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Characterization of secophalloidin-induced force loss in cardiac myofibrils

  • Anna E. Bukatina
  • Gary C. Sieck
  • Kenneth B. Campbell
  • Marek Belohlavek
Original Paper
  • 60 Downloads

Abstract

Secophalloidin (SPH) is known to cause in cardiac myofibrils force without Ca2+ (half-maximal effect ~2 mM) followed by irreversible loss of Ca2+-activated force. At maximal Ca2+ activation, SPH increases force (half-maximal effect < 0.1 mM). We found that SPH at low concentration (0.5 mM) did not cause either force activation or force loss at pCa 8.7, but both of these effects did occur when force was activated by Ca2+. The force loss was prevented when SPH was applied during rigor or in the presence of 2,3-butanedione monoxime (85 mM). Furthermore, studying muscle in which the force was previously reduced by SPH (up to 50%) did not reveal significant changes in Ca2+ sensitivity and cooperativity of Ca2+ activation or qualitative alterations in SPH-induced changes in Ca2+-activated contraction. Data suggest that the force loss is mediated by cycling cross-bridges, and might reflect a reduction in force generated by individual cross-bridges.

Keywords

Actomyosin Ca2+ regulation Cardiac muscle Muscle contraction Myosin Myocardium Secophalloidin 

Abbreviations

AM

Actomyosin

AMP-PNP

Adenosine 5′-(β,γ-imido)triphosphate

A2P5

P1,P5-di(adenosine-5′) pentaphosphate

BDM

2,3-Butanedione monoxime

TnC

Troponin C

Tm

Tropomyosin

SPH

Secophalloidin

Introduction

Contraction of striated myofibrils starts by Ca2+ binding to troponin C (TnC), which triggers conformational changes in the thin filament regulatory complex resulting in tropomyosin (Tm) shift over the actin surface towards an “on” position. This allows formation of myosin cross-bridges strongly bound to actin, which generate force. Once formed, myosin cross-bridges participate in feedback regulation maintaining Tm in the “on” position and increasing Ca2+ binding to TnC that is critically important in cardiac myofibrils. Numerous studies have investigated contraction regulation in myofibrils, but intricate signal pathways in a multi-protein complex with many feedbacks remain to be clarified (Gordon et al. 2000; Moss et al. 2004; Fuchs and Martyn 2005; Hinken and Solaro 2007).

Secophalloidin (SPH), a phalloidin derivative weakly binding to the phalloidin site on actin (Wieland 1986), drastically changes the thin filament regulation in cardiac myofibrils (Bukatina et al. 2002). SPH is the first allosteric drug which can activate cross-bridge cycling in myofibrils without Ca2+, causing force comparable to that at maximal Ca2+ activation. SPH-induced activation is reversible since the muscle relaxes upon washing out SPH. However, the following Ca2+-activated force is considerably reduced. The SPH-induced force loss develops slowly and appears to be irreversible as we could not restore the initial force by washing. Also, this force loss is not mediated by loss or changes in TnC (the most obvious mechanism underlying such a force loss). Thus, SPH has a dual effect: Ca2+-independent muscle activation followed by irreversible loss of Ca2+-activated force. Both SPH effects come from binding to sites distinct from the phalloidin sites on actin. SPH and Ca2+ do not compete for binding sites but synergistically interplay in force activation. Ca2+ increases both the force maximally activated by SPH and SPH sensitivity of force (SPH concentration causing a half-maximal effect decreases from ~2 mM without Ca2+ to less than 0.1 mM at maximal Ca2+ activation). Conversely, SPH increases force at maximal Ca2+ activation and Ca2+ sensitivity of force (Bukatina et al. 2000, 2002, 2006; Bukatina and Sieck 2003a, b). This shows that SPH affects critical steps in the thin filament regulatory mechanism and, thus, provides a promising tool for cardiac muscle research. However, the mechanism of SPH action, including the myofibrillar domain containing SPH binding site(s), and the relationship between the force activating and inhibiting SPH-induced effects remain uncertain.

To further characterize the interaction between SPH and myofibrils in skinned cardiac muscles, we studied the relationship between the two main SPH effects, the ability to activate force and to cause irreversible force loss, exploring the fact that the force increase caused by low SPH concentrations is strongly regulated by Ca2+. In addition, we studied Ca2+ activation and SPH effects in the muscles, the force of which had been previously reduced by SPH. We found that, in the absence of Ca2+, SPH (0.5 mM) neither activates muscle force nor induces the force loss. Moreover, Ca2+ promoted both force activation by SPH and the SPH-induced force loss, an effect that is mediated by cycling cross-bridges, since neither Ca2+ itself nor rigor myosin cross-bridges advanced the force loss. These findings suggest that SPH is a novel probe for the intermediate state(s) of strongly bound cross-bridges. Furthermore, we did not find significant changes in Ca2+ sensitivity and cooperativity of Ca2+ activation or qualitative alterations in SPH-induced changes in Ca2+-activated contraction in the muscles, the force of which had been previously suppressed (up to 50%) by SPH. These data might suggest a reduction in force generated by individual cross-bridges.

