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1 Department of Pharmacology, Georgetown University Medical Center, 3900 Reservoir Road NW, Washington, DC 20057, USA
2 College of Pharmacy, Chungnam National University, Daejeon 305-764, South Korea
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Abstract |
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60% of cells), Rc/p converged on 0.2, while in another population (‘group 2’, 40%), Rc/p had a Gaussian distribution with a mean value of 0.625. The fast central release component of group 2 cells appeared to result from in-focus Ca2+ sparks on activation of ICa. In group 1 cells intracellular membranes associated with t-tubular structures were never seen using short exposures to membrane dyes. In most of the group 2 cells, a faint intracellular membrane staining was observed. Quantification of caffeine-releasable Ca2+ pools consistently showed larger central Ca2+ stores in group 2 and larger peripheral stores in group 1 cells. The Rc/p was larger at more positive and negative voltages in group 1 cells. In contrast, in group 2 cells, the Rc/p was constant at all voltages. In group 1 cells the gain of peripheral Ca2+ release sites (
[Ca2+]/ICa) was larger at –30 than at +20 mV, but significantly dampened at the central sites. On the other hand, the gains of peripheral and central Ca2+ releases in group 2 cells showed no voltage dependence. Surprisingly, the voltage dependence of the fast central release component was bell-shaped and similar to that of ICa in both cell groups. Removal of extracellular Ca2+ or application of Ni2+ (5 mM) suppressed equally ICa and Ca2+ release from the central release sites at +60 mV. Depolarization to +100 mV, where ICa is absent and the Na+–Ca2+ exchanger (NCX) acts in reverse mode, did not trigger the fast central Ca2+ releases in either group, but brief reduction of [Na+]o to levels equivalent to [Na+]i facilitated fast peripheral and central Ca2+ releases in group 2 myocytes, but not in group 1 myocytes. In group 2 cells, long-lasting (> 1 min) exposures to caffeine (10 mM) or ryanodine (20 µM) significantly suppressed ICa-triggered central and peripheral Ca2+ releases. Our data suggest significant diversity of local Ca2+ signalling in rat atrial myocytes. In one group, ICa-triggered peripheral Ca2+ release propagates into the interior triggering central Ca2+ release with significant delay. In a second group of myocytes ICa triggers a significant number of central sites as rapidly and effectively as the peripheral sites, thereby producing more synchronized Ca2+ releases throughout the myocytes. The possible presence of vestigial t-tubules and larger Ca2+ content of central sarcoplasmic reticulum (SR) in group 2 cells may be responsible for the rapid and strong activation of central release of Ca2+ in this subset of atrial myocytes.
(Received 8 June 2005;
accepted after revision 13 July 2005;
first published online 14 July 2005)
Corresponding author M. Morad: Department of Pharmacology, Georgetown University Medical Center, 3900 Reservoir Road NW, Washington, DC 20057, USA. Email: moradm{at}georgetown.edu
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Introduction |
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In atrial myocytes RyRs are also found in extended junctional or corbular SR throughout the central regions of the cell with no associated DHPRs as the t-tubular system is either absent or poorly developed (Sommer & Jennings, 1992; Carl et al. 1995). The mode of gating and regulatory mechanisms of this set of RyRs is not fully understood. It has been proposed that Ca2+ release initiated at the peripheral junctional sites of an atrial myocyte might propagate into the interior of the cell by local diffusion of Ca2+ from the peripheral to more central release sites by saltatory conduction (Berlin, 1995; Hüser et al. 1996; Mackenzie et al. 2001; Kockskämper et al. 2001; Woo et al. 2002). In rat, it has been reported that a subset of atrial myocytes have transverse or longitudinal tubular structures in the cell interior, though poorly developed when compared to that of ventricular myocytes (Forssmann & Girardier, 1970; Kirk et al. 2003). In such rat atrial cells, functional importance of longitudinal, centrally located membrane structures was implied from the simultaneous occurrence of centrally located Ca2+ sparks measured in the transverse line-scan mode following field stimulation (Kirk et al. 2003). Using rapid two-dimensional (2-D) confocal microscopy in voltage-clamped rat atrial myocytes, we have already shown that peripheral Ca2+ release in rat atrial cells is composed of focal Ca2+ releases that are under the direct control of ICa (Woo et al. 2002). In the centre of the cell, on the other hand, both fast (Ca2+-buffer-resistant) and slow (buffer-sensitive) components of ICa-dependent local Ca2+ transients were found (Woo et al. 2002).
