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Originally published In Press as doi:10.1074/jbc.M314095200 on February 16, 2004

J. Biol. Chem., Vol. 279, Issue 17, 17348-17360, April 23, 2004
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Mutations of Hydrophobic Residues in the N-terminal Domain of Troponin C Affect Calcium Binding and Exchange with the Troponin C-Troponin I96–148 Complex and Muscle Force Production*

Jonathan P. Davis{ddagger}, Jack A Rall, Catalina Alionte, and Svetlana B. Tikunova

From the Department of Physiology and Cell Biology, The Ohio State University, Columbus, Ohio 43210

Received for publication, December 23, 2003 , and in revised form, February 4, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Interactions between troponin C and troponin I play a critical role in the regulation of skeletal muscle contraction and relaxation. We individually substituted 27 hydrophobic Phe, Ile, Leu, Val, and Met residues in the regulatory domain of the fluorescent troponin CF29W with polar Gln to examine the effects of these mutations on: (a) the calcium binding and dynamics of troponin CF29W complexed with the regulatory fragment of troponin I (troponin I96–148) and (b) the calcium sensitivity of force production. Troponin I96–148 was an accurate mimic of intact troponin I for measuring the calcium dynamics of the troponin CF29W-troponin I complexes. The calcium affinities of the troponin CF29W-troponin I96–148 complexes varied ~243-fold, whereas the calcium association and dissociation rates varied ~38- and ~33-fold, respectively. Interestingly, the effect of the mutations on the calcium sensitivity of force development could be better predicted from the calcium affinities of the troponin CF29W-troponin I96–148 complexes than from that of the isolated troponin CF29W mutants. Most of the mutations did not dramatically affect the affinity of calcium-saturated troponin CF29W for troponin I96–148. However, the Phe26 to Gln and Ile62 to Gln mutations led to >10-fold lower affinity of calcium-saturated troponin CF29W for troponin I96–148, causing a drastic reduction in force recovery, even though these troponin CF29W mutants still bound to the thin filaments. In conclusion, elucidating the determinants of calcium binding and exchange with troponin C in the presence of troponin I provides a deeper understanding of how troponin C controls signal transduction.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Troponin C (TnC)1 regulates striated muscle contraction and relaxation through the binding and release of Ca2+ (for review see Refs. 1–3). Skeletal muscle TnC (~18 kDa) consists of globular N- and C-terminal domains connected by a 31-residue {alpha}-helix (for review see Refs. 4 and 5). Both domains bind two Ca2+ ions through a pair of EF hand Ca2+-binding motifs. Each pair of EF hands interacts with one another through a short antiparallel {beta}-sheet connecting the two Ca2+-binding loops (Ref. 6 and references within). The EF hands are numbered I–IV, and the helices flanking the loops are designated A–H, with an additional N-terminal 14-residue {alpha}-helix (Fig. 1, N-helix), which is absent in the closely related EF hand Ca2+-binding protein calmodulin.



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  		FIG. 1.
Cartoon representation of the regulatory domain of TnCF29W. The cartoon depicts the amino acids in the regulatory domain of TnCF29W that form the two Ca2+-binding sites (I and II) and the various helices (N–D). The black amino acids represent the hydrophobic residues that were individually mutated to Gln, excluding Trp29.

 
Much is known about the cation binding properties of TnC in solution. Each EF hand system binds Ca2+ and Mg2+ competitively, with the two C-terminal EF hands possessing higher Ca2+ and Mg2+ affinities (6–8). In fact, the Ca2+-binding sites of the C-domain of TnC possess ~10-fold higher Ca2+ affinity with a greater than 100-fold slower Ca2+ dissociation rate compared with those in the N-domain (6, 9). In part because of its high Ca2+ and Mg2+ affinities and slow Ca2+ exchange rates (as compared with the kinetics of muscle contraction and relaxation), the C-domain is thought to play a structural role in muscle function by anchoring TnC into the Tn complex. In contrast, the Ca2+ exchange rates of the N-domain of TnC are rapid enough to be involved in the dynamic Ca2+-dependent regulation of muscle mechanics (for review see Refs. 3 and 10).

Skeletal muscle contraction begins when cytoplasmic [Ca2+] rises and binds to the N-terminal EF hands of TnC. The entire N-domain of TnC subsequently undergoes a large tertiary conformational change, in which helices B and C move away as a unit from helices N, A, and D, exposing a buried hydrophobic pocket to the solvent (Ref. 5 and references within). The newly formed hydrophobic pocket is thought to allow the N-domain of TnC to interact with the C-terminal of TnI transferring the inhibitory domain of TnI away from actin (11). Concurrently or subsequently, tropomyosin changes its position on the actin filament and myosin then binds cyclically to actin causing muscle contraction (for review see Refs. 1–3). As cytoplasmic [Ca2+] lowers, the sequence of events above reverses (not necessarily in the same order), and the muscle relaxes. One of the steps that may influence the rate of muscle relaxation is Ca2+ dissociation from the N-domain of TnC.

The influence that TnC has on the kinetics of muscle relaxation is controversial and incompletely understood (Ref. 12 and references within). The actual rate that Ca2+ dissociates from TnC in muscle fibers has not been measured and thus must be inferred. Ca2+ dissociates from the regulatory domain of isolated TnC ~20–30 times faster than skeletal muscle relaxation and thus has been speculated to be too rapid to influence the rate of relaxation (for review see Ref. 3). However, the antiparallel binding of TnI to TnC increases the Ca2+ sensitivity of the N-domain of TnC ~10-fold and slows the Ca2+ dissociation rate ~30-fold, with little additional change upon the formation of the whole Tn complex (Refs. 8 and 13–16; for review see Ref. 10). Thus, the rate of Ca2+ dissociation from the Tn complex and not TnC alone may be the more meaningful rate when considering factors that control muscle relaxation kinetics. Consistent with this idea, exchanging a TnC mutant with an ~2-fold slower N-terminal Ca2+ dissociation rate into skeletal muscle fibers slowed the rate at which the fibers relaxed ~2-fold (12). However, in the same study exchange of an ~1.5-fold faster TnC mutant into muscle did not statistically increase the rate of relaxation. Clearly, a broader range of Ca2+ dissociation rates from TnC mutants is required to further probe the role of TnC in tuning the rate of striated muscle relaxation.

To better understand the regulation of muscle mechanics, it is important to elucidate the Ca2+-dependent interactions of TnC with TnI because TnC regulates muscle contraction as a part of the Tn complex and not in isolation. TnI residues 96–116 (TnI96–116) bind actin and are primarily responsible for the ability of TnI to inhibit the ATPase activity of actomyosin, which can be reversed upon TnC-Ca2+ binding to TnI96–116 (17–19). In conjunction with residues 96–116, residues 117–148 of TnI are required for the complete inhibitory activity and regulatory interactions with actin and TnC (20–23). Furthermore, the complete enhancement of the Ca2+ sensitivity and the slowing of the Ca2+ dissociation rate from the regulatory domain of TnC in the presence of intact TnI were mimicked by a peptide of TnI corresponding to residues 96–148 (TnI96–148) (8). Thus, the Ca2+-dependent binding of the regulatory domain of TnC to TnI96–148 may be a good model system to study the Ca2+-dependent interactions between TnI and TnC that regulate muscle mechanics.

