Fast stretching of skeletal muscle fibres abolishes residual force enhancement

In 1952, Abbott and Aubert showed that the steady-state force obtained after active stretch (eccentric contraction) of frog, toad and dogfish muscles was greater than the corresponding isometric force at the same final length and level of activation. This force increase was termed ‘residual force enhancement’ (rFE). rFE has been systematically observed in mammalian and amphibian striated muscle, and on all structural levels of muscle including single sarcomere, single myofibril, single fibre, muscle bundle and whole muscle preparations in situ and in vivo (Edman et al., 1978, 1982; Herzog and Leonard, 2002; Peterson et al., 2004; Leonard et al., 2010; Leonard and Herzog, 2010; Hahn et al., 2010; Pinnell et al., 2019).

The energy cost of force is substantially reduced in rFE (Joumaa and Herzog, 2013), suggesting that eccentric contractions could be an effective mechanism to reduce the cost of everyday movement. However, despite more than half a century of research, the detailed molecular mechanisms underlying rFE remain unknown. There is strong evidence in the literature that rFE could result from an engagement of a passive element upon muscle activation and stretch (Edman et al., 1982; Noble, 1992; Herzog and Leonard, 2002; Pinniger et al., 2006; Nocella et al., 2014; Tomalka et al., 2017; Nishikawa et al., 2020; Hessel et al., 2021). Titin, a protein in the sarcomere that is responsible for most of the passive force produced by single fibres and myofibrils (Bartoo et al., 1997; Granzier et al., 2000; Linke, 2000), has been suggested as a possible candidate for this increase in force. It has been shown that calcium binds to titin (Yamasaki et al., 2001; DuVall et al., 2017; Dutta et al., 2018) and increases its stiffness (Labeit et al., 2003; Joumaa et al., 2008; DuVall et al., 2017). Furthermore, it has been suggested that titin interacts with actin upon muscle activation, which results in a reduction in its free spring length, and an increase in its contribution to force when actively stretched compared with passive stretching (Linke et al., 1997; Yamasaki et al., 2001; Leonard and Herzog, 2010; Nishikawa et al., 2012; DuVall et al., 2017; Zhou et al., 2021). However, results using 2,3-butanedione monoxime (BDM) exposure and troponin C deletion showed that when full cross-bridge cycling was inhibited, the calcium induced increase in titin's stiffness had little contribution to rFE (Joumaa et al., 2008; Powers et al., 2014). Furthermore, it has been shown that when myofibrils were actively stretched while cross-bridges were not allowed to cycle, rFE was substantially reduced, suggesting that calcium activation is not a primary cause eliciting rFE and that cross-bridge cycling appears essential to produce rFE (Leonard and Herzog, 2010; Powers et al., 2014). Combined, these experimental results suggest that titin–calcium interaction alone cannot produced rFE and proper cross-bridge cycling is a prerequisite for rFE to occur.

If proper cross-bridge cycling is indeed required to produce rFE, then rFE might not occur in muscles that are stretched so fast as to produce muscle slippage (Sugi, 1972; Griffiths et al., 1980). Slippage is a sudden perturbation in active force when muscles are stretched at a very high speed and is thought to be caused by a disruption in cross-bridge cycling (Griffiths et al., 1980). The purpose of the present study was to further investigate the role of cross-bridge cycling in rFE by testing whether fast stretching in muscle fibres that disrupts cross-bridge cycling is associated with a loss of the rFE property. We hypothesized that rFE will be vastly reduced or abolished after fast active stretching of skeletal muscle preparations for stretch magnitudes that under normal, slow stretch conditions produce substantial rFE.

To test the hypothesis stated above, experiments were performed using skinned muscle fibre bundles from female New Zealand white rabbit (12–14 months old) psoas muscles. Rabbits were euthanized by pentobarbital (2 ml 4.5 kg⁻¹ body mass) through an intravenous injection. We used a disposable scalpel blade to open the abdomen region of the rabbit, and strips of muscle were dissected and tied onto wooden sticks (Leonard and Herzog, 2010). Muscle strips were stored in a rigor solution for 24 h and transferred to a rigor:glycerol (50:50) solution containing protease inhibitors for skinning at –20°C. Connective tissues and membranes were permeabilized by the rigor:glycerol solution for 10 to 14 days before use in experiments. Relaxing solution (no calcium) and activating solution (high calcium concentration) containing adenosine triphosphate (ATP) were used to deactivate and activate the muscle bundles, respectively, during the experimental protocols (Joumaa et al., 2008).

On the day of experiments, small muscle bundles (mean±s.e.m.: length=1.70±0.06 mm and diameter=317±12 µm) were isolated from the skinned muscle strips and incubated in the relaxing solution. Each end of the muscle bundle was attached to a hook connected to a force transducer (model 400B, Aurora Scientific, ON, Canada) and a length controller (Aurora Scientific, model 322C), respectively. Experiments were performed at 4°C.

