Arrhythmogenic late Ca²⁺ sparks in failing heart cells and their control by action potential configuration

Significance

Sudden cardiac death in heart failure is a major unsolved clinical problem that is linked to the development of a spontaneous arrhythmia. Early afterdepolarizations (EADs) are an arrhythmogenic mechanism, but the cellular trigger for EADs in heart failure is unclear. We show that the reduction in synchronous Ca²⁺ release early in the action potential (AP) of failing cardiac myocytes promotes the appearance of late Ca²⁺ sparks which can propagate, forming Ca²⁺ ripples and waves. These, in turn, produce an inward sodium–calcium exchange current which opposes AP repolarization. Restoration of AP phase 1 repolarization improved Ca²⁺ release synchrony and reduced late Ca²⁺ spark rate, suggesting a different approach to reducing the risk of sudden death in heart failure.

Abstract

Sudden death in heart failure patients is a major clinical problem worldwide, but it is unclear how arrhythmogenic early afterdepolarizations (EADs) are triggered in failing heart cells. To examine EAD initiation, high-sensitivity intracellular Ca²⁺ measurements were combined with action potential voltage clamp techniques in a physiologically relevant heart failure model. In failing cells, the loss of Ca²⁺ release synchrony at the start of the action potential leads to an increase in number of microscopic intracellular Ca²⁺ release events (“late” Ca²⁺ sparks) during phase 2–3 of the action potential. These late Ca²⁺ sparks prolong the Ca²⁺ transient that activates contraction and can trigger propagating microscopic Ca²⁺ ripples, larger macroscopic Ca²⁺ waves, and EADs. Modification of the action potential to include steps to different potentials revealed the amount of current generated by these late Ca²⁺ sparks and their (subsequent) spatiotemporal summation into Ca²⁺ ripples/waves. Comparison of this current to the net current that causes action potential repolarization shows that late Ca²⁺ sparks provide a mechanism for EAD initiation. Computer simulations confirmed that this forms the basis of a strong oscillatory positive feedback system that can act in parallel with other purely voltage-dependent ionic mechanisms for EAD initiation. In failing heart cells, restoration of the action potential to a nonfailing phase 1 configuration improved the synchrony of excitation–contraction coupling, increased Ca²⁺ transient amplitude, and suppressed late Ca²⁺ sparks. Therapeutic control of late Ca²⁺ spark activity may provide an additional approach for treating heart failure and reduce the risk for sudden cardiac death.

Worldwide, ∼26 million people suffer from heart failure (HF) (1). More than 50% of HF patients die suddenly, and this sudden cardiac death is most likely due to the spontaneous emergence of arrhythmias (2, 3). At the cellular level, two identified initiators of cardiac arrhythmias are delayed afterdepolarizations (DADs) and early afterdepolarizations (EADs) (4). DADs occur in the resting period between heart beats (diastole) and are due to spontaneous Ca²⁺ release from the sarcoplasmic reticulum (SR) in the form of Ca²⁺ sparks (5), which summate to form propagating Ca²⁺waves (6, 7). These waves cause a depolarizing inward current via the Na⁺/Ca²⁺ exchange (NCX) mechanism (6, 8, 9). In contrast to DADs, EADs are less well understood and occur during the repolarization phase of the cardiac action potential (AP) where several ionic currents interact to control repolarization (10). EADs can be produced by reactivation of ionic currents during AP repolarization when the potassium currents forming the “repolarization reserve” (11, 12) are insufficient to maintain the repolarization trajectory of the AP, although why this should occur spontaneously within a steady train of APs is uncertain (13). Spontaneous Ca²⁺ waves have also been implicated in EAD generation (9), but it is unclear how such waves might be initiated when the SR should be depleted and/or refractory after SR Ca²⁺ release is triggered by the upstroke of the AP (14).

We recently showed that “late” Ca²⁺ sparks (LCS) can occur during the decay of the Ca²⁺ transient in normal rabbit cardiac myocytes (15). Here, we report that stochastic LCS and low-amplitude “Ca²⁺ ripples” are promoted in HF and can generate sufficient NCX current (I_NCX) to trigger EADs. In addition, normalizing the early AP waveform in HF promotes a faster and larger SR Ca²⁺ release (which should improve contractility) and at the same time decreases arrhythmogenic LCS activity.

