Effects of the mutation R145G in human cardiac troponin I on the kinetics of the contraction–relaxation cycle in isolated cardiac myofibrils
1 Department of Vegetative Physiology, University of Cologne, Koeln, Germany
2 Department of Cardiovascular Medicine, University of Oxford, UK
3 Department of Pediatrics, Division of Molecular Cardiovascular Biology, Cincinnati Children's Hospital, Cincinnati, OH 45229-3039, USA
Abstract
Familial hypertrophic cardiomyopathy (FHC) has been linked to mutations in sarcomeric proteins such as human cardiac troponin I (hcTnI). To elucidate the functional consequences of the mutation hcTnIR145G on crossbridge kinetics, force kinetics were analysed in murine cardiac myofibrils carrying either the mutant or the wild-type protein. The mutation was introduced into the myofibrils in two different ways: in the first approach, the endogenous Tn was replaced by incubation of the myofibrils with an excess of reconstituted recombinant hcTn containing either hcTnIWT or hcTnIR145G. Alternatively, myofibrils were isolated either from non-transgenic or transgenic mice expressing the corresponding mcTnIR146G mutation. In myofibrils from both models, the mutation leads to a significant upward shift of the passive force–sarcomere length relation determined at pCa 7.5. Addition of 5 mM BDM (2,3-butandione-2-monoxime), an inhibitor of actomyosin ATPase partially reverses this shift, suggesting that the mutation impairs the normal function of cTnI to fully inhibit formation of force-generating crossbridges in the absence of Ca2+. Maximum force development (Fmax) is significantly decreased by the mutation only in myofibrils exchanged with hcTnIR145G in vitro. Ca2+ sensitivity of force development was reduced by the mutation in myofibrils from transgenic mice but not in exchanged myofibrils. In both models the rate constant of force development kACT is reduced at maximal [Ca2+] but not at low [Ca2+] where it is rather increased. Force relaxation is significantly prolonged due to a reduction of the relaxation rate constant kREL. We therefore assume that the impairment in the regulatory function of TnI by the mutation leads to modulations in crossbridge kinetics that significantly alter the dynamics of myofibrillar contraction and relaxation.
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Introduction
Familial hypertrophic cardiomyopathy (FHC) is a disease characterized by ventricular hypertrophy and interstitial fibrosis, often resulting in arrhythmias, severe cardiac dysfunctions and sudden cardiac death (Marian & Roberts, 1998). FHC is linked to dominant missense mutations in the genes of almost all sarcomeric proteins including the cardiac troponin complex (cTn) (Kimura et al. 1997; Seidman & Seidman, 2001; Fatkin & Graham, 2002). Cardiac Tn is a Ca2+ sensor and regulatory element of the contraction–relaxation cycle in cardiac muscle. It is a heterotrimer consisting of cardiac troponin C (cTnC), the Ca2+-binding subunit, cardiac troponin I (cTnI), the inhibitory subunit that binds to actin and prevents crossbridge cycling at submicromolar [Ca2+] and cardiac troponin T (cTnT), which binds to cTnI, cTnC and tropomyosin (Tm), thereby anchoring the cTn complex to the thin filament. When intracellular [Ca2+] is high during systole, Ca2+ binds to cTnC, which then binds to cTnI with a high affinity, thereby releasing it from its inhibitory site on actin. This Ca2+-induced sequence of protein–protein interactions also leads to a change in the position of Tm on the actin filament, which results in turning actin filaments to an ‘on’ state (Solaro & Rarick, 1998). Relaxation is initiated when [Ca2+] falls to diastolic levels which then, in a reverse process, returns the thin filaments to an ‘off’ state. Central to the inhibitory function of cTnI is the so-called inhibitory region, an evolutionarily conserved 11 amino acid motif in the centre of the molecule (Gordon et al. 2000).
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To date, more than 15 missense mutations on the cTnI molecule have been identified in patients suffering from FHC (Kimura et al. 1997; Seidman & Seidman, 2001). To identify the underlying pathological mechanism of disease development, a number of investigators have studied the effects of FHC-related mutations on actomyosin ATPase and on contractile steady-state properties of skinned fibres and cardiomyocytes (Elliott et al. 2000; Takahashi-Yanaga et al. 2001; Burton et al. 2002; Lang et al. 2002; Westfall et al. 2002; Kohler et al. 2003). The most intensively studied cTnI mutation is located within the inhibitory region at position 145, replacing a positively charged arginine by an uncharged glycine (Kimura et al. 1997). Controversial data for Ca2+ sensitivity of force development aside, in vitro studies consistently show that the mutation R145G leads to an increase in ATPase activity at resting [Ca2+] (Elliott et al. 2000; Takahashi-Yanaga et al. 2001; Lang et al. 2002).
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Most of the pathological manifestations of this mutation observed in the human heart could be reproduced in a transgenic mouse model expressing the corresponding mutation R146G in the murine cTnI (mcTnI) (James et al. 2000). Interestingly, the phenotype of the transgenic mice depends on the amount of expressed mcTnIR146G. If the mutated protein makes up more than 50% of the total mcTnI protein the mice develop cardiomyocyte disarray, interstitial fibrosis and suffer premature death. In contrast, if the mutated protein contributes to the total mcTnI by only 40%, no pathological alterations in cellular and organ morphology were reported. Surprisingly, working heart analyses of these animals revealed impaired isovolumetric relaxation, indicated by a more than 2-fold prolonged lifetime of pressure fall (), a relatively load-independent measure of diastolic function (James et al. 2000). Coexistent to impaired relaxation, systolic function was improved as indicated by significantly increased values of +dP/dt. These data and similar data on transgenic mice overexpressing FHC-related mutations in other sarcomeric proteins (Evans et al. 2000) reflect the clinical observation in FHC patients, which typically exhibit normal and sometimes even supra-normal contractile function, while lusitropy is severely impaired (Kass et al. 2004).
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Studies performed on intact trabeculae revealed that the rate-limiting steps of cardiac contraction and relaxation reside in the myofilaments (Janssen et al. 2002). This makes crossbridge kinetics and altered kinetics of regulatory proteins possible candidates for the altered haemodynamics observed in FHC. To date the effects of FHC mutations in myosin heavy chain, TnT and Tm on crossbridge kinetics have been investigated in skinned fibres where kinetics are derived from force transients induced by mechanical stretch or release (Palmiter et al. 2000). To our knowledge, no study has investigated the effect of FHC-related mutations on force kinetics in a myofibrillar system, in which isometric contraction and relaxation are induced by the physiological activator [Ca2+]. Thus the question of whether the impaired diastolic function observed in FHC might be related to impaired force kinetics during myofibrillar relaxation remains to be answered.
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In this study we investigated the effects of the FHC mutation R145G in cTnI on the force kinetics of isolated myofibrillar bundles, in which contraction and relaxation is induced by changes in [Ca2+] via rapid solution change (Stehle et al. 2002b). We used two different strategies to introduce the mutation into the myofibrils. In one approach, we applied a recently developed biochemical technique to exchange the native Tn in myofibrils with exogenous, recombinant troponin complexes (Kruger et al. 2003). In the second method we investigated force kinetics in cardiac myofibrils isolated from mcTnIR146G transgenic mice. By comparing the data we obtained a detailed analysis of the impact of cTnIR145G on the kinetics of force activation and relaxation. Furthermore, we suggest possible adaptational processes in myofibrillar function that could be manifested during the development of heart failure in the transgenic animals.
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Methods
Preparation of recombinant human cardiac troponin complex
The three human cardiac troponin subunits hcTnCwt, hcTnTwt and hcTnI (either hcTnIwt or hcTnIR145G) were expressed separately in E. coli and purified as previously described (Kruger et al. 2003). The identity of the plasmids was verified by sequencing. The purified subunits were reconstituted to the heterotrimeric hcTn complex (Potter, 1982) that was then stored at 4°C for up to 1 week.
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Isolation of cardiac myofibrils
For the experiments on myofibrils from mcTnIR146G transgenic, adult FVB/N mice in which 40% of the endogenous cTnI was replaced with mcTnIR146G were used (James et al. 2000). For the hcTn exchange experiments myofibrils were prepared from adult Him: OF 1 mice. Mice were killed by cervical dislocation. All experiments were approved by the Institutional Animal Care and Use Committee.
The heart was removed and papillary muscles were dissected from the left ventricle, skinned with 1% v/v Triton-X 100 in rigor buffer (132 mmol l–1 NaCl, 5 mmol l–1 KCl, 1 mmol l–1 MgCl2, 10 mmol l–1 Tris pH 7.1, 5 mmol l–1 EGTA, 1 mmol l–1 NaN3 and a protease inhibitor cocktail) for 2 h and stored at 4°C in rigor buffer without Triton-X 100. Myofibrillar suspensions were prepared immediately before experiments by homogenizing skinned papillary muscles with a blender (Ultra Turrax) for 10 s at 4°C.
