AJP - Heart Calcium Transients and Cell-Sarcomere
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Am J Physiol Heart Circ Physiol 293: H654-H659, 2007. First published March 30, 2007; doi:10.1152/ajpheart.01314.2006
0363-6135/07 $8.00
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
293/1/H654    most recent
01314.2006v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via ISI Web of Science (2)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Rao, V. S.
Right arrow Articles by Guilford, W. H.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Rao, V. S.
Right arrow Articles by Guilford, W. H.

Alterations to myofibrillar protein function in nonischemic regions of the heart early after myocardial infarction

Vijay S. Rao,* Laura R. La Bonte,* Yaqin Xu, Zequan Yang, Brent A. French, and William H. Guilford

Department of Biomedical Engineering, University of Virginia, Charlottesville, Virginia

Submitted 30 November 2006 ; accepted in final form 28 March 2007


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Remote-zone left ventricular dysfunction (LVD) contributes to global reductions in contractile function after localized myocardial infarction (MI). However, the molecular mechanisms underlying this form of LVD are not clear. This study tested the hypothesis that myofibrillar protein function is directly affected in remote-zone LVD early after MI. Cardiac myosin and native thin filaments were purified from mouse myocardium taken from both the nonnecrotic zone adjacent to and the nonischemic zone remote from an infarct induced by 1 h of coronary occlusion followed by 24 h of reperfusion. Thin filament velocities were measured using the in vitro motility assay. Results showed that overall function was significantly reduced in samples from both the adjacent (43 ± 12% of control, n = 7) and remote (53 ± 8% of control, n = 13) zones when compared with control proteins (P < 0.05). Myosin from the remote zone propelled control thin filaments at reduced velocities similar to those measured above. In contrast, the Ca2+ sensitivity of remote-zone thin filaments over control myosin was unchanged from control thin filaments (half-maximal at pCa 6.32 ± 0.08 and 6.27 ± 0.06, respectively) but showed a 20% increase in velocity at saturating Ca2+ that parallels an increase in tropomyosin phosphorylation. Myosin dysfunction may be related to oxidation of cysteines in the myosin heavy chains or carbonylation of myosin binding protein-C. We hypothesize that phosphorylation of tropomyosin may serve a compensatory role, augmenting contraction during periods of oxidative stress when myosin function is compromised.

thin filaments; calcium sensitivity; contractility; left ventricular dysfunction; motility assay

LEFT VENTRICULAR DYSFUNCTION (LVD) as a result of myocardial ischemia may be classified as either irreversible (as in the case of infarct) or as potentially reversible (as in the cases of hibernation, stunning, and remote-zone LVD). Remote-zone LVD is unique in that it occurs in regions of the heart that never experienced ischemia. Remote-zone LVD has been documented in humans (3, 21) and in mice (7) using cardiac magnetic resonance imaging (MRI). The mechanisms underlying remote-zone dysfunction are unclear, but they contribute to global reductions in contractile function after localized myocardial infarction (MI). Compensatory hyperkinesis of the remote zone immediately after MI gives way to reduced contractility within 24 h (21), and this condition may persist for several days. This time course overlaps that of excess mortality resulting from pump failure in patients surviving initial MI, suggesting the possibility of a causal relationship. With nearly eight million MI occurring annually in the United States alone, the potential impact of remote-zone LVD is significant.

Mechanisms that may play key roles in remote-zone LVD include mechanical tethering to the infarcted region, changes in mechanical load, reduced coronary vasodilator function, oxidative stress and inflammation, altered Ca2+ handling, and alterations to the contractile apparatus. Any or all of these may contribute to LVD at different stages of myocardial response to ischemia or infarction. Indeed, inflammation and oxidative stress are believed to play major roles in stunning, hibernation, and remote-zone LVD. Ca2+ overload and elaboration of reactive oxygen and nitrogen species may lead to a loss of sensitivity of contractile filaments to Ca2+ (4), although the molecular mechanisms of this effect remain the subject of debate. In fact, a lowering of Ca2+ sensitivity after MI has yet to be observed in isolated thin filaments. Only in failing hearts has Ca2+ sensitivity been studied in isolated thin filaments, and the results of these two studies are contradictory (19, 30). Similarly, myosin function is normal in failing hearts (29) but has never been studied early after MI. The present study was undertaken to determine whether alterations to cardiac myosin or thin filament proteins may contribute to remote-zone LVD early after MI.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Proteins. All animal studies were approved by the Institutional Animal Care and Use Committee of the University of Virginia. Thirteen C57Bl/6 male mice were infarcted by a 1-h ligation of the left anterior descending coronary artery (7, 46) followed by 24 h of reperfusion. In brief, mice were anesthetized with pentobarbital sodium (100 mg/kg ip) and intubated with an endotracheal tube (PE-20). Respiration was controlled with a rodent ventilator. An incision was made parasternally between the third and fourth ribs and intercostal muscles with a cautery pen. The infarct was produced by passing a 7-0 silk suture beneath the left anterior descending artery at a point 1–2 mm inferior to the left auricle and then tightening it over the length of tubing. After 60 min, the suturing and tubing were removed to allow reperfusion. After reperfusion, the chest was closed in layers, and the endotracheal tube was removed. This procedure results in a well-characterized and spatially reproducible MI (7, 46). After 24 h, hearts were harvested, and the left ventricle (LV) including the interventricular septum was excised. The total LV mass of the MI hearts was ~120 mg.

