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Originally published In Press as doi:10.1074/jbc.M609499200 on November 22, 2006

J. Biol. Chem., Vol. 282, Issue 7, 4336-4344, February 16, 2007
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H2O2-induced Kinetic and Chemical Modifications of Smooth Muscle Myosin

CORRELATION TO EFFECTS OF H2O2 ON AIRWAY SMOOTH MUSCLE*Formula

Alan R. Penheiter{ddagger}, Michelle Bogoger§, Patricia A. Ellison§, Barbara Oswald{ddagger}, William J. Perkins{ddagger}, Keith A. Jones, and Christine R. Cremo§1

From the §Department of Biochemistry and Molecular Biology, University of Nevada School of Medicine, Reno, Nevada 89557, the {ddagger}Department of Anesthesiology, Mayo Clinic, Rochester, Minnesota 55905, and the Department of Anesthesiology, University of Alabama-Birmingham, Birmingham, Alabama 35249-6810

Received for publication, October 10, 2006


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The effect of H2O2 on smooth muscle heavy meromyosin (HMM) and subfragment 1 (S1) was examined. The number of molecules that retained the ability to bind ATP and the actinactivated rate of Pi release were measured by single-turnover kinetics. H2O2 treatment caused a decrease in HMM regulation from 800- to 27-fold. For unphosphorylated and phosphorylated heavy meromyosin and for S1, ~50% of the molecules lost the ability to bind to ATP. H2O2 treatment in the presence of EDTA protected against ATPase inactivation and against the loss of total ATP binding. Inactivation of S1 versus time correlated to a loss of reactive thiols. Treatment of H2O2-inactivated phosphorylated HMM or S1 with dithiothreitol partially reactivated the ATPase but had no effect on total ATP binding. H2O2-inactivated S1 contained a prominent cross-link between the N-terminal 65-kDa and C-terminal 26-kDa heavy chain regions. Mass spectral studies revealed that at least seven thiols in the heavy chain and the essential light chain were oxidized to cysteic acid. In thiophosphorylated porcine tracheal muscle strips at pCa 9 + 2.1 mM ATP, H2O2 caused a ~50% decrease in the amplitude but did not alter the rate of force generation, suggesting that H2O2 directly affects the force generating complex. Dithiothreitol treatment reversed the H2O2 inhibition of the maximal force by ~50%. These data, when compared with the in vitro kinetic data, are consistent with a H2O2-induced loss of functional myosin heads in the muscle.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Contraction of smooth muscle is controlled largely by phosphorylation of the 20-kDa myosin regulatory light chain (RLC),2 resulting in the cyclic attachment and detachment of myosin to actin (cross-bridge cycling) and the hydrolysis of ATP by actin-activated myosin ATPase (1). The level of RLC phosphorylation depends on the balance between the activities of MLCK and SMM phosphatase. MLCK is primarily activated (through Ca2+-calmodulin) by an increase in [Ca2+]i from intracellular stores or influx of extra-cellular Ca2+, whereas inhibition of myosin phosphatase, which contributes to the amount of force at a given [Ca2+]i (Ca2+ sensitivity), occurs through a receptor-G protein-catalyzed signal transduction cascade (2–5).

Reactive oxygen species, such as H2O2, modulate smooth muscle contractility (6) and contribute to the severity of numerous diseases, including acute lung injury, asthma, pulmonary hypertension, ischemia-reperfusion, and arthritis (7). H2O2 reversibly inhibits receptor agonist-induced contraction of both vascular (8–10) and ASM (11–15).

The mechanisms for H2O2 inhibition of smooth muscle contraction are not well understood but have been proposed to involve effects on kinase-mediated signaling cascades, cytoskeletal effects, ionotropic effects, lipid oxidation, direct inhibitory effect on smooth muscle contractile proteins, and potentially confounding effects on epithelial cells (11–20). In general, healthy smooth muscle is quite resistant to exogenous H2O2. The most extreme example is agonist activated bladder smooth muscle (bladder concentrations of H2O2 often reach levels of 0.2–0.4 mM in healthy adults (21), which was shown to exhibit an EC50 for relaxation of 36 mM H2O2 (22)). Airway smooth muscle, although considerably more sensitive than bladder, still requires nearly mM concentrations of H2O2 for relaxation of agonist induced contraction (23). Previously we showed that 0.1–3 mM H2O2-induced relaxation of ASM was caused by a reduction in the amount of force produced at given [Ca2+]i, (Ca2+ sensitivity), whereas [Ca2+]i actually increased in response to H2O2 treatment (23). Additionally, we showed that the inhibitory effect of peroxide on ASM contraction persisted after Triton X-100 permeabilization and RLC thiophosphorylation. Because the levels of RLC thiophosphorylation were unaffected by H2O2 treatment, we proposed that H2O2 directly inhibits the acto-myosin contractile apparatus (24).

Here we address whether or not myosin oxidation underlies the physiological effect of H2O2 treatment in ASM as mentioned above. This is an important question because in terms of contractility it may not matter what the effects of H2O2 are on signaling cascades upstream of myosin activation, because even pro-contractile-signaling would be nullified by direct inhibition of acto-myosin cycling. A better understanding of effects of peroxide on SMM will ultimately clarify the potential mechanisms at play in previous studies of H2O2-induced pathogenesis.

