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(Circulation Research. 2004;95:e1.)
© 2004 American Heart Association, Inc.

UltraRapid Communication

Differential Modulation of L-type Ca²⁺ Current by SR Ca²⁺ Release at the T-Tubules and Surface Membrane of Rat Ventricular Myocytes

Fabien Brette^*, Laurent Sallé^*, Clive H. Orchard

From the School of Biomedical Sciences (F.B., C.H.O.), University of Leeds, Leeds, UK; and the Laboratoire de Physiologie Cellulaire (L.S.), Université de Caen, Caen, France.

Correspondence to Dr Fabien Brette, School of Biomedical Sciences, University of Leeds, Leeds LS2 9JT, UK. E-mail f.brette{at}leeds.ac.uk

	Abstract

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We have characterized modulation of I_Ca by Ca²⁺ at the t-tubules (ie, in control cells) and surface sarcolemma (ie, in detubulated cells) of cardiac ventricular myocytes, using the whole-cell patch clamp technique to record I_Ca. I_Ca inactivation was significantly slower in detubulated cells than in control cells (27.1±7.8 ms, n=22, versus 16.4±7.9 ms, n=22; P<0.05). In atrial myocytes, which lack t-tubules, I_Ca inactivation was not changed by the treatment used to produce detubulation. In the presence of ryanodine or BAPTA, or when Ba²⁺ was used as the charge carrier, the rate of inactivation was not significantly different in control and detubulated cells. Frequency-dependent facilitation occurred in control cells but not in detubulated cells, and was abolished by ryanodine. These results suggest that Ca²⁺ released from the SR has a greater effect on I_Ca in the t-tubules than at the surface sarcolemma. This does not appear to be due to differences in local Ca²⁺ release from the SR, because the gain of Ca²⁺ release was not significantly different in control and detubulated cells. These data suggest that the t-tubules are a key site for the regulation of transsarcolemmal Ca²⁺ flux by Ca²⁺ release from the SR; this could play a role in altered Ca²⁺ homeostasis in pathological conditions. The full text of this article is available online at http://circres.ahajournals.org.

Key Words: transverse tubules • calcium channel • inactivation • facilitation

	Introduction

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Calcium influx via the L-type Ca²⁺ current (I_Ca) is the major trigger for Ca²⁺ release from the sarcoplasmic reticulum (SR) of cardiac ventricular myocytes.¹ This influx activates a cluster of adjacent SR Ca²⁺ release channels (ryanodine receptors; RyRs); the consequent systolic Ca²⁺ transient is the spatial and temporal sum of such local Ca²⁺ releases.² The transverse (t-) tubules of ventricular myocytes play an important role in this process. These tubules are invaginations of the sarcolemma that occur at the Z-line, perpendicular to the longitudinal axis of the cell (see review).³ Functional and immunohistochemical data suggest that I_Ca occurs predominantly in the t-tubules, adjacent to RyRs, which are also located predominantly at the t-tubules.^4–6 Thus, it appears that the t-tubules are the major site of Ca²⁺ entry and hence Ca²⁺ release in cardiac ventricular myocytes. Conversely, Ca²⁺ released by the SR can modulate I_Ca; this plays an important role in cellular Ca²⁺ homeostasis, controlling Ca²⁺ entry via negative feedback.^7,8 However, it is unknown whether the efficacy of coupling between SR Ca²⁺ release and I_Ca is the same in the t-tubule and surface membranes, so that the relative importance of these sites in cellular Ca²⁺ homeostasis is unknown. We have, therefore, investigated the regulation of I_Ca by Ca²⁺ released from the SR in normal ventricular myocytes, in which I_Ca triggers Ca²⁺ release predominantly at the t-tubules, and in myocytes in which the t-tubules have been physically and functionally uncoupled from the surface membrane (detubulated),⁵ in which Ca²⁺ release occurs predominantly at the surface membrane.⁹

	Materials and Methods

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This study conforms to the Guide for the Care and Use of Laboratory Animals published by the US National Institutes of Health (NIH Publication No. 85-23, revised 1996).

