ATP provides the energy in our muscles to generate force, through its use by myosin ATPases, and helps to terminate contraction by pumping Ca2+ back into the sarcoplasmic reticulum, achieved by Ca2+ ATPase. The capacity to use ATP through these mechanisms is sufficiently high enough so that muscles could quickly deplete ATP. However, this potentially catastrophic depletion is avoided. It has been proposed that ATP is preserved not only by the control of metabolic pathways providing ATP but also by the regulation of the processes that use ATP. Considering that contraction (i.e. myosin ATPase activity) is triggered by release of Ca2+, the use of ATP can be attenuated by decreasing Ca2+ release within each cell. A lower level of Ca2+ release can be accomplished by control of membrane potential and by direct regulation of the ryanodine receptor (RyR, the Ca2+ release channel in the terminal cisternae). These highly redundant control mechanisms provide an effective means by which ATP can be preserved at the cellular level, avoiding metabolic catastrophe. This Commentary will review some of the known mechanisms by which this regulation of Ca2+ release and contractile response is achieved, demonstrating that skeletal muscle fatigue is a consequence of attenuation of contractile activation; a process that allows avoidance of metabolic catastrophe.
Typically, when exercise scientists think of control of skeletal muscle contraction, they consider motor unit recruitment and rate coding, the primary means by which the brain regulates the magnitude of muscle contraction. However, the magnitude of skeletal muscle contraction can also be regulated at the cellular level. This level of control is necessary to avoid cellular metabolic catastrophe, i.e. the situation in which ATP depletion reaches a level that is damaging to the fiber.
ATP is the common final source for energy in the cell, and its concentration [along with the concentrations of ADP and inorganic phosphate (Pi)] affects the magnitude of energy that is made available from its hydrolysis. At rest, skeletal muscle has a low metabolic rate, not unlike many other passive tissues, and the intracellular concentration of ATP is in the 5–7 mM range (Hochachka and Matheson, 1992). However, during muscle activity, there are three major ATPases that require ATP for their function (Box 1); during exercise, skeletal muscle can increase its rate of ATP use by more than 100 times (Hochachka and McClelland, 1997), a rate that is much faster than can be replenished by aerobic metabolism. This latter point is clear from the fact that the muscle power output can greatly exceed the level that can be supported by aerobic metabolism (MacIntosh et al., 2000). To accommodate this additional energy requirement, ATP is provided by non-aerobic means (through phosphocreatine hydrolysis and glycolysis resulting in lactate formation), but this alternative source of ATP has limited capacity (Monod and Scherrer, 1965). For this reason, a high rate of ATP use cannot be sustained in muscle and ATP use must thus be regulated (Hochachka, 1994; Hochachka and Matheson, 1992; Myburgh, 2004). We know that muscle fibers can preserve ATP because the ATP level usually decreases by only 20–25% during exercise (Argov et al., 2000; Dawson et al., 1978) and rarely falls below 50% of its initial value (Karatzaferi et al., 2001; Spriet et al., 1987).
We have proposed that the cellular regulation of ATP use is the principal mechanism for the decreased contractile response that is associated with acute repetitive muscle use, typically called muscle fatigue (MacIntosh and Shahi, 2011). We also have recognized that regulating Ca2+ release from the sarcoplasmatic reticulum (SR), and thus the intracellular Ca2+ concentration ([Ca2+]i), dictates the rate of ATP use in the muscle cell because it affects Ca2+ ATPase and myosin ATPase activity, which together account for 90% of ATP use (Box 1). It should be kept in mind that a multitude of highly redundant factors contribute to this regulation of ATP usage. We will not present all of these here. Instead, this Commentary concentrates on the following three major areas: (1) regulation of the contractile proteins actin and myosin by Ca2+ and myosin light chain phosphorylation, (2) regulation of membrane excitability through K+ and Cl− channels, and (3) the regulation of Ca2+ availability and the SR Ca2+ release channel. A summary of the process of excitation–contraction coupling is presented in Fig. 1 and described in detail in Box 1. This figure gives the context for the remaining discussion in this Commentary.
