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The balance of Ca2+ influx and efflux regulates the Ca2+ load of cardiac myocytes, a process known as autoregulation. Previous work has shown that Ca2+ influx, via L-type Ca2+ current (ICa), and efflux, via the Na+/Ca2+ exchanger (NCX), occur predominantly at t-tubules; however, the role of t-tubules in autoregulation is unknown. Therefore, we investigated the sarcolemmal distribution of ICa and NCX current (INCX), and autoregulation, in mouse ventricular myocytes using whole cell voltage-clamp and simultaneous Ca2+ measurements in intact and detubulated (DT) cells. In contrast to the rat, INCX was located predominantly at the surface membrane, and the hysteresis between INCX and Ca2+ observed in intact myocytes was preserved after detubulation. Immunostaining showed both NCX and ryanodine receptors (RyRs) at the t-tubules and surface membrane, consistent with colocalization of NCX and RyRs at both sites. Unlike INCX, ICa was found predominantly in the t-tubules. Recovery of the Ca2+ transient amplitude to steady state (autoregulation) after application of 200 µM or 10 mM caffeine was slower in DT cells than in intact cells. However, during application of 200 µM caffeine to increase sarcoplasmic reticulum (SR) Ca2+ release, DT and intact cells recovered at the same rate. It appears likely that this asymmetric response to changes in SR Ca2+ release is a consequence of the distribution of ICa, which is reduced in DT cells and is required to refill the SR after depletion, and NCX, which is little affected by detubulation, remaining available to remove Ca2+ when SR Ca2+ release is increased.
NEW & NOTEWORTHY This study shows that in contrast to the rat, mouse ventricular Na+/Ca2+ exchange current density is lower in the t-tubules than in the surface sarcolemma and Ca2+ current is predominantly located in the t-tubules. As a consequence, the t-tubules play a role in recovery (autoregulation) from reduced, but not increased, sarcoplasmic reticulum Ca2+ release.
t-tubules are invaginations of the surface membrane of cardiac ventricular myocytes that play a central role in excitation-contraction coupling. Contraction is initiated by Ca2+ influx [Ca2+ current (ICa)] through L-type Ca2+ channels (LTCCs); this activates ryanodine receptors (RyRs) in the adjacent sarcoplasmic reticulum (SR) membrane to cause Ca2+ release from the SR [Ca2+-induced Ca2+ release (CICR)]. ICa, RyRs, and thus CICR occur predominantly at t-tubules (6, 8, 14), which results in a near-synchronous rise in cytosolic Ca2+ throughout the cell to levels sufficient to activate the contractile proteins.
For myocytes to relax, Ca2+ must be removed from the cytosol. This is achieved by Ca2+ reuptake into the SR and Ca2+ efflux from the cell. Although SR Ca2+ uptake is the main route of Ca2+ removal from the cytosol, sarcolemmal Ca2+ efflux pathways, the Na+/Ca2+ exchanger (NCX) and sarcolemmal Ca2+ ATPase, also play an important role (1, 25). Evidence largely from rat cardiac myocytes suggests that, like influx, Ca2+ efflux also occurs predominantly at the t-tubules (11, 14, 37) where, it has been proposed, NCX has privileged access to Ca2+ released from the SR (2, 20, 41).
The balance between sarcolemmal Ca2+ influx and efflux determines the Ca2+ load of the cell and thus the amplitude of the Ca2+ transient and is maintained by a process called “autoregulation,” which involves regulation of both Ca2+ influx and efflux by cytoplasmic Ca2+. For example, sensitizing CICR has only a short-lived effect on the Ca2+ transient amplitude (17, 39, 42) because the resulting increase in SR Ca2+ release decreases ICa by Ca2+-dependent inactivation of ICa and increases Ca2+ efflux by stimulating NCX (17, 36). These changes reduce the Ca2+ transient amplitude back to baseline levels with an accompanying decrease in SR Ca2+ content (17, 39, 42). The role of t-tubules in autoregulation is unknown; however, because ICa and its inactivation by Ca2+ as well as NCX current (INCX) and its stimulation by Ca2+ released from the SR have been reported to occur predominantly at the t-tubules (6, 14, 30), it seems likely that they play an important role in autoregulation. Therefore, this study was designed to determine the sarcolemmal distribution of ICa and INCX and the consequences for the role of the t-tubules in autoregulation in mice.
