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The Ca2+ influx through the mammalian skeletal muscle dihydropyridine receptor is irrelevant for muscle performance

Abstract

Skeletal muscle excitation–contraction (EC) coupling is initiated by sarcolemmal depolarization, which is translated into a conformational change of the dihydropyridine receptor (DHPR), which in turn activates sarcoplasmic reticulum (SR) Ca2+ release to trigger muscle contraction. During EC coupling, the mammalian DHPR embraces functional duality, as voltage sensor and l-type Ca2+ channel. Although its unique role as voltage sensor for conformational EC coupling is firmly established, the conventional function as Ca2+ channel is still enigmatic. Here we show that Ca2+ influx via DHPR is not necessary for muscle performance by generating a knock-in mouse where DHPR-mediated Ca2+ influx is eliminated. Homozygous knock-in mice display SR Ca2+ release, locomotor activity, motor coordination, muscle strength and susceptibility to fatigue comparable to wild-type controls, without any compensatory regulation of multiple key proteins of the EC coupling machinery and Ca2+ homeostasis. These findings support the hypothesis that the DHPR-mediated Ca2+ influx in mammalian skeletal muscle is an evolutionary remnant.

Introduction

Excitation–contraction (EC) coupling in skeletal muscle can be understood only in an evolutionary context. The phylogenetic earliest mechanism involves muscle membrane depolarization, which triggers pore openings of the voltage-gated sarcolemmal l-type Ca2+ channel, or dihydropyridine receptor (DHPR), leading to a substantial and fast activating Ca2+ inward current1,2,3. This sudden increase in intracellular Ca2+ concentration triggers Ca2+-sensitive Ca2+ release channels or ryanodine receptors (RyR) to open and release massive amounts of Ca2+ ions from sarcoplasmic reticulum (SR) stores into the cytoplasm, sufficient to finally induce muscle contraction4,5,6. This phylogenetically ancient EC-coupling mechanism, which is fully dependent on Ca2+-induced Ca2+ release (CICR), is still extant in skeletal muscles of invertebrates and protochordates7,8,9,10,11,12, and interestingly, in all cardiac and smooth muscles up to the evolutionary most derived vertebrates3, 6, 13.

However, during evolution to vertebrates, novel skeletal-muscle-specific DHPR and RyR isoforms emerged, giving rise to a new and unique mechanism of EC coupling: Vertebrate skeletal muscle EC coupling is based on a Ca2+ influx-independent, inter-channel protein–protein interaction between the DHPR and skeletal-muscle-specific RyR type-1 (RyR1)14. Upon this physical interaction, the voltage-induced conformational change of the DHPR is transmitted to RyR1, which subsequently opens, releases large amounts of Ca2+ from the SR stores and ultimately leads to muscle contraction15, 16. This phylogenetically most derived skeletal-muscle-specific mechanism of depolarization-induced Ca2+ release combines the advantage of accelerated EC coupling efficacy along with lower energy consumption due to reduced Ca2+ extrusion across the sarcolemma17. A prerequisite for this Ca2+ -independent physical DHPR–RyR1 interaction was a transition in the ultrastructural positioning of the DHPRs from random clustering around the RyR (the protochordate/cardiac-type) to an arrangement into tetrads, i.e., four DHPRs positioned strictly adjacent to the homotetrameric RyR11, 18.

Interestingly, in addition to their role in conformational EC coupling, mammalian skeletal muscle DHPRs retained their ancient function as Ca2+ channels19. However, the physiological significance of this DHPR Ca2+ current, which is certainly not (immediately) required for EC coupling14, is still unresolved despite being under investigation since nearly half a century19. So far, one can only speculate that this DHPR Ca2+ inward current, might contribute in the form of excitation-coupled Ca2+ entry (ECCE)19,20,21,22 and have a crucial role in Ca2+ homeostasis, e.g., by SR store filling, or is essential for skeletal muscle development via regulation of acetylcholine receptor pre-patterning and formation of neuromuscular junctions23, 24, or has a negative inotropic effect25,26,27,28,29, which is crucial for overall muscle health and fibre integrity29, or finally, is just an evolutionary remnant of the ancient CICR stage2, 3, 30. In other words, it is unclear whether the skeletal muscle DHPR Ca2+ current is exaptational or vestigial. However, to date it was not possible to answer any of these questions due to the unavailability of an appropriate animal model system with a fully occluded skeletal muscle DHPR.

Recently, a mouse model carrying a mutation in the DHPR selectivity filter (E1014K) was generated31. This mutation leads to reduced Ca2+ pore-binding affinity and thus changes the Ca2+ selectivity of the DHPR by allowing flow of monovalent cations instead of Ca2+ 32, 33. The E1014K mouse displays reduced muscle endurance, decreased muscle fibre size and a shift in fibre-type distribution towards fast fibres31. The authors concluded that Ca2+ permeation and/or binding via/to the DHPR pore plays an important modulatory role in muscle performance31. However, a drawback of this interpretation is the negligence of the presence of a persisting permeation of monovalent cations through the mutant EK channel31, 32, 34, which might, for instance, affect the membrane potential during muscle activity32.

With the intention to create a mouse model with a fully occluded skeletal muscle DHPR, we used a DHPR pore-blocking mechanism refined by evolution. Previously, we found that zebrafish and all other euteleost fishes express non-Ca2+-conducting DHPRs in their skeletal muscles2, 35. Consequently, we engineered the selectivity-filter-adjacent point mutation N617D, previously identified to be responsible for DHPR non-conductivity in zebrafish fast, glycolytic muscle2 into the mouse genome. Electrophysiological investigations on cultured skeletal muscle primary myotubes and adult fibres from the resultant homozygous mutant mouse with non-conducting (nc) DHPR showed complete block of DHPR Ca2+ influx but unaltered intracellular SR Ca2+ release. Notably, and in contrast to the E1014K model, the ncDHPR pore does not show permeation of monovalent cations36.

Here we use the ncDHPR mouse model to explore the specific role of the DHPR-mediated Ca2+ current in skeletal muscle via a range of muscle performance tests on whole animals and isolated muscles, as well as at the cellular and molecular level with biophysical, immunohistochemical, biochemical and quantitative PCR (qPCR) techniques. Surprisingly, these experiments did not point to any differences between ncDHPR mice and their wild-type (wt) littermates, irrespective of whether mice were young or aged, or the isolated muscles were fast- or slow-twitch type. Overall, our results strongly support the hypothesis that the Ca2+ influx through the mammalian skeletal muscle DHPR is irrelevant for muscle performance and consequently can be considered as an evolutionary remnant (vestigial) of the phylogenetic stages of early chordates, where DHPR Ca2+ influx was the exclusive trigger for RyR activation in skeletal-muscle EC coupling.

Results

Creation of the non-Ca2+ -conducting (nc)DHPR knock-in mouse

To thoroughly investigate the physiological relevance of the DHPR Ca2+ influx in skeletal muscle, we generated a non-Ca2+-conducting (nc) DHPR knock-in mouse. Mutation N617D in pore loop II, previously identified to be responsible for the complete loss of Ca2+ conductance in zebrafish DHPR α1S-b[2] was introduced into the murine CACNA1S gene (Supplementary Fig. 1a). Strategy and targeting construct used for generating the N617D mutant mouse are shown in Supplementary Fig. 1b. The genotype was confirmed by restriction fragment length polymorphism (RFLP) and sequencing.

