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Effects of Thin and Thick Filament Proteins on Calcium Binding and Exchange with Cardiac Troponin C

Jonathan P. Davis ^*, Catalina Norman ^*, Tomoyoshi Kobayashi , R. John Solaro , Darl R. Swartz  and Svetlana B. Tikunova ^*

^* Department of Physiology and Cell Biology, The Ohio State University, Columbus, Ohio; Department of Physiology and Biophysics, University of Illinois at Chicago, Chicago, Illinois; and Department of Animal Sciences, Purdue University, West Lafayette, Indiana

Correspondence: Address reprint requests to Jonathan P. Davis, Dept. of Physiology and Cell Biology, The Ohio State University, 209 Hamilton Hall, 1645 Neil Ave., Columbus, OH 43210. Tel.: 614-688-4467; Fax: 614-292-4888; E-mail: davis.812{at}osu.edu.

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
Understanding the effects of thin and thick filament proteins on the kinetics of Ca²⁺ exchange with cardiac troponin C is essential to elucidating the Ca²⁺-dependent mechanisms controlling cardiac muscle contraction and relaxation. Unlike labeling of the endogenous Cys-84, labeling of cardiac troponin C at a novel engineered Cys-53 with 2-(4'-iodoacetamidoanilo)napthalene-6-sulfonic acid allowed us to accurately measure the rate of calcium dissociation from the regulatory domain of troponin C upon incorporation into the troponin complex. Neither tropomyosin nor actin alone affected the Ca²⁺ binding properties of the troponin complex. However, addition of actin-tropomyosin to the troponin complex decreased the Ca²⁺ sensitivity (7.4-fold) and accelerated the rate of Ca²⁺ dissociation from the regulatory domain of troponin C (2.5-fold). Subsequent addition of myosin S1 to the reconstituted thin filaments (actin-tropomyosin-troponin) increased the Ca²⁺ sensitivity (6.2-fold) and decreased the rate of Ca²⁺ dissociation from the regulatory domain of troponin C (8.1-fold), which was completely reversed by ATP. Consistent with physiological data, replacement of cardiac troponin I with slow skeletal troponin I led to higher Ca²⁺ sensitivities and slower Ca²⁺ dissociation rates from troponin C in all the systems studied. Thus, both thin and thick filament proteins influence the ability of cardiac troponin C to sense and respond to Ca²⁺. These results imply that both cross-bridge kinetics and Ca²⁺ dissociation from troponin C work together to modulate the rate of cardiac muscle relaxation.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
Cardiac muscle utilizes troponin C (cTnC), an EF-hand Ca²⁺ binding protein, to sense and translate the myoplasmic Ca²⁺ signal into the cascade of events that ultimately leads to force production and/or shortening (for review see Kobayashi and Solaro (1)). In the muscle, cTnC is a subunit of the troponin (cTn) complex, consisting also of cTnI (inhibitory subunit) and cTnT (tropomyosin (cTm) binding subunit). The cTn complex is anchored to the thin filament through multiple protein interactions with actin and cTm. It is the dynamic, Ca²⁺-dependent interplay between these thin filament proteins that permits myosin to strongly bind to actin and produce force.

It is generally assumed that the thin filament system equilibrates with Ca²⁺ very rapidly (much faster than the mechanical events of contraction and relaxation) and thus must possess fast Ca²⁺ association and dissociation rates (for review Gordon et al. (2)). This definitely is the case for the Ca²⁺ binding properties of isolated cTnC (3). However, the Ca²⁺ sensitivity of the regulatory domain of cTnC is increased at least an order of magnitude by the binding of cTnI and cTnT (4,5). The increase in Ca²⁺ sensitivity of cTnC is primarily due to a slowing of the Ca²⁺ dissociation rate from cTnC as cTnI and then cTnT is added to cTnC (from >1000/s to 120/s then 35/s, respectively, at 15°C (6,7)). Thus, both cTnI and cTnT influence the Ca²⁺ binding properties of cTn in solution, by decreasing the rate of Ca²⁺ dissociation from cTnC. Furthermore, in the heart the cTn complex does not function in isolation but as part of the thin filament system.

It is unclear if cTm or actin ± myosin can further affect the kinetics of Ca²⁺ dissociation from cTnC. There are conflicting reports on the effects of cTm and actin-cTm on the steady-state Ca²⁺ binding properties of cTnC (8,9). The most common approach to follow Ca²⁺ binding to cTnC has been to label one or both endogenous Cys residues with a fluorescent probe. Part of the confusion in the literature may arise from the fact that fluorescent labeling of the endogenous Cys residues in cTnC alters the biochemical properties of cTn (8,10,11). Thus, to adequately address these questions, a more accurate reporter of Ca²⁺ binding and dissociation from cTn must be developed.

