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J. Biol. Chem., Vol. 277, Issue 51, 49716-49726, December 20, 2002

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Engineering Competitive Magnesium Binding into the First EF-hand of Skeletal Troponin C^*

Jonathan P. Davis, Jack A. Rall, Peter J. Reiser^¶, Lawrence B. Smillie, and Svetlana B. Tikunova^**

From the Departments of  Physiology and Cell Biology, ^** Molecular and Cellular Biochemistry, and ^¶ Oral Biology, The Ohio State University, Columbus, Ohio 43210 and the  Department of Biochemistry, the University of Alberta, Edmonton, Alberta T6G 2H7, Canada

Received for publication, August 19, 2002, and in revised form, October 17, 2002

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

The goal of this study was to examine the mechanism of magnesium binding to the regulatory domain of skeletal troponin C (TnC). The fluorescence of Trp²⁹, immediately preceding the first calcium-binding loop in TnC^F29W, was unchanged by addition of magnesium, but increased upon calcium binding with an affinity of 3.3 µM. However, the calcium-dependent increase in TnC^F29W fluorescence could be reversed by addition of magnesium, with a calculated competitive magnesium affinity of 2.2 mM. When a Z acid pair was introduced into the first EF-hand of TnC^F29W, the fluorescence of G34DTnC^F29W increased upon addition of magnesium or calcium with affinities of 295 and 1.9 µM, respectively. Addition of 3 mM magnesium decreased the calcium sensitivity of TnC^F29W and G34DTnC^F29W ~2- and 6-fold, respectively. Exchange of G34DTnC^F29W into skinned psoas muscle fibers decreased fiber calcium sensitivity ~1.7-fold compared with TnC^F29W at 1 mM [magnesium]_free and ~3.2-fold at 3 mM [magnesium]_free. Thus, incorporation of a Z acid pair into the first EF-hand allows it to bind magnesium with high affinity. Furthermore, the data suggests that the second EF-hand, but not the first, of TnC is responsible for the competitive magnesium binding to the regulatory domain.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

The EF-hand is the most common Ca²⁺ binding motif found in nature (1). EF-hand proteins and their functions are numerous and diverse (for review see Ref. 2). In general, EF-hand domains can be classified functionally into two groups, those that regulate cellular activities through a reversible change in structure upon Ca²⁺ binding and release (such as the N-terminal regulatory domain of troponin C (TnC)¹ and both domains of calmodulin (CaM)) and those that simply buffer/transport Ca²⁺ (such as parvalbumin and calbindin D_9K) or anchor protein complexes (such as the C-terminal domain of TnC). Often the nonregulatory EF-hand domains bind Mg²⁺ competitively and display higher Ca²⁺ affinity than do the regulatory domains. However, it is becoming clear that regulatory EF-hand proteins, such as the N-terminal of CaM can also bind Mg²⁺ with a physiologically relevant affinity (3, 4). Thus, it is important to elucidate the mechanisms behind EF-hand Mg²⁺ binding.

The canonical EF-hand consists of 29 consecutive residues, with two helices flanking a 12-residue loop. The chelating loop residues in positions 1(+x), 3(+y), 5(+z), 7 (y), 9(x), and 12 (z) ligate Ca²⁺ through seven oxygen atoms arranged three dimensionally on the axes of a pentagonal bipyramid (for review see Refs. 5 and 6). Factors that control Ca²⁺ affinity are complex and involve residues within and outside of the Ca²⁺-binding loop (7-11). The mechanisms utilized for EF-hand Mg²⁺ binding are less understood. The smaller Mg²⁺ cation is typically complexed by some of the same loop residues used for Ca²⁺ binding, although through six oxygen atoms arranged in an octahedral geometry (for review see Refs. 11 and 12). Interestingly, synthetic EF-hand peptides were observed to bind Mg²⁺ when their Ca²⁺-binding loops contained a Z acid pair, chelating residues at loop positions +z and z (13, 14).

The metal binding properties of TnC have been studied extensively using various biophysical and biochemical techniques (15-19). These studies have demonstrated that the two C-domain EF-hands have ~10-fold higher Ca²⁺ affinity and greater than 100-fold slower Ca²⁺ exchange rates than the two N-domain EF-hands (18, 20). In addition to Ca²⁺, the C-domain sites also competitively bind Mg²⁺ with a physiologically relevant affinity. Because of their high Ca²⁺ and Mg²⁺ affinities and slow exchange rates, the C-domain sites are thought to play a structural role in muscle function by anchoring TnC into the Tn Complex.

The N-domain sites of TnC are considered to be Ca²⁺-specific under physiological Mg²⁺ concentrations, and generally accepted to be directly involved in the Ca²⁺-dependent regulation of muscle contraction (for review see Refs. 21 and 22). However, addition of Mg²⁺ has been shown to decrease the Ca²⁺ sensitivity of the regulatory domain of fluorescent TnCs in isolation (19, 23-27), in the Tn complex (23, 28), and in reconstituted muscle fibers (26, 29). Furthermore, several groups have demonstrated that increased [Mg²⁺] caused a decrease in Ca²⁺ sensitivity of Tn-regulated actomyosin ATPase and force development (26, 27, 30, 31). The question whether Mg²⁺ competes with Ca²⁺ for the N-domain sites remains unresolved and controversial. Some research groups have suggested the presence of auxiliary Mg²⁺ binding sites in TnC (15, 23), whereas others hypothesized Mg²⁺ as a direct Ca²⁺ competitor for the N-terminal regulatory sites (19, 27, 32).

Because TnC regulates muscle contraction as a part of the troponin complex and not in isolation, it is important to understand the interactions of TnC with TnI. The binding of TnI to TnC increases the Ca²⁺ sensitivity of the N- and C-domains of TnC ~10-fold (15, 33). TnC and TnI interact in an antiparallel orientation (34, 35) such that the Ca²⁺-dependent binding of the N-terminal regulatory domain of TnC to the C-domain of TnI is an early step in the generation of force in skeletal muscle (21). Previous studies demonstrated that region 96-116 of TnI interacts with actin and was largely responsible for the ability of TnI to inhibit the activity of the actomyosin ATPase that could be reversed upon TnC-Ca²⁺ binding (36-38). Recent studies implicated additional residues within 116-148 of TnI as being important for the complete inhibitory activity and regulatory interactions with actin and TnC (39-42). Thus, the Ca²⁺-dependent binding of the regulatory domain of TnC to TnI-(96-148) may be a good model system to study the Ca²⁺-dependent interactions between TnI and TnC.

The F29W mutation in chicken skeletal TnC (TnC^F29W) has been frequently used to study metal and ligand interactions with the regulatory N-domain sites of TnC (9, 20, 43-46). TnC^F29W is a physiologically active protein that produced maximal isometric tension with a Ca²⁺ sensitivity indistinguishable from that of recombinant TnC, when reconstituted into skinned skeletal fibers (47). Interestingly, whereas Ca²⁺ causes a large increase in TnC^F29W fluorescence, Mg²⁺ does not alter the fluorescence properties of Trp²⁹ (44, 45, 48). Previously, we have demonstrated that an endogenous Z acid pair was required for high affinity Mg²⁺ binding to the first EF-hand of fluorescent CaM^F19W (3). We wanted to test if Mg²⁺ binding could be engineered into the first EF-hand of TnC^F29W by substituting Gly in position 34 with Asp, thus introducing a Z acid pair (Fig. 1). We also wanted to determine how competitive Mg²⁺ binding to the first EF-hand of TnC would affect the physiological properties of G34DTnC^F29W reconstituted into skinned skeletal muscle fibers.

View larger version (29K): [in this window] [in a new window]   Fig. 1.   Predicted structures of the first Ca2+-binding loops of TnCF29W and G34DTnCF29W. Loop residues 29 through 41 are shown in a ribbon/stick configuration for TnCF29W and G34DTnCF29W as rendered by RASMOL (72). The F29W and G34D mutations were predicted by the computer software program WHAT IF (73) and constructed from the coordinates of the N-terminal domain crystal structure of chicken skeletal TnC (Protein Data Bank number 1AVS) (74). These cartoons are not meant to represent the actual structures but are used simply to demonstrate the potential effect of the G34D mutation and to demonstrate the location of the F29W mutation adjacent to the first Ca2+-binding loop. The spherical Ca2+ ion is shown to be coordinated by residues at positions 30 (+x), 32 (+y), 34 (+z), 36 (y), 38 (x), and 41 (z) that form the base and apexes of the pentagonal bipyramid geometry.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Materials-- Phenyl-Sepharose CL-4B, EDTA, and EGTA were purchased from Sigma. Quin-2 was purchased from Calbiochem (La Jolla, CA). All other chemicals were of analytical grade. The TnI-(96-148) peptide was synthesized and purified by the Alberta Peptide Institute (Edmonton Alberta, Canada).