Here, using rapid 2-D confocal microscopy, we have characterized the fast component of central Ca2+ release in voltage-clamped rat atrial myocytes where the diffusion-dependent slower Ca2+ release is minimized by Ca2+ buffers. We found that a subset of atrial myocytes have a much higher level of rapid central Ca2+ release following depolarization. Such cells tend to show faint intracellular membrane staining possibly related to vestigial t-tubular-like structures, which may mediate the depolarization-induced release of Ca2+ from the central SR. Our findings may provide the physiological basis for the intriguing observations that atrial contraction develops faster than the ventricular myocytes (Lüss et al. 1999), where the dyadic Ca2+ release sites of t-tubules are abundant and well organized throughout the myocytes (Shacklock et al. 1995; Cleemann et al. 1998).
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Methods |
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Rat atrial myocytes were enzymatically isolated from male Wistar rats (WKY, 200–300 g) as previously described (Woo et al. 2002). Briefly, rats were deeply anaesthetized with sodium pentobarbital (150 mg kg–1, I.P.), the chest cavity was opened and hearts were excised. This surgical procedure was carried out in accordance with national and institutional ethical guidelines. The excised hearts were retrogradely perfused at 7 ml min–1 through the aorta, first for 5 min with Ca2+-free Tyrode solution composed of (mM): 137 NaCl, 5.4 KCl, 10 Hepes, 1 MgCl2, 10 glucose, pH 7.3, at 37°C and then with Ca2+-free Tyrode solution containing collagenase (1.4 mg ml–1) and protease (0.16 mg ml–1) for 12 min, and finally with Tyrode solution containing 0.2 mM CaCl2 for 6 min. The atria of the digested heart were then cut into several sections and subjected to gentle agitation to dissociate the cells. The freshly dissociated cells were stored at room temperature in Tyrode solution containing 0.2 mM CaCl2.
Measurement of membrane currents
Myocytes were whole-cell clamped (Hamill et al. 1981) with patch pipettes (tip resistance 2.5–3.5 M
) and dialysed with a Cs+-rich solution (see below) containing 1 mM fluo-3 and 2 mM EGTA. cAMP was added to the pipette solution to enhance ICa and prevent rundown during long (30–40 min) experimental times. Membrane currents were measured with a DAGAN (model 8900, Dagan Co., Minneapolis, MN, USA) patch-clamp amplifier. Generation of voltage-clamp protocol and acquisition of data were carried out using pCLAMP software (version 5.5-1; Axon Instruments, Inc., Foster City, CA, USA). The current signals were filtered at 10 kHz before digitization and storage.
Two-dimensional confocal imaging and image analysis
Cells were loaded with the Ca2+ indicator dye fluo-3 via the patch pipette (see above) and were imaged using a Noran Odyssey XL rapid 2-D laser scanning confocal microscopy system (Noran Instruments, Madison, WI, USA) attached to a Zeiss Axiovert TV135 inverted microscope fitted with a x40 water-immersion objective lens (Zeiss, 440052 C-Apochromat, NA 1.2). The excitation wavelength of the argon ion laser was set to 488 nm (Omnichrome), and fluorescence emission (wavelengths greater than 510 nm) was detected by a high-efficiency photomultiplier tube (Hamamatsu, Middlesex, NJ, USA). The y direction (vertical direction in the figures) was scanned at 240 Hz to produce frames with 225 pixels x 90 pixels sampled on a square, 0.207 µm grid. The confocal slit, stretching in the x direction, was set to values corresponding to a width of 0.6 µm in the confocal plane of the objective. The data were acquired by the Intervision program in a workstation computer (IRIX-operating system, Indy, Silicon Graphics) and were analysed with Intervision software and our own computer program written in Visual Basic 6.0 (Microsoft).
Fluorescence measurement was carried out following 6–7 min after rupture of the membrane with the patch pipette. After this period of dialysis, the intracellular fluo-3 concentration was typically at equilibrium throughout the atrial cells (data not shown). Approximately 3 min after rupture of the membrane 10 conditioning voltage pulses from –90 to –10 mV were applied at 0.1 Hz to maintain the Ca2+ load of the SR. To reduce photobleaching of the dyes and possible phototoxic effects to the cells, the laser was electronically shuttered and triggered to open by the command of the patch-clamp program (pCLAMP) only during the data acquisition period. The average resting fluorescence intensity (F0) was calculated from several frames measured immediately before depolarization. We show raw confocal Ca2+ images in Figs 1, 2 and 11. Ca2+ fluorescence images in some figures (Figs 3, 4, 7A, 7D, 8A, 8D and 10A) were quantitatively illustrated by the sequences of frames showing the change in fluorescence (F), such that the fluorescence images were obtained by subtracting the resting average fluorescence (F0) from the raw images (F). The subtracted images were filtered by 3 pixel x 3 pixel averaging. This method of normalization was used to show Ca2+ sparks and local Ca2+ changes as clearly as possible without resorting to the use of contrast enhancement. We also relied, in part, on a colour scale that presents total fluorescence (F) with a large dynamic range. Tracings of local Ca2+ transients were measured from subcellular regions indicated as coloured masks (see online figures) in the computer program (‘Con2’), and were shown as the average fluorescence of each frame normalized relative to the average resting fluorescence (F/F0). This type of normalization was used to compare results from different cells. The area up to 1.5 µm immediately underneath the cell membrane was denoted as the peripheral region (Woo et al. 2002).