Ventricular myocytes were isolated from the hearts of male C57BL/6 mice aged between 11 and 13 wk. All procedures were performed in accordance with United Kingdom legislation and approved by the University of Bristol Ethics Committee. Mice were injected with heparin (500 IU by intraperitoneal injection) and killed by cervical dislocation. The heart was excised and washed in isolation solution supplemented with 0.1 mM CaCl2 and 10 U/ml heparin. The heart was then Langendorff perfused with isolation solution for 4 min followed by enzyme solution (isolation solution plus 0.1 mM CaCl2, 265 U/ml collagenase, and 0.3 U/ml protease) for ~15 min. The ventricles were then removed and shaken in enzyme solution for 4–6 min before being filtered and centrifuged. Cells were resuspended in isolation solution (pH 7.4) plus 0.1 mM CaCl2 and stored for 2–8 h before use on the day of isolation. Detubulation (DT), the physical and functional uncoupling of the t-tubules from the surface membrane, was achieved using formamide-induced osmotic shock, as previously described, by incubating cells with 1.5 M formamide for 2 min before centrifugation and resuspending the cells in Tyrode solution (5). Data from intact and DT myocytes were obtained from separate groups of cells.
All reagents were obtained from Sigma-Aldrich (Poole, UK) unless otherwise specified. The isolation solution contained (in mM) 130 NaCl, 5.4 KCl, 0.4 NaH2PO4, 4.2 HEPES, 10 glucose, 1.4 MgCl2, 20 taurine, and 10 creatine (pH 7.6 using NaOH). Tyrode solution used for experiments contained (in mM) 133 NaCl, 1 MgSO4, 1 CaCl2, 1 Na2HPO4, 10 d-glucose, and 10 HEPES (pH 7.4 using NaOH) plus 5 CsCl to inhibit K+ currents. The pipette solution contained (in mM) 100 CsCl, 20 TEACl, 10 NaCl, 0.5 MgCl2, 5 MgATP, 10 HEPES, 0.4 GTP-Tris (pH 7.2 using CsOH), and 0.1 pentapotassium salt of the fluorescent Ca2+ indicator fluo-4 (Life Technologies, Paisley, UK).
Myocytes were placed in a chamber mounted on a Diaphot inverted microscope (Nikon UK, Kingston-upon-Thames, UK). Membrane currents and cell capacitance were recorded using the whole cell patch-clamp technique using an Axopatch 200B patch-clamp amplifier, a Digidata 1440A analog-to-digital converter, and pClamp 10 software (Molecular Devices, Reading, UK), which was also used for data acquisition (at 2 kHz) and analysis. Pipette tip resistances were typically 1.2–2.0 MΩ when filled with pipette solution. All experiments were performed at room temperature.
To monitor Ca2+ influx and efflux, and thus Ca2+ balance, during a Ca2+ transient, holding potential was set to −80 mV; a 500-ms ramp to −40 mV was used to inactivate Na+ current followed by step depolarization to 0 mV for 100 ms to activate ICa at a frequency of 1 Hz. ICa was measured as the difference between peak inward current and current at the end of the pulse to 0 mV, and the integral was taken as a measure of Ca2+ influx. Inactivation of ICa was quantified by measuring the time to 50% inactivation (T50%). The current representing Ca2+ removed by NCX after the step depolarization (INCX,tail) was measured by fitting a single-exponential function to 350 ms of the current trace starting 20 ms after repolarization from 0 to −80 mV and extrapolating back to when the membrane was repolarized. The integral of the exponential was taken as a measure of Ca2+ efflux during the Ca2+ transient (13, 15). This analysis was performed using MATLAB R2015a (Mathworks, Natick, MA).
To determine the distribution of INCX between the surface and t-tubule membranes, INCX was measured in intact and DT myocytes during the application of 10 mM caffeine to cause spatially and temporally uniform release of SR Ca2+ (4); the resulting inward current due to Ca2+ extrusion via NCX was recorded at −80 mV, and INCX was taken as the difference between the peak current and the current after caffeine washout.
