The Na,K-ATPase α2 isoform is the predominant Na,K-ATPase in adult skeletal muscle and the sole Na,K-ATPase in the transverse tubules (T-tubules). In quiescent muscles, the α2 isozyme operates substantially below its maximal transport capacity. Unlike the α1 isoform, the α2 isoform is not required for maintaining resting ion gradients or the resting membrane potential, canonical roles of the Na,K-ATPase in most other cells. However, α2 activity is stimulated immediately upon the start of contraction and, in working muscles, its contribution is crucial to maintaining excitation and resisting fatigue. Here, we show that α2 activity is determined in part by the K⁺ concentration in the T-tubules, through its K⁺ substrate affinity. Apparent K⁺ affinity was determined from measurements of the K_1/2 for K⁺ activation of pump current in intact, voltage-clamped mouse flexor digitorum brevis muscle fibers. Pump current generated by the α2 Na,K-ATPase, Ip, was identified as the outward current activated by K⁺ and inhibited by micromolar ouabain. Ip was outward at all potentials studied (−90 to −30 mV) and increased with depolarization in the subthreshold range, −90 to −50 mV. The Q₁₀ was 2.1 over the range of 22–37°C. The K_1/2,K of Ip was 4.3 ± 0.3 mM at −90 mV and was relatively voltage independent. This K⁺ affinity is lower than that reported for other cell types but closely matches the dynamic range of extracellular K⁺ concentrations in the T-tubules. During muscle contraction, T-tubule luminal K⁺ increases in proportion to the frequency and duration of action potential firing. This K_1/2,K predicts a low fractional occupancy of K⁺ substrate sites at the resting extracellular K⁺ concentration, with occupancy increasing in proportion to the frequency of membrane excitation. The stimulation of preexisting pumps by greater K⁺ site occupancy thus provides a rapid mechanism for increasing α2 activity in working muscles.

Initiation of skeletal muscle contraction is triggered by rapid activation of RYR1 channels in response to sarcolemmal depolarization. RYR1 is intracellular and has no voltage-sensing structures, but it is coupled with the voltage-sensing apparatus of Ca_V1.1 channels to inherit voltage sensitivity. Using an opto-electrophysiological approach, we resolved the excitation-driven molecular events controlling both Ca_V1.1 and RYR1 activations, reported as fluorescence changes. We discovered that each of the four human Ca_V1.1 voltage-sensing domains (VSDs) exhibits unique biophysical properties: VSD-I time-dependent properties were similar to ionic current activation kinetics, suggesting a critical role of this voltage sensor in Ca_V1.1 activation; VSD-II, VSD-III, and VSD-IV displayed faster activation, compatible with kinetics of sarcoplasmic reticulum Ca²⁺ release. The prominent role of VSD-I in governing Ca_V1.1 activation was also confirmed using a naturally occurring, charge-neutralizing mutation in VSD-I (R174W). This mutation abolished Ca_V1.1 current at physiological membrane potentials by impairing VSD-I activation without affecting the other VSDs. Using a structurally relevant allosteric model of Ca_V activation, which accounted for both time- and voltage-dependent properties of Ca_V1.1, to predict VSD-pore coupling energies, we found that VSD-I contributed the most energy (~75 meV or ∼3 kT) toward the stabilization of the open states of the channel, with smaller (VSD-IV) or negligible (VSDs II and III) energetic contribution from the other voltage sensors (<25 meV or ∼1 kT). This study settles the longstanding question of how Ca_V1.1, a slowly activating channel, can trigger RYR1 rapid activation, and reveals a new mechanism for voltage-dependent activation in ion channels, whereby pore opening of human Ca_V1.1 channels is primarily driven by the activation of one voltage sensor, a mechanism distinct from that of all other voltage-gated channels.

Mutations of Ca_V1.1, the pore-forming subunit of the L-type Ca²⁺ channel in skeletal muscle, are an established cause of hypokalemic periodic paralysis (HypoPP). However, functional assessment of HypoPP mutant channels has been hampered by difficulties in achieving sufficient plasma membrane expression in cells that are not of muscle origin. In this study, we show that coexpression of Stac3 dramatically increases the expression of human Ca_V1.1 (plus α₂-δ_1b and β_1a subunits) at the plasma membrane of Xenopus laevis oocytes. In voltage-clamp studies with the cut-open oocyte clamp, we observe ionic currents on the order of 1 μA and gating charge displacements of ∼0.5–1 nC. Importantly, this high expression level is sufficient to ascertain whether HypoPP mutant channels are leaky because of missense mutations at arginine residues in S4 segments of the voltage sensor domains. We show that R528H and R528G in S4 of domain II both support gating pore currents, but unlike other R/H HypoPP mutations, R528H does not conduct protons. Stac3-enhanced membrane expression of Ca_V1.1 in oocytes increases the throughput for functional studies of disease-associated mutations and is a new platform for investigating the voltage-dependent properties of Ca_V1.1 without the complexity of the transverse tubule network in skeletal muscle.