Consistent with the strong influence of cTnI on the Ca²⁺ binding properties of cTnC, developmental, transgenic, viral mediated, and possibly disease related incorporation of slow skeletal TnI (ssTnI) into cardiac muscle substantially affects cardiac muscle performance (12–22). For instance, in the presence of ssTnI, cardiac muscle force is: 1), more sensitive to Ca²⁺; 2), less sensitive to ß-adrenergic stimulation; 3), more resistant to pH changes; and 4), slower to relax. These physiological phenomena can occur in the presence of cTnC and cTnT, and can be attributed almost entirely to the incorporation of ssTnI into the myofilaments. The biochemical mechanisms behind the Ca²⁺ sensitizing effects of ssTnI have not been previously investigated.

In this article we report the design of a cTnC, fluorescently labeled at a novel position, that minimally affects the biological activity or the Ca²⁺ binding properties of cTnC in isolated cTn and ssTn. The fluorescent cTnC also reports the Ca²⁺ binding properties of the Tn complexes incorporated into a more physiologically relevant biochemical system, reconstituted thin filaments, in the absence and presence of myosin S1.

The objective of these studies was to systematically determine the effects of ssTnI, cTm, actin, and myosin S1 on Ca²⁺ binding and dissociation from the regulatory domain of cTnC in the Tn complex. The ultimate goal of these studies is to be able to better relate the biochemical behavior of cTnC to its physiological function.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
Materials
Phenyl-sepharose CL-4B, Bradford reagent, Tween-20, and EGTA were purchased from Sigma Chemical (St. Louis, MO). Quin-2 was purchased from Calbiochem (La Jolla, CA). 2-(4'-Iodoacetamidoanilo)napthalene-6-sulfonic acid (IAANS) and phalloidin were purchased from Invitrogen (Carlsbad, CA). Affi-Gel 15 affinity media was purchased from Bio-Rad (Hercules, CA).

Protein isolation/purification and mutagenesis
Recombinant human cTnC, cTnI, and cTnT3 (the most abundant cTnT isoform in adult cardiac muscle (23)); recombinant rat ssTnI; rabbit fast skeletal actin and myosin S1; and bovine cTm were isolated, purified, and quantified by standard laboratory procedures (7,8,24,25). The mutations C35S, T53C, and C84S in cTnC (herein denoted cTnC^T53C) were constructed from the cTnC pET3a plasmid utilizing techniques previously described (7).

Fluorescent labeling of cTnC^T53C
cTnC^T53C was dialyzed against 50 mM Tris, 6 M urea, 90 mM KCl, 1 mM EGTA, pH 7.5. Labeling was initiated by the addition of three- to fivefold molar excess IAANS to cTnC^T53C. The labeling reaction was allowed to proceed in the dark for 5–8 h with gentle rocking at 4°C. The reaction was stopped by addition of 2 mM DTT, and unreacted IAANS was removed by exhaustive dialysis against 10 mM MOPS, 90 mM KCl, pH 7.0. cTnC^T53C was determined to be 88 ± 7% (two different batches; all data are shown as a mean ± SE of the mean) labeled with IAANS (herein denoted ) utilizing the Bradford assay and the extinction coefficient for IAANS at 325nm of 24,900 M^–1cm^–1 (5).

Reconstitution of the Tn complexes
The Tn complexes were prepared following a modified protocol previously described by Tobacman and Lee (26). Briefly, the Tn subunits (TnC, TnI, and TnT) were first dialyzed separately against 10 mM MOPS (pH 7.0), 4.6 M urea, 1 mM DTT, and 0.01% NaN₃. After dialysis, the subunits were mixed at a molar ratio of 1:1.5:1.5 (TnC/TnI/TnT) and incubated at room temperature for 20 min. The complexes were subsequently dialyzed in sequential steps against 4 L of 10 mm MOPS (pH 7.0), 0.5 mM DTT, 3 mM MgCl₂, 0.01% NaN₃, and 1), 2 M urea, 1 M KCl; 2), 1 M KCl; and 3), 0.15 M KCl (this final buffer was repeated three times) at 4°C. Excess TnI and TnT that precipitated during dialysis were removed by centrifugation at 14,000 rpm for 20 min at 4°C (27).