The distribution of ICa, INCX, and membrane capacitance (a function of membrane area), and, thus, current density, between the surface and t-tubular membranes was calculated from measurements in intact (whole cell) and DT (surface membrane only) myocytes, as previously described (7, 8). In brief, the currents and capacitance of the surface sarcolemma were calculated from those measured in DT myocytes, corrected for incomplete DT assessed using confocal imaging of Di-8-ANEPPS-stained cells as 12.7%; t-tubular currents and capacitance were calculated from the difference between those in intact cells and those in the surface sarcolemma.
Fluo-4 fluorescence was excited at 450–488 nm, and emitted fluorescence was collected at wavelengths <560 nm. Fluorescence was normalized to fluorescence just before application of caffeine (F/F0). The rate of decay of Ca2+ transients was obtained by fitting single exponential functions to the declining phase of the ICa- and caffeine-induced Ca2+ transients.
NCX hysteresis loops were produced by plotting INCX against F/F0 during application of 10 mM caffeine to release SR Ca2+, as previously described (14, 41). Loops were quantified by calculating the area within the loop and normalizing to the rectangle defined by maximum and minimum INCX and F/F0 (41).
Cells were fixed with 4% paraformaldehyde for 10 min before being permeabilized with 0.1% Triton X-100 and stained with primary antibodies for RyR (MA3-916, Thermo Fisher) or NCX (R3F1, Swant) overnight. Cells were then incubated in Alexa fluor 488-conjugated anti-mouse secondary antibody before being mounted with ProLong Gold. Cells were imaged on an LSM 880 confocal microscope (Zeiss) in Airyscan “super-resolution” mode, with a 1.4 numerical aperture ×63 oil-immersion objective.
Staining at the cell surface and in the center of the cell was determined from a binary cell image obtained using Otsu’s method (27). The perimeter of the cell was outlined manually, and staining within a band that extended 2 µm inside this outline was taken as the cell edge; staining from the image inside this band, excluding the nuclei, was taken as the cell center.
Staining was quantified as follows:
where %bright pixels is the percentage of bright pixels in a given area relative to the total number of bright pixels in the cell and %total pixels is the percentage of pixels in a given area relative to the total number of pixels in the cell.
Data are expressed as means ± SE. Errors of derived variables (e.g., t-tubule ICa and INCX densities) and the subsequent statistical analysis were calculated using propagation of errors from the constituent measurements (8). Student’s t-tests and two-way ANOVA with the Bonferroni post hoc test were used as appropriate and performed using GraphPad Prism 7 (GraphPad Software, San Diego, CA). The limit of statistical confidence was P < 0.05. All statistical tests were performed on the number of cells. Sample sizes (n numbers) are given as c/h, where c is the number of cells used from h number of hearts.
The mathematical model of autoregulation described by Eisner et al. (12) was used to simulate the data obtained in intact and DT myocytes. Baseline values for intact cells were those used by Eisner et al. (12) except that the transsarcolemmal Ca2+ efflux fraction (r) was decreased from 10% to 8% (the value measured in the current experiments). The relative changes obtained experimentally in DT cells were used to model the data in these cells: ICa was decreased from 10 to 4.1, fractional SR Ca2+ release (f) was decreased from 0.7 to 0.287, and r was maintained at 8%, the value measured in DT cells. Changes in INCX relative to steady state obtained from the experimental data were incorporated for intact and DT myocytes.
DT significantly decreased cell capacitance from 181 ± 9 (n = 28/13) to 128 ± 6 pF (n = 24/11, P < 0.0001), with no change in cell volume, calculated as previously described previously (3) [54 ± 4 pl (intact) vs. 58 ± 4 pl (DT)]. After correction for incomplete DT, this suggests that 41% of the cell membrane is t-tubular.
INCX was measured in intact and DT myocytes during application of 10 mM caffeine to determine its distribution. Figure 1 shows representative records of intracellular Ca2+, monitored as fluo-4 fluorescence, and the associated INCX in intact (Fig. 1A) and DT (Fig. 1B) myocytes during application of caffeine. DT had no significant effect on either the amplitude or rate of decay (kCaff) of the caffeine-induced Ca2+ transient (Fig. 1, C and D), suggesting that SR Ca2+ content and sarcolemmal Ca2+ efflux are unchanged by DT of mouse cells. Figure 1E shows whole cell INCX density and the calculated density of INCX at the surface and t-tubular membranes, showing that the density of INCX in the t-tubules is about half that in the surface membrane (P < 0.01). Thus, ~25% of total INCX appears to occur in the t-tubules, consistent with the small, although statistically nonsignificant, decrease in mean kCaff on DT (Fig. 1D).