Materials and methods

Muscle preparations

Cardiac muscle strips were dissected from the left ventricular free wall of fresh bovine hearts (obtained from the local abattoir) for the majority of experiments. In some experiments with rigor muscles, fresh pig hearts (obtained from animals euthanized after completion of an unrelated experimental study approved by the Mayo Clinic Institutional Animal Care and Use Committee) were used. Muscle strips were chemically skinned with 1% (v/v) Triton X-100 in relaxing (Bukatina et al. 2002) or rigor (Bukatina et al. 1998) solution and stored in solutions containing 50% (v/v) glycerol at −20°C.

Solutions

Relaxing solution contained (in mM): 75 KCl, 5 MgCl2, 5 ATP, 100 MOPS (pH 7.04), 2 EGTA, 10 creatine phosphate, and 250–300 U/ml creatine phosphokinase. Rigor solution contained (in mM): 128 KCl, 2 MgCl2, 100 MOPS (pH 7.04), and 2 EGTA. In both solutions, pH was adjusted with 1 M KOH. To adjust Ca2+ concentration, 0.1 M CaCl2 aliquots were added directly to solutions in the experimental well. Due to such a protocol, we could record the whole force–pCa curve using only 100 μl of solution, and, thereby, greatly reduce expenditure of SPH that was available only in very restricted amounts. However, addition of CaCl2 to our solutions caused pH changes (from 7.04 to 6.97 at maximal Ca2+ activation). These changes in pH were taken into account for pCa calculations (see below).

In some experiments, Mg · ATP analogs, Mg · adenosine 5′-(β,γ-imido)triphosphate (Mg · AMP-PNP) or Mg · ADP, were added to rigor solutions. In that case, 2 mM nucleotide was added to solution along with raising MgCl2 concentration to 5 mM to increase content of the Mg2+ · nucleotide complex. Such solutions also contained an ATP exhausting system, 10 mM glucose and 20 U/ml hexokinase along with a myokinase inhibitor, 0.2 mM P1,P5-di(adenosine-5′) pentaphosphate, to remove any possible ATP contaminations and to prevent ATP formation by myokinase. Before their use, these solutions were incubated at 20°C for 15 min. Ionic composition of solutions was calculated using a modified program COMIX (Perrin and Sayce 1967) with equilibrium constants as tabulated in (Fabiato 1981; Pettit and Siddiqui 1976) and 10−5 M Ca2+ contaminations at pH calculated for each Ca2+ concentration. The calculated value of d(pH)/d([Ca2+]total) = −0.0349 (pH units)/mM for \( {\text{Ca}}_{\text{total}}^{ 2+ } \) concentrations from 0 to 2 mM, was in a good agreement with experimentally determined pH changes (− 0.07 pH units) caused by addition 2 mM CaCl2 to a relaxing solution.

SPH (hydroxy-acid) synthesized by NeoRx Corporation (Seattle, WA) was the same batch that was used in our previous studies (Bukatina et al. 2002; Bukatina and Sieck 2003a, b). The preparation was stored in a desiccator with silica gel at −20°C. To add SPH to experimental solutions, aliquots of water SPH solutions were initially dried in Eppendorf tubes in a vacuum chamber. Subsequently, dried SPH was dissolved in an experimental solution just prior to use.

Mechanical experiments

The experimental set up and experimental protocols for studying isometric contraction were described earlier (Bukatina et al. 1998). Briefly, skinned muscle strips (maximal diameter 0.1–0.25 mm, length 1.5–2 mm) were glued between a force transducer (model AE801, Akers, Norway) and a post, lowered into a temperature controlled well (20°C) containing 100 μl of a solution, and sarcomere length was adjusted to 2.1–2.3 μm in a relaxing solution. During experiments, solutions were strongly agitated with a vibrating stainless steel wire. Prior to each experiment, several contraction–relaxation cycles were performed to evaluate the muscle preparation and to ensure stability. To activate muscle, aliquots of 0.1 M CaCl2 were added to the well solution near the vibrating wire. For other solution changes, the well was drained through a special port connected to a vacuum line and refilled with a new solution, or the muscle was transferred to a new well.

To measure Ca2+-dependence of force, cumulative force-calcium data was collected during activation cycles starting from relaxing solution followed by sequential increases in Ca2+ concentration until maximal force was reached, after that muscle was returned to a relaxing solution (Bukatina et al. 1998). Throughout the experiment, force was sampled each second using a custom built software based on LabView from National Instruments.