Isolation and Detubulation of Rat Ventricular Myocytes
Myocytes were isolated from the ventricles of Wistar rat hearts (Central Biomedical Services, University of Leeds, UK) and detubulated using formamide as described previously.^5,9 All experiments were performed at room temperature (22 to 25°C).

Recording I_Ca
Most of the experiments described in this article used the whole-cell (ruptured) patch clamp technique¹⁰ using Na⁺ and K⁺-free internal and external solutions to avoid contamination of I_Ca by overlapping ionic currents and to allow us to use a physiological holding potential. An Axopatch 200B (Axon Instruments) voltage clamp amplifier was used, controlled by a Pentium PC connected via a Digidata 1322A A/D converter (Axon Instruments), which was also used for data acquisition and analysis using pClamp software (Axon Instruments). Signals were filtered at 2 kHz using an 8-pole Bessel low-pass filter before digitization at 10 kHz and storage. Low resistance (1 to 3 M) patch pipettes were used and the junction potential between the pipette solution and the reference electrode was cancelled before obtaining a tight gigaseal (>1 G). Cell membrane capacitance was measured by integrating the capacitance current recorded during a 10-mV hyperpolarizing pulse from –80 mV. Cell capacitance and series resistance were compensated (>70%) so that the maximum voltage error was <3.5 mV. I_Ca was recorded during a 200-ms test pulse to 0 mV from a holding potential of –80 mV. Trains of depolarizing pulses were applied at 0.1 or 1 Hz.

The experiments shown in Figure 4 were performed using the perforated patch clamp technique¹¹ to record I_Ca and the Ca²⁺ transient simultaneously. Cells were loaded with the Ca²⁺ indicator Fura2-acetoxymethyl ester (AM, 3 µmol/L; Molecular Probes) for 10 minutes at room temperature. Fura2 fluorescence was elicited by alternate (every 2 ms) illumination with 340 and 380 nm light, obtained using a rotating filter wheel (Cairn Research Ltd) in front of a Xenon excitation lamp. The fluorescence emitted at 510 nm was monitored using a photomultiplier tube (Cairn Research Ltd). The ratio of fluorescence emitted at 510 nm during excitation at 340 nm to that emitted during excitation at 380 nm (R) is a function of [Ca²⁺]_i, and was converted to [Ca²⁺]_i as described in Data Analysis section. The tip of the patch electrode was filled with pipette solution and then back filled with the same solution containing 250 to 400 µg/mL amphotericin-B (Sigma). After seal formation and pipette capacitance compensation, holding potential was set to –40 mV, and 5 mV, 20-ms depolarizing pulses were applied to monitor pore formation. The pipette solution contained 1 mmol/L CaCl₂ to ensure cell death on accidental rupture of the membrane. Electrical access was typically obtained within 10 minutes and measurements were made after the capacitance transients became constant. I_Ca was recorded during a 500-ms test pulse to 0 mV from a holding potential of –40 mV. Trains of depolarizing pulses were applied at 0.2 Hz.

View larger version (29K): [in this window] [in a new window]   Figure 4. Relationship between ICa and [Ca2+]i in control and detubulated rat ventricular myocytes. A, Ca2+ transient (top trace in each panel) and ICa (bottom trace) in representative control (left) and detubulated (right) myocytes, elicited at 0 mV. B, Mean (±SEM) for control (black bars; n=14) and detubulated (white bars; n=16) myocytes: a, ICa density (left) and T0.37 (right); b, [Ca2+]i (left) and d[Ca2+]i/dtmax (right); c, time to d[Ca2+]i/dtmax (left) and (d[Ca2+]i/dtmax)/[Ca2+]i (right); and d, Gain measured as [Ca2+]i/ICa (left) and (d[Ca2+]i/dtmax)/ICa (right). See text for further details. *P<0.05 between cell types.

Solutions
Myocytes were studied in a chamber mounted on the stage of an inverted microscope (Nikon Diaphot). Cells were initially superfused with a normal physiological salt solution containing (in mmol/L) NaCl 113, KCl 5, MgSO₄ 1, CaCl₂ 1, Na₂HPO₄ 1, Na acetate 20, glucose 10, HEPES 10, and insulin 5 U/L; pH set to 7.4 with NaOH.