Recently, we investigated the effect of hydrophobic residue substitutions on the Ca2+ binding properties of the regulatory domain of TnC with the Phe29 -> Trp mutation (TnCF29W). The global N-terminal Ca2+ affinities of the TnCF29W mutants varied 2340-fold, whereas the Ca2+ association and dissociation rates varied less than 70-fold and more than 45-fold, respectively (6). In the present study we have determined how these mutations affect the Ca2+ binding properties and dynamics of the TnCF29W-TnI96–148 complex and located hydrophobic residues essential for high affinity binding of TnI96–148. Furthermore, we have tested whether the TnCF29W-TnI96–148 complex is a better predictor than isolated TnCF29W for the Ca2+ binding properties of the Tn complex in muscle and the potential for a particular TnCF29W mutant to support force production.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Materials—Phenyl-Sepharose CL-4B and EGTA were purchased from Sigma. Quin-2 was purchased from Calbiochem (La Jolla, CA). All other chemicals were of analytical grade. The TnI96–148 peptide was synthesized and purified by the Alberta Peptide Institute (Edmonton, Canada).

Protein Mutagenesis and Purification—The construction and expression of intact chicken skeletal TnCF29W in pET3a has been described (24). Chicken skeletal fast TnI was prepared as described for the rabbit protein (25). The TnCF29W mutants were constructed from the TnCF29W plasmid by primer based site-directed mutagenesis using a Stratagene Quik-Change site-directed mutagenesis kit. The mutations were confirmed by DNA sequence analysis. The plasmids for TnCF29W and its mutants were transformed into Escherichia coli BL21(DE3)pLysS cells (Novagen) and purified as described previously (6). Aliquots of TnCF29W and I62QTnCF29W were labeled with the Cys-specific fluorescent probe 5-((((2-iodoacetyl)amino)ethyl)amino) naphthalene-1-sulfonic acid (IAEDANS) at position Cys101 for the myofibril studies. Each TnC was reacted with 3–5-fold molar excess of IAEDANS for ~6 h at room temperature with constant shaking in 50 mM Tris, 90 mM KCl, 1 mM EGTA, 6 M urea, pH 7.5. The labeling reaction was stopped by the addition of 2 mM DTT, and the labeled proteins were exhaustively dialyzed against 10 mM MOPS, 90 mM KCl, pH 7.0, at 4 °C to remove unreacted label.

Determination of Ca2+ Affinities—All steady-state fluorescence measurements were performed using a Perkin-Elmer LS5 Spectrofluorimeter at 15 °C. Trp fluorescence was excited at 275 nm and monitored at 345 nm as microliter amounts of CaCl2 were added to 1 ml of each TnCF29W mutant (0.3 µM) plus TnI96–148 (3 µM) in 200 mM MOPS (to prevent pH changes upon the addition of metal), 90 mM KCl, 2 mM EGTA, 1 mM DTT, pH 7.0, at 15 °C. The [Ca2+]free was calculated using the computer program EGCA02 developed by Robertson and Potter (26). The Ca2+ affinities are reported as dissociation constants (Kd). Each Kd represents the mean of 3–5 titrations fit with a logistic sigmoid function mathematically equivalent to the Hill equation, as previously described (6).

Determination of TnI96–148 Peptide Affinities—Trp fluorescence was monitored as described in the previous paragraph. Microliter amounts of TnI96–148 were added to 1 ml of each TnCF29W mutant (0.6 µM) in 200 mM MOPS, 90 mM KCl, 2 mM EGTA, 1 mM [Ca2+]free, 1mM DTT, pH 7.0, at 15 °C. Each peptide affinity, reported as a dissociation constant, represents the mean of three to five titrations fit to the root of a quadratic equation for binary complex formation as previously described (27).

Determination of Ca2+ Dissociation Rates—Ca2+ dissociation rates (koff) were measured using an Applied Photophysics Ltd. (Leatherhead, UK) model SX.18 MV stopped flow instrument with a dead time of 1.4 ms at 15 °C. The samples were excited using a 150-watt Xenon arc source. Ca2+ dissociation from the N-terminal regulatory domain of TnCF29W and its mutants when complexed with TnI96–148 were obtained utilizing Trp fluorescence excited at 275 nm with emission monitored through a UV transmitting black glass filter (UG1 from Oriel, Stratford, CT). koff was also measured using the fluorescent Ca2+ chelator Quin-2 (6, 8). Quin-2 was excited at 330 nm with its emission monitored through a 510-nm broad band pass interference filter (Oriel, Stratford, CT). The buffer used in all stopped flow experiments was 10 mM MOPS, 90 mM KCl, 1 mM DTT, pH 7.0.

Calculation of Ca2+ Association Rates—The Ca2+ association rates (kon) were calculated using the simple relationship kon = koff/Kd, where koff represents the concerted release of two Ca2+ ions, and Kd represents the binding event of two Ca2+ ions to the N-domain of TnC in the presence of TnI96–148, as previously described (8).

Muscle Fiber Experiments—Single fibers were isolated the day of use from bundles of rabbit psoas muscle that had been stored in a glycerinating solution at -20 °C no longer than 1 month. Solutions and the mechanical setup utilized for force measurements were as previously described (28). Briefly, a single fiber was soaked in relaxing solution containing 1% (v/v) Triton X-100 for 5 min to remove any residual sarcolemma and sarcoplasmic reticulum. The fiber was then tied down in troughs attached to a servo-controlled DC torque motor (Cambridge Technologies, Watertown, MA) and an isometric force transducer (model 403A, Cambridge Technologies) as previously described (29). Fiber sarcomere length, width, and depth were measured with a video camera (Sony model XC-ST70) and an image analysis system (Simple PCI; Compix Inc., Cranberry Township, PA). Resting sarcomere length was set between 2.50 and 2.60 µm. The fiber was then activated in a pCa 4.0 solution and rapidly slackened after isometric force reached plateau. The analogue output of the force transducer was digitized using a DaqBoard/2000 and Daqview software (Iotech Inc., Cleveland, OH). The total force was measured between the plateau and base-line levels. The same procedure was utilized to obtain the resting force level of the fiber in a pCa 9.0 solution. The active force generated by the fiber in the various pCa solutions was calculated as the total force minus the resting force. Three active force measurements were performed in pCa 4.0 with the final activation taken as the maximal force generated by the native fiber (i.e. prior to extraction of endogenous TnC), which led to an average force per cross-sectional area of 85 ± 5 kN/m2. The fiber was then soaked for 2 min in a TnC extraction solution containing 5 mM EDTA, 10 mM HEPES, and 0.5 mM trifluoperazine dihydrochloride at pH 7.0 (30). The fiber was then washed three times in pCa 9.0 solution to remove any residual trifluoperazine dihydrochloride. If the residual force in pCa 4.0 solution was >10% of the maximal force, the extraction process was repeated. The fibers were then soaked for 2 min in a pCa 9.0 solution containing 16.7 µM recombinant TnCF29W or its mutant. All of the reconstituted fibers were then exposed to a series of pCa solutions varying from pCa 9.0 to 4.0, and the active force versus pCa was measured. Every fourth activation was performed at pCa 4.0, to which each adjacent and randomized pCa was normalized.

