Cardiac troponin C (TnC) and a site I skeletal TnC mutant alter Ca²⁺versus crossbridge contribution to force in rabbit skeletal fibres

Abstract

We studied the relative contributions of Ca²⁺ binding to troponin C (TnC) and myosin binding to actin in activating thin filaments of rabbit psoas fibres. The ability of Ca²⁺ to activate thin filaments was reduced by replacing native TnC with cardiac TnC (cTnC) or a site I-inactive skeletal TnC mutant (xsTnC). Acto-myosin (crossbridge) interaction was either inhibited using N-benzyl-p-toluene sulphonamide (BTS) or enhanced by lowering [ATP] from 5.0 to 0.5 mm. Reconstitution with cTnC reduced maximal force (F_max) by ∼1/3 and the Ca²⁺ sensitivity of force (pCa₅₀) by 0.17 unit (P < 0.001), while reconstitution with xsTnC reduced F_max by ∼2/3 and pCa₅₀ by 0.19 unit (P < 0.001). In both cases the apparent cooperativity of activation (n_H) was greatly decreased. In control fibres 3 μm BTS inhibited force to 57% of F_max while in fibres reconstituted with cTnC or xsTnC, reconstituted maximal force (rF_max) was inhibited to 8.8% and 14.3%, respectively. Under control conditions 3 μm BTS significantly decreased the pCa₅₀, but this effect was considerably reduced in cTnC reconstituted fibres, and eliminated in xsTnC reconstituted fibres. In contrast, when crossbridge cycle kinetics were slowed by lowering [ATP] from 5 to 0.5 mm in xsTnC reconstituted fibres, pCa₅₀ and n_H were increased towards control values. Combined, our results demonstrate that when the ability of Ca²⁺ binding to activate thin filaments is compromised, the relative contribution of strong crossbridges to maintain thin filament activation is increased. Furthermore, the data suggest that at low levels of Ca²⁺, the level of thin filament activation is determined primarily by the direct effects of Ca²⁺ on tropomyosin mobility, while at higher levels of Ca²⁺ the final level of thin filament activation is primarily determined by strong cycling crossbridges.

In striated muscle, contraction is initiated by Ca²⁺ binding to troponin (Tn), specifically at the Ca²⁺-binding subunit of Tn (TnC), ultimately enabling strong myosin binding to actin, force generation and shortening. The level of force is controlled by the availability of strong myosin binding sites on the thin filament (the ‘activation’ level) that is in turn, controlled by the position of tropomyosin (Tm) on the surface of actin. As reviewed in detail by Lehrer & Geeves (1998) and more recently by Gordon et al. (2000), the inhibitory subunit of Tn (TnI) binds to actin in the absence of Ca²⁺, anchoring Tm in a position where myosin binding is inhibited. When Ca²⁺ binds to TnC, it induces strong TnI–TnC interactions and weakens the TnI–actin interactions, resulting in increased mobility of Tm and exposure of strong myosin binding sites on actin. In addition, myosin binding to actin (crossbridge formation) is thought to further displace Tm from blocking positions and is necessary for maximal activation of thin filaments in skeletal muscle (Lehman et al. 2000). However, the relative contribution of Ca²⁺ binding to TnC versus strong crossbridges to activation is unknown.

There is some structural (Lehman et al. 2000) and biochemical (Lehrer & Geeves, 1998) evidence of the relative role of strong crossbridges and TnC Ca²⁺ binding in thin filament activation, but few studies have looked at these contributions in fibre preparations. Moss et al. (1985, 1986a) found that partial extraction of native TnC reduced maximal force, Ca²⁺ sensitivity and the apparent cooperativity of the force–pCa relation, but they did not independently manipulate strong crossbridges. Swartz & Moss (1992) reported that exogenous non-cycling strong crossbridges (NEM-S1) elevated force at low [Ca²⁺]. Fitzsimons et al. (2001) extended this study by using NEM-S1 to enhance crossbridge formation after partially extracting native TnC from muscle fibres. They found that decreased Ca²⁺ sensitivity of force following partial TnC extraction is returned to control values in the presence of 6 μm NEM-S1. However, to determine the relative role of myosin binding in thin filament activation, studies need to be done with cycling crossbridges. Additionally, to understand the relative role of Ca²⁺ in thin filament activation, studies need to be done where the complex interactions that occur between TnI, TnT, Tm and actin are not altered by regions of thin filaments with incomplete Tn complexes. We recently reported reconstitution of Tn complexes in demembranated rabbit psoas fibres with mixtures of native TnC and a TnC mutant that does not bind Ca²⁺ at either N-terminal site (Regnier et al. 2002). This reconstitution dramatically decreases the number of activatable Tn complexes and maximal force (F_max), eliminates most of the apparent cooperativity (n_H) of thin filament activation and greatly reduces the Ca²⁺ sensitivity (pCa₅₀) of force (Regnier et al. 2002). An alternative approach is to reduce the ability of each Tn complex to activate the thin filament. Morris et al. (2001) accomplished this by reconstituting skeletal fibre Tn complexes with cardiac TnC, while Sheng et al. (1990) used a site I inactive sTnC mutant. These studies also found a decrease in maximal steady-state force, pCa₅₀ and n_H of the force–pCa relation. However, these studies did not independently manipulate strong crossbridges to determine their role in the activation process.