When I_Ca was measured using the ruptured whole-cell patch clamp configuration, the pipette solution contained (in mmol/L) CsCl 110, TEACl 20, MgCl₂ 0.5, Mg-ATP 5, EGTA 5, HEPES 10, and GTPTris 0.4; set to pH 7.2 with CsOH. A fast perfusion system placed close to the cell was used to deliver the external solution, which contained (in mmol/L) 4AP 5, TEACl 130, MgCl₂ 0.5, HEPES 10, Glucose 10, and CaCl₂ 1. At least 5 minutes was allowed for cell dialysis by the pipette solution before experiments were initiated.

When I_Ca was measured using the perforated whole-cell patch clamp configuration, the pipette solution contained (in mmol/L) CsCl 130, NaCl 10, MgCl₂.6H2O 1, HEPES 10, and CaCl₂ 1; pH set to 7.2 using CsOH. CsCl (5 mmol/L) was also added to the normal physiological (extracellular) solution to prevent I_K contamination.

Data Analysis
I_Ca was measured as the difference between the peak inward current and the current at the end of the depolarizing pulse. Currents are expressed as current density (pA/pF). Because the decay of I_Ca varied between cell types and experimental conditions, the kinetics of inactivation of I_Ca were characterized by the time required for the current to decay to 0.37 of the peak amplitude (T_0.37). Most currents decayed monoexponentially, in which case T_0.37 is equivalent to the time constant of decay. However in control cells, the decay was biexponential, in which case T_0.37 is used as a simple measure to compare the time course of decay in these cells with that in detubulated and treated cells. Frequency-dependent facilitation was analyzed by integrating I_Ca (pA·ms) during the 200 ms test pulse to obtain total Ca²⁺ influx during the pulse.

[Ca²⁺]_i was calculated from [Ca²⁺]_i=K_dß(R–R_min)/(R_max–R).¹² R_max, R_min, and ß were determined experimentally and K_d taken as 200 nmol/L.¹³ Traces were averaged (10 to 15 transients); the difference between diastolic and peak [Ca²⁺]_i was taken as [Ca²⁺]_i, and the maximum rate of change of [Ca²⁺]_i (d[Ca²⁺]_i/dt_max) was calculated using Origin software. [Ca²⁺]_i/I_Ca and (d[Ca²⁺]_i/dt_max)/I_Ca were taken as commonly used and useful approximations¹ of the gain of the Ca²⁺ release process as defined by Wier et al.¹⁴

Statistics
Data are presented as mean±SD in the text and mean±SEM in the figures. Statistical analysis was performed using SigmaStat software. A two-tailed unpaired t test was used to compare data from control and formamide treated cells when normal distribution and equal variance were confirmed, otherwise a nonparametric test (Mann-Whitney rank sum) was used. Paired t tests were used to test the effect of ryanodine or Ba²⁺ within the same group of cells. A value of P<0.05 was taken as significant.

	Results

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Detubulation Slows I_Ca Inactivation
Figure 1A shows recordings of I_Ca elicited by test pulses from –80 to 0 mV in representative control and detubulated ventricular myocytes. Detubulation decreased I_Ca density, confirming that this current is concentrated in the t-tubules.⁵ These traces also show that I_Ca recorded from control myocytes showed biphasic inactivation whereas I_Ca recorded from detubulated myocytes showed monophasic inactivation. Detubulation of ventricular myocytes significantly decreased cell capacitance and I_Ca density (193±22 pF and –12.6±5.7 pA/pF respectively in control, n=22; 137±34 pF and –5.3±2.2 pA/pF in detubulated cells, n=22; P<0.05; Figure 1B). The time required for the current to decay to 0.37 of its peak amplitude (T_0.37) was significantly longer in detubulated cells (27.1±7.8 ms, n=22) than in control cells (16.4±7.9 ms, n=22, P<0.05; Figure 1B).

View larger version (22K): [in this window] [in a new window]   Figure 1. ICa in control and detubulated rat ventricular myocytes. A, ICa elicited at 0 mV in representative control (left) and detubulated (right) myocytes. B, Mean±SEM cell capacitance (left), ICa density at 0 mV (middle), and time for ICa to decay to 0.37 of peak amplitude (T0.37; right) in control (n=22, black bars) and detubulated (n=22, open bars) myocytes. *P<0.05 between cell types.

