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LETTERS |
Firstly, they suggest that ‘for isolated rabbit myofibrils, activated with pCa = 4.5 at 10°C, it is not at all clear that activation was maximal’ and hence the data may be from submaximally activated responses. The available experimental evidence from a number of studies on skinned mammalian muscle fibres does not support their claim (Stephenson & Williams, 1985; Goldman et al. 1987; Brenner, 1988). Moreover, thin filaments are fully activated at pCa 4.5, even at temperatures below 10°C (Brenner et al. 1999). Steady-state force–pCa relations and force kinetics data of Piroddi et al. (2003), Poggesi et al. (2005) and Stehle et al. (2002) clearly show that isolated rabbit psoas and cardiac myofibrils are fully Ca2+-regulated and maximally activated at pCa 4.5. We have confirmed that the fluorescent antibody labelling as such does not alter the functional properties of rabbit psoas and cardiac myofibrils (Telley et al. 2006a). The rate constant of Ca2+-induced force development, kACT, is very sensitive to changes in [Ca2+], especially at high levels of Ca2+ activation in the rabbit psoas (e.g. see Fig. 4D in Poggesi et al. 2005). The value of kACT indicated in our study (Telley et al. 2006b) is in agreement with those reported in the literature for rabbit psoas myofibrils and fibres at saturating [Ca2+] at this temperature.
Secondly, they point out that since ‘rabbit body temperature is normally close to 38°C, activation is likely to be well below maximum at 10°C’. The studies referred to above showed that the Ca2+ sensitivity of thin filament activation is actually greater at lower than at higher temperatures; the pCa–tension relation is shifted to the left (lower Ca2+ levels) with cooling (see Fig. 1 in Stephenson & Williams, 1985 and Fig. 4 in Goldman et al. 1987). Therefore, we believe that the detailed critical comments made in the letter with regard to myofibril activation not being maximal in our experiments lack experimental support.
Thirdly, they say ‘comparison of Telley et al.'s quoted specific tension with published values from the literature suggests that activation was only about 50%’. The tension was 
170 nN µm–2 (i.e. kN m–2) and this is within the range of values in the literature. This is expected to be about half that at 35–37°C, as shown from tetanically activated rat muscle experiments (Ranatunga & Wylie, 1983) but this is not due to submaximal activation; similar temperature dependence is obtained for maximally Ca2+-activated tension in skinned mammalian fibres. Thus, although the number of attached cross-bridges remains approximately similar at different temperatures the proportion of force-holding cross-bridges increases with temperature. This is well known and is true for muscles of different species and it is due to the endothermic nature of muscle force generation (see references in Coupland et al. 2005).
Fourthly, it is suggested in their letter that ‘the choice by Telley et al. of half-sarcomere lengths only just beyond optimum overlap (1.2 µm) suggests that these isolated myofibrils have a high passive tension’. It is true that we have not examined the length–tension relation in detail in our study. However, the zero-force level in our figures was determined by a slack test carried out before each contraction and the passive tension at the initial hSL of 1.2 µm was small, 3–8 nN µm–2 or 2–3% P0. Additionally, detailed analysis of force transients showed that after deactivation at the longer length (i.e. after 20%
L0 stretch to a mean hSL 1.3–1.4 µm), the passive force estimated at steady-state is only 5–8% P0 higher than before the stretch (see the analysis illustrated in Fig. 1). Even though this seems slightly higher than values given in the literature (see Minajeva et al. 2002 and references therein), the value does not constitute a high resting tension in myofibrils; moreover, a resting tension of < 10% P0 is more in accord with the modelled tension curve at 100% activation than at 50% activation, shown in the letter.
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Finally, there have been a number of studies on intact and skinned frog and mammalian fibres (Bagni et al. 1995; Mutungi & Ranatunga, 1996) that clearly showed that the response to stretch in resting muscle fibres is complex and displays velocity-dependent (viscoelastic) components; extension of the Huxley (1980) calculations to take account of other sarcomeric features (e.g. titin filaments) could partly account for this complex tension response to stretch in resting muscle fibres (see Ranatunga, 2001). Therefore, incorporation of parallel visco-elasticity in our model was realistic and essential; hence our model (Telley et al. 2003), although incomplete, is not based entirely on steady state force–length relations but rather it contains dynamic length- and time-dependent behaviour features. Additionally, there is accumulating evidence now that titin stiffness is Ca2+ sensitive and increases on activation (Labeit et al. 2003). Thus, there may be mechanisms within sarcomeres that can prevent sarcomere popping during lengthening (e.g. titin filaments, C-protein, etc., see Pinniger et al. 2006) so that force and work generated by cross-bridges during muscle function are efficiently transmitted across sarcomeres in muscle fibres. From our experiments we cannot exclude the possibility that sarcomere popping may occur under other conditions, e.g. at longer sarcomere length, higher temperature and after injury. On the other hand, despite the theoretical attractiveness of the sarcomere popping idea, there is a dearth of clear experimental evidence that sarcomere popping is a physiological process of importance in lengthening muscle; our modelling indicates that sarcomere dynamics during and after stretch are slow so that if sarcomere popping occurs it would be a rapid transient process that cannot be adequately described with steady-state relationships of force, velocity and length. The challenge for the future is to elucidate the molecular mechanisms involved in preventing sarcomere popping during lengthening of active muscle; it appears that the role of other sarcomeric structures in this regard deserves more careful attention.
Laboratory for BiomechanicsETH Zürich8093 Zürich, SwitzerlandEmail: jdenoth{at}ethz.ch
Institute for Vegetative PhysiologyUniversity of Cologne50931 Cologne, Germany
Department of PhysiologyUniversity of BristolBristol BS8 ITD, UK
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