Protein Mutagenesis and Purification-- The construction and expression of intact chicken skeletal TnC^F29W and isolated N-domain residues 1-90, TnC, both in pET3a, have been described (44, 49-50). Chicken skeletal fast TnI was prepared as described for the rabbit protein (51). The G34DTnC^F29W mutant was constructed from the TnC^F29W plasmid by primer-based site-directed mutagenesis using the QuikChange site-directed mutagenesis kit from Stratagene (La Jolla, CA). The mutation was confirmed by DNA sequence analysis. The plasmids for TnC^F29W and G34DTnC^F29W were transformed into Escherichia coli BL21(DE3)pLysS cells (Novagen) and purified as described previously (9).

Determination of Ca²⁺ and Mg²⁺ Affinities-- All steady-state fluorescence measurements were performed using a PerkinElmer Life Sciences LS5 spectrofluorimeter at 15 °C. Trp fluorescence was excited at 275 nm and monitored at 345 nm as microliter amounts of CaCl₂ or MgCl₂ were added to 1 ml of each TnC^F29W mutant (0.3 µM) in 200 mM MOPS (to prevent pH changes upon 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 (52). The Ca²⁺ and Mg²⁺ affinities are reported as dissociation constants K_d(Ca) and K_d(Mg), respectively. Each K_d represents a mean of 3-5 titrations fit with a logistic sigmoid function mathematically equivalent to the Hill equation, as previously described (8, 9).

Determination of Apparent TnI-(96-148) Peptide Affinities-- Trp fluorescence was monitored as described in the previous paragraph. Microliter amounts of TnI-(96-148) were added to 1 ml of each TnC^F29W mutant (0.6 µM) in 200 mM MOPS, 90 mM KCl, 2 mM EGTA, 1 mM [Ca²⁺]_free, 1 mM DTT, pH 7.0, at 15 °C. Each apparent peptide affinity represents a mean of 3 titrations fit with a logistic sigmoid function.

Determination of Ca²⁺ and Mg²⁺ Association and Dissociation Rates-- Ca²⁺ and Mg²⁺ association (k_on(Ca) and k_on(Mg)) and dissociation rates (k_off(Ca) and k_off(Mg)) 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⁺ and Mg²⁺ binding kinetics of the N-terminal domain within intact TnC^F29W and G34DTnC^F29W were obtained utilizing Trp fluorescence changes excited at 275 nm with emission monitored through a UV transmitting black glass filter (UG1 from Oriel, Stratford, CT). Ca²⁺ dissociation rates in the absence of Mg²⁺ were also measured using the fluorescent Ca²⁺ chelator Quin-2. Whereas the fluorescence of Trp²⁹ was selective for N-terminal Ca²⁺ dissociation, Quin-2 fluorescence reported Ca²⁺ dissociation from both the N- and C-domains within TnC^F29W. However, the Ca²⁺ dissociation rate from the N-terminal domain within TnC^F29W was easily distinguished from the rate of Ca²⁺ dissociation from the C-terminal domain because the latter rate was >95-fold slower in the absence and presence of TnI-(96-148) or intact TnI. Quin-2 fluorescence was excited at 330 nm, with its emission monitored through a 510-nm broad band-pass interference filter (Oriel). Each data set represents an average of 10-15 traces, fit with a single exponential (variance <2 × 10⁴). The curve fitting program (by P. J. King, Applied Photophysics Ltd.) uses the nonlinear Levenberg-Marquardt algorithm. The buffer used in all the Ca²⁺ stopped-flow experiments was 10 mM MOPS, 90 mM KCl, 1 mM DTT, pH 7.0.

Calculation of Ca²⁺ and Mg²⁺ Association Rates-- The Ca²⁺ and Mg²⁺ association rates were calculated using the equation k_on = k_off/K_d, where k_off represents the concerted release of two Ca²⁺ or single Mg²⁺ ions and K_d represents the binding event of two Ca²⁺ or one Mg²⁺ to the N-domain of TnC, as previously described (3, 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 (53). 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 with pens 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 (54). 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 (-log[Ca²⁺]) 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 baseline 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 TnC^endogenous), which led to an average force per cross-sectional area of 89 ± 4 kilo newton/m². Force versus pCa was then established or 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 (55). The average force in pCa 4.0 generated after TnC^endogenous extraction was 5 ± 1% of the maximal force generated by the native fibers. The fiber was then washed three times in pCa 9.0 solution to remove any residual trifluoperazine dihydrochloride and then soaked for 2 min in a pCa 9.0 solution containing 16.7 µM recombinant TnC^F29W or G34DTnC^F29W. The maximal force generated by the recombinant TnCs averaged 93 ± 1 and 62 ± 5% for TnC^F29W and G34DTnC^F29W, respectively. Because additional fibers reconstituted with 33.4 µM G34DTnC^F29W or TnC^F29W did not improve the maximal amount of force recovery or affect the force versus pCa relationship, these data were then combined with the 16.7 µM measurements. All of the reconstituted fibers were then exposed to a series of pCa solutions varying from pCa 9.0 to 3.0 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. Because fibers reconstituted with recombinant TnC did not completely recover maximal force, force versus [Ca²⁺] curves were also performed on fibers in which TnC was partially extracted. Four fibers were soaked in TnC extraction solution for variable times to partially extract TnC^endogenous.

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Effect of Ca²⁺ and Mg²⁺ on the Fluorescence Spectra of the N-terminal Domains within TnC^F29W and G34DTnC^F29W-- TnC^F29W undergoes an ~2.4-fold increase in Trp fluorescence upon Ca²⁺ binding to its N-domain sites at 15 °C (Fig. 2A). No change in Trp fluorescence was observed upon addition of 50 mM Mg²⁺ to TnC^F29W (Fig. 2A). We previously demonstrated that high Mg²⁺ binding to the first EF-hand of CaM required the presence of an endogenous Z acid pair (3). When a Z acid pair was introduced into the first EF-hand of TnC^F29W (i.e. G34DTnC^F29W), addition of Ca²⁺ led to an ~2.2-fold increase in Trp fluorescence (Fig. 2B). Furthermore, G34DTnC^F29W also underwent an ~1.4-fold increase in its Trp fluorescence upon addition of Mg²⁺ (Fig. 2B). These results demonstrate that substitution of Gly-34 with Asp in the first EF-hand of TnC led to Mg²⁺ binding to this EF-hand.

View larger version (19K): [in this window] [in a new window]   Fig. 2.   Effect of Ca2+ and Mg2+ on the fluorescence spectra of TnCF29W and G34DTnCF29W. Fluorescence emission spectra of TnCF29W (panel A) or G34DTnCF29W (panel B) in the apo, Ca2+ saturated, or Mg2+ saturated states. The spectra were recorded with an excitation wavelength of 275 nm. Protein concentration was 1 µM in 1 ml of 200 mM MOPS, 90 mM KCl, 2 mM EGTA, 1 mM DTT, pH 7.0, at 15 °C. Trp fluorescence spectra were recorded before the addition of metals (Apo), after the addition of either 50 mM Mg2+ (+Mg2+) or 1 mM [Ca2+]free (+Ca2+).

Measurement of the Mg²⁺ Binding Affinity for TnC and the N-terminal Domains within TnC^F29W and G34DTnC^F29W-- Fig. 3A shows the Mg²⁺-dependent increase in Trp fluorescence of G34DTnC^F29W (). G34DTnC^F29W exhibited a half-maximal increase in Trp fluorescence at 295 ± 10 µM. However, addition of Mg²⁺ (up to 50 mM) caused no change in the Trp fluorescence signal of TnC^F29W (Fig. 3A, ). Thus, as expected, the G34D mutation in TnC^F29W incorporates physiological Mg²⁺ binding to the first EF-hand of TnC^F29W.