The LV tissue was characterized as infarct, adjacent, and remote zones and separated into individual tissue sections (Fig. 1). Adjacent zone tissue consisted of a 1-mm cut around the infarcted region and comprised ~25% of the LV mass. The remote-zone tissue consisted of a region along the basal septum remote to the infarct and comprised ~35% of the LV mass. Proteins and samples from these regions will henceforth be referred to simply as "adjacent" and "remote." The adjacent region was identified visually and may have therefore contained infarct. It thus served as a buffer zone between the infarct and the remote myocardium. Subsections of each ~30 mg sample were used to prepare cardiac myosin and fluorescently labeled native thin filaments. Thin filaments and myosin were similarly prepared from noninfarcted surgical sham hearts (n = 5) and 18 nonsurgical control hearts (one for each of the experiments and shams).


Figure 1
View larger version (26K):
[in this window]
[in a new window]

  		Fig. 1. Diagrams depicting the tissue sources of myosin and thin filament proteins relative to the infarct. A: whole heart view of the ischemia-repurfused (I/R) heart. Dark gray, the area of infarct; light gray, the adjacent zone of the heart as defined by a 1-mm cut around the area of infarct; white, the remote zone; dotted lines, the cut made to remove right ventricle, which was excluded in preparations. B: a short-axis cross section of the heart depicting the areas of infarct, adjacent, remote, and right ventricle (RV) cut line. Infarct, adjacent, and remote tissue was exclusively collected around the left ventricle (LV) and included the interventricular septum. C: a flat view of the cut section used in preparations. Dotted line represents the fold line between the free wall and the interventricular septum. The remote-zone tissue was collected on left side of the fold line.

 
Cardiac myosin was prepared similarly to Shiverick et al. (37). Briefly, each sample (~15 mg wet wt) was homogenized using a glass-bead tissue grinder in 0.5 ml of 0.3 M KCl, 10 mM HEPES, 10 mM Na4P2O7, 4 mM MgCl2, 1 mM ATP, 10 mM dithiothreitol (DTT), and tosyl phenylalanyl chloromethyl ketone/tosyl lysyl chloromethyl ketone (TPCK/TLCK) as protease inhibitors (pH 6.45). The homogenate was centrifuged at 140,000 g for 60 min in a Sorvall M120XG ultracentrifuge to remove insoluble residue and actin. The supernatant was diluted with 0.01 M DTT and allowed to stand on ice for 1 h to precipitate myosin. The myosin was collected by centrifugation at 20,000 g for 20 min, and the pellet was dissolved in "myosin buffer" (0.3 M KCl, 25 mM imidazole, 1 mM EGTA, 4 mM MgCl2, and 10 mM DTT, pH 7.4).

Native thin filaments were isolated from the remaining tissue according to Lehman et al. (22). Each sample was homogenized using a glass-bead tissue homogenizer in 0.5 ml of 25 mM imidazole, 1 mM EGTA, 100 mM KCl, 5 mM MgCl2, 5 mM ATP, 10 mM DTT, TPCK/TLCK, and 1% Triton X (pH 6.45). The homogenate was centrifuged for 20 min at 40,000 g, and the pellet was discarded. Thin filaments were removed from the supernatant by centrifugation for 45 min at 200,000 g. The resulting pellet was resuspended in 0.2 ml of 25 mM imidazole, 1 mM EGTA, 100 mM KCl, 5 mM MgCl2, 5 mM ATP, and 10 mM DTT (pH 7.9). This mixture was again clarified for 5 min at 40,000 g, and the thin filaments were collected by a 45-min centrifugation at 200,000 g. The pellet was allowed to swell overnight at 4°C in 30 µl of 25 mM imidazole, 1 mM EGTA, 100 mM KCl, 5 mM MgCl2, 10 mM DTT, and 0.6 nmol of tetramethyl rhodamine isothiocyanate phallodin (pH 6.45). On day 2, the pellet was resuspended in 200 µl of 25 mM imidazole, 1 mM EGTA, 100 mM KCl, 4 mM MgCl2, 5 mM ATP, and 10 mM DTT (pH 7.9) and clarified by centrifugation for 10 min at 40,000 g. The supernate was used in motility experiments.

Motility assay. The in vitro motility assay was performed at 30°C and 1 mM ATP essentially as described by Warshaw et al. (42) with minor modifications (13). Thin filament velocities were determined by cross-correlation (5) using a custom plug-in for ImageJ (33). The total number of filaments per visual field and the fraction moving were determined manually. The "overall function," which is the velocity of all filaments, moving and nonmoving, was calculated as the product of mean velocity and fraction moving.