Regulation of the interaction between actin and myosin
Muscle contraction is the consequence of the interaction between actin and myosin. When the myosin head binds to actin, forming
Box 1. Excitation–contraction coupling and ATP usage
Contractions in skeletal muscle are triggered when action potentials are generated in the outer cell membrane (Fig. 1). Under normal conditions, an action potential consists of a depolarization from a resting membrane potential (Em) of −80 mV to +30 mV, followed by a repolarization, returning Em to its resting value. The depolarization depends on Na+ influx through voltage-sensitive Na+ channels (Nav), and the repolarization involves K+ efflux through voltage-sensitive K+ channels (Kv). Each action potential propagates along the surface membrane and into the transverse tubules (Fig. 1). Depolarization of the transverse tubules is detected by voltage-sensitive Ca2+ channels (Cav), also known as dihydropyridine receptors (DHPRs), and these communicate directly with Ca2+ channels, also known as ryanodine receptors (RyRs), to release Ca2+ from the sarcoplasmic reticulum. At low [Ca2+]i, tropomyosin blocks the myosin-binding site on actin. When [Ca2+]i increases in response to an action potential, it binds to troponin and a segment of actin, seven monomers long, becomes available to myosin as one tropomyosin is moved away from the myosin-binding site on actin. The number of such available segments is a function of the [Ca2+]i. The subsequent binding of myosin to actin generates force and/or shortening of sarcomeres.
There are three major proteins that contribute to the increased ATP use during contraction: Na+/K+-ATPase, myosin ATPase and Ca2+ ATPase (Fig. 1). The Na+/K+-ATPase pumps Na+ back out and K+ back into the fiber after an action potential. The myosin ATPase uses ATP to generate force and do work, whereas the Ca2+ ATPase is responsible for pumping Ca2+ back into the SR to allow muscle relaxation. Each of these ATPases, contributes ~10, ~60 and ~30% of total ATP use, respectively (Homsher, 1987).
a cross-bridge, it undergoes a sequence of reactions referred to as the cross-bridge cycle (Box 2). The force of isometric contraction is primarily a function of the number of cross-bridges that are engaged simultaneously in the strong-binding state. This number is governed, at a given sarcomere length, by the proportional occupation of troponin with Ca2+ and the rate constants for the transition to the strong-binding state, as well as the dissociation of the cross-bridge. Troponin is a regulatory protein associated with the thin filament. As explained in Box 1, binding of Ca2+ to troponin allows cross-bridge cycling to occur.
When force is plotted as a function of [Ca2+], a sigmoid curve is obtained. At very low [Ca2+], no force is generated (Fig. 2). Resting [Ca2+] is in this range. Next, there is a range for which increases in [Ca2+] are associated with increases in force, as additional binding of Ca2+ to troponin reveals an increased number of actin myosin interaction sites. This range of [Ca2+] is referred as submaximal [Ca2+]. Finally, there is a range, for which increases in [Ca2+] produce no further increase in force because, in that range, all Ca2+-binding sites of troponin are occupied; i.e. the contractile apparatus is maximally activated. The [Ca2+], at which force is half-maximum is known as Ca50 and reflects the Ca2+ sensitivity of the contractile components. A decrease in Ca50 is observed when the force–[Ca2+] curve is shifted towards lower [Ca2+] and represents an increased Ca2+ sensitivity.
There are two primary mechanisms that affect Ca2+ sensitivity, altered binding of Ca2+ to troponin and altered cross-bridge kinetics. Cross-bridge kinetics is the more dominant of these mechanisms and its kinetics is affected by several factors, including: temperature, pH, sarcomere length, myosin regulatory light chain (RLC) phosphorylation, and concentration of Pi. How these factors affect cross-bridge kinetics and Ca2+ sensitivity has been discussed in detail elsewhere (MacIntosh, 2003). Here, we will discuss the mechanism for increased Ca2+ sensitivity that is affected by myosin RLC phosphorylation.
Myosin RLC phosphorylation increases Ca2+ sensitivity
There are two myosin light chains (small peptides) associated with each myosin head. One of these light chains, the RLC, can be phosphorylated by myosin light chain kinase (MLCK). In resting muscle, the RLCs of myosin are mostly unphosphorylated (MacIntosh et al., 1993). When [Ca2+]i increases during contraction, Ca2+ binds to another small peptide, calmodulin, and the Ca2+–calmodulin complex subsequently binds to MLCK, thereby activating it. Therefore, the repetitive activation of muscle leads to a progressive phosphorylation of RLCs (MacIntosh et al., 1993). MLCK is eventually deactivated when [Ca2+]i decreases to resting levels and RLC is dephosphorylated over the course of a few minutes by the slow activity of myosin light chain phosphatase (Hartshorne et al., 1998).