View larger version (15K): [in this window] [in a new window]   Fig. 3.   Measurement of Mg2+ binding and dissociation from G34DTnCF29W and TnCF29W. Panel A shows the Mg2+-dependent change in Trp fluorescence as a function of -log[Mg2+] for G34DTnCF29W () or TnCF29W (). For panels A and B, the protein, buffer, and experimental conditions were as described in the legend to Fig. 2. Trp fluorescence was monitored at 345 nm, with excitation at 275 nm. Each data point represents an average ± S.E. of 3-5 titrations. Panel B shows the Mg2+-dependent decrease in Trp fluorescence of Ca2+ saturated G34DTnCF29W (), TnCF29W (), or TnC () as a function of -log[Mg2+]. Initially each protein was subjected to 10 µM [Ca2+]free (pCa 5.0) and then titrated with increasing concentrations of Mg2+. 100% Trp fluorescence corresponds to the Ca2+ bound state, whereas 0% fluorescence corresponds to the Mg2+ saturated state in the presence of 10 µM Ca2+. The Trp fluorescence decreased 71, 88, and 98% of the total fluorescence change between the Ca2+ bound and apo states for G34DTnCF29W, TnCF29W, and TnC, respectively. The Ca2+ affinity of TnC was shown to be nearly identical to that of TnCF29W (44). Panel C shows Mg2+ dissociation from G34DTnCF29W and TnCF29W. The time course of the change in Trp fluorescence is shown as EDTA dissociates Mg2+ from the N-terminal domains of G34DTnCF29W and TnCF29W. Each protein (2 µM) in the same buffer as Fig. 2 plus 10 mM Mg2+ was rapidly mixed with an equal volume of 20 mM EDTA in the same buffer at 15 °C. Trp fluorescence was monitored through a UV-transmitting black glass filter (UG1 from Oriel) with excitation at 275 nm. The traces have been staggered for clarity. Each trace is an average of at least 15 traces, and the data were fit with a single exponential (excluding TnCF29W, which was flat, variance <2 × 104). The kinetic traces were triggered at time 0, the first ~2 ms of premixing time is shown (the apparent lag phase), and the average trace was fit after mixing was complete. Control experiments in which each protein (2 µM) in the same buffer as Fig. 2 plus 10 mM Mg2+ was rapidly mixed with an equal volume of the same buffer were flat lines.

Fig. 3B shows that in the presence of 10 µM [Ca²⁺]_free the Trp fluorescence signal of TnC (), TnC^F29W (), and G34DTnC^F29W () was greater than 90% saturated, which Mg²⁺ subsequently decreased in a concentration-dependent manner. Mg²⁺ half-maximally decreased the Trp fluorescence of TnC, TnC^F29W and G34DTnC^F29W at 7.3 ± 0.8, 9 ± 1, and 1.9 ± 0.2 mM, respectively. Thus, the Ca²⁺-dependent fluorescence of TnC, TnC^F29W, and G34DTnC^F29W was reversed by Mg²⁺ binding, presumably because of the closing of the N-terminal hydrophobic pocket of TnC and not because of any type of interference from Mg²⁺ binding to the C-terminal domain. Knowing the K_d(Ca) for each N-terminal domain protein (determined as described later) and assuming competitive Mg²⁺ binding, the K_d(Mg) of TnC, TnC^F29W, and G34DTnC^F29W was calculated to be ~1.8 mM, 2.2 mM, and 303 µM, respectively. Thus, the calculated Mg²⁺ affinity for G34DTnC^F29W was identical to that measured directly as described above. Furthermore, even though TnC^F29W does not undergo a change in fluorescence upon Mg²⁺ binding (Fig. 3A, ) its N-terminal domain appears to bind Mg²⁺ and displace Ca²⁺. Table I summarizes the data for Mg²⁺ binding to TnC^F29W and G34DTnC^F29W.

Measurement of the Mg²⁺ Dissociation Rate from the N-terminal Domain within G34DTnC^F29W-- Fluorescence stopped-flow measurements were done to determine the rate of Mg²⁺ dissociation from G34DTnC^F29W. Fig. 3C shows the time course of the EDTA-induced decrease in Trp fluorescence for G34DTnC^F29W. At 15 °C, EDTA dissociated Mg²⁺ from G34DTnC^F29W at 570 ± 15 s¹. There was no change in Trp fluorescence when the same experiment was performed with TnC^F29W because Mg²⁺ binding to the N-terminal sites of TnC^F29W does not lead to a change in the Trp fluorescence.

Measurements of Ca²⁺ Binding Affinities for the N-terminal Domains within TnC^F29W and G34DTnC^F29W in the Absence and Presence of 3 mM Mg²⁺-- The Ca²⁺ binding affinities of TnC^F29W and G34DTnC^F29W were measured following the Ca²⁺ induced increases in Trp fluorescence in the absence and presence of 3 mM Mg²⁺ at 15 °C. Fig. 4A shows that in the absence of Mg²⁺, TnC^F29W () and G34DTnC^F29W () exhibited half-maximal Ca²⁺-dependent increases in Trp fluorescence at 3.3 ± 0.1 and 1.9 ± 0.1 µM Ca²⁺, respectively. Therefore, in the absence of Mg²⁺, G34DTnC^F29W exhibited an ~1.7-fold increase in its Ca²⁺ affinity, relative to that of TnC^F29W. Fig. 4A also shows that in the presence of 3 mM Mg²⁺, TnC^F29W () and G34DTnC^F29W () exhibited half-maximal increases in Trp fluorescence at 5.9 ± 0.4 and 11.6 ± 0.9 µM Ca²⁺, respectively. Therefore, in the presence of 3 mM Mg²⁺, G34DTnC^F29W exhibited an ~2-fold decrease in its Ca²⁺ sensitivity, relative to that of TnC^F29W. Furthermore, 3 mM Mg²⁺ shifts the Ca²⁺ sensitivity of TnC^F29W and G34DTnC^F29W ~1.8- and 6-fold, respectively. Again, assuming competitive Mg²⁺ binding, the K_d(Mg) of TnC^F29W and G34DTnC^F29W was ~3.8 mM and 590 µM, respectively. These values are in good agreement with the direct and competitive Mg²⁺ binding studies described above (see Table I). Thus, in the presence of 3 mM Mg²⁺ both proteins are capable of opening their N-terminal hydrophobic pockets in a Ca²⁺-dependent manner, albeit with decreased Ca²⁺ sensitivity. Table II summarizes the data for Ca²⁺ binding to TnC^F29W and G34DTnC^F29W.

View larger version (17K): [in this window] [in a new window]   Fig. 4.   Measurement of Ca2+ binding to TnCF29W, G34DTnCF29W, the TnCF29W·TnI-(96-148) complex, or the G34DTnCF29W·TnI-(96-148) complex in the presence and absence of 3 mM Mg2+. Panel A shows the Ca2+-dependent increase in Trp fluorescence as a function of -log[Ca2+] for TnCF29W (), G34DTnCF29W (), TnCF29W + 3 mM Mg2+ (), or G34DTnCF29W + 3 mM Mg2+ (). One-hundred percent fluorescence corresponds to a 2.4-, 2.2-, 2.4-, and 1.6-fold fluorescence increase for TnCF29W, G34DTnCF29W, TnCF29W + 3 mM Mg2+, or G34DTnCF29W + 3 mM Mg2+, respectively. Experimental conditions were the same as described in the legend to Fig. 3, panel A. Panel B shows the TnI-(96-148)-dependent decrease in Trp fluorescence as a function of -log[TnI-(96-148)] for Ca2+-saturated TnCF29W () or G34DTnCF29W (). 100% Trp fluorescence corresponds to the Ca2+-saturated state, whereas 0% Trp fluorescence corresponds to the Ca2+-TnI-(96-148)-saturated state of each TnCF29W. The Trp fluorescence decreased 55 and 49% of the total fluorescence change between the Ca2+-saturated and apo states for TnCF29W and G34DTnCF29W, respectively. Panel C shows the Ca2+-dependent increase in Trp fluorescence as a function of -log[Ca2+] for the TnCF29W·TnI-(96-148) complex (), the G34DTnCF29W·TnI-(96-148) complex (), the TnCF29W·TnI-(96-148) complex + 3 mM Mg2+ (), or the G34DTnCF29W·TnI-(96-148) complex + 3 mM Mg2+ () under the same buffer conditions as panel A. The TnC protein concentration was 0.3 µM, whereas the TnI-(96-148) peptide concentration was 3 µM. One-hundred percent fluorescence corresponds to a 1.4-, 1.4-, 1.4-, and 1.1-fold fluorescence increase for the TnCF29W·TnI-(96-148) complex, the G34DTnCF29W·TnI-(96-148) complex, the TnCF29W·TnI-(96-148) complex + 3 mM Mg2+, or the G34DTnCF29W·TnI-(96-148) complex + 3 mM Mg2+, respectively.