Fibre bundles were set at an average sarcomere length (SL) of 2.4 µm using laser diffraction (Joumaa and Herzog, 2013). Fibre bundle diameters were then measured, and cross-sectional areas were calculated assuming a cylindrical shape. Forces were then normalized to cross-sectional area to obtain engineering stress.

Twelve fibre bundles were tested for the fast and slow stretch protocols. Each fibre bundle was exposed to two stretch experiments. In one stretch experiment, bundle stretching was slow (rFE after slow stretching), and in the other, stretching was fast (rFE after fast stretching) to induce cross-bridge slippage. The fast and slow stretching experiments were performed in a random order. To confirm that our results were reproducible, we repeated the slow stretch protocol after the fast stretch for the experiments in which the slow stretch was performed before the fast stretch. Slow rFE was reproduced in all tested bundles. The fibre bundles were given a rest period of 5 min between tests.

Fibre bundles were passively stretched from an average SL of 2.4 µm to an average SL 3.2 µm, then activated and held at steady state for 200 s. Note that there is no fatigue (loss of force) in skinned fibre preparations over the 200 s experimental period (Fig. 1) as the calcium handling is controlled within the activating solution.

Fibre bundles were activated at an average SL of 2.4 µm and actively stretched to an average SL of 3.2 µm in 10 s (0.08 µm s⁻¹). The steady-state isometric force following the stretch was compared with the reference force at corresponding times to determine rFE.

The protocol for this reference contraction was the same as the one described for rFE after slow stretching.

Fibre bundles were activated at an average SL of 2.4 µm and actively stretched to an average SL of 3.2 µm in 1 ms (800 µm s⁻¹) (Griffiths et al., 1980). The steady-state isometric force reached following the stretch was compared with the reference force to determine rFE.

Reference isometric forces and forces after slow or fast active stretches were recorded and compared. rFE was determined as the difference between the steady-state force obtained following active stretch and the corresponding reference force, and was expressed as a percentage of the reference force. The Wilcoxon signed rank test was used to identify differences in rFE between the slow and fast stretch conditions. Statistical significance was set at 0.05.

Stress at isometric steady state following the slow stretch condition (95.4±6.5 kPa, with a force value of 7.3±0.3 mN) was greater than for the corresponding isometric reference contraction (69.0±4.1 kPa, 5.3±0.2 mN), resulting in rFE of 38.1±3.7% (Figs 1 and 2). For the fast stretch condition, the isometric reference stress was 62.4±4.9 kPa (4.7±0.2 mN) and after the fast stretch, the isometric stress was 58.8±4.8 kPa (4.5±0.2 mN), similar to the isometric reference stress (P=0.071), resulting in the absence of rFE (Figs 1 and 2).

The purpose of this study was to investigate the potential role of cross-bridge cycling in rFE by testing whether fast stretching of muscle fibres that disrupts proper cross-bridge cycling with actin is associated with a decrease or loss of the rFE property. Our main finding was that slow active stretching (with an average SL stretch rate of 0.08 µm s⁻¹) resulted in a substantial rFE of 38.1±3.7%, whereas fast active stretching (with an average SL stretch rate of 800 µm s⁻¹) abolished rFE. This result indicates that proper cross-bridge kinetics during the stretch phase of muscle contraction is required to produce the molecular conditions that allow for rFE to occur.

Since the discovery and systematic description of rFE in 1952 (Abbott and Aubert, 1952), the mechanism underlying rFE has been the subject of intense scientific debate (Huxley, 1980; Edman et al., 1982; Morgan, 1994; Herzog and Leonard, 2002; Edman, 2012; Seiberl et al., 2015). It has been suggested that rFE may have two components: an active component associated with cross-bridge kinetics, and a passive component associated with titin (Rassier and Herzog, 2004). The active component has been thought to be associated with changes in the conformation of the contractile filaments, which give rise to changes in cross-bridge kinetics and the force produced per cross-bridge without changing the proportion of attached cross-bridges (Joumaa et al., 2021). The passive component has been associated with a change in the stiffness of titin, either by calcium binding to titin (Yamasaki et al., 2001; Labeit et al., 2003; DuVall et al., 2013), or by titin's proximal segment interacting with actin (Herzog and Leonard, 2002; Rode et al., 2009; Leonard and Herzog, 2010; Powers et al., 2014; Nishikawa et al., 2012; Zhou et al., 2021). However, like the active component of rFE, the passive component has been thought to depend on proper cross-bridge cycling (Leonard and Herzog, 2010; Fukutani et al., 2019).

Exposure of myofibrils to BDM and troponin C deletion have been used to identify the contribution of calcium-triggered titin–actin interactions in the absence of cross-bridge formation on rFE (Joumaa et al., 2008; Powers et al., 2014). BDM allows for weak but not strong cross-bridge binding to actin (Herrmann et al., 1992), thus disrupting the normal cross-bridge kinetics and not allowing for force production via actin–myosin interactions. Troponin C deletion prevents calcium from attaching to the troponin complex, thereby completely supressing cross-bridge formation during muscle activation. When strong cross-bridge binding to actin is inhibited with BDM or troponin C, titin stiffness increases imperceptibly resulting in a minute titin-based force enhancement (Joumaa, et al., 2008; Leonard and Herzog, 2010; Powers et al., 2014) that cannot explain the substantial rFE observed in preparations where cross-bridge cycling occurs normally. This observation led to the hypothesis that strong cross-bridge binding may be required to allow for significant amounts of rFE.

