We employed single myofibril techniques to test whether the presence of slow skeletal troponin-I (ssTnI) is sufficient to induce increased myofilament calcium sensitivity (EC50) and whether modulation of EC50 affects the dynamics of force development. Studies were performed using rabbit psoas myofibrils activated by rapid solution switch and in which Tn was partially replaced for either recombinant cardiac Tn(cTn) or Tn composed of recombinant cTn-T (cTnT) and cTn-C (cTnC), and recombinant ssTnI (ssTnI-chimera Tn). Tn exchange was performed in rigor solution (0.5 mg/ml Tn; 20°C; 2 h) and confirmed by SDS-PAGE. cTnI exchange induced a decrease in EC50; ssTnI-chimera Tn exchange induced a further decrease in EC50 (in μM: endogenous Tn, 1.35 ± 0.08; cTnI, 1.04 ± 0.13; ssTnI-chimera Tn, 0.47 ± 0.03). EC50 was also decreased by application of 100 μM bepridil (control: 2.04 ± 0.03 μM; bepridil 1.35 ± 0.03 μM). Maximum tension was not different between any groups. Despite marked alterations in EC50, none of the dynamic activation-relaxation parameters were affected under any condition. Our results show that 1) incorporation of ssTnI into the fast skeletal sarcomere is sufficient to induce increased myofilament Ca2+ sensitivity, and 2) the dynamics of actin-myosin interaction do not correlate with EC50. This result suggests that intrinsic cross-bridge cycling rate is not altered by the dynamics of thin-filament activation.
- single myofibril
- contraction kinetics
- muscle mechanics
contraction of striated muscle is initiated upon binding of calcium to troponin (Tn). Tn is located on the thin filament and is composed of three subunits: Tn-C (TnC) that binds calcium, Tn-I (TnI) that inhibits cross-bridge formation, and Tn-T (TnT) that anchors the Tn complex to the tropomyosin filament. Tn-tropomyosin is located in the grooves that are formed within the double-stranded actin filament. Substantial progress has been made in elucidating the general molecular mechanisms by which calcium binding to Tn activates the thin filament and the formation of cycling cross-bridges, which leads to force development and muscle shortening. Myofilament activation is a highly cooperative process in that the relation between activating calcium ion concentration and force development is much steeper than would be predicted on the basis of activation of a single actin segment by Tn. The mechanisms that underlie this phenomenon are incompletely understood but undoubtedly involve transmission of signals along the thin filament, possibly involving tropomyosin (12, 15, 34, 44).
The properties of the thin-filament regulatory system are determined, among others, by the isoform composition of the Tn subunits and, in the heart, by phosphorylation of TnI and TnT (12, 34, 44, 49). In many species, but not all, the neonatal heart expresses exclusively slow skeletal TnI (ssTnI), which is completely replaced by cardiac TnI (cTnI) early during development (6, 16, 26, 39). It should be noted that although this is true in mice, rats, and humans, it is now clear there are species that express high levels of cTnI well before birth (22). Experiments on isolated chemically permeabilized myocardium have demonstrated enhanced myofilament calcium sensitivity and diminished pH sensitivity in the neonatal myocardium compared with adult myocardium (23, 42). Recent investigations have suggested that these phenomena may be due, in large part, to the presence of ssTnI in the neonatal cardiac sarcomere (1, 11, 13, 21, 28, 30, 53, 54). However, whether the presence of ssTnI alone is sufficient to induce enhanced myofilament calcium sensitivity, that is, regardless of thin or thick filament protein isoform composition, is not known. In addition, phosphorylation of TnI or TnT greatly affects myofilament calcium sensitivity in the heart (12, 34, 44, 49). Hence, these studies suggest that the signal transduction pathway leading from TnC calcium binding to thin-filament activation and cross-bridge formation are modulated by the structural state of the Tn complex.