Results

LCS occur in a wide variety of species (SI Appendix, Fig. S1), but, to study repolarization mechanisms, an AP similar to human is desirable, and we therefore used an established rabbit model of chronic myocardial infarction that causes HF. When left ventricular ejection fraction had fallen to 44 ± 3% (SI Appendix, Fig. S2 and Table S1), indicative of moderate to severe HF, ventricular myocytes exhibited t-tubule remodeling (Fig. 1 A and B) similar to that seen in most models of HF (16). This “t-tubule disease” (17) decreases the efficiency of excitation–contraction coupling and the rate of rise of the Ca²⁺ transient (SI Appendix, Fig. S3) (18, 19). Although there was no change in I_Ca density (Fig. 1C) in this HF model, the AP lost phase 1 repolarization (red arrow, Fig. 1D) and became prolonged (SI Appendix, Fig. S3). All of these changes are also seen in human HF (17, 20). The loss of AP phase 1 contributes to the reduced SR Ca²⁺ release rate (Fig. 1E and SI Appendix, Fig. S3) and increased SR release latency (Fig. 1F) (16, 18) because this phase of the AP affects the magnitude and time course of I_Ca which triggers SR Ca²⁺ release (21, 22).

Fig. 1.

Changes in structure, AP time course, t-tubule morphology, and increased LCS in HF. (A) di-8-ANEPPS staining shows reduced tubule regularity and (B) reorientation of t-tubules in HF cells compared to control (CON). n/N = 9/4 CON 12/5 HF. (C) I_Ca density was unchanged in HF. n/N = 20/4 CON 10/3 HF Representative currents at the indicated test potentials (mV) are shown (Inset). Error bars not shown where error is less than the size of symbols. (Scale bars: 5 pA/pF, 10 ms.) (D) APs in HF myocytes had less AP phase 1 repolarization (arrow). (E) Ca²⁺ transients in CON myocytes showed synchronous Ca²⁺ release, while HF showed less uniform Ca²⁺ release and (F) an increase in latency from electrical stimulation to detectable Ca²⁺ increase. (G) Image processing reveals increased LCS frequency in HF with some Ca²⁺ ripples (yellow dashed box). (H) LCS frequency was greater in HF cells than in CON. n/N = 16/6 CON; 14/5 HF. **P < 0.01, ***P < 0.001, unpaired t tests.

The reduction in synchronous Ca²⁺ release in HF, seen as a less uniform increase in Ca²⁺ after the AP upstroke (Fig. 1E, arrow), was associated with an increase in the number of LCS (Fig. 1 G and H). This ∼3-fold increase in LCS frequency can be explained by an increase in the number of Ca²⁺ release sites that were not triggered earlier during the AP (15, 22). The increased availability of Ca²⁺ release sites also promotes sequential activation of LCS, resulting in Ca²⁺ ripples (15), an example of which can be seen within the yellow dashed box in Fig. 1G.

Linkage between LCS and Arrhythmogenesis.

LCS activity depends on SR Ca²⁺ content (15) which generally increases with AP frequency (10, 23). A pacing-pause protocol increases SR Ca²⁺ and may provoke EADs (24). With this protocol, LCS activity increased, and EADs appeared (Fig. 2A). The high rate of LCS production (and appearance of Ca²⁺ ripples; see SI Appendix, Fig. S4) in these conditions clearly opposed the normal decline of Ca²⁺ during the Ca²⁺ transient.

Fig. 2.