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Exchange of hcTn complex in mouse cardiac myofibrils
Replacement of native mcTn by recombinant hcTn was performed with a protocol adapted and modified for cardiac myofibrils (Kruger et al. 2003), which was initially described for skeletal muscle fibres by Brenner et al. (1999) and also for skinned cardiac fibres (Kohler et al. 2003). In brief, reconstituted hcTn complex was dialysed against rigor buffer, centrifuged at 13 000 g for 10 min and filtered through a polypropylene mesh (Millex-HV, 0.45 μm pore, Durapore) to remove aggregates. The freshly prepared myofibril suspension was centrifuged for 5 min at 380 g (10°C), the myofibril pellet resuspended in rigor buffer containing hcTn complex (1 mg ml–1) and incubated at 21°C for 60 min. To remove excess Tn, myofibrils were washed by centrifugation in rigor buffer (5 min at 380 g, 10°C). Exchange of the troponin subunits was confirmed by Western blot analysis as reported previously (Kruger et al. 2003). The protein components were separated on SDS-PAGE, transferred onto a nitrocellulose membrane (Schleicher & Schuell GmbH, Dassel, Germany) by standard tank transfer Western blot and probed by a monoclonal antibody against cTnI (clone 6F9, Dunn Labortechnik GmbH, Asbach, Germany) which also recognizes sTnI and a monoclonal anti-TnT antibody (JLT12, Sigma-Aldrich, Taufkirchen, Germany). The membranes were incubated with anti-mouse IgG-HRP (Sigma-Aldrich, Taufkirchen, Germany) and enzymatic activity was detected using an ECL-kit (Amersham Biosciences, Freiburg, Germany).
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Myofibrillar force measurements
Myofibrillar force kinetics were measured as previously described (Stehle et al. 2002a,b; Kruger et al. 2003). Composition of relaxing (pCa 7.5) and activating solution (pCa 4.5) was as in Stehle et al. (2002b). Using silicone adhesive, small myofibrillar bundles 5–10 μm in diameter and 40–70 μm in length were attached in relaxing solution to a stiff micro-needle at one end and to the tip of an atomic force cantilever at the other end. After mounting, the bundles were stretched to a sarcomere length of 2.3 μm. To avoid possible errors caused by phase contrast, determination of sarcomere length and cross-sectional area (CSA) were performed in bright field. Rapid changes ( 10 ms) in Ca2+ concentration were applied to the myofibrils using the solution change technique described in Stehle et al. (2002b). Force transients were recorded by monitoring the deflection of the cantilever by laser light reflection (Stehle et al. 2002a). All experiments were performed at 10°C.
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Data analysis and statistics
To determine force kinetic parameters, original force transients were fitted by either a mono-exponential function to derive kACT or by a function consisting of a linear and an exponential term to derive kLIN, tLIN and kREL (Stehle et al. 2002b). Statistical analysis was performed by subjecting the data to Student's t test. Significance was determined as *P < 0.05, **P < 0.01 and ***P < 0.001. Unless otherwise noted, all data are given as mean ± S.E.M. (standard error of the mean) of n experiments.
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Results
Incubation of isolated myofibrils with an excess of recombinant hcTn containing either hcTnIR145G or hcTnIWT results in almost complete replacement of the endogenous cTn (see Fig. 1 for Western blot analysis). From these results we conclude that the content of the mutant protein is above 80% in biochemically exchanged myofibrils. It should be noted that the transgenic mice only express about 40% mcTnIR146G with respect to their total mcTnI expression.
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A myofibrillar suspension from mouse papillary muscle was tested for cardiac troponin I. Lane 1: unexchanged control suspension; lane 2: myofibrillar suspension exchanged with hcTn. The molecular weight standard is shown on the right. After exchange no more mcTnI can be detected in hcTn-exchanged myofibrils.
Force development in the absence of Ca2+
Force–sarcomere length relationships were measured by stretching myofibrils (pCa 7.5) from slack sarcomere length of 2.0 μm to defined sarcomere length (SL) ranging from 2.2 to 2.6 μm. In this SL range, regular sarcomere patterns were observed in all myofibrils. In both experimental models, myofibrils containing mutant cTnI showed an increased upward shift of the force–SL relationship compared with either non-transgenic (ntg) (Fig. 2A) or hcTnWT (Fig. 2B). It is noteworthy that the slopes of the force–SL relationships were not altered by the mutant protein as would be expected if passive length-dependent stiffness properties were changed.
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Force is plotted as a function of sarcomere length (SL). A, mean ± S.E.M. of force–SL relationship for myofibrils isolated from mcTnIR146G transgenic mice (filled circles) (n = 16) and the corresponding non-transgenic controls (open circles) (n = 11). B, force–SL relationship of myofibrils exchanged with either hcTnIR145G (filled symbols) or hcTnIWT (open symbols) before (circles) and after (squares) incubation with 5 mM BDM, respectively. Results are representative of 3 independent experiments.
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Addition of 5 mM BDM (2,3-butandione-2-monoxime), an inhibitor of actomyosin ATPase, to the hcTnIR145G-exchanged myofibrils partially abolished the upward shift of the force–SL relationship. In contrast, BDM had no effect in hcTnIWT-containing myofibrils (Fig. 2B). This indicates that the mutation-induced force enhancement in the absence of Ca2+ at least partly results from force-generating crossbridges.
Ca2+ dependence of force development and Ca2+-induced force development kinetics
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Maximum Ca2+-activated force at pCa 4.5 normalized to cross-sectional area (Fmax/CSA) is significantly decreased by 24% in mouse myofibrils exchanged with hcTnIR145G compared with hcTnIWT-exchanged controls (see Table 1). In contrast, no difference in Fmax/CSA was observed for myofibrils isolated from transgenic mice when compared with those isolated from non-transgenic controls. The values for Fmax/CSA are given in Table 1.
The Ca2+ sensitivity of force development was significantly (P < 0.05) lower in myofibrils from the transgenic mice expressing mcTnIR146G (pCa50 5.73 ± 0.03; n = 12) when compared with the non-transgenic control mice (pCa50 5.79 ± 0.01 n = 4, Fig. 3). Cooperativity of force development was significantly reduced (P < 0.0001) as indicated by the reduced Hill coefficient of 6.2 ± 1.4 in myofibrils from the transgenic animals compared with 18 ± 2 of those from the non-transgenic control animals. In contrast, no significant reduction in Ca2+ sensitivity of force development was observed in myofibrils treated with the exchange protocol (data not shown).
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pCa50 is 5.73 ± 0.03 in myofibrils from the transgenic mcTnIR146G mice (n = 12) and 5.79 ± 0.01 in myofibrils from non-transgenic control mice (n = 4) (P < 0.05). Hill coefficients are 6.2 ± 1.4 in myofibrils from the transgenic animals and 18 ± 2 in those from the non-transgenic control (P < 0.0001).
Myofibrils from both models were analysed for their kinetics of force development following a rapid change from pCa 7.5 to pCa 4.5 (e.g. Fig. 4). As shown previously for untreated cardiac myofibrils from different species (Stehle et al. 2002b), rapid increases in [Ca2+] induced a mono-exponential increase in force in myofibrils from both models with a rate constant kACT, whereby kACT increases with increasing [Ca2+] (Fig. 5). We noted that the Ca2+-dependent increase in kACT of cardiac myofibrils exchanged with hcTnIWT (Fig. 5A) is similar to that of myofibrils from the non-transgenic mice (Fig. 5B). This indicates that Ca2+-induced force kinetics of murine myofibrils is not affected by the exchange of their native cTn complex with the recombinant human wildtype isoform. In both models, the mutation caused qualitatively similar changes in the kACT–pCa relationship (Fig. 5A and B): at low [Ca2+] (pCa 5.83), kACT is increased by the mutation, whereas at maximally activating [Ca2+] (pCa 4.5), kACT is significantly reduced from controls by 30% in the hcTnIR145G myofibrils and by 15% in myofibrils isolated from mcTnIR146G transgenic mice. The values for kACT after Ca2+ activation with pCa 4.5 are given in Table 1.
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The left side shows the kinetics of Ca2+-induced force development upon a switch from pCa 7.5 to 4.5. The right panel shows relaxation induced by a switch from pCa 4.5 to 7.5 at t = 1.5 s. The mathematical fit is shown in black, and kinetic parameters are given as kACT, kLIN, tLIN and kREL for each curve. Force transients were normalized to the same amplitude to illustrate the differences in kinetics; force at pCa 7.5 was subtracted before normalization.
A, kACT–pCa relationship for myofibrils exchanged in vitro with either hcTnIR145G (filled circles) or hcTnIWT (open circles) (n = 8–22). B, kACT–pCa relationship for myofibrils isolated from the transgenic mice expressing mcTnIR146G (filled circles) and for myofibrils isolated from the non-transgenic control mice (open circles) (n = 9–18). All data are means ± S.E.M.
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Myofibrillar relaxation kinetics
Rapidly deactivating myofibrils by changing the [Ca2+] from pCa 4.5 to 7.5 induced a biphasic relaxation (see Fig. 4 for typical force transients): an initial linear force decay followed by a fast exponential decay. The slow linear phase has a rate constant kLIN and a duration tLIN (Stehle et al. 2002b) while the exponential phase is described by the rate constant kREL. Figure 6 shows the values obtained for the parameters kLIN and tLIN of the initial force decay. Neither myofibrils exchanged with hcTnIR145G nor myofibrils isolated from mcTnIR146G transgenic mice differed in their values of kLIN compared with their hcTnIWT or non-transgenic controls, respectively (Fig. 6A). In myofibrils isolated from the transgenic mice the duration of the linear phase tLIN was unaltered compared with the corresponding non-transgenic mice (Fig. 6B), whereas myofibrils exchanged with hcTnIR145G showed a 1.7-fold longer tLIN (P < 0.005) than those exchanged with hcTnIWT.