Reconstitution of thin filaments
Purified actin filaments, myosin S1, and cTm were exhaustively dialyzed separately in 10 mM MOPS, 150 mM KCl, 3 mM MgCl₂, 1 mM DTT at pH 7.0. After dialysis actin was mixed with an equal molar ratio of phalloidin to stabilize the actin filaments. Control experiments determined that the addition of phalloidin to actin only marginally decreased the rates of Ca²⁺ dissociation from (-cTnI-cTnT; 1.3-fold) under the experimental conditions described below (data not shown). Actin-phalloidin (4 µM) and cTm (0.57 µM) were mixed in 10 mM MOPS, 150 mM KCl, 3 mM MgCl₂, 1 mM DTT at pH 7.0 and kept on ice for 20 min. Tn complexes (0.5 µM) were subsequently added to the actin-cTm mixture and kept on ice for an additional 15 min before use. Slightly substoichiometric Tn (7:1:0.88 (actin/cTm/Tn)) was utilized in the experiments to avoid the possibility of contaminating free Tn. Control experiments verified that the Ca²⁺ dissociation rates from the reconstituted thin filaments were not affected by increasing the actin/Tn ratios (ranging from 7:1:0.88 to 7:1:0.26 actin/cTm/Tn). Because the rates at the different actin/Tn ratios were indistinguishable, all the data were pooled and included in the average value (see Table 1). To address the effects of myosin binding on the Ca²⁺ binding properties of Tn, myosin S1 (1.14 µM) was added and allowed to equilibrate with the reconstituted thin filaments for 3 min. For all the data shown in the figures, the stoichiometry for the reconstituted thin filaments was 7:1:0.88:2 (actin/cTm/Tn/Mysoin S1). Initial experiments were performed utilizing bovine cardiac actin. However, due to contamination of cardiac actin with endogenous cTn and cTm, only 0.3 µM could be added to 4 µM of cardiac actin before unbound was observed via stopped-flow kinetics. This problem was not observed with rabbit fast skeletal actin. Moreover, incorporation of into reconstituted thin filaments containing cardiac or skeletal actin led to indistinguishable Ca²⁺ sensitivities and dissociation rates (data not shown). Thus, skeletal actin was utilized for the collection of data, because similar results were obtained and near stoichiometric concentrations of proteins could be used. Nonetheless, potential problems could also arise from using unmatched filament proteins (human cTn subunits, rat ssTnI, bovine cTm, and rabbit skeletal myosin S1). However, the biochemical results obtained in this work are consistent with previously observed physiological outcomes (14,28).

Determination of Ca²⁺ dissociation kinetics (k_off)
Ca²⁺ dissociation rates were measured using an Applied Photophysics (Leatherhead, UK) model SX.18 MV stopped-flow instrument at 15°C. The Ca²⁺ dissociation rates from the unlabeled Tn complexes were directly measured using the fluorescent Ca²⁺ chelator Quin-2. Quin-2 (150 µM before mixing) was excited using a 150-W xenon arc source at 330 nm with emission monitored through a 510-nm broad band-pass interference filter (Oriel, Stratford, CT). As in the steady-state measurements, IAANS fluorescence was excited at 330 nm. IAANS emission for the labeled Tn complexes was monitored through a 420–470 nm band-pass interference filter (Oriel) or the 510-nm broad band-pass interference filter. IAANS emission for the reconstituted thin filaments ± myosin S1 was monitored through the 510-nm broad band-pass interference filter, excluding myosin S1 experiments with in which the emission was monitored through a 415–490 nm band-pass interference filter (Newport, Irvine, CA). Each k_off represents an average of at least five traces, fit with a single exponential and repeated 14 times. The buffer used for the stopped-flow experiments was 10 mM MOPS, 150 mM KCl, 3 mM MgCl₂, 1 mM DTT, pH 7.0. To obtain the data traces, 200 µM Ca²⁺ was added to the Tn complexes (0.3 µM), or to the reconstituted thin filaments ± S1 and rapidly mixed with buffer containing 10 mM EGTA in the stopped-flow apparatus. Control experiments were also conducted, in which the reconstituted thin filaments ± myosin S1 with Ca²⁺ were rapidly mixed with buffer containing Ca²⁺, or reconstituted thin filaments ± myosin S1 in the presence of EGTA were rapidly mixed with buffer containing EGTA. These control traces were subtracted from the data traces to correct for scattering artifacts. The relative amplitudes of the scattering artifacts to the measured signals were 6.9 ± 0.6%, 16 ± 1%, and 6 ± 1% for the reconstituted thin filaments, in the presence of S1 and the subsequent addition of Mg-ATP, respectively (n 11).

Skinned cardiac trabecula experiments
Solutions for the trabecula force measurements were prepared as previously described (30). All experiments were performed at 15°C. Unbranched trabecula were dissected from the right ventricle of male Lewis-Brown Norway F1 rats and placed overnight at 4°C in relaxing solution containing 1% Triton-X. All trabeculae were used within 48 h of harvest. A reticule on the eyepiece of the dissecting microscope was used to measure the width and depth of the trabecula to calculate the cross-sectional area of the muscle by assuming an elliptical circumference. Aluminum T-clips were used to mount the trabecula between the arms of a high-speed length controller (model 322C, Aurora Scientific, Ontario, Canada) and an isometric force transducer (model 403A, Aurora Scientific) in relaxing solution. The resting sarcomere length was adjusted to 2.2 µm utilizing the first-order diffraction pattern from a HeNe laser. The trabecula was then activated in a pCa 4.0 solution and rapidly slackened after isometric force reached a plateau. The position of the high-speed length controller and the analog output of the force transducer were monitored by LabView 7.0 software (National Instruments, Austin, TX). The total force was measured between the plateau and baseline levels. The same procedure was utilized to obtain the resting force level of the trabecula in a pCa 9.0 solution. The active force generated by the trabecula in various pCa solutions was calculated as the total force minus the resting force. Average maximal active force per cross-sectional area before extraction of endogenous TnC was calculated to be 43 ± 7 kN/m², in which resting force accounted for 9 ± 1% of the total force (n = 21).