LCS can trigger EADs. (A) EADs evoked by pacing-pause protocol. During repolarization, an initial V_m inflection at ∼ −20 mV generates EADs (Top), and numerous LCS create oscillatory Ca²⁺ release (Bottom). (B) Calculation of I_net for a typical HF AP. The filled region indicates where the greatest number of LCS occur (A and Fig. 4E) and corresponds to an I_net of ∼0.18 pA/pF. Any balancing current that counteracts I_net (i.e., −I_net, Bottom) will stop repolarization (SI Appendix, Fig. S5). (C) Failing AP with voltage-clamp steps used to probe Ca²⁺ and currents underlying EADs. (D) During the AP steps, LCS-triggered Ca²⁺ waves (Top) and inward currents (black trace) appear. Dashed line shows the inward current that balances I_net in B and could therefore stop repolarization. This current occurs when LCS are about to initiate Ca²⁺ waves. (E) Example of apparently chaotic Ca²⁺ during EADs in an HF cell (Top). Image processing reveals many LCS and Ca²⁺ ripples during V_m oscillations (red trace). The filled regions show where the depolarizing current (I_net) is increasing, and this corresponds to increasing Ca²⁺ (blue) and LCS activity (orange bars).

In order to generate an EAD, the net cellular repolarizing current (I_net) and/or repolarization reserve (11, 12) must be decreased and/or counteracted by depolarizing membrane current. While I_net is the sum of all inward and outward currents, the rate of change of membrane potential (dV_m/dt) is directly proportional to I_net because I_net = −C_m·dV_m/dt (where C_m is membrane capacitance). Applying this equation allows us to determine how much current is responsible for the repolarization phase of the AP, and Fig. 2B illustrates the magnitude and time course of I_net.. In this exemplar, I_net was only ∼0.18 pA/pF during the late AP repolarization phase, and the average I_net during AP repolarization was 0.26 ± 0.08 pA/pF (mean ± SD n/N = 15/4), showing that a small change in membrane currents will affect repolarization trajectory. It follows that any additional inward current of similar amplitude to I_net could stop AP repolarization (SI Appendix, Fig. S5) and thereby initiate an EAD.

An increase in intracellular Ca²⁺ moves the electrochemical gradient for NCX Ca²⁺ transport outward, causing an inward shift of I_NCX (25) which can be up to ∼1 pA/pF in magnitude (26). To examine whether the observed LCS activity could produce sufficient I_NCX to oppose I_net, we voltage-clamped cells with a failing AP which included steps to different holding potentials around the times (and V_m) where EADs occur (Fig. 2C). During these V_m steps, many LCS occurred which triggered Ca²⁺ ripples and Ca²⁺ waves. The average Ca²⁺ (blue trace) and evoked inward current (black trace) are shown in Fig. 2 D, Lower. This inward current had a linear relationship to mean Ca²⁺ with a slope of −1.5 pA/pF/µmol/L (SI Appendix, Fig. S6), compatible with the expected Ca²⁺ dependence of I_NCX (26). The critical inward current (from Fig. 2B) that will balance I_net and stop repolarization is indicated by the dotted line which, after projection onto the Ca²⁺ records, reveals the Ca²⁺ changes associated with this critical current density. It is notable that this current density is developed just before the onset of Ca²⁺ waves which appear as chevron patterns in line scans (5, 7), showing that LCS and/or Ca²⁺ ripples can be the initiating events for EADs.

Fig. 2E shows that multiple Ca²⁺ ripples, rather than Ca²⁺ waves, can also give rise to EADs in current-clamped cells. In this exemplar, the line scan Ca²⁺image (upper panel) is rather chaotic with no obvious Ca²⁺ waves being present. However, after image processing, numerous LCS events are seen, some of which form propagating Ca²⁺ ripples (cf. SI Appendix, Fig. S4). V_m and I_net can be linked to Ca²⁺ by two mechanisms: (1) Either Ca²⁺ activates inward I_NCX producing an inward shift of I_net and depolarization of V_m or (2) V_m activates L-type Ca²⁺ channels (LTCCs) which then trigger SR Ca²⁺ release. As shown by the colored bars in Fig. 2E, the rise in Ca²⁺ occurred in synchrony with inward I_net (peak value ∼ 0.15 pA/pF) which also supports the idea that the initiation of EADs can result from the summation of LCS and Ca²⁺ ripples which activate inward I_NCX.

Model Analysis of the Role of I_Ca and I_NCX in EADs.