The molecular switch model of Tn/tropomyosin regulation of thin-filament activation is sufficient to explain variations in steady-state myofilament calcium sensitivity (7, 15, 27). However, whether alterations in Tn structure affect the dynamics of myofilament force generation is less clear. Relaxation from the active state in muscle commences when the concentration of activator calcium ions declines due to the action of calcium pump SERCA in the sarcoplasmic reticulum (5). Simultaneous measurement of cytosolic calcium concentration and sarcomere shortening in isolated twitching cardiac myocytes has suggested that the rate of relaxation is limited either by the rate of calcium dissociation from TnC or by the rate of calcium removal from the cytosol (5, 45); similar experiments in isolated mechanically loaded twitching cardiac trabeculae, suggest that relaxation is not limited by the rate of calcium sequestration into the sarcoplasmic reticulum (2). Consistent with this notion, a decrease in myofilament calcium sensitivity, such as seen, for example, in response to β-adrenergic stimulation induced activation of protein kinase A (PKA) (43), is generally associated with an enhanced rate of pressure relaxation in the intact heart (5, 34, 38, 43, 44). Likewise, increased myofilament calcium sensitivity is generally associated with a decreased rate in pressure relaxation, for example, in the ssTnI replacement transgenic murine model (13) or following application of a myofilament calcium-sensitizing agent, such as bepridil (24). On the other hand, studies employing rapid flash photolysis of a calcium chelator have shown either an accelerated rate of myofilament force relaxation (19, 56) or no change upon PKA-mediated phosphorylation (18). In addition, enhanced myofilament force relaxation was reported in ssTnI-containing myocardium, despite increased myofilament Ca2+ sensitivity (19). Thus, whether myofilament calcium sensitivity and myofilament activation/relaxation dynamics are correlated is not known at present. Some groups have presented evidence that differences between fast and slow myosin heavy chain isoforms are exclusively kinetic in nature (35) and that tropomyosin and Tn isoforms are the major determinants of myofilament Ca2+ sensitivity (10, 32, 33), at least in cardiac muscle. In contrast, other groups have argued that the Ca2+ sensitivity of force does depend on the myosin heavy-chain isoform (14). Accordingly, here, we measured kinetics of force development and relaxation in single rabbit psoas myofibril preparations using solution-switching techniques to rapidly vary activator Ca2+ concentration ([Ca2+]). Myofilament Ca2+ sensitivity was altered by Tn exchange or application of the drug bepridil. Despite marked alterations in myofilament Ca2+ sensitivity, none of the dynamic activation-relaxation parameters were affected under any condition. Our results suggest that intrinsic cross-bridge cycling rate is not altered by the dynamics of thin-filament activation.
Isolation and mounting of myofibrils.
Single myofibrils or bundles of two to three myofibrils were prepared from fast skeletal muscle by homogenization of glycerinated rabbit psoas muscles, as described previously (36, 51). Rabbits were killed by intravenous administration of pentobarbital (120 mg/kg) through the marginal ear vein. All of the procedures were conducted in accordance with the official regulations of the European Community Council on use of laboratory animals (directive 86/609/EEC); the Ethical Committee for Animal Experiments of the University of Florence approved the study. All solutions to which the myofibrils were exposed contained a cocktail of protease inhibitors including leupeptin (10 μM), pepstatin (5 μM), phenylmethylsulphonyl fluoride (200 μM), and E64 (10 μM), as well as NaN3 (500 μM) and 10 mM DTT. Myofibril preparations were stored at 0–4°C and used for up to 5 days after preparation. An aliquot of myofibril suspension in rigor solution was injected into a chamber filled with ∼3 ml relaxing solution. The bath was mounted on the stage of an inverted microscope (Nikon TMD or Olympus IX-70). Myofibrils were attached to glass tools that were mounted on micromanipulators. Sarcomere length was determined by video microscopy and set to 2.4 μm. Experiments were performed at 15°C.
The compositions of the activating and relaxing solutions were calculated as described previously (36, 51). The solutions contained (in mM): 10 total EGTA, 5 MgATP, 10 free Mg2+, 10 MOPS; pH = 7.0. The CaEGTA-to-EGTA ratio was varied so as to obtain different values of free [Ca2+] in the range between 0.01 μM and 316 μM. Potassium propionate was added to adjust the final solution to an ionic strength of 200 mM and a monovalent cation concentration of 155 mM. Although continuous solution flow minimizes alterations in the concentration of MgATP and its hydrolysis products in the myofibrillar space, the measurements were made in the presence of 10 phosphocreatine and 200 U/ml creatine kinase to prevent any ADP gradients. Contaminant phosphate (around 170 μM in standard solutions) was reduced to <5 μM by a phosphate-scavenging enzyme system (purine-nucleoside phosphorylase with substrate 7-methyl-guanosine) (36, 51).