Here we characterize the effects of H2O2 on the kinetics of purified SMM and correlate those data to changes in force generation in ASM tissue. We have examined the steady-state and single-turnover ATPase kinetics of purified SMM in both the unphosphorylated and phosphorylated states. We have used two soluble myosin subfragments that are suitable for kinetic experiments. HMM is double-headed and lacks two-thirds of the tail and therefore cannot form filaments. However, it is fully regulated by light chain phosphorylation. S1 is a single head domain that cannot form filaments and is not regulated by light chain phosphorylation. The effects of EDTA during the H2O2 treatment and the effect of DTT after the H2O2 treatment have been analyzed. In addition, we have analyzed the effect of H2O2 on the total number of reactive thiols and the formation of intersubunit disulfide bonds. Mass spectral studies revealed which residues were oxidized and to which oxidation state. The effect of H2O2 on the rate and amplitude of force development in permeabilized porcine tracheal smooth muscle strips was also examined. Remarkably, the specific effects on myosin kinetics observed for purified SMM were paralleled by the effects observed in the muscle, strongly suggesting that elevated H2O2 levels during diseased states could alter myosin kinetics and partially explain altered contractility.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Protein Preparations—Protein preparations were essentially as described (25). HMM and S1 were prepared by digestion of gizzard SMM (26) with Staphylococcus aureus protease V8 (Sigma). Both HMM (E (0.1%) = 0.65) and S1 (E (0.1%) = 0.75) are mixtures of full-length heavy chains and clipped heavy chains with an internal cleavage at the actin binding loop ~26 kDa from the head-tail junction. The light chains were unaffected. F-actin was prepared from rabbit muscle according to the method of Spudich and Watt (27) and was dialyzed versus 10 mM MOPS (pH 7.0), 0.1 mM EGTA, 50 mM NaCl, 0.8 mM MgCl2, 0.2 mM ATP. MLCK (E (0.1%) = 1.14) was prepared (28) with the modifications described (25) and was stored at –80 °C in single aliquots with 10 mM DTT. 10 mM DTT was added to MLCK immediately after thawing and then stored on ice.

Thiophosphorylation of HMM—HMM was thiophosphorylated as described (29). This preparation is referred to as pHMM. The extent of thiophosphorylation was verified using 10% Tris-glycine gels (10 cm x 10 cm, 12 lanes; Invitrogen) with standard Tris-glycine running buffer. This type of gel gave superior results to urea and/or urea-glycerol gels (30, 31). The samples were precipitated with 3 volumes of cold acetone prior to the addition of sample buffer (8 M urea (ultrapure, Research Organics), 33 mM Tris-glycine (pH 8.6), 0.17 mM EDTA, 10 mM DTT (added immediately before use), bromphenol blue) to 6–7 mg/ml protein. Approximately 25–30 µg of HMM was applied to the gel. The samples were not heated above room temperature.

Peroxide Treatment of HMM and S1—HMM samples were thiophosphorylated where indicated and spun through a 5-ml buffer exchange column (32) prepared with Sephadex G-50–80 resin (Sigma) in nonreducing buffer (10 mM MOPS, 50 mM NaCl (pH 7.0)). The samples (typically 1–2 mg/ml) were incubated with H2O2 under specified conditions (see legends). To remove H2O2, the samples were either spun through buffer exchange columns to nonreducing buffer or treated with catalase, as indicated.

Single Turnover of HMM-MantATP in the Presence of Actin—This assay measures the rate of phosphate release from the acto-HMM MantADPPi state (25). Because phosphate release is rate-limiting, the decrease in fluorescence as MantADP dissociates from HMM can be used to measure phosphate release rates. The single-turnover assay allows populations of molecules or heads with different turnover rates to be elucidated. The assays were performed at 25 °C in a temperature controlled stopped flow fluorimeter (Hi-Tech, SF-61DX2, Salisbury, UK) equipped with a 75-watt Xe-Hg lamp. The mixing dead time was ~2 ms. The excitation wavelength was 365 nm, and the excitation bandwidth was 4 nm. Emission was collected through a 389 low cut-off filter (Corion). For upHMM, formycin triphosphate was used instead of MantATP. The excitation wavelength was 313 nm, and the excitation bandwidth was 4 nm. Emission was collected through a 370 low cut-off filter (Corion). The experiment was done with two syringes. The first syringe contained 0.8 µM HMM heads in 10 mM MOPS (pH 7.0), 50 mM NaCl. The second syringe contained 10 µM actin, 200 µM ATP, 10 mM MOPS (pH 7.0), 50 mM NaCl, 0.1 mM EGTA, 0.8 mM MgCl2,1mM DTT. A stopwatch was started upon adding MgMantATP (1.6 µM MgCl2, 0.8 µM MantATP) to the contents of the first syringe. The fluorophore binds maximally to pHMM in about 45 s (data not shown). After flushing the cuvette three times, 50 µl was shot from each syringe (at 45 s), and the data were collected with the anti-bleaching shutter engaged for upHMM (1000–1400 s; 1024 points) or without the shutter for shots under 60 s for pHMM and S1 (1024 points). All of the data were analyzed with Kaleidagraph software (Synergy Software, Reading, PA). The data were fit to a double exponential model (25). KATPase (Km for actin) and Vmax for pHMM were determined by varying the actin concentration from 5 to 150 µM and fitting the curve to the Michaelis-Menten equation as described (29).

Steady-state ATPase Assay—Steady-state [{gamma}-32P]ATP hydrolysis of S1 was measured in 10 mM MOPS (pH 7.0), 1 mM ATP, 2 mM MgCl2, and 50 µM F-actin at 25 °C. Released 32Pi was separated from [{gamma}-32P]ATP with acidified, activated charcoal (33) and quantified by scintillation counting. The background [{gamma}-32P]ATP hydrolysis rates of actin alone and S1 in the absence of actin were subtracted.