Endogenous cTnC was extracted by soaking the trabecula for 30 min in an extraction solution containing 10 mM HEPES, 5 mM EDTA, and 0.5 mM trifluoperazine dihydrochloride (TFP) at pH 7.0. The trabecula was then washed extensively with a pCa 9.0 solution to remove residual TFP. The average postextraction force was 2 ± 1% (N = 11) of the maximal preextraction force. cTnC was reconstituted into the trabecula by soaking the extracted trabecula for 30 min in a pCa 9.0 solution containing 16.7 µM wild-type TnC or The reconstituted trabecula was then exposed to a series of pCa solutions ranging from pCa 9.0 to pCa 4.0, to measure the Ca²⁺ sensitivity of force development. The maximally activated tension of the trabecula was obtained at pCa 4.0 before the extraction. pCa 4.0 activations were performed at the beginning, middle, and end of each force versus pCa experiment, averaged, and used to normalize the submaximal activations. During the course of the force versus pCa experiments, the rundown in force (% reduction in force of the last pCa 4.0 contraction versus the initial pCa 4.0 contraction) was 3 ± 1% (N = 21). For each protein, the force versus pCa experiment was repeated five to nine times.

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
Characterization of the biochemical and physiological properties of
To study thin and thick filament effects on Ca²⁺ binding and kinetics with cTnC, we developed a novel fluorescent cTnC, Thr-53 was mutated to Cys for the following reasons: 1), a Thr to Cys mutation is relatively conservative; 2), Cys can be selectively labeled with environmentally sensitive fluorophores, such as IAANS; 3), Thr-53, within the BC subdomain (consisting of helices B and C), moves away from the NAD subdomain (consisting of helices N, A, and D) as the regulatory domain of cTnC binds Ca²⁺ and interacts with cTnI, potentially changing its environment for spectroscopic analysis (Fig. 1, A and B); 4), Thr-53 does not directly ligate Ca²⁺ and is spatially separated from the Ca²⁺ binding loop, thus upon mutation should not directly interfere with Ca²⁺ binding; 5), Thr-53 does not interact with the residues within cTnC that contribute to the core of the hydrophobic pocket utilized to bind cTnI and that have been shown to modulate Ca²⁺ sensitivity of cTnC (7); and 6), Thr-53 itself does not interact with cTnI or cTnT (as can be observed from the Ca²⁺ saturated cTn structures (31)). Thus, substitution of Thr-53 with Cys and subsequent labeling with IAANS was expected to minimally affect cTnC function, and report the structural changes that occur in the regulatory domain of cTnC upon Ca²⁺ binding and dissociation.

View larger version (40K): [in this window] [in a new window]   FIGURE 1  behaves biochemically and physiologically similar to wild-type and endogenous cTnC. Panel A shows a ribbon representation of the regulatory domain of cTnC in the apo state (1SPY (53)) and in Ca2+ saturated cTn (1J1E; TnI, and TnT have been omitted for clarity (31)) utilizing the software Rasmol (54). The regulatory domain of cTnC contains five helices denoted as N, A, B, C, and D. As shown in panel A, helices B and C (BC subdomain) move away from the N, A, and D helices (NAD subdomain) upon Ca2+ and TnI binding. The D-helix is pointing out of the page, with the NAD subdomain colored light gray, the BC subdomain colored dark gray, and the Ca2+ binding loop indicated by an asterisk (*). Thr-53 is depicted in a stick representation. Panel B shows the IAANS emission spectra of the apo state (solid line) and Ca2+ saturated state (dashed line, pCa 3.0) for in the cTn complex (B1), in reconstituted thin filaments (B2), plus myosin S1 (B3), and plus ATP (B4). The emission fluorescence of the spectra was calculated relative to the peak fluorescence of each respective apo state, which was considered 100%. Panel C shows the time course of Ca2+ dissociation from wild-type cTn directly followed by an increase in quin-2 fluorescence. Panel C also shows the EGTA-induced time courses of Ca2+ dissociation from and reported by an increase in IAANS fluorescence. Overlaid with the kinetic traces are the fitted exponential curves to the data (smooth curves, which may be difficult to discern). Panel D shows the Ca2+-dependent increase in force development in skinned rat trabeculae containing endogenous cTnC (), wild-type cTnC (), and ().