Myofibril Preparation and Experiments—Rabbit skeletal psoas myofibrils were prepared and stored as previously described (31). Endogenous TnC was extracted from a sample of the stock myofibrils by first washing the myofibrils three times in a myofibril TnC extraction solution (10 mM MOPS, 90 mM KCl, 5 mM EDTA, 2 mM DTT, 0.02% Tween 20, pH 8.0) to remove any residual glycerol. The myofibrils were then soaked in the TnC extraction solution for approximately 10 min at room temperature, pelleted, and resuspended in fresh TnC extraction solution an additional three times. The TnC extracted myofibrils were then washed three times in a pCa 9 solution after which the concentration of the myofibrils was determined (31). 0.1 mg/ml aliquots of the TnC extracted myofibrils in the pCa 9.0 solution were then exposed to 1 µM TnCF29W or I62QTnCF29W labeled with IAEDANS for approximately 5 min. The myofibrils were then diluted with a pCa 3.0 solution to bring the final pCa of the solution to 4.0 according to a mixing table. The reconstituted myofibrils were then plated on a glass slide with a coverslip and imaged as previously described (31, 32). Briefly, the images were collected using a Zeiss Axiovert TV (Thornwood, NJ) epifluorescence microscope equipped with a 100x oil immersion phase contrast lens and a Chroma filter set #11000UV (360 nm broad excitation, 400-nm-long pass dichroic and 420-nm-long pass emission). The digital images were obtained using a 12-bit intensity resolution CCD camera (Kodak KAF 1300 chip; Photometrics, Tucson, AZ) controlled by a Matrox board and IPLab Spectrum software (version 3.0, Signal Analytics, Vienna, VA) run by a Macintosh 840AV.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Effect of Ca2+ on the Fluorescence Spectra of TnCF29W and Mutants in the Presence of TnI96–148 at 15 °C—The influence of individual hydrophobic to Gln mutations in the regulatory domain of TnCF29W on Ca2+ binding and exchange was previously characterized (6). However, TnC does not function in muscle in isolation but as part of the Tn complex, primarily interacting with TnI. Thus, we determined the influence of 27 TnCF29W mutants in which all N-terminal hydrophobic residues were individually substituted with Gln (Fig. 1) on Ca2+ binding and exchange in the presence of the regulatory peptide of TnI, TnI96–148. The N-terminal domain of TnCF29W undergoes a large increase in Trp fluorescence upon forming the Ca2+-TnCF29W-TnI96–148 complex (8, 20). In the absence of Ca2+, the Trp fluorescence of TnCF29W upon the addition of TnI96–148 marginally decreased <1.1-fold at 345 nm (Fig. 2A, dashed line). Subsequent addition of 1 mM [Ca2+]free to TnCF29W plus TnI96–148 blue-shifted (~8 nm) and increased the Trp fluorescence ~2.2-fold at 335 nm (Fig. 2A, dotted line). Fluorescence spectra of the other TnCF29W mutants were similar to TnCF29W (data not shown) except for I37QTnCF29W. The addition of TnI96–148 to apo I37QTnCF29W increased the Trp fluorescence ~1.3-fold, which subsequently decreased ~1.3-fold upon the addition of 1 mM [Ca2+]free at 345 nm with a similar blue shift in the maximum fluorescence as observed with TnCF29W. The reason for this atypical behavior of I37QTnCF29W is currently unknown. Ile37 is part of the first Ca2+-binding loop located in the middle of the small {beta}-sheet connecting the two N-terminal EF hands. Thus, Ile37 may be critical for proper Ca2+ binding and coordination of subsequent structural changes.



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  		FIG. 2.
Effect of Ca2+ on the fluorescence spectra of TnCF29W and I37QTnCF29W in the presence of TnI96–148. Fluorescence emission spectra for TnCF29W (A) or I37QTnCF29W (B) are shown in the apo (solid lines), apo + TnI96–148 (dashed lines), and Ca2+ + TnI96–148 (dotted lines) states. The Trp fluorescence spectra were recorded with an excitation wavelength of 275 nm in 200 mM MOPS, 90 mM KCl, 2 mM EGTA, 1 mM DTT, pH 7.0, at 15 °C. The concentrations of the TnCF29W proteins, TnI96–148 peptide and [Ca2+]free were 0.3 µM, 3 µM, and 1 mM, respectively.

 
Measurement of Ca2+ Binding Affinities for the N-terminal Domains of TnCF29W and Mutants in the Presence of TnI96–148 at 15 °C—The Ca2+ binding affinities (Kd) for TnCF29W and each hydrophobic mutant were measured by following the Ca2+-induced changes in Trp fluorescence in the presence of TnI96–148. Examples of the Ca2+-dependent increases in N-terminal TnCF29W and mutant Trp fluorescence are shown in Fig. 3 for L49QTnCF29W (•), TnCF29W ({blacktriangleup}), M81QTnCF29W ({square}), I73QTnCF29W ({triangleup}), and F26QTnCF29W ({circ}). Table I summarizes the Ca2+ binding data for these and the remaining TnCF29W mutants. In the presence of TnI exhibited a half-maximal increase in its Trp fluorescence upon the addition of Ca2+ at 267 ± 3 nM. The Ca2+ affinities for the mutants ranged from 70 ± 1 nM for L49QTnCF29W to 17 ± 3 µM for F26QTnCF29W. Therefore, substitution of hydrophobic residues with polar Gln produced N-domain TnCF29W mutants that exhibited ~243-fold variation in their Ca2+ affinities in the presence of TnI96–148. The Hill coefficients for all but two of the TnCF29W-TnI96–148, mutant complexes (F26QTnC and I37QTnCF29W) were between 1.6 and 2.8 (see Table I), implying cooperative binding of Ca2+ and TnI96–148 to TnCF29W and its mutants.



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  		FIG. 3.
Ca2+ binding to TnCF29W and its mutants in the presence of TnI96–148. The Ca+-dependent increases in Trp fluorescence are shown for L49QTnCF29W (•), TnCF29W ({blacktriangleup}), M81QTnCF29W ({square}), I73QTnCF29W ({triangleup}), and F26QTnCF29W ({circ}) as a function of -Log[Ca2+]. Microliter amounts of Ca2+ were added to 1 ml of each protein (0.3 µM) plus TnI96–148 (3 µM) in the same buffer and temperature as described in the legend of Fig. 2. Trp fluorescence emission was monitored at 345 nm with excitation at 275 nm. 0% Trp fluorescence corresponds to the apo state fluorescence, whereas 100% Trp fluorescence corresponds to the highest fluorescent state in the presence of Ca2+ for each individual TnCF29W protein. Each data point represents the mean ± S.E. of three to five titrations fit with a logistic sigmoid equation.