Thin filament Ca2+ sensitivity was measured over a range of Ca2+ concentrations in the motility buffer. Appropriate Ca2+ levels from pCa 8.0 to pCa 5.0 were calculated using MaxC software (32). Calculations were based on a buffer with pH 7.4 and ionic strength 60 using 1 mM EGTA as a Ca2+/Mg2+ chelator. Motility buffer is highly viscous because of the presence of 0.5% methyl cellulose, making it difficult to consistently aliquot fixed volumes. Therefore, aliquots of motility buffer were dispensed using a positive-displacement pipette (Rainin) and weighed. Density was determined using an analytical balance and a vendor-supplied density measuring kit (Mettler-Toledo). The volume of 100 mM CaCl2 to be added was determined based upon the weight and the density of motility buffer (1.007 g/cm3 at 20°C).

Immunochemistry and phosphoprotein analysis. SDS-PAGE and concurrent phosphoprotein and total protein staining were performed on each set of extracted thick and thin filament proteins (n = 5) as stated above. Precast NuPage-3-(N-morpholino)propanesulfonic acid 12% bis-Tris gels (Invitrogen) were used for electrophoresis to ensure sufficient separation of lower-molecular-weight thin filament proteins. Total protein concentrations were measured (Advanced Protein Assay, Cytoskeleton) to ensure consistent protein loading. Gels were washed and fixed before the application of stains. Pro-Q Diamond (Molecular Probes) phosphoprotein staining solution was applied to the gels and imaged using a Bio-Rad FX fluorescent scanner. Subsequently, gels were stained with Sypro Ruby (Molecular Probes) total protein stain and imaged as above. Sixteen-bit images of each gel were collected (Bio-Rad FX) and analyzed using the ImageJ gel analysis toolkit. Relative phosphorylation was based on the ratio of phosphoprotein to total protein for the same band.

Carbonylation was measured using an Oxyblot derivatization kit (Chemicon) according to the manufacturer's instructions. Protein nitration and glutathionation were measured by Western blot using antibodies against 3-nitrotyrosine (Upstate) and glutathione (Chemicon), respectively. Glutationation was assayed both in normally prepared proteins and a separate set prepared without reducing agents. Nitration was assessed both in native proteins and in trypsin-digested myosin fragments (1:200 trypsin-myosin at room temperature for 15 min). Blots were developed using the Western Breeze chromogenic and chemiluminescent systems (Invitrogen).

Cysteine oxidation. The relative number of accessible cysteines in proteins was assayed by purifying myosin and thin filaments without DTT from additional, independent tissue samples (as above for glutathionation) and incubating the proteins with 5-iodoacetomido fluorescein (5-IAF; MGT Inc), a thiol-reactive fluorescent dye. 5-IAF was added to protein samples at a 4:1 molar excess in the dark for 30 min at room temperature. A 10:1 molar excess of DTT was added to the mixture to quench excess 5-IAF. 5-IAF-reacted proteins were separated by SDS-PAGE in the dark. Gels were imaged using a Bio-Rad FX Molecular Imager using a 488-nm wavelength scanning laser and 530 nm emission filter. Gels were subsequently stained using Simply Blue for total protein and imaged. 5-IAF-to-total protein ratios were determined as the ratio of the integrated band densities for each protein using these two stains. ImageJ software was used for band quantification. A reduction in the 5-IAF-to-total protein ratio implies cysteine oxidation.

Statistics. The sample number (n) for statistics was taken as the number of independent animal experiments for a given treatment. Statistical comparison of means was by z-test, with a significance level of P < 0.05. For the in vitro motility assay, at least 50 thin filaments were tracked for each condition in each animal. When determining Ca2+ sensitivity, each condition was normalized against daily control thin filament velocities at 10–5 M Ca2+ and errors propagated (38).


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Myosin and thin filament function. Control experiments performed in the absence of Ca2+ resulted in negligible motility, confirming that the purified thin filaments were Ca2+ regulated. There were no regional differences in thin filament or myosin function in control hearts at pCa 5, comparing tissue from the anterolateral wall with the basal septum (data not shown), nor were there differences in thin filament or myosin function comparing tissue from the septal ventricular wall of surgical sham hearts (analogous to the remote zone after infarct) with control hearts (Fig. 2D). The mean velocity of protein from control hearts was 4.9 ± 0.2 µm/s.


Figure 2
View larger version (31K):
[in this window]
[in a new window]

  		Fig. 2. Myosin and thin filament function as assessed by thin filament velocity in the in vitro motility assay: mean thin filament velocity (black), mean fraction moving (white), and overall function (gray: a velocity obtained by the product of fraction moving and velocity). A: the function of contractile proteins matched by zone was compromised in both the adjacent and remote zones. B: thin filament function tested against control myosin exhibited only a modest decrease in fraction moving in the adjacent zone. C: myosin function was significantly altered in both the adjacent and remote zones. D: neither thin filament nor myosin function was altered in the septal myocardium of surgical shams. Error bars indicate SE. *P < 0.05 vs. control.