Measurement of Ca²⁺ Binding Affinities for the N-terminal Domains within TnC^F29W·TnI-(96-148) and G34DTnC^F29W·TnI- (96-148) Complexes in the Absence and Presence of 3 mM Mg²⁺-- A partial decrease in Ca²⁺-saturated TnC^F29W and G34DTnC^F29W fluorescence occurring upon binding of TnI-(96-148) was utilized to determine the apparent peptide binding affinity (39). Fig. 4B demonstrates that the apparent TnI-(96-148) affinities for Ca²⁺ saturated TnC^F29W and G34DTnC^F29W were nearly identical at 457 ± 16 and 508 ± 9 nM, respectively. Thus, any differences in Ca²⁺ sensitivity observed between the two proteins in the presence of the peptide could not be because of differences in peptide affinities. The Ca²⁺ binding affinities for the TnC^F29W·TnI-(96-148) and G34DTnC^F29W·TnI-(96-148) complexes were measured following the Ca²⁺-induced increases in Trp fluorescence in the absence and presence of 3 mM Mg²⁺ at 15 °C. Fig. 4C shows that in the absence of Mg²⁺, the TnC^F29W·TnI-(96-148) () and G34DTnC^F29W·TnI-(96-148) () complexes exhibited half-maximal Ca²⁺-dependent increases in Trp fluorescence at 267 ± 3 and 147 ± 2 nM Ca²⁺, respectively. Therefore, in the absence of Mg²⁺, the G34DTnC^F29W·TnI-(96-148) complex exhibited an ~1.7-fold increase in its Ca²⁺ affinity, relative to that of the TnC^F29W·TnI-(96-148) complex. Consistent with intact TnI binding to TnC, TnI-(96-148) enhanced the Ca²⁺ sensitivity of the regulatory domain of TnC^F29W ~12-fold (14, 32) and to G34DTnC^F29W ~13-fold (see Table II). Fig. 4C also shows that in the presence of 3 mM Mg²⁺, the TnC^F29W-TnI-(96-148) () and G34DTnC^F29W·TnI-(96-148) () complexes exhibited half-maximal increases in Trp fluorescence at 397 ± 13 and 765 ± 41 nM Ca²⁺, respectively. Therefore, in the presence of 3 mM Mg²⁺, the G34DTnC^F29W·TnI-(96-148) complex exhibited ~1.9-fold decrease in Ca²⁺ sensitivity, relative to that of the TnC^F29W·TnI-(96-148) complex. Furthermore, 3 mM Mg²⁺ shifts the Ca²⁺ sensitivity of the TnC^F29W·TnI-(96-148) and G34DTnC^F29W·TnI-(96-148) complexes ~1.5- and 5.2-fold, respectively. Again, assuming competitive Mg²⁺ binding, the K_d(Mg) of TnC^F29W and G34DTnC^F29W in complex with TnI-(96-148) was ~6.2 mM and 714 µM, respectively. Thus, unlike Ca²⁺ binding to the regulatory sites of TnC^F29W and G34DTnC^F29W, Mg²⁺ binding is not enhanced by TnI-(96-148) binding and may actually be slightly decreased (see Table I). Furthermore, in the presence of 3 mM Mg²⁺, TnI-(96-148) increased the Ca²⁺ sensitivity to the regulatory domains of both TnC^F29W and G34DTnC^F29W ~15-fold. Thus, Mg²⁺ did not affect the enhancement of the Ca²⁺ affinity to the regulatory domain of the TnCs caused by TnI-(96-148) binding, consistent with Mg²⁺ acting only as a competitor for Ca²⁺ binding.

Measurement of Ca²⁺ Dissociation Rates from the N-terminal Domains within TnC^F29W and G34DTnC^F29W Induced by EGTA, Quin-2, or Mg²⁺-- Fluorescence stopped-flow measurements, utilizing the EGTA-induced decreases in Trp fluorescence, were conducted to determine the rates of Ca²⁺ dissociation from the N-terminal sites of TnC^F29W and G34DTnC^F29W. Fig. 5A shows the time course of the EGTA-induced decreases in Trp fluorescence as Ca²⁺ was dissociated from TnC^F29W and G34DTnC^F29W (Trp EGTA traces). At 15 °C, excess EGTA removed Ca²⁺ from TnC^F29W and G34DTnC^F29W at 342 ± 5 and 403 ± 8 s¹, respectively. Under saturating Ca²⁺ conditions, the presence of 3 mM Mg²⁺ did not affect the rates of Ca²⁺ dissociation from TnC^F29W or G34DTnC^F29W (data not shown, Table II). These results were consistent with the Ca²⁺ dissociation rate from the regulatory sites of the fluorescent TnC^Danz not being affected by Mg²⁺ (56).

View larger version (21K): [in this window] [in a new window]   Fig. 5.   Measurement of Ca2+ dissociation from TnCF29W or G34DTnCF29W in the presence or absence of either TnI-(96-148) or intact TnI using EGTA, Quin-2, or Mg2+. Panel A shows the rate of Ca2+ dissociation from TnCF29W and G34DTnCF29W following the decreases in Trp fluorescence or the increase in Quin-2 fluorescence. The time course of the decrease in Trp fluorescence is shown as EGTA dissociates Ca2+ from the N-terminal domains of G34DTnCF29W and TnCF29W (Trp EGTA traces). Each protein (2 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 5 mM EGTA at 15 °C. Trp fluorescence was monitored as described in the legend to Fig. 3, panel C. Panel A also shows the time course of the increase in Quin-2 fluorescence as Quin-2 dissociates Ca2+ from the N-terminal domains of G34DTnCF29W and TnCF29W (Quin-2 traces). Each protein (6 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. Quin-2 fluorescence was monitored through a 510-nm broad band-pass interference filter with excitation at 330 nm. All the traces have been normalized and displaced vertically for clarity. Panel B shows the time course of decrease in Trp fluorescence as Mg2+ displaced Ca2+ from the N-terminal domains of TnCF29W or G34DTnCF29W (Trp Mg2+ traces). Each protein (4 µM) in the same buffer as panel A plus 15 µM Ca2+ was rapidly mixed with an equal volume of the same buffer plus 150 mM Mg2+. Trp fluorescence was monitored as described in the legend to Fig. 3, panel C. The traces have been normalized and staggered for visual clarity. Panel C shows the rate of Ca2+ dissociation from TnCF29W and G34DTnCF29W following the decreases in Trp fluorescence in the presence of TnI-(96-148) or the increase in Quin-2 fluorescence in the presence of intact TnI. The time course of the decrease in Trp fluorescence is shown as EGTA-dissociated Ca2+ from the regulatory Ca2+ binding sites of the G34DTnCF29W·TnI-(96-148) and the TnCF29W·TnI-(96-148) complexes (Trp TnI96-148 EGTA traces). Each TnC protein (0.6 µM) plus TnI-(96-148) peptide (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 as described in the legend to Fig. 3, panel C. Panel C also shows the time course of the increase in Quin-2 fluorescence as Quin-2 dissociates Ca2+ from the regulatory Ca2+ binding sites of the G34DTnCF29W·TnI and the TnCF29W·TnI complexes (Quin-2 intact TnI traces). Each TnC (3 µM) plus intact TnI (30 µM) in 10 mM MOPS, 90 mM KCl, 1 mM DTT, pH 7.0, plus 10 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 as described in panel A. All the traces have been normalized and displaced vertically for clarity. All the traces in Panels A-C are an average of at least 15 traces fit with a single exponential as described in the legend to Fig. 3, panel C (variance <2 × 104).