No Ca2+ influx but unaltered EC coupling in the ncDHPR mouse

As a first step, to test whether introduction of the N617D mutation eliminates inward Ca2+ currents via the mouse skeletal muscle DHPR, we measured whole-cell Ca2+ currents from ncDHPR myotubes isolated from 3 to 4 days old pups and interosseous muscle fibres isolated from young mice. Recordings from ncDHPR myotubes display complete loss of DHPR Ca2+ influx (with I max of −0.02 ± 0.01 pA pF−1) compared (P < 0.001, unpaired t-test) with wt myotubes (I max = −4.87 ± 0.31 pA pF−1) (Fig. 1a–c). However, voltage dependence and size of intracellular Ca2+ release was indistinguishable (P > 0.05, unpaired t-test) between ncDHPR and wt myotubes (Fig. 1d–f), pointing to intact EC coupling. Next, we examined whether elevating the recording temperature to the physiological value might evoke inward Ca2+ currents in ncDHPR myotubes. Interestingly, at ~37 °C, we found a twofold augmentation of Ca2+ currents in wt myotubes (I max = −9.55 ± 0.70 pA pF−1), but again no inward Ca2+ current in ncDHPR myotubes (I max = −0.07 ± 0.04 pA pF−1) (Supplementary Fig. 2a). However, contrary to the enhancement of DHPR Ca2+ influx in wt, there was no effect (P > 0.05, unpaired t-test) of elevated temperature on intracellular Ca2+ release, neither in wt nor ncDHPR myotubes (Supplementary Fig. 2b). Consistent with the outcome from primary myotubes, adult interosseous muscle fibres isolated from young ncDHPR mice also displayed complete loss of DHPR Ca2+ influx (I max = 0.06 ± 2.02 nA) compared (P < 0.001, unpaired t-test) with wt fibres (I max = −19.1 ± 2.62 nA) (Supplementary Fig. 3). The SR Ca2+ release showed the typical phasic-tonic response (Fig. 2a) with indistinguishable (P > 0.05, unpaired t-test) voltage-dependence of peak Ca2+ release flux (Fig. 2b) and Ca2+ permeability (Fig. 2c) between ncDHPR and wt fibres. As well, kinetic parameters characterizing the shape of the signals viz. time to reach peak activation and the ratio of permeability peak to plateau were comparable between both genotypes at all test potentials (Supplementary Fig. 4). Altogether, these results indicate a complete elimination of skeletal muscle Ca2+ influx but unaltered voltage-induced SR Ca2+ release in the ncDHPR mouse model. Thus, the ncDHPR model serves as a unique system for investigating the physiological relevance of the DHPR Ca2+ influx in skeletal muscle EC coupling.

Fig. 1
figure1

N617D mutation completely abolishes DHPR Ca2+ influx without altering EC coupling. a, b Representative whole cell Ca2+ current recordings on myotubes isolated from 3 to 4 days old (a) wt and (b) ncDHPR pups, indicating the complete loss of inward Ca2+ current in ncDHPR mice in contrast (P < 0.001) to wt control mice. Scale bars, 50 ms (horizontal), 2 pA pF−1 (vertical). c Current–voltage relationships for DHPR-mediated Ca2+ currents recorded from ncDHPR (n = 13) and wt (n = 7) myotubes. d, e Representative intracellular SR Ca2+ release recordings from (d) wt and (e) ncDHPR myotubes. Scale bars, 50 ms (horizontal), ∆F/F 0 = 2 (vertical). f In contrast to DHPR Ca2+ influx, voltage dependence of maximal Ca2+ release is indistinguishable (P > 0.05) between ncDHPR ((ΔF/F 0)max = 3.75 ± 0.39; n = 12) and wt ((∆F/F 0)max = 3.39 ± 0.26; n = 8) myotubes, implying unaltered EC coupling in the ncDHPR mouse. Data are represented as mean ± s.e.m.; P determined by unpaired Student’s t-test

Fig. 2
figure2

Unaltered voltage dependence of SR Ca2+ release in ncDHPR adult muscle fibres. a Representative traces of Ca2+ release flux signals determined from isolated adult toe fibres (musculus interosseus) at 100-ms depolarizing voltage steps. The flux exhibits an early peak and a rapid decline to a lower level (plateau). Scale bars, 200 ms (horizontal), 50 mM s−1 (vertical). b Voltage dependence of normalized peak Ca2+ release flux is indistinguishable (P > 0.05) between ncDHPR (n = 12) and wt (n = 10) adult muscle fibres. c Ca2+ release flux traces were converted to permeability with the assumption that the slow decline during the plateau phase results from SR depletion60. The calculated peak Ca2+ release permeability as a function of voltage showed no significant difference (P > 0.05) between ncDHPR and wt fibres. Data are represented as mean ± s.e.m.; P determined by unpaired Student’s t-test

SR Ca2+ store content is unaltered in ncDHPR mice

To assess the importance of DHPR Ca2+ influx in SR store filling, Fura-FF-AM loaded adult interosseous muscle fibres were challenged with RyR agonist 4-CmC (4-chloro-m-cresol) under conditions that inhibited Ca2+ uptake by sarco/endoplasmic reticulum Ca2+-ATPase(SERCA) (viz. the addition of CPA, cyclopiazonic acid) (Fig. 3a). SERCA block could be verified by a slow increase in resting Ca2+ concentration when CPA was applied alone. Figure 3b depicts the comparison of the peak Fura-FF fluorescence ratios and demonstrates that 4-CmC-induced Ca2+ release is not significantly different (P > 0.05, unpaired t-test) between ncDHPR and wt fibres, indicating identical steady state SR Ca2+ loading in both genotypes.

Fig. 3
figure3

Absence of DHPR-mediated Ca2+ influx has no impact on SR Ca2+ store content. a Representative trace of SR Ca2+ release in intact interosseous muscle fibres triggered by application (green arrow) of 500 µM of RyR agonist 4-CmC in the presence of 30 µM of SERCA blocker CPA. The resulting Ca2+ signals were measured with the low affinity indicator Fura-FF-AM to avoid dye saturation during Ca2+ release stimulation. b Comparison of 4-CmC-induced Ca2+ signals, measured as peak fluorescence ratio of Fura-FF (340/380) showed no significant difference (P > 0.05) between ncDHPR (1.0 ± 0.02; n = 53) and wt muscle fibres (1.04 ± 0.02; n = 48), indicating identical SR Ca2+ store filling. Bars represent mean ± s.e.m.; P determined by unpaired Student’s t-test

Basic phenotypic characterization of the ncDHPR mouse

The absence of DHPR Ca2+ influx in ncDHPR mice did not cause embryonic lethality and animals had normal posture and gait, and could not be visually distinguished from wt littermates. No difference (P > 0.05, unpaired t-test) in male (Fig. 4a) and female (Fig. 4b) body weight development was observed between ncDHPR and wt mice. In addition, litter size (Fig. 4b, inset) was indistinguishable (P > 0.05, unpaired t-test) between both genotypes.

Fig. 4
figure4

Lack of DHPR Ca2+ influx has no impact on body weight development and fertility. Growth curve of (a) male and (b) female ncDHPR and wt mice was monitored weekly for 150 days. a Male ncDHPR (n = 30) and wt (n = 35) siblings showed identical (P > 0.05) body weight development. b Similarly, female ncDHPR (n = 29) showed no difference (P > 0.05) in body weight development compared with their wt control siblings (n = 31). At the end of the observation period, the maximum body weight of male ncDHPR and wt mice was 29.76 ± 0.55 and 30.74 ± 0.54 g, respectively, and of female ncDHPR and wt mice was 28.96 ± 0.73 and 28.98 ± 0.60 g, respectively. b (Inset) The mean litter size of ncDHPR mice (7.64 ± 0.21; n = 100) was similar (P > 0.05) to wt mice (7.31 ± 0.37; n = 49). Data are represented as mean ± s.e.m.; P determined by unpaired Student’s t-test

To identify possible muscle-type-specific differences we chose extensor digitorum longus (EDL) muscle, which primarily consists of fast-twitch fibres and soleus (SOL) muscle, which is more a slow-twitch muscle compared with EDL. Basic physical parameters such as wet weight, length and diameter of SOL and EDL muscles, isolated from young (Supplementary Fig. 5a, b) or aged mice (Supplementary Fig. 5c, d) were not different (P > 0.05, unpaired t-test) between ncDHPR and wt mice.