To ensure that the slowed inactivation of I_Ca was not due to a direct effect of formamide, the effect of formamide treatment on I_Ca was determined in atrial cells, which lack t-tubules.⁹ I_Ca was recorded in atrial myocytes using the same solutions and experimental conditions as for ventricular myocytes except that the holding potential was set to –40 mV in order to inactivate T-type Ca²⁺ current. In contrast to ventricular myocytes, I_Ca elicited at 0 mV showed biphasic inactivation in control and formamide-treated atrial myocytes (Figure 2A), and formamide treatment of atrial cells had no significant effect on cell capacitance, I_Ca density or T_0.37 (Figure 2B).

View larger version (19K): [in this window] [in a new window]   Figure 2. ICa in control and formamide-treated rat atrial myocytes. A, ICa elicited at 0 mV in representative control (left) and formamide-treated (right) atrial myocytes. B, Mean (±SEM) cell capacitance (left), ICa density at 0 mV (middle), and time for ICa to decay to 0.37 of peak amplitude (T0.37; right) in control (n=7, black bars) and formamide-treated (n=7, open bars) atrial myocytes. There were no significant differences between control and formamide-treated atrial myocytes.

Thus it appears that in ventricular myocytes the rate of inactivation of I_Ca is different at the t-tubules and the surface membrane. Subsequent experiments were designed to investigate the mechanism(s) underlying this difference.

I_Ca Inactivation Due to Ca²⁺ Released From the SR
The inactivation phase of I_Ca is due to two mechanisms: voltage-dependent inactivation (VDI) and Ca²⁺-dependent inactivation (CDI).⁷ To investigate whether VDI is altered by detubulation we recorded I_Ca using Ba²⁺ as the charge carrier, to eliminate CDI.⁷ I_Ba was recorded at –10 mV to compensate for screening of surface charge.¹⁵ The top panel of Figure 3A shows representative currents from control and detubulated ventricular myocytes recorded using Ca²⁺ (gray trace) or Ba²⁺ (black trace) as the charge carrier; the currents have been normalized to allow comparison of their time course. Inactivation of I_Ba was not significantly different in control and detubulated myocytes (T_0.37: 72±7 ms, n=7 control, versus 79±15, n=8 detubulated; P>0.05; Figure 3B), suggesting that VDI is the same at the t-tubules and the surface membrane. These results, and the observation that the rapid phase of inactivation, which is due to CDI,⁷ is reduced in detubulated myocytes (eg, Figure 3A, top panel), suggest that CDI rather than VDI is reduced in detubulated myocytes.

View larger version (27K): [in this window] [in a new window]   Figure 3. Effects of Ba2+, ryanodine, and BAPTA on inactivation of ICa in control and detubulated rat ventricular myocytes. A, ICa in representative control (left) and detubulated (right) myocytes. Top, ICa with 1 mmol/L Ca2+ (gray traces) or 1 mmol/L Ba2+ (black traces) as the charge carrier in the same cell. Middle, ICa in the absence (gray traces) or presence (black traces) of 30 µmol/L ryanodine in the same cell. Bottom, ICa when 5 mmol/L EGTA has been replaced with 10 mmol/L BAPTA in the pipette solution. Currents are normalized and elicited at 0 mV except when using Ba2+ as a charge carrier (–10 mV; see text). B, Mean (±SEM) time for ICa to decay to 0.37 of peak amplitude in control (black bar) and detubulated (open bar) myocytes when Ba2+ was used as the charge carrier (n=7 and 8, respectively; left), in the presence of ryanodine (30 µmol/L; n=13 and 13; middle), and with BAPTA in the pipette solution (n=8 and 9; right). There were no significant differences between control and detubulated myocytes.