The importance of I_NCX for EAD generation has also been demonstrated in murine cells by heterozygous NCX knockdown and in failing rabbit ventricle by acute NCX blockade (27, 28). To further test the idea that sufficient I_NCX can be generated by LCS to trigger an oscillatory EAD, we modified a new computer model of spatially distributed Ca²⁺ signaling (29) to calculate membrane currents that should occur in response to experimental V_m and Ca²⁺ signals (Fig. 3A). This approach breaks the intrinsic feedback loop(s) that otherwise make experimental dissection of causation problematic. Utilizing the recorded Ca²⁺ and V_m data shown in Fig. 2 as exemplar controlling variables, the model exhibited two types of behavior: when Ca²⁺ and LCS activity was high (Fig. 3Ai), I_NCX was in phase with dV/dt and of sufficient magnitude to explain the start of the V_m oscillations (cf. Fig. 2 D and E). However, when SR Ca²⁺ release was slightly reduced, changes in I_NCX were much smaller because the change in V_m opposed the change in electrochemical gradient for Ca²⁺ extrusion by NCX (Fig. 3Aii). The relative roles of I_NCX and I_Ca in initiating the EAD can be examined by cross-correlation with V_m. The magnitude of the negative peak in the cross-correlogram shows the strength of the linkage between these inward currents and V_m, while the time delay associated with the peak in the correlogram indicates whether they are the consequence of a change in V_m (by lagging behind V_m) or a potential cause (by leading the change in V_m). When the Ca load was high, inward I_NCX clearly preceded the change in V_m (Fig. 3Bi), consistent with the idea that the EADs were triggered by LCS activity and the consequent change in I_NCX. In comparison, I_Ca was both more weakly correlated and almost in phase with V_m. When SR load was decreased, the reduction in LCS activity led to a smaller inward I_NCX being less well correlated with V_m and with inappropriate phase to explain EAD initiation (Fig. 3Bii). However, I_Ca became more strongly correlated with V_m (due to a decrease in Ca²⁺-dependent LTCC inactivation (10)) and had appropriate timing to explain EAD initiation. Thus, while both mechanisms for EAD generation were present in the model, an increase in Ca²⁺ release caused the dominant inward current trigger for EAD generation to shift from I_Ca reactivation to LCS-evoked I_NCX.

Fig. 3.

Computer simulations of experimental traces using a model that generates LCS (29). Panels are ordered to indicate the flow of causation. (Ai) Increased LCS frequency is in phase with inward I_NCX and dV_m/dt, which precedes V_m and I_Ca forming an oscillatory feedback pathway as shown by the arrows. (Aii) When Ca²⁺ load is reduced, V_m oscillates because I_Ca (and possibly I_Na) supply the inward current to increase dV_m/dt and cause the EAD, as shown. In this case, LCS (and Ca²⁺) are the consequence of the reactivation of I_Ca. Note that in this simulation, I_NCX bears no obvious relation to dV_m/dt (unlike the results shown in Ai). Cross-talk between the oscillators shown in Ai and Aii can occur because they contain common mechanisms. (B) Cross-correlograms between V_m and both I_Ca and I_NCX from the data shown in part A. The negative peaks in the cross-correlogram (black arrows) reveal both the strength of association (amplitude) and temporal relationship (offset from V_m) between V_m and I_Ca or I_NCX (see SI Appendix). (Bi) When Ca²⁺ load was high, inward I_NCX preceded V_m oscillations and was more strongly correlated with V_m than I_Ca. (Bii) Under reduced Ca²⁺ load, I_NCX lags behind V_m oscillations. I_Ca reactivation preceded V_m and so has the required temporal relationship to explain EAD initiation, although some inward I_NCX may still contribute (13).

Improving Ca²⁺ Signaling and Suppressing LCS in HF Cells.