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Shown are means ± S.E.M. obtained for kLIN (A) and tLIN (B) in myofibrils exchanged with hcTnIWT (n = 18) or hcTnIR145G (n = 22) and in myofibrils isolated from transgenic mice expressing mcTnIR146G (n = 14) or non-transgenic control animals (n = 18). *P < 0.05, Student's t test.
Figure 7 shows the values of the rate constant of the fast exponential relaxation phase kREL plotted as a function of the activating pCa. In hcTnIR145G-exchanged myofibrils following maximum activation with pCa 4.5, kREL is decreased by 40% of the values for hcTnIWT-exchanged myofibrils. In myofibrils from the transgenic mice kREL is reduced by 27% compared with controls. In both models kREL was also significantly reduced in relaxations initiated from partial activations with pCa 5.75 and 5.51 (Fig. 7). The values for kLIN, tLIN and kREL in relaxation experiments following maximum Ca2+ activation are summarized in Table 1.
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A, kREL–pCa relationship for myofibrils exchanged in vitro with either hcTnIR145G (filled circles) or hcTnIWT (open circles). B, kREL–pCa relationship for myofibrils isolated from the transgenic mice expressing mcTnIR146G (filled circles) and from the non-transgenic control mice (open circles). All data are means ± S.E.M. for n = 22 (hcTnIR145G), n = 17 (hcTnIWT), n = 18 (mcTnIR146G tg) and n = 14 (ntg control).
Discussion
In this study we analysed the contractile behaviour of cardiac myofibrils isolated from cTnIR146G transgenic mice and of cardiac myofibrils from wild-type mice, in which the endogenous Tn complex had been biochemically replaced with the corresponding human mutation hcTnIR145G. Common to both models we find an upward shift of the passive length–tension relationship, at low concentrations of Ca2+ an increase and at high concentrations of Ca2+ a decrease in the rate of contraction kACT, as well as a decrease in the rate of the fast exponential phase of the relaxation. Differences between the two models were observed for Ca2+-activated steady-state force and the duration of the linear phase of relaxation, which is prolonged in the exchanged but not in myofibrils taken from the transgenic mice. Several factors may account for the differences: (i) the exchanged myofibrils contained the human troponin isoform which despite its high homology with the mouse troponin isoform may nevertheless lead to subtle changes in the protein–protein interactions within the mouse sarcomeric environment; (ii) lower dosage of the mutant cTnI in the myofibrils taken from the transgenic mouse compared with the exchanged myofibrils; (iii) effect of the transgene on the expression of the myosin heavy chain or other sarcomeric proteins.
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We find it unlikely that the differences are due to the hybrid system for the following reasons: in recent studies we compared steady-state and kinetic contractile parameters in native murine myofibrils with those exchanged with hcTn and with native human myofibrils (Stehle et al. 2002b; Kruger et al. 2003). The kinetic parameters were an order of magnitude higher in the native as well as in the exchanged murine myofibrils compared with the native human myofibrils indicating that the kinetic parameters are not sensitive to the Tn isoform. Fmax/CSA was lower in the exchanged myofibrils as also seen in this study. At present we do not know whether this is due to the exchange procedure per se or is a reflection of the hybrid system. Irrespective of the cause of the depression in force, it is larger in myofibrils exchanged with hcTnIR145G indicating that this mutation has a specific effect on Fmax/CSA. Therefore we propose that in the transgenic myofibrils the gene dosage may be too low to affect Fmax/CSA.
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The impact of transgenic expression of R145G on myosin isoform content has been explored at both the RNA levels, via - and myosin heavy chain-specific oligonucleotide probes (Ng et al. 1991) and by titrating the two proteins directly using a general and myosin-specific antibody (Krenz et al. 2003). Transcript levels were up slightly (less than 10%) but no detectable changes in the relative or absolute levels of the myosin isoforms were detected. We therefore conclude that altered expression of MHC isoforms does not contribute to the observed differences between the transgenic mice and the myofibrils from normal mice exchanged with the troponin mutant. It remains to be seen whether there are adaptational changes in the expression of other sarcomeric proteins, which may account for the differences. Taken together our results indicate that biochemically exchanged murine cardiac myofibrils are a suitable model to study the direct effects of hcTnI mutations on the activation–relaxation cycle.
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Increased force–sarcomere length relationship at low [Ca2+]
Regardless of the model system used – transgenic or biochemical exchange – the cTnI mutation R145G caused a significant upward shift of the force–SL relationship at pCa 7.5 when compared with myofibrils containing the wild-type isoform. It is known from previous studies that isoform switching of titin, one major element determining myofibrillar stiffness, can alter passive myofibrillar force properties during cardiomyopathy (Neagoe et al. 2002). However, since myofibrils exchanged with hcTnIR145G in vitro also exhibit increased force in the absence of Ca2+ we can exclude that any compensatory alterations in elastic elements such as titin contribute to this effect. Instead, it strengthens our hypothesis that the increased force–SL relationship observed here is caused by incomplete inhibition of force-generating crossbridges by the troponin complex. This is further supported by the finding that addition of 5 mM BDM, an inhibitor of myosin ATPase (Zhao et al. 1995), can partially reverse the upward shift of the force–SL relationship. Our data are in agreement with previous findings reporting increased ATPase activity and increased isometric force under relaxing conditions (pCa 9) in myofilaments and exchanged skinned fibre preparations carrying the same cTnI mutation (Takahashi-Yanaga et al. 2001; Burton et al. 2002; Lang et al. 2002).
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Ca2+ sensitivity of force activation
In myofibrils from mcTnIR146G transgenic mice we observed a slight, but significant decrease in Ca2+ sensitivity of force development, manifested by a reduction of pCa50 values. Myofibrils exchanged with hcTnIR145G in vitro did not show any significant effects on Ca2+ sensitivity of force development. While this is somewhat puzzling, there are conflicting data in the literature for the influence of cTnIR145G on the Ca2+ sensitivity of force development. Incorporation of cTnIR145G in skinned fibre preparations from guinea-pig heart had no effect on Ca2+ sensitivity (Burton et al. 2002), while in skinned fibre preparations from porcine left ventricle the R145G mutation induced a significant increase in Ca2+-sensitivity and ATPase-activity (Takahashi-Yanaga et al. 2001; Lang et al. 2002). Similarly, in skinned fibre preparations from mcTnIR146G transgenic mice the Ca2+ sensitivity of force development is increased (James et al. 2000). The question of what causes the different effects of the FHC mutation R145G on Ca2+ sensitivity in the different preparations still remains to be answered. Species-specific differences aside, many factors could possibly influence the effect of the mutation on the Ca2+ sensitivity, such as pH or the mutant-to-wild-type ratio (Elliott et al. 2000). Much more pronounced than the change in pCa50 observed in this study was the strong loss of cooperativity indicated by the gradient of the force–pCa relationship (Fig. 2). Thus, at very low activating [Ca2+], we actually observed an increase in Ca2+-activated force instead of a decrease, which correlates with the differences observed in the pCa–kACT relationship discussed below. These results parallel findings previously described for skinned fibre preparations exchanged with the R145G mutation, showing relative isometric force to be increased at low [Ca2+], whereas force development at high [Ca2+] is significantly decreased (Burton et al. 2002; Lang et al. 2002).
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We note that the cooperativity of Ca2+ activation in the analysed non-transgenic myofibrils (Hill coefficient >10) is much higher than those which had been found in skinned trabeculae from this mouse model (James et al. 2000). High Hill coefficients had been also reported for single skinned cardiomyocytes (Brandt et al. 1998). One reason for this could be that variability in Ca2+ sensitivity among individual myocytes generating the force in a multicellular preparation would add together to yield a less steep force–pCa relationship than those of the individual myocyte. The subcellular myofibrils investigated here might be therefore a more sensitive model than skinned trabeculae to detect changes in cooperativity.