When muscles are elicited to contract with a single stimulus, force rapidly rises then falls in a contraction called a twitch. Peak twitch force represents about 5–20% of the maximum force a muscle can generate when stimulated at high frequency (>150 Hz, yielding a completely fused tetanic contraction). When a twitch is elicited immediately after a 1-second long tetanus, the peak force is much higher than that measured before the tetanic contraction; this phenomenon is known as post-tetanic twitch potentiation. When muscles are stimulated at frequencies below 10 Hz, each subsequent contraction develops more force for the first several seconds, a phenomenon known as staircase. During staircase and post-tetanic potentiation, the twitch peak rate of force development increases proportionally to the increase in active force, whereas the timecourse of the twitch is not altered (Hainaut and Desimedt, 1968; MacIntosh and Gardiner, 1987). Both staircase and post-tetanic twitch potentiation are thought to be due to RLC phosphorylation (MacIntosh et al., 1993; MacIntosh and Rassier, 2002; Moore and Stull, 1984). Skinned fiber experiments have confirmed that RLC phosphorylation, as achieved with addition of MLCK and calmodulin, results in increased Ca2+ sensitivity (Persechini et al., 1985), which now has been related to changes in cross-bridge kinetics, as explained below.
Sweeney and Stull (Sweeney and Stull, 1990) addressed the question of whether potentiation relies on an increased number of engaged cross-bridges or an increased force per cross-bridge. They used the quick longitudinal stretch of the muscle to measure stiffness, which is assumed to be proportional to the number of cross-bridges engaged in the strong-binding state. They reported that longitudinal stiffness increases in proportion to the increased active force at submaximal [Ca2+]i when myosin light chains are phosphorylated by MLCK. This was interpreted as indicating that potentiation is a function of increased number of cross-bridges engaged, not an increase in force per cross-bridge. They also reported that myosin ATPase activity increases in proportion to the increased active force, indicating that the cycle time per cross-bridge is not altered. This allowed them to conclude that the increase in the number of cross-bridges is a function of an increased rate of engagement, not a decrease in the rate of dissociation of cross-bridges (Sweeney and Stull, 1990).
Levine and colleagues have shown that RLC phosphorylation is associated with a change in the way in which the myosin heads are ordered along the thick filament backbone (Levine et al., 1996). In the unphosphorylated state, the myosin heads are packed tightly along the thick filament. However, phosphorylation of the RLC results in disorder of the myosin heads. This disorder was interpreted as an increased mobility, such that the myosin head can swing away from the thick filament, bringing the myosin head into close proximity to actin and, hence, increasing the probability of its interaction with actin.
So, the regulation of [Ca2+]i during contraction and phosphorylation of myosin light chain can have significant impact on ATP usage. Although RLC phosphorylation is typically associated with potentiation, this phosphorylation persists when fatigue begins (MacIntosh et al., 1993). The same Ca2+–calmodulin complex that activates MLCK also inhibits Ca2+ release (Boschek et al., 2007; Xiong et al., 2006). The muscle is capable of decreasing the release of Ca2+, while
Box 2. Cross-bridge cycle
In the resting state, most of the myosin heads exist in a primed state [step (ii) in the figure]; that is, with the ATP partially hydrolyzed and the myosin head cocked, ready for the power stroke. Although it is thought that a weak binding of the myosin head to actin can occur, even when the tropomyosin is in the ‘blocked’ position [step (i) in the figure], the binding of Ca2+ to troponin moves tropomyosin to the ‘closed’ position, thereby increasing the probability of this weak binding and permitting the rapid transition from weak to strong binding. The existence of one myosin head in the strong binding state causes the tropomyosin to move further from the blocked position to a position referred to as ‘open’. This is a form of cooperativity. The rate constant for the transition to strong binding is low without Ca2+ bound to troponin, but increases with binding of Ca2+ to troponin. On going from a weak-binding state to a strong-binding state [step (iii) in the figure], inorganic phosphate (Pi) is released from the myosin head and a swinging lever action generates force and/or permits relative motion of the thick and thin filaments. This strong-binding state will persist until ATP replaces the ADP that remains after the Pi is released. ATP binding permits dissociation of the myosin head from actin [step (iv) in the figure]. It is generally accepted that each cycle as described here results in the hydrolysis of one ATP molecule (Barclay, 2003; Gordon et al., 2000).
preserving its contractile function through increasing Ca2+ sensitivity. Such a mechanism helps in preserving ATP and prolonging the duration of any muscle activity. The importance of this process is further illustrated by the fact that muscle activation by motor neurons is at frequencies (Hennig and Lømo, 1985) that do not give rise to maximal increases in [Ca2+]i and full force development (Westerblad and Allen, 1991); i.e. most of the time [Ca2+]i is submaximal.