To estimate Ca2+ sensitivity and cooperativity of activation, values of steady-state Ca2+-activated force vs. Ca2+ concentration were fitted with the Hill equation \( F/F_{\max } = 1/(1 + 10^{{ ( {\text{pCa - pCa}}_{ 5 0} )n_{\text{H}}}}),\) where F is the muscle force, F max is the force at maximal Ca2+ activation, pCa = −log([Ca2+]), pCa50 is the Ca2+ sensitivity of myofibrils that equals to pCa at half-maximal force, and n H is the slope of the force–pCa curve (the Hill parameter characterizing cooperativity of Ca2+ activation).

Experimental results are reported as mean ± standard error (SE).

Results

Factors affecting development of SPH-induced loss of Ca2+-activated force

Effects of a number of factors on development of the SPH-induced loss of Ca2+-activated force were studied using the experimental protocols shown in Fig. 1. At the beginning of each experiment, force was maximally activated by Ca2+ (pCa 4.2) to measure a reference Ca2+-activated force (F 1, see Fig. 1a), followed by relaxation at pCa 8.7. Later in the experiment, an active, relaxed, or rigor muscle was exposed to SPH, and SPH was washed out with a relaxing solution. After that, the maximal Ca2+-activated force was re-measured during the testing activation (F 2, see Fig. 1a). The ratio F 2/F 1, a fraction of retained Ca2+-activated force after SPH treatment, was calculated for each experiment.
Fig. 1

Representative records illustrating experimental protocols for study the effects of Ca2+ and cross-bridges on SPH-induced force loss. Conditions during SPH application: a maximal Ca2+ activation (pCa 4.2); b intermediate Ca2+ activation (Ca2+ concentration in the SPH solution was adjusted to match initial maximal Ca2+-activated force); c relaxed muscle (pCa 8.7); d force was suppressed by 85 mM BDM at maximal Ca2+ activation (pCa 4.2); e rigor muscle. Control muscles underwent the same procedure, but SPH was not added, except a common control for the groups “a” and “b”, where muscles were maximally activated by Ca2+ for 15 min without SPH as shown in the right plot on panel b. In each record force was normalized to the maximal force during the reference Ca2+ activation. The bursts on the records reflect mechanical artifacts during solution change. In all panels, the lower drawings indicate pCa values calculated as described in text

Ca2+-activation of myofibrils promotes the SPH-induced force loss

To evaluate the role of Ca2+ activation in the SPH-induced force loss, muscle preparations in activating and relaxing solutions (Fig. 1, a–c) were exposed to low SPH concentration (0.5 mM) for 15 min, which did not activate appreciable muscle force in the absence of Ca2+ (in a relaxing solution, SPH-activated force was only 3.7 ± 0.2% (n = 6) of maximal Ca2+-activated force (Fig. 1c). Exposure to SPH in a relaxing solution (at pCa 8.7, when TnC does not contain bound Ca2+, the thin filament is in “off” position, and the cross-bridges are weakly bound to actin) did not cause any effect on Ca2+ activated force (Figs. 1c and 2c). In contrast, SPH treatment at maximal Ca2+ activation (started after 3 min equilibration with SPH in a relaxing solution) resulted in ~30% force loss (Figs. 1a and 2a). This was considerably more than the 2% force loss after contraction at maximal Ca2+ activation without SPH in control experiments. However, in agreement with our earlier data (Bukatina et al. 2000), 0.5 mM SPH considerably increased force generated at pCa 4.2 (by 32 ± 5%, n = 4). To exclude muscle deterioration caused by prolonged contraction at excessive force as a possible reason for observed force loss, we conducted force-matched experiments, where force in the presence of SPH was only partially activated by Ca2+ in order not to exceed the reference level in the absence of SPH (Fig. 1b). We found a 22 ± 1%, (n = 6) force loss in such experiments (Fig. 2b) despite the fact that force did not exceed the reference value at any time. Thus, Ca2+ activation was shown to promote SPH-induced force loss.
Fig. 2

Ca2+-activated force retained after interaction with SPH in active, relaxing and rigor conditions. a maximal Ca2+ activation; b intermediate Ca2+ activation; c relaxed muscle; d 85 mM BDM at maximal Ca2+ activation; e muscle in rigor (for more details see Legend Fig. 1). Data are presented as means ± SE. Symbol * indicates significant difference from control (P < 0.05, t-test). Numbers over columns denote number of experiments

Ca2+ does not promote the SPH-induced force loss, if force generation is prevented