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Kinetics of Ca2+-induced force activation
By investigating values for kACT after activation with different [Ca2+] we showed that regardless of the experimental method used, the mutation cTnIR145G affects kACT at both low and high [Ca2+] (Fig. 5). At low [Ca2+] kACT is increased, whereas at high [Ca2+] it is decreased. There is experimental evidence that kACT is rate-limited by crossbridge turnover kinetics (Stehle et al. 2002a; Tesi et al. 2002; Palmer & Kentish, 1998; Moss et al. 2004 and references cited within). Application of a fast slack–restretch sequence to an activated (pCa 4.5) myofibril results in tension redevelopment with kinetic values kTR, which are similar to kACT. This implies that the Ca2+-induced switch-on of the regulatory system is a very rapid process occurring before the onset of tension development and therefore cannot directly rate-limit the rate constant of force development kACT. This is further corroborated by our previous report that human cardiac myofibrils exhibit 10-fold lower values of kACT than cardiac myofibrils isolated from mice (Stehle et al. 2002a). If kACT was rate-limited by the switch-on kinetics of the Tn complex, one would expect exchange of human cTn into murine myofibrils to slow down myofibrillar force kinetics. However, here we report that incorporation of human wild-type troponin complex into murine cardiac myofibrils has no impact on the values of kACT when compared with values found in native myofibrils isolated from non-transgenic mice (Table 1). It therefore seems unlikely that the troponin isoforms directly contribute to the observed effects of the mutation on the activation kinetics. Instead, we favour a model in which force development kinetics are primarily rate-limited by turnover kinetics of the crossbridge cycle (Stehle et al. 2002a; Martin et al. 2004). The cTn–Tm complex rapidly fluctuates between off (Ca2+ bound) and on (no Ca2+ bound) states (Brenner & Chalovich, 1999). It thereby regulates the probability and thus the apparent rate constant for the formation of force-generating crossbridges. The effects of the cTnI mutation on the Ca2+ dependences of force and kACT could therefore be explained by an increased fraction of cTn–Tm units in on-states at low [Ca2+], leading to increased values of kACT, and a reduced fraction at high [Ca2+], leading to the significantly decreased values of kACT after activation with pCa 4.5. This implies that the mutation impairs two different functions of the TnI molecule: first, to fully inhibit force-generating actomyosin interactions at low [Ca2+], and second, to release this inhibition at high [Ca2+]. This leads to the hypothesis that the amino acid at position R145 is required for the proper interaction of TnI with TnC in the Ca2+-bound state. This is supported by studies on the inhibitory peptide, which has been shown to be sufficient to partially restore Ca2+ regulation of actomyosin ATPase (Tripet et al. 1997) and force (Van Eyk et al. 1993). In particular, Li et al. (2003) demonstrated that the R145G mutation decreases the affinity of the isolated inhibitory peptide for TnC.
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Kinetics of force relaxation
During relaxation, kLIN, the rate constant of the initial slow linear phase, was not significantly affected by the cTnI mutation in both the in vitro exchange preparations and the transgenic myofibrils. Although kLIN is difficult to determine accurately in murine cardiac myofibrils because of their extremely rapid turnover kinetics, we can exclude that the mutation changes this parameter by more than 20%. Previous studies revealed that during the initial slow linear phase of relaxation all sarcomeres remain isometric (Stehle et al. 2002a) and several lines of evidence in other species suggest that no significant recruitment of new force-generating crossbridges occurs during this phase (for review see Poggesi et al. 2005). We therefore assume that also in the mouse the rate constant kLIN is predominantly determined by the apparent rate by which crossbridges leave force-generating states under isometric conditions (Stehle et al. 2002a; Tesi et al. 2002). The fact that kLIN is not altered suggests that the cTnI mutation has no effect on the decay of force-generating crossbridges during this relaxation phase.
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The initial linear relaxation phase ends at a time tLIN after Ca2+ removal. As reported previously (Stehle et al. 2002a), this is the time point when the first, mechanically weakest, sarcomere in a myofibril elongates, thereby initiating the fast exponential relaxation phase. During sarcomere relaxation in this phase crossbridge detachment rates increase in all sarcomeres, regardless of their lengthening or shortening, and crossbridges leave force-generating states by increased rates of backward turnover kinetics in currently relaxing sarcomeres. This releases the strain on the other sarcomeres and allows the remaining crossbridges to leave force-generating states by rapid forward kinetics (Stehle et al. 2002a). Different to the rate constant kLIN during the linear relaxation phase, kREL is therefore determined by the rapid crossbridge detachment in the presence of sarcomere dynamics (Stehle et al. 2002a).
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We showed that incorporation of hcTnIR145G into isolated myofibrils leads to a prolongation of tLIN by 70% compared with hcTnIWT-exchanged myofibrils. This implies that the sarcomeres remain isometric for a longer time during relaxation. It is not quite clear at present by which mechanism the mutation delays the initiation of the fast relaxation phase. One factor may be the mechanical strain on the individual crossbridges (cf. Poggesi et al. 2005 for review). Reduced force leads to reduced strain inhomogeneities within the elastic elements of the sarcomeres and thereby reduces the probability of relaxation onset in the weakest sarcomere. Hence, the increase in tLIN may be related to the decreased Ca2+-activated force (Fmax/CSA) in the myofibrils exchanged with hcTnIR145G. Consistent with this interpretation, tLIN was not altered in myofibrils isolated from the transgenic cTnIR145G mice, in which Fmax/CSA remained unchanged, compared with non-transgenic control mice. However, as this interpretation is consistent only within each of the two models but not between them, additional, but as yet unknown, factors must come into play.
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The most prominent effect of the mutation common to both models was the decelerated fast exponential phase of force relaxation. In both models, the mutation decreased the rate constant kREL of this phase to 60–70% of the wild-type or non-transgenic control, respectively. Tesi et al. (2002) showed in skeletal myofibrils that kREL depends very sensitively on the final steady-state force after relaxation. They found kREL to be decreased 3- to 4-fold whereas kLIN remains unaffected if the [Ca2+] is not reduced to fully relaxing levels but to slightly activating levels leading to a residual steady-state force of only about 10% of maximum force. We recently confirmed these findings also for cardiac myofibrils (R. Stehle, unpublished data). The detailed molecular mechanism of the dependence of relaxation kinetics on the final force remains unclear. However, we presume that the observed decrease in kREL by the mutation does not directly reflect a slower switch-off of the thin filament (koff), which rate-limits the force decay, since this should even more sensitively affect the rate constant of the initial phase (kLIN) which was not the case. Instead we suggest that the reduction of kREL relates somehow to the effect of the mutation to increase the steady-state force at low [Ca2+] as indicated by the increased force–SL relation. The strong decrease in kREL with increasing final force might indicate a load dependence of sarcomere dynamics as known, for example, for the shortening velocity, which varies most steeply with load at low levels. Regardless of the molecular mechanism for the mutant-induced decrease in kREL, this slow-down in mechanical relaxation of myofibrils is expected to directly affect the rate of the isovolumetric pressure decay in the heart and could therefore be one of the main determinants of the decreased value of tau () found in the working heart analysis of cTnIR146G transgenic mice (James et al. 2000). This effect on the overall duration of force decline is even more pronounced in hcTnIR145G-exchanged myofibrils, since they additionally exhibit a prolonged linear phase (tLIN), which contributes to the overall relaxation period. Clinically, the impaired relaxation abilities of the contractile apparatus could explain the diastolic dysfunction of the heart, as typical for patients suffering from hypertrophic cardiomyopathy (Kass et al. 2004).
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Maximum Ca2+-induced force development (Fmax/CSA)
In myofibrils exchanged in vitro the maximum force development at pCa 4.5 (Fmax/CSA) was significantly reduced. Interestingly, no changes were observed for Fmax/CSA in myofibrils isolated from the transgenic mice. Previous studies have already described a reduction of maximum force development in cTnIR145G-exchanged skinned fibre preparations and a significant reduction of maximum ATPase activity after incorporation of cTnIR145G in reconstituted thin filaments (Elliott et al. 2000; Takahashi-Yanaga et al. 2001; Burton et al. 2002). This is in agreement with our own results, as impaired maximum ATPase activity could be the underlying cause of the decreased Fmax/CSA in hcTnIR145G-exchanged myofibrils described in the present study.
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Surprisingly, despite the decrease in kACT no changes were observed for Fmax/CSA in myofibrils from the transgenic mice. According to the model of Brenner (1988), kACT = fapp + gapp and force is proportional to fapp/(fapp + gapp), where fapp and gapp are the crossbridge attachment and detachment rates, respectively. Hence, changes in kACT should relate to changes in force and vice versa. As kLIN is not affected by the mutation we assume that the crossbridge detachment rate gapp is not altered. The mutation therefore can be regarded as affecting the Ca2+-regulated crossbridge attachment rate fapp. However, because the relation of kACT versus relative force is very steep at high forces, at high activating [Ca2+] changes in the crossbridge attachment rate fapp are more sensitively detected by force development kinetics than by the force itself. This is the most likely reason why we do not find significant changes in Fmax/CSA between myofibrils from non-transgenic and transgenic mice. Future studies will have to show whether gene doses of the mutation can contribute to an effect of the mutation on Fmax/CSA in myofibrils from the transgenic animals compared with the biochemically exchanged myofibrils.
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While kACT was significantly decreased in isolated cardiac myofibrils from the transgenic mice, James et al. (2000) reported an enhanced contractile function in working heart analysis, described by increased values of +dP/dt. We assume that the enhanced contractility observed on the organ level could at least in part be due to adaptational processes aiming to compensate the impaired contractility of the myofilaments.
In summary, to the best of our knowledge our work provides the first direct experimental evidence that FHC can impair kinetic properties of Ca2+-induced force activation and relaxation at the myofibrillar level. The present results lead to the conclusion that the cTnIR145G mutation primarily perturbs the interactions between cTnI–actin and cTnI–cTnC, thereby affecting crossbridge turnover kinetics, which could then underlie the impaired pressure dynamics found in the working heart of cTnIR146G transgenic animals. We therefore suggest that impaired myofibrillar relaxation could be a fundamental cause for FHC-induced diastolic dysfunction. As these changes were observed in a mouse line that exhibited no gross morphological pathology of the heart (James et al. 2000), our study also indicates that functional changes at the myofibrillar level may precede morphological changes. Whether the functional changes cause the hypertrophy and in particular whether they are relevant for the development of the disease in human patients remains to be seen. In addition we propose that the increased contractility detected in the working heart analysis of the transgenic animals may be due to an over-compensation of reduced myofibrillar contractility. More functional and clinical studies will be needed to further understand the physiological impacts of FHC-associated cTnI mutants and the regulatory processes finally leading to hypertrophy and heart failure.