Repetitive activation of skeletal muscle fibers leads to a reduced transmembrane K⁺ gradient. The resulting membrane depolarization has been proposed to play a major role in the onset of muscle fatigue. Nevertheless, raising the extracellular K⁺ (\( {\text{K}}_{\text{o}}^{ + } \)) concentration (\( [ {\text{K}}^{ + } ]_{\text{o}} \)) to 10 mM potentiates twitch force of rested amphibian and mammalian fibers. We used a double Vaseline gap method to simultaneously record action potentials (AP) and Ca²⁺ transients from rested frog fibers activated by single and tetanic stimulation (10 pulses, 100 Hz) at various \( [ {\text{K}}^{ + } ]_{\text{o}} \) and membrane potentials. Depolarization resulting from current injection or raised \( [ {\text{K}}^{ + } ]_{\text{o}} \) produced an increase in the resting [Ca²⁺]. Ca²⁺ transients elicited by single stimulation were potentiated by depolarization from −80 to −60 mV but markedly depressed by further depolarization. Potentiation was inversely correlated with a reduction in the amplitude, overshoot and duration of APs. Similar effects were found for the Ca²⁺ transients elicited by the first pulse of 100 Hz trains. Depression or block of Ca²⁺ transient in response to the 2nd to 10th pulses of 100 Hz trains was observed at smaller depolarizations as compared to that seen when using single stimulation. Changes in Ca²⁺ transients along the trains were associated with impaired or abortive APs. Raising \( [ {\text{K}}^{ + } ]_{\text{o}} \) to 10 mM potentiated Ca²⁺ transients elicited by single and tetanic stimulation, while raising \( [ {\text{K}}^{ + } ]_{\text{o}} \) to 15 mM markedly depressed both responses. The effects of 10 mM \( {\text{K}}_{\text{o}}^{ + } \) on Ca²⁺ transients, but not those of 15 mM \( {\text{K}}_{\text{o}}^{ + } \), could be fully reversed by hyperpolarization. The results suggests that the force potentiating effects of 10 mM \( {\text{K}}_{\text{o}}^{ + } \) might be mediated by depolarization dependent changes in resting [Ca²⁺] and Ca²⁺ release, and that additional mechanisms might be involved in the effects of 15 mM \( {\text{K}}_{\text{o}}^{ + } \) on force generation.

Mutations in CAPN3 cause limb girdle muscular dystrophy type 2A (LGMD2A), a progressive muscle wasting disease. CAPN3 is a non-lysosomal, Ca-dependent, muscle-specific proteinase. Ablation of CAPN3 (calpain-3 knockout (C3KO) mice) leads to reduced ryanodine receptor (RyR1) expression and abnormal Ca2+/calmodulin-dependent protein kinase II (Ca-CaMKII)-mediated signaling. We previously reported that Ca2+ release measured by fura2-FF imaging in response to single action potential stimulation was reduced in old C3KO mice; however, the use of field stimulation prevented investigation of the mechanisms underlying this impairment. Furthermore, our prior studies were conducted on older animals, whose muscles showed advanced muscular dystrophy, which prevented us from establishing whether impaired Ca2+ handling is an early feature of disease. In the current study, we sought to overcome these matters by studying single fibers isolated from young wild-type (WT) and C3KO mice using a low affinity calcium dye and high intracellular ethylene glycol-bis(2-aminoethylether)-n,n,n′,n′-tetraacetic acid (EGTA) to measure Ca2+ fluxes. Muscles were subjected to both current and voltage clamp conditions. Standard and confocal fluorescence microscopy was used to study Ca2+ release in single fibers enzymatically isolated from hind limb muscles of wild-type and C3KO mice. Two microelectrode amplifier and experiments were performed under current or voltage clamp conditions. Calcium concentration changes were detected with an impermeant low affinity dye in the presence of high EGTA intracellular concentrations, and fluxes were calculated with a single compartment model. Standard Western blotting analysis was used to measure the concentration of RyR1 and the α subunit of the dihydropyridine (αDHPR) receptors. Data are presented as mean ± SEM and compared with the Student’s test with significance set at p < 0.05. We found that the peak value of Ca2+ fluxes elicited by single action potentials was significantly reduced by 15–20 % in C3KO fibers, but the kinetics was unaltered. Ca2+ release elicited by tetanic stimulation was also impaired in C3KO fibers. Confocal studies confirmed that Ca2+ release was similarly reduced in all triads of C3KO mice. Voltage clamp experiments revealed a normal voltage dependence of Ca2+ release in C3KO mice but reduced peak Ca2+ fluxes as with action potential stimulation. These findings concur with biochemical observations of reduced RyR1 and αDHPR levels in C3KO muscles and reduced mechanical output. Confocal studies revealed a similar decrease in Ca2+ release at all triads consistent with a homogenous reduction of functional voltage activated Ca2+ release sites. Overall, these results suggest that decreased Ca2+ release is an early defect in calpainopathy and may contribute to the observed reduction of CaMKII activation in C3KO mice.

Ion movements across biological membranes, driven by electrochemical gradients or active transport mechanisms, control essential cell functions. Membrane ion movements can manifest as electrogenic currents or electroneutral fluxes, and either process can alter the extracellular and/or intracellular concentration of the transported ions. Classic electrophysiological methods allow accurate measurement of membrane ion movements when the transport mechanism produces a net ionic current; however, they cannot directly measure electroneutral fluxes and do not detect any accompanying change in intracellular ion concentrations. Here, we developed a method for simultaneously measuring ion movements and the accompanying dynamic changes in intracellular ion concentrations in intact skeletal muscle fibers under voltage or current clamp in real time. The method combines a two-microelectrode voltage clamp with ion-selective and reference microelectrodes (four-electrode system). We validate the electrical stability of the system and the viability of the preparation for periods of ∼1 h. We demonstrate the power of this method with measurements of intracellular Cl⁻, H⁺, and Na⁺ to show (a) voltage-dependent redistribution of Cl⁻ ions; (b) intracellular pH changes induced by changes in extracellular pCO₂; and (c) electroneutral and electrogenic Na⁺ movements controlled by the Na,K-ATPase. The method is useful for studying a range of transport mechanisms in many cell types, particularly when the transmembrane ion movements are electrically silent and/or when the transport activity measurably changes the intracellular activity of a transported ion.