In the current study we compromised the ability of Ca²⁺ to activate skeletal thin filaments by replacing native TnC with either cardiac TnC (cTnC) or a recombinant sTnC mutant that only binds Ca²⁺ at site II of the N-terminus (like the cardiac isoform). Ca²⁺ binding to site I of the N-terminal region was eliminated by a single amino acid mutation (D28A) at the x position (xsTnC). The crossbridge component of thin filament activation was altered in two different ways. The number of cycling crossbridges and force was decreased with N-benzyl-p-toluene sulphonamide (BTS), which was recently shown to inhibit steady-state force of skeletal fibres by specifically (and reversibly) inhibiting crossbridge formation at low micromolar concentrations (Cheung et al. 2002). Crossbridge interactions were also modified by lowering [ATP] from 5.0 mm to 0.5 mm. We previously reported that this [ATP] decreases crossbridge cycling rate by ∼30%, elevates maximal force slightly, has no significant effect on pCa₅₀ or n_H in skeletal muscle, and does not significantly increases rigor crossbridges (Regnier & Homsher, 1998; Regnier et al. 1998a, 1998b). While rigor crossbridges activate the thin filament in skeletal and cardiac muscle (Metzger, 1995), the long life of actomyosin interaction creates an artificial condition that may not be relevant in the normal activation processes. Measurements of the force–pCa relation demonstrated that replacement of native TnC with TnC containing a single Ca²⁺ binding site at the N-terminus (cTnC or xsTnC) reduces F_max, pCa₅₀ and n_H. Under these conditions, the sensitivity of force to inhibition by BTS was greater in fibres reconstituted with cTnC or xsTnC than in control conditions. In fibres reconstituted with xsTnC, lowered [ATP] caused a recovery of pCa₅₀ and n_H to control values. Our results indicate that when the Ca²⁺ component of thin filament activation is compromised, thin filament activation by crossbridges is more critical for maintenance of force development. The data also suggest that the main determinant of the overall level of thin filament activation varies, such that at higher levels, it is determined by strong crossbridges and at lower levels by both Ca²⁺ binding to TnC and near-neighbour interactions along the thin filament. A preliminary report of this work was published previously (Moreno Gonzalez et al. 2003).

Methods

Fibre preparation

Single rabbit psoas fibre segments were prepared as previously described (Regnier et al. 2002). Male New Zealand rabbits were housed in the Department of Comparative Medicine at the University of Washington (UW) and were cared for in accordance with the US National Institute of Health Policy on Humane Care and Use of Laboratory Animals. The animals were killed with pentobarbital (120 mg kg⁻¹) administered through the marginal ear vein. All protocols were approved by the UW Animal Care Committee. Isolated fibres were treated with 1% Triton X-100 (v/v) in relaxing solution to remove membranous residue. In most experiments, fibre segment ends were chemically fixed by focal application of 1% glutaraldehyde in H₂O to minimize compliance (Chase & Kushmerick, 1988).

Experimental solutions

Compositions of relaxing and activating solutions were calculated and prepared as previously described (Martyn et al. 1994; Regnier et al. 2002). Solutions were maintained at 0.17 m ionic strength and pH 7.0 at 15°C (experimental temperature), and contained (mm): 5 MgATP or 0.5 MgATP, 15 phosphocreatine (PCr), 15 EGTA, at least 40 Mops, 1 free Mg²⁺, 135 Na⁺+ K⁺, 1 dithiothreitol (DTT), at least 250 units ml⁻¹ creatine kinase (CK), 4% w/v Dextran T-500. Ca²⁺ levels (given as pCa = −log[Ca²⁺]) were established by varying the amount of calcium (propionate)₂. In some solutions, 1–40 μm N-benzyl-p-tholuene sulphonamide (BTS) (Sigma) was added as indicated.

Preparation of mutant rabbit xsTnC and native sTnC and cTnC

The rabbit sTnC gene was cloned as previously described for rat cardiac cTnC (Dong et al. 1996). The mutation of xsTnC was introduced at the x coordinating position of the low-affinity, N-terminal Ca²⁺ binding site I (D28A) by site-directed mutagenesis using T7-GEN In Vitro Mutagenesis Kit (USB, Cleveland, OH, USA). In our previous report we introduced single amino acid substitutions at the x position of both binding sites I and II (Regnier et al. 2002) but the amino acid positions were labelled as D27A, D63A. In this report the single site I mutation was introduced at the same x coordinating position of binding site I, and here the amino acid position is labelled D28A as determined by sequence analysis (University of Washington Department of Biochemistry DNA Sequencing Facility). The protein was extracted and purified from bacterial cells as described for rat cardiac cTnC (Dong et al. 1996). Native rat cTnC was purified according to Dong et al. (1996). Rabbit skeletal TnC was purified according to Potter (1982). The purity of native sTnC, xsTnC and cTnC was assessed by SDS-PAGE and protein concentration was determined by UV spectroscopy and Bio-Rad protein assay.

Data acquisition

Mechanical measurements on individual fibres were performed using two sets of apparatus as previously described (Regnier et al. 2002). In all experiments, sarcomere length was set initially to 2.5 μm using a He–Ne laser. For most experiments fibres were periodically (every 5 s) unloaded by rapid (10 L_F s⁻¹) 15% release of total fibre segment length (L_F) for ∼40 ms, followed by rapid restretch to L_F (Brenner, 1983; Sweeney et al. 1987; Chase & Kushmerick, 1988). This procedure maintains structural and functional integrity of muscle fibre segments. The resulting force transients are evident as vertical lines in Fig. 1. Force and L_F signals were digitized and analysed as previously described using custom data acquisition software (Chase et al. 1994; Regnier et al. 2002). Isometric force was measured just prior to the release/restretch mechanical transient. In the subgroup of unfixed fibres, force was estimated from the chart records. Fibre stiffness was measured from small amplitude (0.05%) high frequency (1000 Hz) sinusoidal oscillations as previously described (Regnier & Homsher, 1998).