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In some experiments the measurements of Ca2+-induced fluo-3 fluorescence were followed by examination of surface and t-tubular membrane structures in the same myocyte, labelled with the fluorescent indicator dyes, di-2-ANEPEQ or di-4-ANEPPS (Molecular Probes). Effective staining required superfusion with 4 µM di-2-ANEPEQ for 3–5 min or with 10 µM di-4-ANEPPS for 5 min.
Solutions and data analysis
The external solution used for cellular equilibrium and formation of the gigaseal contained (mM): 137 NaCl, 5.4 KCl, 2 CaCl2, 10 Hepes, 1 MgCl2, 10 glucose, buffered to pH 7.4 with NaOH. Patch pipettes were filled with solutions containing (mM): 110 CsOH, 110 aspartic acid, 5 NaCl, 20 TEACl, 20 Hepes, 5 Mg-ATP, 0.2 cAMP, 1 K5fluo-3 (Molecular Probes Inc.), 2 EGTA, with the pH adjusted to 7.2 with CsOH. At 2 min following rupture of the membrane, myocytes were superfused with K+-free Tyrode solution containing 10–30 µM tetrodotoxin and 5 mM Ca2+ to eliminate K+ and Na+ current, and to load SR. Drugs were dissolved in the external experimental solutions, and applied rapidly using a concentration-clamp device (Cleemann & Morad, 1991). All experiments were carried out at room temperature (22–24°C). Numerical results are given as ‘means ± S.E.M. (n =)’, where S.E.M. is the standard error of the mean and n is the number of cells. One-way analysis of variance (ANOVA) was used to verify statistical significance with P < 0.05 taken as significant.
Precautions
Before a confocal plane was fixed for measurements, whole-cell images were routinely monitored in a vertical (z) direction from the top to the bottom of a cell (average thickness: 11.1 ± 0.72 µm, n
= 13) to determine the middle of the cell (defined as the image with sharp edges and most prominent nucleus). Currents were neither leak nor capacitance subtracted (to obtain high quality data and to control for myocyte viability), but the series resistance (1.5–3 times the pipette resistance) was electronically compensated (90%). Imaging of the myocytes in 2-D required 6–7 min of dye dialysis. Only cells with low leak current and clear edges were included in the final analysis of the results.
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Results |
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Figure 1A and C shows sequential 2-D rapid confocal images (240 Hz) of Ca2+ release triggered by the activation of L-type Ca2+ current (ICa), obtained from the mid-sections of two representative rat atrial myocytes. Myocytes were dialysed with internal solution containing 2 mM EGTA plus 1 mM fluo-3, which normally suppressed the delayed slow component of central Ca2+ releases on activation of ICa (Woo et al. 2002). Under these conditions a depolarizing pulse to +20 mV showed two different patterns of Ca2+ release among atrial myocytes. In one population of atrial myocytes (Fig. 1A and B) full activation of ICa (Fig. 1B, upper traces) triggered strong Ca2+ release in the cell periphery (Fig. 1A, see 4 ms image), which then induced a smaller rise of Ca2+ in the centre of myocyte. In the second population of myocytes the central release of Ca2+ occurred as early and as strongly as the peripheral Ca2+ releases (Fig. 1C, see 4 ms image). The peripheral and central Ca2+ releases were quantified from the series of confocal Ca2+ images (Fig. 1B and D, lower traces), and the ratio (Rc/p) of the magnitude of central to peripheral release was estimated. The Rc/p at +20 mV, measured in a total of 122 atrial myocytes, showed bimodal distributions, such that Rc/p of 71 atrial myocytes (57% of the myocytes, Rc/p
= 0.45, ‘group 1’) converged to values close to 0.2. In another population of cells (51 or 43% of the myocytes, Rc/p > 0.45, ‘group 2’) Rc/p had a Gaussian distribution with mean value of 0.625 (Fig. 1E). The data suggest that Ca2+-buffered rat atrial myocytes may exist in two subsets based on their local Ca2+ signalling patterns.