Analysis of the force–Ca2+ relationship also suggests that the attenuation of Ca2+ release and thus lower [Ca2+]i can also be an effective way of decreasing the activity of myosin ATPases as well as Ca2+ ATPases. Myosin ATPase activity decreases because less Ca2+ will bind to troponin, and therefore there will be a reduced level of myosin–actin interaction. A reduction in Ca2+ ATPase activity occurs because there would be less Ca2+ to pump back into the SR. Decreased Ca2+ release can therefore be an effective way to preserve ATP during a metabolic stress. There is considerable evidence that a reduction in Ca2+ release does occur during repeated muscle activation, either through reduced Ca2+ availability (Allen et al., 2011; MacIntosh and Kupsh, 1987) or by regulating the process of its release (Li et al., 2002). In the next sections, we will look at how changes in membrane excitability affect Ca2+ release. This is followed by a discussion of the regulation of the ryanodine receptor (RyR), which is the Ca2+ release channel of the SR.
Control of membrane excitation
As mentioned above, the release of Ca2+ is triggered by an action potential propagating through transverse tubules (Fig. 1). The magnitude of Ca2+ release can be modulated by the amplitude and shape of the action potential. Shape and amplitude are affected by both ion concentration gradients and by modulation of ion channel activity. During metabolic stress, it is desirable to decrease Ca2+ release. Here, we will present the mechanisms by which the cell modifies the amplitude and shape of the action potential to decrease Ca2+ release. Through the discussion below, we will see that some of the changes are meant to preserve contractile function, but when ATP levels are threatened, the changes result in decreased excitability, decreased Ca2+ release and preservation of ATP.
To facilitate an understanding of the impact of ion concentration gradients and permeabilities on the action potential, it is worthwhile to begin with the resting condition. The resting state of the cell and the changes in ion permeability that occur during the action potential are presented in Box 3.
The role of K+ in membrane excitability
At rest, the normal extracellular [K+] ([K+]e) varies between 3.5 and 4.0 mM. However, this increases to 10–14 mM during muscle activity (Juel et al., 2000; Mohr et al., 2004; Nielsen et al., 2004; Street et al., 2005). In vitro at 37°C, maximal tetanic force is not affected until the [K+]e reaches 10 mM; at or above that concentration, tetanic force decreases (Cairns et al., 2011). Twitch and submaximal tetanic forces, are potentiated when [K+]e is increased up to 12 mM (Holmberg and Waldeck, 1980), whereas above that concentration twitch force is depressed (Yensen et al., 2002). The mechanism of the K+-induced potentiation is unknown, but is no doubt beneficial for maximizing muscle performance at the onset of exercise. The K+-induced force depression, on the other hand, is due to a K+-induced depolarization of the cell membrane resulting in decreased action potential overshoot as Na+ channels are inactivated (Yensen et al., 2002) (Fig. 3; Box 3). There is indirect evidence that the amount of Ca2+ that is released by SR is reduced once the action potential overshoot decreases below 5 mV; twitch force is unaffected when overshoot varies between 5 and 30 mV (Cairns et al., 2003; Yensen et al., 2002). It is also important to note that the force–[K+]e relationship is dynamic; i.e. the concentration at which K+ potentiates or depresses force is modulated by several factors. Two of these factors are the activities of the voltage-dependent chloride channel protein (ClC-1, also known as CLC1 and CLCN1) and the ATP-sensitive K+ (KATP) channel, which are discussed in the following two sections. Ion channel activity can influence action potential amplitude because the amplitude of the action potential is dependent not only on the resting membrane potential but, more importantly, also on the exchange of ions during the action potential through channels in addition to voltage-gated Na+ and K+ channels (Nav and Kv, respectively), such as the Cl− and the KATP channels (Box 3).
Regulation and impact of the ClC-1
ClC-1 is the main channel in skeletal muscle by which Cl− crosses the cell membrane; this channel therefore controls the muscle membrane Cl− permeability. In the resting state, Cl− permeability largely exceeds that of K+ (Palade and Barchi, 1977; Pedersen et al., 2009a; Pedersen et al., 2005; Pedersen et al., 2009b) and plays a major role in the maintenance of resting membrane potential (Em). For example, when [K+]e increases, the resulting membrane depolarization is slower and smaller when Cl− is present in the muscle bathing solution than in its complete absence (Dulhunty, 1978). This observation indicates that inward
Box 3. Action potential and its modulation by Cl2 and K+ permeability
Transmembrane movement of ions is affected by three factors, ion permeability, concentration gradient and membrane potential (Em). At rest, Em is about −80 mV and the cell membrane is impermeable to Na+, somewhat permeable to K+ and most permeable to Cl−. Na+ and Cl− are at a high concentration outside the membrane and a low concentration inside. [K+] is high inside and low outside. The action potential results from increased permeability to Na+ and K+, while Cl− permeability does not change. The depolarization phase involves the activation of Na+ channels (Nav) and the resulting influx brings Em to +30 mV. In normal conditions, this occurs despite a Cl− influx due to the predominance of Nav activity. Inactivation of Nav channels shortly after they open, and increasing activation of K+ channels (Kv) resulting in K+ efflux, and the continuing Cl− influx allow the repolarization of the membrane back to −80 mV. Repolarization of the membrane reprimes Nav channels, so they can be activated for another action potential. However, small membrane depolarizations (i.e. less than 15 mV) at rest (e.g. an increased [K+]e during muscle activity) inactivates a portion of the Nav channels, decreasing their availability. As a consequence of lower Nav channel availability there is less predominance of the Na+ inward movement during the depolarization phase. Now, any changes in Cl− and K+ channel activity will substantially impact the rate of depolarization and the amplitude of action potentials. For example, closure of Cl− channels will decrease the Cl− influx during depolarization allowing for faster and greater depolarization. By contrast, activation of Cl− and K+ channels (to increase Cl− influx and K+ efflux) will slow down the depolarization and decrease the action potential amplitude.
