INTRODUCTION

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CA²⁺-induced Ca²⁺ release (CICR) is the major mechanism for Ca²⁺ release from the sarcoplasmic reticulum (SR) of cardiac muscle (1, 8, 9). Ca²⁺ entering the cell via L-type Ca²⁺ channels (1) and, under some conditions, the Na⁺/Ca²⁺ exchange mechanism (18), triggers the release of further Ca²⁺ from the SR. Current evidence suggests that, of these two pathways, Ca²⁺ entering via the L-type Ca²⁺ channels is the main trigger for Ca²⁺ release (e.g., Ref. 7 and see DISCUSSION) and that the transverse (T) tubules are a particularly important site for the coupling of Ca²⁺ entry across the cell membrane to Ca²⁺ release from the SR (28).

T tubules are invaginations of the sarcolemma that occur perpendicular to the longitudinal axis of the cell at intervals of ~1.8 to 2.0 µm (23, 25). The percentage of the total surface membrane contained in the T tubules depends on the species (24) and the region of the heart (e.g., see Ref. 13). By analogy with skeletal muscle, it has been suggested that T tubules help the spread of excitation to the center of the cell so that activation of Ca²⁺ release is synchronized to produce homogenous Ca²⁺ release throughout the cell (3). This suggestion is supported by three lines of evidence. First, Ca²⁺ sparks evoked by electrical stimulation occur predominantly close to T tubules (28), suggesting an important role for the T tubules in excitation-contraction coupling. Second, atrial cells (that lack a T tubular system) exhibit Ca²⁺ release that is initially localized near the subsarcolemmal space and subsequently spreads to the central regions of the cell (13). Finally, cultured ventricular myocytes, which lose their T tubules, exhibit biphasic Ca²⁺ transients that may result from spatially nonuniform SR Ca²⁺ release (20). However, in the latter two cases, other aspects of cell function (e.g., protein expression) are also different. Thus direct evidence for the role of T tubules is still sparse.

Previous studies designed to elucidate the role of the T tubules in skeletal muscle have used membrane-permeant solutes such as glycerol (12) and formamide (26) to produce an osmotic shock that disrupts the normal coupling between the T tubules and the surface membrane (detubulation). Such methods have not been successful in multicellular cardiac preparations. However, we have used formamide to produce osmotic shock in single isolated cardiac cells to investigate the functional role of the T tubules in excitation-contraction coupling.

	METHODS

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Isolation of myocytes and measurement of cell length and width. Ventricular myocytes were isolated from female adult Wistar rat hearts. The animals were stunned and then killed by cervical dislocation, in accordance with the Home Office Guidance on the Operation of the Animals (Scientific Procedures) Act of 1986. Myocytes were isolated as described previously (10) and allowed to settle in the recording chamber, which was mounted on an inverted microscope (Diaphot; Nikon). The setup was configured for the simultaneous recording of cell length or width, fluorescence transients, and membrane currents (10, 14).

Myocytes were illuminated with red light (>610 nm) to generate an image of the cell, which was detected by a camera mounted on the side limb of the microscope and displayed on a monitor. The length or width of this image was monitored using an edge detection system (Crescent Electronics). These measurements were made on myocytes that had been loaded with the Ca²⁺ chelator 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA)-AM (5 µmol/l for 15 min) to avoid formamide-induced changes in intracellular Ca²⁺ concentration ([Ca²⁺]_i; see Fig. 4) that could also alter the dimensions of the cell.

Measurement of Ca²⁺ current and [Ca²⁺]_i. Two series of voltage-clamp experiments were carried out to determine the effects of formamide on Ca²⁺ current (I_Ca). In the first, I_Ca and [Ca²⁺]_i were monitored simultaneously in myocytes loaded with fura 2-AM (5 µmol/l for 15 min). The membrane potential was controlled using the perforated patch-clamp technique, with amphotericin B (250 or 500 µg/ml) as the pore-forming agent, as described previously (14). The pipette solution used for these recordings contained (in mmol/l) 120.0 cesium glutamate, 20.0 KCl, 10.0 NaCl, 1.0 MgCl₂, 1.0 CaCl₂, and 10.0 HEPES, adjusted to pH 7.3 with tetraethylammonium hydroxide (TEA-OH). In the second series of experiments, membrane capacitance and I_Ca were monitored using conventional whole cell voltage-clamp in BAPTA-loaded cells. The pipette solution for these experiments contained (in mmol/l) 120.0 cesium glutamate, 20.0 KCl, 10.0 NaCl, 5.0 Mg-ATP, 1.0 BAPTA, and 10.0 HEPES, adjusted to pH 7.3 with TEA-OH. The amplitude of I_Ca was measured as the difference between peak inward current and the current remaining at the end of the 300-ms voltage-clamp pulse. Cs-based pipette solutions and 20.0 mmol/l CsCl in the external solution were used in both sets of experiments to avoid contamination of I_Ca by potassium currents.