During Ca2+ activation, the thin filament is activated by both Ca2+ and by cross-bridges, and the contractile apparatus contains cycling cross-bridges, which include various states of weakly and strongly bound cross-bridges. To separate the effects of Ca2+ and myosin cross-bridges during Ca2+ activation, we suppressed formation of force-generating cross-bridges with 2,3-butanedione monoxime (BDM), a myosin directed muscle relaxant (Herrmann et al. 1992), which inhibits the power stroke step of the cross-bridge cycle (Zhao and Kawai 1994; Regnier et al. 1995). Exposure to SPH at maximal Ca2+ activation (pCa 4.2) in the presence of 85 mM BDM, when muscle developed only 12 ± 1% (n = 6) of the reference force (Fig. 1d), did not cause any change in force compared to control (Fig. 2d). Based on this result, we suggested, that strongly bound cross-bridges, but not Ca2+ itself, are responsible for promotion of the SPH-induced force loss by Ca2+ activation.

Rigor cross-bridges do not promote the force loss

Based on our findings, we expected that SPH interaction with rigor muscles, where all cross-bridges are strongly bound to actin and the thin filament is in “on” position, would also result in a force loss. A rigor state was induced by washing out ATP from relaxed muscles with a rigor solution (Fig. 1e). To increase sensitivity, a higher SPH concentration (2 mM) was used in experiments with rigor muscles where there were no force changes upon SPH application (Fig. 1e). There was also no irreversible force loss after SPH application to rigor muscle at any pCa (Fig. 1e). In this experiment, cardiac muscle was incubated with 2 mM SPH for 20 min including 5 min at pCa 8.7, two 5-min incubations at intermediate Ca2+ concentrations, and 5 min at saturating Ca2+ concentration. Since SPH concentration was 4 times more than in experiments shown in Fig. 1 (panels a and b), the force loss would be seen at the end of the experiment, if irreversible force loss were to occur under any of these conditions. In agreement with this experiment, we did not find any significant force loss in settings where rigor muscle was incubated only at pCa 8.7 or pCa 4.2 during SPH administration. The irreversible force loss was also not observed in the presence of Mg · ADP (1.3 mM) or Mg · AMP-PNP (1.5 mM), Mg · ATP analogs which induce some conformational changes in cross-bridges resulting in a rigor force reduction (Marston et al. 1976; Bukatina et al. 1984; Horiuti et al. 2003). Pig heart samples were used in some of the experiments with rigor muscles. We confirmed that in pig muscles, SPH also produced a pronounced force loss after application to Ca2+-activated muscle: in force-matched experiments similar to that shown in Fig. 1b, the retained force was 59 ± 1% (n = 6). As no significant force loss was found in any experiment with rigor muscle, we combined data from all experiments for illustration in Fig. 2e. Based on these combined results, we concluded that the rigor cross-bridges do not advance SPH-induced force loss.

Thus, the results of previous sections showed that SPH induces the irreversible force loss in activated muscle (Fig. 2a, b), an effect prevented by BDM (Fig. 2d). The force loss also was prevented when SPH was applied to a relaxing (Fig. 2c) or rigor (Fig. 2e) muscle. Taken together, these results suggest the SPH-induced force loss is mediated by cyclic cross-bridges.

Contractile properties of muscles, force of which has been partially suppressed with SPH

To estimate changes in contractile properties associated with SPH-induced force loss, we studied the force–Ca2+ relationship and SPH-induced changes in Ca2+ activation as functions of Ca2+-activated force retained after incubation with SPH (1–4 mM) in the presence of Ca2+. Usually, muscle underwent several such incubations alternating with contractions in the absence of SPH (Fig. 3a). In this way, we could get more than 50% force loss.
Fig. 3

A representative experiment illustrating changes in contraction during progressive force loss caused by SPH. a A record of force throughout the experiment. During the experiment, muscle has undergone many Ca2+ activation cycles, including several stepwise increases in Ca2+ concentration until saturation. Force responses to the Ca2+ concentration increases are clearly seen on extended time scale in insertions, where each step in force reflects changes in pCa. Arrows depict moments of some changes in pCa to values indicated by numbers near the arrows. These data were used to estimate the force–pCa relationship. Time intervals of muscle activation and relaxation are showed by black and white areas, respectively, in the Ca2+ bar under the force record. Some of Ca2+ activation cycles (#4, 7, 9, and 15) were carried out in the presence of 1 mM SPH after several minutes of equilibration with a relaxing solution containing SPH. To continue experiment, SPH was washed out with a relaxing solution for at least 15 min, solution being renewed 3–4 times. Data were filtered using a 3rd order, 0.1 Hz low pass filter. The force bursts in the upper part of the curve, originating from solution substitution to a relaxing one, were removed manually; b Maximal active force (upper panel), the Hill parameter, n H (middle panel), and Ca2+ sensitivity, pCa50, in the course of the experiment. Open and filled symbols used for data collected without SPH and in the presence of 1 mM SPH, respectively. In the presence of SPH, maximal active force is shown, i.e. maximal force minus force before either SPH or Ca2+ application. In the absence of SPH, maximal Ca2+-activated force is indicated; c Ca2+ sensitivity and the Hill parameter as functions of F max in the absence of SPH. The regression lines equations are pCa50 = −0.0516F + 5.8893 (R 2 = 0.0301) and n H = −0.7003F + 3.195 (R 2 = 0.1179); d Activation of force by 1 mM SPH at maximal Ca2+ activation as a function of F max in the absence of SPH. SPH activation was determined as a ratio of maximal active force in the presence of 1 mM SPH to F max in the following activation. If several measurements of F max were available, average values were used. There is no correlation between force and SPH-induced force activation (P > 0.05, Pearson Product Moment Correlation). The regression lines equation is y = −0.001x + 1.6785 (R 2 = 0.4535)