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2 Department of Cardiovascular Medicine, University of Oxford, UK
3 Department of Pediatrics, Division of Molecular Cardiovascular Biology, Cincinnati Children's Hospital, Cincinnati, OH 45229-3039, USA
Abstract
Familial hypertrophic cardiomyopathy (FHC) has been linked to mutations in sarcomeric proteins such as human cardiac troponin I (hcTnI). To elucidate the functional consequences of the mutation hcTnIR145G on crossbridge kinetics, force kinetics were analysed in murine cardiac myofibrils carrying either the mutant or the wild-type protein. The mutation was introduced into the myofibrils in two different ways: in the first approach, the endogenous Tn was replaced by incubation of the myofibrils with an excess of reconstituted recombinant hcTn containing either hcTnIWT or hcTnIR145G. Alternatively, myofibrils were isolated either from non-transgenic or transgenic mice expressing the corresponding mcTnIR146G mutation. In myofibrils from both models, the mutation leads to a significant upward shift of the passive force–sarcomere length relation determined at pCa 7.5. Addition of 5 mM BDM (2,3-butandione-2-monoxime), an inhibitor of actomyosin ATPase partially reverses this shift, suggesting that the mutation impairs the normal function of cTnI to fully inhibit formation of force-generating crossbridges in the absence of Ca2+. Maximum force development (Fmax) is significantly decreased by the mutation only in myofibrils exchanged with hcTnIR145G in vitro. Ca2+ sensitivity of force development was reduced by the mutation in myofibrils from transgenic mice but not in exchanged myofibrils. In both models the rate constant of force development kACT is reduced at maximal [Ca2+] but not at low [Ca2+] where it is rather increased. Force relaxation is significantly prolonged due to a reduction of the relaxation rate constant kREL. We therefore assume that the impairment in the regulatory function of TnI by the mutation leads to modulations in crossbridge kinetics that significantly alter the dynamics of myofibrillar contraction and relaxation.
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Introduction
Familial hypertrophic cardiomyopathy (FHC) is a disease characterized by ventricular hypertrophy and interstitial fibrosis, often resulting in arrhythmias, severe cardiac dysfunctions and sudden cardiac death (Marian & Roberts, 1998). FHC is linked to dominant missense mutations in the genes of almost all sarcomeric proteins including the cardiac troponin complex (cTn) (Kimura et al. 1997; Seidman & Seidman, 2001; Fatkin & Graham, 2002). Cardiac Tn is a Ca2+ sensor and regulatory element of the contraction–relaxation cycle in cardiac muscle. It is a heterotrimer consisting of cardiac troponin C (cTnC), the Ca2+-binding subunit, cardiac troponin I (cTnI), the inhibitory subunit that binds to actin and prevents crossbridge cycling at submicromolar [Ca2+] and cardiac troponin T (cTnT), which binds to cTnI, cTnC and tropomyosin (Tm), thereby anchoring the cTn complex to the thin filament. When intracellular [Ca2+] is high during systole, Ca2+ binds to cTnC, which then binds to cTnI with a high affinity, thereby releasing it from its inhibitory site on actin. This Ca2+-induced sequence of protein–protein interactions also leads to a change in the position of Tm on the actin filament, which results in turning actin filaments to an ‘on’ state (Solaro & Rarick, 1998). Relaxation is initiated when [Ca2+] falls to diastolic levels which then, in a reverse process, returns the thin filaments to an ‘off’ state. Central to the inhibitory function of cTnI is the so-called inhibitory region, an evolutionarily conserved 11 amino acid motif in the centre of the molecule (Gordon et al. 2000).
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To date, more than 15 missense mutations on the cTnI molecule have been identified in patients suffering from FHC (Kimura et al. 1997; Seidman & Seidman, 2001). To identify the underlying pathological mechanism of disease development, a number of investigators have studied the effects of FHC-related mutations on actomyosin ATPase and on contractile steady-state properties of skinned fibres and cardiomyocytes (Elliott et al. 2000; Takahashi-Yanaga et al. 2001; Burton et al. 2002; Lang et al. 2002; Westfall et al. 2002; Kohler et al. 2003). The most intensively studied cTnI mutation is located within the inhibitory region at position 145, replacing a positively charged arginine by an uncharged glycine (Kimura et al. 1997). Controversial data for Ca2+ sensitivity of force development aside, in vitro studies consistently show that the mutation R145G leads to an increase in ATPase activity at resting [Ca2+] (Elliott et al. 2000; Takahashi-Yanaga et al. 2001; Lang et al. 2002).
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Most of the pathological manifestations of this mutation observed in the human heart could be reproduced in a transgenic mouse model expressing the corresponding mutation R146G in the murine cTnI (mcTnI) (James et al. 2000). Interestingly, the phenotype of the transgenic mice depends on the amount of expressed mcTnIR146G. If the mutated protein makes up more than 50% of the total mcTnI protein the mice develop cardiomyocyte disarray, interstitial fibrosis and suffer premature death. In contrast, if the mutated protein contributes to the total mcTnI by only 40%, no pathological alterations in cellular and organ morphology were reported. Surprisingly, working heart analyses of these animals revealed impaired isovolumetric relaxation, indicated by a more than 2-fold prolonged lifetime of pressure fall (), a relatively load-independent measure of diastolic function (James et al. 2000). Coexistent to impaired relaxation, systolic function was improved as indicated by significantly increased values of +dP/dt. These data and similar data on transgenic mice overexpressing FHC-related mutations in other sarcomeric proteins (Evans et al. 2000) reflect the clinical observation in FHC patients, which typically exhibit normal and sometimes even supra-normal contractile function, while lusitropy is severely impaired (Kass et al. 2004).
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Studies performed on intact trabeculae revealed that the rate-limiting steps of cardiac contraction and relaxation reside in the myofilaments (Janssen et al. 2002). This makes crossbridge kinetics and altered kinetics of regulatory proteins possible candidates for the altered haemodynamics observed in FHC. To date the effects of FHC mutations in myosin heavy chain, TnT and Tm on crossbridge kinetics have been investigated in skinned fibres where kinetics are derived from force transients induced by mechanical stretch or release (Palmiter et al. 2000). To our knowledge, no study has investigated the effect of FHC-related mutations on force kinetics in a myofibrillar system, in which isometric contraction and relaxation are induced by the physiological activator [Ca2+]. Thus the question of whether the impaired diastolic function observed in FHC might be related to impaired force kinetics during myofibrillar relaxation remains to be answered.
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In this study we investigated the effects of the FHC mutation R145G in cTnI on the force kinetics of isolated myofibrillar bundles, in which contraction and relaxation is induced by changes in [Ca2+] via rapid solution change (Stehle et al. 2002b). We used two different strategies to introduce the mutation into the myofibrils. In one approach, we applied a recently developed biochemical technique to exchange the native Tn in myofibrils with exogenous, recombinant troponin complexes (Kruger et al. 2003). In the second method we investigated force kinetics in cardiac myofibrils isolated from mcTnIR146G transgenic mice. By comparing the data we obtained a detailed analysis of the impact of cTnIR145G on the kinetics of force activation and relaxation. Furthermore, we suggest possible adaptational processes in myofibrillar function that could be manifested during the development of heart failure in the transgenic animals.
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Methods
Preparation of recombinant human cardiac troponin complex
The three human cardiac troponin subunits hcTnCwt, hcTnTwt and hcTnI (either hcTnIwt or hcTnIR145G) were expressed separately in E. coli and purified as previously described (Kruger et al. 2003). The identity of the plasmids was verified by sequencing. The purified subunits were reconstituted to the heterotrimeric hcTn complex (Potter, 1982) that was then stored at 4°C for up to 1 week.
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Isolation of cardiac myofibrils
For the experiments on myofibrils from mcTnIR146G transgenic, adult FVB/N mice in which 40% of the endogenous cTnI was replaced with mcTnIR146G were used (James et al. 2000). For the hcTn exchange experiments myofibrils were prepared from adult Him: OF 1 mice. Mice were killed by cervical dislocation. All experiments were approved by the Institutional Animal Care and Use Committee.
The heart was removed and papillary muscles were dissected from the left ventricle, skinned with 1% v/v Triton-X 100 in rigor buffer (132 mmol l–1 NaCl, 5 mmol l–1 KCl, 1 mmol l–1 MgCl2, 10 mmol l–1 Tris pH 7.1, 5 mmol l–1 EGTA, 1 mmol l–1 NaN3 and a protease inhibitor cocktail) for 2 h and stored at 4°C in rigor buffer without Triton-X 100. Myofibrillar suspensions were prepared immediately before experiments by homogenizing skinned papillary muscles with a blender (Ultra Turrax) for 10 s at 4°C.