There was no significant difference in inactivation kinetics or peak amplitude of the ICa between the two groups of myocytes (Fig. 1B and D; Table 1). Table 1 describes in more detail the Ca2+ signalling parameters of each group of atrial myocytes. Interestingly the membrane capacitance of group 2 myocytes was bigger than that of group 1 cells.
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Although we were careful to place the focal plane at the central plane of the myocyte along the z-axis, one could argue that the peripheral release events occurring near the surface membrane (above or below the focal plane) might complicate imaging in a subset of cells. To test for such a possibility, we measured Ca2+ releases triggered by the same voltage pulse at two different depths along the z dimension, i.e. at the middle and 2 µm above the middle planes of group 2 myocytes. In the mid-plane of the myocyte, depolarization from –60 to +80 mV activated a central Ca2+ release that was larger than at the periphery (Fig. 2A and C, box 1). In contrast, 2 µm above the mid-plane of the myocyte, where the edges of the cell were less distinct, depolarization now produced equally weak signals at the midline or the edges of the myocyte (Fig. 2B and D, box 1), inconsistent with the notion that the Ca2+ release sites imaged at the central plane resulted from the reflection of those occurring at the surface. Similar relationships held following repolarization from +80 mV when the larger and faster Ca2+ releases were triggered by the large and rapidly inactivating ICa tail currents (cf. Fig. 10C). In the central plane of the cell, the peripheral Ca2+ release was now larger than at mid-line (Fig. 2A and C, box 2), while at the cell surface equally large Ca2+ releases were recorded medially and peripherally (Fig. 2B and D, box 2). Thus we consistently found that Ca2+ releases were similar at the upper and lateral aspects of the cell membrane, and that the distinct properties of the central Ca2+ release stood out most clearly when confocal measurements were performed along a central line in the mid-plane of the cell. A statistical comparison of parameters of central Ca2+ sparks detected from the two different focal planes shows that central sparks detected from the middle focal plane were significantly brighter (larger amplitude) and bigger (size) with similar full width at half amplitude (FWHA) and release times compared to the sparks measured close to the cell surface (Table 2). This result supports the idea that the depolarization-induced central sparks originate from the cell interior.
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We examined whether membrane invaginations imaged using membrane potential dyes were more prevalent in the group 2 subset of atrial cells that generate the fast central Ca2+ release component. In such experiments, following the measurement of depolarization-triggered Ca2+ releases (using fluo-3), the atrial myocyte was stained with di-2-ANEPEQ or di-4-ANEPPS dyes. Among 15 rat atrial myocytes stained with di-2-ANEPEQ or di-4-ANEPPS, six cells showed large Rc/p > 0.45 (group 2). Among the six cells with strong central Ca2+ release, one cell showed little or no indications of intracellular membrane staining. Five cells, however, did have very weak central fluorescence signals (Fig. 3C, ‘group 2’). None of the group 1 cells (n = 9) showed any membrane staining in the cell interior (Fig. 3C, ‘group 1’). The data suggest that there was 93% correlation between the intracellular membrane staining and depolarization-induced focal Ca2+ releases in the cell interior. Note, however, that we did not find exact colocalizations of the membrane fluorescence signals with the central Ca2+ release sites in group 2 myocytes (symbols in Fig. 3D). The result suggests that the distinct Ca2+ signalling patterns observed in the two groups of rat atrial myocytes may be correlated with differential staining of intracellular structures.
Voltage dependence of local Ca2+ releases in group 1 and group 2 atrial myocytes
Figure 4 displays three series of 2-D confocal Ca2+ images recorded from two representative atrial myocytes (one from group 1 and the other from group 2) on depolarizing the cell to –30, +60 or +100 mV from a holding potential of –60 mV (see Fig. 5A for simultaneously measured currents). The first confocal image in each set, represents resting Ca2+ at –60 mV and had low levels of fluorescence. Depolarization induced two types of Ca2+ signalling patterns in the two cell types. In group 1 cells (left panels) Ca2+ release appeared to be more confined to the cell periphery with little propagation into the cell interior, whereas in the group 2 cells (right panels) well-defined central Ca2+ release sites activated as early as, or prior to, the activation of peripheral release sites. It may be noted that at –30 mV a few focal release sites were also activated early in group 1 cells, although they were mostly quiescent at +60 mV (Fig. 4A and B, left panels). Clamp pulses to +100 mV, approaching the Ca2+ reversal potential, failed to trigger significant Ca2+ release in either group 1 or group 2 cells (Fig. 4C).