movement of Cl− counteracts the depolarizing effects of the accumulation of K+ outside the cell.
ClC-1 can, under some conditions, directly influence the amplitude of the action potential (Box 3). Under normal conditions (i.e. 4 mM K+), removal of extracellular Cl− does not affect the kinetics of an action potential (Cairns et al., 2004), because Na+ and K+ fluxes largely exceed that of Cl−. However, the same is not true at high [K+]e. At 11 mM K+, ~50% of soleus fibers are completely unexcitable and the remaining fibers generate action potentials with peaks that only reach −10 mV (Pedersen et al., 2005). Reducing Cl− permeability by 50% allows for a recovery of membrane excitability; the number of fibers generating action potentials increases to 95% and action potential peak reaches +10 mV. This recovery occurs because at 11 mM K+, a large number of Na+ channels are inactivated resulting in a considerably reduced Na+ influx during the action potential compared with that at 4 mM K+. This inactivation of Na+ channels allows the Cl− influx to strongly counteract the smaller Na+ depolarizing effect, decreasing the action potential amplitude. Lowering Cl− permeability by closing Cl− channels lowers Cl− influx, thus diminishing its ability to counteract the Na+ depolarization and allowing for greater action potential amplitude. Associated with the increase in membrane excitability is an increase in force production (Pedersen et al., 2005). Therefore, Cl− has two opposing effects on membrane excitability and force when the [K+]e is elevated; a complete removal of Cl− from the extracellular fluid exaggerates the K+-induced membrane depolarization, loss of membrane excitability and, subsequently, force, whereas the small decreases in Cl− permeability that are achieved when up to 50% of the Cl−channels are closed improves membrane excitability and force generation.
The modulation of the effect of K+ by regulation of ClC-1 discussed above is based on studies that use resting unfatigued muscle fibers. If the observed modulation has any physiological significance, one would expect that there would be some regulation of the Cl− permeability by controlling the ClC-1 activity. In fact, during repeated contractions in rat extensor digitorum longus (EDL) fibers, Cl− permeability initially decreases by 50% within 1 minute (Pedersen et al., 2009a). As the stimulation is prolonged, Cl− permeability remains constant for 3 minutes and then suddenly increases by threefold above the pre-stimulation level, corresponding with opening of a very large number of ClC-1 channels. Associated with the increase in Cl− permeability is a decrease in membrane excitability (Pedersen et al., 2009a). Thus, ClC-1 is controlled during muscle activity.
Here, we consider an exercise example to illustrate the impact of the regulation of Cl− channels. Interstitial [K+] reaches 10–11 mM within 5 minutes during 30-watt single leg extension exercise (Nielsen et al., 2004). Such [K+] largely depresses tetanic force in vitro (Cairns et al., 2011; Pedersen et al., 2003) and yet the exercise continues without difficulty for 30 minutes. Perhaps, the decreased Cl− permeability at the onset of muscle activity is essential in preventing any K+-induced force depression, while at the same time favoring a K+-induced force potentiation (Fig. 3). In other words, the combined increase in [K+]e and decrease in Cl− permeability might help to maximize muscle performance at the onset of muscle activity. Conversely, any increase in Cl− permeability might be linked to a fatigue process triggered by a metabolic stress to increase muscle sensitivity to the K+-induced force depression (Pedersen et al., 2009b). In the latter situation, membrane excitability decreases, thereby lowering Ca2+ release and force generation (i.e. fatigue) in order to lower the activity of the Ca2+ and myosin ATPases. The ultimate benefit is a reduced ATP demand, avoiding metabolic catastrophe.