In ventricular myocytes, CDI is due in part to Ca²⁺ released from the SR.^16,17 The SR inhibitor ryanodine (30 µmol/L) was therefore used to investigate the role of SR Ca²⁺ release in the decrease of CDI observed in detubulated myocytes. Ryanodine significantly slowed inactivation and prolonged I_Ca in control and detubulated myocytes (Figure 3A, middle panels); however, this effect was greater in control cells, so that in the presence of ryanodine, T_0.37 was not significantly different in control and detubulated myocytes (29.7±4.8 ms, n=13, versus 32.9±8.3, n=13, respectively; P>0.05; Figure 3B; cf, Figure 1B, right). This suggests, in agreement with previous work,^16–20 that Ca²⁺ release from the SR inactivates I_Ca; however, this effect is reduced after detubulation, suggesting that SR Ca²⁺ release has a greater effect on I_Ca inactivation in the t-tubules than at the surface membrane. Including the fast Ca²⁺ chelator BAPTA (10 mmol/L) in the pipette solution had similar effects (Figure 3A, bottom, and 3B).

The decreased effect of SR Ca²⁺ release on I_Ca inactivation in detubulated cells could be explained if I_Ca triggered less SR Ca²⁺ release at the surface membrane. To address this hypothesis, we simultaneously recorded I_Ca and intracellular Ca²⁺.

Effect of Detubulation on I_Ca-Induced Ca²⁺ Release
The left traces in Figure 4A show representative recordings of [Ca²⁺]_i (top) and I_Ca (bottom) from a control myocyte during a pulse to 0 mV. Under these conditions, the majority of L-type Ca²⁺ current and Ca²⁺ release occur at the t-tubule (see Introduction). The right traces in Figure 4A show corresponding recordings from a detubulated cell. In these cells, the amplitude of I_Ca and the Ca²⁺ transient are reduced, as reported previously,⁵ and are determined predominantly by Ca²⁺ entry and release at the surface membrane.⁹ The decay of the Ca²⁺ transient is also slower because Na⁺-Ca²⁺ exchange occurs predominantly in the t-tubules.^21,22

Figure 4B shows mean data from 14 control and 16 detubulated cells. Figure 4B, a, shows that I_Ca density was significantly reduced (left) and T_0.37 for I_Ca inactivation significantly prolonged (right) in detubulated cells, as in the previous set of experiments (cf, Figure 1). [Ca²⁺]_i and d[Ca²⁺]_i/dt_max were significantly smaller after detubulation (Figure 4B, b), although the time to d[Ca²⁺]_i/dt_max, and d[Ca²⁺]_i/dt_max normalized to [Ca²⁺]_i ((d[Ca²⁺]_i/dt_max)/[Ca²⁺]_i) were the same in the two cell types (Figure 4B, c) The gain of the Ca²⁺ release process was calculated in two ways (see Materials and Methods); Figure 4B, d, shows that there was a modest ( 20%) but nonsignificant decrease in gain, determined using either method in detubulated myocytes. Thus, the effectiveness of I_Ca in causing SR Ca²⁺ release is not significantly different in the two cell types and, therefore, at the t-tubules and surface membrane. Thus, it appears that the slowing of I_Ca inactivation after detubulation cannot be explained by a decrease in local SR Ca²⁺ release; it is more likely that a given SR Ca²⁺ release has less effect on I_Ca inactivation at the surface membrane than in the t-tubules.

Effect of Detubulation on I_Ca Facilitation
To investigate this idea further, we studied the response of control and detubulated myocytes to an increase in stimulation rate, which causes frequency-dependent facilitation (FDF) of I_Ca that is dependent, in part, on SR Ca²⁺ release.¹⁸ FDF in a representative control cell is shown in Figure 5A (top, left) as increased amplitude and slowed inactivation of I_Ca during the fourth voltage clamp pulse from –80 to 0 mV after an increase in stimulation rate from 0.1 to 1 Hz. The SR inhibitor ryanodine (30 µmol/L) slowed the time course of inactivation, and abolished FDF (Figure 5A, bottom left). The right traces in Figure 5A show that FDF was absent in detubulated cells in either the absence (top) or presence (bottom) of ryanodine. Figure 5B shows that I_Ca, expressed as the integral of the current, increased significantly in control cells when stimulation rate was increased but not in detubulated cells (I₄/I₁=1.34±0.23, n=11, and 0.97±0.08, n=13, respectively), nor was there any significant change in either control or detubulated cells in the presence of ryanodine (I₄/I₁=0.96±0.06 and 0.98±0.09, respectively). Inclusion of BAPTA in the patch pipette, or use of Ba²⁺ as the charge carrier, also abolished FDF (not shown). These data support the idea that the effect of SR Ca²⁺ release on I_Ca is smaller at the surface membrane than in the t-tubules.