Recombinant Tn production and exchange into myofibrils.
Recombinant Tn subunits were cloned into pET-17b vector, expressed in BL21 (DE3) pLysS Escherichia coli, purified by column chromatography, and lyophilized as described previously (9, 50). The Tn complex was formed by reconstitution of the Tn subunits in 6 mol/l urea, followed by multiple dialysis steps at increasing lower strength, the final step of which was against rigor exchange solution. Exchange of Tn was as described previously with some modifications (8, 41, 50). Briefly, myofibrils were sedimented at 800 g, followed by resuspension into a rigor exchange solution containing (in mM): 1 EDTA, 3 Ca2+, 100 KCl, 10 MOPS, and 1 MgCl2. Next, myofibrils were exposed to either cTn or ssTnI-chimera Tn at 0.5 mg/ml for 2 h at room temperature, followed by two sedimentation/resuspension wash cycles with rigor solution. Control myofibrils were treated identically but without exposure to Tn.
The extraction/reconstitution of Tn in myofibrils was assessed using SDS-PAGE (12%) (36). Briefly, myofibrils containing endogenous fast skeletal Tn (fsTn), exchanged cTn, or exchanged ssTnI-chimera Tn were placed in a Laemmli sample buffer. Total protein content was determined with a colorimetric assay; each lane was loaded with 10 μg total protein. Gels were run for 4 h at 20°C, subsequently stained with Coomassie blue, and scanned, and the relative distribution of the various isoforms of TnI was determined by densitometry. Identification of TnI bands on the gels was confirmed by comigration with recombinant cTn or ssTnI-chimera Tn.
Apparatus for mechanical measurements and rapid solution changes.
Myofibrils selected for use were mounted horizontally between two glass microtools: a calibrated cantilevered force probe and a length-control motor (cf. Fig. 1) (36, 51). The myofibrils adhered strongly to the glass tools, which were positioned to maximize the attachment area using micromanipulators. The length of attached myofibrils between attachments was ∼50 μm. Isometric force was measured from the deflection of the bright-field shadow of the force probe projected onto a split photodiode (approximately ×100 magnification) and the known compliance of the force probe (1–3 nm/nN; ∼5 kHz resonance frequency). Myofibril shortening associated with force-probe compliance was kept ≤3% of initial length. Myofibrils were activated and relaxed by rapid translation of two continuous streams of relaxing and activating solutions flowing by gravity from a stepper motor-controlled double-barreled glass pipette placed within 0.5–1 mm of the preparation (cf. Fig. 1). The gravity-driven flow of each solution was ∼200 μl/min, flowing past the myofibril at ∼2 cm/s. Solution changes occurred with a time constant of 2–4 ms and were complete within 10 ms (36, 51). A release-restretch protocol was used to measure the rate of force redevelopment (ktr) at saturating [Ca2+] (7).
Data analysis and statistics.
Values of the rate of rise of tension following a step increase in [Ca2+] by fast solution switching (kact) or following a release-restretch maneuver (ktr) were estimated by fitting the force data to a single exponential by using iterative nonlinear least squares approximation (Kaleidograph). A similar procedure was employed to derive the rate of the second, fast phase of force relaxation (krel). The rate of slow force relaxation was estimated by linear regression. Force-[Ca2+] relationships were fit to a modified Hill equation, F(%) = 100% * [Ca2+]nhill/(EC50nhill + [Ca2+]nhill), also by nonlinear least squares curve fitting, where, F is force normalized to [Ca2+]-saturated force development, EC50 is [Ca2+] at which F is 50% (myofilament calcium-sensitivity parameter), and nhill is Hill coefficient (a measure of the level of cooperativity). Differences between means were analyzed statistically by one-way or two-way ANOVA as appropriate. Data are presented as means ± SE. P < 0.05 was considered significant.