Figure 1. Example chart record of extraction–reconstitution protocol in a single permeabilized rabbit psoas muscle fibre.

Force records for an example fibre prior to extraction of endogenous TnC (A) and after reconstitution with 100% xsTnC (A and B). Following determination of F_max (force at pCa 4.5), Ca²⁺ concentration (pCa) was varied as indicated below the force records. Force transients occur every 5 s because of ramp release–restretch cycles (see Methods). Cyclical extraction incubations (see Methods) were performed until pCa 4.5 force was reduced to 1.8% of F_max (** in A) for extraction of TnC. Tn complexes were then reconstituted with xsTnC until pCa 4.5 force no longer increased (§ in A) to assure all Tn sites were filled. Initial maximal force (F_max) was 312.8 mN mm⁻² and fibre dimensions were 1.8 mm (length) and 59 μm (diameter). Maximal force after xsTnC reconstitution (rF_max) decreased to 32% of F_max. Force scale was increased after determination of rF_max for better visualization of pCa curve (B). Scale bar: 62 mN mm⁻² (A) and 33 mN mm⁻² (B) (y-axis) and 68 ms (x-axis).

Maximal Ca²⁺-activated force (F_max) was determined at pCa 4.5 or 4.0. Passive force was determined at pCa 9.0 and was subtracted from total force at the various pCa levels to obtain Ca²⁺-activated force. Force was normalized to fibre cross-sectional area in the preparations with fixed ends and was calculated from the diameter assuming circular geometry. For these fibres (n = 22), diameter was 57.8 ± 1.3 μm and F_max was 318.9 ± 19.1 mN mm⁻² prior to extraction of native sTnC. Most fibres included in this study had < 10% rundown of F_max. A few fibres with > 10% (but < 15%) rundown showed the same trend in the data and were included.

Extraction of troponin C and reconstitution of Tn complexes

TnC was selectively extracted from fibres as previously described using trifluoperazine (TFP) extracting solution (Regnier et al. 1999, 2002). Fibres were placed in extracting solution for 30 s followed by 10 s in pCa 9.0 solution. This procedure was repeated in sets of five, F_max was then tested and the procedure repeated until F_max was nearly or completely eliminated. This took an average of 9 min in the extracting solution. Residual force (n = 56) was 1.0 ± 0.2% of pre-extracted F_max.

Reconstitution of Tn complexes with cTnC or xsTnC was achieved by 2 or 3 min incubations in 1 mg ml⁻¹ (total) TnC in pCa 9.2 without CK or Dextran (Hannon et al. 1993; Regnier et al. 1999). Reconstitution was considered complete when maximal force (rF_max) no longer increased with subsequent incubations, which occurred within 4–9 min of incubation (Fig. 1). An additional 1 min incubation with purified sTnC (1 mg ml⁻¹) in a subset of fibres (n = 17) resulted in minimal force increase suggesting incubations with cTnC or xsTnC reconstituted most or all Tn complexes (data not shown). In a separate group of fibres (n = 12) Tn complexes were reconstituted by 1–6 min incubation with purified sTnC and force recovered to near pre-extracted F_max (0.94 ± 0.03) (data not shown). In separate set of experiments we determined that the extraction procedure eliminates the band associated with sTnC on SDS gels, and we have also verified the appearance of a cTnC band after the reconstitution protocol (data not shown). For xsTnC reconstituted fibres the band corresponding to xsTnC migrates to a similar location as native sTnC, probably because there is only a single amino acid difference between these two proteins.

Curve fitting and statistical analyses

BTS titration curves were fitted to equation 1 (Cheung et al. 2002) and the IC₅₀ (K_i, [BTS] that inhibits force to 50% of pre-inhibited level) was estimated from the fitted curve.

(1)

Similarly, for each fibre the force–pCa relation was fitted to the Hill equation:

(2)

using non-linear least squares regression analysis (SigmaPlot). Individual fibre force was normalized to F_max or rF_max for estimation of the regression parameters (pCa₅₀ and n_H). Reported pCa₅₀, n_H, F_max and rF_max values for each group are means ± s.e.m. from fits of individual fibre data. Comparisons were made using paired t tests and are considered significant for P < 0.01. For easier visualization, plots are presented as the average of each data point fitted to the Hill equation.

Results

Replacement of native sTnC with either cTnC or xsTnC in psoas fibres

To study the effects of cTnC and xsTnC replacement on force development, endogenous TnC was extracted from single psoas fibres, and the troponin (Tn) complexes reconstituted with either protein. For the example muscle fibre in Fig. 1, following measurement of maximum force (F_max) a control force–pCa curve was obtained (Fig. 1A). Native TnC was then extracted and the residual force at pCa 4.5 was 1.8% of F_max (** in Fig. 1A) (average residual force for all fibres was 1% of F_max – see Methods). The fibre was then incubated 2 × 2 min in solution containing xsTnC (1.0 mg ml⁻¹ protein) to reconstitute Tn complexes. Further incubations did not increase force at pCa 4.5, therefore reconstitution was considered complete (§ in Fig. 1A). Reconstituted F_max (rF_max) was 32% of control (pre-extracted) F_max. A second force–pCa curve was then obtained for comparison with control conditions (Fig. 1B). For this example fibre, Ca²⁺ sensitivity (pCa₅₀) was reduced from 6.13 ± 0.01 to 5.92 ± 0.02 and the Hill coefficient (n_H) decreased from 3.7 ± 0.3 to 2.0 ± 0.2, which was representative of the values obtained for all fibres reconstituted with xsTnC. The protocol described for Fig. 1 was performed for most experiments. For some fibres a force–pCa curve in the presence of 3 μm BTS or reduced [ATP] (0.5 mm) was obtained following the control force–pCa curve and/or following the force–pCa curve obtained after Tn reconstitution.