Furthermore, transverse cryosections of SOL and EDL muscle immunostained with antibodies against specific myosin heavy-chain isoforms indicated that the lack of DHPR Ca2+ influx in the ncDHPR mouse did not induce any change (P > 0.05, unpaired t-test) in the fibre-type composition, neither in the slower-twitch SOL muscle nor the fast-twitch EDL (Supplementary Fig. 6a, b). Similarly, no significant differences (P > 0.05, unpaired t-test) in the fibre cross-sectional area (CSA) of SOL and EDL muscles were observed between both genotypes (Supplementary Fig. 6c).

ncDHPR mice show unaltered voluntary locomotor activity

As the ncDHPR mouse did not show any evident basic phenotypic differences from wt, we next tested for consequences of DHPR Ca2+ non-conductivity on voluntary locomotor activity37 in young (3–6 months old) mice. Home-cage ambulatory activity recorded over 60 h was indistinguishable (P > 0.05, unpaired t-test) between ncDHPR and wt mice (average age: 4.0 months). Both the genotypes displayed normal behaviour, with higher activity during the dark cycle and lower activity during the light cycle (Fig. 5a). Furthermore, to record enhanced voluntary locomotor activity37, the mice (average age: 4.1 months) were challenged with a voluntary activity wheel experiment for 21 days. Consistent with above results, the cumulative average distance covered by ncDHPR mice (19.29 ± 6.31 km) was similar (P > 0.05, unpaired t-test) to that of wt control mice (21.28 ± 3.71 km) (Fig. 5b).

Fig. 5
figure5

Voluntary locomotor activity is identical in ncDHPR and wt mice. a Home cage ambulatory activity was measured for 2 days and 3 nights with an infrared video monitoring system. Young (3–6 months old) ncDHPR mice (n = 10) displayed high nighttime activity (41,520 ± 3,934 a.u.) and low daytime activity (5,794 ± 790 a.u) similar (P > 0.05) to the control wt mice (37,620 ± 1,079 a.u. and 6,247 ± 1,152 a.u., respectively; n = 6). b Average cumulative distance covered during 21 days of voluntary activity wheel task (binned every 24 h) was indistinguishable (P > 0.05) between young ncDHPR (n = 18) and wt (n = 17) mice. Data are represented as mean ± s.e.m.; P determined by unpaired Student’s t-test

Ex vivo contractile properties are unaltered in ncDHPR mice

To test the impact of the loss of DHPR Ca2+ influx on skeletal muscle functioning in intact muscles independent of other parameters and under high frequency stimulations38, we compared ex vivo contractile properties, viz. force and fatigability of SOL and EDL muscles isolated from ncDHPR and wt mice. For this part of the study, young (3–7 months old; average age: 4.4 months) and aged (17–22 months old; average age: 19.3 months) mice were used to examine putative age-related accumulative degenerative effects on muscle performance due to the loss of DHPR Ca2+ influx. On being subjected to force-frequency tests, SOL and EDL muscles did not show any differences (P > 0.05, unpaired t-test) in maximum twitch force between young ncDHPR and wt mice (Fig. 6a, left graph). Furthermore, the maximum twitch force observed in both muscle types of aged mice was indistinguishable (P > 0.05, unpaired t-test) between ncDHPR and wt (Fig. 6a, right graph). The tetanic force exhibited by SOL and EDL at increasing stimulation frequencies (for exemplar recordings see Supplementary Fig. 7a, b) were similar (P > 0.05, unpaired t-test) in young (Fig. 6b, left graph) or aged (Fig. 6b, right graph) ncDHPR and wt mice.

Fig. 6
figure6

Absence of DHPR Ca2+ influx has no impact on ex vivo muscle force and fatigue. Soleus (SOL) and extensor digitorum longus (EDL) muscles isolated from young and aged ncDHPR and wt control mice were subjected to (a, b) force-frequency and c repetitive tetanic fatigue tests (see also Supplementary Fig. 7). a The average maximum twitch force (at supramaximal voltage, 25 V for 1 ms) is identical (P > 0.05) in SOL and EDL muscles between young (left graph) ncDHPR (SOL: 16.78 ± 1.80 mN; n = 12; EDL: 55.33 ± 3.31 mN; n = 19) and wt (SOL: 17.39 ± 1.49 mN: n = 9; EDL: 55.14 ± 3.39 mN: n = 18) mice, as well as between aged (right graph) ncDHPR (SOL: 11.66 ± 0.91 mN; n = 19 and EDL: 9.76 ± 1.04 mN; n = 20) and wt (SOL: 13.33 ± 1.35 mN; n = 18 and EDL: 10.09 ± 1.06 mN; n = 22) mice. b Frequency dependence of average tetanic force in SOL and EDL muscles is also unaltered (P > 0.05) between age-matched ncDHPR and wt mice (left and right graphs). c Under high-frequency repetitive stimulations, the number of tetani till the tetanus amplitude decreased to 80% (T80%) of the initial force in case of SOL and 50% (T50%) for EDL was taken as an index of fatigue. No difference in susceptibility to fatigue was observed in SOL and EDL muscles isolated from either young (left graph) ncDHPR (SOL: T80% = 50.12 ± 14.65 n; n = 12; EDL: T50% = 25.95 ± 1.04 n; n = 19) and wt (SOL: T80% = 48.0 ± 16.66 n; n = 9; EDL: T50% = 25.33 ± 1.05 n; n = 18) mice or aged (right graph) ncDHPR (SOL: T80% = 27.22 ± 4.89 n; n = 18; EDL: T50% = 18.12 ± 1.14 n; n = 17) and wt (SOL: T80% = 25.5 ± 3.31 n; n = 18; EDL: T50% = 21.42 ± 2.61 n; n = 19) mice. All recordings were performed at room temperature (~26 °C). Data are represented as mean ± s.e.m.; P determined by unpaired Student’s t-test

As the EC coupling mechanism is affected in muscle fatigue38, the implications of the loss of DHPR Ca2+ influx in the ncDHPR mouse might become evident only under extreme fatiguing conditions. Hence, we subjected isolated SOL and EDL muscles to fatigue-inducing repetitive high-frequency tetanic stimulations (Supplementary Fig. 7c, d). SOL is markedly more fatigue resistant than EDL due to its higher oxidative capacity38. Consistent with the above findings, susceptibility to fatigue was not different (P > 0.05, unpaired t-test) between muscles isolated from young ncDHPR and wt mice (Fig. 6c, left graph). Likewise, there was no significant difference in fatigability between muscle derived from aged ncDHPR and wt mice (Fig. 6c, right graph). Akin to the physiological muscle parameters, kinetic parameters of single twitches such as time to peak, time to half-relaxation and twitch duration were similar (P > 0.05, unpaired t-test) in muscles from ncDHPR and wt, regardless of whether isolated from young or aged mice (Supplementary Fig. 8).

ncDHPR mice display normal muscle coordination and strength

DHPR-mediated Ca2+ influx has no relevance in muscle force production, as can be concluded from the above ex vivo isometric contraction experiments on isolated SOL and EDL muscles from ncDHPR and wt mice. However, to assess whether this also holds true at the whole animal level, we confronted the animals with a battery of tests to measure muscle coordination and strength37. In the beam walk test, the average time to traverse the 120-cm-long beam was similar (P > 0.05, unpaired t-test) in young ncDHPR (8.40 ± 0.31 s; n = 10) and wt (8.83 ± 0.60 s; n = 6) mice. Next, we compared muscle strength of all four limbs37 by letting the mice hang upside down on a cross-wired grid for a maximum of 600 s. The latency to fall from the inverted grid was indistinguishable (P > 0.05, unpaired t-test) between young ncDHPR and wt control mice (Fig. 7a, left two bars). Mice that achieved the maximum hanging time were also similar for both genotypes (14 ncDHPR and 12 wt mice). Likewise, no significant difference (P > 0.05, unpaired t-test) in performance was observed between aged ncDHPR and wt mice (Fig. 7a, right two bars). None of the aged mice reached the maximum hanging time of 600 s. Finally, we performed a highly specific grip strength test to exclude other aspects of behaviour and coordination37, and exclusively measure the maximal muscle strength of forelimbs. In this test, the mouse was allowed to grasp onto a trapezoidal steel grip and after a gentle pull, the maximal force (in g) exerted before it releases the grip was recorded. Consistent with results obtained with other tests, young and aged ncDHPR mice displayed similar (P > 0.05, unpaired t-test) muscle strength to their age-matched control wt siblings (Fig. 7b). Average body weight was homogenous across all the cohorts.