Previous work in rat ventricular myocytes has shown that when caffeine is applied to release Ca2+ from the SR, INCX is greater for a given cytoplasmic Ca2+ concentration as Ca2+ increases than during the subsequent decrease. It has been suggested that this hysteresis arises because Ca2+ released from the SR has “privileged” access to NCX due to the proximity of NCX to RyRs. Thus, during Ca2+ release, NCX is responding to a higher local Ca2+ concentration than that reported by a Ca2+ indicator in the cytoplasm, while during the falling phase, the local Ca2+ concentration surrounding NCX is closer to bulk cytoplasmic Ca2+ concentration (41). Hysteresis is lost after DT in rat ventricular myocytes, consistent with the hysteresis arising at the t-tubules as a result of localization of INCX to the t-tubules, close to the site of Ca2+ release, in these cells (14, 41). Because the majority of INCX appears to be located at the surface sarcolemma in mouse ventricular myocytes, we investigated whether the hysteresis and its response to DT were different in these cells by plotting INCX against F/F0. Figure 2A shows that intact cells showed a marked hysteresis that was not abolished by DT: INCX density for a given Ca2+ was greater during the rising phase than the declining phase of the caffeine-induced transient in both cell types, with no significant difference in the area ratio of the loop (see Fig. 2B and materials and methods). However, the loop was shifted in DT cells, with a greater INCX density for a given Ca2+, consistent with a similar Ca2+ release but greater INCX density at the cell surface. These data are consistent with the majority of NCX being located at the surface sarcolemma in the mouse and the location of INCX determining the hysteresis, which arises at the site of highest INCX density: the t-tubules in the rat and the surface sarcolemma in the mouse.
Immunohistochemistry was used to investigate the distribution of NCX and RyR. Figure 2C shows confocal images of a representative mouse myocyte stained for NCX, showing marked striations within the cell and continuous staining at the cell surface, consistent with NCX being present at the t-tubular and surface membranes and thus with the measured distribution of INCX. Figure 2D shows a representative cell stained for RyRs, which showed marked striations, consistent with t-tubular localization, with less staining at the cell surface, although two or sometimes three distinct areas of RyRs were observed coinciding with the mouth of t-tubules (Fig. 2D,ii). Staining of NCX and RyRs at the cell edge and cell center, quantified using Eq. 1, are shown in Fig. 2E; the apparent density of NCX staining was higher than that of RyR at the cell surface (P < 0.0001, Bonferroni post hoc test) but similar in the cell center, although both proteins had a greater density at the cell edge than at the cell center (P < 0.0001, two-way ANOVA). However, although the images were obtained using Airyscan “super-resolution” to minimize the contribution of out of focus light, there may be more surface membrane than t-tubular membrane in the optical field, since the former is likely to be present in the full depth of the field. This could lead to a higher apparent NCX density at the cell surface, although its effect on measured RyR density is unclear. However, although quantification is difficult, these data show that Ca2+ release sites and NCX are present at the cell edge, consistent with the observed distribution of INCX and the hysteresis observed in DT myocytes.
Since INCX distribution appears to be different in mouse and rat myocytes, we also investigated the distribution of ICa in these cells. Figure 3 shows representative ICa and the elicited Ca2+ transients in intact (Fig. 3A) and DT (Fig. 3B) myocytes. Consistent with previous reports in rat myocytes, the Ca2+ transient amplitude (Fig. 3C) and rate of decay (Fig. 3D) were significantly decreased by DT (5, 6, 21).
ICa in intact and DT cells was used to calculate the distribution of ICa between the surface sarcolemma and t-tubule membranes. Figure 3E shows that ICa density was about four times greater in the t-tubule membrane compared with the surface sarcolemma (P < 0.0001), as in the rat (6, 8, 14). However, in contrast to the rat, there was no significant change in the rate of inactivation of ICa after DT (Fig. 3F), suggesting that inactivation was similar at the surface and t-tubular membranes.