The present study employed isolated rabbit psoas myofibrils to obtain both steady-state force-[Ca2+] relationships and rapid contraction dynamics. The method is illustrated in Fig. 1A, which shows a myofibril (here a bundle of ∼3 myofibrils) that was attached to an ink-coated glass force probe and a glass holding pipette. Fig. 1B illustrates the positioning of the attached myofibril close to the outflow opening of a double-barreled perfusion pipette. A fast stepper motor controlled the perfusion pipette's position, thus allowing the myofibril to be rapidly (∼5 ms) exposed to either solution stream, as is illustrated in Fig. 1C; the white and black symbols indicate the approximate positions within the laminar solution streams where the myofibrillar preparation is located upon solution switching. An example of a contraction-relaxation cycle is shown in Fig. 1D. The bar above the force trace indicates the rapid switch from relaxing conditions (white bar; [Ca2+] < 0.01 μM) to full activation (black bar; [Ca2+] = 316 μM) and back again to relaxing conditions, as induced by the rapid movement of the perfusion pipette (cf. Fig. 1C). Upon activation, force developed exponentially (kact parameter). During steady-state force development, the length of the attached myofibril was rapidly (∼1 ms) reduced by 20%, followed by a rapid restretch 20 ms later; this maneuver caused a rapid drop in force, followed by an exponential rise in force back to steady state (ktr parameter). Force relaxation commenced upon switching back to the relaxation solution in two distinct phases: a slow and close-to-linear phase, followed by final, more rapid exponential decay of force (slow phase relaxation parameters: duration and krel; fast-phase relaxation parameter: krel).
We used the whole Tn exchange technique (8, 41, 50) to partially replace the endogenous Tn complex in rabbit psoas myofibrils for cTn and a chimera complex composed of cTnT, cTnC, and ssTnI (ssTnI-chimera Tn). SDS-PAGE was used to confirm Tn exchange as illustrated in Fig. 2. Control, rabbit psoas myofibrils that were subjected to the exchange solutions but not recombinant proteins, were loaded in lane 1; ssTnI-chimera Tn exchanged myofibrils were loaded in lane 2, while cTnI exchanged myofibrils were loaded in lane 3. Lanes 4 and 5 show the purified cTn and ssTnI-chimera Tn, respectively. Exchange with cTn resulted in a reduction of endogenous fsTnI content and concomitant increase in cTnI content (cf. arrows; compare lanes 1 and 3). A similar exchange was observed upon treatment with ssTnI-chimera Tn (compare lanes 1 and 2). On average, recombinant Tn exchange resulted in ∼90% replacement of the native Tn in rabbit psoas myofibrils, consistent with previous reports where this technique was employed in skinned muscle (8, 36, 41, 50).
The impact of Tn exchange on the steady-state force-[Ca2+] relationship is summarized in Fig. 3. Fig. 3A shows an example of five activation-relaxation cycles, starting at saturating [Ca2+], followed by three consecutive activations at intermediate [Ca2+] and ending with a contraction again at saturating [Ca2+]. Steady-state force development in this series was a sigmoidal function of [Ca2+], as illustrated in Fig. 3B. Furthermore, this typical example illustrates that several contraction-relaxation cycles could be imposed to an attached myofibril with little rundown (cf. compare first and last contraction at [Ca2+] = 316 μM Fig. 3A). On average, force rundown was 8.4 ± 2.6% in the control nonexchanged group, 9.1 ± 1.0% in the cTnI-exchanged group, and 6.1 ± 1.1% in the ssTnI-chimera Tn group. Fig. 3C summarizes the average impact of Tn exchange on the steady-state force-[Ca2+] relationship. Consistent with previous reports, exchange of native fsTn for cTn resulted in an increase in myofilament Ca2+ sensitivity (left shift of the relationship), as well as a reduction in the level of cooperative activation (a reduction in the “slope” of the force-[Ca2+] relationship) (8, 31, 36). Exchange for ssTnI-chimera Tn further increased myofilament Ca2+ sensitivity without a further reduction in cooperativity. The average Hill-fit parameters are summarized in the legend to Fig. 3. Thus, as in some neonatal myocardium (1, 13, 21, 22), presence of ssTnI within the context of a sarcomere composed of fast skeletal thick and thin filaments was sufficient to induce an increase in myofilament calcium sensitivity beyond that induced by exchange with cTn. Maximum Ca2+ saturated force development, on the other hand, was not different between the groups (cf. Table 1).
