HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS		QUICK SEARCH:   [advanced] Author: Keyword(s): Year:  Vol:  Page:

This Article Abstract Full Text (PDF) All Versions of this Article: 288/5/H2527    most recent 00188.2004v1 Alert me when this article is cited Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in ISI Web of Science Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via HighWire Citing Articles via ISI Web of Science (9) Citing Articles via Google Scholar Google Scholar Articles by Pérez, C. G. Articles by Mejía-Alvarez, R. Search for Related Content PubMed PubMed Citation Articles by Pérez, C. G. Articles by Mejía-Alvarez, R.

Ryanodine receptor function in newborn rat heart

¹Department of Physiology, Stritch School of Medicine, Loyola University Chicago, Maywood, Illinois; ²Departmento de Fisiología, Biofísica y Neurosciencias, Centro de Investigación y de Estudios Avanzados del Instituto Politécnico Nacional, Mexico City, Mexico; and ³Department of Physiology, Texas Tech University, Lubbock, Texas

Submitted 26 February 2004 ; accepted in final form 11 December 2004

	ABSTRACT

TOP ABSTRACT METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
The role of ryanodine receptor (RyR) in cardiac excitation-contraction (E-C) coupling in newborns (NB) is not completely understood. To determine whether RyR functional properties change during development, we evaluated cellular distribution and functionality of sarcoplasmic reticulum (SR) in NB rats. Sarcomeric arrangement of immunostained SR Ca²⁺-ATPase (SERCA2a) and the presence of sizeable caffeine-induced Ca²⁺ transients demonstrated that functional SR exists in NB. E-C coupling properties were then defined in NB and compared with those in adult rats (AD). Ca²⁺ transients in NB reflected predominantly sarcolemmal Ca²⁺ entry, whereas the RyR-mediated component was 13%. Finally, the RyR density and functional properties at the single-channel level in NB were compared with those in AD. Ligand binding assays revealed that in NB, RyR density can be up to 36% of that found in AD, suggesting that some RyRs do not contribute to the Ca²⁺ transient. To test the hypothesis that RyR functional properties change during development, we incorporated single RyRs into lipid bilayers. Our results show that permeation and gating kinetics of NB RyRs are identical to those of AD. Also, endogenous ligands had similar effects on NB and AD RyRs: sigmoidal Ca²⁺ dependence, stronger Mg²⁺-induced inhibition at low cytoplasmic Ca²⁺ concentrations, comparable ATP-activating potency, and caffeine sensitivity. These observations indicate that NB rat heart contains fully functional RyRs and that the smaller contribution of RyR-mediated Ca²⁺ release to the intracellular Ca²⁺ transient in NB is not due to different single RyR channel properties or to the absence of functional intracellular Ca²⁺ stores.

Ca²⁺ release; sarcoplasmic reticulum; excitation-contraction coupling

In this work, developmental changes in the cellular distribution of the SR were defined by immunostaining SR Ca²⁺-ATPase (SERCA2a) in NB and AD cells. Intracellular Ca²⁺ transients were examined using confocal microscopy in acutely isolated NB rat ventricular myocytes. The density of RyR protein was defined with ryanodine binding assays in both SR microsomes and whole heart homogenate from AD and NB rat cardiac muscle. The single-channel functional properties of NB rat RyR were also determined for the first time in planar lipid bilayer studies and were compared with those of AD. Our results indicate that in acutely isolated myocytes from the NB rat heart, the contribution of RyR-mediated CICR to the intracellular Ca²⁺ transient is negligible. This is interesting, because there are sizeable amounts of RyR protein present in the NB cell, and at least some of this protein forms RyR channels that release Ca²⁺ from intracellular stores when stimulated by caffeine. Furthermore, single-channel studies demonstrate that RyR channels in AD and NB cells have nearly identical permeation properties, gating kinetics, and ligand regulation characteristics. Thus the NB cells have functional Ca²⁺ stores containing working AD-like RyR channels, yet these stores do not contribute substantially to the intracellular Ca²⁺ transient during E-C coupling.