View larger version (25K): [in this window] [in a new window]   Figure 5. Frequency-dependent facilitation of ICa in control and detubulated myocytes. A, ICa recorded in response to the first (1) and the fourth (4) pulse to 0 mV after increasing stimulation rate from a steady state at 0.1 Hz to 1 Hz (schematic protocol shown above the traces) in representative control (left) and detubulated (right) myocytes in the absence (top) and presence (bottom) of ryanodine in the same cell. B, Mean (±SEM) changes in ICa charge (integral) between the 1st and 4th ICa after increasing stimulation rate in control (n=11, black bars) and detubulated (n=13, open bars) myocytes in the absence (left) and presence (right) of ryanodine. *P<0.05 between cell types;, #P<0.05 between treatments.

Quantification of CDI
The principal findings of this study are summarized in the Table. To quantify inactivation of I_Ca, we measured the fraction of current remaining 20 ms after its peak (I_R20). This time was chosen because the peak of SR Ca²⁺ release occurs 5 ms after peak I_Ca, with a time to 90% decay of 45 ms.²³

View this table: [in this window] [in a new window]   Table 1. Fraction of ICa Remaining 20 ms After its Peak, and Proportion of CDI in Control and Detubulated Myocytes

When Ba²⁺ is used as the charge carrier, I_Ca inactivates almost exclusively by VDI, and I_R20Ba was not significantly different between control and detubulated myocytes (Table). When Ca²⁺ is used as the charge carrier, I_Ca inactivates by VDI and CDI. Thus the difference between I_Ca and I_Ba represents the current inactivated by CDI. By normalizing to I_R20Ba{ie, [(I _R20Ba–I_R20Ca)/I _R20Ba]x100]}, we estimate total CDI to be 62% in control and 40% in detubulated myocytes (Table).

To separate SR-induced CDI from total CDI, we used a similar analysis, comparing I_R20Ca in the presence and absence of ryanodine. The difference current between I_Ca and I_Ca in the presence of ryanodine represents the fraction of I_Ca inactivated by SR CDI. By normalizing to total CDI {ie, [(I_R20Rya–I_R20Ca)/(I_R20Ba–I_R20Ca)]x100}, we estimate the SR CDI to be 40% of total CDI in control myocytes and 8% in detubulated myocytes (table 1). Correcting for the 10% of cells that remain non-detubulated following formamide treatment, we can calculate that 20% of I_Ca is located at the surface membrane (see Discussion) and thus that SR-dependent CDI at the t-tubules is 50%, and 5% at the surface membrane.

	Discussion

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Experimental Approach
The method used to detubulate rat ventricular myocytes has been described and validated previously.⁹ The observed effects are unlikely to be due to formamide treatment per se because no changes were observed in atrial cells (Figure 2 and Brette et al).⁹

The decrease in I_Ca and cell capacitance after formamide treatment, and correction for the presence of 10% nondetubulated myocytes (assessed as in Despa et al²¹), indicate that 80% of I_Ca is within the t-tubules, so that the density of I_Ca in the t-tubules is 6 fold higher than in the surface membrane. Thus, Ca²⁺ signaling in control cells is dominated by the t-tubules, whereas Ca²⁺ signaling in detubulated cells occurs predominantly at the cell surface.