Mass Spectrometry—UpS1 was treated with 1 mM H2O2 for 30 min, alkylated with 10 mM iodoacetamide for 30 min, and subjected to nonreducing SDS-PAGE. The bands were stained with Coomassie Safe Stain, excised, reduced with DTT, treated with vinyl-pyridine, and subjected to in-gel trypsinolysis with modified sequencing grade trypsin (Promega). Following digestion, the samples were subjected to nanobore LC with on-line tandem mass spectrometry performed on a ThermoFinnigan LTQ-FT. This instrument is a hybrid of a linear ion trap with a Fourier transform-infrared cyclotron resonance detector (34, 35). Searches for oxidative modifications of Cys and Met, alkylation of Cys, and N-acetyl Cys (ELC) were conducted with Mascot (Matrix Science).


Figure 1
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  		FIGURE 1.
MantATP single-turnover kinetics of H2O2-treated pHMM. HMM samples were treated with the indicated H2O2 concentrations for 30 min at 37 °C in the absence or presence of 5 mM EDTA, 5.1 mM MgCl2. H2O2 was then removed by centrifugal gel filtration. Rates of Pi release (expressed as percentages of a heated control) from the acto-HMM-MantADP-Pi complex were measured as described under "Experimental Procedures" at 5 µM actin. The data were fitted to a double exponential model. The fast rate is typically 80% of the total amplitude and has a rate of ~1.2–2.1 s–1. The slow rate has a rate of ~0.04–0.06 s–1. A, rates of the fast (squares, no EDTA; circles, with EDTA) and the slow phases (diamonds, no EDTA; triangles, with EDTA) versus H2O2. The data are from two independent experiments. Each point was run in triplicate, and the data shown are the means and standard error of the six measurements. The turnover rate for the unheated control was 103 ± 5% of the heated control. B, plot of the amplitudes (as percentages of total amplitude) of the fast (squares) and slow (circles) rates of turnover from the experiments (no EDTA) in A. C, the weighted rate equals (rate1)(amplitude1 %) + (rate2)(amplitude2 %). No EDTA (squares) and with EDTA (circles) are for a single experiment with triplicate data points. An identical independent experiment gave similar results. Standard errors were calculated according to Ref. 59. D, total amplitude (percentage of control) versus H2O2 for two independent experiments. No EDTA (squares) and with EDTA (circles). For all plots, the lines through the data are either a linear fit or interpolated.

 
Tracheal Smooth Muscle Experiments—See the supplemental materials and the legend for Fig. 9 for information regarding the tracheal smooth muscle experiments.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
To evaluate the effects of H2O2 on the ATPase kinetics of pHMM (Fig. 1) and upHMM (Fig. 2), samples were heated for 30 min at 37 °C in nonreducing buffer with H2O2 or H2O2 plus 5mM EDTA. After rapidly removing H2O2, the transient kinetics of phosphate release was measured in the presence of 5 µM actin. The data were fit to a double exponential model. This assay has two advantages over the more typical steady-state assay. First, it can monitor populations of molecules with different turnover rates, and second, it can monitor the total number of ATP-binding sites by analyzing the extent of fluorescence increase upon binding of a fluorescent ATP analog. The rate of the fast phase of turnover (as a percentage of an identically treated control without H2O2) for pHMM is plotted versus H2O2 concentration in Fig. 1A. Unheated controls (4 °C) behaved essentially identically to heated controls (37 °C exposure without H2O2, data not shown). The fast rate decreased to ~50%, and the slow rate increased to about 120% of control at 2.5 mM H2O2. An identical H2O2 treatment in the presence of 1 mM MgADP (data not shown) gave a similar result, suggesting that nucleotide binding at the active site did not protect against the inactivation. Fig. 1B shows that the fast rate contributes ~80% and the slow rate contributes ~ 20% of the total turnover amplitude prior to H2O2 treatment. H2O2 treatment caused the fractional amplitude of the fast and slow rate to decrease and increase, respectively. These data might suggest that untreated HMM contains ~20% oxidized heads or heads that behave like H2O2-oxidized heads. However, we have found that the highly similar nonmuscle IIB HMM (expressed in insect cells) (36) and the single head fragment S1 (see later) also show the 80% fast to 20% slow turnover behavior. This suggests that the biphasic turnover is an intrinsic property of smooth and nonmuscle myosin heads. The molecular basis of this phenomenon is under further investigation. The weighted rate plotted in Fig. 1C takes into account both rates and amplitudes and is indicative of the steady-state ATPase rate of the population of molecules that bind to nucleotide. In the absence of EDTA, the weighted rate decreases to ~40% of control (squares). Similarly, the total amplitude (Fig. 1D), which reflects the total number of nucleotide-binding sites, decreases to ~50% of control (squares). This was confirmed in a separate experiment monitoring the rate and amplitude of MantATP binding under pseu-do-first order conditions (data not shown). Only the amplitude and not the rate of binding was altered by H2O2. Therefore treatment with 2.5 mM peroxide results in two populations of heads, 50% with 40% activity and 50% with ~0 activity. If this sample had been measured by a steady-state assay, the activity would be 20% of control. In summary, H2O2 affects the kinetics of pHMM in two ways. First, for those molecules that still bind to nucleotide, it decreases the steady-state ATPase by decreasing the rate and the fractional amplitude of a fast component and increasing the rate and fractional amplitude of a slow component. This is an effect upon Vmax, not Km. Actin titrations showed that the KATPase (of Km for actin) of control pHMM was 83 ± 18 µM, and that of peroxide-treated pHMM was 80 ± 21 µM (data not shown). Second, H2O2 causes ~50% of the molecules to either lose the ability to bind to nucleotide or drastically weaken their affinity for nucleotide.