The data for fibres reconstituted with cTnC or xsTnC are summarized in Fig. 2, showing that maximal Ca²⁺-activated force, pCa₅₀ and n_H are compromised. In fibres reconstituted with cTnC (n = 15), rF_max was 0.71 ± 0.01 of control F_max (Fig. 2A), pCa₅₀ decreased from 6.01 ± 0.04 to 5.81 ± 0.04 (P < 0.001), and n_H was reduced from 3.8 ± 0.4 to 2.2 ± 0.2 (P < 0.001) (Fig. 2C). Similar results have been previously reported by other groups using cTnC (Moss et al. 1991; Morris et al. 2001, 2003). There are a few reports where force was reconstituted close to 100% with cTnC, but in these studies native TnC was only partially extracted (Moss et al. 1986b, 1991; Metzger, 1996). In an independent set of fibres, we studied the effect on rF_max of complete versus partial TnC extraction to ∼30% of F_max. We found that reconstitution with cTnC was significantly lower in fibres where native TnC was fully extracted (72% of F_max(n = 9)), compared with partially extracted fibres (86 ± 1% of F_max (n = 5, data not shown)) (P < 0.01), which was close to rF_max for fibres reconstituted with purified sTnC (94 ± 3% of F_max, see Methods). These results and our control experiments where cTnC reconstituted fibres were incubated in 1 mg ml⁻¹ sTnC (see Methods) indicate that most or all Tn complexes had bound TnC following the reconstitution protocol. Therefore, compromises in force after cTnC reconstitution mostly result from a reduced ability of Ca²⁺ bound to cTnC to activate the thin filament. This may be due to differences in cTnC–sTnI–sTnT interactions, as previously suggested (Piroddi et al. 2003).

Figure 2. Effects of cTnC or xsTnC reconstitution on thin filament activation.

Summary of force–pCa data, normalized relative to pre-extracted F_max (•, control), for fibres reconstituted with cTnC (▪, 9 fibres) (A) and xsTnC (▾, 10 fibres) (B). Reconstitution with cTnC reduced F_max by ∼28% and xsTnC reduced F_max by ∼80%. Values are shown as mean ±s.e.m. and data were fit by the Hill equation (lines). In C and D, data from A and B were normalized relative to F_max for each condition to visualize changes in pCa₅₀ and n_H (see text for values). Both cTnC and xsTnC reconstitution significantly decreased Ca²⁺ sensitivity and slope of the force–pCa relation.

In fibres reconstituted with xsTnC (n = 16), rF_max was 0.21 ± 0.02 of control F_max (Fig. 2B), pCa₅₀ decreased from 6.11 ± 0.01 to 5.86 ± 0.03 (P < 0.001), and n_H was reduced from 3.1 ± 0.2 to 1.9 ± 0.1 (P < 0.001) (Fig. 2D). Sheng et al. (1990) used a chicken isoform of xsTnC and reported similar reconstituted F_max and a greater decrease in pCa₅₀ and n_H. Interestingly, the ability to activate fibres maximally with saturating [Ca²⁺] was compromised to a greater extent by the skeletal site I mutant than by cTnC, while both manipulations reduced the Ca²⁺ sensitivity and apparent cooperativity of force by a similar amount.

Inhibition of isometric force with N-benzyl-p-tholuene sulphonamide (BTS)

To study if isometric force is more affected by reduced strong crossbridges when the Ca²⁺ binding component of thin filament activation has been compromised, we compared the effect of BTS on steady-state force prior to extraction of native sTnC and following reconstitution with cTnC or xsTnC. Maximal Ca²⁺-activated force was inhibited hyperbolically as [BTS] was increased from 0 to 40 μm, and force recovered to 0.89 ± 0.02 of pre-inhibited levels (n = 28) following BTS wash out. Under control (pre-extracted) conditions, BTS inhibited force with an IC₅₀ ([BTS] for 50% inhibition) of ∼5 μm (Fig. 3A), similar to previously reported values (Cheung et al. 2002). Interestingly, BTS suppressed force (normalized to rF_max for each condition) to a much greater extent at each [BTS] in either cTnC or xsTnC reconstituted fibres (Fig. 3A), compared with control measurements. For example, at 5 μm BTS level, force was inhibited to ∼11% and 9% of pre-inhibited force with cTnC and xsTnC, respectively, compared with ∼50% for control conditions. A proportional decrease of stiffness and force as [BTS] was increased, both under control conditions and when native TnC was replaced by cTnC (Fig. 3B), suggests that force inhibition resulted from a decreased number of strongly bound crossbridges. To determine if the extraction–reconstitution procedure affected BTS inhibition, a separate set of fibres (n = 3) were reconstituted with purified sTnC. The BTS titration performed after sTnC reconstitution was only slightly steeper (IC₅₀= 3.2 μm) (data not shown) than under control conditions (IC₅₀= 4.6 μm). This indicates the much greater force inhibition by BTS with cTnC or xsTnC resulted primarily from a reduced ability of Ca²⁺ to maintain thin filament activation.