To verify that the Trp signal changes were accurately reporting the true Ca²⁺ dissociation rates and not a slower or faster structural change, Ca²⁺ dissociation rates in the absence of Mg²⁺ were also measured with the fluorescent Ca²⁺ chelator Quin-2. Fig. 5A also shows the time course of the Quin-2-induced increases in fluorescence as Ca²⁺ was dissociated from TnC^F29W and G34DTnC^F29W (Quin-2 traces). Nearly identical Ca²⁺ dissociation rates were measured using Quin-2 fluorescence for TnC^F29W and G34DTnC^F29W at 346 ± 3 and 397 ± 2 s¹, respectively, as were measured by the EGTA-induced Trp changes. Therefore, G34DTnC^F29W exhibited an ~1.2-fold faster N-terminal Ca²⁺ dissociation rate, relative to that of TnC^F29W. Similarly, the C-terminal Ca²⁺ dissociation rates from TnC^F29W and G34DTnC^F29W were nearly identical at 0.48 ± 0.01 and 0.56 ± 0.01 s¹, respectively (data not shown).

Fig. 3B demonstrated that Mg²⁺ could reverse the Ca²⁺-induced increase in Trp fluorescence for both TnC^F29W and G34DTnC^F29W. If Mg²⁺ binding to the N-terminal domains of TnC^F29W and G34DTnC^F29W was truly competitive with Ca²⁺ binding, then it would be expected that Mg²⁺ could not bind these Ca²⁺ saturated sites until Ca²⁺ dissociates, as has been demonstrated for the Ca²⁺- and Mg²⁺-binding protein parvalbumin (57, 58). Fig. 5B shows the time course of Mg²⁺ binding to Ca²⁺-saturated TnC^F29W and G34DTnC^F29W (Trp Mg²⁺ traces) following the decreases in Trp fluorescence in a stopped-flow apparatus at 15 °C. Mg²⁺ was able to displace Ca²⁺ from TnC^F29W and G34DTnC^F29W at 348 ± 2 and 399 ± 6 s¹, respectively. Because these values are virtually identical to the Ca²⁺ dissociation rates in the absence of Mg²⁺, it is clear that Mg²⁺ binding to Ca²⁺-saturated TnC^F29W and G34DTnC^F29W was limited by their respective Ca²⁺ dissociation rates, supporting the hypothesis that Mg²⁺ binds competitively with Ca²⁺.

Measurement of Ca²⁺ Dissociation Rates from the N-terminal Domains within TnC^F29W and G34DTnC^F29W Induced by EGTA or Quin-2 in the Presence of TnI-(96-148) or Intact TnI-- The rates of Ca²⁺ dissociation from TnC^F29W and G34DTnC^F29W induced by EGTA were also measured in the presence of TnI-(96-148) (Fig. 5C, Trp TnI-(96-148) EGTA traces). At 15 °C, EGTA removed Ca²⁺ from the TnC^F29W·TnI-(96-148) and G34DTnC^F29W·TnI-(96-148) complexes at 10.6 ± 0.2 and 6.0 ± 0.3 s¹, respectively. Nearly identical rates were measured using Quin-2 to remove Ca²⁺ from the TnC^F29W·TnI-(96-148) and G34DTnC^F29W·TnI-(96-148) complexes at 10.6 ± 0.2 and 6.3 ± 0.1 s¹, respectively (data not shown). Thus, the N-terminal Trp signal change for both proteins in the absence and presence of TnI-(96-148) accurately reports Ca²⁺ binding and dissociation.

To verify that TnI-(96-148) 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 also shows the time course of the Quin-2-induced increases in fluorescence as Ca²⁺ was dissociated from the TnC^F29W·TnI complex at 9.1 ± 0.8 s¹ and the G34DTnC^F29W·TnI complex at 5.4 ± 0.5 s¹ (Quin-2 intact TnI traces). Thus, the Ca²⁺ dissociation rates from the TnC^F29W·TnI and G34DTnC^F29W·TnI complexes were similar to the Ca²⁺ dissociation rates when complexed with TnI-(96-148). Furthermore, G34DTnC^F29W complexed with either TnI or TnI-(96-148) exhibited an ~1.7-fold slower Ca²⁺ dissociation rate, relative to that of the TnC^F29W complexes. Thus, the binding of TnI to the regulatory domain of G34DTnC^F29W slows Ca²⁺ dissociation ~73-fold, whereas this effect on TnC^F29W was slowed only ~37-fold. However, both TnI-(96-148) and intact TnI similarly slowed the Ca²⁺ dissociation rate from the C-terminal domains within TnC^F29W and G34DTnC^F29W only ~4-fold (data not shown).

Following the Trp fluorescence signal, in the presence of TnI-(96-148), 3 mM Mg²⁺ did not affect the rate of Ca²⁺ dissociation from TnC^F29W but increased the Ca²⁺ dissociation rate of the G34DTnC^F29W·TnI-(96-148) complex to that of the TnC^F29W·TnI-(96-148) complex (data not shown, Table II). The reason for this Mg²⁺ effect on the G34DTnC^F29W·TnI-(96-148) complex is currently unknown. One possibility is that the increased negative charge of Asp-34 decreased the rate of Ca²⁺ dissociation from the regulatory domain of G34DTnC^F29W in the presence of TnI-(96-148), which becomes screened by the positive Mg²⁺ ions.

Calculated Ca²⁺ Association Rates to the N-terminal Domains within TnC^F29W and G34DTnC^F29W-- Knowing the affinity of Ca²⁺ (K_d(Ca)) from the Ca²⁺ dependence of the increase in TnC^F29W and G34DTnC^F29W Trp fluorescence and the rate of Ca²⁺ dissociation (k_off(Ca)) from these same sites, we could calculate the rate of Ca²⁺ association (k_on(Ca)) from the equation: k_on = k_off/K_d. The calculated values of k_on(Ca) for TnC^F29W and G34DTnC^F29W were 1.0 × 10⁸ and 2.1 × 10⁸ M¹ s¹, respectively at 15 °C. Thus, G34DTnC^F29W possesses a ~2.1-fold faster calculated Ca²⁺ association rate than does TnC^F29W. Furthermore, the k_on(Mg) calculated for G34DTnC^F29W was ~1.9 × 10⁶ M¹ s¹, consistent with the slower dehydration rate of Mg²⁺ compared with Ca²⁺ that tends to limit the rate of Mg²⁺ binding to proteins (12).

Measurement of Ca²⁺ Association Rates to the N-terminal Domains within TnC^F29W and G34DTnC^F29W in the Absence and Presence of 3 mM Mg²⁺-- To verify these rapid, nearly diffusion controlled Ca²⁺ association rates, the Ca²⁺-induced increase in Trp fluorescence was measured in a stopped-flow apparatus. Fig. 6A shows the time course of the rapid Ca²⁺-induced increase in Trp fluorescence of G34DTnC^F29W as the [Ca²⁺] was increased from 0 to 5 µM. As the [Ca²⁺] increases, the observed rate of Ca²⁺ binding increases as expected for a second order reaction. At 1, 2.5, and 5 µM Ca²⁺, the reaction occurred at 903 ± 37, 1103 ± 56, and 1660 ± 126 s¹, respectively. A plot of the observed rate versus [Ca²⁺] was fit by a linear regression (r² = 0.985, Fig. 6C, ) from which the Ca²⁺ association rate to G34DTnC^F29W was estimated at 1.9 ± 0.2 × 10⁸ M¹ s¹. Similar studies with TnC^F29W yielded a Ca²⁺ association rate of 1.57 ± 0.06 × 10⁸ M¹ s¹ (r² = 0.995, Fig. 6C, ). Thus, the measured Ca²⁺ association rates to the N-terminal domains of TnC^F29W and G34DTnC^F29W were extremely rapid and essentially the same as the calculated Ca²⁺ association rates.

View larger version (23K): [in this window] [in a new window]   Fig. 6.   Measurement of the Ca2+ association rates to TnCF29W and G34DTnCF29W in the absence and presence of 3 mM Mg2+. Panel A shows the time course of the increase in Trp fluorescence of G34DTnCF29W as increasing concentrations of Ca2+ were rapidly mixed with the protein. 4 µM G34DTnCF29W in the same buffer as described in the legend to Fig. 5 was rapidly mixed with the same buffer in the presence of increasing [Ca2+] at 15 °C. The indicated [Ca2+] represents the amount of Ca2+ that was presented to the apo-TnC immediately after mixing was complete. Trp fluorescence was monitored as described in the legend to Fig. 3, panel C. Each trace is an average of at least 10-15 traces fit with a single exponential as described in Fig. 3, panel C (except No Ca2+, which was flat, variance <2 × 104). Panel B shows the time course of the increases in Trp fluorescence of G34DTnCF29W as increasing concentrations of Ca2+ were rapidly mixed with the protein in the presence of 3 mM Mg2+. The experimental conditions and analysis were identical to the reactions as demonstrated in panel A, except 3 mM Mg2+ was added to the buffer and greater concentrations of Ca2+ were required to achieve similar fluorescence values. Panel C shows plots of the observed Ca2+ association rate versus the [Ca2+] in the mixing chamber immediately after mixing was complete for TnCF29W (), G34DTnCF29W (), TnCF29W plus 3 mM Mg2+ (), or G34DTnCF29W plus 3 mM Mg2+ (). Each point represents the average ± S.E. of at least three determinants. The data were fit with a linear regression where the slope represents the calculated Ca2+ association rate, except for G34DTnCF29W in the presence of 3 mM Mg2+, which demonstrated no Ca2+ dependence of its association rate as, described under "Results."