 
Regional reductions in thin filament velocity were observed in experimental protein preparations (n = 13), even in those from nonischemic regions remote from the infarct (Fig. 2A). The overall function of contractile proteins from myocardium adjacent to the infarct was 0.43 ± 0.12 of control, whereas that remote from the infarct was 0.54 ± 0.13 of control. The major effect was a large reduction (0.55 of control) in the fraction of filaments that moved, with less of an effect on the velocity of moving filaments (0.85 of control). There was also a notable drop in the number of thin filaments bound to the motility surface, suggesting an impairment of actin-myosin binding. These differences were not because of an absolute loss of protein, since there were no differences in the protein concentrations of myosins isolated from control, remote, and adjacent myocardium (250 µg/ml).

When control myosin was paired with thin filaments prepared from the adjacent and remote myocardium of infarcted hearts, no statistically significant differences in overall function were found at 10–5 M Ca2+ (0.79 and 1.09 of control, respectively; Fig. 2B). However, the fraction of filaments moving from the adjacent myocardium was significantly reduced, and there was a trend toward higher velocity in remote myocardium.

Thin filaments from control hearts were then paired with myosin from the adjacent and remote myocardium of infarcted hearts. In contrast to the function of thin filaments, overall function of myosin from infarcted hearts was significantly reduced to 0.56 ± 0.11 and 0.60 ± 0.16 of control (P < 0.05) when isolated from the adjacent and remote myocardium, respectively (Fig. 2C). These reductions in overall function did not differ significantly from those observed with regionally matched proteins, above. Both mean velocity and fraction of filaments moving were significantly affected.

Although the velocities of thin filaments from experimental hearts were unaffected at saturating Ca2+, Ca2+ sensitivity might still have been affected. We therefore measured the Ca2+ sensitivity of thin filaments from the remote zone. Thin filament proteins isolated from control hearts (n = 6) and the remote zones of infarcted hearts (n = 6) were run against control myosin in the in vitro motility assay over a range of Ca2+ concentrations ([Ca2+]). Figure 3 shows the resulting Ca2+ sensitivity curves. Neither the pCa50 (6.27 ± 0.06 vs. 6.32 ± 0.08) nor the Hill coefficients (1.1 ± 0.1 vs. 0.9 ± 0.1) were altered, comparing control with remote thin filaments, respectively. The results fail to show a statistically significant change in Ca2+ sensitivity between the control and remote-zone samples. However, there was a statistically significant 20% increase in velocity at saturating [Ca2+], verifying the trend observed in previous experiments.


Figure 3
View larger version (17K):
[in this window]
[in a new window]

  		Fig. 3. Thin filament Ca2+ sensitivity. Overall velocity was measured at various Ca2+ concentrations ([Ca2+]) for control (bullet) and remote-zone ({circ}) thin filaments and the Hill equation fitted (lines). There was no significant difference in Ca2+ sensitivity for control and remote thin filaments in the presence of control myosin. Maximum velocity measured at saturating [Ca2+] was significantly increased in the remote zone. Velocities are shown normalized to control velocities at saturating Ca2+. Error bars indicate SE.

 
Posttranslational modifications. SDS-PAGE gels of extracted proteins (n = 5) were stained for phosphoproteins. Among the phosphoproteins normally present in purified myosin and thin filaments, only tropomyosin exhibited a significant change in phosphorylation comparing the remote zone with control protein preparations (2.5x control; Fig. 4). The regulatory light chain of myosin, cardiac myosin binding protein-C (MyBP-C), and troponin I were phosphorylated but unchanged from controls.


Figure 4
View larger version (13K):
[in this window]
[in a new window]

  		Fig. 4. Relative phosphorylation of thick and thin filament proteins. Phosphorylation and total protein were assayed by Pro-Q Diamond (phosphoprotein) and Sypro Ruby (total) stains of purified myosin, and native thin filaments were separated by electrophoresis. The relative level of phosphorylation was determined as the ratio of band densities from the two stains for each phosphoprotein. Tropomyosin (TM) phosphorylation was significantly increased over control (*P < 0.05). Troponin I (TnI), cardiac myosin binding protein-C (MyBP-C), and the cardiac myosin regulatory light chain (RLC) were not changed significantly. Myosin heavy chain served as an internal negative control. Error bars indicate SE.

 
Western blots against commonly assayed markers of oxidative stress suggest an increase in carbonylation of MyBP-C in the remote-zone samples (1.7x control), although the difference is not statistically significant. Western blots of proteins revealed no protein glutathionation. There was also no measurable nitration of myosin heavy chain or tryptic digests of myosin as determined by Western blot. Neither was there any detectable nitration of the thin filament regulatory proteins. In contrast, actin was endogenously nitrated, but experimental samples were unchanged from controls. Similarly, although both myosin and actin are carbonylated in vivo, no significant change in carbonylation of these proteins was observed between control and experimental animals.

Staining of myosin SDS-PAGE gels with 5-IAF demonstrated a 25 ± 1% decrease in myosin heavy chain reactive cysteine-sulfhydryls in the remote zone compared with controls, suggesting that some available reactive cysteines were oxidized (Fig. 5). However, no significant change in cysteine modification was measured in the copurified proteins, MyBP-C, and the myosin light chains.