Regulation and impact of the KATP channel
The KATP channel is an ATP-sensitive K+ channel. The channel was named after it was discovered that ATP binding closes the channel (Noma, 1983). It is now established that KATP channels are primarily activated by the changes in metabolite levels that occur during metabolic stress, including decreases in intracellular ATP and pH, and increases in intracellular ADP and extracellular adenosine (Barrett-Jolley et al., 1996; Davies, 1990; Noma, 1983; Vivaudou et al., 1991). The KATP channel is thus considered to be an energy sensor that is activated during metabolic stress. Being an ion channel, it also links the excitability of the sarcolemma to the metabolic state of the fiber. There is now clear evidence that activation of KATP channels during repetitive stimulation coincides with the activation of Cl− channels as discussed above (Pedersen et al., 2009a), and that the channel is crucial in preventing fiber damage and severe muscle dysfunction during exercise and fatigue by decreasing muscle cell excitability (Cifelli et al., 2008; Cifelli et al., 2007; Kane et al., 2004; Stoller et al., 2009; Thabet et al., 2005).
One mechanism by which the KATP channel prevents fiber damage was revealed by studies in which channel openers were used, which allows for a large K+ efflux during the action potential (Box 3) and results in a reduction in the action potential amplitude (Gong et al., 2003; Matar et al., 2000). The resulting decreases in membrane excitability and action potential amplitude lead to a decrease in Ca2+ release and force (Burton and Smith, 1997; Duty and Allen, 1995; Gong et al., 2003; Matar et al., 2001; Weselcouch et al., 1993; Wickenden et al., 1996). The physiological benefit of a reduced Ca2+ release would be reduced activity of Ca2+ ATPases and myosin ATPases, thereby preserving ATP (Fig. 3). Indeed, decreases in ATP levels during metabolic inhibition are faster in the presence KATP channel blockers (Gramolini and Renaud, 1997; Weselcouch et al., 1993). However, modulation of KATP channel activity during muscle fatigue has no effect on ATP levels in mouse EDL compared with that in control conditions, possibly as other regulatory mechanisms prevail. Nevertheless, in soleus muscle, blocking the KATP channel accentuates ATP loss compared with that in control conditions, whereas channel activation prevents a reduction of ATP (Matar et al., 2000). These results therefore support the hypothesis that KATP channels affect energy metabolism, but more studies are necessary to fully understand how exactly these effects prevent fiber damage and muscle dysfunction.
A second mechanism by which KATP channels prevent fiber damage and dysfunction is through the maintenance of resting Em. This finding came from studies in which the KATP channel activity is abolished by either exposing normal muscles to channel blockers or by using muscles from Kcnj11-null mice [mice in which the Kcnj11 (Kir6.2) gene that encodes the channel pore protein has been deleted]. Normally, resting Em depolarizes at between 10 and 15 mV during repeated muscle contractions (Cifelli et al., 2008; Comtois et al., 1995; Light et al., 1994), whereas it does not change during metabolic inhibition (Gramolini and Renaud, 1997). In the absence of KATP channel activity, the depolarization can be as high as 50 mV under both conditions. This depolarization has been shown to be sufficiently large enough to activate the Ca2+ channels in transverse tubules, leading to an uncontrolled Ca2+ influx, large increases in resting intracellular Ca2+ and in force (Cifelli et al., 2008; Cifelli et al., 2007; Gong et al., 2000; Light et al., 1994; Matar et al., 2000). As a consequence of the increases in resting [Ca2+], there is greater use of ATP by the Ca2+ and myosin ATPases, which worsen the possibility of a metabolic catastrophe. Furthermore, chronic elevation of [Ca2+]i is a well known factor that causes fiber damage (Jackson et al., 1984; Jones et al., 1984).
Thus, the regulation of membrane excitability is complex and depends not only on changes in extra- and intra-cellular ion concentrations, such as that of K+, but also on changes in ion channel activity. At the onset of exercise, [K+]e increases rapidly and is likely to potentiate force generation similar to that upon an increased RLC phosphorylation. At the same time, the activity of ClC-1 decreases, possibly to prevent any K+-induced force depression. When metabolic stress occurs, the activity of ClC-1 increases and this results in a depression of membrane excitability, which is further enhanced by a concomitant activation of KATP channels. As membrane excitability is reduced by these two channels, less Ca2+ is released and less force is generated. The outcome of these mechanisms is the preservation of the limited supply of ATP through reducing its use by the Ca2+ and myosin ATPases.