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Exchange of hcTn complex in mouse cardiac myofibrils
Replacement of native mcTn by recombinant hcTn was performed with a protocol adapted and modified for cardiac myofibrils (Kruger et al. 2003), which was initially described for skeletal muscle fibres by Brenner et al. (1999) and also for skinned cardiac fibres (Kohler et al. 2003). In brief, reconstituted hcTn complex was dialysed against rigor buffer, centrifuged at 13 000 g for 10 min and filtered through a polypropylene mesh (Millex-HV, 0.45 μm pore, Durapore) to remove aggregates. The freshly prepared myofibril suspension was centrifuged for 5 min at 380 g (10°C), the myofibril pellet resuspended in rigor buffer containing hcTn complex (1 mg ml–1) and incubated at 21°C for 60 min. To remove excess Tn, myofibrils were washed by centrifugation in rigor buffer (5 min at 380 g, 10°C). Exchange of the troponin subunits was confirmed by Western blot analysis as reported previously (Kruger et al. 2003). The protein components were separated on SDS-PAGE, transferred onto a nitrocellulose membrane (Schleicher & Schuell GmbH, Dassel, Germany) by standard tank transfer Western blot and probed by a monoclonal antibody against cTnI (clone 6F9, Dunn Labortechnik GmbH, Asbach, Germany) which also recognizes sTnI and a monoclonal anti-TnT antibody (JLT12, Sigma-Aldrich, Taufkirchen, Germany). The membranes were incubated with anti-mouse IgG-HRP (Sigma-Aldrich, Taufkirchen, Germany) and enzymatic activity was detected using an ECL-kit (Amersham Biosciences, Freiburg, Germany).
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Myofibrillar force measurements
Myofibrillar force kinetics were measured as previously described (Stehle et al. 2002a,b; Kruger et al. 2003). Composition of relaxing (pCa 7.5) and activating solution (pCa 4.5) was as in Stehle et al. (2002b). Using silicone adhesive, small myofibrillar bundles 5–10 μm in diameter and 40–70 μm in length were attached in relaxing solution to a stiff micro-needle at one end and to the tip of an atomic force cantilever at the other end. After mounting, the bundles were stretched to a sarcomere length of 2.3 μm. To avoid possible errors caused by phase contrast, determination of sarcomere length and cross-sectional area (CSA) were performed in bright field. Rapid changes ( 10 ms) in Ca2+ concentration were applied to the myofibrils using the solution change technique described in Stehle et al. (2002b). Force transients were recorded by monitoring the deflection of the cantilever by laser light reflection (Stehle et al. 2002a). All experiments were performed at 10°C.
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Data analysis and statistics
To determine force kinetic parameters, original force transients were fitted by either a mono-exponential function to derive kACT or by a function consisting of a linear and an exponential term to derive kLIN, tLIN and kREL (Stehle et al. 2002b). Statistical analysis was performed by subjecting the data to Student's t test. Significance was determined as *P < 0.05, **P < 0.01 and ***P < 0.001. Unless otherwise noted, all data are given as mean ± S.E.M. (standard error of the mean) of n experiments.
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Results
Incubation of isolated myofibrils with an excess of recombinant hcTn containing either hcTnIR145G or hcTnIWT results in almost complete replacement of the endogenous cTn (see Fig. 1 for Western blot analysis). From these results we conclude that the content of the mutant protein is above 80% in biochemically exchanged myofibrils. It should be noted that the transgenic mice only express about 40% mcTnIR146G with respect to their total mcTnI expression.
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A myofibrillar suspension from mouse papillary muscle was tested for cardiac troponin I. Lane 1: unexchanged control suspension; lane 2: myofibrillar suspension exchanged with hcTn. The molecular weight standard is shown on the right. After exchange no more mcTnI can be detected in hcTn-exchanged myofibrils.
Force development in the absence of Ca2+
Force–sarcomere length relationships were measured by stretching myofibrils (pCa 7.5) from slack sarcomere length of 2.0 μm to defined sarcomere length (SL) ranging from 2.2 to 2.6 μm. In this SL range, regular sarcomere patterns were observed in all myofibrils. In both experimental models, myofibrils containing mutant cTnI showed an increased upward shift of the force–SL relationship compared with either non-transgenic (ntg) (Fig. 2A) or hcTnWT (Fig. 2B). It is noteworthy that the slopes of the force–SL relationships were not altered by the mutant protein as would be expected if passive length-dependent stiffness properties were changed.
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Force is plotted as a function of sarcomere length (SL). A, mean ± S.E.M. of force–SL relationship for myofibrils isolated from mcTnIR146G transgenic mice (filled circles) (n = 16) and the corresponding non-transgenic controls (open circles) (n = 11). B, force–SL relationship of myofibrils exchanged with either hcTnIR145G (filled symbols) or hcTnIWT (open symbols) before (circles) and after (squares) incubation with 5 mM BDM, respectively. Results are representative of 3 independent experiments.
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Addition of 5 mM BDM (2,3-butandione-2-monoxime), an inhibitor of actomyosin ATPase, to the hcTnIR145G-exchanged myofibrils partially abolished the upward shift of the force–SL relationship. In contrast, BDM had no effect in hcTnIWT-containing myofibrils (Fig. 2B). This indicates that the mutation-induced force enhancement in the absence of Ca2+ at least partly results from force-generating crossbridges.
Ca2+ dependence of force development and Ca2+-induced force development kinetics
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Maximum Ca2+-activated force at pCa 4.5 normalized to cross-sectional area (Fmax/CSA) is significantly decreased by 24% in mouse myofibrils exchanged with hcTnIR145G compared with hcTnIWT-exchanged controls (see Table 1). In contrast, no difference in Fmax/CSA was observed for myofibrils isolated from transgenic mice when compared with those isolated from non-transgenic controls. The values for Fmax/CSA are given in Table 1.
The Ca2+ sensitivity of force development was significantly (P < 0.05) lower in myofibrils from the transgenic mice expressing mcTnIR146G (pCa50 5.73 ± 0.03; n = 12) when compared with the non-transgenic control mice (pCa50 5.79 ± 0.01 n = 4, Fig. 3). Cooperativity of force development was significantly reduced (P < 0.0001) as indicated by the reduced Hill coefficient of 6.2 ± 1.4 in myofibrils from the transgenic animals compared with 18 ± 2 of those from the non-transgenic control animals. In contrast, no significant reduction in Ca2+ sensitivity of force development was observed in myofibrils treated with the exchange protocol (data not shown).
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pCa50 is 5.73 ± 0.03 in myofibrils from the transgenic mcTnIR146G mice (n = 12) and 5.79 ± 0.01 in myofibrils from non-transgenic control mice (n = 4) (P < 0.05). Hill coefficients are 6.2 ± 1.4 in myofibrils from the transgenic animals and 18 ± 2 in those from the non-transgenic control (P < 0.0001).
Myofibrils from both models were analysed for their kinetics of force development following a rapid change from pCa 7.5 to pCa 4.5 (e.g. Fig. 4). As shown previously for untreated cardiac myofibrils from different species (Stehle et al. 2002b), rapid increases in [Ca2+] induced a mono-exponential increase in force in myofibrils from both models with a rate constant kACT, whereby kACT increases with increasing [Ca2+] (Fig. 5). We noted that the Ca2+-dependent increase in kACT of cardiac myofibrils exchanged with hcTnIWT (Fig. 5A) is similar to that of myofibrils from the non-transgenic mice (Fig. 5B). This indicates that Ca2+-induced force kinetics of murine myofibrils is not affected by the exchange of their native cTn complex with the recombinant human wildtype isoform. In both models, the mutation caused qualitatively similar changes in the kACT–pCa relationship (Fig. 5A and B): at low [Ca2+] (pCa 5.83), kACT is increased by the mutation, whereas at maximally activating [Ca2+] (pCa 4.5), kACT is significantly reduced from controls by 30% in the hcTnIR145G myofibrils and by 15% in myofibrils isolated from mcTnIR146G transgenic mice. The values for kACT after Ca2+ activation with pCa 4.5 are given in Table 1.
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The left side shows the kinetics of Ca2+-induced force development upon a switch from pCa 7.5 to 4.5. The right panel shows relaxation induced by a switch from pCa 4.5 to 7.5 at t = 1.5 s. The mathematical fit is shown in black, and kinetic parameters are given as kACT, kLIN, tLIN and kREL for each curve. Force transients were normalized to the same amplitude to illustrate the differences in kinetics; force at pCa 7.5 was subtracted before normalization.
A, kACT–pCa relationship for myofibrils exchanged in vitro with either hcTnIR145G (filled circles) or hcTnIWT (open circles) (n = 8–22). B, kACT–pCa relationship for myofibrils isolated from the transgenic mice expressing mcTnIR146G (filled circles) and for myofibrils isolated from the non-transgenic control mice (open circles) (n = 9–18). All data are means ± S.E.M.
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Myofibrillar relaxation kinetics
Rapidly deactivating myofibrils by changing the [Ca2+] from pCa 4.5 to 7.5 induced a biphasic relaxation (see Fig. 4 for typical force transients): an initial linear force decay followed by a fast exponential decay. The slow linear phase has a rate constant kLIN and a duration tLIN (Stehle et al. 2002b) while the exponential phase is described by the rate constant kREL. Figure 6 shows the values obtained for the parameters kLIN and tLIN of the initial force decay. Neither myofibrils exchanged with hcTnIR145G nor myofibrils isolated from mcTnIR146G transgenic mice differed in their values of kLIN compared with their hcTnIWT or non-transgenic controls, respectively (Fig. 6A). In myofibrils isolated from the transgenic mice the duration of the linear phase tLIN was unaltered compared with the corresponding non-transgenic mice (Fig. 6B), whereas myofibrils exchanged with hcTnIR145G showed a 1.7-fold longer tLIN (P < 0.005) than those exchanged with hcTnIWT.