The kinetic parameters of contraction and relaxation were measured in contractions initiated from relaxing conditions ([Ca2+] = 0.01 μM) to maximum activation ([Ca2+] = 316 μM) in the three groups of myofibrils in a separate series of experiments. Fig. 4 shows, superimposed, typical individual recordings obtained in an attached myofibril from each group. Fig. 4A shows the complete activation, release-restretch, and relaxation force response on a slow time scale, while Fig. 4, B–D, shows Ca2+ activation kinetics, force redevelopment, and relaxation kinetics, respectively, on a faster time scale. The arrows in this figure indicate the time at which the solution switch reached the attached myofibril. None of the activation and relaxation kinetic parameters were affected by either cTn or ssTnI-chimera Tn exchange. The average kinetic parameters derived from all of the data are summarized in Table 1. These data demonstrate that despite the marked impact of Tn composition on steady-state myofilament Ca2+ sensitivity (cf. Fig. 3), the dynamics of contractile activation and relaxation are invariant under these conditions.
To test further whether increased myofilament Ca2+ sensitivity decreases the rate of force relaxation, we also tested the impact of bepridil, a calcium-sensitizing agent known to retard Ca2+ dissociation from TnC (24). Because of this, the drug delays thin-filament deactivation upon switching from activating to relaxing conditions. Consistent with previous reports (24), application of 100 μM bepridil induced a marked increase in steady-state myofilament Ca2+ sensitivity as revealed by a left shift of the force-[Ca2+] relationship (cf. Fig. 5A). A set of two typical dynamic force relaxation recordings is shown superimposed in Fig. 5B; the average kinetic fit parameters obtained in all rabbit psoas myofibrils are summarized in Table 2. As was the case with Tn exchange (cf. Figs. 3 and 4), increased myofilament Ca2+ sensitivity upon application of bepridil did not significantly affect either the fast or slow phase of force relaxation.
In the current study, we employed single myofibril technique to test 1) whether the presence of ssTnI is sufficient to induce increased myofilament calcium sensitivity and 2) whether modulation of myofilament Ca2+ sensitivity affects the dynamics of force development. Studies were performed using rabbit psoas myofibrils in which Tn was partially replaced for either cTn or a chimera Tn composed of cTnT, cTnC, and ssTnI. In addition, myofilament Ca2+ sensitivity was increased by application of bepridil, a myofilament Ca2+- sensitizing agent. We found that incorporation of ssTnI into the fast skeletal sarcomere is sufficient to induce an increase in myofilament Ca2+ sensitivity. Furthermore, neither activation nor relaxation kinetics were affected under any of the conditions tested. These results suggest that myofilament Ca2+ sensitivity does not correlate directly with myofilament activation-relaxation dynamics.
Steady-state contractile activation.
Tension development at saturating activating [Ca2+] (Po) by control nonexchanged rabbit psoas myofibrils in the present study was comparable to that reported in our previous studies (36, 51) but lower than that reported by Bartoo et al. (4). Furthermore, exchange of endogenous Tn for either cTn or ssTnI-chimera Tn did not affect Po (cf. Table 1), consistent with our previous report on cTn exchange (36). Hence, assuming similar force generation by cycling cross-bridges, these data suggest that fully activated Tn induces a similar activation state of the fast skeletal thin filament, regardless of constituent Tn isoform composition. Previous work has suggested that substitution of cTnC for endogenous fsTnC in fast skeletal muscle leads to reduced levels of Po (31, 36). Therefore, the molecular interactions between the Tn subunits may be specific for each muscle type. Here, we found that this restriction does not apply to slow skeletal TnI in the setting of fast skeletal thick and thin filaments, an observation that is likely related to the fact that this Tn subunit is the predominant isoform found in the neonatal heart (6, 16, 26, 39). Nevertheless, it should be noted that the procedure used to exchange only TnC (31, 36) differs from the whole Tn exchange procedure employed in the present study. In addition, the neonatal heart may also express fsTnI (48). Hence, whether Tn isoform “mismatch” leads to reduced levels of Po remains to be determined.