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When the ‘gain of Ca2+ release’ (ratio of the magnitude of Ca2+ release, measured at 12 ms after the depolarization relative to the peak ICa; Wier et al. 1994; Adachi-Akahane et al. 1996) was compared in the two groups of atrial myocytes, group 1 cells showed significantly higher gain of peripheral Ca2+ release compared to group 2 cells (Fig. 6). The peripheral gain in group 1 cells was larger at –30 mV than at +20 mV (Fig. 6, left panel, n = 8), which is similar to the voltage dependence of gain reported for ventricular myocytes (Wier et al. 1994; Adachi-Akahane et al. 1996). Central gain in group 1 cells also showed a similar tendency of voltage dependence as the peripheral sites. This is consistent with the notion that the small central Ca2+ increase in group 1 cells may reflect the diffusion of peripherally released Ca2+. In contrast, in group 2 cells, there was no significant difference in the peripheral or central gains at –30 and +20 mV (Fig. 6, right panel, n = 7). The result suggests a regional difference in functional couplings between the Ca2+ channels and the Ca2+ release sites in the two groups of myocytes. In addition, the data indicate that the coupling between the central sites and Ca2+ channels in group 2 cells may not be as tight as peripheral couplings in group 1 cells.
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We tested whether influx of extracellular Ca2+ is required to trigger the fast central focal Ca2+ releases in highly Ca2+-buffered group 2 cells. Figure 7A compares the fast and brief removal of extracellular Ca2+ in a myocyte step depolarized to +60 mV for 80 ms. Peripheral and central Ca2+ sparks activated within 4 ms of depolarization in the presence of external Ca2+ (Fig. 7A, Control) were completely suppressed within 1 s of removal of Ca2+ (Fig. 7A, Zero Cao), as was the fast central and peripheral component of the Ca2+ transient (Fig. 7C, 0 Cao). In a total of nine cells examined, removal of extracellular Ca2+ almost completely suppressed ICa (Fig. 7B) and the central and peripheral local Ca2+ releases, as well as occurrence of sparks at +60 mV, suggesting that Ca2+ influx during the depolarization plays a critical role in the activation of the fast central Ca2+ releases.
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Effect of removal of extracellular Na+ on local Ca2+ releases
To examine whether Ca2+ influx on NCX may contribute to the activation of the fast central Ca2+ release sites we examined if: (1) larger depolarizations (+100 mV) could trigger Ca2+ releases, and (2) rapid and short reductions of extracellular Na+ could potentiate Ca2+ release. In both group 1 and group 2 cells depolarization to +100 mV normally failed to induce Ca2+ releases either in the centre or the periphery of myocytes (upper series of images in Fig. 8A and D). Using an electronically controlled solution switcher, the extracellular Na+ concentration was lowered to 5 mM by sucrose substitution 1 s prior to the onset of Ca2+ imaging and depolarization to +100 mV. The low Na+ concentration increased the resting Ca2+ level at –70 mV in the central and peripheral zones of group 1 and group 2 myocytes (compare left and right traces in Fig. 8C and F), but failed to cause a significant sustained increase in the frequency of spontaneous sparks at –70 mV (first image in the lower series of images in Fig. 8A and D). Note, however, that when cells exposed to low [Na+]o for 1 s were subsequently voltage clamped to +100 mV, large Ca2+ releases were consistently observed only in group 2 cells both at the periphery and at the centre (Fig. 8D, last 4 images in the lower images; Fig. 8F, right traces), but not in group 1 cells (Fig. 8A, last 4 lower images; Fig. 8C, right traces). These results suggest that although Ca2+ influx via the NCX at +100 mV is hardly capable of triggering central release sites, the activation of central release sites in group 2 cells may be facilitated by intracellular Ca2+ loading via NCX.