Since the distribution of Ca2+-handling proteins determines the role of the t-tubules in Ca2+ handling (4, 6, 14, 38), we investigated the effect of detubulation on the recovery of systolic Ca2+ transient amplitude and Ca2+ flux via ICa and INCX after depletion of SR Ca2+ by a high concentration (10 mM) of caffeine, and during and after application of a low concentration (200 µM) of caffeine to sensitize CICR (26, 42), both of which elicit autoregulation.
Figure 4 shows that after application of 10 mM caffeine, Ca2+ transient amplitude was initially small and gradually recovered to steady state with successive beats in both intact (Fig. 4A,i) and DT (Fig. 4B,i) myocytes. Recovery was accompanied by a decrease in the amplitude and integral of ICa, and an increase in the amplitude and integral of INCX in both intact (Fig. 4A,ii) and DT (Fig. 4B,ii) myocytes. However, steady-state Ca2+ transient amplitude was significantly smaller in DT cells (P < 0.0001), consistent with reduced ICa and loss of t-tubules, and the half-time (t1/2) to reach steady state was significantly longer (8.7 ± 1.0 s for intact cells vs. 12.6 ± 0.4 s for DT cells, P < 0.01); thus, the rate of recovery due to SR refilling was slower in DT myocytes (Fig. 4C). Figure 4D shows that recovery of Ca2+ transient amplitude was accompanied by a small reduction in Ca2+ influx via ICa in both intact and DT cells, although Ca2+ influx, and thus the rate of Ca2+ accumulation, was significantly smaller in DT myocytes (Fig. 4, D and E). In contrast, Ca2+ efflux via INCX gradually increased with continued stimulation in both cell types (Fig. 4F), reflecting an increase in SR Ca2+ content and release, although Ca2+ efflux was significantly smaller in DT cells (Fig. 4, F and G), which is likely to reflect the decrease in Ca2+ transient amplitude due to reduced ICa rather than loss of NCX (above).
The ratio between Ca2+ influx and efflux was calculated to compare Ca2+ balance in intact and DT cells (Fig. 4H). In both cell types, the ratio was initially greater than 1 after caffeine, reflecting net Ca2+ influx, and gradually decreased toward 1, which represents the steady-state balance of influx and efflux. However, the initial ratio was slightly lower in DT than intact myocytes. These data suggest that reduced Ca2+ influx via ICa, and therefore slower Ca2+ accumulation, underlies the slower recovery of Ca2+ transient amplitude in DT cells and thus that the t-tubules play an important role in autoregulation.
To test this idea further, Ca2+ autoregulation was investigated during application and washout of 200 µM caffeine to intact (Fig. 5A) and DT (Fig. 5B) myocytes. Application of caffeine to intact myocytes caused a transient increase in Ca2+ transient amplitude, which recovered to steady state, while washout of caffeine caused a transient decrease in Ca2+ transient amplitude, which also recovered to steady state, consistent with previous work (e.g., Ref. 39). DT cells showed a similar response, although the Ca2+ transient amplitude was smaller; mean data are shown in Fig. 5C. Interestingly, the half-time to recover to steady state during application of caffeine was not changed by DT (5.7 ± 0.7 s in intact cells vs. 5.3 ± 0.6 s in DT cells), but the recovery to steady state on washout of caffeine was significantly slowed in DT cells (t1/2: 3.1 ± 0.3 s in intact cells vs. 5.4 ± 0.4 s in DT cells, P < 0.001), similar to the recovery after 10 mM caffeine.
Application of 200 µM caffeine caused similar changes in the rate of inactivation of ICa in intact and DT myocytes (Fig. 5D) and decreased Ca2+ influx in both cell types, whereas washout caused only a small increase in both cell types (Fig. 5E). Ca2+ efflux increased on application of 200 µM caffeine, while on washout efflux was reduced (Fig. 5F). Although both influx and efflux were significantly (P < 0.001) reduced in DT cells, the overall balance was not significantly different (Fig. 5G), whether during application 200 µM caffeine, where the balance favored Ca2+ efflux, or whether during washout of caffeine, where the balance favored net Ca2+ influx.