Figure 2
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  		FIGURE 2.
Formycin triphosphate single-turnover kinetics of H2O2-treated upHMM. A, rates of fast (squares) and slow (circles) phases versus H2O2 concentration. B, corresponding amplitudes of the fast (squares) and slow (circles) phases. C, weighted rate as described in Fig. 1. D, total amplitude (percentage of control) versus H2O2 concentration. The error bars and replicates are similar to those in Fig. 1.

 
The presence of EDTA during the H2O2 treatment (5.0 mM) generally diminished the H2O2-induced inactivation of pHMM with respect to the fast rates and diminished the H2O2-induced activation with respect to the slow rates (Fig. 1A), as reflected in the weighted rates (Fig. 1C). EDTA also protected against the loss of ATP-binding sites (Fig. 1D), although the effect was not large.

The effect of H2O2 on the kinetics of upHMM is shown in Fig. 2. The assay was the same as in Fig. 1, except that formycin triphosphate instead of MantATP was used as the fluorescent nucleotide. Fig. 2 (A and B) shows that the effect of H2O2 is 2-fold. Without H2O2, the turnover data can be fit to a single exponential with a rate constant of ~0.0015 s–1. H2O2 causes this rate to increase slightly, and a new phase appears with a rate at about 10 times the control rate. The relative contribution of this faster rate increases with increasing H2O2 concentration (Fig. 2B). Fig. 2C shows that H2O2 increases the weighted rate by ~15-fold at 2 mM peroxide. Fig. 2D shows that treatment with 2 mM peroxide caused a loss of ~50% of the total ATP-binding sites as was found for pHMM (Fig. 1D). Peroxide-treated upHMM was capable of being phosphorylated (data not shown). As in pHMM, the effect of peroxide on upHMM was to alter the kinetics of a population of molecules that can still bind to nucleotide and to render about 50% of the molecules incapable of binding to nucleotide.

The effect of H2O2 on the kinetics of upS1 is shown in Fig. 3. This subfragment has no tail and only a single head domain. It is unregulated in that both the unphosphorylated and phosphorylated forms have a high activity. The pattern of H2O2-induced kinetic effects is very similar to those observed for pHMM (Fig. 1). This is expected, as pHMM and upS1 have similar high catalytic activity and other similarities with respect to RLC structure (37). The observed changes in the two rate constants are similar, although the changes in amplitudes (Fig. 3B) are not as great. Again, ~50% of the total nucleotide-binding sites have been destroyed (Fig. 3D).

Fig. 4 shows the protective effect of EDTA on the peroxide-mediated inactivation of upS1 as measured with a steady-state ATPase assay. At 3 mM H2O2 without EDTA, about 20% of the activity remains. This is consistent with the single-turnover data in Fig. 3, which predicts that at 2.5 mM H2O2 the steady-state activity should be ~30% of control. EDTA (5 mM) protected against the inactivation, with about 70% of the activity remaining after a 3.0 mM H2O2 treatment. These data are similar to those observed for pHMM (Fig. 1).


Figure 3
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  		FIGURE 3.
MantATP single-turnover kinetics of H2O2-treated upS1. A, rates of fast (squares) and slow (circles) phases versus H2O2 concentration. B, corresponding amplitudes of the fast (squares) and slow (circles) phases. C, weighted rate as described in the legend to Fig. 1. D, total amplitude (percentage of control) versus H2O2 concentration. The error bars and replicates are similar to those in Fig. 1.

 


Figure 4
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  		FIGURE 4.
Effect of H2O2 on the upS1 actomyosin steady-state ATPase activity. After incubating upS1 (1.75 mg ml–1) with H2O2 (0.1–3.0 mM, 30 min, 30 °C), the reactions were terminated with catalase (250 units) to remove excess H2O2, and steady-state ATPase activity was measured as described under "Experimental Procedures." The EC50 for H2O2 was 0.61 ± 0.12 mM. The activities are expressed relative to control (typically 130–175 nmol min–1 mg–1). The basal ATPase activity (no actin) was 32.6 ± 5.3 nmol min–1 mg–1. The actin concentration was 50 µM, and EDTA concentration was 5 mM.

 
Fig. 5 shows a time course of the reaction of upS1 with 3 mM H2O2. The percentage of inhibition of the steady-state ATPase activity correlated with the total number of thiols as measured by Ellman’s reagent, DTNB (38). There is an initial rapid inactivation phase in which about two thiols are modified, followed by a slower phase in which approximately six or seven thiols (about one-half of the total) are modified after 30 min. At the same time the ATPase was inhibited to ~70% of control. The tight correlation between the inactivation and the loss of reactive thiols suggests that oxidized thiols cause the ATPase inactivation. The final thiol oxidation states could be disulfides or higher oxidation states such as sulfenic or sulfonic acids.