At 3 mM Mg²⁺, G34DTnC^F29W was greater than 90% saturated with Mg²⁺ (Fig. 3A, ). Fig. 6B shows the time course of the Ca²⁺-induced increase in Trp fluorescence of G34DTnC^F29W in the presence of 3 mM Mg²⁺ as the [Ca²⁺] was increased from 0 to 25 µM. As the [Ca²⁺] increased, the rate of the reaction did not increase as expected for a second order reaction but remained fairly static at 440 ± 24 s¹ (Fig. 6, B and C, ). These static observed Ca²⁺ association rates, close to the Mg²⁺ dissociation rate from G34DTnC^F29W (see Table I), were consistent with the interpretation of competitive Ca²⁺ and Mg²⁺ binding to the N-terminal domain of G34DTnC^F29W. Thus, in the presence of saturating [Mg²⁺], Ca²⁺ cannot bind to the regulatory domain of G34DTnC^F29W until Mg²⁺ dissociates. Similar studies with TnC^F29W yielded a Ca²⁺ association rate in the presence of 3 mM Mg²⁺ of 0.45 ± 0.03 × 10⁸ M¹ s¹ (r² = 0.999, Fig. 6C, ). Thus, at these [Mg²⁺] and [Ca²⁺] the Mg²⁺ dissociation rate from the regulatory domain of TnC^F29W was not rate-limiting for Ca²⁺ binding, although it was slower than in the absence of Mg²⁺. Mathematical modeling of the Ca²⁺ association rate experiments with TnC^F29W in the presence of 3 mM Mg²⁺ predicted the Mg²⁺ dissociation rate to be ~6000 s¹ assuming a k_on(Mg) of 2 × 10⁶ M¹ s¹ leading to a K_d(Mg) of 3 mM (data not shown). Thus, 3 mM Mg²⁺ slows the Ca²⁺ association rate to TnC^F29W only ~3.5-fold but drastically slows the Ca²⁺ association rate to G34DTnC^F29W by ~4.3 × 10⁵-fold. Table II also compares the Ca²⁺ binding properties of TnC^F29W and G34DTnC^F29W in the absence or presence of 3 mM Mg²⁺, with or without TnI-(96-148).

Over longer times (0-5 s), in the absence of Mg²⁺, as the [Ca²⁺] was increased from 0 to 5 µM slow decreases in the Trp fluorescence signal were observed (~1-2 s¹) for both TnC^F29W and G34DTnC^F29W (less than 5% of the total Trp change, data not shown). The amplitudes of these slow decreases in Trp fluorescence decreased with increasing [Ca²⁺] and were absent when the [Ca²⁺] exceeded 5 µM. Computer modeling of these reactions predicted that these decreases in Trp fluorescence were associated with Ca²⁺ removal from the N-domain sites of the TnCs by the high affinity C-domain sites that possess an ~100-fold slower Ca²⁺ association rate and ~10-fold higher Ca²⁺ affinity (20). Furthermore, the rapid increase in Trp fluorescence for both TnC^F29W and G34DTnC^F29W became too fast to observe as the [Ca²⁺] exceeded 5 µM. Unexpectedly, as the [Ca²⁺] was increased from 5 µM up to 1 mM another slow second order rate constant was observed for TnC^F29W and G34DTnC^F29W at ~1.0 × 10⁶ and 2.0 × 10⁶ M¹ s¹, respectively (less than 10% of the total Trp signal change, data not shown). This second Ca²⁺ association rate process was abolished in the presence of 3 mM Mg²⁺ for both TnCs and was not observed with TnC, which was missing the C-terminal domain Ca²⁺ binding sites (data not shown). Thus, these slow Ca²⁺ association rates were associated with the C-terminal domain of the TnCs and were consistent with previously calculated Ca²⁺ association rates to the C-terminal Ca²⁺/Mg²⁺ sites of TnC (18, 20). Thus, the Trp²⁹ fluorescence was marginally influenced by C-terminal Ca²⁺ binding as has been reported for TnC^Danz, TnC^F52W, and TnC^F78W (17, 24, 59).

Functional Properties of TnC^F29W and G34DTnC^F29W-- To investigate the effects of increased competitive binding of Mg²⁺ to the regulatory domain of G34DTnC^F29W compared with TnC^F29W, both proteins were reconstituted into skinned rabbit psoas muscle fibers. Fig. 7A shows the Ca²⁺-dependent increase in skinned psoas muscle force with TnC^endogenous (), TnC^F29W (), or G34DTnC^F29W () in the presence of physiological 1 mM [Mg²⁺]_free (60). Half-maximal force occurred at 331 ± 7, 531 ± 62, and 1830 ± 460 nM Ca²⁺ for TnC^endogenous, TnC^F29W, and G34DTnC^F29W, respectively. Thus, 1 mM [Mg²⁺]_free shifted the Ca²⁺ sensitivity of force development with G34DTnC^F29W ~3.4-fold compared with TnC^F29W, consistent with physiological competitive Mg²⁺ binding to G34DTnC^F29W.

View larger version (21K): [in this window] [in a new window]   Fig. 7.   Ca2+ sensitivity of force development with unextracted and reconstituted psoas muscle fibers with TnCF29W and G34DTnCF29W in the presence of 1 or 3 mM [Mg2+]free. Panel A shows the Ca2+ dependence of isometric force development in unextracted (TnCendogenous, ) and reconstituted psoas muscle fibers with TnCF29W () or G34DTnCF29W () in the presence of 1 mM [Mg2+]free. Each trace represents the mean ± S.E. of at least three separate fibers fit with a logistic sigmoid curve mathematically equivalent to the Hill equation. The conditions for the experiments are as described under "Experimental Procedures." Panel B shows the Ca2+ dependence of isometric force development in unextracted (TnCendogenous, ) and reconstituted psoas muscle fibers with TnCF29W () or G34DTnCF29W () in the presence of 3 mM [Mg2+]free. Analysis and experimental conditions were the same as described in panel A, except for the increased [Mg2+]free. Panel C shows the -fold decreases in the [Ca2+] that produced half-maximal force ([Ca2+]50) for reconstituted fibers with TnCF29W (in 1 mM [Mg2+]free () or 3 mM [Mg2+]free () or G34DTnCF29W (in 1 mM [Mg2+]free () or 3 mM [Mg2+]free ()) versus percent of maximal force recovery compared with the mean [Ca2+]50 of the unextracted fibers ([Ca2+]50 endogenous). Each point represents the mean ± S.E. for all the individual reconstituted fibers. Panel C also shows the -fold decrease in [Ca2+]50 for fibers in which TnCendogenous was partially extracted versus percent of maximal force recovery compared with [Ca2+]50 endogenous (TnCendogenous Extraction, ). The data from four fibers were fit with a linear regression to determine the relationship between incomplete force recovery and decreased Ca2+ sensitivity as described under "Results."