	RESULTS

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Effects of formamide on cell volume. We first determined the effects of formamide on cell volume, because detubulation is thought to occur in response to the increase in volume that occurs on washout of a membrane-permeable solute (e.g., Refs. 12 and 26 and see DISCUSSION).

Figure 1A, top trace, shows that addition of 1.5 mol/l formamide to the bathing solution caused a rapid (<1 min) decrease in resting cell length, from 106.6 ± 2.0 to 96.8 ± 2.1 µm (n = 5, P < 0.001). This was followed by a further slower decrease, to 84.0 ± 2.1 µm, during the remainder of the 15-min exposure. The second trace shows that changes in cell width occurred over a similar time course, although the initial decrease, from 30.2 ± 2.0 to 28.2 ± 2.1 µm (P < 0.01, n = 5), was followed by an increase (rather than a further decrease) in the continued presence of formamide. Washout of formamide caused the opposite effects. These data were used to calculate cell volume (v), as described previously (2) where l and w are cell length and width, respectively. Cell depth (d) was derived by assuming the cell to be an elliptical cylinder with a minor-to-major axis ratio of 1:3 (2). Cell volume calculated using the top two traces in Fig. 1A is shown in the third trace. Figure 1A, bottom, shows that average cell volume decreased by ~21.0% within 1 min of adding 1.5 mol/l formamide and increased by 17.8% (n = 5) after washout of formamide, again in <1 min of washout, before recovery toward control levels.

View larger version (36K): [in this window] [in a new window]   Fig. 1.   Effects of formamide on cell length, width, volume (A), and membrane staining with di-8-aminonaphthylethenylpyridinium (Di-8-ANNEPS; B). A: top two traces show the effect of formamide (1.5 mol/l for 15 min) on the length and width of representative BAPTA-loaded rat ventricular myocytes; because only one of these measurements can be made at a time, these traces are from 2 different cells. These were used to estimate the changes in cell volume (see text for details). Bottom trace shows average changes of calculated cell volume in 5 cells. B: confocal image (512 × 512 pixels) through the center of a control cell stained with Di-8-ANNEPS, showing clear staining of the T tubules. C: similar confocal image taken from a representative myocyte stained with Di-8-ANNEPS after washout of formamide. White scale bar in B represents 10 µm and is also applicable to C.

Effects of formamide on membrane staining with Di-8-ANNEPS. To investigate whether these changes in volume caused detubulation, we used confocal microscopy to visualize staining of the cell membrane with the lipophilic fluorescent indicator Di-8-ANNEPS (13, 20). The confocal images in Fig. 1 show Di-8-ANNEPS staining of a representative control myocyte (Fig. 1B), characterized by regular cross-striations at intervals of 1.82 ± 0.02 µm (n = 9) along the longitudinal axis of the cell, as described previously for staining of the T tubules (23, 25). Similar staining patterns were observed in all the sections taken at 1.0-µm intervals through the depth of the cell. In contrast, there were no such cross-striations in 609 out of 700 cells examined after treatment with formamide (Fig. 1C). These data suggest that the Di-8-ANNEPS no longer had access to the T tubules; this could be either because the T tubule membranes had been lost or because the T tubules had been uncoupled from the surface membrane. However, because labeling with Di-8-ANNEPS was carried out after the osmotic shock treatment (see METHODS), we cannot distinguish between these possibilities.

Effects of formamide on I_Ca and Ca²⁺ transient in voltage-clamped cells. If detubulation occurs during the rapid swelling of the cell upon washout of formamide (26), changes in I_Ca and the Ca²⁺ transient might also be expected to occur during this period. Figure 2A shows a portion of the cell volume record (taken from Fig. 1) on an expanded time scale and records of I_Ca and Ca²⁺ transients recorded simultaneously using the perforated patch-clamp technique and fura 2, respectively (Fig. 2B), showing that cell expansion after washout of formamide is accompanied by a rapid decrease in the amplitude of I_Ca and the accompanying Ca²⁺ transient. Figure 2C shows these changes on a faster time base, showing more clearly the parallel decrease in these parameters. These data are compatible with the idea that detubulation occurs during cell swelling upon washout of the solute (26).