Taken together, these data suggest that t-tubules play an important role in recovery from a decrease in SR Ca2+ load, since DT cells recover more slowly, but not in recovery from an increased SR Ca2+ release, since intact and DT cells recover at similar rates. To test this idea further, we incorporated the data from intact and DT myocytes into the model of Eisner et al. (12), as described in materials and methods. The results of these simulations are shown in Fig. 6, which shows that after SR Ca2+ depletion (Fig. 6A), DT cells recovered much more slowly than intact cells (t1/2: 10.7 stimuli in intact cells vs. 26.8 stimuli in DT cells), as observed experimentally (cf. Fig. 4C). In contrast, the rate of recovery during sensitization of CICR (Fig. 6B) was similar in intact and DT myocytes (t1/2: 8.8 stimuli in intact cells vs. 12.8 stimuli in DT cells), consistent with the data from the experiments (cf. Fig. 5C).
The present study was designed to investigate the role of the t-tubules in Ca2+ autoregulation in mouse ventricular myocytes. The data show that the majority of ICa occurs in the t-tubules, whereas, in contrast to rat myocytes, INCX occurs mainly at the surface sarcolemma and the hysteresis between Ca2+ and INCX persists after DT. Although autoregulation to steady state occurred after DT, the time course of recovery was slower during recovery from decreased SR Ca2+ release but, interestingly, not from increased SR Ca2+ release. This suggests that t-tubules play a role in recovery from decreased, but not increased, SR Ca2+ release, which may reflect the localization of ICa and INCX.
Previous work has shown that ICa and INCX occur predominantly in the t-tubules in rat ventricular myocytes (6, 8, 11, 14, 21, 37, 44). The present study demonstrated a slightly higher t-tubular membrane fraction (41%) than that previously reported for rat myocytes (6, 8, 30), consistent with the higher t-tubule density reported previously for the mouse (18, 29); it also showed that ICa is located predominantly in the t-tubules of mouse myocytes, consistent with a previous report using DT in mouse myocytes (19). The presence of ICa at the t-tubules in both species, in close proximity to the majority of RyRs, allows the tight coupling between Ca2+ influx and SR Ca2+ release that underlies CICR, while the higher t-tubule density in the mouse may reflect its high heart rate and the consequent need for rapid activation.
However, the present work also demonstrated that, in contrast to rat ventricular myocytes, INCX is located predominantly at the cell surface in mouse myocytes. Although this might explain some of the discrepancies in the literature regarding the location of NCX determined using immunological techniques (35, 37), it is unclear why the distribution of INCX should be different in the two species, which have similar Ca2+-handling properties, with similar action potential configurations, fractions of SR and trans-sarcolemmal Ca2+ flux (1, 10, 22, 25), and kinetics of contraction and relaxation (23), although computer modeling suggests that NCX distribution has relatively little effect on whole cell Ca2+ handling (31). One possibility is that location of NCX at the cell surface is energetically favorable, which might be important at the higher heart rates in the mouse, because it will avoid the futile Ca2+ cycling that results from NCX being in close proximity to the main site of SR Ca2+ release, by decreasing the amount of released Ca2+ that is immediately removed via NCX, enabling more of the released Ca2+ to activate the contractile proteins. The small t-tubular INCX is, however, associated with a small (nonsignificant) decrease in the rate of decay of the caffeine-induced Ca2+ transient after DT in the mouse, in contrast to the marked decrease observed in the rat (14).
This species difference in the sarcolemmal distribution of Ca2+-handling protein function raises questions about the distribution in other species. Although, as far as we are aware, corresponding data are not available for large mammals, previous work in cultured guinea pig myocytes (28) showed no significant change in INCX density with time in culture, whereas cell capacitance decreased (correlating with loss of t-tubules), suggesting a similar INCX density in the surface sarcolemma and t-tubules of guinea pig myocytes (28). Nevertheless, the maintained INCX density might have been due to other mechanisms upregulating INCX in culture.