Fig. 7
figure7

DHPR Ca2+ influx has no impact on whole animal muscle coordination and strength. a ncDHPR mice performed equally well similar to control littermates in wire hang test set for 600 s. The mean latency to fall from the cross-wired grid was similar between young ncDHR (453.85 ± 39.97 s; n = 26) and wt (453.04 ± 38.06 s; n = 25) mice, as well as between aged ncDHPR (153.12 ± 24.82 s; n = 25) and wt (100.44 ± 15.06 s; n = 27) mice. b The mean grip strength, an index of maximal muscle force of forelimbs, was not different in young (185.68 ± 4.71 g; n = 15) and aged (136.39 ± 3.59 g; n = 23) ncDHPR mice compared with their age-matched wt littermates (young wt: 181.38 ± 3.88 g; n = 6; aged wt: 142.29 ± 4.34 g; n = 27 mice). Normal age-dependent deterioration in muscle strength is evident in both genotypes. Bars represent mean ± s.e.m.; P determined by unpaired Student’s t-test

Similar whole animal muscle endurance/fatigue in ncDHPR mice

Ex vivo measurements on isolated SOL and EDL muscles showed no alterations in fatigability in the absence of DHPR-mediated Ca2+ influx in ncDHPR mice. To confirm these findings and fully characterize the skeletal muscle functioning in its physiological environment in the ncDHPR mice, we performed involuntary fatigue and endurance tests at the whole animal level. As a first measure of sensorimotor skills and fatigue, animals were confronted to a standard Rotarod test with steady acceleration37 from 4 to 40 rpm over 300 s. Young and aged ncDHPR mice remained on the accelerating rod for the same (P > 0.05, unpaired t-test) time such as the age-matched wt control mice (Fig. 8a). Likewise, the maximum acceleration at which the mice fell from the rotating rod was indistinguishable (P > 0.05, unpaired t-test) between young ncDHPR and wt mice (26.74 ± 1.67 rpm and 25.03 ± 1.79 rpm, respectively). Consistently, aged ncDHPR and wt mice displayed equal performance (20.07 ± 0.81 rpm and 20.01 ± 1.22 rpm, respectively). Next, we designed a prolonged Rotarod protocol (600 s) at constant speed (20 rpm for young and 15 rpm for aged mice) to access endurance. Latency to fall displayed by young ncDHPR mice was comparable (P > 0.05, unpaired t-test) to their wt counter mates (Fig. 8b, right two bars). Likewise, endurance level was indistinguishable (P > 0.05, unpaired t-test) between aged mice of both genotypes (Fig. 8b, left two bars).

Fig. 8
figure8

DHPR Ca2+ influx has no impact on whole animal muscle endurance and fatigue. a In Rotarod performance test, the mean latency to fall from a rotating rod under steady acceleration (4–40 rpm for 300 s) was indistinguishable (P > 0.05) when young (188.78 ± 13.95 s; n = 16) and aged ncDHPR (133.36 ± 6.73 s; n = 26) mice were compared with their wt (174.50 ± 15.02 s; n = 16 and 132.85 ± 10.19 s; n = 28, respectively) counter mates. b The mean latency to fall during 600-s Rotarod endurance task at constant speed, 20 rpm or 15 rpm in young or aged mice, respectively, was indifferent (P > 0.05) between age-matched ncDHPR (young: 351.54 ± 58.64 s; n = 16 and aged: 312.22 ± 50.76 s; n = 25) and wt (young: 263.43 ± 64.66 s; n = 16 and aged: 220.67 ± 47.32 s; n = 27) mice. The number of mice that remained on the rotating rod for the entire duration of the task was similar for both genotypes (5 mice each for the young group and 10 aged ncDHPR versus 6 aged wt mice). c Mean cumulative resting time during an intense treadmill run (0.9 km in 1 h) in young ncDHPR (30.45 ± 8.75 s; n = 17) was similar (P > 0.05) to wt control (50.15 ± 12.36 s; n = 17) mice, indicating unaltered susceptibility to muscle fatigue and endurance. The cumulative number of rests at the end of the task was also indistinguishable (P > 0.05) between both genotypes (139.94 ± 37.97 for ncDHPR and 232.18 ± 52.89 for wt mice). d Mean cumulative resting time during 1-h treadmill run (0.55 km) in aged ncDHPR mice (3.54 ± 0.34 s; n = 23) was analogous to the wt (5.04 ± 0.85 s; n = 27) siblings. Cumulative number of rests at the end of the task was again comparable (P > 0.05) between aged ncDHPR and wt mice (19.65 ± 2.27 s and 24.52 ± 3.82 s, respectively). Data are represented as mean ± s.e.m.; P determined by unpaired Student’s t-test

Resistance to fatigue is an optimal, comprehensive measure of muscle performance and one of the most powerful tools for analysing fatigue resistance is a treadmill task. An acute exhaustive treadmill protocol was used to measure the enforced maximum endurance limit of ncDHPR mice39. Interestingly, young mice of both genotypes accomplished the entire 0.9 km run with similar efficiency. The total average resting time cumulated during the 1-h task, as well as the cumulative number of rests at the end of the task were indistinguishable (P > 0.05, unpaired t-test) between both genotypes (Fig. 8c). Finally, due to age-related depreciation in muscle strength (see Fig. 6), aged mice were challenged with a milder protocol (see Methods). Consistent with the above findings, aged ncDHPR mice completed the entire 0.55 km treadmill run in 1 h with similar (P > 0.05, unpaired t-test) competence to their age-matched wt siblings (Fig. 8d). The comparable total average cumulative resting time reached by ncDHPR and wt mice indicates no age-related accumulative adverse effects on skeletal muscle fatigability despite the life-long absence of DHPR Ca2+ influx in ncDHPR mice. Akin to young mice, there was again no difference (P > 0.05, unpaired t-test) in the cumulative number of rests between aged ncDHPR and wt mice. Altogether, results from all the above experiments indicate that ncDHPR mice exhibit equivalent muscle performance to their wt siblings, which convincingly points to the insignificance of DHPR Ca2+ influx for skeletal muscle function.

No compensatory regulation of triad proteins in ncDHPR mice

The absence of DHPR Ca2+ influx has no obvious impact on general physiological parameters and whole animal muscle performance, neither in young nor aged ncDHPR mice. Apparently, the question arose whether up- or downregulation of any of the key triadic proteins responsible for either influx or efflux of Ca2+ ions compensates for the lack of DHPR-mediated Ca2+ influx in the ncDHPR mice. To address this issue, we first opted for qPCR analysis to quantify the messenger RNA levels transcribed from a selected panel of genes encoding for proteins involved in EC coupling and Ca2+ homeostasis, with β-actin (ACTB) and eukaryotic translation elongation factor 1 alpha 2 (EEF1A2) as reference genes. Total RNA was isolated from skeletal muscle of mice at different developmental stages viz. neonatal mice and adult SOL and EDL muscles. Starting with the main players of EC coupling, DHPR and RyR1, we found no differences (P > 0.05, unpaired t-test) in transcript levels between neonatal ncDHPR and wt mice (Fig. 9a). This lack of compensatory transcriptional DHPR upregulation is congruent with the density of DHPR Ca2+ currents recorded from heterozygous wt/ncDHPR myotubes, which did not reach more than half the current size compared (P < 0.001, unpaired t-test) with that recorded from wt myotubes (I max = −2.39 ± 0.15 and −4.87 ± 0.31 pA pF−1, respectively; Supplementary Fig. 9). Furthermore, expression profiles of mRNAs that code for critical Ca2+-handling proteins viz. TRPC1, Orai (predominant isoforms Orai1, Orai2 and Orai3), STIM1, SERCA1, PMCA1, NCX (isoforms NCX1 and NCX3) and CSQ (isoforms CSQ1 and CSQ2) were unaltered in skeletal muscles of neonatal ncDHPR mice compared (P > 0.05, unpaired t-test) with wt mice (Fig. 9). All PCRs were performed in triplicates on 18 first-strand replicates from 9 pups, with EEF1A2 as reference gene. Comparable results were obtained with ACTB as the reference gene (Supplementary Table 1). Likewise, there was no significant difference (P > 0.05, unpaired t-test) at the transcriptional level of all the above mentioned proteins in adult SOL or EDL muscles between both genotypes with EEF1A2 as reference gene (Supplementary Fig. 10).