Figure 5
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  		FIGURE 5.
Time course of H2O2 treatment of upS1. H2O2 (3 mM) was added to upS1 (1.75 mg ml–1) at time 0 and incubated at 30 °C. The aliquots were removed at the indicated times, and the reaction was terminated with catalase (250 units) followed by Sephadex G-25 gel filtration. The inhibition of the steady-state ATPase activity (solid line, left axis) relative to control activity (typically 130–175 nmol of Pi min–1 mg–1) and the moles of DTNB-reactive thiols (SH) per mol of S1 (dotted line, right axis) are plotted. For the DTNB assay, S1 was diluted to 0.67 mg/ml in 8 M urea, 50 mM Tris, pH 7.0, 10 mM EDTA, 0.1 M KCl and incubated at 25 °C for 10 min prior to addition of DTNB to 1 mM from a 0.1 M stock made in 1 M Tris, pH 8.0. A412 measurements were taken immediately. The ratio of DTNB reactive thiols per S1 was calculated using the extinction coefficient for the released anion of 14,140 M–1 cm–1 (38). The data points and error bars are the means and standard error, respectively, from three or four independent experiments.

 


Figure 6
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  		FIGURE 6.
Effect of treatment with the thiol-specific reducing compound DTT on H2O2-induced inhibition of upS1 steady-state ATPase activity. After incubating upS1 (1.75 mg ml–1) with 0 (control) or 1 mM H2O2 (30 min, 30 °C), both reactions were terminated with catalase (250 units) and then were divided. One portion of each divided sample was then treated (25 °C) with 0 or 50 mM DTT for 1 or 2 h, and the ATPase activity was measured. The data are the means ± S.E. from four to seven independent experiments.

 
Fig. 6 shows the effect of treatment with the reducing agent, DTT, on H2O2-inhibited upS1. Treatment with DTT (50 mM) for 1 or 2 h caused reversal of the inhibition from 40% of control to ~66 and 77%, respectively, and suggests that the mechanism for at least some of the loss of ATPase activity is by thiol oxidation. Because it is known that DTT readily reduces disulfides and sulfenic acid(s) (R-SOH) (39) to thiols, it is likely that these modifications are responsible for some of the inhibited ATPase activity. Under these robust DTT treatment conditions the ATPase activity was not completely reactivated. This suggests that H2O2 treatment causes some of the thiols to become oxidized further to DTT-unreactive states such as sulfinic (R-SO2H) or sulfonic acids (R-SO3H) or that methionine(s) is also oxidized.

Fig. 7 shows the effects of DTT on the single-turnover kinetics of H2O2-treated pHMM. In this example the H2O2 treatment inhibited the ATPase activity (of those heads that can still bind nucleotide) to ~30% of control (Fig. 7C). At the same time, ~54% of the heads lost the ability to bind to nucleotide and therefore have 0 activity (Fig. 7D). Treatment with up to 40 mM DTT did not increase the total ATP-binding sites (Fig. 7D) but did increase the weighted rate (Fig. 7C) to ~55% of control and partially reversed the pattern of kinetic rates (Fig. 7A) and amplitudes (Fig. 7B) characteristic of the H2O2-treated S1. These data suggest that DTT can partially restore the native kinetics of molecules still capable of binding nucleotide but has no effect upon the molecules that lost the ability to bind nucleotide. This suggests that the loss of ATP binding ability is not correlated to disulfide bond formation but perhaps to further thiol oxidation or to methionine oxidation (40).

To test for H2O2-induced intersubunit disulfide bond formation, we analyzed a nonreducing SDS gel of H2O2-treated upS1 (Fig. 8A). S1 prepared in this manner has most of the heavy chain internally cleaved into a ~65-kDa N-terminal fragment and a C-terminal 26-kDa fragment. A small amount of uncleaved heavy chain still remains at ~90 kDa. H2O2 internally cross-linked the ELC, causing it to smear to higher Rf. In addition, the 26-kDa heavy chain fragment was cross-linked to the 65-kDa heavy chain fragment (C-terminal heavy chain to N-terminal heavy chain). This latter cross-link was identified by excising, reducing, and subsequently electrophoresing the band onto a reducing gel (Fig. 8A, right lane). Mass spectral analysis of a tryptic digest of the ~90-kDa band identified heavy chain peptides from both the 65- and 26-kDa fragments (data not shown). The nonreducing gel in Fig. 8B shows that the presence of EDTA during H2O2 treatment largely inhibits disulfide cross-linking of the 65/26-kDa peptide pair. Under these conditions EDTA does not fully protect against the loss of ATPase activity. Therefore peroxide-treated S1 can be partially inactivated without the above disulfide bonding, suggesting that the disulfide bond is not the only modification to the protein. Fig. 8C shows that the reductant in the sample buffer was sufficient to largely reverse the cross-link, as expected for a disulfide bond. Therefore, gel analysis revealed that there are at least two SH groups, one on the 65-kDa fragment and the other on the 26-kDa fragment that become disulfide-bonded (reversible by DTT) upon H2O2 treatment and that EDTA can inhibit disulfide bond formation.

Table 2 shows the results of a mass spectral analysis of H2O2-treated upS1. Listed are the peptides that were found to contain cysteine thiols completely oxidized to the sulfonic acid group (cysteic acid). Sulfonic acids were not detected in the control. It is likely that one or more of these cysteine oxidations are responsible for the loss of tight ATP binding because sulfonic acids cannot be reduced by DTT. In all cases where a sulfonic acid was identified, corresponding peptides with carbamidomethyl-Cys (iodoacetamide-treated) and/or pyridylethyl-Cys (vinyl pyridine-treated) were also detected, indicating that only a fraction of S1 molecules were oxidized completely. Cys121, Cys481, Cys524, Cys545, and Cys717 (SH1) were not reliably identified because of the inappropriate size and/or hydrophobicity of the tryptic fragments containing these residues. The remaining 14 of 18 total Cys in the S1 head and ELC were reliably identified (not listed in Table 2) and were either carbamidomethyl-Cys or pyridylethyl-Cys. With our experimental approach it should have been possible to identify Cys residues in the disulfide-linked 26–68 kDa (Fig. 8) that obligatorily participated in disulfide bonds. These Cys should not react with iodoacetamide but should react with vinyl pyridine following reduction with DTT. However, for every tryptic fragment, peaks were detected with carbamidomethyl-Cys, indicating that these SDS-PAGE resolved disulfide-linked dimers contained populations of alternate disulfide linkages. All Cys in the ELC were reliably identified. For the disulfide-linked 26–68-kDa band, it is also possible that two of the Cys in the myosin head, which were not identified by our mass spectral analysis, may have been responsible for the H2O2-induced disulfides. It is likely that Cys717 in the 26-kDa fragment is involved because its alkylation can alter SMM regulation and ability to move actin in an in vitro assay (41).