It is well known that there is bidirectional cross-talk between I_Ca and RyRs in cardiac myocytes.¹ This local Ca²⁺ signaling occurs in a restricted diffusion space; the use of a low concentration of slow Ca²⁺ buffer (EGTA)²⁴ as in the present study allows Ca²⁺ in the bulk cytosol to be "clamped" (indicated by the absence of cell contraction) while permitting Ca²⁺ in the dyadic space to change (indicated by the effect of ryanodine on I_Ca), whereas a faster Ca²⁺ chelator (BAPTA) inhibits this local signaling, slowing the rate of inactivation of I_Ca to the same level in control and detubulated myocytes (Figure 3). This local Ca²⁺ signaling has been observed in numerous studies recording I_Ca using EGTA to buffer bulk Ca²⁺,^{16,18–20,23} and Song et al²³ have demonstrated such signaling directly using confocal microscopy to show I_Ca-induced Ca²⁺ release in the dyadic space ("Ca²⁺ spikes") in the presence of EGTA. The solutions used in the present study allowed I_Ca to be recorded without contamination by Na⁺-Ca²⁺ exchange current, which is concentrated in the t-tubules.^21,22 Such contamination could alter the shape of I_Ca, particularly from the physiological holding potential used (–80 mV), which is necessary to elicit FDF.⁷

Modulation of I_Ca by SR Ca²⁺ Release Is Smaller at Surface Sarcolemma Than T-Tubule
Ca²⁺ entering the cell through the L-type Ca²⁺ channel, and Ca²⁺ released from the SR, appear to bind to calmodulin (CaM) that is prebound to the Ca²⁺ channel, causing channel inactivation.²⁵ Figure 3A shows such Ca²⁺-dependent inactivation; inhibition of SR Ca²⁺ release by ryanodine slowed the time course of inactivation in control cells. In detubulated cells, the time course of inactivation of I_Ca was slower than in control cells; this is probably due to lack of feedback of Ca²⁺ released from the SR rather than to smaller Ca²⁺ influx, because T_0.37 was little affected by ryanodine in detubulated cells, and was the same in control and detubulated cells in the presence of ryanodine (Figure 1B) despite differences in the amplitude of I_Ca. It is also unlikely that this prolongation of T_0.37 occurs because of the solutions used for cell dialysis, because T_0.37 was also longer in detubulated myocytes in perforated patch experiments (Figure 4B, a), suggesting that this phenomenon exists in physiological conditions. Although absolute T_0.37 was slower in both cell types compared with ruptured patch experiments, probably due to the difference in holding potential, the ratio of T_0.37 in detubulated:control myocytes in perforated patch (1.53) was similar to that in the ruptured patch experiments (1.65).

The idea that Ca²⁺ released from the SR has less effect on inactivation of I_Ca in detubulated cells is supported by the observations that FDF did not occur in control cells in the presence of ryanodine (showing that it is due to SR Ca²⁺ release), that it did not occur in detubulated cells and that ryanodine had no effect on the response of I_Ca to increasing stimulation rate in detubulated cells. FDF may be a consequence of Ca²⁺ binding to CaM on the Ca²⁺ channel,²⁵ phosphorylation of the channel after activation of Ca²⁺/calmodulin–dependent protein kinase II (CaMKII)^26,27 or to less Ca²⁺-dependent inactivation of I_Ca.¹⁸

I_Ca-Induced SR Ca²⁺ Release Is the Same at the T-Tubules and Surface Membrane
Although SR Ca²⁺ release had a greater effect on I_Ca at the t-tubules than at the surface membrane, the gain of the Ca²⁺ release process was not significantly altered by detubulation. Our experimental conditions precluded analysis of gain as first defined by Wier et al;¹⁴ instead, we used two methods widely accepted as useful approximations:¹ [Ca²⁺]_i/I_Ca and (d[Ca²⁺]_i/dt_max)/I_Ca. [Ca²⁺]_i, and hence [Ca²⁺]_i/I_Ca might be overestimated in detubulated myocytes because of the spatially and temporally inhomogeneous Ca²⁺ transient,^22,28 so we also used d[Ca²⁺]_i/dt_max. However, neither method showed a significant change in gain, nor was the time to d[Ca²⁺]_i/dt_max altered after detubulation. This suggests that coupling of Ca²⁺ release to Ca²⁺ influx is the same in control and detubulated cells, and hence at the t-tubules and surface membrane; differences in such coupling would be expected to alter these measures.²⁹ These data agree with previous work showing that the rate of rise of Ca²⁺ is the same at the cell periphery after detubulation as at the periphery and center of control myocytes, using confocal microscopy.^9,28

Thus, it is unlikely that the smaller effect of SR Ca²⁺ release on I_Ca at the surface membrane is due to less local Ca²⁺ release, because feed-forward (I_Ca to SR Ca²⁺ release) is the same but feedback (SR Ca²⁺ release to I_Ca) is smaller at the surface membrane than in the t-tubules. A possible explanation for this dyadic rectification is considered next.