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Recent evidence suggests that SR Ca2+ release is controlled not only by cytosolic Ca2+, but also by Ca2+ in the lumen of the SR, such that a larger luminal Ca2+ concentration appears to lower the threshold for activation of RyRs (Györke & Györke, 1998). To test whether higher SR Ca2+ loading contributes to the rapid activation of central sites on depolarization, we examined the magnitude of SR Ca2+ contents (measured as the amount of local Ca2+ release triggered by rapid application of 10 mM caffeine at –80 mV). Figure 9A and B shows caffeine-triggered Ca2+ transients measured from the periphery and centre of two representative atrial myocytes (group 1, A; group 2, B). We noted significant differences in the local SR Ca2+ loading status between the two groups of cells. Group 1 cells (inset of Fig. 9A; Rc/p
0.2 at +20 mV) had a somewhat larger SR Ca2+ content in the periphery than in the centre (Fig. 8A), while group 2 cells (inset of Fig. 9B; Rc/p
0.67 at +20 mV) had a larger Ca2+ load of central SR as compared to the periphery (Fig. 9B). The ratio of central to peripheral release at depolarizations to +20 mV (Rc/p) did not correlate with the absolute value of peripheral or central Ca2+ content (Fig. 9C), but was dependent on the relative size of the Ca2+ load of central SR as compared to the peripheral SR load in each cell (Fig. 9D). Among 13 cells examined 8 cells had a low Rc/p of 0.25, where central Ca2+ load was consistently smaller than the peripheral Ca2+ load. Five other cells showed a much higher Rc/p, and significantly larger central SR Ca2+ load than the peripheral Ca2+ load. This result suggests that the larger Ca2+ content of the central SR in group 2 cells may in part be responsible for the higher sensitivity of central release sites to Ca2+, thereby causing faster activation.
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In atrial myocytes electrically evoked peripheral and central Ca2+ releases appear to originate primarily from the junctional and non-junctional RyRs, respectively (Hüser et al. 1996; Hatem et al. 1997). We examined whether the depolarization-induced fast Ca2+ releases in the cell centre were mediated by the RyR-gated SR store, using caffeine to directly activate RyRs and deplete the SR Ca2+ stores (Sitsapesan & Williams, 1990; Adachi-Akahane et al. 1996). Preincubation of atrial cells with 10 mM caffeine-containing solutions for 3 min mostly inhibited (86 ± 9%) central or peripheral focal Ca2+ releases induced by the pulse from –80 to +60 mV (Fig. 10A) even though caffeine increased basal Ca2+ signals (Fig. 10C; by 0.27 ± 0.07, n = 9) and ICa was enhanced from 1.32 ± 0.20 to 1.70 ± 0.22 pA pF–1 (n = 9, P < 0.01). Nevertheless in some cells (5 out of 9 cells) pretreated with caffeine, an increase in Ca2+ signal (grey traces in Fig. 10C, or red traces in the online figure), close to the Ca2+ release sites in the centre and periphery, was observed on depolarization (Fig. 10A, see arrowheads). Such weak Ca2+ signals in the presence of caffeine did not appear to be as distinct or bright as focal Ca2+ releases in the control condition, reflecting either Ca2+ influx or Ca2+ release from the incompletely depleted SR.
In another set of experiments, where 20 µM ryanodine was used to inhibit ryanodine receptors, we found almost complete blockade of central and peripheral focal Ca2+ releases at +60 mV in group 2 cells (Fig. 11; n = 3). These results suggest that the rapid central Ca2+ increase on depolarization is related to SR Ca2+ release and that the slight Ca2+ rises in the cell interior in the caffeine-pretreated cell (Fig. 10A) may reflect the release of the remaining SR Ca2+ rather than Ca2+ influx through the invaginated cell membrane.
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Discussion |
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Two subsets of rat atrial myocytes
Depolarization-activated Ca2+ transients in mammalian atrial myocytes have a fast release component triggered by ICa at peripheral junctions and a delayed central component activated by diffusion of Ca2+ to the non-junctional SR (Lipp et al. 1990, 1996; Berlin, 1995; Hüser et al. 1996; Mackenzie et al. 2001; Kockskämper et al. 2001; Woo et al. 2002; Sheehan & Blatter, 2003). Ultrastructural studies in support of this idea show a well-developed non-junctional SR containing electron-dense feet structures (RyRs) in the complete absence (guinea pig: Forbes & Van Neil, 1988; cat: Hüser et al. 1996) or poor development (rat: Ayettey & Navaratnam, 1978; rabbit: Mitcheson et al. 1997) of a t-tubular system. Immunolabelled imaging of RyRs (Carl et al. 1995; Lipp et al. 2000; Mackenzie et al. 2001; Kockskämper et al. 2001) and triadin (Carl et al. 1995) shows ordered sarcomeric distribution throughout rabbit atrial cells, while immunolabelled DHPRs are confined to the surface membrane and are colocalized only with RyRs (Carl et al. 1995; Scriven et al. 2000). In rat atrium, it has been specifically reported that a subset of myocytes have a poorly developed t-tubular structure in the cell interior when compared to ventricular t-tubules (Forssmann & Girardier, 1970; Ayettey & Navaratnam, 1978; Kirk et al. 2003). The punctuate internal membrane staining we report here in type 2 cells (Fig. 3B) was, however, quite different from the centrally located longitudinal membrane staining observed by Kirk et al. (2003). In this context it may be emphasized that the fluorescent lipophilic membrane dyes (ANEPPS, ANEPEQ, etc.), that are intended to stain surface and t-tubular membranes, all have a tendency to penetrate to SR, mitochondria and membranes surrounding and extending from the nucleus given enough time. It should also be noted that narrower longitudinal tubules in rat atrial myocytes may have poor accessibility to the dye producing possibly inhomogeneous staining of such structures. The patterns of internal membrane staining should therefore be cautiously interpreted and, whenever possible, correlated with functional data.