It has been suggested that the hysteresis between INCX and bulk cytoplasmic Ca2+ is due to the proximity of NCX to the site of Ca2+ release, so that the exchanger responds to a higher Ca2+ concentration than monitored in the bulk cytosol during Ca2+ release. The hysteresis is abolished after DT of rat myocytes (14), suggesting that this occurs at the t-tubules, where the majority of NCX is located close to the site of SR Ca2+ release, and that surface sarcolemmal INCX responds mainly to changes in global Ca2+ concentration. The present observation that the hysteresis is present in both intact and DT mouse myocytes is consistent with the observed distribution of INCX in these cells and suggests that INCX at the surface sarcolemma is located close to sites of Ca2+ release and responding to local changes of Ca2+ concentration during Ca2+ release (and to changes in global Ca2+ concentration during the descending phase). The staining data support this idea: RyRs were observed not only in striations in the cell interior, consistent with localization at the t-tubules, but also at the surface membrane in clusters at the mouth of t-tubules, close to NCX, which, in agreement with the INCX measurements, was also observed at both the t-tubules and cell surface. These data suggest that RyRs, and thus Ca2+ release, occur at the t-tubules and cell surface and thus that it is the distribution of INCX that determines the site of the hysteresis in these cells. These data also suggest that the site of origin of arrhythmogenic delayed afterdepolarizations, which are caused by activation of INCX by SR Ca2+ release, will depend on the location of INCX and Ca2+ release and may thus be different in different species.
The present data show that, in agreement with previous studies in other species (17, 39), recovery of Ca2+ transient amplitude following altered SR Ca2+ release is associated with reciprocal changes in Ca2+ influx via ICa and Ca2+ efflux via NCX.
Changes in Ca2+ transient amplitude result in changes in Ca2+ influx through Ca2+-dependent inactivation of ICa. In rabbit cells, Ca2+ influx can increase by 50–60% in the absence of SR Ca2+ release (16, 33); in the rat, Ca2+ influx can double after SR Ca2+ depletion (40). Although Ca2+-dependent inactivation also occurs in mouse myocytes, as seen by the faster inactivation during application of 200 µM caffeine and slower inactivation on wash off in the present study, only small differences in the rate of inactivation between intact and DT myocytes were observed. This suggests, in contrast to rat myocytes in which Ca2+-dependent inactivation occurs predominantly at the t-tubules (6), that inactivation is similar at the surface and t-tubular membranes. The reason for this species difference is unknown but may reflect differences in local Ca2+ and/or local LTCC regulation at the two sites. The lack of change in the rate of inactivation of ICa after DT, despite the smaller Ca2+ transient, could be explained by the absence of basal Ca2+-dependent inactivation of ICa or by rapid inactivation, which is manifested as a change in peak ICa rather than its duration. However, studies using intracellular Ca2+ chelators or barium as the charge carrier for ICa to inhibit Ca2+-induced inactivation have suggested that there is substantial basal ICa inactivation, which mainly affects the rate of inactivation rather than peak ICa (32, 34, 43), making these explanations unlikely. Thus, the most likely cause for ICa inactivation being unaffected by DT appears to be that local Ca2+ release, and thus inactivation, at the cell surface is similar to that at the t-tubules. In this case, the smaller, slower Ca2+ transient in DT myocytes is due to loss of the quantitatively more important t-tubular CICR and loss of synchronization of Ca2+ release. A similar local Ca2+ release at the cell surface and t-tubules can explain why the hysteresis occurs in DT myocytes given the observed distribution of INCX. Thus, it appears that, in contrast to the rat, both INCX density and Ca2+-dependent inactivation of ICa are similar at the t-tubular and surface membranes and that the decrease in Ca2+ influx after DT is predominantly due to loss of t-tubular ICa rather than an altered rate of inactivation.
In contrast to ICa, regulation of INCX appears similar to that previously reported, with an increase in INCX associated with an increase in Ca2+ transient amplitude in intact and DT myocytes. DT resulted in changes in kCaff and INCX consistent with loss of the t-tubular fraction of INCX (25%), while the hysteresis between INCX and cytosolic Ca2+ showed a larger current for a given Ca2+ in DT myocytes, with no loss of hysteresis, consistent with the greater density of INCX in the surface membrane and similar Ca2+ release at the surface and t-tubular membranes. Thus, the observed changes are consistent with the observed distribution of INCX, with no evidence for altered regulation.