Possible Mechanisms for Differential Modulation of I_Ca by SR Ca²⁺ Release
There are a number of possible explanations for why I_Ca is affected more by SR Ca²⁺ release at the t-tubules than at the surface membrane: (1) Ca²⁺-dependent modulation of I_Ca depends on local Ca²⁺ entry and local Ca²⁺ release¹⁶; the present data could be explained if either is different in detubulated cells. It is unlikely that differences in local Ca²⁺ entry can account for the present observations, because in the presence of ryanodine, the rate of inactivation of I_Ca was the same in control and detubulated cells (Figure 3). It is also unlikely that differences in local Ca²⁺ release are responsible, because this appears to be the same at the t-tubules and surface membrane (above), consistent with previous work showing that SR Ca²⁺ content (assessed using caffeine) is not altered by detubulation.^22,28 (2) Ca²⁺ entering the cytoplasm via I_Ca and from the SR enters a restricted diffusion space in which high [Ca²⁺] occurs before Ca²⁺ diffuses to the bulk cytosol.¹ The present data could be explained if the rise of Ca²⁺ in this space, and sensed by the Ca²⁺ channel, were lower at the surface sarcolemma. This could occur if the dyadic space is less restricted at the surface membrane than at the t-tubules, allowing Ca²⁺ released from the SR to diffuse away rapidly, or if RyR and I_Ca are less well colocalized at the surface membrane; either could result in a rise of Ca²⁺ around the Ca²⁺ channel that is too small or brief to affect I_Ca. However, this seems unlikely because electron microscopy has shown no difference in the size of the dyadic space at the surface sarcolemma and t-tubules,³⁰ and immunocytochemistry has shown similar colocalization of I_Ca and RyR^4,6 at the two sites. In addition, any differences in the geometry and/or structure of the dyadic cleft would be expected to alter the gain of Ca²⁺ release and the rate of rise of Ca²⁺,²⁹ both of which appear to be unaltered after detubulation. (3) The sensitivity of I_Ca to Ca²⁺ could be different at the 2 sites. This could be due to different channel isoforms; although several different isoforms have been found in ventricular myocytes,³¹ we are unaware of any evidence that they are differentially located between the t-tubule and surface membranes. Alternatively, CaM or CaMKII may regulate I_Ca more effectively at the t-tubules as a consequence of concentration of these proteins at the t-tubules²⁶ or closer colocalization with the channel. This could explain why I_Ca-induced SR Ca²⁺ release is the same at both sites, whereas the feedback is different. We have recently shown such localized regulation of I_Ca by protein kinase A at the t-tubules.²⁸

Physiological Significance
Previous work has shown that Na⁺-Ca²⁺ exchange is concentrated in the t-tubules and is more sensitive to Ca²⁺ released from the SR than to bulk cytoplasmic Ca²⁺.^8,22,32 The present work shows that I_Ca is concentrated in the t-tubules and is regulated more effectively by SR Ca²⁺ release at the t-tubules than at the surface membrane. These data suggest that Ca²⁺ released from the SR will stimulate Ca²⁺ efflux via Na⁺-Ca²⁺ exchange, and inhibit Ca²⁺ influx via I_Ca, most effectively at the t-tubules. Thus, the t-tubules appear to be the primary site at which changes in SR Ca²⁺ release bring about compensatory changes in transsarcolemmal Ca²⁺ flux⁸ and are therefore central to cellular Ca²⁺ homeostasis.²² This may also be important during development and in pathological conditions (see review);³ for example, during heart failure, t-tubule density decreases³ and FDF is blunted^33,34; this could be explained by the present work.

	Acknowledgments

 
This study was supported by the Wellcome Trust and British Heart Foundation, including a travel grant to L.S. from the Wellcome Trust.

	Footnotes

 
*Both authors contributed equally to this study.

Original received July 1, 2003; resubmission received February 25, 2004; revised resubmission received May 28, 2004; accepted June 3, 2004.

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