Fig. 9
figure9

ncDHPR mice do not show compensatory regulation of key triadic EC coupling proteins. a Schematic representation of the skeletal muscle triad with T-tubular invagination (T-tubule) of the sarcolemma adjacent to the sarcoplasmic reticulum Ca2+ store (SR). TaqMan qPCR assay (comparative C T method) on skeletal muscle from neonatal mice revealed that none of the investigated key EC coupling and Ca2+ handling proteins is transcriptionally up- or down-regulated (P > 0.05) in ncDHPR mice (n = 18) compared to wt controls (n = 18). All PCRs were performed in triplicates on 18 first strand replicates from 9 pups. For each gene of interest, the expression level was normalized to the expression of the reference gene EEF1A2 (Eukaryotic translation elongation factor 1 alpha 2)66. Comparable results were obtained from SOL and EDL muscles of adult mice with EEF1A2 as the reference gene (Supplementary Fig. 10). Abbreviations: CSQ, calsequestrin; DHPRα1S, dihydropyridine receptor α1S subunit; NCX, Na+–Ca2+ exchanger; Orai, Ca2+ release-activated Ca2+ channel; PMCA, plasma membrane Ca2+-ATPase; RyR, ryanodine receptor; SERCA, sarco/endoplasmic reticulum Ca2+-ATPase; STIM, stromal interaction molecule; TRPC, transient receptor potential cation channel. b Quantitative western blot analysis revealed no compensatory translational regulation (P > 0.05) of crucial proteins involved in EC coupling and Ca2+ handling in the ncDHPR mouse (n = 3) compared with wt controls (n = 3). Band densities, quantified by densitometry, were standardized to GAPDH and normalized to the wt control sample on each gel. The corresponding western blots are shown in Supplementary Fig. 11 and in toto in Supplementary Fig. 12. Bars represent mean ± s.e.m. fold-change relative to wt; P determined by unpaired Student’s t-test

Subsequently, to assess putative changes at the translational level, we implemented quantitative western blot analysis of selected key triadic EC-coupling proteins in tibialis anterior (TA) muscles derived from adult ncDHPR and wt mice. Densitometric analyses of immunoreactive bands with glyceraldehyde 3-phosphate dehydrogenase (GAPDH) as internal reference, indicated comparable protein levels (P > 0.05, unpaired t-test) between both genotypes (Fig. 9b and Supplementary Figs. 11 and 12). Thus, our results clearly indicate that no adaptive transcriptional or translational regulation of key Ca2+-handling proteins occurs in the ncDHPR mouse, which could account for its unaltered muscle performance.

Overall, the surprising outcome of our study from the plethora of in vitro, ex vivo and whole-animal tests underlines the fact that DHPR Ca2+ influx is irrelevant for proper skeletal muscle functioning.

DHPR Ca2+ influx does not reduce skeletal muscle contraction

Previous work proposed29 that DHPR Ca2+ influx has a negative regulatory effect on skeletal muscle contraction by inhibition of the Ca2+-sensitive adenylyl cyclase (AC) isoforms, AC5 and AC6. Apparently, our knock-in ncDHPR mouse provides the perfect system to re-investigate this hypothesis. As intracellular SR Ca2+ release was unaltered in ncDHPR mice (refer Fig. 1d–f) in contrast to the described augmentation of Ca2+ release29, we first compared expression levels of AC5 and AC6 RNAs by qPCR assay with EEF1A2 as reference gene. Conversely, we did not find any adaptive transcriptional downregulation of either of the AC isoforms in skeletal muscles of neonatal ncDHPR mice compared (P > 0.05, unpaired t-test) with wt mice (Supplementary Fig. 13a). Likewise, there was no significant difference (P > 0.05, unpaired t-test) in expression levels of AC5 and AC6 in adult SOL and EDL muscles between both genotypes (Supplementary Fig. 10). The negative inotropic effect of Ca2+ influx was demonstrated in the diaphragm muscle by ex vivo blocking of DHPRs with the chemically distinct l-type Ca2+ channel blockers, nifedipine and verapamil29. Using an analogous approach, we compared the isometric contractility of diaphragm muscles, isolated from ncDHPR and wt mice, after treatment with the above mentioned drugs. Consistent with the previous findings29, 10 µM of nifedipine or 50 µM of verapamil induced a robust increment in muscle twitch contraction force in wt mice (Supplementary Fig. 13b, c, respectively). Surprisingly, and in stark contrast to the previous hypothesis29, application of 10 µM nifedipine to the ncDHPR diaphragm muscle also exhibited ~ 30% increase in isometric contraction force, which was indistinguishable (P > 0.05, unpaired t-test) from control wt muscle (Supplementary Fig. 13b). Similarly, the positive inotropic effect of verapamil ( ~ 45% increment in twitch force) was indistinguishable (P > 0.05, unpaired t-test) between ncDHPR and wt diaphragm muscles (Supplementary Fig. 13c). The increment in twitch force with both drugs could be reversed by washing. Altogether, in stark contrast to the previous findings29, our data from the ncDHPR mouse model provide compelling evidence that DHPR Ca2+ influx does not attenuate skeletal muscle contraction.

Discussion

In this study we investigated the physiological consequences of the elimination of skeletal muscle inward Ca2+ current in our newly generated k.i.-mutant mouse model (ncDHPR), which expresses a non-Ca2+-conducting (nc) DHPR in skeletal muscle. Despite more than 40 years of research, the role of this Ca2+ influx in mammalian skeletal muscle which occurs during conformational EC coupling is still obscure and controversial19. Theoretically, there are three possible roles for this enigmatic DHPR Ca2+ influx: (i) it still marginally contributes to skeletal muscle EC coupling via a rudimentary CICR effect; (ii) it is important as an exaptational current, i.e., during evolution it underwent a change in function from the crucial initiator of CICR to a Ca2+ current that serves another function now; or lastly, (iii) it is just a vestigial current, viz., an evolutionary remnant Ca2+ current from phylogenetically ancestral CICR stages3, 7,8,9,10,11,12 but without any functional importance.

Nevertheless, contemplating these three options it seems very unlikely that this DHPR Ca2+ current still contributes to EC coupling via rudimentary CICR, because it is small, slowly activating and considerably lags behind the activation of SR Ca2+ release40, 41. Moreover, previous biophysical experiments on skeletal muscles ex vivo, muscle fibres and myotubes from frogs to rodents, explicitly showed that pharmacological blocking of the DHPR or elimination of extracellular Ca2+ did not obstruct EC coupling14, 42,43,44,45 and, consequently, it is widely accepted that the DHPR Ca2+ influx is not an (immediate) prerequisite for the contractile activation19, 30.

However, substantial arguments against a complete and long-term insignificance of the DHPR Ca2+ influx (i.e., being vestigial) also exist, because it is apparently regulated in different ways. For example, the accessory DHPRα2δ-1 subunit functions as major determinant of the characteristic slow l-type Ca2+ current kinetics46, 47 and the DHPRγ1 acts similar to an endogenous Ca2+ antagonist that reduces Ca2+ entry under conditions promoting modest membrane depolarization48. DHPR Ca2+ influx is also massively ( ~ 8-fold) enhanced in developing fetal mouse skeletal muscle due to the expression of an alternatively spliced DHPRα1S subunit49, 50 and even more substantially amplified ( ~ 30-fold) by retrograde RyR1-DHPR coupling51, 52. Hence, the probability of this regulated skeletal muscle DHPR Ca2+ influx to function as an exaptational Ca2+ current cannot be ruled out. In this case its character would have completely changed, from a crucial EC coupling signal transmitter to a supporter of muscle cell Ca2+ homeostasis (e.g., SR store filling), tightly linked to the EC coupling process.