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  		TABLE 2
Peptides containing cysteine thiols oxidized to sulfonic acids (cysteic acid) as determined by mass spectrometry of trypsin-digested upS1

Peptides from tryptic digest of upS1 following oxidation with 1 mM H2O2 for 30 min at 37 °C. Following peroxide treatment, cysteines were treated with iodoacetamide, and bands were resolved on SDS-PAGE, excised, then reduced with DTT, and treated with vinyl pyridine prior to trypsin digestion. Experimental Mr is based on a search of b and y ions from the tandem mass spectrometry analysis. The masses listed are for the precursor ion prior to tandem mass spectrometry fragmentation.

 


Figure 7
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  		FIGURE 7.
Effect of DTT on the single-turnover kinetics of H2O2 treated pHMM. Control samples were not treated with H2O2. All other samples were treated with DTT at the indicated concentrations for 10 min at 42 °C after treating with and removing 2.5 mM H2O2 and assayed for the rate of phosphate release as described in Fig. 1. A, rate values for the slow and fast phases are shown next to eachbar. The statistics are similar to Fig. 1. B, percentage of total amplitude of the fast (top bar) and slow phases (bottom bar), respectively. C, the weighted rate (see Fig. 1 for definition) is plotted as a percentage of control (100%). The solid line is a fit to a hyperbola. D, the total amplitude (%).

 


Figure 8
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  		FIGURE 8.
Gel analysis of H2O2-induced intersubunit cross-linking of S1. A, Coomassie-stained nonreducing (–DTT) SDS-PAGE (Invitrogen 4–20%) of S1 (10 µg) treated with or without H2O2 for 30 min at 30 °C. H2O2 concentrations are indicated. HC, heavy chain. The emerging cross-linked band migrating at the position corresponding to the unclipped heavy chain (boxed; ~90 kDa) was excised (box) and ground in reducing buffer containing 200 mM DTT, 50 mM Tris, pH 7.0, and 2%SDS. The homogenate was incubated overnight at 37 °C followed by centrifugation at 13,000 x g for 1 min. Glycerol and bromphenol blue were added to the supernatant in preparation for electrophoresis on another 4–12% gel (reducing; right lane). B, nonreducing gel of S1 treated with EDTA alone (left) or with 3 mM H2O2 in the presence of 5 mM EDTA. C, reducing gel (200 mM DTT) of S1 treated with 3 mM H2O2,3mM H2O2 + 5mM EDTA, or EDTA alone.

 
Fig. 9 shows the effects of H2O2 treatment of intact porcine tracheal smooth muscle upon the rate and amplitude of force generation at pCa 9 + 2.1 mM ATP following Triton X-100 permeabilization and RLC thiophosphorylation (see the supplemental materials for a representative trace of the entire protocol). H2O2 caused a ~50% decrease in the amplitude but not in the rate of force generation. Note that the levels of thiophosphorylation were not affected by H2O2 treatment (24). Because the effect of H2O2 persists after permeabilization and complete RLC thiophosphorylation, this suggests that H2O2 directly affects the force-generating complex. These data are consistent with a H2O2-induced loss of functional myosin heads. The fact that the rate of force generation is unaffected suggests that oxidation does not slow all myosin molecules but rather abolishes the activity of some myosin molecules. This is remarkably consistent with our finding that H2O2 treatment disables some heads in regard to nucleotide binding in vitro. In the muscle (Fig. 9), DTT treatment at 10 mM for 10 min reversed the H2O2 inhibition by ~50%. Further DTT treatment had little effect (data not shown). This suggests that DTT can reactivate a population of heads that were rendered incapable of force generation but does not rescue all heads. Again this is consistent with our in vitro finding that DTT cannot restore the heads that have lost the ability to bind to nucleotide (Fig. 7D).