Because the Mg²⁺ affinity for the regulatory domain of G34DTnC^F29W is higher than that of TnC^F29W, as the [Mg²⁺] in the fiber increases there should be a larger shift in the Ca²⁺ sensitivity of force for muscle fibers containing G34DTnC^F29W, compared with those containing TnC^F29W. Fig. 7B shows the Ca²⁺-dependent increase in skinned psoas muscle force with TnC^endogenous (), TnC^F29W (), or G34DTnC^F29W () in the presence of 3 mM [Mg²⁺]_free. Half-maximal force occurred at 540 ± 25, 770 ± 18, and 5770 ± 800 nM Ca²⁺ for TnC^endogenous, TnC^F29W, and G34DTnC^F29W, respectively. Thus, in the presence of 3 mM [Mg²⁺]_free the Ca²⁺ sensitivity of force development was ~7.5-fold lower for G34DTnC^F29W compared with TnC^F29W. Furthermore, as the [Mg²⁺]_free was increased from 1 to 3 mM, the Ca²⁺ sensitivity of force development with G34DTnC^F29W was decreased ~3.2-fold. However, under the same conditions the Ca²⁺ sensitivity of force development with TnC^endogenous or TnC^F29W was decreased only ~1.5-fold. These results are consistent with competitive Mg²⁺ binding to the regulatory domains of all the TnC proteins, with the largest effect occurring with G34DTnC^F29W because of its higher Mg²⁺ affinity. Table III summarizes the skinned muscle results.

After TnC was extracted, force was decreased to 5 ± 1% of the maximal force generated with TnC^endogenous (data not shown). TnC^F29W was capable of recovering maximal force to 93 ± 1%, whereas G34DTnC^F29W recovered only 62 ± 5% of the maximal force. It has been demonstrated that partial extraction of TnC can reduce maximal force recovery and decrease the Ca²⁺ sensitivity of force development (61-64). We were concerned that the decreased Ca²⁺ sensitivity of force development generated by G34DTnC^F29W could be explained by its incomplete force recovery and not by its increased Mg²⁺ affinity. Fig. 7C shows that upon partial extraction of TnC^endogenous () to levels of 87, 69, 44, and 37%, maximal force recovery shifted the Ca²⁺ sensitivity of force development 2.1-, 2.5-, 4.4-, and 5.4-fold, respectively. This decrease in Ca²⁺ sensitivity of force development caused by partial extraction of TnC^endogenous was nearly identical to that determined by Brandt et al. (62). A linear regression fit to our data (r² = 0.934) predicted that at the average force recovery observed with G34DTnC^F29W of 62%, the Ca²⁺ sensitivity of force development would be decreased ~3.3-fold due only to incomplete force recovery. Fig. 7C also shows that the average decrease in Ca²⁺ sensitivity of force development for G34DTnC^F29W compared with TnC^endogenous at 1 () and 3 mM () [Mg²⁺]_free was 5.5- and 10.7-fold, respectively. However, after taking into account incomplete force recovery, Mg²⁺ actually decreased the Ca²⁺ sensitivity of force generated by G34DTnC^F29W at 1 and 3 mM ~1.7- and 3.2-fold, respectively, compared with TnC^endogenous. Treating the data similarly for TnC^F29W predicted that the shifts in Ca²⁺ sensitivity of force development with TnC^F29W compared with TnC^endogenous could be entirely explained by incomplete force recovery. Therefore, in the presence of 3 mM [Mg²⁺]_free, the ~3.2-fold shift in Ca²⁺ sensitivity of force development with G34DTnC^F29W was similar to the ~2.0-fold shift in Ca²⁺ affinity of G34DTnC^F29W compared with TnC^F29W either in isolation or in the presence of TnI-(96-148).

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Whether Mg²⁺ competes with Ca²⁺ for the N-terminal regulatory domain of TnC is unresolved (15, 19, 23, 27, 32). Several groups demonstrated that as the [Mg²⁺] was increased, the Ca²⁺ sensitivity of TnC, Tn-activated actomyosin ATPase, and force development decreased (19, 23-31). These results suggested that Mg²⁺ binding to the regulatory domain of TnC, competitive or otherwise, affected its biochemical and physiological properties. We studied Mg²⁺ binding and exchange with the regulatory domain of TnC and its mutant, utilizing the Trp fluorescence of TnC^F29W, which possesses a Phe  Trp mutation immediately preceding the first Ca²⁺-binding loop.

We have previously demonstrated that an endogenous Z acid pair in the first EF-hand of CaM^F19W was required for physiologically relevant Mg²⁺ binding to this EF-hand (3). Because the first EF-hand of TnC has a Gly at the +z position, we hypothesized that substitution of Gly with Asp should enable the first EF-hand of TnC to bind Mg²⁺. Indeed, when Gly-34 at the +z position was substituted with Asp, the Trp fluorescence of G34DTnC^F29W increased ~1.4-fold upon addition of Mg²⁺, with a K_d(Mg) ~ 295 µM. This same mutation in the absence of Mg²⁺ led to an ~1.8-fold increase in Ca²⁺ affinity of the regulatory domain. However, in the presence of 3 mM Mg²⁺, the Ca²⁺ sensitivity of G34DTnC^F29W decreased ~6-fold. From the K_d(Ca) obtained in the absence and presence of Mg²⁺, we calculated the K_d(Mg) to be ~588 µM for G34DTnC^F29W. Thus, the measured K_d(Mg) was in good agreement with the calculated K_d(Mg). These experiments demonstrate that introduction of an Asp residue at the +z position in the first EF-hand of TnC^F29W enabled this EF-hand to competitively bind Mg²⁺ with a physiologically relevant affinity. Consistent with our results, introduction of a Z acid pair into the CD site of oncomodulin led to an ~12- and 52-fold increased Ca²⁺ and Mg²⁺ affinity to this site, respectively (65). Similarly, in CaM^F19W, replacement of Asp with Asn at the +z position of the first EF-hand led to an ~5- and 58-fold decreased Ca²⁺ and Mg²⁺ affinity to the N-terminal sites, respectively (3, 8). Thus, it would appear that modification of the +z position in some EF-hand proteins modulates the Mg²⁺ affinity more so than the Ca²⁺ affinity.

Interestingly, the fluorescence of TnC^F29W did not change upon binding Mg²⁺ but increased upon binding Ca²⁺ to the regulatory sites (43, 44, 48). However, we have demonstrated that Mg²⁺ can competitively bind and displace Ca²⁺ from TnC^F29W at the Ca²⁺ dissociation rate of ~350 s¹, with a K_d(Mg) in the range of 2.2-3.8 mM at 15 °C. This low Mg²⁺ affinity of TnC^F29W was consistent with previously calculated values of 0.8-5 mM (averaging to 3.7 ± 0.6 mM) using other fluorescent TnCs in which competitive Mg²⁺ binding was assumed (23-26, 28, 29, 56). We propose that Mg²⁺ binds to the second regulatory Ca²⁺ binding site of TnC^F29W because the Trp residue adjacent to the first Ca²⁺ binding site was unperturbed by the direct Mg²⁺ binding studies. Recent structures of the Ca²⁺/Mg²⁺-binding proteins calbindin D_9K and CaM have demonstrated that Mg²⁺ binding primarily affects only the local structural environment of the cation-binding loop with the effects in CaM primarily occurring in the N-terminal portion of the loop (4, 66, 67). Thus, if Mg²⁺ bound to the first EF-hand of TnC^F29W Trp²⁹ should have been affected, as we have demonstrated for G34DTnC^F29W. Therefore, Mg²⁺ must be binding to the second N-domain EF-hand of TnC^F29W. In support of this conclusion, preliminary experiments in our laboratory utilizing the Trp fluorescence of cardiac TnC^F27W (cTnC^F27W) have demonstrated that direct Mg²⁺ titrations did not affect the fluorescence of this protein, whereas Ca²⁺ binding enhances the fluorescence as previously reported (68). However, as was the case with TnC^F29W, addition of increasing [Mg²⁺] to Ca²⁺-saturated cTnC^F27W reversed the Ca²⁺-induced increase in the Trp fluorescence with a K_d(Mg) of ~3.6 mM. For cTnC^F27W only the second N-terminal EF-hand is functional and appears to bind both Ca²⁺ and Mg²⁺. In any regard, the regulatory domain of TnC under physiological [Mg²⁺] of ~1 mM and resting [Ca²⁺] of ~100 nM (60) would be ~20-30% saturated (whereas G34DTnC^F29W would be 60-70% saturated) and thus is not as Ca²⁺ specific as the current dogma claims. Thus, researchers should consider that interaction of Mg²⁺ with the regulatory domain of TnC may affect the outcome of experimental studies. The reason for the low affinity and competitive Mg²⁺ binding to the regulatory domain of TnC might be to ensure that at resting [Ca²⁺], or slight fluctuations thereof, the muscle would not be activated until neurally stimulated.