During this work, maximal force loss (61%) was reached in an experiment similar to that showed in Fig. 3a, but using higher SPH concentration (4 mM) for 30 min during the 4th SPH application. This value is, likely, somehow overestimated since there is a natural force loss during prolonged experiments. Average force loss was 0.4 ± 0.2% (n = 3) per each Ca2+ activation cycle during prolonged experiments including many consecutive activation–relaxation cycles without SPH. Accepting this value, we concluded that SPH can cause a ~50% force loss.

Graphs of the Hill equation parameters, determined in each Ca2+ activation cycle in the representative experiment (Fig. 3b, middle and lower panels), show that, during the experiment, values of pCa50 and n H are rather stable in spite of considerable loss in force. This is seen even better from Fig. 3c, where the Hill equation parameters in the absence of SPH are presented as functions of retained F max in the course of the experiment. We did not find correlation between either pCa50 or n H with force in this experiment (P > 0.05, Pearson Product Moment Correlation).

This conclusion was confirmed by analysis of combined data collected from four muscle preparations (Fig. 4). Because the Hill equation parameters, pCa50 and n H, in different muscle samples varied over a significant range 5.79–5.87 and 2.64–2.97, respectively, we used normalized data for analysis. Normalized values, pCa50–pCa 50 control and n H/n H control , characterize the ratio of Ca2+ sensitivities (concentrations causing a half maximal force) and the ratio of the force–pCa slopes to those values before 1st SPH application, respectively. This analysis did not reveal statistically significant correlation.
Fig. 4

SPH-induced irreversible force loss occurs without significant changes in Ca2+ sensitivity or cooperativity. We did not find correlation between percentage of retained force and changes in Ca2+ sensitivity (pCa50–pCa 50 control ) or cooperativity (n H/n H control ) (P > 0.05, Pearson Product Moment Correlation). Data collected on four muscle preparations and values determined before 1st SPH application were taken as controls Regression lines equations are y = −0.0008x + 1.0735 (R 2 = 0.0899) for n H/n H control , and y = −0.0002x + 0.0023 (R 2 = 0.0093) for pCa50–pCa 50 control

As we published earlier (Bukatina and Sieck 2003b), partial activation by SPH causes a considerable increase in Ca2+ sensitivity and reduction in cooperativity (Fig. 3b, middle and lower panels, activation cycles #4, 7, 9, 15). Moreover, one can see that this difference in the values of Ca2+ sensitivity and cooperativity persists after a significant force loss, thus indicating that the SPH action on the mechanism of Ca2+ activation is not changed critically by the force loss. However, we did not analyze the Hill equation parameters of Ca2+ activation as functions of retained force in the presence of SPH because (1) SPH-induced force loss during each recording of cumulative force-Ca2+ data may have distinctive influence in different records since both the force loss magnitude and timing during collection of the data somehow differ, and (2) we have too little such data as they were not collected in each SPH application.

However, it is possible to estimate changes in force activation by SPH during the experiment. For this, we used the ratio of the maximal active force in the presence of SPH (1 mM) and saturating Ca2+ concentration to the F max in the following Ca2+ activation cycle without SPH (for example, the ratio of maximal active force during activation cycle #4 to F max during activation cycle #5). Determined this way, SPH activation for this preparation (162 ± 1%, n = 4) does not change much (changes were within 3%) throughout the experiment (Fig. 3d). Estimation using linear regression gives an increase in the SPH activation associated with 10% force loss equaled 0.6% for this experiment (Fig. 1d) or averaged 1.4 ± 1% for 3 muscle preparations. Thus, SPH effects on steady state Ca2+ activated contraction appear to be only weakly (if at all) sensitive to the SPH-induced force loss.