View larger version (19K): [in this window] [in a new window]   Fig. 2.   Effects of osmotic shock on the time course of changes in cell volume, Ca2+ current (ICa), and Ca2+ transient in rat ventricular myocytes. A: part of the record from the cell volume trace in Fig. 1 shown on an expanded time scale. B: records of membrane current (top) and Ca2+ transients (bottom) on the same time scale as A, obtained during perforated patch-clamp recordings from fura 2-loaded myocytes, during washout of formamide, as indicated by the end of the bar above the traces. Upward deflections on the current record are capacity transients, and downward deflections are ICa (downward-going capacity transients have been removed using a sample-and-hold circuit to enable changes in ICa to be seen more easily). F340/380, ratio of fluorescence excited at 340 and 380 nm. C: section of traces from B (indicated by bar labeled C above lower trace in B) on a faster time base. It should be noted that, since the Ca2+ transients and ICa change during exposure to formamide (see text), some of the changes shown in this figure on washout of formamide will be due to reversal of these effects; this does not, however, alter the data presented in the text, since the control data were not obtained in the presence of formamide (see RESULTS).

View larger version (20K): [in this window] [in a new window]   Fig. 3.   Effects of formamide treatment on ICa in rat ventricular myocytes. Superimposed records of ICa obtained when the membrane potential was stepped in 4-mV increments from a holding potential of 40 mV to between 36 and +52 mV for 300 ms at 0.5 Hz in a representative control cell (A) and in a representative cell after treatment with formamide (B). C: mean ± SE (n = 5-6) current-voltage relationship for ICa in control () and formamide-treated () cells. D: mean ICa density (± SE) after ICa was normalized to cell capacitance in control () and formamide-treated () cells.

Effects of formamide on Ca²⁺ transients in field-stimulated myocytes. Figure 4A shows that addition of formamide caused an initial decrease and then an increase in the amplitude of the Ca²⁺ transient. This was followed by a slower decrease in amplitude to near control levels during the continued presence of formamide. Diastolic Ca²⁺ remained elevated throughout the exposure to formamide.

View larger version (18K): [in this window] [in a new window]   Fig. 4.   Effects of formamide on Ca2+ transients in field-stimulated rat ventricular myocytes. A: slow time base record of fura 2 fluorescence before, during, and after 15-min exposure to 1.5 mol/l formamide, as indicated above the record. Electrical stimulation was briefly interrupted before and 5, 10, 15, and 30 min after the application of formamide, and 20 mmol/l caffeine was pressure ejected onto the cell to assess the Ca2+ content of the sarcoplasmic reticulum. B: fast time base records of Ca2+ transients from a different cell before and 5 and 30 min after exposure to formamide. C: response of intracellular Ca2+ to maintained exposure to caffeine from another cell. Unlike the experiment shown in A, in which caffeine was applied rapidly and briefly to ascertain the peak of the caffeine-induced release, in C the application of caffeine was maintained until Ca2+ had returned close to baseline levels (for 8-10 s).

	DISCUSSION

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Detubulation of single ventricular myocytes. The high osmotic pressure of the formamide solution (1,780 mosmol), compared with the control solution (286 mosmol), would be expected to cause the cell to lose water and shrink (Fig. 1A). However, formamide is membrane permeant and therefore enters the cell down its concentration gradient, accompanied by water, so the cell subsequently expands in the continued presence of formamide (Fig. 1A). During washout, the intracellular concentration of formamide is initially high so that water enters the cell, causing the rapid expansion (Figs. 1A and 2A) that is thought to cause the T tubules to break from the surface membrane while the surface membrane reseals (26). The data shown in Figs. 1 and 2 are compatible with this idea. Formamide will then leave the cell, taking water with it, to allow the cell volume and shape to recover slowly.

Several lines of evidence suggest that formamide-induced changes in cell volume are effective in detubulating cardiac myocytes. First, the absence of cross-striations from the center of cells stained with the lipophilic dye Di-8-ANNEPS after formamide treatment (Fig. 1C) is strong evidence that T tubules were either disrupted or uncoupled from the cell surface in these cells. Second, detubulation would be expected to occur predominantly during the rapid expansion that occurs upon washout of formamide. The data in Figs. 1 and 2 show that cell expansion does occur rapidly and that this is accompanied by a decrease in the amplitude of functional parameters such as I_Ca and the Ca²⁺ transient (Fig. 2B). Third, membrane capacitance (and presumably membrane area, if capacitance is uniformly 1.0 µF/cm²) was decreased by 26.4% after detubulation. This is close to previous estimates of the proportion of the membrane contained in the T tubules in rat ventricular myocytes (33%; see Ref. 24) and therefore also suggests that T tubules have been largely removed. Finally, the biphasic Ca²⁺ transients shown in Fig. 4B, right, are similar to the transients seen in cultured myocytes in which such changes were suggested to result from the detubulation that occurs in culture (Ref. 20 and see below).