 


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  		TABLE I
Summary of Ca2+ binding properties for TnCF29W and its mutants in the presence of TnI96–148

 
Similar to the binding of TnI to TnC, the binding of TnI96–148 to TnCF29W increases the Ca2+ sensitivity of the regulatory domain of TnCF29W ~12-fold (Table I and Refs. 8 and 13–15). On average, the Ca2+ sensitivity of the TnCF29W mutants increased ~11-fold (Table I). However, the Ca2+ sensitivities of F26QTnCF29W, V80QTnCF29W, and M81QTnCF29W increased >23-fold, whereas for F22QTnCF29W, L42QTnCF29W, V45QTnCF29W, M46QTnCF29W, L49QTnCF29W, I61QTnCF29W, F78QTnCF29W, and M82QTnCF29W, the Ca2+ sensitivities increased <5.5-fold when compared with the isolated TnCF29W mutant (Table I and Ref. 6). Thus, the hydrophobic to Gln mutations in TnCF29W not only affect the overall Ca2+ sensitivity of the TnCF29W-TnI96–148 complex but also modulate the effectiveness of TnI96–148 to increase the Ca2+ affinity of the regulatory domain of TnCF29W.

Measurement of TnI96–148 Binding Affinities for the Ca2+-saturated N-terminal Domains of TnCF29W and Mutants at 15 °C—The binding of TnI96–148 to Ca2+-saturated TnCF29W and its mutants caused on average a 26.6 ± 0.4% decrease in the Trp fluorescence (excluding I37QTnCF29W, which displayed a 23 ± 1% increase), which can be utilized to determine the peptide binding affinity (8, 20). Examples of the TnI96–148-dependent decrease in N-terminal TnCF29W and mutant Trp fluorescence are shown in Fig. 4 for L49QTnCF29W (•), TnCF29W ({blacktriangleup}), M81QTnCF29W ({square}), I73QTnCF29W ({triangleup}), and F26QTnCF29W ({circ}). Table I summarizes the TnI96–148 binding data for these and the remaining TnCF29W mutants. In the presence of a saturating concentration of Ca2+, TnCF29W bound to TnI96–148 with an affinity of 146 ± 19 nM, which is in agreement with a previously reported value (20). Hydrophobic interactions are important for the Ca2+-dependent binding of the N-domain of TnC to the C-domain of TnI. Thus, it seemed logical that substitution of hydrophobic residues with polar Gln in the N-domain of TnC would likely decrease its affinity for TnI96–148. However, Fig. 4 and Table I also demonstrate that the TnI96–148 affinities for the Ca2+-saturated mutants of TnCF29W were only marginally decreased (<2-fold with an average value of 230 ± 15 nM), except for F26QTnCF29W and I62QTnCF29W, which displayed ~10- and 14-fold lower affinities, respectively. Thus, all of the Ca2+-saturated N-terminal domains of the TnCF29W mutants bind TnI96–148, with the majority of them binding with similar affinity.



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  		FIG. 4.
TnI96–148 binding to Ca2+-saturated TnCF29W and its mutants. The TnI96–148-dependent decreases in Trp fluorescence are shown for Ca2+-saturated L49QTnCF29W (•), TnCF29W ({blacktriangleup}), M81QTnCF29W ({square}), I73QTnCF29W ({triangleup}), and F26QTnCF29W ({circ}) as a function of TnI96–148 concentration. Microliter amounts of TnI96–148 were added to each TnC (0.6 µM) in the same buffer and temperature as in Fig. 2 in the presence of 1 mM [Ca2+]free (or 10 mM [Ca2+]free in the case of F26QTnCF29W). 100% Trp fluorescence corresponds to the Ca2+-saturated state, whereas 0% represents the Ca2+-TnI96–148-saturated state for each individual TnCF29W protein. Each data point represents the mean ± S.E. of three to five titrations fit to the root of a quadratic equation for binary complex formation.

 
Measurement of Ca2+ Dissociation Rates from the N-terminal Domains of TnCF29W and Mutants in the Presence of TnI96–148 at 15 °C Using Trp and Quin-2 Fluorescence—Utilizing fluorescence stopped flow technology, the rates of Ca2+ dissociation from the regulatory domain of TnCF29W and its mutants in the presence of TnI96–148 were determined. Fig. 5A shows examples of the EGTA-induced decreases in Trp fluorescence for M81QTnCF29W (5 s-1), L49QTnCF29W (8 s-1), TnCF29W (11 s-1), I73QTnCF29W (29 s-1), and F26QTnCF29W (169 s-1) complexed with TnI96–148. The rates of Ca2+ dissociation for the remaining mutant complexes fell between that of M81QTnCF29W and that of F26QTnCF29W (Table I). Therefore, substitution of hydrophobic residues with polar Gln in the regulatory domain of TnCF29W increased (~16-fold) and decreased (~2-fold) the Ca2+ dissociation rate from the TnCF29W-TnI96–148 complex, creating an ~33-fold variation.



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  		FIG. 5.
Rates of Ca2+ dissociation from TnCF29W and its mutants in the presence of TnI96–148 or intact TnI. A shows the time course of decrease in Trp fluorescence as Ca2+ was removed by EGTA from the regulatory Ca2+-binding sites of M81QTnCF29W, L49QTnCF29W, TnCF29W, I73QTnCF29W, and F26QTn-CF29W in the presence of TnI96–148. Each TnCF29W protein (0.6 µM) plus TnI96–148 (6 µM) in 10 mM MOPS, 90 mM KCl, 1 mM DTT, pH 7.0, plus 100 µM Ca2+ was rapidly mixed with an equal volume of the same buffer plus 10 mM EGTA at 15 °C. Trp fluorescence was monitored through a UV-transmitting black glass filter (UG1 from Oriel) with excitation at 275 nm. B shows the time course of increase in Quin-2 fluorescence as Ca2+ was removed by Quin-2 from the regulatory Ca2+-binding sites of M81QTnCF29W, L49QTnCF29W, TnCF29W, I73QTnCF29W, and F26QTnCF29W in the presence of TnI96–148. Each TnCF29W protein (3 µM) plus TnI96–148 (30 µM)in10mM MOPS, 90 mM KCl, 1 mM DTT, pH 7.0, plus 30 µM Ca2+ was rapidly mixed with an equal volume of the same buffer plus 150 µM Quin-2 at 15 °C. Quin-2 fluorescence was monitored through a 510-nm broad band pass interference filter with excitation at 330 nm. C shows the time course of increase in Quin-2 fluorescence as Ca2+ was removed by Quin-2 from the regulatory Ca2+-binding sites of M81QTnCF29W, L49QTnCF29W, TnCF29W, I73QTnCF29W, and F26QTnCF29W in the presence of intact TnI. Each TnCF29W protein (3 µM) plus intact TnI (30 µM) in 10 mM MOPS, 90 mM KCl, 1 mM DTT, pH 7.0, plus 30 µM Ca2+ was rapidly mixed with an equal volume of the same buffer plus 150 µM Quin-2 at 15 °C. All of the traces have been staggered and normalized for clarity. Each trace is an average of at least 15 traces fit with a single exponential equation (variance < 2 x 10-4). The kinetic traces were triggered at time 0 with the first ~2 ms of premixing time shown (the apparent lag phase). The traces were fit after the mixing time was complete.