One can note, that the value of SPH-induced force at maximal Ca2+ activation (~60%) is considerably higher than reported by us earlier ~25% (Bukatina et al. 2000). The main reason of this difference is that earlier we compared the force in the presence of SPH with the force without SPH before SPH addition instead of with that after washing out SPH, as in the present study. The present method is less sensitive to the force loss between measurements of force with and without SPH, since after maximal force in the presence of SPH had been reached, the solution was quickly substituted with a relaxing one that not only washes out SPH, but also slows down the force loss, as it was shown in this study. Using the previous estimation method, the force activation during the 1st SPH application would be the ratio of maximal active force during activation cycle #4 to F max during the activation cycle #3 instead of that in the 5th activation cycle. The latter value is considerably less, thereby increasing the ratio. For the data shown on Fig. 3a, the mean SPH-induced force increase would be 34 ± 6%, instead of 62 ± 1%. Importantly, the SE is much larger than with the previous method (6 vs. 1) reflecting the differences in the force loss during activation cycles in the presence of SPH (activation cycles #4, 7, 9, and 15 in Fig. 3a). In addition, the present estimation of the force increase by SPH is in agreement with our previous data, where this value was within 50–75%, while active force at maximal Ca2+-activation in the presence of SPH was similar to the initial F max (Bukatina et al. 2002, Figs. 2 and 3). Thus, the present estimation method appears to be more correct.

Discussion

The first aim of this study was to get more insight on the relationship between two major SPH effects, the force activation and the irreversible force loss. We found that these two effects are closely connected. The SPH-induced force activation caused by an increase in concentration of either SPH or Ca2+ results in the loss of Ca2+-activated force. Our force-matched experiments suggest that the mechanical deteriorating effects hardly play a leading role in the force loss. This conclusion is supported by our unpublished observations from previous study (Bukatina et al. 2002) that the force loss (20–30%) can follow Ca2+-independent SPH activation causing only 70–90% of initial Ca2+ activated force.

Using a low SPH concentration that did not cause force without Ca2+, we found that (1) Ca2+ activation promotes SPH-induced force loss, an effect that is mediated by myosin cross-bridges but not by Ca2+ itself, and (2) The force loss is promoted by cycling cross-bridges but not by rigor cross-bridges (either in apo-form or in complex with Mg · ADP). The former result is in agreement with the force loss by high SPH concentrations without Ca2+. The latter result suggests that SPH probes the post-power stroke biochemical intermediate(s) of the cross-bridge cycle which precede the step of the irreversible AM** · ADP isomerization followed by a rapid release of ADP. These intermediates, the force generating states AM** · ADP · Pi and AM** · ADP, are structurally different from other cross-bridge intermediates (Gordon et al. 2000). Furthermore, this result may suggest that the SPH binding site is more available in the post-power stroke intermediates and located within the AM complex. In support of the latter idea, SPH binding site does exist within AM, as SPH (2 mM) causes a reduction in actin-activated ATPase activity of myosin S1 (S.S. Lehrer, personal communication).

Previously we found, that both SPH-induced force activation and the irreversible force loss (1) arise from sites distinct from those for phalloidin (Bukatina et al. 2002), and (2) are not mediated by TnC (Bukatina et al. 2002; Bukatina and Sieck 2003a). Taken together with our new results about a close connection between the two major SPH effects, these results are supportive of the hypothesis (Bukatina and Sieck 2003b) that both SPH effects could be consecutive steps of interaction at the same site where the first step (quick, reversible) results in the activating effect and the second step (slow, irreversible) leads to a loss of muscle force.

The second aim was to characterize properties of fibers, the force of which had been previously reduced by SPH. We did not find considerable changes in steady state isometric contraction using the force–Ca2+ relationship, or SPH-induced force increase.

A muscle force decrease may reflect a reduction in (1) the number of force generating cross-bridges or (2) the force generated by each force-bearing cross-bridge. Due to cooperative interactions within the contractile apparatus there is a positive correlation between the number of strongly bound cross-bridges and the Ca2+ sensitivity of force. Therefore, a reduction in the number of strongly bound cross-bridges is usually associated with a reduction in Ca2+ sensitivity. In cardiac muscle, cooperative Ca2+ activation is determined mainly by events within regulatory units of the thin filament and Ca2+ sensitivity of activation depends strongly on cross-bridges interaction with the turned “on” regulatory units (Metzger 1995; Fitzsimons et al. 2001; Gordon et al. 2003; Gillis et al. 2007).

Force suppression (to 31%) by inorganic phosphate, that reduces the number of strongly-bound cross-bridges by changes in kinetics of cycling cross-bridges, is associated with a reduction in Ca2+ sensitivity by 0.38 pCa units (Kentish 1986). Reduction in number of cycling cross-bridges by blebbistatin (Straight et al. 2003) via blocking of myosin heads in a low actin affinity state (Kovacs et al. 2004) reduces Ca2+ sensitivity by 0.34 pCa units at force reduction to ~25% (Dou et al. 2007) or by 0.16 pCa units at 50% force suppression (Baudenbacher et al. 2008). Reduction in the number of strongly bound cross-bridges by inactivation of functional units of the thin filament also associated with significant, though lesser, reduction in Ca2+ sensitivity (by ~0.09 pCa units at 50% force suppression as estimated by linear regression analysis using data in Table 1 from Gillis et al. 2007). These mechanisms can hardly underlay the force loss induced by SPH, as we did not find even a tendency to reduction in sensitivity.