Fig. 4A shows that a normal AP applied to HF cells increased the rate of rise and amplitude of the Ca²⁺ transient, and this was associated with an increase in Ca²⁺ release synchrony. On average, the normal AP increased the rate of rise of the Ca²⁺ transient from 30 ± 8 nmol/L/ms to 40 ± 9 nmol/L/ms (P < 0.01, paired t test), while the time to peak decreased from 75 ± 9 ms to 65 ± 9 ms (P < 0.01, paired t test). This improvement in excitation–contraction coupling efficiency was also reflected by a decrease in the latency for Ca²⁺ release (Fig. 4B). With the improvement in synchrony of early release, the duration of the Ca²⁺ transient became shorter (Fig. 4C) due to an ∼50% decrease in LCS frequency (Fig. 4 D and E). These results confirm that at least a part of the nonuniform and delayed SR Ca²⁺ release seen in the HF model is due to AP phase 1 effects on I_Ca.

Fig. 4.

Applying a normal AP to HF cells improves excitation–contraction coupling and reduces LCS frequency. (A) Ca²⁺ transient upstroke velocity, amplitude (orange), and Ca²⁺ release synchrony all increase in an HF cell (Left) when voltage-clamped with a normal AP (Right). (Lower) Improved synchrony in Ca²⁺ release. (B) Mean latency for Ca²⁺ release and (C) Ca²⁺ transient duration were reduced in HF cells clamped with a normal AP. (D) Processed Ca²⁺ images show fewer LCS in HF after applying normal AP. (E) Number of LCS detected in HF with failing AP waveform (HF AP, 404 LCS) and with a normal AP (+Norm AP, 217 LCS) from 13 HF cells binned by V_m during the AP repolarization. **P < 0.01, paired t test n/N = 13/5.

It may not be possible to completely restore the maximum rate of rise, time to peak Ca²⁺, and Ca²⁺ transient duration to CON values with phase 1 modification alone because (1) the t-tubule disruption associated with HF would still be present and (2) there is reduced SERCA2a expression in HF which decreases SR Ca²⁺ uptake rate (30). Nevertheless, it is apparent that AP restoration can improve the defective Ca²⁺ signaling seen in HF and, importantly, reduce the LCS activity that can trigger EADs.

Discussion

Reactivation of I_Ca and I_Na to overcome the repolarization reserve and initiate EADs has been widely considered to be the primary mechanism for EAD generation (illustrated by the inner feedback loop shown in Fig. 5), e.g., refs. 11, 31, and 32. Indeed, when SR Ca²⁺ release was inhibited by blocking LTCCs, it was still possible to evoke EADs by a simulated I_Ca (33). Here we show that LCS-activated inward NCX current can initiate EADs and, we suggest, act as a coupled oscillator to further increase the risk for development of multiple EADs (outer loop in Fig. 5). This Ca²⁺ oscillator arises from two connected mechanisms: (1) Diffusion of Ca²⁺ from an initiating LCS may trigger additional LCS due to the gain inherent in Ca²⁺-induced Ca²⁺ release (CICR). This SR load-dependent process manifests as propagating Ca²⁺ ripples and, if enough LCS sites are available, more synchronous and larger Ca²⁺ waves. (2) LCS will increase inward NCX current to depolarize V_m (34) while V_m couples back onto LCS activity via the V_m dependence of LTCC gating (which may explain the V_m dependence of LCS frequency shown in Fig. 4E). The stochastic nature of LCS provides an explanation for the sudden appearance of arrhythmogenic EADs, and, once started, an EAD will promote additional LCS and EADs due to the increased SR Ca²⁺ load arising from I_Ca reactivation.

Fig. 5.

Flow diagram for two interacting positive-feedback mechanisms that can drive V_m and Ca²⁺ oscillations during EADs. The red inner cycle represents the well-established V_m oscillator, wherein an increase in dV_m/dt (caused by a relative increase in depolarizing currents compared to repolarizing currents, e.g., text in orange box at Lower Right) leads to voltage-dependent reactivation of I_Ca, which in turn causes further depolarization. The blue outer cycle represents the stochastic LCS-mediated Ca²⁺ oscillator mechanism for EAD initiation. Reduced early Ca²⁺ release and/or increased SR load increases LCS production, and the resulting increase in inward I_NCX tends to depolarize the membrane which then feeds onto I_Ca to trigger additional LCS. Green arrows indicate possible factors contributing to EAD initiation. β-AR, beta-adrenoreceptor.