To determine whether the fluorescence of reported the Ca²⁺-dependent structural changes associated with the regulatory domain of cTnC, stopped-flow studies were performed. Ca²⁺ dissociated from the regulatory domain of wild-type human cTn at a rate of 36.5 ± 0.4/s utilizing the fluorescent Ca²⁺ chelator quin-2 (Fig. 1 C), which was similar to what we have previously reported for the regulatory domain of wild-type human cTn under slightly different environmental conditions (6). As the [Ca²⁺]_free was rapidly chelated by EGTA, subsequently causing Ca²⁺ to dissociate from the IAANS fluorescence increased at a rate of 41.9 ± 0.4/s (Fig. 1 C and Table 1). Over a longer time (200 s) there was an additional 14% increase in fluorescence associated with the C-terminal domain of that occurred at 0.04 ± 0.01/s (data not shown). Thus, the decrease in IAANS fluorescence that occurs as binds Ca²⁺ can be largely attributed to the regulatory N-terminal domain of cTnC. Furthermore, it appears that the initial and major IAANS fluorescence increase occurs at the actual Ca²⁺ dissociation rate from the regulatory domain of cTn. This was not the case for the rate of IAANS fluorescence increase that occurred with the endogenously labeled Cys-35 of (10 ± 0.3/s; Fig. 1 C). Thus, as predicted and unlike it appears that the fluorescence of follows and minimally affects the Ca²⁺ binding properties of the regulatory domain of cTn. Consistent with this idea, skinned trabeculae reconstituted with developed force with nearly identical Ca²⁺ sensitivity (pCa 5.68 ± 0.03) and Hill coefficient (4.7 ± 0.5) values as wild-type cTnC (pCa 5.70 ± 0.02 and 5.5 ± 0.8 Hill coefficient) and endogenous cTnC (pCa 5.70 ± 0.02 and 6.5 ± 0.5 Hill coefficient; Fig. 1 D). Furthermore, was able to substantially recover force (79 ± 4%) to a similar extent as wild-type cTnC (79 ± 7%). Thus, behaved biochemically and physiologically similar to wild-type and endogenous cTnC, at 15°C.

Effects of ssTnI on the emission spectra of in systems of increasing complexity
We also wanted to test whether could be utilized to follow Ca²⁺ binding and dissociation from ssTn. Similar to Fig. 2 A shows that the fluorescence emission intensity of ( -ssTnI-cTnT) was sensitive to Ca²⁺ binding, and decreased 41% upon Ca²⁺ saturation at 450 nm. The Ca²⁺-dependent decrease in fluorescence intensity of was also only marginally affected by addition of cTm (data not shown). Again similar to Ca²⁺ saturation of incorporated into reconstituted actin-cTm increased the fluorescence intensity 39% at 450 nm (Fig. 2 B). In the presence of myosin S1, Ca²⁺ saturation of the reconstituted thin filaments induced a red shift in the maximal emission fluorescence and decreased the area under the emission spectra between 370 and 440 nm by 25% (Fig. 2 C), which could be reversed to the thin filament state by addition of Mg-ATP (Fig. 2 D). Thus, the Ca²⁺-dependent structural changes that occur in the regulatory domain of ssTn can be followed by the fluorescence of in the ssTn complex, and in reconstituted thin filaments in the absence and presence of myosin S1.

View larger version (18K): [in this window] [in a new window]   FIGURE 2  Effects of ssTnI on the IAANS emission spectra of in systems of increasing complexity. Panels A–D show the IAANS emission spectra of the apo state (solid line) and Ca2+-saturated state (dashed line, pCa 3.0) for in the ssTn complex (A), in reconstituted thin filaments (B), plus myosin S1 (C), and plus ATP (D). The emission fluorescence of the spectra was calculated relative to the peak fluorescence of each respective apo state, which was considered 100%.

View larger version (16K): [in this window] [in a new window]   FIGURE 3  Ca2+ sensitivities and dissociation rates from and Panel A shows the Ca2+-dependent decrease in IAANS fluorescence of () and (•). Panel B shows the time courses of Ca2+ dissociation from unlabeled cTnT53C and ssTnT53C directly followed by an increase in quin-2 fluorescence. Panel B also shows the EGTA-induced time courses of Ca2+ dissociation from and reported by an increase in IAANS fluorescence.

Ca²⁺ binding properties and Ca²⁺ dissociation rates from the Tn complexes in the presence of cTm, or actin ± myosin S1 without cTm
Because the Tn complex does not function in muscle in isolation but as an integral part of the thin filament, it is essential to understand the effects of actin and other relevant proteins on the behavior of cTn and ssTn. Fig. 4 A and Table 1 show that the addition of cTm to either or had only marginal effects on their Ca²⁺ sensitivities and apparent cooperativities. Fig. 4, B and C, also show that the addition of cTm had little effect on the rate of structural change in the fluorescent Tn complexes as Ca²⁺ was rapidly chelated by EGTA (for comparison see Table 1). Additionally, in the absence of cTm, neither actin alone nor actin in the presence or absence of myosin S1 affected the apparent rate of Ca²⁺ dissociation from (Fig. 4 B) or (Fig. 4 C). Thus, at the molar ratios used, neither cTm, actin, nor myosin S1 have nonspecific effects on the Tn complexes.