Control of Ca2+ release
Action potentials on the membrane are translated into a release of Ca2+ in a process referred to as excitation–contraction coupling (Box 1). The dihydropyridine receptors (DHPRs) are specialized channels that detect the action potential in the transverse tubule membrane (Fig. 1) and, through a direct connection (in skeletal muscle), open and close the Ca2+ release channels in the SR, the RyRs. The release of Ca2+ into the myofibril initiates muscle contraction and, as a result, increases ATP demand through the activity of the myosin and Ca2+ ATPases. Although an action potential is the main controlling factor detected by the DHPR, and therefore the main determinant that results in opening and closing of the RyR, this ion channel is further regulated by a large number of ionic and protein ligands that influence the opening probability (Po). We will examine further a small well-known collection of these ligands and how they prevent ATP depletion. It is also known that reactive oxygen species can modulate Ca2+ release through the RyR (Oba et al., 2002), but this will not be discussed further here.
Regulation of RyR by ATP, Ca2+ and Mg2+
The three most studied regulatory ligands of the RyR are ATP, Ca2+ and Mg 2+ (Lamb and Stephenson, 1992; Laver et al., 1997; Meissner, 1984; Meissner et al., 1986). There are two distinct sites on RyR for these ligands that either enhance or reduce its Po. These sites are referred to as the high-affinity A-site, regulating activation, and the low affinity I-site, leading to inhibition (Lamb, 2000). Both Ca2+ and ATP bind to the A-site to increase the Po of RyR (Meissner, 1986), whereas both Ca2+ and Mg2+ bind to the lower affinity I-site to reduce its Po (Laver et al., 1997; Meissner et al., 1986). In the absence of Mg2+, Ca2+ opens RyR by binding to the A-site if it is present at a concentration of 1 μM and closes the receptor through binding at the I-site at high concentrations of 1 mM (Meissner et al., 1986). Under normal resting conditions with [Mg2+] being present at 1 mM, the I-site is occupied with Mg2+ and the subsequent inhibition of the channel overrides any activating effect of Ca2+ and ATP. Mg2+ also competes with Ca2+ for binding to the A-site, but this does not result in an activation of the channel (Laver et al., 1997). The strong activating effect of ATP is most clearly observed when Mg2+ is removed from the solution (Lamb and Stephenson, 1991). ATP, in the absence of external Ca2+, by itself is capable of fully activating RyR and initiating Ca2+ release (Meissner et al., 1986). This result further emphasizes how important the inhibitory effect of Mg2+ at rest is.
Considering the effects of Mg2+ described above, it would appear that the RyR would never be activated! However, with each action potential, some RyRs are activated and Ca2+ is released into the cell. The fact that Mg2+ reduces the Po of RyR becomes more important when the concentration of ATP decreases, because as shown in Fig. 1, a net ATP hydrolysis results in an increase in [Mg2+]. This occurs because Mg2+ has a lower affinity for ADP than for ATP. This increase in [Mg2+] and decrease in the concentration of ATP combine to cause a decrease in Po, attenuating Ca2+ release in subsequent activations.
Several studies have examined the effect of higher [Mg2+] on excitation–contraction coupling (Blazev and Lamb, 1999; Lamb and Stephenson, 1991; Laver et al., 1997; Owen et al., 1997). The general consensus of these studies is that [Mg2+] at rest is a strong inhibitor of RyR, which keeps the channel closed and unresponsive to the strong activating effect of ATP and Ca2+ until an action potential activates the cell. Ca2+-induced activation of RyR, a key mechanism in cardiac muscle, is apparently not relevant in skeletal muscle (Endo, 2009). At 3 mM [Mg2+], a concentration that can be reached during fatiguing contractions (Westerblad and Allen, 1992), the magnitude of Ca2+ release achieved by transverse tubule depolarization is reduced (Blazev and Lamb, 1999; Laver et al., 1997; Owen et al., 1996). Thus, inhibition by Mg2+ is important to prevent spontaneous Ca2+ release at rest and it provides another mechanism, by which less Ca2+ is released during metabolic stress.
The reduced activation of RyR, which results from low concentrations of ATP, has also been explored. When the concentration of ATP is held artificially low (<0.5 mM ATP), Ca2+ release is significantly reduced when RyR is directly activated by DHPR (Blazev and Lamb, 1999; Owen et al., 1996). Although this ATP decrease might appear extreme, it has been reasoned that a localized decrease in the concentration of ATP to below 1 mM could occur in the vicinity of RyR (Owen et al., 1996). Therefore, both the concomitant decrease in ATP concentration and increase in [Mg2+] upon an impending metabolic catastrophe can contribute to a decrease in Ca2+ release by lowering RyR Po to eventually preserve ATP. The sensitivity of the RyR to changes in [Mg2+] and ATP concentration, can also make it an energy sensor, in a similar manner to the KATP channel.