The computer model clearly showed that both the electrical (V_m) and Ca²⁺ oscillators can couple and synergize (Fig. 4A). While the phase relationship between LCS-dependent NCX currents and dV_m/dt directly supports the idea that I_NCX can initiate Ca²⁺ dominant oscillations, I_Ca reactivation is also important (35) because late LTCC openings are a potent trigger for LCS (15). However, when Ca²⁺ levels are lower, and Ca²⁺ ripples and waves cannot form, EADs can still arise from instability in the repolarization process due to recruitment of noninactivated inward currents in the presence of insufficient repolarization reserve (11). In the latter case, smaller numbers of LCS may still play a lesser role in EAD initiation, and this is reminiscent of the role played by diastolic Ca²⁺ sparks in cardiac pacemaking by sino-atrial node cells (36).

Ca²⁺ Ripples, Ca²⁺ Waves, and V_m Oscillator Coupling.

Although Ca²⁺ waves (Fig. 2D) have been implicated in EAD genesis (37), it is notable that the fluctuations in Ca²⁺ shown in Fig. 2 A and E are not typical Ca²⁺ waves (as seen in Fig. 2D) (6, 7); instead, the Ca²⁺ fluctuations came from low-amplitude Ca²⁺ ripples. In such cases, the local propagation of Ca²⁺ release (short-range Ca²⁺ ripples) may not be able to initiate cell-wide Ca²⁺ waves because the effective amplification of Ca²⁺ release by CICR is insufficient for full Ca²⁺ wave support. This lack of sufficient amplification could be explained by the stochastic nature of LCS, coupled with local SR refractoriness (15, 38), decreasing the recruitment of adjacent LCS sites. Nevertheless, some synchronization in Ca²⁺ ripple initiation must occur for average Ca²⁺ to oscillate. The mathematical basis for the emergence of macroscopic Ca²⁺ oscillations from large numbers of independent LCS oscillators is beyond the scope of this study, but Kuramoto model analysis has shown that macroscopic oscillations can develop even when coupling across individual oscillators is weak (39). The relative importance of the Ca²⁺ and V_m oscillators in EAD generation will be variable because they depend on many factors such as the actual V_m trajectory, K⁺ current availability, the level of SR Ca²⁺ loading in the cell, and I_Ca availability, as well as RYR2 Ca²⁺ sensitivity, as illustrated in Fig. 5. In connection with this point, increased LTCC activity and SR load associated with β-adrenergic stimulation (10) would almost certainly increase the risk for LCS-stimulated EADs.

It should be noted that the LCS activity seen in our confocal line scans reflects only a small fraction of the actual number of LCS occurring in the entire cell; the confocal line scan surveys ∼2% of the cell volume, so ∼1 LCS/ms (Fig. 2E) would correspond to ∼50 LCS/ms cell-wide, and this much larger number underlies the measured NCX current that can initiate the EAD. This estimate is in reasonable agreement with the computer model used here which predicts that 107 LCS/ms may generate 0.26 pA/pF NCX current (29).

Since LTCC activity is common to both the V_m and Ca²⁺ oscillators, the idea that LTCC gating modification could be a therapeutic target for EAD prevention (33) becomes even more attractive. In addition, if modifying the late component of I_Ca is able to reduce net Ca²⁺ influx into the cell, SR Ca²⁺ content might be reduced (40), and this would also inhibit LCS activity (15). The improvement in Ca²⁺ signaling produced by applying a normal AP to HF myocytes is remarkable. This suggests that new therapies should be developed with the aim of improving early Ca²⁺ release by restoring phase 1 repolarization and/or restoring t-tubule regularity. This would reduce LCS frequency and thereby reduce the risk for potentially lethal LCS-triggered arrhythmias as well as mitigate the defective excitation–contraction coupling seen in HF (41).