 
To verify that the time course of the EGTA-induced Trp fluorescence decreases for the TnCF29W-TnI96–148 complexes followed Ca2+ dissociation and not a slower structural change, Ca2+ dissociation was also measured using the fluorescent Ca2+ chelator Quin-2. Whereas the fluorescence of Trp was selective for the events of N-terminal Ca2+ dissociation, Quin-2 fluorescence reported Ca2+ dissociation from both the N- and C-domains of TnCF29W and its mutants complexed with TnI96–148. However, the Ca2+ dissociation rates from the N-terminal domain of TnCF29W and its mutants were easily distinguished from the rates of Ca2+ dissociation from the C-terminal domain (on average 0.159 ± 0.007 s-1) because the latter rates were >30-fold slower in the presence of TnI96–148 or intact TnI. Fig. 5B demonstrates that for all of the mutants, the Ca2+ dissociation rate reported by Trp was in excellent agreement with the N-terminal rate determined by Quin-2. Therefore, the fluorescent Trp signal accurately reports Ca2+ binding and dissociation from the TnCF29W-TnI96–148 mutant complexes.

To verify that TnI96–148 is a satisfactory model system for the regulatory domain binding of TnC to TnI, stopped flow studies were also conducted with intact chicken skeletal TnI. Fig. 5C shows the time course of the increases in Quin-2 fluorescence as Ca2+ was dissociated from the N-terminal domains of M81QTnCF29W (4 s-1), L49QTnCF29W (5 s-1), TnCF29W (9 s-1), I73QTnCF29W (22 s-1), and F26QTnCF29W (135 s-1) complexed with intact TnI. The Ca2+ dissociation rates measured from the regulatory domain of TnCF29W and its mutants in the presence of TnI96–148 were similar to that measured in the presence of intact TnI. Therefore, the TnCF29W-TnI96–148 complex is a good model system to study the regulatory mechanisms of the Ca2+-dependent binding of TnC to TnI.

Calculation of Ca2+ Association Rates—The Ca2+ association rates to TnCF29W and its mutants in the presence of TnI96–148 were calculated using the Ca2+ Kd and koff values determined by Trp (kon = koff/Kd; Table I). The calculated kon for TncF29W in the presence of TnI96–148 was ~4.5 x 107 M-1 s-1, which was ~2.4-fold slower than the kon calculated or measured in the absence of the peptide (6, 8). In the presence of TnI96–148, kon varied ~38-fold between the TnCF29W mutants with L49QTnCF29W (~1.1 x 108 M-1 s-1) exhibiting the fastest kon and L42QTnCF29W (~3 x 106 M-1 s-1) exhibiting the slowest kon. Clearly, some of the hydrophobic mutations in the regulatory domain of TnCF29W significantly alter the Ca2+ association rate in the presence of TnI96–148.

Ca2+ Binding to TnCF29W in the Presence of TnI96–148 as a Predictor for the Ca2+ Dependence of Force Development in Skeletal Muscle—Fig. 6A shows that V45QTnCF29W and M46QTnCF29W possess ~19- and 3.6-fold higher Ca2+ affinity than TnCF29W in the absence of TnI96–148, respectively (Table II and Ref. 6). On the other hand, M81QTnCF29W and F78QTnCF29W display ~5.9- and 8.4-fold lower Ca2+ affinity than TnCF29W in the absence of TnI96–148, respectively (Table II and Ref. 6). However, Fig. 6B shows that only the Ca2+ sensitivities of V45QTnCF29W and F78QTnCF29W in the presence of TnI96–148 remain higher (~3-fold) and lower (~19-fold) than TnCF29W, respectively (see also Table II). Thus, the qualitative and quantitative changes in N-terminal Ca2+ sensitivities for several of the TnCF29W mutants compared with TnCF29W in the presence or absence of TnI96–148 were not the same.



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  		FIG. 6.
Comparison of the Ca2+ binding properties of TnCF29W and mutants in the absence and presence of TnI96–148 with their Ca2+ dependence of skeletal muscle force generation. A shows the Ca2+-dependent increases in Trp fluorescence for V45QTnCF29W (•), M46QTnCF29W ({blacksquare}), TnCF29W ({blacktriangleup}), M81QTnCF29W ({square}), and F78QTnCF29W ({circ}) as a function of -Log[Ca2+]. The data have been adapted from Tikunova et al. (6). Because these data were collected under identical experimental conditions as were used here, we have for comparative purposes reproduced these data in this figure and Table II. Microliter amounts of Ca2+ were added to 1 ml of each protein (0.3 µM) in the same buffer and temperature as described in the legend to Fig. 2. Trp fluorescence was monitored as described in the legend to Fig. 3. The data sets were normalized individually for each mutant. B shows the Ca2+ dependent increases in Trp fluorescence in the presence of TnI96–148 for V45QTnCF29W (•), M46QTnCF29W ({blacksquare}), TnCF29W ({blacktriangleup}), M81QTnCF29W ({square}), and F78QTnCF29W ({circ}) as a function of -Log[Ca2+]. The experimental details are identical to those described in the legend to Fig. 3. C shows the Ca2+ dependence of isometric force generation in single skinned psoas fibers reconstituted with V45QTnCF29W (•), M46QTnCF29W ({blacksquare}), TnCF29W ({blacktriangleup}), M81QTnCF29W ({square}), and F78QTnCF29W ({circ}) as a function of -Log[Ca2+]. The experimental conditions are described under "Experimental Procedures." Each data point represents the mean ± S.E. from at least three separate fibers individually normalized and fit with a logistic sigmoid equation mathematically equivalent to the Hill equation. their Ca +

 


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  		TABLE II
Comparison of the Ca2+ binding properties of TnCF29W and mutants in the absence and presence of TnI96–148 with their Ca2+ dependence of skeletal muscle force generation

 
To test which TnCF29W system (with or without TnI96–148) better represents the Ca2+ sensitivity of force production in muscle, the endogenous TnC in psoas muscle fibers was extracted and then replaced with TnCF29W or its mutants, and force versus pCa was measured. After TnC extraction, the average force generated by the single skinned muscle fibers was 2.3 ± 0.5% of the maximal force (data not shown). Subsequent reconstitution of the muscle fibers with V45QTnCF29W, M46QTnCF29W, TnCF29W, M81QTnCF29W, or F78QTnCF29W recovered 82 ± 5, 73 ± 4, 90 ± 3, 65 ± 8, and 80 ± 2% of the maximal force at pCa 4, respectively. Fig. 6C demonstrates that the Ca2+ dependence of force generation with TnCF29W or its mutants followed qualitatively more closely to the Ca2+ sensitivities of the mutant TnCF29W-TnI96–148 complexes and not to that of the isolated TnCF29W proteins (see also Table II). Thus, the Ca2+-dependent behavior of the TnCF29W-TnI96–148 complex is a better indicator for how the TnC mutant will control the Ca2+ dependence of force generation when incorporated into muscle fibers.