Figure 9
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  		FIGURE 9.
Isometric force measurements of tracheal smooth muscle. These data are collected during a set of experiments that were similar to that shown in the supplemental data that describes the entire protocol. Here, only the final ATP-induced force segment of the experiment is shown. Specifically for this experiment, tracheal smooth muscle strips were perfused with or without 3 mM H2O2 for 10 min and permeabilized with Triton X-100, and the RLC was thiophosphorylated (not reversed by phosphatases) during 3 10-min perfusion segments with ATP{gamma}S in high calcium rigor buffer as described in the supplemental data. For the DTT treated strips, 10 mM DTT was included in the second 10 min thiophosphorylation perfusion. Force was induced with 2.1 mM ATP. Note that H2O2 treatment decreased the force even though the level of myosin thiophosphorylation was the same. H2O2 treatment does not affect thiophosphorylation levels of RLC under these conditions (24). All of the traces were normalized to percentages of force generated by 1 µM acetylcholine in the same muscle strip prior to permeabilization. Traces shown are the averages of six experiments with standard error for each condition denoted by flanking dashed lines. Force data were fit to a single exponential equation. [Force = Forcemax*(1 – exp(–K*X))]. Forcemax was 30.1 ± 0.18, 43.0 ± 0.23, and 52.6 ± 0.22% of acetylcholine-induced force for H2O2,H2O2 + DTT, and DTT control, respectively. The rate of force generation (K) for all three conditions was 0.04 s–1.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The data in Figs. 1 and 2 demonstrate that the effect of H2O2 upon the actin-activated ATPase kinetics of molecules that retain nucleotide binding affinity is to alter regulation, which is defined as the phosphorylated/unphosphorylated rate (Table 1). Treatment with 2.5 mM H2O2 decreased the pHMM rate by 65% and increased the upHMM rate by 90%. There was no effect on KATPase (Km), only on Vmax. Regulation decreased from 800- to 27-fold under these conditions. Constructs of SMM with altered regulation typically have abnormally high ATPase activities in the unphosphorylated state and an abnormally low ATPase in the phosphorylated state (36, 42–47). Alkylation of SMM thiols causes a similar effect (41, 48, 49) most likely attributed to reaction of SH1(Cys717). The effects of H2O2 on pHMM and upS1 were very similar in our studies, suggesting that they are not mediated by modification of the most reactive thiol on SMM, which is in the S2 region of the rod (Cys892) (49) and which is not present in S1.


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  		TABLE 1
Summary of the effects of H2O2 treatment on HMM

 
An important finding of this work is that H2O2 causes a large population of the SMM heads to lose the ability to tightly bind to nucleotide. Myosin subfragments have been shown to contain variable relative amounts of weak ATP-binding heads (50) even under control conditions. However, we are unaware of any treatment known to cause loss of tight ATP binding prior to this work. It is known that an uncharacterized process can cause myosin to form ATP-resistant actomyosin complexes often termed "dead heads" (25, 29). Indeed, ATP-unreactive heads must often be removed by actin selection prior to single molecule force and displacement measurements to allow the active heads to be observed. Skeletal myosin that is extensively modified with the thiol-selective reagent N-ethylmaleimide forms ATP-resistant actomyosin complexes (51), but it is not known whether this is due to the loss of tight ATP binding.

It is known that alkylation of SMM thiols can cause changes in the regulation of the ATPase activity of SMM (41, 48) similar to those observed here. However, because prior work measured ATPase activities by steady-state methods, it is not clear whether or not a population of the heads was rendered incapable of ATP binding.

We show that a robust DTT treatment cannot revive the ability to bind to ATP, suggesting that disulfide bonding is not the cause but rather further oxidized thiols to sulfonic acid or oxidation of methionines. Mass spectrometric analysis confirmed that seven total cysteine thiols, four on the heavy chain (Cys753, Cys707, Cys388, and Cys582) and three on the ELC (Cys31, Cys84, and Cys137) were oxidized to sulfonic acids. This was observed only for the H2O2-treated sample. Thiols are most likely initially oxidized to the unstable sulfenic acids (R-SOH) (52) and further oxidized to the sulfonic acids (R-SO3H). These data are consistent with a study of sulfenic acid formation in H2O2-perfused heart muscle (53). Sulfenic acid was detected in the myosin heavy chain after initial H2O2 treatment but diminished as further oxidation proceeded, most likely because of conversion to sulfinic then sulfonic acids. Such oxidation occurred at ~10–50-fold lower effective H2O2 concentrations than used here, suggesting that the thiols in question are highly susceptible to oxidation. We also detected oxidized methionines but cannot assess the significance because they are commonly found in unoxidized samples digested from gels. Overall, the data suggest that the loss of nucleotide binding affinity is most likely due to an oxidation of one or more thiols to sulfonic acids.

The location of the observed oxidized thiols (Table 2) does not immediately suggest a mechanism for the loss of ATP binding ability. The ELC is not required for ATP binding. The locations of Cys388 in the upper 50-kDa subdomain, Cys582 in the lower 50-kDa subdomain, and Cys753 in the converter domain do not appear to be critical to the ATP-binding site. Cys582 is in loop 3 of the heavy chain. Loop 3 is known to be involved in actin binding in some myosin isoforms but is not known to be important to ATP binding affinity (54). Cys707 is "SH2" in the SH1-SH2 helix, which when modified in other myosin isoforms inhibits the ATPase activity but is not known to abolish ATP binding. However, its oxidation is likely to affect the rearrangement of this important connector helix and could trap the motor in a conformation that could alter ATP binding. Note that the highly reactive SH1 (Cys717) is also likely to be modified, but we were unable to detect this peptide in the mass spectrometry experiment. SH1 alkylation is known to affect the ATPase activity but does not abolish ATP binding for iodoacetamide-labeled skeletal myosin (55) nor for spin-labeled skeletal myosin (although the rate of binding was slowed) (56). Alternatively, these modification(s) collectively may be sufficient to partially unfold the myosin. Interestingly, skeletal myosin oxidized by peroxynitrite loses the ability to form the ADP-Vi complex in addition to partially unfolding (57). Further studies will be required to understand the mechanism of the observations from our study.