Figure 5
View larger version (19K):
[in this window]
[in a new window]

  		Fig. 5. Iodoacetomido fluorescein (IAF) staining for cysteine oxidation. A: IAF staining of the myosin heavy chain (MHC) and MyBP-C from control (Ct) and remote (Rm) hearts was performed with subsequent Simply Blue (Coomassie) staining for total protein (B). C: the level of cysteine oxidation was determined by a decrease in the ratio of band density of IAF/total protein. MHC exhibited a significant decrease in remote samples compared with controls.

 
Finally, SDS-PAGE gels show no change in cardiac myosin isoform content in our samples, ruling out a shift to the slower beta-isoform as an explanation for slowed velocities. However, an isoform change would not have explained the decrease in the fraction of filaments moving that we observed (Fig. 2). Furthermore, the half-life of myosin heavy chain in cardiac myocytes is ~5.5 days (25), so it is unlikely that a change in isoform could have happened on such a short time scale.


   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
These data suggest that LVD in the remote zone after MI may be due, in part, to posttranslational modification of myosin that is not reversible by reducing agents. Thin filament velocity on cardiac myosin, which is generally reflective of unloaded shortening velocity, is significantly reduced after MI, even in nonischemic areas of the myocardium. Reported deficits in circumferential shortening [0.81 of control (7)] and wall thickening [0.44 of control (45)] as measured by cardiac MRI in remote regions of mouse hearts after MI bracket the overall deficit in function observed here (0.6 of control). However, one must take care in drawing correspondence between thin filament velocities in vitro and in vivo measures of myocardial strain. Aside from differences in the constrained in situ geometry of myofibrillar proteins, ionic strength affects the kinetics of the cross-bridge cycle in fibers (36, 43) and in the motility assay (13). The in vitro motility assay is normally conducted at lower than physiological ionic strength.

Myosin dysfunction. The mechanism underlying the deficit in remote-zone myosin function is probably related to oxidative stress leading to modifications to the myosin heavy chain and MyBP-C. Nitric oxide, peroxynitrite (ONOO–), superoxide, and hydroxyl radicals have been implicated in myocardial dysfunction following MI (8, 12, 27, 48). Indeed, common products of reactions between these species and proteins, such as 3-nitrotyrosine and carbonyl groups, have been detected in diseased myocardium (23, 28, 47) and even in remote tissues following myocardial ischemia-reperfusion injury (23). Although not all of our proteins displayed markers of oxidative stress, MyBP-C was increasingly carbonylated in the remote zone, and the number of available reactive cysteines in myosin heavy chains was reduced.

Myosin possesses two particularly reactive cysteines (18), Cys-707 and Cys-697, among nine in the motor domain of the heavy chain. Both are located in a critical {alpha}-helix in myosin (34) where covalent modifications of either cysteine typically lead to complete inhibition of myosin (24, 35). Tiago et al. (39) reported that myosin may be inactivated by ONOO– through oxidative modification of one of the reactive cysteines, perhaps to sulfenic acid (1), leading to partial unfolding of the protein. Consistent with this, our data show a decrease in reactive cysteines in myosin heavy chain purified from remote-zone samples. Because we purified proteins in the presence of a strong reducing agent, loss of cysteines is not because of nitrosation but rather irreversible oxidation.

Carbonylation of MyBP-C may result from side-chain radical formation, perhaps via hydroxyl radicals (6). MyBP-C is thought to have two major functions in the thick filament. First, the carboxy terminal of MyBP-C serves to stabilize alignment along the thick filament through strong binding to LMM myosin and titin (14). Second, the amino terminal of MyBP-C may regulate contractility through binding to myosin subfragment 2 (S2) on the myosin heavy chain, thereby limiting myosin flexibility at the hinge region (20). An increase in carbonylation of MyBP-C may therefore impact myosin function by modifying these interactions. However, the standard motility assay eliminates thick filaments; therefore, any effects of MyBP-C oxidation in our assays are most likely because of its interactions with S2.

Tropomyosin phosphorylation. Although no change in Ca2+ sensitivity was observed in the remote zone, maximum velocity as assayed at saturating Ca2+ was significantly increased in parallel with phosphorylation of tropomyosin. It has been shown that phosphorylation of tropomyosin in regulated thin filaments increases actin-activated ATPase activity (15, 16). ATPase rates are positively correlated with velocity of actin filaments in the motility assay (44), and so increased tropomyosin phosphorylation would be expected to manifest as increased velocities of actin filaments in vitro. Notably, an increase in tropomyosin phosphorylation may also be related to oxidative stress. Houle et al. (17) showed an increase in nonmuscle tropomyosin-1 phosphorylation via a kinase downstream of the extracellular signal-regulated kinase pathway in response to oxidative stress in endothelial cells. This increase in tropomyosin-1 phosphorylation was correlated with an increase in cellular contractility. We should note, however, that the Ca2+ sensitivities of force and shortening velocity are significantly different measures and do not necessarily correlate (10). Thus measurements of force from isolated thin filaments in remote-zone LVD may show a change in Ca2+ sensitivity that we have not yet assayed.