The transmission of the activation signal from Tn to the fast skeletal thin filament did not appear to differ between the various muscle-type Tns, suggesting that the interaction between Tn and tropomyosin is not muscle-type specific. In contrast, myofilament Ca2+ sensitivity, as indexed by the EC50 parameter, significantly depended on the Tn type. Consistent with previous observations (36), exchange of endogenous fsTn by cTnI resulted in a marked increase in myofilament Ca2+ sensitivity and reduced level of cooperativity, as indexed by the Hill coefficient (cf. Fig. 3). We recently reported a comparative study on the force-[Ca2+] relationship between muscle types where we demonstrated that fast skeletal muscle is more sensitive to activator Ca2+ than cardiac muscle, but with similar levels of cooperativity and length-dependent activation (20). The mechanisms that lead to enhanced myofilament Ca2+ sensitivity upon cTn exchange into fast skeletal muscle are not known. In general, Ca2+ binding to the NH2-terminal domain of fsTnC exposes a “hydrophobic” patch to which a regulatory domain within TnI binds (15, 34, 44, 52). This binding is thought to detach the inhibitory peptide of fsTnI from a binding site on actin and allows movement of Tn and tropomyosin to expose myosin-binding sites on actin. In cardiac muscle, Ca2+ binding does not induce exposure of the “hydrophobic patch”; rather, residues within the cardiac regulatory domain bind to hydrophobic side chains on cTnI, resulting in detachment of this domain from actin, thereby activating the thin filament (15, 34, 44, 46). The binding of the inhibitory region of TnI to the NH2-terminal region of TnC is ∼6 times stronger in fsTn compared with cardiac Tn, but how these molecular interactions result in enhanced myofilament Ca2+ sensitivity in cTn exchanged myofibrils is less clear. It may be that, as previously suggested (36), due to the tight binding of fsTnI to fsTnC described above, the fast skeletal thin filament is kept in a lower state of activation than is possible when regulation is via cTn; less activator Ca2+ would then be required to release this inhibition, thus leading to enhanced myofilament Ca2+ sensitivity. A problem with this theory is that it is not clear how such a mechanism would lead to a reduced level of cooperativity as seen in the present study (cf. Fig. 3) and in previous studies (31, 36).
As in many neonatal hearts, presence of ssTnI in the present study on fast skeletal muscle induced enhanced myofilament Ca2+ sensitivity compared with exchange with cTn. As discussed above, it has been shown that the affinity of ssTnI to cTnC is higher than the affinity of cTnI to cTnC. This difference may be related to differences in amino-acid composition in the COOH terminus of TnI between these two isoforms (11, 28, 53, 54). There are charge differences between ssTnI and cTnI, making the slow muscle variant a less positively charged protein than the cardiac variant. Moreover, there are charge differences in functionally important microdomains between these molecules that may cause the neonatal sarcomere to be less sensitive to deactivation at low pH (23). How these differences lead to enhanced myofilament Ca2+ sensitivity when Tn contains ssTnI is not clear, however. The current data demonstrate that ssTnI-chimera Tn is equally effective when the signal is transmitted to fast skeletal contractile proteins (tropomyosin, actin, myosin, etc.). Hence, regardless of the precise molecular mechanisms that underlie the enhanced response to activator Ca2+ in the cardiac sarcomere when cTnI is replaced by ssTnI, these mechanisms are likely confined to the Tn complex itself.
Dynamic contractile activation.