Figure 3. BTS inhibition of maximum force and stiffness.

A, BTS titration of maximum force (F_max) prior to extraction of endogenous TnC (•, control), and after reconstitution with either cTnC (▪, 7 fibres) or xsTnC (▾, 3 fibres) (some of the symbols overlap). Since each individual fibre showed similar inhibition curve under control conditions, pre-extracted data of both fibre groups was pooled together for ease of viewing. Data were fitted to eqn (1) (see Methods) and IC₅₀ estimated values are 4.6 μm for control conditions (continuous line), 1.1 μm for cTnC (long dash line), and 1.0 μm for xsTnC (short dash line). Inset bar graph shows F_max relative to control F_max, both in the absence of BTS (filled bar) or presence (open bar) of 3 μm[BTS] for each condition. B, stiffness–force relation as [BTS] was increased to inhibit force. Under both conditions (•, control and ▪, cTnC), stiffness decreased linearly as force decreased due to inhibition by BTS. Data were fitted to a straight line for each condition (control, continuous line; cTnC, long dash line). Values were normalized relative to maximum force (or stiffness) in the absence of BTS for each condition and are presented as mean ± s.e.m.

To study whether crossbridge inhibition affects the Ca²⁺ dependence of force, a single concentration of 3 μm BTS was selected because it significantly inhibited maximal Ca²⁺-activated force (Fig. 3A, inset) yet it was still possible to measure force at low [Ca²⁺] in cTnC or xsTnC reconstituted fibres. Under all conditions 3 μm BTS inhibited force at all levels of Ca²⁺ activation, but this inhibition was much greater following reconstitution with either cTnC or xsTnC. Figure 4 shows that BTS reduced the Ca²⁺ sensitivity of force (pCa₅₀) under control conditions (0.26 pCa unit), but this effect was diminished for fibres with cTnC (0.15 pCa unit) and completely eliminated for fibres with xsTnC (Table 1). Interestingly, either BTS inhibition or replacement of native TnC with cTnC or xsTnC shifted the force–pCa curve to the right by ∼0.20 pCa unit; however, these effects were not additive. The apparent cooperativity of force (n_H) was significantly increased by BTS under control conditions, but the effect was eliminated when n_H was already diminished by reconstitution with cTnC or xsTnC (Fig. 4 and Table 1). Taken together, these results indicate that when the Ca²⁺ component of thin filament activation is compromised, crossbridge inhibition with BTS has a greater effect on maximal force, but little or no further effect on the Ca²⁺ dependence of force.

Figure 4. Effect of 3 μM BTS on the force–pCa relation in fibres prior to extraction of endogenous TnC and after reconstitution with cTnC or xsTnC.

Summary of force–pCa data for a subgroup of fibres in the absence (filled symbols) and presence (open symbols) of 3 μm BTS. Under pre-extracted conditions (control) (circles, A) 3 μm BTS decreased pCa₅₀ by ∼0.26 pCa unit, while after reconstitution with cTnC (5 fibres, squares, B) the effect of BTS was only an ∼0.15 pCa unit decrease or disappeared after reconstitution with xsTnC (4 fibres, downward triangles, C) (see Table 1 for values). Data were normalized relative to F_max for each condition to visualize changes in pCa₅₀ and n_H. Since the regression parameters of the pre-extracted condition (control) with and without BTS of both subgroups were not statistically significant, control data for both fibre groups were pooled together for ease of viewing. Values are shown as means ±s.e.m. and data were fitted to the Hill equation (lines).

Table 1.

Effect of reducing force-producing crossbridges with BTS on the force–pCa relation for a subgroup of fibres

	Relative Fmax	pCa50	nH
Control (9 fibres)
No BTS	1	6.12 ± 0.02	3.2 ± 0.2
3 μm BTS	0.57 ± 0.02	5.86 ± 0.02*	4.7 ± 0.4*
cTnC (5 fibres)
No BTS	0.68 ± 0.01	5.94 ± 0.02	1.9 ± 0.1
3 μm BTS	0.06 ± 0.01	5.79 ± 0.04*	2.3 ± 0.2
xsTnC (4 fibres)
No BTS	0.28 ± 0.03	5.91 ± 0.02	2.1 ± 0.2
3 μm BTS	0.04 ± 0.01	5.87 ± 0.06	1.7 ± 0.8

Values are average of individual regression parameters. Regression parameters of two subgroups of fibres are presented: one group of cTnC reconstituted fibres and one group of xsTnC reconstituted fibres. Since the regression parameters of the pre-extracted condition (Control) for both subgroups were not significantly different, control data for both fibre subgroups was pooled together. Relative F_max represents force relative to that under control conditions without BTS.

^*P < 0.01versus the same parameter measured in the absence of BTS under the same condition: control or reconstituted. All values are given as mean ± s.e.m.