Alternative explanations. Several explanations for the dysfunction in myosin and increase in thin filament velocity can be excluded. A change in cardiac myosin essential light chain to the atrial isoform would be expected to increase, not decrease, velocity (9) and would not result in a decreased fraction of moving filaments. Ischemia-reperfusion injury in rats has been reported to lead to proteolysis (40) or phosphorylation (2) of myosin light chain-1 in the ischemic zone, although their relationship to dysfunction in nonischemic tissue is unknown. Also, phosphorylation of troponin I, troponin T, MyBP-C, and myosin light chain-2 may serve a regulatory role in contractility (11, 31, 41), but our results show no significant change in phosphorylation of these proteins in the remote zone compared with controls. Troponin I truncation has been reported to occur in a rat model of myocardial stunning (26) but is associated with an increase in Ca2+ sensitivity of sliding velocity (10), which we did not observe. Furthermore, stunning occurs in transiently ischemic myocardium, while we examined nonischemic myocardium. We have not, however, assayed directly for troponin I truncation.

In conclusion, these findings suggest that the phosphorylation of tropomyosin may serve a compensatory role, augmenting contraction during periods of oxidative stress when myosin function is compromised. Unfortunately, it is not known how tropomyosin phosphorylation affects the force, velocity, or Ca2+ sensitivity in muscle. Neither is it known what kinase or series of kinases endogenously phosphorylate tropomyosin. These data hint at exciting new mechanisms for thin filament regulation, as well as for novel and potentially common noninherited dysfunctions of myosin.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by National Institutes of Health Grants AR-45604 (to W. H. Guilford), HL-58582, and HL-69494 (to B. A. French) and the generous support of the Department of Biomedical Engineering at the University of Virginia.


   ACKNOWLEDGMENTS
 
We thank the laboratories of Drs. Klaus F. Ley and Joel Linden for providing control mouse hearts and the Cardiovascular Research Center at the University of Virginia for scholarly support.


   FOOTNOTES
 

Address for reprint requests and other correspondence: W. H. Guilford, Dept. of Biomedical Engineering, Univ. of Virginia, Box 800759, Charlottesville, VA 22908 (e-mail: guilford{at}virginia.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.