As in previous studies (4, 14), DT had no effect on SR Ca2+ content, as assessed by releasing SR Ca2+ using 10 mM caffeine, but did result in a smaller and slower voltage-stimulated Ca2+ transient, due to decreased ICa and thus CICR and loss of synchronization of SR Ca2+ release. The smaller Ca2+ transient, in turn, decreased INCX, even though it was present at the surface sarcolemma.
Although DT had no effect on the rate of recovery of Ca2+ transient amplitude to steady state when SR Ca2+ release was increased using low-dose caffeine, recovery was slower from a decreased SR Ca2+ content after caffeine-induced SR Ca2+ depletion or on washout of 200 µM caffeine. This asymmetric response to changes in SR Ca2+ release in DT cells was not due to the nonlinear response of fluo-4 to Ca2+ because it was still present after the conversion of fluorescence to Ca2+ (not shown), an idea supported by the model. It may, however, be explained by the differential distribution of ICa and INCX. In this case, when an increase in SR Ca2+ release occurs, Ca2+ efflux via NCX is the main mechanism to decrease Ca2+ transient amplitude and return the cell to steady state; thus, in DT mouse cells, in which the majority of INCX occurs at the surface, recovery is similar in intact and DT myocytes and is not affected by the loss of ICa. However, when SR Ca2+ release is decreased, Ca2+ influx via ICa is necessary to refill the SR and increase Ca2+ transient amplitude to steady state and is thus essential for recovery. Since steady-state SR Ca2+ content is similar in intact and DT myocytes but Ca2+ influx via ICa is smaller in DT cells, more stimuli are required to refill the SR to steady state, so that recovery is slower in DT than in intact cells, even though the majority of NCX is present and quantitatively shows greater changes over time than ICa. Slowing of recovery from caffeine has also been reported after DT of rat myocytes (4), consistent with a key role for ICa, which, like INCX, occurs predominantly in the t-tubules in this species.
Although t-tubules uncoupled from the surface membrane by DT may reseal within the cell and, in principle, sequester and release Ca2+ (5, 24), it seems unlikely that they can explain the current data, since they are electrically uncoupled from the surface membrane. Therefore, Ca2+ channels in such resealed tubules will not undergo activation (and thus inactivation), an idea supported by the lack of Ca2+ release in the center of DT myocytes (5). Because they do not release Ca2+ and have a small volume, they are also unlikely to take up Ca, and previous work inhibiting surface NCX in DT rat myocytes during application of caffeine showed that only ~2% of Ca2+ removal occurs into non-SR sinks within the cell (9), which includes mitochondria, and this is likely to be even less in mouse myocytes, which have a smaller fraction of NCX within the t-tubules. Thus, resealed t-tubules are unlikely to play an important role in normal Ca2+ cycling or autoregulation, an idea supported by the effects of DT on the Ca2+ transient and autoregulation.
Thus, it appears that the t-tubules play an important role in autoregulation in mouse myocytes during recovery from decreased SR Ca2+ release, because of the high t-tubular ICa density, but play little role in the recovery from increased Ca2+ release. These data also suggest that the response of a particular species to such changes will depend on the distribution of ICa and INCX between the t-tubules and surface sarcolemma. Although the distribution of Ca2+ fluxes in human myocytes is unknown, the current work shows that the distribution of ICa and INCX determines cell function and enhances our understanding of how this occurs. Thus, when human data become available, it should be possible to better predict the consequent changes in cell function in both physiological conditions and after t-tubule disruption and changes in the distribution of ICa and INCX, which have been reported in heart failure (14) and which may also affect autoregulation.
This work was supported by British Heart Foundation Grants PG/14/65/31055 and RG/12/10/29802.
No conflicts of interest, financial or otherwise, are declared by the authors.
H.C.G. performed experiments; H.C.G. and C.H.K. analyzed data; H.C.G., S.M.B., A.F.J., and C.H.O. interpreted results of experiments; H.C.G. prepared figures; H.C.G. and C.H.O. drafted manuscript; H.C.G., C.H.K., S.M.B., A.F.J., and C.H.O. edited and revised manuscript; A.F.J. and C.H.O. conceived and designed researc h; A.F.J. and C.H.O. approved final version of manuscript.