Because TnC regulates muscle contraction as a part of the Tn complex, we have also studied Ca²⁺ and Mg²⁺ binding to TnC^F29W and G34DTnC^F29W in the presence of the TnI-(96-148) peptide. The addition of the TnI-(96-148) peptide to TnC^F29W and G34DTnC^F29W produced ~12- and 14-fold increases in their N-domain Ca²⁺ binding affinities, respectively, as has been previously reported for intact TnI (15, 33). In the absence of Mg²⁺, the Ca²⁺ affinity of the G34DTnC^F29W·TnI-(96-148) was 1.7-fold higher than that of the TnC^F29W·TnI-(96-148) complex. However, in the presence of 3 mM Mg²⁺ the Ca²⁺ affinity of the TnC^F29W·TnI-(96-148) and G34DTnC^F29W·TnI-(96-148) complexes decreased ~1.5- and 5-fold, respectively. From the K_d(Ca) obtained in the absence and presence of Mg²⁺, we estimated K_d(Mg) to be ~6 and 0.7 mM, for the TnC^F29W·TnI-(96-148) and G34DTnC^F29W·TnI-(96-148) complexes, respectively. These results suggest that TnI-(96-148) peptide binding increased the Ca²⁺, but not Mg²⁺, affinities of the N-domain sites of TnC^F29W and G34DTnC^F29W. A similar result had been used previously to argue that Mg²⁺ must bind at an auxiliary N-terminal site(s) of TnC^Danz because the Mg²⁺ affinity was not enhanced in the presence of TnI (23). However, Mg²⁺ binding to the N-domain of TnC^F29W, or G34DTnC^F29W, appears not to open the N-terminal hydrophobic pocket of TnC required for TnI binding that subsequently increases the Ca²⁺ affinity. Furthermore, it would appear that competitive Mg²⁺ binding and displacement of Ca²⁺ from the regulatory domain of TnC^F29W and G34DTnC^F29W actually closes the hydrophobic pocket.

Functionally, Mg²⁺ did not cause force development in reconstituted skeletal muscle fibers at 1 or 3 mM [Mg²⁺]_free with G34DTnC^F29W or TnC^F29W, consistent with Mg²⁺ binding not inducing the same structural changes as does Ca²⁺. In the absence of Mg²⁺, the Ca²⁺ affinity of G34DTnC^F29W was ~2-fold higher than that of TnC^F29W, yet the Ca²⁺ sensitivity of force development in the presence of 1 and 3 mM [Mg²⁺]_free with G34DTnC^F29W was decreased compared with TnC^F29W, consistent with competitive Mg²⁺ binding to G34DTnC^F29W. Furthermore, the Ca²⁺ sensitivity of force was decreased only ~1.5-fold with TnC^endogenous and TnC^F29W, but was decreased ~3.2-fold with G34DTnC^F29W as the [Mg²⁺]_free was increased from 1 to 3 mM. These results were also consistent with competitive Mg²⁺ binding to the regulatory domains of TnC^endogenous and TnC^F29W. After taking incomplete force recovery into consideration, in the presence of 3 mM Mg²⁺, G34DTnC^F29W was ~3.2-fold less sensitive to Ca²⁺ compared with TnC^endogenous or TnC^F29W. However, the magnitude of the shift in Ca²⁺ sensitivity could vary depending upon the cause of the reduced force recovery (partial TnC extraction versus substitution by a mutant). In any case, the force produced by G34DTnC^F29W was less sensitive to Ca²⁺ than that of TnC^F29W, consistent with G34DTnC^F29W possessing a higher Mg²⁺ affinity.

Unexpectedly, G34DTnC^F29W recovered only ~62% of maximal force. Our results demonstrated that reduced affinity of the regulatory domain within G34DTnC^F29W for the inhibitory region of TnI was not the cause for the decreased force recovery. However, we cannot rule out the possibility that the conformation of the G34DTnC^F29W·TnI complex was somehow different from that of the TnC^F29W·TnI complex, leading to reduced force. The extent of force recovery by G34DTnC^F29W was similar to that of psoas muscle fibers reconstituted with cTnC (69, 70). The G34D mutation is in the first EF-hand of TnC, which is the same EF-hand in cTnC that does not bind Ca²⁺. It may be that the mechanism(s) that underlie the decreased force recovery of fast twitch skeletal muscle reconstituted with cTnC are the same for G34DTnC^F29W. It has been speculated that the decreased force recovery with cTnC may be due in part to its decreased actomyosin-ATPase rate in reconstituted skeletal muscle systems and lack of Ca²⁺ binding cooperativity because of its single regulatory Ca²⁺ binding site (71). There does appear to be decreased cooperativity of force development with the G34DTnC^F29W reconstituted muscle fibers (Table I). However, a N-helix deleted TnC mutant also displayed decreased actomyosin-ATPase activity but recovered near maximal force in reconstituted psoas muscle (47). Thus, the reason for the lack of maximal force recovery for G34DTnC^F29W remains unresolved.

A major consequence of the high Mg²⁺ affinity and relatively slow Mg²⁺ dissociation rate of G34DTnC^F29W (at a rate comparable with its Ca²⁺ dissociation rate) would be to drastically slow the Ca²⁺ association rate in the presence of physiological Mg²⁺ by a factor of ~4 × 10⁵. Our laboratory is interested in delineating the influence that the Ca²⁺ kinetics of the regulatory domain of TnC has on striated muscle contraction and relaxation. G34DTnC^F29W could be used as a molecular tool to test whether slower Ca²⁺ association rates to the regulatory domain of TnC could slow Ca²⁺-induced rates of muscle contraction.

In conclusion, the data suggests that the second EF-hand, but not the first, of TnC^F29W was able to competitively bind Mg²⁺. Physiologically relevant Mg²⁺ binding could be engineered into the first EF-hand of TnC^F29W by substituting Gly³⁴ with Asp creating a Z acid pair. In the absence of Mg²⁺, the Gly³⁴  Asp mutation also increased the Ca²⁺ affinity of TnC^F29W and the TnC^F29W·TnI-(96-148) complex. However, in the presence of Mg²⁺, the Gly³⁴  Asp mutation decreased the Ca²⁺ affinity of TnC^F29W and the TnC^F29W·TnI-(96-148) complex. Consistent with competitive Mg²⁺ binding to the first EF-hand of G34DTnC^F29W, exchange of this protein into skinned skeletal psoas muscle fibers led to lower Ca²⁺ sensitivity of force development.

	ACKNOWLEDGEMENTS

We thank Warren Erdahl and Clifford Chapman for technical assistance with atomic absorption measurements of our Ca²⁺ stocks, Catalina Alionte for help in preparation of the manuscript, and Dr. Ruth Altschuld for helpful discussion of the data.

	FOOTNOTES

^* This work was supported in part by National Institutes of Health Grants AR20792 (to J. A. R.) and DK33727 (to R. A. A.), by a grant from the Canadian Institutes for Health Research (to L. B. S.), and 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. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

To whom correspondence should be addressed: Dept. of Physiology and Cell Biology, The Ohio State University, 304 Hamilton Hall, 1645 Neil Ave., Columbus, OH 43210. Tel.: 614-292-6137; Fax: 614-292-4888; E-mail: davis.812@osu.edu.

Published, JBC Papers in Press, October 22, 2002, DOI 10.1074/jbc.M208488200

	ABBREVIATIONS

The abbreviations used are: TnC, intact chicken skeletal troponin C; Tn, troponin; TnC^F29W, intact TnC mutant with the Phe²⁹  Trp mutation; G34DTnC^F29W, intact TnC^F29W mutant with the Gly³⁴  Asp mutation; TnC, isolated N-terminal domain representing residues 1-90 of TnC^F29W; cTnC^F27W, intact human cardiac TnC mutant with the Phe²⁷  Trp mutation; TnI, chicken skeletal troponin I; TnI-(96-148), chicken skeletal troponin I peptide corresponding to residues 96-148; CaM, calmodulin; CaM^F19W, CaM mutant with the Phe¹⁹  Trp mutation; pCa, -log[Ca²⁺]; [Ca²⁺]₅₀, [Ca²⁺] that produced half-maximal force; DTT, dithiothreitol; MOPS, 3-(N-morpholino)propanesulfonic acid; Quin-2, 2-((bis(carboxymethyl)amino)-5-methylphenoxy)-methyl)-6- methoxy-8-(bis(carboxymethyl)amino)-quinoline.

	REFERENCES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

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