Eventually, to gain deeper and clearer insight into the role of DHPR Ca2+ conductivity in mammalian skeletal muscle, we engineered point mutation N617D from pore loop II of zebrafish DHPRα1S-b, identified to be responsible for the complete lack of Ca2+ conductivity in fast, glycolytic muscle2, into the murine gene CACNA1S (Supplementary Fig. 1). Whole-cell Ca2+ current measurements revealed that indeed this novel ncDHPR mouse model expresses a non-conducting Ca2+ channel in skeletal muscle (Fig. 1 and Supplementary Figs. 2 and 3). Survival of the ncDHPR mice and thriving into adulthood provided us a unique opportunity to study muscle physiology, development and performance in the absence of DHPR Ca2+ influx in young and aged skeletal muscle. Elimination of skeletal muscle DHPR Ca2+ influx might cause an imbalance in the intracellular Ca2+ levels. This would consequently lead to reduced muscle performance, which on the long term (with aging) might induce gross physiological effects such as myotonia or paralysis. However, very surprisingly, our results clearly demonstrate that the absence of the physiological DHPR Ca2+ influx in ncDHPR mice is irrelevant for SR store filling (Fig. 3). DHPR Ca2+ influx, in particular the massive conductance via the embryonic splice variant of the DHPRα1S subunit, has also been proposed to regulate acetylcholine receptor pre-patterning and formation of neuromuscular junction and thus to be crucial for skeletal muscle development23, 24. Contrary to this proposal, we found no motor coordination defects in ncDHPR mice. Notably, our findings also provide clear and firm evidences that the lack of DHPR Ca2+ influx in ncDHPR mice is inconsequential for body weight development, fertility, locomotor activity, muscle fibre-type composition, fibre CSA, muscle strength and susceptibility to fatigue, irrespective of the age. Thus, we can undoubtedly exclude putative age-related accumulative effects of the lack of DHPR Ca2+ influx on overall muscle health and performance. In addition, a reduction to half the size of whole-cell Ca2+ current density in heterozygous wt/ncDHPR mice compared with wt (Supplementary Fig. 9), points to a lack of compensatory upregulation of the DHPR activity. Furthermore, qPCR and quantitative western blot analyses of multiple key proteins of the EC coupling machinery and Ca2+ homeostasis did not indicate any transcriptional or translational up- or downregulation to compensate for the lack of DHPR Ca2+ influx in ncDHPR mice (Fig. 9 and Supplementary Figs. 1012). This is especially interesting for the proteins involved in SR Ca2+ store refilling via store-operated Ca2+ entry (SOCE) such as TRPC, STIM and Orai53, because it shows that despite the lack of ECCE in the ncDHPR mouse muscle the SOCE machinery is not upregulated to compensate for the loss of ECCE. However, analyses of mRNA or protein levels of these major players in EC coupling and Ca2+ homeostasis do not necessarily equate to their activity levels since a number of these proteins are posttranslationally regulated. Still, the evidence that these key proteins remain uninfluenced by the lack of DHPR Ca2+ influx is very high.

ECCE was described as store-independent Ca2+ entry pathway20 via the skeletal muscle DHPR22, activated by repetitive or prolonged depolarizations and to be essential for store repletion20, 53. The complete lack of Ca2+ influx in ncDHPR mouse myotubes even after 2-s prolonged depolarizations in the presence of the 1,4-dihydropyridine (DHP) agonist BayK864436, presents convincing evidence that skeletal muscle of the ncDHPR mouse does not require ECCE for its proper functioning.

Although the hypothesis that the DHPR Ca2+ current is vestigial seems strongly and clearly confirmed by all our findings, earlier studies on frog25, 26, crab27 and rat28 skeletal muscles pointed to positive inotropic effects of different l-type Ca2+ channel blockers. Recently, analogous inotropic effects were described by ex vivo Ca2+ channel-blocking experiments on mouse diaphragm muscle29. It was hypothesized that DHPR Ca2+ influx attenuates skeletal muscle contraction via inhibition of Ca2+-sensitive AC isoforms, AC5 and AC6, which in turn leads to reduced intracellular cAMP levels, known to potentiate muscle contraction mainly through PKA-dependent RyR1 phosphorylation and thus preserving muscle fibre integrity29. With our ncDHPR mouse model we re-examined this apparently paradoxical Ca2+ antagonist effect on muscle contraction. As demonstrated explicitly in the Results section, perfusion of isolated diaphragms of ncDHPR mice with two chemically distinct organic l-type Ca2+ channel blockers, the DHP nifedipine and the phenylalkylamine (PAA) verapamil, surprisingly and in stark contrast to the hypothesis of Menezes-Rodrigues et al.29, induced identical positive inotropic effects in wt and ncDHPR mice (Supplementary Fig. 13). With this experimental series, we could clearly exclude that Ca2+ influx via the DHPR has a negative regulatory effect on skeletal muscle contraction and thus it is not essential for overall muscle health and fibre integrity as proposed in the earlier study29. However, our results cannot provide an explanation for this (Ca2+-influx independent) apparently paradoxical Ca2+ antagonist effect on muscle contraction. The most likely reason that we can speculate is that the antagonist concentrations (10 µM nifedipine and 50 µM verapamil) used in all these ex vivo experiments are ~ 6,000- and 25,000-fold, respectively, beyond the K d-values observed for skeletal muscle membrane binding54, 55 and thus might bind and interact non-specifically to low-affinity binding sites of other muscle proteins besides its specific high-affinity binding pocket on the DHPRα1S 56. It is quite possible that such undefined low-affinity protein–DHP/PAA complexes are responsible for the observed amplification of skeletal muscle contraction.

Furthermore, our results and conclusions are also incongruent with a study on another mouse model (E1014K mouse) with a non-Ca2+-conducting DHPR31. With this mouse model, it has been proposed that Ca2+ permeation through (or just Ca2+ binding to) the DHPR has a significant impact on muscle function. In contrast to the ncDHPR mouse model, the E1014K mouse displayed decreased SR Ca2+ store refilling, increased fatigue, decreased muscle fibre size, increased type-IIb fibre count31 and an obese phenotype57. The most probable reason for this disparity might be different biophysical consequences due to the implementation of distinct DHPR pore mutations. In the ncDHPR mouse, the N617D pore mutation adds a negative charge in close vicinity of the selectivity filter glutamate in repeat II of the DHPRα1S, which induces an aberrant high-affinity Ca2+ binding to the pore36. This results in complete occlusion of the pore for all ions under physiological conditions. Conversely, in the E1014K mouse model, the E to K charge conversion in repeat III leads to impairment of the Ca2+ selectivity filter, enabling only low-affinity (mM) Ca2+ binding to the pore32, 33. Thus, the E1014K-DHPR does not allow Ca2+ permeation but shows a strong conductivity for monovalent cations31, 32, which consequently turns it into a junctionally targeted, slowly activating, non-inactivating Na+ channel34.

It is quite possible that this E1014K-specific leakiness for monovalent cations causes the observed alterations in muscle performance and is responsible for the differences between the two mouse models. Alternatively (or additionally), a proper conformation might be required for fine-tuning the DHPR signalling function, which is stabilized by high-affinity Ca2+ pore-binding (as in the case of wt or N617D-DHPR) and destabilized by low-affinity Ca2+ pore-binding (as in the case of E1014K-DHPR). Only this option allows discussing the claim of the study on the E1014K model that Ca2+ permeation through and/or binding to the DHPR pore has an important modulatory role in muscle performance31. As the ncDHPR mouse does not show any of the aberrant muscle phenotypes found with the E1014K model, we can undoubtedly exclude the importance of DHPR Ca2+ permeation for skeletal muscle performance. However, as Ca2+ binds firmly to the N617D DHPR pore36 in contrast to the E1014K channel, a putative crucial sole Ca2+ binding effect to the DHPR to sustain skeletal muscle function cannot be excluded.

We are aware of the innate dilemma of a study, which intends to describe the non-existence of a phenotype and thus, despite an exhaustive scope of experiments, the fact that a stone was left unturned cannot be excluded. Nevertheless, all our results strongly suggest that the DHPR-mediated Ca2+ current is fully dispensable in mammalian skeletal muscle and is most likely a non-functional, tolerated evolutionary remnant (vestigial Ca2+ current) from the ancestral Ca2+-influx-dependent skeletal muscle EC coupling found in early chordates3, 7, 10. The final switch-off of this vestigial skeletal muscle DHPR Ca2+ influx took place only in the phylogenetically most derived clade among the vertebrates—the euteleost fishes2, 3.