Another effect observed in the reconstituted muscle, which could not be predicted by the Ca2+ binding properties of the isolated TnC, was the maximal amount of force recovered by a particular mutant. Fig. 7A shows the Ca2+-dependent increases in force recovered by TnCF29W ({blacktriangleup}), L42QTnCF29W (*), I73QTnCF29W ({triangleup}), and I62QTnCF29W ({blacksquare}). At pCa 4.0, TnCF29W, L42QTnCF29W, I73QTnCF29W, and I62QTnCF29W recovered 90 ± 3, 79 ± 3, 45 ± 6, and 12 ± 4% of the force generated by the endogenous TnC prior to extraction, respectively. Similar to I62QTnCF29W, I37QTnCF29W and F26QTnCF29W also recovered force poorly at 15 ± 1 and 13 ± 1% of the maximal amount of force generated by the fiber prior to endogenous TnC extraction, respectively (data not shown). Thus, the amount of maximal force sustained by the TnCF29W mutants was variable, with three of the mutants only marginally allowing any force production. As will be discussed, the binding of TnI96–148 to the Ca2+-saturated TnCF29W mutants may offer clues as to why some of the mutants support little force.



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  		FIG. 7.
Comparison of TnCF29W and mutants with varied maximal force recoveries in TnC reconstituted muscle fibers and myofibrils. A shows the Ca2+ dependence of isometric force generation in single skinned psoas fibers reconstituted with TnCF29W ({blacktriangleup}), L42QTnCF29W (*), I73QTnCF29W ({triangleup}), and I62QTnCF29W ({blacksquare}) as a function of -Log[Ca2+]. The experimental conditions are described under "Experimental Procedures." Each data point represents the mean ± S.E. from at least three separate fibers fit with a logistic sigmoid equation mathematically equivalent to the Hill equation. Information regarding the parameters of the fit for TnCF29W can be found in Table II. L42QTnCF29W displayed half-maximal isometric force at 1.3 ± 0.2 µM Ca2+ with a Hill coefficient of 2.1 ± 0.2. I73QTnCF29W displayed half-maximal isometric force at 3.2 ± 0.4 µM Ca2+ with a Hill coefficient of 1.0 ± 0.1. I62QTnCF29W displayed half-maximal isometric force at 1.5 ± 0.1 µM Ca2+ with a Hill coefficient of 1.2 ± 0.1. B, the open squares show the time course decay of maximal isometric force as I62QTnCF29W displaces TnCF29W from single skinned psoas fibers (t1/2 = 3.3 ± 0.4 min). Time 0 represents the maximal isometric force generated in the pCa 4.0 solution for TnCF29W reconstituted muscle fibers. Subsequently the fibers were soaked in a resting solution containing 16.7 µM I62QTnCF29W and periodically contracted in a pCa 4.0 solution every 3 min for the first 12 min then every 5 min thereafter. The closed squares show the time course increase of maximal isometric force as TnC displaces I62QTnCF29W from single skinned psoas fibers (t1/2 = 2.2 ± 0.1 min). Time 0 represents the maximal isometric force generated in the pCa 4.0 solution for I62QTnCF29W reconstituted muscle fibers. Subsequently the fibers were soaked in a resting solution containing 16.7 µM TnC and periodically contracted in a pCa 4.0 solution every 3 min for the first 12 min then every 5 min thereafter. Each data point represents the mean ± S.E. from three separate fibers fit with a single exponential equation. C shows the phase contrast (top panels) and IAEDANS fluorescence (bottom panels) images obtained from TnC extracted rabbit psoas myofibrils reconstituted with TnCF29W-IAEDANS (left panels) or I62QTnCF29W-IAEDANS (right panels). The vertical line designates the location of the Z lines, and the horizontal lines designate the locations of the A bands (actin-myosin filament overlap).

 
To test whether I62QTnCF29W was actually binding to the thin filaments in the TnC extracted muscle fibers, additional TnC exchange experiments were performed on the muscle. Fig. 7B ({square}) at time 0 shows the maximal force recovered by TnCF29W at pCa 4.0 in a reconstituted muscle fiber. The fiber was then transferred to a relaxing solution containing 16.7 µM I62QTnCF29W, and the force generated at pCa 4.0 was measured at several time intervals. The amount of force production decreased with time, eventually reaching a value similar to that generated by fibers solely reconstituted with I62QTnCF29W. The data indicate that I62QTnCF29W was able to bind to the thin filaments and competitively displace TnCF29W from the Tn complex. Furthermore, when a muscle fiber was initially reconstituted with I62QTnCF29W (Fig. 7B, time 0, {blacksquare}) and then competitively displaced with 16.7 µM TnC in relaxing solution, maximal force increased with a time course similar to that at which I62QTnCF29W inhibited the force generated with TnCF29W. When the TnC-binding sites in the fiber were vacant (i.e. after endogenous TnC extraction), the addition of TnC at the concentration used for the competitive binding studies caused force to be maximal within 2 min (data not shown). Results similar to that of I62QTnCF29W were obtained in the displacement studies when I37QTnCF29W or F26QTnCF29W were tested (data not shown). Thus, the data supports the hypothesis that the TnCF29W mutants that minimally support force (<15%) bind to the thin filament and form the Tn complex.

To directly visualize whether I62QTnCF29W was able to incorporate into the TnC-depleted muscle fiber, both TnCF29W and I62QTnCF29W were labeled with the extrinsic fluorescent probe IAEDANS and reconstituted into psoas myofibrils. Fig. 7C shows representative phase contrast images of TnCF29W-IAEDANS (top left panel) and I62QTnCF29W-IAEDANS (top right panel) reconstituted myofibrils. As can be seen from the fluorescent images (Fig. 7C, bottom left panel for TnCF29W-IAEDANS and bottom right panel for I62QTnCF29W-IAEDANS), both IAEDANS-labeled TnC proteins incorporate into the myofibril at the myosin-actin filament overlap and nonoverlap space. Thus, as predicted from the physiological competition experiments, I62QTnCF29W binds to the thin filament at a similar location, as does TnCF29W, and forms the Tn complex, albeit in an inactive state.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The goal of the present study was to examine the effect of the hydrophobic mutations on the Ca2+ binding properties of the TnCF29W-TnI96–148 complex and on the affinity of TnCF29W for TnI96–148. Furthermore, we wanted to examine whether the effect of hydrophobic mutations on the Ca2+ sensitivity of force development could be better predicted by the Ca2+ and TnI96–148 binding properties of the TnCF29W mutants than by that of the isolated TnCF29W. Because the regulatory domain of chicken TnC is spectroscopically silent, the Phe29 -> Trp mutation was utilized to follow the structural changes in the N-domain of TnC induced by changes in Ca2+ concentration (6, 8, 9, 12, 24, 33–35). In our previous study, all 27 Phe, Ile, Leu, Val, and Met residues were individually mutated to polar Gln to examine the role of hydrophobic residues in Ca2+ binding and exchange with the regulatory domain of intact TnCF29W in isolation (6). The hydrophobic TnCF29W mutants exhibited ~2340-fold variation in their Ca2+ binding affinities. Indicative of the Ca2+ affinity changes, the hydrophobic TnCF29W mutants also exhibited less than 70-fold and more than 45-fold variation in their Ca2+ association rates and dissociation rates, respectively (6). The data indicated that the local side chain interactions of the hydrophobic residues within the tertiary structures of the apo and Ca2+-bound regulatory domain of TnCF29W played an important role in dictating the Ca2+ binding properties of the protein.