In this context, our functional data showing that a subset of rat atrial cells, in addition to their delayed central Ca2+ release, also have a fast component of Ca2+ release in the cell centre (Fig. 1), are somewhat consistent with the previously observed ultrastructural data on rat atrial myocytes (Forssmann & Girardier, 1970; Ayettey & Navaratnam, 1978; Yamasaki et al. 1997) and line-scan Ca2+ imaging data in field-stimulated rat atrial myocytes (Kirk et al. 2003). Our observation that most of the group 2 cells had rudimentary membrane staining in the cell interior (Fig. 3), while group 1 cells did not show such membrane staining, suggests the possible existence of two groups of rat atrial myocytes, possibly related to differential expression of vestigial t-tubules. In support of such a possibility are also the findings that group 2 cells show: (1) bell-shaped voltage dependence of central Ca2+ release (Fig. 5), (2) the release blockade of the central sites on removal of Ca2+ or application of Ni2+ (Fig. 7), and (3) a significantly larger cell membrane capacitance (73 ± 4.5 pF, n = 51) compared to group 1 cells (59 ± 4.4 pF, n = 71; Table 1).
The degree of differentiation of the t-tubules among the rat atrial myocytes appears to have regional specificity as t-tubules are absent in the specialized nodal and conducting systems (Ayettey & Navaratnam, 1978) but are more developed in left atrium (4 left atria, 10 of 15 cells; 7 right atria, 12 of 30 cells; Kirk et al. 2003). On the other hand, although t-tubular structures are expressed in a subset of rat atrial myocytes, it is unclear whether they have similar dyadic junctions and distributions to those of the surface membrane. In this regard, in group 2 cells: (1) we rarely found colocalization of central release sites with the faint membrane-staining signals (Fig. 3D), (2) central release sites were less sensitive to short exposure to extracellular Ni2+ (Fig. 7D), (3) with short exposures to 5 mM Na+ there was strong depolarization-induced Ca2+ release at +100 mV, but not in group 1 cells (Fig. 8), and (4) the gain of ICa-induced Ca2+ release did not display the voltage-dependent characteristics of ventricular or type 1 atrial myocytes (Fig. 6, Wier et al. 1994; Adachi-Akahane et al. 1996). It is difficult to explain how the removal of Na+ for 1 s would be more effective in depleting the Na+ within the t-tubules as compared to the surface of myocytes, especially since the reported vestigial t-tubules are much narrower in rat atrial myocytes (longitudinal tubules of 67 nm in diameter; Ayettey & Navaratnam, 1978) than the t-tubules of ventricular myocytes (130 nm in diameter; Ayettey & Navaratnam, 1978), yet cause larger central Ca2+ release signals in group 2 cells (Fig. 8).
In the present study we observed that Rc/p does not depend strongly on the membrane potential (Fig. 5E). In fact Rc/p in group 2 cells showed no significant change in the range of potentials from –30 to +60 mV. On the other hand, in group 1 cells, Rc/p at –30 or +60 mV showed a modest, but significant increase compared to the value at +20 mV (Fig. 5E). The reason for the voltage dependence of Rc/p only in group 1 is not clear, but it may be related to the diffusion of Ca2+ from the periphery into the centre, which in turn may occur to varying degrees depending on the rate at which Ca2+ released at the periphery is extruded from the cell by NCX or re-sequestered by the SR.