An alternative that should be considered, however, is the possibility that formamide has direct effects on cardiac cells that might account for the present data. During exposure to formamide, the amplitude of the Ca²⁺ transient changes (Fig. 4A) and, in the cells in which it was monitored throughout exposure to formamide, I_Ca initially increased and subsequently decreased. Although these changes could be due to formamide itself or secondary to the induced changes of volume (Fig. 1A), they raise the possibility that formamide has direct effects on cardiac cells. There are a number of lines of evidence, however, that make it unlikely that such effects can account for the present results. First, the changes of I_Ca and the Ca²⁺ transient reported here occurred on wash off, not wash on, of formamide (Fig. 2), which is unexpected for a direct effect of formamide, and occurred with a similar time course to the change in cell volume (Fig. 2), consistent with this being important in the observed changes. Second, the characteristics of I_Ca, including its configuration and voltage dependence (Fig. 3), and its response to BAY K 8644 and isoprenaline (not shown) were unchanged after formamide treatment, as was the response to caffeine (Fig. 4), so that any direct effects of formamide would have to be very specific to account for the observed changes, and it seems unlikely that direct effects of formamide would be so specific as to produce a series of changes so compatible with detubulation (see above).

Previous attempts to use this technique in cardiac muscle have been unsuccessful (5), possibly because multicellular preparations were used in which diffusion delays could have slowed changes in cell volume and hence decreased the efficacy of the shock. It is also possible that the most commonly used solute (glycerol) crosses the cell membrane too slowly to increase intracellular concentration sufficiently to induce osmotic shock; in support of this idea, in an earlier series of experiments we were unable to achieve effective detubulation using glycerol (data not shown). Although excitation-contraction coupling remains disrupted 30 min after washoff of formamide (e.g., Fig. 4), it is unclear whether the observed changes reverse over longer periods than those monitored in the present study.

Functional consequences of detubulation on Ca²⁺ handling in cardiac muscle. The percentage decrease in the amplitude of I_Ca was greater than the percentage decrease in membrane capacitance (and hence membrane area). Thus the density of I_Ca decreased, consistent with previous suggestions that L-type Ca²⁺ channels are concentrated in the T tubules (19). This has implications for excitation-contraction coupling in cardiac cells. Current models of this process suggest that local Ca²⁺ release, from a cluster of ryanodine receptors, depends on local trans-sarcolemmal Ca²⁺ influx, which increases local Ca²⁺ concentration, thus increasing the open probability of adjacent ryanodine receptors (4, 21, 22, 27). Such Ca²⁺ release has previously been shown to be triggered by the L-type I_Ca in a stochastic manner (4, 21, 22), with a voltage dependence similar to that of the L-type I_Ca (22, 27); such release does not, however, normally appear to be triggered by Ca²⁺ influx via the Na⁺/Ca²⁺ exchanger (22). These data suggest that the main trigger for release is normally Ca²⁺ influx via the L-type Ca²⁺ channels, rather than influx via the Na⁺/Ca²⁺ exchanger. Labeling studies have shown that the L-type Ca²⁺ channel and the ryanodine receptor are concentrated in, and close to, the T tubules, respectively, and are closely juxtaposed (19), suggesting a possible mechanism for the potency of Ca²⁺ influx via I_Ca as a Ca²⁺ trigger. However some studies also show a concentration of the Na⁺/Ca²⁺ exchange protein in the T tubules (6, 11); in this case, the relative lack of potency of Ca²⁺ influx via the exchanger as a trigger of SR Ca²⁺ release could be due to the position of the exchanger relative to the ryanodine receptor or to lower Ca²⁺ flux rate through the exchanger, since the size and rate of change of Ca²⁺ at the surface of the SR is known to determine the effectiveness of the Ca²⁺ trigger (8). However, other studies show a more even distribution of the exchange protein throughout the T tubules and surface membrane (17). The present observation that the rate of decline of caffeine-induced Ca²⁺ release decreases by approximately the same proportion as cell surface area after treatment with formamide is compatible with such a distribution (see below); in this case, the relative ineffectiveness of Ca²⁺ influx via the exchanger to trigger SR Ca²⁺ release might be because the Na⁺/Ca²⁺ exchanger is not concentrated near, or closely juxtaposed to, the ryanodine receptors.