The Ca2+ affinities of the mutant TnCF29W-TnI96–148 complexes varied ~243-fold. However, the variation in the Ca2+ sensitivity of the mutants in the absence of TnI96–148 was an order of magnitude larger (6). It would appear that Ca2+ binding is optimized when the regulatory domain of TnC is in the open state (helices B and C swing away from helices N, A, and D) (6). The binding of TnI or C-terminal peptides of TnI to the regulatory domain of TnC help to lock TnC into the open state and thus enhance the Ca2+ binding affinity of TnC ~10–12-fold (8, 13–15). The high Ca2+ affinity mutants of TnCF29W (F22QTnCF29W, V45QTnCF29W, M46QTnCF29W, L49QTnCF29W, and M82QTnCF29W) may mimic TnI binding to TnC by shifting the equilibrium of the regulatory domain of TnC into the open state and away from the closed state (6). Thus, binding of TnI96–148 to the high Ca2+ affinity mutants of TnCF29W increases their Ca2+ sensitivities less than 5-fold as compared with an ~12-fold increase in Ca2+ sensitivity for TnCF29W. The reduced Ca2+ sensitivity enhancement to the high affinity Ca2+-binding mutants by TnI96–148 is the primary reason for the decreased variation in the Ca2+ binding affinities of the TnCF29W-TnI96–148 mutant complexes.

On the other hand, hydrophobic mutations to polar Gln in the regulatory domain of TnCF29W that may impede the formation of the open state (Phe26, Ile37, Leu42, Ile62, Val65, Ile73, Phe78, Leu79, Val80, or Met81) led to large (more than ~2-fold) decreases in the Ca2+ affinity of the isolated proteins (6). The Ca2+ affinities of these TnCF29W-TnI96–148 complexes were dramatically lower (more than 2-fold) than that of the TnCF29W-TnI96–148 complex (excluding M81QTnCF29W). Thus, in all cases but one, TnI96–148 was not able to correct for the decrease in Ca2+ binding affinity caused by the hydrophobic mutation. Interestingly, analysis of the NMR structure of the Ca2+-TnC-TnI115–131 complex (36) or a modeled structure of Ca2+-TnC-TnI (37) indicates that none of these residues interacts with TnI96–148. Thus, a direct loss in TnI96–148 contact cannot be the explanation for the low Ca2+ affinity of these mutants in the TnCF29W-TnI96–148 complex. Most of these hydrophobic residues (excluding Leu79, Val80, and Met81) are located in the {beta}-sheet connecting the two N-terminal EF hands (Ile37 and Ile73) or form a network of hydrophobic interactions among themselves and with the {beta}-sheet hydrophobic side chains in both the closed and open state of TnC (Ref. 6 and Fig. 8). Thus, maintenance of this {beta}-sheet and {beta}-sheet interacting hydrophobic core within the regulatory domain of TnCF29W is critical for high affinity Ca2+ binding to TnC in both the absence and the presence of TnI96–148.



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  		FIG. 8.
Hydrophobic {beta}-sheet and {beta}-sheet interacting residues in the Ca2+ bound state of TnC. Helices are shown as ribbons. The hydrophobic {beta}-sheet (Ile37 and Ile73) and {beta}-sheet interacting residues (Phe26, Leu42, Val65, Ile62, and Phe78) are labeled and shown in a stick format. The Protein Data Bank file used in this figure was generated by NMR (1TNX [PDB] ) (43) and was rendered using Rasmol (44).

 
Surprisingly, at saturating [Ca2+], the TnCF29W mutants bound to TnI96–148 with a similar affinity as TnCF29W, with exception of F26QTnCF29W and I62QTnCF29W, which exhibited ~10- and 14-fold lower affinity, respectively. One possible explanation for the consistency of TnCF29W mutant binding to TnI96–148 is that numerous interaction sites between TnC and TnI96–148 compensate for the loss of one potential hydrophobic side chain interaction. Consistent with this idea, analysis of the NMR structure of the Ca2+-TnC-TnI115–131 complex (36) or a modeled structure of TnC-TnI (37) indicates that there are nine different hydrophobic residue side chains within the N-domain of TnC that come within 4 Å of six different hydrophobic side chains within TnI115–131. All of the high Ca2+ affinity mutant hydrophobic residue side chains (Phe22, Val45, Met46, Leu49, and Met82) come in close contact to TnI115–131 and modestly decrease TnI96–148 binding ~1.7–3-fold. However, as mentioned above, neither Phe26 nor Ile62 come in close contact with TnI but interact with the Ca2+-binding loop {beta}-sheet residues Ile37 and Ile62 (Fig. 8). Thus, Phe26 and Ile62 may help maintain the open state in such a way as to allow high affinity TnI96–148 binding to the regulatory domain of TnCF29W. Consistent with this idea, inhibiting the Ca2+-dependent opening of the regulatory domain of TnC by the introduction of a disulfide bond between the NAD and BC units decreased the affinity of TnI binding ~15-fold (38).

The ~12-fold increase in TnCF29W Ca2+ affinity upon binding of TnI or TnI96–148 is primarily reflected by an ~30-fold slower rate of Ca2+ dissociation from the TnCF29W-TnI96–148 complex (8). Hydrophobic residue substitutions to polar Gln in the regulatory domain of TnCF29W varied the Ca2+ dissociation rates from the TnCF29W-TnI96–148 complex ~33-fold. This broad range in Ca2+ dissociation rates is primarily reflected by the ability of some of the hydrophobic mutations to speed the rate of Ca2+ dissociation, up to ~15-fold compared with theTnCF29W-TnI96–148 complex. Consistent with the inability of TnI96–148 to enhance the Ca2+ sensitivity of the TnCF29W mutants with increased Ca2+ affinity, the Ca2+ dissociation rate from the mutant TnCF29W-TnI96–148 complexes could only be slowed ~2-fold.

The TnCF29W-TnI96–148 complex may be a good model system to study how different Tn complexes respond to changes in Ca2+ concentration in muscle. Interestingly, the rate of Ca2+ dissociation from the TnCF29W-TnI96–148 complex, and not isolated TnCF29W, is similar to the rate of fast twitch skeletal muscle relaxation (Fig. 9). Previous experiments with TnC mutants in which the Ca2+ sensitivity of the regulatory domain in solution was increased or decreased demonstrated similar qualitative shifts in the force-pCa relationship upon reconstitution in skeletal muscle fibers (8, 12, 34, 35, 39, 40). This may be the case only in those circumstances when the Ca2+ sensitivity of the isolated TnC and the TnC-TnI complex are shifted in a similar direction. To test this hypothesis we reconstituted skinned rabbit psoas fibers with TnCF29W mutants that displayed the same qualitative changes in Ca2+ sensitivities in the absence or presence of TnI96–148 (V45QTnCF29W and F78QTnCF29W) and two that did not (M46QTnCF29W and M81QTnCF29W). Comparison of the effects of these mutations on the force-pCa relationship suggests that the TnCF29W-TnI96–148 complex is a better predictor than isolated TnCF29W for how changes in Ca2+ binding to TnC modulate the Ca2+ sensitivity of force production. For instance, even though both Val45 -> Gln and Met46 -> Gln mutations increase the Ca2+ affinity of isolated TnCF29W, only the Val45 -> Gln mutation increased the Ca2+ sensitivity of force development. These results are consistent with the fact that only Val45 -> Gln, but not Met46 -> Gln increases the Ca2+ affinity of the TnCF29W-TnI96–148 complex.