The increased myofilament calcium sensitivity seen in the neonatal heart has been recapitulated in a transgenic murine model where cTnI is completely replaced by ssTnI (13). In that animal model, a reduced rate of pressure relaxation compared with wild-type animals is observed, which may be caused by enhanced myofilament Ca2+ sensitivity. On the other hand, a recent study employing rapid light flash-induced liberation of a calcium chelator, reported a significantly faster rate of force relaxation in transgenic ssTnI containing myocardium compared with wild-type cTnI containing myocardium (19). The modulation of [Ca2+] that can be achieved by flash photolysis, however, is somewhat limited in that full relaxation can only be accomplished in contractions starting at ∼70% of maximum activation. Thus, when studying contractions in muscles with varied myofilament Ca2+ sensitivity, either relaxation is initiated at a constant [Ca2+] but from varied levels of myofilament force development, or at different concentrations of [Ca2+] so as to achieve a constant level of force development. Interpretation of these data, therefore, may be limited due to technical shortcomings of the flash photolysis technique. Here, we employed a rapid solution-switching technique to activate and deactivate attached rabbit psoas myofibrils. This technique allows for the determination of activation and deactivation kinetic force measurements over the full range of activator [Ca2+]. In addition, the small diameter of the myofibrillar preparation virtually eliminates diffusion barrier-induced chemical gradients within the skinned muscle preparation (e.g., phosphate, ATP, etc.) (4, 17, 25, 36, 51).
Using this technique, we tested here whether increased myofilament Ca2+ sensitivity leads to altered rates of kact, ktr, or the two phases of krel (fast and slow). Myofilament Ca2+ sensitivity was altered either by Tn isoform composition or by application of bepridil, an agent known to enhance myofilament Ca2+ sensitivity by altering Ca2+ binding to TnC (24). None of these kinetic parameters were different between the various groups. This result suggests that the rate of force development following maximal Ca2+ activation of Tn is governed primarily by the intrinsic kinetics of the interaction between actin and myosin. Likewise, rapid removal of Ca2+ led to force relaxation from maximum activation at a rate (or rates) that were invariable and not related to myofilament Ca2+ sensitivity, implying that this kinetic step also is governed, primarily, by the intrinsic kinetic properties of the actin-myosin reactions, as has been suggested previously both by us (36, 51) and others (47). A similar result has recently been reported by Schoffstall et al. (40) employing the in vitro motility assay. Implicit in this theory is the assumption that Ca2+ binding/unbinding kinetics and the subsequent thin-filament activation dynamics are much faster than the kinetics of the actin-myosin interaction. Consistent with this notion, force relaxation is not dependent on the level of thin-filament activation prior to Ca2+ removal, but rather depends on the final [Ca2+] following the rapid solution switch (47, 51). Likewise, here, as in previous studies (36, 47, 51), both kact and ktr parameters were virtually identical, suggesting that both parameters are governed primarily by the transition rate of cross bridges from a weak, nonforce-generating state to a strong, force-generating state and not by the dynamics of Ca2+ binding and unbinding to TnC. In addition, these activation parameters have been shown to critically depend on the level of thin-filament activation (3, 7, 12, 15, 29, 34, 36, 37, 44, 47, 51, 55), a phenomenon that also can be explained by such a theory. The current experiments were not designed to specifically test whether myofilament Ca2+ sensitivity impacts on the modulation of kact by thin-filament activation state. Nevertheless, Fig. 6 shows a plot of the relative rate of force development vs. the relative level of steady-state force development obtained in control, nonexchanged myofibrils, cTn exchanged myofibrils, and ssTnI-chimera Tn exchanged myofibrils; this plot reveals no apparent differences between the groups. These data, albeit derived from relatively noisy force records (cf. Fig. 3), as well as the kinetic data obtained at saturating [Ca2+] (cf. Tables 1 and 2; Figs. 4 and 5) strongly suggest that myofilament Ca2+ sensitivity is predominantly determined by the properties of the Tn-tropomyosin-thin filament interaction, while the kinetics of force activation and relaxation are predominantly determined by the kinetics of the actin-myosin interaction.
These studies were supported, in part, by National Heart, Lung, and Blood Institute Grants PO1-HL-62426 and HL-75494, Telethone-Italy Grant GGP-02428, and European Union Grant HPRN-CT-2000-00091.
The authors thank Edward Allen for help with the Tn purification.
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.
- Copyright © 2007 the American Physiological Society