses of the guinea pig trachea to hydrogen peroxide. J Appl Physiol 74: 2105-2111, 1993[Abstract].

13.   Genet, S, Kale RK, and Baquer NZ. Effects of free radicals on cytosolic creatine kinase activities and protection by antioxidant enzymes and sulfhydryl compounds. Mol Cell Biochem 210: 23-28, 2000[ISI][Medline].

14.   Gunst, SJ, al-Hassani MH, and Adam LP. Regulation of isotonic shortening velocity by second messengers in tracheal smooth muscle. Am J Physiol Cell Physiol 266: C684-0C691, 1994[Abstract/Free Full Text].

15.   Gupta, JB, and Prasad K. Mechanism of H2O2-induced modulation of airway smooth muscle. Am J Physiol Lung Cell Mol Physiol 263: L714-L722, 1992[Abstract/Free Full Text].

16.   Guth, K, and Wojciechowski R. Perfusion cuvette for the simultaneous measurement of mechanical, optical and energetic parameters of skinned muscle fibres. Pflugers Arch 407: 552-557, 1986[ISI][Medline].

17.   Hanazaki, M, Jones KA, Perkins WJ, and Warner DO. Halothane increases smooth muscle protein phosphatase in airway smooth muscle. Anesthesiology 94: 129-136, 2001[ISI][Medline].

18.   Iesaki, T, Okada T, Shimada I, Yamaguchi H, and Ochi R. Decrease in Ca2+ sensitivity as a mechanism of hydrogen peroxide- induced relaxation of rabbit aorta. Cardiovasc Res 31: 820-825, 1996[ISI][Medline].

19.   Iesaki, T, Okada T, Yamaguchi H, and Ochi R. Inhibition of vasoactive amine induced contractions of vascular smooth muscle by hydrogen peroxide in rabbit aorta. Cardiovasc Res 28: 963-968, 1994[ISI][Medline].

20.   Ikebe, M, and Hartshorne DJ. Effects of Ca2+ on the conformation and enzymatic activity of smooth muscle myosin. J Biol Chem 260: 13146-13153, 1985[Abstract/Free Full Text].

21.   Ikebe, M, and Hartshorne DJ. Proteolysis of smooth muscle myosin by Staphylococcus aureus protease: preparation of heavy meromyosin and subfragment 1 with intact 20 000-dalton light chains. Biochemistry 24: 2380-2387, 1985[ISI][Medline].

22.   Jones, KA, Lorenz RR, Prakash YS, Sieck GC, and Warner DO. ATP hydrolysis during contraction of permeabilized airway smooth muscle. Am J Physiol Lung Cell Mol Physiol 277: L334-L342, 1999[Abstract/Free Full Text].

23.   Jones, KA, Lorenz RR, Warner DO, Katusic ZS, and Sieck GC. Changes in cytosolic cGMP and calcium in airway smooth muscle relaxed by 3-morpholinosydnonimine. Am J Physiol Lung Cell Mol Physiol 266: L9-L16, 1994[Abstract/Free Full Text].

24.   Jones, KA, Perkins WJ, Lorenz RR, Prakash YS, Sieck GC, and Warner DO. F-actin stabilization increases tension cost during contraction of permeabilized airway smooth muscle in dogs. J Physiol 519: 527-538, 1999[Abstract/Free Full Text].

25.   Kai, T, Yoshimura H, Jones KA, and Warner DO. Relationship between force and regulatory myosin light chain phosphorylation in airway smooth muscle. Am J Physiol Lung Cell Mol Physiol 279: L52-L58, 2000[Abstract/Free Full Text].

26.   Kamm, KE, and Stull JT. The function of myosin and myosin light chain kinase phosphorylation in smooth muscle. Annu Rev Pharmacol Toxicol 25: 593-620, 1985[ISI][Medline].

27.   Kim, JR, Yoon HW, Kwon KS, Lee SR, and Rhee SG. Identification of proteins containing cysteine residues that are sensitive to oxidation by hydrogen peroxide at neutral pH. Anal Biochem 283: 214-221, 2000[ISI][Medline].

28.   Kojima, S, Fujiwara K, and Onishi H. SH1 (cysteine 717) of smooth muscle myosin: its role in motor function. Biochemistry 38: 11670-11676, 1999[ISI][Medline].

29.   Konishi, M, Olson A, Hollingworth S, and Baylor SM. Myoplasmic binding of fura-2 investigated by steady-state fluorescence and absorbance measurements. Biophys J 54: 1089-1104, 1988[Abstract].

30.   Lorenz, RR, Warner DO, and Jones KA. Hydrogen peroxide decreases Ca2+ sensitivity in airway smooth muscle by inhibiting rMLC phosphorylation. Am J Physiol Lung Cell Mol Physiol 277: L816-L822, 1999[Abstract/Free Full Text].