View larger version (16K): [in this window] [in a new window]   FIGURE 4  Ca2+ binding properties and Ca2+ dissociation rates from the Tn complexes in the presence of cTm, or actin ± myosin S1 without cTm. Panel A shows the Ca2+-dependent decrease in IAANS fluorescence of () and (•) in the presence of cTm. Panel B shows the EGTA-induced time courses of Ca2+ dissociation from (0.3 µM) in the presence of cTm (0.9 µM) or actin (4 µM) ± myosin S1 (0.57 µM) reported by an increase in IAANS fluorescence. Panel C shows the EGTA-induced time courses of Ca2+ dissociation from (0.3 µM) in the presence of cTm (0.9 µM) or actin (4 µM) ± myosin S1 (0.57 µM) reported by an increase in IAANS fluorescence. Increasing the concentration of or to 0.5 µM did not alter the results (data not shown).

View larger version (16K): [in this window] [in a new window]   FIGURE 5  Ca2+ sensitivities and dissociation rates from and reconstituted thin filaments. Panel A shows the Ca2+-dependent increase in IAANS fluorescence of () and () reconstituted thin filaments. Panel B shows the EGTA-induced time courses of Ca2+ dissociation from and reconstituted thin filaments reported by a decrease in IAANS fluorescence.

Effects of myosin S1 on the Ca²⁺ binding properties and Ca²⁺ dissociation rates from the Tn complexes in reconstituted thin filaments
It has long been known that strong cross-bridge binding increases the apparent Ca²⁺ sensitivity of cTnC in muscle (28). Consistent with this effect, Fig. 6 A shows that addition of myosin S1 to the reconstituted thin filaments increased the Ca²⁺ sensitivity of and 6.2-fold and 29-fold, respectively (Table 1). Similar to that of the isolated Tn complexes, the apparent Hill coefficients of the reconstituted thin filaments bound with myosin S1 indicated a negative cooperative Ca²⁺ binding process (Table 1). The effect of myosin S1 could be completely reversed by the addition of Mg-ATP, causing the Ca²⁺ sensitivity of the Tn complexes to be similar to that of the reconstituted thin filaments in the absence of myosin S1 (Fig. 6 A; Table 1). Furthermore, dissociation of attached myosin S1 from actin by addition of Mg-ATP increased the Hill coefficients back to that of the thin filament alone, which again indicated a positive cooperative Ca²⁺ binding process (Table 1). Additionally, in the presence of rigor myosin S1, displayed 6.5-fold higher Ca²⁺ sensitivity than However, the Ca²⁺ sensitivity of was only 1.8-fold higher than that of after addition of Mg-ATP (Table 1).

View larger version (16K): [in this window] [in a new window]   FIGURE 6  Effects of myosin S1 on the Ca2+ sensitivities and dissociation rates from and reconstituted thin filaments. Panel A shows the Ca2+-dependent decrease in IAANS fluorescence of () and (•) reconstituted thin filaments in the presence of myosin S1. Panel A also shows the Ca2+-dependent increase in IAANS fluorescence of () and () reconstituted thin filaments in the presence of myosin S1 and ATP. Panel B shows the EGTA-induced time courses of Ca2+ dissociation from and reconstituted thin filaments in the presence of myosin S1 reported by changes in IAANS fluorescence. The EGTA-induced decrease in fluorescence as opposed to the increase in fluorescence is due to the different emission filters used to capture the signals. Panel C shows the EGTA-induced time courses of Ca2+ dissociation from and reconstituted thin filaments in the presence of myosin S1 and ATP reported by an increase in IAANS fluorescence.

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
To determine the effects of myofilament proteins on cTnC function, we have fluorescently labeled cTnC at a novel position. This was accomplished by mutating the endogenous Cys residues to Ser and Thr-53 to Cys, which was then labeled with the environmentally sensitive fluorophore IAANS, resulting in When incorporated into the cTn complex, biochemically and physiologically responded to changes in [Ca²⁺] similarly to wild-type and endogenous cTnC. This cannot be stated for the functional behavior of cTnC labeled with IAANS on either endogenous Cys-35 or 84. Labeling of the endogenous Cys residues on cTnC has been shown to modify the Ca²⁺ binding properties of cTn and the Ca²⁺-dependent ATPase activity of cardiac reconstituted thin filaments (Fig. 1 C and (8,10,11)). Furthermore, the ability of to remain spectroscopically sensitive to Ca²⁺ binding in the Tn complex and on the thin filament is unique and different from that observed with single labeling of either endogenous Cys residue of cTnC (8,11). Thus, is a very useful and novel sensor to test the effects of thin and thick filament proteins on the Ca²⁺ binding properties of the cTn and ssTn complexes.