Differences in the voltage-dependent Ca2+ signalling of type 1 and type 2 cells were seen more clearly by measuring the gain of Ca2+-induced Ca2+ release. In rat ventricular cells, the calibrated gain, or amplification factor, was typically 18 at positive or slightly negative potentials, but increased progressively to more than 40 at –30 and –40 mV (Adachi-Akahane et al. 1996). In the present study, the uncalibrated gains ([Ca2+]/ICa) of group 1 cells displayed a similar increase at –30 mV compared to +20 mV (Fig. 6, left panel). Furthermore, this was observed for both the peripheral and central Ca2+ release, consistent with the observation that Rc/p has only minor voltage dependence (Fig. 5E). On this background it is striking to find that the gain of Ca2+ release sites in group 2 cells, either in the centre or at the periphery, shows no sign of increasing at negative potentials (Fig. 6, right panel). These findings, together with those illustrated in Fig. 8, suggest that the release sites both at the centre and at the periphery of group 2 atrial cells are distinctly different from those found at the periphery of group 1 cells, as well as those of ventricular cells. Considering the relative low gain of ICa-gated release in group 2 cells compared to group 1 cells (Fig. 6) and the likelihood that they are more sensitive to Ca2+ entry via NCX (Fig. 8E
versus
C), it seems possible that their RyRs are less tightly coupled to the DHPRs, but instead are posed more advantageously for sensing Ca2+ influx via NCX.
Role of Na+–Ca2+ exchanger and SR Ca2+ in the two groups of atrial myocytes
Our data indicate that in control conditions the reverse mode of NCX cannot trigger Ca2+ release sites in the centre or periphery of atrial myocytes (Figs 5D and 8). Brief removal of extracellular Na+, however, did lead to the release of Ca2+ in the centre of only group 2 myocytes at positive voltages. Decreasing extracellular Na+ to 5 mM may have increased the basal cytosolic Ca2+ levels (Fig. 8C and F), but such elevations in the resting Ca2+ level occurred in both group 1 and group 2 myocytes, making the finding that depolarization to +100 mV caused central Ca2+ release only in the group 2 cells intriguing, especially since the washout of such narrow longitudinal spaces may be diffusion limited when extracellular solutions are rapidly exchanged for 1–2 s. Since Ca2+ current is negligible at +100 mV a possible mediator for Ca2+ entry in the presence of low [Na+]o is the reverse mode of NCX. The use of a 5 mM Na+-containing external solution (comparable to the internal Na+ concentrations, Despa et al. 2004) would be likely to favour the influx of Ca2+ via NCX at the resting potential and even more so on depolarization to +100 mV. What is surprising is that the same intervention (1 s exposure to low [Na+]o) does not appear to activate the peripheral junctional release sites in group 1 cells (Fig. 8A and D). Of course, one may postulate a markedly higher density of NCX proteins in vestigial t-tubules of atrial cells, or conversely a much lower density of NCX protein within the surface membrane. There is little evidence presently to suggest such a differential distribution of NCX proteins in rat atrial myocytes.
We found that the two groups of rat atrial myocytes produced significantly different patterns of local SR Ca2+ contents (Fig. 9). This result may suggest distinct local distributions or densities of Ca2+ regulatory proteins in the two groups of cells. It has been shown that in the rabbit atrial cells lacking t-tubules peripheral couplings are well developed and SR density is higher in the cell periphery than in the centre (Carl et al. 1995). In contrast, in ventricular myocytes the peripheral couplings are less developed compared to the dyadic junctions (Carl et al. 1995). Thus it is possible that a population of rat atrial myocytes expressing t-tubule-like structures (group 2) have fewer peripheral couplings and a higher density of central SR. Such differential SR density could explain the different ratio of central to peripheral SR Ca2+ contents measured in the two groups of rat atrial myocytes (Fig. 9D). In addition, sensitization of central release sites in group 2 cells by the larger central SR Ca2+ may serve as a cofactor in activating Ca2+ releases from central release sites at +100 mV in the presence of low [Na+]-containing extracellular solution (Fig. 8).
Our data clearly show the significant contribution of the central release sites to the fast component of Ca2+ release early in depolarization in a large population of atrial myocytes. The fast (no-delay) release from the cell interior is likely to be responsible for the faster activation of atrial contraction compared to the ventricular contraction (Lüss et al. 1999). The early activation of release sites in the centre of a subset of atrial myocytes, like the peripheral Ca2+ release, is clearly controlled by ICa mostly in a manner similar to the peripheral release, possibly through the vestigial t-tubules. Interestingly, these sites appear to be more sensitive to NCX activity and have larger Ca2+ content. Studies on the modulation of early versus delayed central focal releases in the two cell types might provide further insight into the functional roles of the two atrial cell types in Ca2+ signalling.
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2)), where a
= 3.7 (magnitude), b
= 0.625 (mean Rc/p) and s
= 0.165 (standard deviation).
P < 0.01, centre versus periphery.