31.   Meyer, TN, Gloy J, Hug MJ, Greger R, Schollmeyer P, and Pavenstadt H. Hydrogen peroxide increases the intracellular calcium activity in rat mesangial cells in primary culture. Kidney Int 49: 388-395, 1996[ISI][Medline].

32.   Mian, KB, and Martin W. Hydrogen peroxide-induced impairment of reactivity in rat isolated aorta: potentiation by 3-amino-1,2,4-triazole. Br J Pharmacol 121: 813-819, 1997[Abstract].

33.   Olson, NJ, Pearson RB, Needleman DS, Hurwitz MY, Kemp BE, and Means AR. Regulatory and structural motifs of chicken gizzard myosin light chain kinase. Proc Natl Acad Sci USA 87: 2284-2288, 1990[Abstract].

34.   Olszewski, MA, Robinson NE, Yu MF, and Derksen FJ. Effects of hydrogen peroxide on isolated trachealis muscle of horses. Am J Vet Res 56: 1479-1485, 1995[ISI][Medline].

35.   Paik, SR, Shin HJ, and Lee JH. Metal-catalyzed oxidation of alpha-synuclein in the presence of Copper(II) and hydrogen peroxide. Arch Biochem Biophys 378: 269-277, 2000[ISI][Medline].

36.   Penefsky, HS. Reversible binding of Pi by beef heart mitochondrial adenosine triphosphatase. J Biol Chem 252: 2891-2899, 1977[Abstract].

37.   Perkins, WJ, Han YS, and Sieck GC. Skeletal muscle force and actomyosin ATPase activity reduced by nitric oxide donor. J Appl Physiol 83: 1326-1332, 1997[Abstract/Free Full Text].

38.   Rabe, KF, Dent G, and Magnussen H. Hydrogen peroxide contracts human airways in vitro: role of epithelium. Am J Physiol Lung Cell Mol Physiol 269: L332-L338, 1995[Abstract/Free Full Text].

39.   Reeve, HL, Tolarova S, Nelson DP, Archer S, and Weir EK. Redox control of oxygen sensing in the rabbit ductus arteriosus. J Physiol 533: 253-261, 2001[Abstract/Free Full Text].

40.   Roveri, A, Coassin M, Maiorino M, Zamburlini A, van Amsterdam FT, Ratti E, and Ursini F. Effect of hydrogen peroxide on calcium homeostasis in smooth muscle cells. Arch Biochem Biophys 297: 265-270, 1992[ISI][Medline].

41.   Shaw, S, Naegeli P, Etter JD, and Weidmann P. Role of intracellular signalling pathways in hydrogen peroxide-induced injury to rat glomerular mesangial cells. Clin Exp Pharmacol Physiol 22: 924-933, 1995[ISI][Medline].

42.   Simpson, JA, Cheeseman KH, Smith SE, and Dean RT. Free-radical generation by copper ions and hydrogen peroxide. Stimulation by Hepes buffer. Biochem J 254: 519-523, 1988[ISI][Medline].

43.   Somlyo, AP, and Somlyo AV. Signal transduction and regulation in smooth muscle. Nature 372: 231-236, 1994[ISI][Medline].

44.   Spudich, JA, and Watt S. The regulation of rabbit skeletal muscle contraction. I. Biochemical studies of the interaction of the tropomyosin-troponin complex with actin and the proteolytic fragments of myosin. J Biol Chem 246: 4866-4871, 1971[Abstract/Free Full Text].

45.   Szarek, JL, and Schmidt NL. Hydrogen peroxide-induced potentiation of contractile responses in isolated rat airways. Am J Physiol Lung Cell Mol Physiol 258: L232-L237, 1990[Abstract/Free Full Text].

46.   Trimm, JL, Salama G, and Abramson JJ. Sulfhydryl oxidation induces rapid calcium release from sarcoplasmic reticulum vesicles. J Biol Chem 261: 16092-16098, 1986[Abstract/Free Full Text].

47.   Ward, CA, and Moffat MP. Role of protein kinase C in mediating effects of hydrogen peroxide in guinea-pig ventricular myocytes. J Mol Cell Cardiol 27: 1089-1097, 1995[ISI][Medline].

48.   Yoshimura, H, Jones KA, Perkins WJ, Kai T, and Warner DO. Calcium sensitization produced by G protein activation in airway smooth muscle. Am J Physiol Lung Cell Mol Physiol 281: L631-L638, 2001[Abstract/Free Full Text].

49.   Zoete, V, Bailly F, Vezin H, Teissier E, Duriez P, Fruchart JC, Catteau JP, and Bernier JL. 4-Mercaptoimidazoles derived from the naturally occurring antioxidant ovothiols 1. Antioxidant properties. Free Radic Res 32: 515-524, 2000[ISI][Medline].


Am J Physiol Lung Cell Mol Physiol 284(2):L324-L332
1040-0605/03 $5.00 Copyright © 2003 the American Physiological Society