Neither cTm nor actin (± S1) affected the Ca²⁺ binding properties of the Tn complexes. However, reconstitution of the thin filaments decreased the Ca²⁺ sensitivity and increased the rate of Ca²⁺ dissociation from both Tn complexes. Thus, actin-cTm drastically attenuated the Ca²⁺ sensitizing effects of cTnI and cTnT on cTnC. This observation is consistent with cTnC and actin competing for binding to the C-terminal domain of cTnI (for review see Kobayashi and Solaro (1)). This competition for TnI binding may be the reason for the reduced Ca²⁺ sensitivity of the troponin complexes upon incorporation into the reconstituted thin filaments.

The positive cooperativity of Ca²⁺ binding observed in the reconstituted thin filaments may arise from Ca²⁺ binding to one stoichiometric unit (seven actins, one cTm dimer, and one cTn complex), allowing cTnC to out compete actin for cTnI, resulting in cTm repositioning on actin. This cTm movement is thought to span over a distance greater than a single stoichiometric unit (for review see Solaro and Rarick (33)) and may weaken the cTnI-actin interactions in neighboring stoichiometric units. These weakened cTnI-actin interactions will effectively increase the probability of cTnC binding to cTnI, increasing the apparent Ca²⁺ sensitivity of the neighboring cTnCs and thus cooperativity. Consistent with this hypothesis, positive cooperativity of Ca²⁺ binding was not observed with the Tn complexes alone or with "mini"-thin filaments that contain only a single stoichiometric unit (8,9,34).

If cTnC and actin truly act as competitive antagonists for cTnI binding, and this was the sole mechanism determining the Ca²⁺ sensitivity of cTnC on the thin filament, then the rate of Ca²⁺ dissociation from cTnC bound to cTnI in the different systems should not change. Alternatively or additionally, the Ca²⁺ sensitivity, cooperativity, and dissociation rate from the Tn complexes on the thin filaments may be modulated by cTnT-cTm interactions or direct cTnT-cTnC interactions (for review see Farah and Reinach (35)), which may change depending on the state of the thin filament. Consistent with cTnT and cTm influencing the Ca²⁺ sensitivity of cTn on the thin filament, isoform variations and disease-related mutations in cTnT and cTm have been shown to affect the Ca²⁺ sensitivity and cooperativity of cardiac muscle force development, even in the presence of ssTnI (6,36,37).

The myosin S1 effects on the reconstituted thin filaments were studied with two myosin S1 per stoichiometric unit. This concentration of myosin S1 is at the upper limit for the physiological range of myosin strongly bound to a stoichiometric unit during maximal contraction in muscle (for review (2,38)). The effect of myosin S1 on the Ca²⁺ sensitivity of the reconstituted thin filaments is consistent with strong binding cross-bridges increasing the Ca²⁺ sensitivity of cTnC in cardiac muscle (11,28,39,40). For cTn, the myosin S1 induced increase in the Ca²⁺ sensitivity of the thin filaments (approximately sixfold) could be explained by a proportionally similar decrease in the rate of Ca²⁺ dissociation from cTnC (approximately eightfold). The effects of myosin S1 binding to the thin filaments could be completely reversed by the addition of Mg-ATP. The molecular mechanism(s) behind these S1 effects on the thin filament are unknown. However, myosin binding to actin moves Tm further on actin than does Ca²⁺ binding alone (for review see Gordon et al. (38)) and drastically increases the affinity of cTm for actin (41). Thus, myosin binding to actin may cause a structural change in actin and move cTm to ultimately increase the probability of cTnI-cTnC binding. Furthermore, the positive cooperativity of Ca²⁺ binding to the thin filaments was abolished in the presence of myosin S1. Because our Ca²⁺ binding studies were performed after the addition of myosin S1, it is likely that the rigor binding of myosin S1 already displaced cTm so that the binding of Ca²⁺ to the Tn complexes had no effect on repositioning cTm (the proposed mechanism for the cooperativity). Consistent with the effects of myosin S1 on the thin filaments, the addition of NEM-S1 (an irreversible rigor like myosin S1) to cardiac muscle increased the Ca²⁺ sensitivity of force development with a reduced cooperativity (for review see Moss et al. (42)).

It is generally assumed that Ca²⁺ dissociation from cTnC in muscle is very rapid and that the rate of cross-bridge detachment is the primary determinant of cardiac muscle relaxation (for review see Gordon et al. (2)). Our data clearly demonstrate that myosin binding to actin drastically slows the rate of Ca²⁺ dissociation from cTn to a rate comparable to that of cardiac muscle relaxation (5–11/s at similar ionic strength and the same temperature used in this article (15°C); (43–45)). The fact that myosin binding slows the Ca²⁺ dissociation rate from cTnC incorporated into the thin filament fits well with recent mathematical models predicting this phenomenon to describe steady-state (46) and kinetic (47) muscle data. Assuming the reconstituted thin filament is a reasonable model system for studying the kinetics of Ca²⁺ dissociation from cTnC in muscle, then it may be that both cross-bridge kinetics and Ca²⁺ dissociation from cTn work together to modulate the rate of cardiac muscle relaxation. However, the reconstituted thin filaments represent only two extreme conditions, unbound myosin and rigor bound myosin. At the physiological salt conditions used in this work it is not possible to address the effects of cycling cross-bridges on cTnC.