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Shown are means ± S.E.M. obtained for kLIN (A) and tLIN (B) in myofibrils exchanged with hcTnIWT (n = 18) or hcTnIR145G (n = 22) and in myofibrils isolated from transgenic mice expressing mcTnIR146G (n = 14) or non-transgenic control animals (n = 18). *P < 0.05, Student's t test.
Figure 7 shows the values of the rate constant of the fast exponential relaxation phase kREL plotted as a function of the activating pCa. In hcTnIR145G-exchanged myofibrils following maximum activation with pCa 4.5, kREL is decreased by 40% of the values for hcTnIWT-exchanged myofibrils. In myofibrils from the transgenic mice kREL is reduced by 27% compared with controls. In both models kREL was also significantly reduced in relaxations initiated from partial activations with pCa 5.75 and 5.51 (Fig. 7). The values for kLIN, tLIN and kREL in relaxation experiments following maximum Ca2+ activation are summarized in Table 1.
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A, kREL–pCa relationship for myofibrils exchanged in vitro with either hcTnIR145G (filled circles) or hcTnIWT (open circles). B, kREL–pCa relationship for myofibrils isolated from the transgenic mice expressing mcTnIR146G (filled circles) and from the non-transgenic control mice (open circles). All data are means ± S.E.M. for n = 22 (hcTnIR145G), n = 17 (hcTnIWT), n = 18 (mcTnIR146G tg) and n = 14 (ntg control).
Discussion
In this study we analysed the contractile behaviour of cardiac myofibrils isolated from cTnIR146G transgenic mice and of cardiac myofibrils from wild-type mice, in which the endogenous Tn complex had been biochemically replaced with the corresponding human mutation hcTnIR145G. Common to both models we find an upward shift of the passive length–tension relationship, at low concentrations of Ca2+ an increase and at high concentrations of Ca2+ a decrease in the rate of contraction kACT, as well as a decrease in the rate of the fast exponential phase of the relaxation. Differences between the two models were observed for Ca2+-activated steady-state force and the duration of the linear phase of relaxation, which is prolonged in the exchanged but not in myofibrils taken from the transgenic mice. Several factors may account for the differences: (i) the exchanged myofibrils contained the human troponin isoform which despite its high homology with the mouse troponin isoform may nevertheless lead to subtle changes in the protein–protein interactions within the mouse sarcomeric environment; (ii) lower dosage of the mutant cTnI in the myofibrils taken from the transgenic mouse compared with the exchanged myofibrils; (iii) effect of the transgene on the expression of the myosin heavy chain or other sarcomeric proteins.
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We find it unlikely that the differences are due to the hybrid system for the following reasons: in recent studies we compared steady-state and kinetic contractile parameters in native murine myofibrils with those exchanged with hcTn and with native human myofibrils (Stehle et al. 2002b; Kruger et al. 2003). The kinetic parameters were an order of magnitude higher in the native as well as in the exchanged murine myofibrils compared with the native human myofibrils indicating that the kinetic parameters are not sensitive to the Tn isoform. Fmax/CSA was lower in the exchanged myofibrils as also seen in this study. At present we do not know whether this is due to the exchange procedure per se or is a reflection of the hybrid system. Irrespective of the cause of the depression in force, it is larger in myofibrils exchanged with hcTnIR145G indicating that this mutation has a specific effect on Fmax/CSA. Therefore we propose that in the transgenic myofibrils the gene dosage may be too low to affect Fmax/CSA.
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The impact of transgenic expression of R145G on myosin isoform content has been explored at both the RNA levels, via - and myosin heavy chain-specific oligonucleotide probes (Ng et al. 1991) and by titrating the two proteins directly using a general and myosin-specific antibody (Krenz et al. 2003). Transcript levels were up slightly (less than 10%) but no detectable changes in the relative or absolute levels of the myosin isoforms were detected. We therefore conclude that altered expression of MHC isoforms does not contribute to the observed differences between the transgenic mice and the myofibrils from normal mice exchanged with the troponin mutant. It remains to be seen whether there are adaptational changes in the expression of other sarcomeric proteins, which may account for the differences. Taken together our results indicate that biochemically exchanged murine cardiac myofibrils are a suitable model to study the direct effects of hcTnI mutations on the activation–relaxation cycle.
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Increased force–sarcomere length relationship at low [Ca2+]
Regardless of the model system used – transgenic or biochemical exchange – the cTnI mutation R145G caused a significant upward shift of the force–SL relationship at pCa 7.5 when compared with myofibrils containing the wild-type isoform. It is known from previous studies that isoform switching of titin, one major element determining myofibrillar stiffness, can alter passive myofibrillar force properties during cardiomyopathy (Neagoe et al. 2002). However, since myofibrils exchanged with hcTnIR145G in vitro also exhibit increased force in the absence of Ca2+ we can exclude that any compensatory alterations in elastic elements such as titin contribute to this effect. Instead, it strengthens our hypothesis that the increased force–SL relationship observed here is caused by incomplete inhibition of force-generating crossbridges by the troponin complex. This is further supported by the finding that addition of 5 mM BDM, an inhibitor of myosin ATPase (Zhao et al. 1995), can partially reverse the upward shift of the force–SL relationship. Our data are in agreement with previous findings reporting increased ATPase activity and increased isometric force under relaxing conditions (pCa 9) in myofilaments and exchanged skinned fibre preparations carrying the same cTnI mutation (Takahashi-Yanaga et al. 2001; Burton et al. 2002; Lang et al. 2002).
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Ca2+ sensitivity of force activation
In myofibrils from mcTnIR146G transgenic mice we observed a slight, but significant decrease in Ca2+ sensitivity of force development, manifested by a reduction of pCa50 values. Myofibrils exchanged with hcTnIR145G in vitro did not show any significant effects on Ca2+ sensitivity of force development. While this is somewhat puzzling, there are conflicting data in the literature for the influence of cTnIR145G on the Ca2+ sensitivity of force development. Incorporation of cTnIR145G in skinned fibre preparations from guinea-pig heart had no effect on Ca2+ sensitivity (Burton et al. 2002), while in skinned fibre preparations from porcine left ventricle the R145G mutation induced a significant increase in Ca2+-sensitivity and ATPase-activity (Takahashi-Yanaga et al. 2001; Lang et al. 2002). Similarly, in skinned fibre preparations from mcTnIR146G transgenic mice the Ca2+ sensitivity of force development is increased (James et al. 2000). The question of what causes the different effects of the FHC mutation R145G on Ca2+ sensitivity in the different preparations still remains to be answered. Species-specific differences aside, many factors could possibly influence the effect of the mutation on the Ca2+ sensitivity, such as pH or the mutant-to-wild-type ratio (Elliott et al. 2000). Much more pronounced than the change in pCa50 observed in this study was the strong loss of cooperativity indicated by the gradient of the force–pCa relationship (Fig. 2). Thus, at very low activating [Ca2+], we actually observed an increase in Ca2+-activated force instead of a decrease, which correlates with the differences observed in the pCa–kACT relationship discussed below. These results parallel findings previously described for skinned fibre preparations exchanged with the R145G mutation, showing relative isometric force to be increased at low [Ca2+], whereas force development at high [Ca2+] is significantly decreased (Burton et al. 2002; Lang et al. 2002).
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We note that the cooperativity of Ca2+ activation in the analysed non-transgenic myofibrils (Hill coefficient >10) is much higher than those which had been found in skinned trabeculae from this mouse model (James et al. 2000). High Hill coefficients had been also reported for single skinned cardiomyocytes (Brandt et al. 1998). One reason for this could be that variability in Ca2+ sensitivity among individual myocytes generating the force in a multicellular preparation would add together to yield a less steep force–pCa relationship than those of the individual myocyte. The subcellular myofibrils investigated here might be therefore a more sensitive model than skinned trabeculae to detect changes in cooperativity.
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Kinetics of Ca2+-induced force activation
By investigating values for kACT after activation with different [Ca2+] we showed that regardless of the experimental method used, the mutation cTnIR145G affects kACT at both low and high [Ca2+] (Fig. 5). At low [Ca2+] kACT is increased, whereas at high [Ca2+] it is decreased. There is experimental evidence that kACT is rate-limited by crossbridge turnover kinetics (Stehle et al. 2002a; Tesi et al. 2002; Palmer & Kentish, 1998; Moss et al. 2004 and references cited within). Application of a fast slack–restretch sequence to an activated (pCa 4.5) myofibril results in tension redevelopment with kinetic values kTR, which are similar to kACT. This implies that the Ca2+-induced switch-on of the regulatory system is a very rapid process occurring before the onset of tension development and therefore cannot directly rate-limit the rate constant of force development kACT. This is further corroborated by our previous report that human cardiac myofibrils exhibit 10-fold lower values of kACT than cardiac myofibrils isolated from mice (Stehle et al. 2002a). If kACT was rate-limited by the switch-on kinetics of the Tn complex, one would expect exchange of human cTn into murine myofibrils to slow down myofibrillar force kinetics. However, here we report that incorporation of human wild-type troponin complex into murine cardiac myofibrils has no impact on the values of kACT when compared with values found in native myofibrils isolated from non-transgenic mice (Table 1). It therefore seems unlikely that the troponin isoforms directly contribute to the observed effects of the mutation on the activation kinetics. Instead, we favour a model in which force development kinetics are primarily rate-limited by turnover kinetics of the crossbridge cycle (Stehle et al. 2002a; Martin et al. 2004). The cTn–Tm complex rapidly fluctuates between off (Ca2+ bound) and on (no Ca2+ bound) states (Brenner & Chalovich, 1999). It thereby regulates the probability and thus the apparent rate constant for the formation of force-generating crossbridges. The effects of the cTnI mutation on the Ca2+ dependences of force and kACT could therefore be explained by an increased fraction of cTn–Tm units in on-states at low [Ca2+], leading to increased values of kACT, and a reduced fraction at high [Ca2+], leading to the significantly decreased values of kACT after activation with pCa 4.5. This implies that the mutation impairs two different functions of the TnI molecule: first, to fully inhibit force-generating actomyosin interactions at low [Ca2+], and second, to release this inhibition at high [Ca2+]. This leads to the hypothesis that the amino acid at position R145 is required for the proper interaction of TnI with TnC in the Ca2+-bound state. This is supported by studies on the inhibitory peptide, which has been shown to be sufficient to partially restore Ca2+ regulation of actomyosin ATPase (Tripet et al. 1997) and force (Van Eyk et al. 1993). In particular, Li et al. (2003) demonstrated that the R145G mutation decreases the affinity of the isolated inhibitory peptide for TnC.