Regulation of RyR by calmodulin and the S100 Ca2+-binding protein A1 (S100A1)
There are several additional ligands that interact with RyR. Here we will discuss two that have known regulatory impact on Po of the RyR, calmodulin and S100 Ca2+-binding protein A1 (S100A1).
The small ubiquitous peptide calmodulin (CaM) was one of the first ligand regulators of RyR to be discovered (Chen and MacLennan, 1994; Hamilton et al., 2000; Yang et al., 1994). At rest, when [Ca2+] is low, calmodulin exists in the apocalmodulin (apoCaM) form, and upon increase in [Ca2+]i binds to Ca2+, becoming Ca2+-bound (CaCaM), and resulting in a change of its shape and its effect as a ligand (Boschek et al., 2007; Boschek et al., 2008; Xiong et al., 2006). At low [Ca2+]i (up to 0.1 μM), the Ca2+-free apoCaM binds to RyR and increases its Po. Conversely, as [Ca2+]i increases, the amount of the CaCaM complex also increases, which binds to RyR but decreases its Po (Boschek et al., 2007; Xiong et al., 2006).
RyR has many distinct sites for ligand binding (Box 1) (Song et al., 2011), although it will probably be many years before we fully understand the range of ligands for RyR and their functions. Recent work has provided some insight. One of the important ligands for RyR is S100A1. Prosser and colleagues used a knockout mouse (S100A1) to demonstrate that S100A1 enhances Ca2+ release (Prosser et al., 2008). Twitch and tetanic contractile amplitudes are attenuated, and the corresponding Ca2+ transients are smaller in the S100A1 mice compared with muscles of wild-type control mice. Yamaguchi et al. contributed to our understanding of ligand control of RyR by using a mouse with an altered RyR CaCaM-binding site (RyRD/D) (Yamaguchi et al., 2011). Similar to the S100A1 mice, the RyRD/D mice produce lower twitch forces, suggesting that they lack the S100A1-binding site. However, Ca2+ release during a maximal tetanic contraction is enhanced and there is a greater peak [Ca2+]i compared with that in muscles of wild-type mice (Yamaguchi et al., 2011). On the basis of these results, the authors suggest that there is one site on the RyR that binds both CaCaM and S100A1, and that with S100A1 bound, this site is important for promoting Ca2+ release at low [Ca2+], such as occurs during a twitch. The same site is also important for inhibiting Ca2+ release during repeated or sustained activation by binding CaCaM at higher [Ca2+]i. In this way, Ca2+ is able to slow energy expenditure later in contraction. It is interesting to note that CaCaM activates MLCK and inhibits Ca2+ release. These two processes will have opposing effects on muscle, enhancing contraction by promoting RLC phosphorylation and attenuating contraction by decreasing Ca2+ release.
This Commentary has aimed to illustrate, with a few examples, how skeletal muscle contraction is regulated at the cellular level. Although most researchers concentrate on the depression of contraction during fatigue, we point out here that there are regulatory processes that can enhance muscle performance. The first example is that of myosin RLC phosphorylation, which allows for increased force production at a given [Ca2+]i. This effect permits a decrease in Ca2+ release without any loss of force or power, but with the advantage of a reduced Ca2+ ATPase activity, as less Ca2+ needs to be pumped back into the SR. Overall, this effect might be important in reducing ATP usage and prolonging the muscle activity. The second example is that of K+-induced force potentiation that occurs when [K+]e increases. The underlying mechanism of this potentiation is still unknown. Finally, a third mechanism that helps maintain a high muscle performance at the onset of exercise is the decrease in Cl− channel activity; this effect is important to counteract the likely depression of force that would otherwise be the result of increasing [K+]e. We also discuss some mechanisms that prevent a metabolic catastrophe when energy demand exceeds the capability to replenish ATP. This regulation is highly redundant and appears to concentrate on diminishing Ca2+ release, because this simultaneously reduces the activity of both Ca2+ ATPase and myosin ATPase and serves to preserve ATP. This regulation occurs at the level of membrane excitability, where the activation of both Cl− and KATP channels reduces excitability, thereby reducing the Ca2+ release by RyR. There is also regulation at the level of the RyR, which responds to a decrease in [ATP] and increase in [Mg2+], which occur during repetitive or sustained contractions. In this case, both of these changes reduce the Po of RyR. RyR Po is also regulated by Ca2+, either directly or through CaM as sustained elevation of [Ca2+]i results in an inhibition of RyR by the CaCaM complex. Collectively, these regulatory processes allow exercise to continue, as long as additional motor units are available for recruitment, while preventing depletion of ATP in individual cells.
The authors would like to thank Barbara Holash for creating Fig. 1. Research in the laboratories of the authors is supported by the Natural Sciences and Engineering Research Council of Canada.