Consistent with the fact that ssTnI increases the Ca²⁺ sensitivity of force development and slows the rate of cardiac muscle relaxation and relengthening (14–17,21), we have demonstrated that biochemically ssTn displayed a higher Ca²⁺ sensitivity and slower Ca²⁺ dissociation rate than cTn, under all experimental conditions. These data further support the hypothesis that the Ca²⁺ dissociation rate from Tn may influence the rate of cardiac muscle relaxation. Interestingly, there are numerous cTnI mutations linked to hypertrophic cardiomyopathy that also display increased myofilament and force Ca²⁺ sensitivities (8,48,49). However, transgenic expression of ssTnI in an adult heart does not trigger a hypertrophic response and actually improves cardiac muscle performance under various pathophysiological conditions (14,50–52). It may be that the hypertrophic cardiomyopathy related cTnI mutants increase the myofilament Ca²⁺ sensitivity through different mechanism(s) from that of ssTnI. For instance, unlike ssTnI, several hypertrophic cardiomyopathy linked cTnI mutants did not directly increase the Ca²⁺ sensitivity of cTnC in the Tn complex (8).

Steady-state affinities are set by the rates of association and dissociation. It is clear from the data that the relative changes in the Ca²⁺ sensitivity of the Tn complexes that occur upon incorporation into the reconstituted thin filaments in the absence and presence of myosin S1 are not always associated with proportional changes in the respective Ca²⁺ dissociation rates. Contrary to dogma, these results imply that the Ca²⁺ association rates to cTn and ssTn in the various biochemical systems must also change. This hypothesis is consistent with the fact that several Ca²⁺ sensitizing mutations in cTnC^F27W increased the Ca²⁺ association rates to the regulatory domain of isolated cTnC (7). As a consequence of this phenomenon, ssTnI has a much larger effect on the Ca²⁺ sensitivity of the Tn complex incorporated into the reconstituted thin filaments in the presence of myosin S1 than on the rate of Ca²⁺ dissociation, compared to that of cTnI. This mechanism would allow cardiac muscle expressing ssTnI to be substantially more sensitive to Ca²⁺, while only marginally affecting the rate of cardiac muscle relaxation. Consistent with our results, cardiac muscle expressing ssTnI exhibits a much larger increase in the Ca²⁺ sensitivity of force development than a decrease in the rate of muscle relaxation or relengthening (14,17,21).

In summary we developed a novel fluorescent cTnC that accurately reports the rate of Ca²⁺ dissociation from cTnC incorporated into cTn and ssTn. Furthermore, the fluorescent cTnC also reports the Ca²⁺ binding properties of the Tn complexes reconstituted into thin filaments in the absence and presence of myosin S1. This study demonstrated: 1), reconstitution of cTnC into the thin filament dramatically increases the Ca²⁺ dissociation rate from cTn; 2), myosin S1 substantially decreases the rate of Ca²⁺ dissociation from cTn reconstituted into thin filaments to a rate similar to reported rates of cardiac muscle relaxation; 3), ssTn possesses a higher Ca²⁺ sensitivity than cTn in all systems studied; and 4), ssTn possesses a slower Ca²⁺ dissociation rate than cTn reconstituted into thin filaments in the absence and presence of myosin S1. Thus, all the major thin filament proteins (TnI, TnT, Tm, and actin) have direct or indirect effects on the Ca²⁺ sensitivity and dissociation rate from cTnC. Even the thick filament protein myosin can drastically alter the sensitivity and kinetics of Ca²⁺ dissociation from cTnC. All these proteins influence the ability of cTnC to sense and respond to the Ca²⁺ signal, allowing cardiac muscle to be finely or grossly tuned to perform mechanically on a beat-to-beat basis.

	ACKNOWLEDGEMENTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
We thank Dr. Lawrence Smillie for the generous gift of the human cTnC and cTnI plasmids, and Dr. James D. Potter and Dr. Aldrin V. Gomes for the generous gift of human cTnT3 protein. We thank Dr. Jack Rall for critical reading of the manuscript and financial support. We also thank Bin Liu, Laszlo Sarkozy, and Chad Warren for technical assistance.

This research was funded, in part, by National Institutes of Health grants HL073828 (D.R.S.), HL022231 (R.J.S.), HL073600 (S.B.T.), and AR020792 (Jack A. Rall), and awards from the American Heart Association (to J.P.D., C.N., S.B.T., and T.K. (0230038N));

Submitted on August 16, 2006; accepted for publication January 17, 2007.

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