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  		FIG. 9.
Comparison of the Ca2+ dissociation rates from the regulatory domain of TnCF29W in the absence and presence of TnI96–148 with the relaxation rate of a skinned rabbit psoas muscle fiber at 15 °C. The data in this figure has been adapted and reproduced from the work of Davis et al. (8) and Luo et al. (12) for comparative purposes. The experimental procedure for the EGTA induced decreases in Trp fluorescence for TnCF29W and the TnCF29W-TnI96–148 complex in a stopped flow apparatus are described by Davis et al. (8). The experimental conditions for the laser-induced flash photolysis of diazo-2 induced relaxation of single skinned rabbit Psoas muscle fibers are described by Luo et al. (12).

 
Furthermore, the Ca2+ sensitivity of force development generated with F78QTnCF29W was dramatically lower than that generated with M81QTnCF29W or TnCF29W, even though both Phe78 -> Gln and Met81 -> Gln mutations in isolated TnCF29W decreased the Ca2+ sensitivity of the regulatory domain to a similar extent (6). These results are consistent with the fact that the Phe78 -> Gln but not Met81 -> Gln mutation leads to a dramatic decrease in Ca2+ affinity of the TnCF29W-TnI96–148 complex. However, the Phe78 -> Gln mutation appears to have a larger effect on the Ca2+ affinity of the TnCF29W-TnI96–148 complex than on the Ca2+ sensitivity of force development. Thus, the Ca2+ binding properties of the TnC-TnI complex are not the only determinants of Ca2+ sensitivity of force development. There is evidence that skeletal troponin T, tropomyosin, and actomyosin can modulate the Ca2+ sensitivity of muscle mechanics either directly through TnC or through mechanisms yet to be explained (for review see Ref. 3).

A striking result observed with I62QTnCF29W was the dramatic reduction of force production generated by muscle fibers reconstituted with this mutant, even though the data shows it is able to bind to the thin filament. The near lack of force production cannot be explained by the low Ca2+ sensitivity of the I62QTnCF29W-TnI96–148 complex because the F78QTnCF29W-TnI96–148 complex has ~ 2-fold lower Ca2+ sensitivity but is able to produce ~80% of maximal force. However, Ca2+-saturated I62QTnCF29W has an ~14-fold decreased affinity for TnI96–148 as compared with TnCF29W. The large decrease in TnI96–148 binding affinity for I62QTnCF29W is the likely reason why this mutant is unable to support force. It appears that Ca2+-I62QTnCF29W in the fiber might not effectively compete with actin binding to the regulatory domain of TnI, thus keeping the muscle fiber in a state of inactivation. Consistent with this interpretation, F26QTnCF29W, which had an ~10-fold lower affinity for TnI96–148, also produced only ~13% of the maximal force upon reconstitution in the muscle fibers (data not shown). The exact opposite effect occurred when the regulatory regions of TnC and TnI were cross-linked, causing a regulated thin filament system to be permanently activated even in the absence of Ca2+ (41). However, Ca2+-saturated I37QTnCF29W (a {beta}-sheet mutant) bound TnI96–148 with only an ~2-fold lower affinity than TnCF29W but still only produced ~15% maximal force upon reconstitution in the muscle fibers (data not shown). Furthermore, Ca2+-saturated I73QTnCF29W, another {beta}-sheet mutant, bound TnI96–148 with an affinity nearly identical to that of TnCF29W but only produced ~45% maximal force. Again, this points out that additional events besides Ca2+ binding and subsequent TnI binding are involved in the signal pathway of force production. Consistent with this idea, a mutant TnC with a decreased skeletal troponin T affinity (but similar affinity for TnI) has been implicated in a loss of reconstituted thin filament ATPase activity (27). However, another mutant TnC with apparently normal Ca2+, TnI, and skeletal troponin T binding also displayed a diminished reconstituted thin filament ATPase activity through an unidentified mechanism apparently important for the Ca2+-dependent regulation of signal transduction (42).

In summary, we utilized TnCF29W to study Ca2+ binding and exchange with a series of hydrophobic N-domain TnC mutants in the presence of TnI96–148 and intact TnI. The TnCF29W-TnI96–148 mutant complexes exhibited ~243-fold variation in their Ca2+ binding affinities, ~38-fold variation in their Ca2+ association rates, and ~33-fold variation in their Ca2+ dissociation rates. The regulatory peptide of TnI, TnI96–148, was an accurate mimic of intact TnI for measuring Ca2+ dissociation rates from the TnC-TnI complexes. Furthermore, the effect of hydrophobic mutations on the Ca2+ sensitivity of force development could be better predicted from the Ca2+ affinities of the TnCF29W-TnI96–148 mutant complexes than from that of the isolated TnCF29W mutants. Interestingly, TnCF29W mutants with >10-fold lower TnI96–148 affinities in the presence of saturating Ca2+, compared with that of TnCF29W, were able to bind to the thin filaments but led to dramatic reduction of force recovery in reconstituted muscle fibers. Thus, not just Ca2+ binding to TnC but the changes in the interactions with other regulatory proteins are critical in the pathway of signal transduction of force development. In conclusion, elucidating the determinants of Ca2+ binding and exchange with TnC in the presence of its target protein TnI may provide a deeper understanding of how TnC and other closely related EF hand proteins respond to Ca2+ and control signal transduction.


   FOOTNOTES
 
* This work was supported in part by National Institutes of Health Grants AR20792 (to J. A. R.) and HL073600 (to S. B. T.) and by an award from the American Heart Association, Ohio Valley Affiliate (to J. P. D.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

{ddagger} To whom correspondence should be addressed: 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.

1 The abbreviations used are: TnC, intact chicken skeletal troponin C; Tn, troponin; TnCF29W, intact TnC mutant with the Phe29 -> Trp mutation; TnI, chicken skeletal troponin I; TnI96–148, chicken skeletal troponin I peptide corresponding to residues 96–148; TnI96–116, skeletal troponin I peptide corresponding to residues 96–116; TnI115–131, skeletal troponin I peptide corresponding to residues 115–131; IAEDANS, 5-((((2-iodoacetyl)amino)ethyl)amino) naphthalene-1-sulfonic acid; DTT, dithiothreitol; MOPS, 3-(N-morpholino)propanesulfonic acid; Quin-2, 2-((bis(carboxymethyl)amino)-5-methylphenoxy)-methyl)-6-methoxy-8-(bis(carboxymethyl)amino)-quionline; N-domain, N-terminal domain; C-domain, C-terminal domain. Back


   ACKNOWLEDGMENTS
 
We thank Dr. Lawrence Smillie for the generous gifts of the chicken fast twitch skeletal muscle TnCF29W plasmid and TnI protein and for help in obtaining the TnI96–148 peptide, Dr. Peter Reiser for the expert advice and training pertaining to the physiological muscle fiber experiments as well as use of his equipment, Dr. Darl Swartz for the expert advice and training pertaining to the myofibril imaging experiments as well as use of his equipment, and Zhenyun Zyang for help and training in preparing the myofibril samples.



   REFERENCES