Methods

Generation of the ncDHPR knock-in mouse

The strategy for transforming the mouse DHPRα1S non conducting via introduction of point mutation N to D into exon 13, simultaneously with a silent mutation for restriction enzyme site PflF1 for RFLP test, was deliberated by our lab. Finally, the ncDHPR mouse on C57BL/6NTac background was generated at the Institut Clinique de la Souris—ICS (Illkirch, France).

Animal care and all experimental procedures were conducted in strict accordance with the guidelines of the European Union Directive 2010/63/EU and approved by the Austrian Ministry of Science (BMWF-5.031/0001-II/3b/2012). Mice were housed on a 12/12-h light/dark schedule, with access to food and water ad libitum. At least 24 h before the start of the experiments mice were housed individually and acclimatized to the behavioural facility. All experiments were performed on mutant and gender- and age-matched wt mice. Mice were organized into two age groups, “young” between 3 and 7 months and “aged” between 14 and 22 months old. Only male mice from both age groups were used in this study, unless indicated otherwise.

Primary culture and whole-cell patch clamp on myotubes

Myoblasts from skeletal muscles of new born up to 5 days old pups of wt, heterozygous or homozygous for non-conducting DHPRs were isolated and plated on 35-mm ECL-coated (Merck Millipore) culture dishes in Dulbecco’s modified Eagle’s medium in a 37 °C incubator with 5% CO2 58. l-type Ca2+ currents simultaneously with intracellular SR Ca2+ release using 0.2 mM Fluo-4 in the patch pipette solution were recorded from these cultured myotubes, as described59. Myosin-II blocker N-benzyl-p-toluene sulphonamide (BTS, 100 µM, Sigma) was continuously present in the bath solution.

Two-electrode voltage clamp on isolated adult muscle fibres

Measurements on voltage-clamped muscle fibres and signal analysis were performed as described60. Briefly, single muscle fibres were enzymatically isolated from interosseous muscles of young mice, voltage-clamped on a two-microelectrode setup and dialysed with an artificial intracellular solution containing 0.2 mM Fura-2 and 15 mM EGTA, to allow parallel fluorescence recordings of Ca2+ currents and SR Ca2+ release. The high intracellular EGTA concentration dominates Ca2+ binding in these experiments and facilitates the calculation of the time course of the Ca2+ release flux from the SR61. Depolarization-induced fluorescence ratio signals (380 and 360 nm excitation) were subjected to a removal analysis using a Ca2+ distribution model fit approach60. The results were used to estimate the voltage-dependent Ca2+ release flux and converted to SR Ca2+ permeability by correcting for putative SR Ca2+ depletion during the depolarizing voltage steps60.

SR Ca2+ content measurements on isolated adult muscle fibres

Primary-cultured interosseous muscle fibres62 were loaded with 5 µM of the ratiometric low-affinity Ca2+ indicator dye Fura-FF-AM in HEPES-buffered standard Krebs–Ringer solution for 30 min at room temperature, washed and equilibrated with Ringer solution containing 100 µM BTS for another 30 min to supress contractions. The bath solution was then replaced with Ca2+-free Ringer solution to eliminate the influence of external Ca2+. Target cells were superfused using Ca2+ releasing solution, i.e., Ca2+-free Ringer solution containing 500 µM of the RyR agonist 4-CmC to trigger Ca2+ release and 30 µM CPA to inhibit SERCA Ca2+ sequestration. A period of 20-s application of the Ca2+ releasing solution was always bracketed with 20-s superfusion with Ca2+-free Ringer solution (Fig. 3a). For Ca2+ measurements, the ratio of fluorescence at 340 and 380-nm excitation was determined using the Zeiss Microscope Photometer System (FFP) based on an inverted microscope (Axiovert 35, Zeiss) equipped for epifluorescence (Fluar ×40/1.3 oil-immersion objective)63.

Fibre-type and fibre CSA analysis

Eight-micrometre-thick cryosections of snap-frozen SOL and EDL muscles from 2 to 3 months old ncDHPR and wt mice were immunostained overnight at 4 °C with antibodies against fibre-type-specific myosin heavy chain (MHC) isoforms. Primary antibodies against MHC isoforms I (BS-D5), IIa (SC-71), IIb (BF-F3) and IIx (6H1) (Developmental Studies Hybridoma Bank, USA) at a dilution of 1:2,000, except 6H1, which was 1:200, were used. Secondary staining was performed for 1 h at room temperature (RT) with anti-mouse IgG Alexa 594 or anti-mouse IgM Alexa 594 (1:4,000, Invitrogen). Images were taken under a fluorescence microscope (Olympus BX53) with a ×20 objective lens. The number of antibody-stained fibres were counted and normalized to the total number of muscle fibres per field64. Saved images were additionally used for calculating the fibre CSA with Metamorph software (Molecular Devices, USA).

Ex vivo force frequency and fatigue measurements

Intact muscle-tendon complexes of EDL and SOL were dissected from the hindlimbs of euthanised young or aged wt or ncDHPR mice65 and mounted vertically between a force transducer (Model FT03, Grass Instruments, Quincy, USA) and static clamp in an organ bath equipped with platinum electrodes and under continuous perfusion with carbogen-(95% O2 + 5% CO2) saturated Krebs–Ringer solution. Optimal muscle stretch was determined by applying single twitches at supramaximal voltage (25 V for 1 ms) and set at the length that generated maximal force. After 10 min of equilibration, EDL and SOL muscles were subjected to different force frequency (tetani with increasing stimulation frequencies but fixed 2-min recovery intervals) and repetitive tetanic fatiguing (decreasing recovery intervals by 27% every 2 min) protocols38 (Supplementary Fig. 7). All experiments were performed at room temperature (~26 °C). Data acquisition and analysis was carried out using custom made software (Delphi, Borland). Muscle length, diameter and wet weight were measured at the end of each experiment.

Voluntary locomotor activity measurements

Spontaneous physical activity of 3–6 months old male mice was measured in their home-cage environment with free access to food and water. Mice were singularized at least 1 week before the start of the experiment.

Spontaneous total activity was monitored by a home cage ambulatory activity test over a period of 2 days and 3 nights with an infrared sensor unit (InfraMot, TSE Systems, Homburg, Germany) mounted on top of every home cage.

For voluntary activity wheel test, mice were individually habituated for 2 days before the start of the experiment in cages furnished with activity wheels (Bioseb, Vitrolles, France). The mice had continuous free access to the wheels, which could be revolved in either direction. Activity wheels were equipped with photoelectric sensors interfaced to a computerized system that automatically recorded the distance travelled every 5 min continuously for 21 days. Up to six mice were run in parallel and for each genotype the distances travelled per day were calculated and plotted as averaged, cumulative values.

Muscle coordination, strength and endurance measurements

Three standard tests were used to compare the motor coordination and muscle strength between ncDHPR and wt mice.

For the beam walk test, ncDHPR or wt mice from the young group were used. The mouse was placed on one end of a 120-cm long beam resting 50 cm above a padded surface. An enclosed safety box was mounted at the other end of the beam. After three pre-training trails, the time needed to traverse the beam and reach the safety box was recorded for every mouse.

For the wire hang test, a custom-designed device consisting of a cross-wired grid, positioned between two poles 35 cm apart and 55 cm above a padded surface was constructed. The mouse was placed in the centre of the grid and shaken gently three times, to ensure firm gripping to the wire (1.7 mm in diameter) with all four paws. The grid was then immediately turned upside down and the latency to fall was recorded. Three trials were done with every mouse, with a recovery interval of some minutes, and only the highest score was used for analysis. The maximum hanging time was set to 600 s and the mouse that did not fall during this period was removed and given the maximum score. Body weight was recorded at the end of the experiments and the average weight was similar between ncDHPR and wt mice.