Effect of lowering [ATP] on thin filament activation

We next tested whether slowing crossbridge detachment by lowering [ATP] affects thin filament activation when the Ca²⁺ component has been greatly compromised following reconstitution with xsTnC. Lowering [ATP] from 5.0 mm to 0.5 mm allowed us to slow cycling rate without a significant increase in rigor crossbridges (Regnier & Homsher, 1998; Regnier et al. 1998a). We previously reported that lowering [ATP] from 5.0 mm to 0.5 mm had no effect on pCa₅₀ or n_H and increased F_max slightly (Regnier et al. 1998b). In this study F_max was also slightly increased under control conditions (1.06 ± 0.01 of F_max, n = 6) by lowering [ATP] from 5 mm to 0.5 mm. Following reconstitution with xsTnC, rF_max was not increased by lowering the [ATP] (0.92 ± 0.05 of 5 mm rF_max, n = 8) (Fig. 5A). However, lowering [ATP] significantly increased pCa₅₀ and n_H, shifting the pCa₅₀ from 5.92 ± 0.02 to 6.06 ± 0.02 and increasing n_H from 1.9 ± 0.1 to 2.9 ± 0.2 (n = 8, P < 0.001) (Fig. 5B). These values were close to control conditions where pCa₅₀ was 6.12 ± 0.02 and n_H 2.9 ± 0.02 indicating that 0.5 mm ATP restores the Ca²⁺ sensitivity and apparent cooperativity of force lost by reconstitution with xsTnC. This contrasts with the effect of crossbridge inhibition on pCa₅₀, where BTS caused a right-ward shift of this value under control conditions but had no effect on pCa₅₀ after reconstitution with xsTnC.

Figure 5. Effect of lowering [ATP] from 5 to 0.5 mM on force–pCa relations in fibres prior to extraction of endogenous TnC and after reconstitution with xsTnC.

Summary of force–pCa data for a subgroup of fibres in the presence of 5.0 mm ATP (filled symbols) or 0.5 mm ATP (open symbols), before endogenous TnC extraction (circles) and after reconstitution with xsTnC (downward triangles, 8 fibres). Values are shown as mean ±s.e.m. and data were fitted to the Hill equation (lines). In A, force was normalized relative to pre-extracted F_max to show the decreased force level after reconstitution with xsTnC with either [ATP]. rF_max with 5.0 mm ATP was 0.17 ± 0.02 of pre-extracted F_max. In B, force was normalized relative to F_max for each condition to visualize changes in pCa₅₀ and n_H. Lowering [ATP] to 0.5 mm after reconstitution with xsTnC increased the Ca²⁺ sensitivity of force (pCa₅₀) by ∼0.14 pCa unit and increased the Hill coefficient by 1 unit (see text for values), bringing the force–pCa relation to overlap with the control curve.

Discussion

The goal of this study was to investigate the relative contribution of Ca²⁺ binding to TnC and strong crossbridges in establishing the level of thin filament activation in rabbit psoas muscle fibres during isometric contraction. The main findings of the present study are that when thin filaments are reconstituted with cTnC or xsTnC: (1) reductions in the Ca²⁺ dependence of force (pCa₅₀ and n_H) are similar for different levels of reconstituted F_max (rF_max), (2) BTS inhibition of strong crossbridges results in greater reduction of rF_max, and (3) at low rF_max (when reconstituted with xsTnC) the Ca²⁺ dependence of force is not further affected by strong crossbridge inhibition with BTS but is enhanced by slowed crossbridge kinetics with decreased [ATP]. Below we discuss how these data provide clues to how Ca²⁺ binding (to thin filaments) and subsequent strong crossbridge formation work together to activate thin filaments and determine the Ca²⁺ dependence of isometric force development in skeletal muscle.

Maximal Ca²⁺-activated force

Replacement of native TnC with cTnC or xsTnC reduced maximal force to different levels (Fig. 2) even though both exogenous proteins have a single regulatory Ca²⁺-binding site. These decreases in maximal Ca²⁺-activated force with cTnC or xsTnC were similar to values previously reported by Morris et al. (2001, 2003) and Sheng et al. (1990), respectively. There are at least two potential explanations for this difference. First, the mutation of site I in xsTnC could cause a proportionately greater decrease in Ca²⁺ affinity at site II, if Ca²⁺ binding to the regulatory sites of sTnC involves cooperative interaction between sites I and II (Gagne et al. 1995). In addition, since cTnC has a single site in the native isoform, and it has been demonstrated that strong crossbridge formation enhances Ca²⁺ binding to site II in cardiac muscle (Fuchs & Wang, 1996; Martyn & Gordon, 2001; Martyn et al. 2001), it is possible that crossbridges cause an increase in Ca²⁺ binding to site II of cTnC compared with site II on xsTnC in reconstituted skeletal fibres. In this study we did not monitor Ca²⁺ binding to TnC and thus from the experiments presented here, we cannot conclude if this explains the difference in rF_max between cTnC and xsTnC reconstituted fibres. Second, the site I mutation in xsTnC could alter interactions with sTnI or sTnT and impair transmission of the Ca²⁺-binding signal to Tm to a greater extent than cTnC (Piroddi et al. 2003), even though the C-terminal end of xsTnC is the same as for native sTnC. There are several differences in the amino acid sequences of sTnC and cTnC that suggest the tertiary structure of site I of TnC may be very different among sTnC, cTnC and xsTnC. This could influence interactions of the protein with TnI when Ca²⁺ binding to the N-terminal end occurs, and could account in part for the differences in steady-state reconstituted force. Takeda et al. (2003) recently reported the crystal structure of troponin and on the interactions of the C-terminal end of TnC with TnT and TnI. Of the seven residues of TnC reported to be interacting with TnT and TnI, only one is different (a conservative substitution) between sTnC and cTnC. In contrast, the coordinating residues of site I are significantly different between sTnC and cTnC. Moyes et al. (1996) suggested that the V28 insertion in the cardiac isoform might contribute to the inactivation of site I, since it is located right before the coordinating residues. This also corresponds to the site of our D28A mutation in skeletal TnC (xsTnC) that eliminates Ca²⁺ binding at site I, and suggests this site may be involved in determining exposure of the hydrophobic patch on the N-terminus of TnC and/or the strength of TnC–TnI interaction. Whichever mechanism is responsible for the relative degrees of thin filament activation with cTnC and xsTnC, these proteins allow us to establish conditions where the Ca²⁺ component of thin filament activation is compromised to varying degrees, to study the role of strong cycling crossbridges in thin filament activation.