Materials and Methods

More extensive details are available in SI Appendix. Briefly, all experiments were performed in accordance with the UK Home Office Animals (Scientific Procedures) Act 1986 and institutional approval by the University of Bristol ethics committee. We used an established coronary artery ligation model that leads to heart failure in adult New Zealand White rabbits (3–3.5 kg) which were daily monitored for health status. The target endpoint for HF was an ejection fraction of 40% (as measured by echocardiography SI Appendix, Fig. S2). Some rabbits did not quite reach this endpoint but presented other indicators of heart failure including dilated left ventricle and lung congestion (SI Appendix, Table S1). This model, due to repolarizing current behavior, can be used to gain insight into repolarization reserve with human relevance (42). It was not possible to wait for arrhythmias to start in this model (as they would be fatal), but EADs can be provoked in vitro by suitable interventions. Rather than use pharmacological manipulation to provoke EADs, we chose a pacing-pause method to minimize other possible perturbations of cellular function.

Cardiac Myocyte Isolation.

Left ventricular epicardial myocytes were obtained from rabbit hearts after full anesthesia (50 mg/kg sodium pentobarbital i.v.) and euthanasia. Enzymatic dissociation was carried out using 1 mg/mL collagenase I (Worthington), 0.05 mg/mL protease (type XIV Sigma), and 0.1 mmol/L Ca²⁺, as described previously (22). Guinea pig, rat, mouse, and zebrafish myocytes were isolated using similar methods to those described previously (22, 43, 44), and methods for zebrafish myocyte isolation are given in SI Appendix.

Electrophysiology.

Electrophysiology experiments were performed in a modified Tyrode’s solution (containing, in mmol/L: 133 NaCl, 5 KCl, 1 NaH₂PO₄, 10 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), 10 glucose, 1.8 CaCl₂, 1 MgCl₂, pH 7.4 with NaOH) at 36 ± 1 °C. Patch pipettes were pulled from borosilicate glass using a P80 micropipette puller (Sutter Instruments). Pipettes were filled with an intracellular solution containing, in mmol/L: 120 aspartic acid, 20 KCl, 10 HEPES, 10 NaCl, 5 glucose, 5 Mg.ATP, 0.05 Fluo-4 pentapotassium salt, with KOH added to adjust to pH 7.2. Tip resistance was typically 1.6–2.0 MΩ when filled with this solution. Membrane potential and currents were recorded using an Axopatch 1D amplifier (Molecular Devices), Power1401 digitizer (Cambridge Electronic Design), and Signal data acquisition software (version 6.04, Cambridge Electronic Design). Cell membrane capacitance was measured by step depolarizations to −75 mV from a holding potential of −80 mV for 25 ms. Series resistance was compensated by ∼70%. Liquid junction potential (10 mV) was subtracted from recordings.

Confocal Imaging.

Ca²⁺ sparks and transients were recorded in line scan mode (45) from the fluo-4 loaded cells using an inverted confocal microscope (LSM 880, Zeiss) with a 1.4 NA 63× oil immersion lens. Excitation light was provided by a 488-nm argon laser, and fluorescence emission was collected at 492–600 nm. Ca²⁺ line scans were recorded with the pinhole set to <2 Airy units, at a pixel size of 0.1–0.2 μm/pixel, and with a scan speed of 1 ms per line. GaAsP photodetectors were used to increase the sensitivity of Ca²⁺ spark detection. The t-tubule system was imaged by labeling the sarcolemma with di-8-ANEPPS from a stock 1 mmol/L solution (in anhydrous dimethyl sulfoxide) added directly to the cell-recording chamber (final concentration 1 μmol/L) for 2–3 min. Excitation was at 488 nm, and emission was collected at >600 nm.

Statistical Analysis.

Statistical analyses were performed at the level of the cell, and statistics on replicates of n individual independent cell experiments from N animals are given in the text as n/N. Data were tested for normality using the Shapiro-Wilk test (Prism7, Graphpad); in any cases where data were skewed, the test was reapplied to log-transformed data. Paired or unpaired t tests were performed on normally and log-normally distributed data. Results are presented as mean ± SEM. The limit of statistical confidence was P < 0.05.

Data Availability Statement.

All data and computer codes are available upon request from the authors.