* V. Rao and L. R. La Bonte contributed equally to this work. Back


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

  1. Alvarez B, Radi R. Peroxynitrite reactivity with amino acids and proteins. Amino Acids 25: 295–311, 2003.[CrossRef][ISI][Medline]
  2. Arrell DK, Neverova I, Fraser H, Marban E, Van Eyk JE. Proteomic analysis of pharmacologically preconditioned cardiomyocytes reveals novel phosphorylation of myosin light chain 1. Circ Res 89: 480–487, 2001.[Abstract/Free Full Text]
  3. Bogaert J, Bosmans H, Maes A, Suetens P, Marchal G, Rademakers FE. Remote myocardial dysfunction after acute anterior myocardial infarction: impact of left ventricular shape on regional function: a magnetic resonance myocardial tagging study. J Am Coll Cardiol 35: 1525–1534, 2000.[Abstract/Free Full Text]
  4. Bolli R. Why myocardial stunning is clinically important. Basic Res Cardiol 93: 169–172, 1998.[CrossRef][ISI][Medline]
  5. Cheezum MK, Walker WF, Guilford WH. Quantitative comparison of algorithms for tracking single fluorescent particles. Biophys J 81: 2378–2388, 2001.[ISI][Medline]
  6. Dean RT, Fu S, Stocker R, Davies MJ. Biochemistry and pathology of radical-mediated protein oxidation. Biochem J 324: 1–18, 1997.[ISI][Medline]
  7. Epstein FH, Yang Z, Gilson WD, Berr SS, Kramer CM, French BA. MR tagging early after myocardial infarction in mice demonstrates contractile dysfunction in adjacent and remote regions. Magn Reson Med 48: 399–403, 2002.[CrossRef][ISI][Medline]
  8. Feng Q, Lu X, Jones DL, Shen J, Arnold JM. Increased inducible nitric oxide synthase expression contributes to myocardial dysfunction and higher mortality after myocardial infarction in mice. Circulation 104: 700–704, 2001.[Abstract/Free Full Text]
  9. Fewell JG, Hewett TE, Sanbe A, Klevitsky R, Hayes E, Warshaw D, Maughan D, Robbins J. Functional significance of cardiac myosin essential light chain isoform switching in transgenic mice. J Clin Invest 101: 2630–2639, 1998.[ISI][Medline]
  10. Foster DB, Noguchi T, VanBuren P, Murphy AM, Van Eyk JE. C-terminal truncation of cardiac troponin I causes divergent effects on ATPase and force: implications for the pathophysiology of myocardial stunning. Circ Res 93: 917–924, 2003.[Abstract/Free Full Text]
  11. Garvey JL, Kranias EG, Solaro RJ. Phosphorylation of C-protein, troponin I and phospholamban in isolated rabbit hearts. Biochem J 249: 709–714, 1988.[ISI][Medline]
  12. Grill HP, Zweier JL, Kuppusamy P, Weisfeldt ML, Flaherty JT. Direct measurement of myocardial free radical generation in an in vivo model: effects of postischemic reperfusion and treatment with human recombinant superoxide dismutase. J Am Coll Cardiol 20: 1604–1611, 1992.[Abstract]
  13. Guo B, Guilford WH. The tail of myosin reduces actin filament velocity in the in vitro motility assay. Cell Motil Cytoskeleton 59: 264–272, 2004.[CrossRef][ISI][Medline]
  14. Harris SP, Rostkova E, Gautel M, Moss RL. Binding of myosin binding protein-C to myosin subfragment S2 affects contractility independent of a tether mechanism. Circ Res 95: 930–936, 2004.[Abstract/Free Full Text]
  15. Heeley DH. Investigation of the effects of phosphorylation of rabbit striated muscle alpha alpha-tropomyosin and rabbit skeletal muscle troponin-T. Eur J Biochem 221: 129–137, 1994.[ISI][Medline]
  16. Heeley DH, Watson MH, Mak AS, Dubord P, Smillie LB. Effect of phosphorylation on the interaction and functional properties of rabbit striated muscle alpha alpha-tropomyosin. J Biol Chem 264: 2424–2430, 1989.[Abstract/Free Full Text]
  17. Houle F, Rousseau S, Morrice N, Luc M, Mongrain S, Turner CE, Tanaka S, Moreau P, Huot J. Extracellular signal-regulated kinase mediates phosphorylation of tropomyosin-1 to promote cytoskeleton remodeling in response to oxidative stress: impact on membrane blebbing. Mol Biol Cell 14: 1418–1432, 2003.[Abstract/Free Full Text]
  18. Kielley WW, Barnett LM. The identity of the myosin subunits. Biochimica et Biophysica Acta 51: 591–593, 1961.
  19. Knott A, Purcell I, Marston S. In vitro motility analysis of thin filaments from failing and non-failing human heart: troponin from failing human hearts induces slower filament sliding and higher Ca(2+) sensitivity. J Mol Cell Cardiol 34: 469–482, 2002.[CrossRef][ISI][Medline]
  20. Korte FS, McDonald KS, Harris SP, Moss RL. Loaded shortening, power output, and rate of force redevelopment are increased with knockout of cardiac myosin binding protein-C. Circ Res 93: 752–758, 2003.[Abstract/Free Full Text]
  21. Kramer CM, Rogers WJ, Theobald TM, Power TP, Petruolo S, Reichek N. Remote noninfarcted region dysfunction soon after first anterior myocardial infarction. A magnetic resonance tagging study. Circulation 94: 660–666, 1996.[Abstract/Free Full Text]
  22. Lehman W, Vibert P, Uman P, Craig R. Steric-blocking by tropomyosin visualized in relaxed vertebrate muscle thin filaments. J Mol Biol 251: 191–196, 1995.[CrossRef][ISI][Medline]
  23. Liu P, Hock CE, Nagele R, Wong PY. Formation of nitric oxide, superoxide, and peroxynitrite in myocardial ischemia-reperfusion injury in rats. Am J Physiol Heart Circ Physiol 272: H2327–H2336, 1997.[Abstract/Free Full Text]
  24. Marriott G, heidecker M. Light-directed generation of the actin-activated ATPase activity of caged heavy meromyosin. Biochemistry 35: 3170–3174, 1996.[CrossRef][Medline]