The steady-state level of force is ultimately determined by the equilibrium distribution of the Tm position on the surface of thin filaments between those that inhibit acto–myosin interaction (‘blocked’), those that allow a weak interaction (‘closed’) and those that allow strong acto–myosin interaction and force (‘open’) (Lehrer & Geeves, 1998). Lehrer & Geeves (1998) suggested that the dominant effect of Ca²⁺ binding to TnC is to shift the equilibrium of thin filament states from the ‘blocked’ towards the ‘closed’ state, with strong crossbridge formation favouring the transition from 'closed' to ‘open’. In this model, strong crossbridges act as allosteric activators of the thin filaments. In demembranated muscle fibres this is manifested in the steady-state force–Ca²⁺ relation which has a high degree of apparent cooperativity (n_H). Additional recent evidence comes from the in vitro protein mechanical assays where the force produced by skeletal heavy meromyosin (HMM) against reconstituted thin filaments is much greater than with actin alone (Homsher et al. 2000; Clemmens & Regnier in press). Given this interpretation, our results (Fig. 2) would suggest that cTnC and xsTnC limit the extent of Ca²⁺-induced tropomyosin movement to the ‘closed’ state, thereby diminishing the probability of transition to the ‘open’ state that allows force generation (even at maximally activating [Ca²⁺]). In this context, the greater F_max with cTnC relative to xsTnC (Fig. 2) may reflect a greater occupancy of the ‘closed’ state for a given level of Ca²⁺ binding than with xsTnC and therefore a greater probability of force generation in the ‘open’ state.

Even though rF_max was almost 3-fold less for fibres reconstituted with xsTnC versus cTnC (Fig. 3A, inset), the relative effect of BTS on rF_max was similar over the entire range of [BTS] tested (Fig. 3A). If strong crossbridge formation results in an allosteric enhancement of the transition of thin filaments from the ‘closed’ to ‘open’ states in the presence of Ca²⁺, as proposed by Lehrer & Geeves (1998), our data could suggest that when the strong crossbridge component of thin filament activation is greatly reduced, the portion of thin filament sites in the ‘open’ state would also be reduced and depend only on the [Ca²⁺]. In other words, there may be some critical level of the Ca²⁺ component of activation required before strong cycling crossbridges can contribute significantly to thin filament activation. This idea is supported by our comparison of BTS effects on muscle fibres reconstituted with cTnC versus xsTnC. In fibres reconstituted with cTnC (rF_max= 0.68 control F_max), 3 μm BTS caused a further decrease in the Ca²⁺ sensitivity of force (pCa₅₀) (Fig. 4; Table 1). In contrast, in fibres reconstituted with xsTnC (rF_max= 0.28 control F_max), pCa₅₀ was not further affected by force inhibition with 3 μm BTS. These observations suggest that impairing the ability of Ca²⁺ to activate the thin filament below a critical level, somewhere between 25 and 70% of control F_max, greatly diminishes the cooperative mechanism of crossbridge-induced increase in crossbridge formation. Below this threshold, strong acto–myosin interactions may have little effect on the level of thin filament activation (i.e. tropomyosin position), and at this point the level of activation is determined primarily by Ca²⁺ binding to TnC. The fact that n_H is still > 1.0 under any condition (Table 1), suggests that some form of cooperative activation still occurs. This could be a crossbridge-induced increase in TnC Ca²⁺ binding or some minimal level of crossbridge-induced increase in crossbridge formation. However, further experiments need to be done to test this hypothesis.

Interestingly, the reduction in maximal Ca²⁺-activated force for fibres reconstituted with cTnC or xsTnC, and inhibited with 3 μm BTS is greater than the fractional reduction of each manipulation combined (Fig. 3– inset). In other words, the effect of Ca²⁺ binding to TnC and strong crossbridge formation at saturating [Ca²⁺] are more than additive in the activation of skeletal muscle fibre thin filaments. Therefore these data support the biochemical model of crossbridges as allosteric activators (Lehrer & Geeves, 1998), as well as the electron microscopy observations of Holmes et al. (1993), and more recently Lehman et al. (2000). Here we have shown the first direct demonstration of this cooperative relationship between the two components of thin filament activation in skeletal muscle fibres.

Ca²⁺ dependence of force

The Ca²⁺ sensitivity of force (pCa₅₀) was decreased to a similar extent by 3 μm BTS under control conditions (Fig. 4A) and following native TnC replacement with cTnC or xsTnC (Fig. 2), suggesting that both the Ca²⁺ component and the crossbridge component of thin filament activation play a role in establishing the Ca²⁺ sensitivity of force. We recently reported that replacing native TnC in rabbit psoas muscle fibres with mixtures of sTnC and a recombinant sTnC mutant that does not bind Ca²⁺ at N-terminal sites I and II (xxsTnC), allows progressive decreases in the number of ‘activatible’ Tn complexes. This results in a reduction of near-neighbour functional regulatory unit interactions along the thin filament, without compromising the remaining ‘activatible’ Tn complexes (Regnier et al. 2002). As the number of functional regulatory units was decreased, there was a progressive decrease in the Ca²⁺ sensitivity of force, but the apparent cooperativity of activation (n_H) was not affected until reconstitution mixtures contained ≤ 40% sTnC. In contrast, in the present study we compromised the Ca²⁺-activation properties of all Tn complexes in the thin filament by reconstituting fibres with cTnC or xsTnC. Therefore, a comparison of the force–pCa relation for these two studies is useful in understanding the mechanisms controlling thin filament activation and force development.