Maximal muscle strength of the forelimbs was measured using a grip strength test (Bioseb). The mouse was gently lowered by the tail and allowed to grip a metal bar (1.3 mm in diameter) with the front paws only. Keeping the torso horizontal, a slow gradual pull was exerted in backward direction until the grip was released. The force applied to the bar by the animal at the time of the release is recorded by the sensor as the maximum force reached. Three measurements separated by 30-s inter-trial intervals were collected from each animal and the maximum value was used for statistical analysis. Mice were weighed at the end of the testing session and there was no difference in average weight between ncDHPR and wt mice.

Acute fatigue measurements

Two standard methods were used to compare fatigability and endurance between ncDHPR and wt mice.

Rotarod tests at accelerating and constant speeds were used to assess muscle fatigue and endurance. Mice were familiarized and trained on a four-lane rotarod (TSE Systems) by running them for 5 min per day for 3 days at a speed of 4 rpm. For testing the fatigability, young and aged ncDHPR and wt mice were placed on the rotarod at an initial speed of 4 rpm for 25 s. After the warmup phase, the rotarod was set to accelerate steadily from 4 to 40 rpm for 300 s. Each mouse was subjected to three trials with 15 min inter-trial intervals. For rotarod endurance task, mice were placed on the rod rotating at a constant speed, 20 rpm for young and 15 rpm for aged mice for 600 s. Latency to fall was recorded and only the highest score among the 3 trails was considered for analysis.

Treadmill test was performed to assess the degree of fatigability under rigorous conditions. Mice were run on a five-lane treadmill with electric stimulus grid (Bioseb). Three days before the experiment, mice were acclimatized and trained on the treadmill by running them for 5 min per day at a speed of 9 cm s−1 at a 15° inclination. Acute exhaustion protocols following an initial 5-min warm up at 5 cm min−1 were designed for young and aged mice. In case of young mice, band speed was increased 2 cm s−1 every minute till 27 cm s−1 and then maintained at the maximum speed for an additional 45 min. The entire treadmill task was 0.9 km in 1 h. For aged mice, a milder protocol with a speed increment of 1 cm s−1 every minute up to 17 cm s−1 was used. The entire run was 0.55 km distance in 1 h. To encourage continuous forward running, the front end of the treadmill was covered with a black cloth and electric stimulus at the back end was set at 0.2 mA. The number of rests and body weight was recorded for each mouse. Finally, the average cumulative resting time collapsed into 5 min bins, was plotted against the duration of the task. In addition, average number of cumulative rests during the entire task was calculated.

RNA isolation and qPCR

Putative transcriptional regulation of key triadic proteins involved in EC coupling and Ca2+ homeostasis was assessed by comparative TaqMan quantitative reverse-transcriptase PCR (qRT-PCR). Total RNA was extracted from skeletal muscle of neonatal pups (up to 2 days old) and adult (7 months old) EDL and SOL muscles using RNeasy Mini Kit (Qiagen). Following RNAse-free DNAse treatment, first-strand complementary DNA synthesis was performed using random primers and M-MLV reverse transcriptase (Promega). For every gene, flanking primer pairs and 5′ FAM (6-carboxyfluorescein) and 3′ BHQ1 (Black Hole Quencher)-labeled TaqMan probe spanning exon–exon boundaries, were designed and purchased from Eurofins, Germany (Supplementary Table 2). qRT-PCR assays were performed in triplicates with GoTaq Probe qPCR master mix (Promega) on a Real-Time thermal cycler (Montania 483, Anatolia Geneworks, Turkey). Relative expression levels were calculated using the comparative C T method with two endogenous reference genes—ACTB (β-actin) and one of the most stable mammalian skeletal muscle housekeeping gene, EEF1A2 (eukaryotic translation elongation factor 1 alpha 2)66.

Western blot analysis

TA muscles from 2 to 3 months old ncDHPR and wt mice were excised, homogenized and lysed in sample buffer containing 250 mM Tris-HCl (pH 8.5), 2% lithium dodecyl sulfate, 10% glycerol and 0.5 mM EDTA and centrifuged at RT, 16,000×g for 5 min. Protein concentration measurements and separation by SDS-polyacrylamide gel electrophoresis on 3–8% Tris-Acetate or 4–12% Bis-Tris NuPAGE gels (Thermo Fisher Scientific) was performed as described previously67. Immunostaining was performed with different primary antibodies followed by appropriate secondary antibodies (Supplementary Table 3). Immunoreactive bands were visualized on the Odyssey scanner (LI-COR). Western blot densitometries were performed using Image Studio Lite software, with GAPDH as a loading control. Uncropped images of western blots are shown in Supplementary Fig. 12.

Ex vivo isometric contraction recordings on diaphragm muscle

Three- to 5-mm strips with rib overhangs were dissected from diaphragm muscle isolated from euthanized 3–6 months old ncDHPR or wt mice. Strips were mounted vertically in the ex vivo contraction setup (as described above) with the central tendon facing up and the rib end towards the bottom and kept under continuous perfusion with carbogen (95% O2 + 5% CO2) saturated Krebs–Ringer solution. Optimal muscle stretch was determined by applying a single twitch at supramaximal voltage (25 V for 2 ms). After 10 min of equilibration under continuous stimulation (0.33 Hz, 2 ms, 25 V), nifedipine (10 µM) or verapamil (50 µM) was added to the bath solution to investigate the effect of l-type Ca2+ channel blockers on muscle contraction29. Nifedipine was washed out after 25 min and stimulation continued for another 15 min, whereas verapamil was washed out after 40 min with further stimulation for 20 min. At the end of each experiment, muscle wet weight, diameter and length were measured. All recordings were performed at room temperature ( ~ 26 °C). Baseline force was calculated by taking the average twitch peak force of the initial 6 min before adding the drug. Finally, the average peak twitch force, collapsed into 2-min bins and expressed as percentage of baseline force, was plotted against the duration of the experiment.

General experimental design and statistical analyses

Sample sizes of mice or tissue are based on previous publications2, 29, 31, 38, 39, 64; hence, power calculations were not necessary. Animals were arbitrarily allocated to the specific groups (according to genotype, sex and age), but without the use of an explicit randomization procedure. Experiments did not require blinding and thus were not performed under blinded conditions. In our study, no data points, samples or mice were excluded from analysis. Based on our previous publications2, 3, 35, 44, 45, 52, 56, 59, all statistical analyses are considered appropriate. n-Values represent the number of mice, muscles, muscle fibres or myotubes, as specified. Variance is similar between comparison groups. All data are expressed as means ± s.e.m. and statistical analyses were conducted using unpaired Student’s t-test. P < 0.05 was considered statistically significant.

Data availability

All relevant data are available from the first/corresponding author on reasonable request.

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Acknowledgements

We thank Dr Christoph Schwarzer for providing support and access to the Home cage and Rotarod setup, Dr Gerald Obermair for help with the qPCR analysis, Drs Zoran Culig and Florian Handle for helping with western blot analysis, Dr Bernhard Flucher and Stefanie Geisler for providing support in examining the muscle fibre-type composition, Dr Veit Flockerzi for providing the anti-Orai1 antibody, Drs Robert T. Dirksen and Lan Wai-LaPierre for most of the other WB antibodies and Achim Riecker for help with the contraction setup. We also thank Birgit Kagerbauer and Karin Fuchs for excellent technical assistance. This study was supported by the Austrian Science Fund (FWF) Grants (P23229, P27392 and DK-W1101 to M.G.) and W.M. received funding from the Deutsche Forschungsgemeinschaft (DFG) Grant (ME-713/18).

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A.D. and M.G. designed research; Y.P. performed electrophysiology on muscle fibres and together with K.F. the SR Ca2+ loading experiments; A.D., M.G. and K.S., W.M. performed all the other experiments; and A.D. and M.G. wrote the manuscript.

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Correspondence to Manfred Grabner.

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Dayal, A., Schrötter, K., Pan, Y. et al. The Ca2+ influx through the mammalian skeletal muscle dihydropyridine receptor is irrelevant for muscle performance. Nat Commun 8, 475 (2017). https://doi.org/10.1038/s41467-017-00629-x

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