  25. Martin AF, Rabinowitz M, Blough R, Prior G, Zak R. Measurements of half-life of rat cardiac myosin heavy chain with leucyl-tRNA used as precursor pool. J Biol Chem 252: 3422–3429, 1977.[Abstract/Free Full Text]
  26. McDonough JL, Arrell DK, Van Eyk JE. Troponin I degradation and covalent complex formation accompanies myocardial ischemia/reperfusion injury. Circ Res 84: 9–20, 1999.[Abstract/Free Full Text]
  27. Mihm MJ, Coyle CM, Schanbacher BL, Weinstein DM, Bauer JA. Peroxynitrite induced nitration and inactivation of myofibrillar creatine kinase in experimental heart failure. Cardiovasc Res 49: 798–807, 2001.[Abstract/Free Full Text]
  28. Mihm MJ, Yu F, Carnes CA, Reiser PJ, McCarthy PM, Van Wagoner DR, Bauer JA. Impaired myofibrillar energetics and oxidative injury during human atrial fibrillation. Circulation 104: 174–180, 2001.[Abstract/Free Full Text]
  29. Nguyen TT, Hayes E, Mulieri LA, Leavitt BJ, ter Keurs HE, Alpert NR, Warshaw DM. Maximal actomyosin ATPase activity and in vitro myosin motility are unaltered in human mitral regurgitation heart failure. Circ Res 79: 222–226, 1996.[Abstract/Free Full Text]
  30. Noguchi T, Kihara Y, Begin KJ, Gorga JA, Palmiter KA, LeWinter MM, VanBuren P. Altered myocardial thin-filament function in the failing Dahl salt-sensitive rat heart: amelioration by endothelin blockade. Circulation 107: 630–635, 2003.[Abstract/Free Full Text]
  31. Noland TA Jr, Kuo JF. Phosphorylation of cardiac myosin light chain 2 by protein kinase C and myosin light chain kinase increases Ca(2+)-stimulated actomyosin MgATPase activity. Biochem Biophys Res Commun 193: 254–260, 1993.[CrossRef][ISI][Medline]
  32. Patton C, Thompson S, Epel D. Some precaustions in using chelators to buffer metals in biological solutions. Cell Calcium 35: 427–431, 2004.[CrossRef][ISI][Medline]
  33. Rasband J. NIH ImageJ. Bethesda, MD: National Institutes of Health, 1997.
  34. Rayment I, Rypniewski WR, Schmidt-Base K, Smith R, Tomchick DR, Benning MM, Winkelmann DA, Wesenberg G, Holden HM. Three-dimensional structure of myosin subfragment-1: a molecular motor. Science 261: 50–58, 1993.[Abstract/Free Full Text]
  35. Root DD, Reisler E. Cooperativity of thiol-modified myosin filaments. ATPase and motility assays of myosin function. Biophys J 63: 730–740, 1992.[ISI][Medline]
  36. Seow CY, Ford LE. High ionic strength and low pH detain activated skinned rabbit skeletal muscle crossbridges in a low force state. J Gen Physiol 101: 487–511, 1993.[Abstract/Free Full Text]
  37. Shiverick KT, Thomas LL, Alpert NR. Purification of cardiac myosin. Application to hypertrophied myocardium Biochim Biophys. Acta 393: 124–133, 1975.
  38. Skoog DA, West DM. Fundamentals of Analytical Chemistry. New York: Saunders, 1982.
  39. Tiago T, Simao S, Aureliano M, Martin-Romero FJ, Gutierrez-Merino C. Inhibition of skeletal muscle S1-myosin ATPase by peroxynitrite. Biochemistry 45: 3794–3804, 2006.[CrossRef][Medline]
  40. Van Eyk JE, Powers F, Law W, Larue C, Hodges RS, Solaro RJ. Breakdown and release of myofilament proteins during ischemia and ischemia/reperfusion in rat hearts: identification of degradation products and effects on the pCa-force relation. Circ Res 82: 261–271, 1998.[Abstract/Free Full Text]
  41. Venema RC, Raynor RL, Noland TA Jr, Kuo JF. Role of protein kinase C in the phosphorylation of cardiac myosin light chain 2. Biochem J 294: 401–406, 1993.[ISI][Medline]
  42. Warshaw DM, Desrosiers JM, Work SS, Trybus KM. Smooth muscle myosin cross-bridge interactions modulate actin filament sliding velocity in vitro. J Cell Biol 111: 453–463, 1990.[Abstract/Free Full Text]
  43. West TG, Ferenczi MA, Woledge RC, Curtin NA. Influence of ionic strength on the time course of force development and phosphate release by dogfish muscle fibres. J Physiol 567: 989–1000, 2005.[Abstract/Free Full Text]
  44. Yamashita H, Sugiura S, Serizawa T, Sugimoto T, Iizuka M, Katayama E, Shimmen T. Sliding velocity of isolated rabbit cardiac myosin correlates with isozyme distribution. Am J Physiol Heart Circ Physiol 263: H464–H472, 1992.[Abstract/Free Full Text]
  45. Yang Z, Berr SS, Gilson WD, Toufektsian MC, French BA. Simultaneous evaluation of infarct size and cardiac function in intact mice by contrast-enhanced cardiac magnetic resonance imaging reveals contractile dysfunction in noninfarcted regions early after myocardial infarction. Circulation 109: 1161–1167, 2004.[Abstract/Free Full Text]
  46. Yang Z, Zingarelli B, Szabo C. Crucial role of endogenous interleukin-10 production in myocardial ischemia/reperfusion injury. Circulation 101: 1019–1026, 2000.[Abstract/Free Full Text]
  47. Zweier JL, Fertmann J, Wei G. Nitric oxide and peroxynitrite in postischemic myocardium. Antioxid Redox Signal 3: 11–22, 2001.[CrossRef][ISI][Medline]
  48. Zweier JL, Kuppusamy P, Williams R, Rayburn BK, Smith D, Weisfeldt ML, Flaherty JT. Measurement and characterization of postischemic free radical generation in the isolated perfused heart. J Biol Chem 264: 18890–18895, 1989.[Abstract/Free Full Text]




This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
293/1/H654    most recent
01314.2006v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via ISI Web of Science (2)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Rao, V. S.
Right arrow Articles by Guilford, W. H.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Rao, V. S.
Right arrow Articles by Guilford, W. H.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online
Copyright © 2007 by the American Physiological Society.