Table 2 summarizes pCa₅₀ and n_H values when rF_max was reduced to similar extents by either approach. When comparing fibres reconstituted with 100% cTnC to fibres reconstituted with 60: 40 sTnC: xxsTnC mixtures, the rF_max is similar and we observe a similar decrease in Ca²⁺ sensitivity of force (pCa₅₀). However, the apparent cooperativity of force development (n_H) was greatly reduced in cTnC, but not in 60: 40 sTnC: xxsTnC reconstituted fibres. In comparison, the rF_max of fibres reconstituted with 100% xsTnC was similar to fibres reconstituted with 15: 85 sTnC: xxsTnC mixtures and both groups of fibres have a low n_H, similar to fibres reconstituted with 100% cTnC. However, while the reduction in pCa₅₀ for 100% xsTnC reconstituted fibres was similar to 100% cTnC and 60: 40 sTnC: xxsTnC reconstituted fibres, the reduction of pCa₅₀ in 15: 85 sTnC: xxsTnC fibres was twice as great. Fibres reconstituted with 15: 85 sTnC: xxsTnC probably have a number of functional regulatory units that are isolated from each other (Regnier et al. 2002), suggesting that interaction between near-neighbour regulatory units is important in determining the Ca²⁺ sensitivity of force development.

Table 2.

Effect on Ca²⁺ dependence of force after reconstitution with different TnC or TnC mixtures

Protein (% of each)§	Relative rFmax	▵pCa50	Reconstituted nH†
cTnC (100)	0.71	0.20	2.2
sTnC: xxsTnC* (60: 40)	0.75	0.21	3.4
xsTnC (100)	0.21	0.25	1.9
sTnC: xxsTnC* (15: 85)	0.14	0.49	1.7

^§Percentage of each TnC protein in the reconstitution solution. Relative rF_max represents reconstituted force relative to F_max under pre-extracted conditions (control). ▵pCa₅₀ represents the change in pCa₅₀ between the control and reconstituted condition.

^†n_H under control conditions is similar among all groups (∼3.6). All values are given as mean.

^*Values from (Regnier et al. 2002).

It is likely that the spread of activation along cTnC or xsTnC reconstituted thin filaments by Ca²⁺ binding is reduced as well, and this may explain the reduction of n_H for 100% cTnC, 100% xsTnC and 15: 85 sTnC: xxsTnC, but not 60: 40 sTnC: xxsTnC reconstituted fibres. A reduced spread of activation along thin filaments would suggest a mechanism that explains why crossbridge inhibition with BTS has a reduced effect on pCa₅₀ in fibres reconstituted with 100% cTnC or xsTnC. In control fibres, the spread of activation along thin filaments by Ca²⁺ binding to TnC and subsequent strong crossbridge formation is substantial (Regnier et al. 2002). Reducing the available pool of strong crossbridges to spread activation would mean that a greater contribution of Ca²⁺ is needed to reach the same relative level of contractile activation. However, in cTnC or xsTnC reconstituted fibres, strong crossbridge formation following Ca²⁺ binding has already been reduced, so a further reduction in strong crossbridge formation with BTS has relatively little effect.

Results where [ATP] was reduced from 5.0 to 0.5 mm (Fig. 5) agree well with this idea. Lowering [ATP] slows acto-myosin kinetics of psoas skeletal fibres by decreasing the rate of crossbridge detachment, as suggested by a slower rate of tension redevelopment (k_tr) and unloaded shortening velocity (Regnier et al. 1998b), though maximum force is only slightly elevated (Cooke, 1997; Regnier et al. 1998b). These observations are consistent with the interpretation that lowering [ATP] to 0.5 mm results in some increase in the time crossbridges spend in the rigor state (Regnier et al. 1998b). Rigor crossbridges have been shown to activate thin filaments in skinned skeletal fibres through an allosteric mechanism that may increase Ca²⁺ binding, as evidenced by increasing the amount of Ca²⁺ bound to sTnC (Fuchs, 1985) and by inducing changes in sTnC structure (Martyn et al. 1999). While slowed crossbridge detachment with low [ATP] does not increase the apparent level of thin filament activation in native skinned skeletal fibres, as evidenced by no change in pCa₅₀ or n_H (Regnier et al. 1998b), in xsTnC reconstituted fibres pCa₅₀ and n_H were increased by low [ATP] towards control values (Fig. 5). Combined with the results of 15: 85 sTnC: xxsTnC reconstituted fibres (discussed above), the data suggest that when the Ca²⁺ binding component of thin filament activation is greatly compromised, longer lived cycling strong crossbridges can recover the Ca²⁺ dependence of force development.

In summary, here we found that the activation level of thin filaments in skeletal muscle fibres depends on the balance between two components: Ca²⁺ binding to TnC and strong crossbridge formation. When the Ca²⁺ component of thin filament activation is compromised, the level of activation can be increased by strong cycling crossbridges. Furthermore, the main determinant of Ca²⁺-dependent force depends on the overall level of thin filament activation. At higher levels it is determined by strong crossbridges and at lower levels by both Ca²⁺ binding to TnC and near-neighbour interactions along the thin filament.