On the other hand, a reduction in force of strongly bound cross-bridges could take place without changes in the Ca2+ sensitivity, at least theoretically. It was shown by study of soluble actomyosin system (Bremel and Weber 1972) or by activation of skinned muscles with a strong-binding, nonforce-generating analog of myosin subfragment-1 (Swartz and Moss 1992) that strongly bound myosin heads activate the thin filament without any force. Thus, a force reduction of strongly bound cross-bridges could be a basis of SPH-induced force loss. Alternatively, a reduction in the number of force generating cross-bridges could underlay the force loss, if suppression of strongly bound cross-bridges would occur in specific, clustered regions.

Notes

Acknowledgments

The authors are thankful to Dr. Eileen McMahon for critical reading of the manuscript and constructive comments. This study was supported in parts by NIH grants HL34817, and HL68555, and by a grant from GE Healthcare.

References

  1. Baudenbacher F, Schober T, Pinto JR, Sidorov VY, Hilliard F, Solaro RJ, Potter JD, Knollmann BC (2008) Myofilament Ca2+ sensitization causes susceptibility to cardiac arrhythmia in mice. J Clin Invest 118:3893–3903PubMedGoogle Scholar
  2. Bremel RD, Weber A (1972) Cooperation within actin filament in vertebrate skeletal muscle. Nat New Biol 238:97–101PubMedGoogle Scholar
  3. Bukatina AE, Sieck GC (2003a) Secophalloidin as a novel activator of skinned cardiac muscle. Biochem Biophys Res Commun 301:646–649CrossRefPubMedGoogle Scholar
  4. Bukatina AE, Sieck GS (2003b) Phallotoxins as an instrument for studying the intramyofibrillar regulation of muscle contraction. [Russian] Biofizika 48:1086–1093 (English version: Bukatina AE, Sieck GC (2003b) Myofibrillar regulation of contraction as revealed with phallotoxins. Biophysics 48:1003–1010)Google Scholar
  5. Bukatina AE, Son’kin BIa, Alievskaia LL (1984) Interactions of vanadate with the contractile system of the rabbit psoas muscle in active, relaxed and rigor states. [Russian] Biofizika. 29:280–283 [English version: Bukatina AE, Son’kin BY, Alievskaia LL (1984) Interactions of vanadate with the contractile system of the rabbit psoas muscle in active, relaxed and rigor states. Biophysics 29:303–313]Google Scholar
  6. Bukatina AE, Kirkpatrick RD, Campbell KB (1998) Dethiophalloidin increases Ca2+ responsiveness of skinned cardiac muscle. J Muscle Res Cell Motil 19:515–523CrossRefPubMedGoogle Scholar
  7. Bukatina AE, Kirkpatrick RD, Campbell KB (2000) Secophalloidin and phalloidin-(S)-sulfoxide as contraction modifiers for comparative study of skeletal and cardiac muscles. Tsitologiia 42:37–41PubMedGoogle Scholar
  8. Bukatina AE, Theodore LJ, Campbell KB, Sieck GC (2002) Calcium-independent activation of skinned cardiac muscle by secophalloidin. FEBS Lett 519:201–204CrossRefPubMedGoogle Scholar
  9. Bukatina AE, Sieck GC, Belohlavek M (2006) Phallotoxins as a novel tool for muscle research. In: Pandalai SG (ed) Recent research development in biophysics, vol 5. Transworld Research Network, Kerala, pp 257–267Google Scholar
  10. Dou Y, Arlock P, Arner A (2007) Blebbistatin specifically inhibits actin-myosin interaction in mouse cardiac muscle. Am J Physiol Cell Physiol 293:C1148–C1153CrossRefPubMedGoogle Scholar
  11. Fabiato A (1981) Myoplasmic free calcium concentration reached during the twitch of an intact isolated cardiac cell and during calcium-induced release of calcium from the sarcoplasmic reticulum of a skinned cardiac cell from the adult rat or rabbit ventricle. J Gen Physiol 78:457–497CrossRefPubMedGoogle Scholar
  12. Fitzsimons DP, Patel JR, Moss RL (2001) Cross-bridge interaction kinetics in rat myocardium are accelerated by strong binding of myosin to the thin filament. J Physiol 530:263–272CrossRefPubMedGoogle Scholar
  13. Fuchs F, Martyn D (2005) Length-dependent Ca2+ activation in cardiac muscle: some remaining questions. J Muscle Res Cell Motil 26:199–212CrossRefPubMedGoogle Scholar