Results from previous research suggested that the magnitude of rFE is independent of the speed of stretch (Edman et al., 1982). However, Fukutani et al. (2019), working with the cat soleus muscle, performed experiments using a wide range of stretching speeds, varying from 2 to 64 mm s⁻¹, to re-examine the effect of speed of stretching on rFE. They found that at the high speeds of 32 and 64 mm s⁻¹, force slippage occurred, which coincided with a significant reduction in rFE. Slippage can be observed as the abrupt drop in force during quick muscle stretching that results in force at the end of stretch that is often smaller than the isometric force at the corresponding length (Fukutani et al., 2019). When a muscle is stretched fast and over a sufficient range, attached cross-bridges are torn forcibly from actin (Huxley, 1957; Kuhn, 1978; Griffiths et al., 1980). Our results on skinned fibre bundles are consistent with the findings by Fukutani et al. (2019). When fibre bundles were stretched from 2.4 to 3.2 µm in 1 ms (567 mm s⁻¹, for our bundles of an average length of 1.7 mm), rFE was abolished, whereas active stretches in 10 s (0.0567 mm s⁻¹) produced significant rFE (38%). Our results and those of Fukutani et al. (2019) support the idea that complete cross-bridge cycling, including strong cross-bridge binding to actin, is required to induce rFE.

The abolishment of rFE after rapid active stretching might be associated with the active and passive origins of rFE. According to the cross-bridge theory (Huxley, 1957), the active origin of rFE could be associated with changes in cross-bridge kinetics, resulting in an increase in the proportion of attached cross-bridges and/or an increase in the force produced per cross-bridge. Joumaa et al. (2021) used the X-ray diffraction technique to provide further insight into the cross-bridge conformation at steady state following active stretch in skinned rabbit psoas muscle bundles. They found that rFE was not accompanied with an increase in the proportion of attached cross-bridges, but with changes in the conformation of cross-bridges, possibly leading to an increase in the force produced per cross-bridge. This finding is consistent with Huxley and Simmons (1971), who suggested that active stretch alters the conformation of cross-bridges. It seems that this stretch-caused alteration in cross-bridge conformation could last beyond the transient phase of the stretch and remain at the steady-state reached following the stretch. Using very rapid active stretching and associated mechanical rupture of cross-bridges from actin, the changes in cross-bridge kinetics usually observed with slow and physiologically relevant active stretching might be lost, resulting in the return of cross-bridges to their un-stretched state, and thus abolishing the increase in force following fast active stretching.

The passive component of rFE has been associated with titin. It has been reported that calcium activation changes the stiffness of titin (Yamasaki et al., 2001; Labeit et al., 2003), thus possibly contributing to the rFE (Joumaa et al., 2008). It has also been suggested that titin might bind to actin during muscle activation and stretching, thereby decreasing its free spring length and increasing its force in actively stretched muscles (Leonard and Herzog, 2010; DuVall et al., 2017; Dutta et al., 2018). The mechanism for possible titin–actin interaction remains unclear, and proposed mechanisms of a calcium/activation regulated binding of titin to actin are speculative and not supported by experimental research using intact muscle, fibre or sarcomere preparations (e.g. Joumaa et al., 2008; Leonard and Herzog, 2010). In the absence of direct experimental evidence for the role of titin in rFE in structurally intact preparations, except possibly for Duvall et al. (2017), and supported by the current findings, we would like to put forward a hypothesis that we hope might encourage experimental testing. (i) Strong cross-bridge binding is required to elicit a muscle's rFE property. (ii) When strong cross-bridge binding is inhibited, for example via troponin C deletion, BDM administration or super-fast stretching, rFE is abolished. (iii) Strong cross-bridge binding is known to move the regulatory protein tropomyosin from its position in the passive and weakly bound cross-bridge state to a new position (Gordon et al., 2001). (iv) This movement of tropomyosin exposes binding sites for titin on actin, thereby permitting titin binding to actin when cross-bridges are allowed to engage in strongly bound states, but not when strong cross-bridge binding is inhibited or disrupted, as we have done here with the super-fast stretch experiments.

Fast stretching of active muscles abolished rFE. We conclude that fast muscle stretching interrupts cross-bridge cycling, specifically strong cross-bridge binding. We propose that the tropomyosin relocation on actin, associated with strong cross-bridge binding, exposes attachment sites for titin on actin, thus allowing titin–actin binding, which results in a shortened free titin spring length and increased titin stiffness, and is the mechanism that produces rFE when active muscles are stretched at physiologically relevant speeds.

The authors acknowledge Dawn Martin for helping with tissue collection.