We have shown that the presence of EDTA can diminish the H2O2-induced alterations in the kinetics of pHMM (Fig. 1), upHMM (Fig. 2), and upS1 (Figs. 3 and 4). EDTA chelates trace transition metals in the buffer, and transition metals are known to increase the rates of thiol oxidation by two general mechanisms (58). Metal-thiol complexes are more nucleophilic than the respective protonated thiols and can therefore react faster with oxidants such as H2O2. Also, transition metals can activate and deliver H2O2 to protein thiols, thus increasing the rate of thiol oxidation. Our data show that EDTA has a protective effect against H2O2-induced changes in kinetics, suggesting that thiol oxidation is a contributing factor.

Gel analysis revealed that there are at least two SH groups, one on the 65-kDa fragment and the other on the 26-kDa fragment, that become disulfide-bonded (reversible by DTT) upon H2O2 treatment and that EDTA can inhibit disulfide bond formation. The SH1 helix, with SH1(Cys717) and SH2(Cys707), is in the 26-kDa fragment. It is likely that SH1 is involved in the disulfide bonding because it is known to be highly reactive. However, our mass spectral data did not reveal the location of this disulfide bond.

An analysis of the correlation between loss of reactive thiols and ATPase activity (Fig. 5) shows that there is a biphasic loss of thiols correlated to the loss of ATPase mediated by H2O2. This biphasic relationship is mirrored in thiol alkylation studies of SMM where the rapid inactivation phase was attributed to reaction of a single ELC or rod thiol (Cys892). Our mass spectral data of upS1 (which does not contain Cys892 on the rod) show that three thiols in the ELC are partially oxidized to sulfonic acids. Therefore ELC oxidation is most likely responsible for the rapid phase of the loss of ATPase activity. The slow phase most likely involves modification of Cys717 (SH1) and the other thiols in Table 2.

An important question is whether or not it is possible that myosin oxidation is responsible for the observed H2O2-dependent decrease in maximal force generation in intact airway smooth muscle tissue (Fig. 9 and supplemental data). Under these conditions, the level of myosin thiophosphorylation is the same for the control and H2O2-treated tissues. Here we correlate the in vitro kinetics obtained with purified SMM to the in situ kinetics of force development. H2O2-treated tissue has diminished amplitude of force development, but the rate is not altered (Fig. 9). DTT treatment only partially restored the amplitude. These properties are remarkably consistent with the kinetic properties of purified SMM. It appears that a population of the myosin heads in both the muscle and purified protein are rendered incompetent. In the tissue this is manifested as a decrease in maximal force generation and in the purified preparations as a loss of tight ATP binding for a population of the myosin heads. The fact that DTT partially restored the maximal force is consistent with the in vitro finding that DTT can restore those myosin heads that had retained ATP binding affinity. This interpretation requires that the heads with altered kinetics can generate force in the muscle at a rate similar to those in untreated muscle. We conclude that myosin oxidation is likely to be at least partially responsible for the altered contractile properties of airway smooth muscle exposed to H2O2. This may be important in diseased states in which H2O2 is produced. Our findings have significance to previous work in the field in which similar levels of H2O2 have been proposed to affect various cellular processes upstream of cross-bridge cycling in smooth muscles. Our study shows that oxidation of myosin alone may be partially responsible for the observations.

In summary, we have shown that H2O2 treatment of pHMM and upS1 causes a decrease in the ATPase activity, whereas for upHMM the activity is increased. Together these effects lead to a loss in phosphorylation-dependent regulatory properties (Table 1). The changes in regulation are mediated by thiol oxidation, as evidenced by the inhibition of inactivation by EDTA, which can chelate heavy metals that enhance the oxidative power of H2O2; by the partial reversal of the inhibition by DTT; and by the fact that we observed interdomain disulfide bonding and loss of DTNB-reactive thiols. A major finding is that a population of myosin heads lost the ability to bind nucleotide, and this effect could not be reversed with DTT. A comparison of the in vitro myosin kinetic data with the rate and amplitude of force development in airway smooth muscle tissue leads to the notion that myosin is oxidized by H2O2 in situ, and the oxidation may be responsible for altering the muscle behavior.


   FOOTNOTES
 
* This work was funded by NIAMS, National Institutes of Health Grant AR 40917 (to C. R. C.), National Institutes of Health Grant HL 54757 (to K. A. J.), and National Institutes of Health Grant 1P20RR018751 (to the COBRE Smooth Muscle Plasticity, Cell to Proteomics Interface Core facility). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

Formula The on-line version of this article (available at http://www.jbc.org) contains supplemental Fig. S1 and supplemental methods. Back

1 To whom correspondence should be addressed: Dept. of Biochemistry and Molecular Biology/330, 1664 N. Virginia St., University of Nevada School of Medicine, Reno, NV 89557. Tel.: 775-784-7033; Fax: 775-784-1419; E-mail: cremo{at}unr.edu.

2 The abbreviations used are: RLC, regulatory light chain; ASM, airway smooth muscle; HMM, heavy meromyosin; S1, subfragment 1; MLCK, myosin light chain kinase; SMM, smooth muscle myosin; upHMM, unphosphorylated HMM; pHMM, thiophosphorylated HMM; MantATP, 2'(3')-methylanthranilyoyl-ATP; ELC, essential light chain; DTT, dithiothreitol; DTNB, 5,5'-dithiobis-(2-nitrobenzoic acid); MOPS, 4-morpholinepropanesulfonic acid; ATP{gamma}S, adenosine 5'-O-(thiotriphosphate). Back


   ACKNOWLEDGMENTS
 
We thank Darrel Loeffler for technical expertise in performing the isometric force measurements, Olivia John for myosin preparations, and Benjamin Madden at the Mayo Expression Proteomics and Protein Chemistry Facility for performing the mass spectral analysis.



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