  14. Gillis TE, Martyn DA, Rivera AJ, Regnier M (2007) Investigation of thin filament near-neighbour regulatory unit interactions during force development in skinned cardiac and skeletal muscle. J Physiol 580:561–576CrossRefPubMedGoogle Scholar
  15. Gordon AM, Homsher E, Regnier M (2000) Regulation of contraction in striated muscle. Physiol Rev 80:853–924PubMedGoogle Scholar
  16. Gordon AM, Rivera AJ, Wang CK, Regnier M (2003) Cooperative activation of skeletal and cardiac muscle. Adv Exp Med Biol 538:371–378PubMedGoogle Scholar
  17. Herrmann C, Wray J, Travers F, Barman T (1992) Effect of 2,3-butanedione monoxime on myosin and myofibrillar ATPases. An example of an uncompetitive inhibitor. Biochemistry 31:12227–12232CrossRefPubMedGoogle Scholar
  18. Hinken A, Solaro R (2007) A dominant role of cardiac molecular motors in the intrinsic regulation of ventricular ejection and relaxation. Physiology 22:73–80CrossRefPubMedGoogle Scholar
  19. Horiuti K, Yagi N, Takemori S, Yamaguchi M (2003) An x-ray diffraction study on the ADP-induced conformational change in skeletal muscle myosin. J Biochem 133:207–210CrossRefPubMedGoogle Scholar
  20. Kentish JC (1986) The effects of inorganic phosphate and creatine phosphate on force production in skinned muscles from rat ventricle. J Physiol 370:585–604PubMedGoogle Scholar
  21. Kovacs M, Toth J, Hetenyi C, Malnasi-Czismadia A, Sellers J (2004) Mechanism of blebbistatin inhibition of myosin II. J Biol Chem 279:35557–35563CrossRefPubMedGoogle Scholar
  22. Marston SB, Rodger CD, Tregear RT (1976) Changes in muscle crossbridges when beta, gamma-imido-ATP binds to myosin. J Mol Biol 104:263–276CrossRefPubMedGoogle Scholar
  23. Metzger JM (1995) Myosin binding-induced cooperative activation of the thin filament in cardiac myocytes and skeletal muscle fibers. Biophys J 68:1430–1442CrossRefPubMedGoogle Scholar
  24. Moss RL, Razumova M, Fitzsimons DP (2004) Myosin crossbridge activation of cardiac thin filaments: implications for myocardial function in health and disease. Circ Res 94:1290–1300CrossRefPubMedGoogle Scholar
  25. Perrin DD, Sayce LG (1967) Computer calculation of equilibrium concentrations in mixtures of metal ions and complexing species. Talanta 14:833–842CrossRefPubMedGoogle Scholar
  26. Pettit LD, Siddiqui KF (1976) The proton and metal complexes of adenyl-5’-yl imidodiphosphate. Biochem J 159:169–171PubMedGoogle Scholar
  27. Regnier M, Morris C, Homsher E (1995) Regulation of the cross-bridge transition from a weakly to strongly bound state in skinned rabbit muscle fibers. Am J Physiol 269:C1532–C1539PubMedGoogle Scholar
  28. Straight AF, Cheung A, Limouze J, Chen I, Westwood NJ, Sellers JR, Mitchison TJ (2003) Dissecting temporal and spatial control of cytokinesis with a myosin II Inhibitor. Science 299:1743–1747CrossRefPubMedGoogle Scholar
  29. Swartz DR, Moss RL (1992) Influence of a strong-binding myosin analogue on calcium-sensitive mechanical properties of skinned skeletal muscle fibers. J Biol Chem 267:20497–20506PubMedGoogle Scholar
  30. Wieland T (1986) Peptides of poisonous amanita mushrooms. Springer, New YorkGoogle Scholar

Copyright information

© Springer Science+Business Media B.V. 2009

Authors and Affiliations

  • Anna E. Bukatina
    • 1
    • 5
  • Gary C. Sieck
    • 2
    • 3
  • Kenneth B. Campbell
    • 4
  • Marek Belohlavek
    • 1
  1. 1.Division of Cardiovascular DiseasesMayo Clinic College of MedicineScottsdaleUSA
  2. 2.Department of AnesthesiologyMayo Clinic College of MedicineRochesterUSA
  3. 3.Department of Physiology & Biomedical EngineeringMayo Clinic College of MedicineRochesterUSA
  4. 4.Department of Veterinary and Comparative Anatomy, Pharmacology and PhysiologyWashington State UniversityPullmanUSA
  5. 5.ScottsdaleUSA

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