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Kinetics of force relaxation
During relaxation, kLIN, the rate constant of the initial slow linear phase, was not significantly affected by the cTnI mutation in both the in vitro exchange preparations and the transgenic myofibrils. Although kLIN is difficult to determine accurately in murine cardiac myofibrils because of their extremely rapid turnover kinetics, we can exclude that the mutation changes this parameter by more than 20%. Previous studies revealed that during the initial slow linear phase of relaxation all sarcomeres remain isometric (Stehle et al. 2002a) and several lines of evidence in other species suggest that no significant recruitment of new force-generating crossbridges occurs during this phase (for review see Poggesi et al. 2005). We therefore assume that also in the mouse the rate constant kLIN is predominantly determined by the apparent rate by which crossbridges leave force-generating states under isometric conditions (Stehle et al. 2002a; Tesi et al. 2002). The fact that kLIN is not altered suggests that the cTnI mutation has no effect on the decay of force-generating crossbridges during this relaxation phase.
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The initial linear relaxation phase ends at a time tLIN after Ca2+ removal. As reported previously (Stehle et al. 2002a), this is the time point when the first, mechanically weakest, sarcomere in a myofibril elongates, thereby initiating the fast exponential relaxation phase. During sarcomere relaxation in this phase crossbridge detachment rates increase in all sarcomeres, regardless of their lengthening or shortening, and crossbridges leave force-generating states by increased rates of backward turnover kinetics in currently relaxing sarcomeres. This releases the strain on the other sarcomeres and allows the remaining crossbridges to leave force-generating states by rapid forward kinetics (Stehle et al. 2002a). Different to the rate constant kLIN during the linear relaxation phase, kREL is therefore determined by the rapid crossbridge detachment in the presence of sarcomere dynamics (Stehle et al. 2002a).
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We showed that incorporation of hcTnIR145G into isolated myofibrils leads to a prolongation of tLIN by 70% compared with hcTnIWT-exchanged myofibrils. This implies that the sarcomeres remain isometric for a longer time during relaxation. It is not quite clear at present by which mechanism the mutation delays the initiation of the fast relaxation phase. One factor may be the mechanical strain on the individual crossbridges (cf. Poggesi et al. 2005 for review). Reduced force leads to reduced strain inhomogeneities within the elastic elements of the sarcomeres and thereby reduces the probability of relaxation onset in the weakest sarcomere. Hence, the increase in tLIN may be related to the decreased Ca2+-activated force (Fmax/CSA) in the myofibrils exchanged with hcTnIR145G. Consistent with this interpretation, tLIN was not altered in myofibrils isolated from the transgenic cTnIR145G mice, in which Fmax/CSA remained unchanged, compared with non-transgenic control mice. However, as this interpretation is consistent only within each of the two models but not between them, additional, but as yet unknown, factors must come into play.
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The most prominent effect of the mutation common to both models was the decelerated fast exponential phase of force relaxation. In both models, the mutation decreased the rate constant kREL of this phase to 60–70% of the wild-type or non-transgenic control, respectively. Tesi et al. (2002) showed in skeletal myofibrils that kREL depends very sensitively on the final steady-state force after relaxation. They found kREL to be decreased 3- to 4-fold whereas kLIN remains unaffected if the [Ca2+] is not reduced to fully relaxing levels but to slightly activating levels leading to a residual steady-state force of only about 10% of maximum force. We recently confirmed these findings also for cardiac myofibrils (R. Stehle, unpublished data). The detailed molecular mechanism of the dependence of relaxation kinetics on the final force remains unclear. However, we presume that the observed decrease in kREL by the mutation does not directly reflect a slower switch-off of the thin filament (koff), which rate-limits the force decay, since this should even more sensitively affect the rate constant of the initial phase (kLIN) which was not the case. Instead we suggest that the reduction of kREL relates somehow to the effect of the mutation to increase the steady-state force at low [Ca2+] as indicated by the increased force–SL relation. The strong decrease in kREL with increasing final force might indicate a load dependence of sarcomere dynamics as known, for example, for the shortening velocity, which varies most steeply with load at low levels. Regardless of the molecular mechanism for the mutant-induced decrease in kREL, this slow-down in mechanical relaxation of myofibrils is expected to directly affect the rate of the isovolumetric pressure decay in the heart and could therefore be one of the main determinants of the decreased value of tau () found in the working heart analysis of cTnIR146G transgenic mice (James et al. 2000). This effect on the overall duration of force decline is even more pronounced in hcTnIR145G-exchanged myofibrils, since they additionally exhibit a prolonged linear phase (tLIN), which contributes to the overall relaxation period. Clinically, the impaired relaxation abilities of the contractile apparatus could explain the diastolic dysfunction of the heart, as typical for patients suffering from hypertrophic cardiomyopathy (Kass et al. 2004).
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Maximum Ca2+-induced force development (Fmax/CSA)
In myofibrils exchanged in vitro the maximum force development at pCa 4.5 (Fmax/CSA) was significantly reduced. Interestingly, no changes were observed for Fmax/CSA in myofibrils isolated from the transgenic mice. Previous studies have already described a reduction of maximum force development in cTnIR145G-exchanged skinned fibre preparations and a significant reduction of maximum ATPase activity after incorporation of cTnIR145G in reconstituted thin filaments (Elliott et al. 2000; Takahashi-Yanaga et al. 2001; Burton et al. 2002). This is in agreement with our own results, as impaired maximum ATPase activity could be the underlying cause of the decreased Fmax/CSA in hcTnIR145G-exchanged myofibrils described in the present study.
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Surprisingly, despite the decrease in kACT no changes were observed for Fmax/CSA in myofibrils from the transgenic mice. According to the model of Brenner (1988), kACT = fapp + gapp and force is proportional to fapp/(fapp + gapp), where fapp and gapp are the crossbridge attachment and detachment rates, respectively. Hence, changes in kACT should relate to changes in force and vice versa. As kLIN is not affected by the mutation we assume that the crossbridge detachment rate gapp is not altered. The mutation therefore can be regarded as affecting the Ca2+-regulated crossbridge attachment rate fapp. However, because the relation of kACT versus relative force is very steep at high forces, at high activating [Ca2+] changes in the crossbridge attachment rate fapp are more sensitively detected by force development kinetics than by the force itself. This is the most likely reason why we do not find significant changes in Fmax/CSA between myofibrils from non-transgenic and transgenic mice. Future studies will have to show whether gene doses of the mutation can contribute to an effect of the mutation on Fmax/CSA in myofibrils from the transgenic animals compared with the biochemically exchanged myofibrils.
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While kACT was significantly decreased in isolated cardiac myofibrils from the transgenic mice, James et al. (2000) reported an enhanced contractile function in working heart analysis, described by increased values of +dP/dt. We assume that the enhanced contractility observed on the organ level could at least in part be due to adaptational processes aiming to compensate the impaired contractility of the myofilaments.
In summary, to the best of our knowledge our work provides the first direct experimental evidence that FHC can impair kinetic properties of Ca2+-induced force activation and relaxation at the myofibrillar level. The present results lead to the conclusion that the cTnIR145G mutation primarily perturbs the interactions between cTnI–actin and cTnI–cTnC, thereby affecting crossbridge turnover kinetics, which could then underlie the impaired pressure dynamics found in the working heart of cTnIR146G transgenic animals. We therefore suggest that impaired myofibrillar relaxation could be a fundamental cause for FHC-induced diastolic dysfunction. As these changes were observed in a mouse line that exhibited no gross morphological pathology of the heart (James et al. 2000), our study also indicates that functional changes at the myofibrillar level may precede morphological changes. Whether the functional changes cause the hypertrophy and in particular whether they are relevant for the development of the disease in human patients remains to be seen. In addition we propose that the increased contractility detected in the working heart analysis of the transgenic animals may be due to an over-compensation of reduced myofibrillar contractility. More functional and clinical studies will be needed to further understand the physiological impacts of FHC-associated cTnI mutants and the regulatory processes finally leading to hypertrophy and heart failure.
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