	METHODS

TOP ABSTRACT METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
Isolation of ventricular myocytes. Ventricular myocytes were enzymatically isolated from hearts of NB (1–5 days old) and AD (8 wk old) Sprague-Dawley rats of either sex. Cell isolation was conducted in accordance with Institutional Animal Care and Use Committee guidelines. Rats were injected intraperitoneally with a mixture of pentobarbital sodium (10 mg/g body wt) and heparin (2.2 U/g body wt). After 10 min, the heart was quickly removed by thoracotomy and rinsed in normal Tyrode solution (in mM: 100 NaCl, 10 KCl, 4 MgSO₄, 50 taurine, 20 glucose, 2 CaCl₂, and 10 HEPES, pH 7.4 adjusted with NaOH) at 37°C. The aorta was cannulated and connected to a Langendorff perfusion apparatus. The heart was then continuously perfused with Ca²⁺-free Tyrode solution for 10 min, followed by a variable perfusion time (5 min for NB and 10 min for AD) with enzymatic solution [0.5 mg/ml collagenase (Yakult) for NB; 1.28 mg/ml collagenase I (Sigma, St. Louis, MO) and 0.12 mg/ml protease XIV (Sigma) for AD]. The heart was then perfused with Kraftbrühe (KB) medium [in mM: 85 KCl, 30 KH₂PO₄, 5 MgSO₄, 5 Na₂ATP, 5 pyruvate, 5 D,L--hydroxybutyrate, 5 creatine, 20 taurine, 20 glucose, 50 g/l polyvinylpyrrolidone (PVP-40), and 1 EGTA, pH 7.2 adjusted with KOH] for 10 min. After washout of the enzyme, both ventricles were separated from the atria and then minced. Single myocytes were obtained by gentle dispersion with a wide-bore pipette and then filtered through a cell strainer (40-µm Nylon). The entire isolation procedure was carried out with solutions equilibrated with 100% O₂ at 37°C. Isolated cells were incubated for at least 30 min in KB medium at 4°C, placed in normal Tyrode solution, and kept at room temperature (RT; 21–23°C) until used. This procedure consistently yielded healthy and Ca²⁺-tolerant NB and AD myocytes. Only quiescent myocytes with clearly visible striations were used for fluorescence experiments.

Immunofluorescence labeling. Double immunofluorescence labeling was conducted on ventricular myocytes from NB (2 days old) and AD rat hearts. Mouse and rabbit monoclonal antibodies directed against SERCA2a (Affinity Bioreagents, CO) and -actinin (Sigma), respectively, were used. Acutely isolated myocytes were fixed for 10 min in 3% buffered paraformaldehyde solution, washed with PBS twice, and permeabilized for 10 min with a PBS solution containing 0.1% Triton X-100. The cells were incubated overnight with the primary antibodies, both at a dilution of 1:250, and were exposed for 2 h to fluorophore-conjugated secondary antibodies (Texas red anti-mouse IgG for SERCA2a and Alexa 488 anti-rabbit IgG for -actinin; Molecular Probes, Eugene, OR) at a dilution of 1:200. To determine nonspecific labeling, we also conducted control experiments without the primary antibody (data not shown). SERCA2a and -actinin immunoreactivity was determined with a Radiance 2000 MP laser scanning confocal microscope (Bio-Rad, Hercules, CA) equipped with a x40 oil immersion objective lens (Plan Fluor, numerical aperture = 1.3; Nikon). The excitation light was generated using two laser sources: green helium-neon (543 nm) and argon ion (488 nm). Emitted fluorescence was sequentially collected by a pair of photomultiplier tubes in bands between 515–530 nm (for Alexa 488) and 600–650 nm (for Texas red). A Kalman averaging filter (N = 3) was used to reduce the noise. Images (1,024 x 1,024 pixels) were acquired in XY frame-scan mode. Zoom values of 4.6 and 10 (pixel sizes of 60 and 30 nm) were used for NB and AD, respectively. A theoretical maximal resolution of 230 nm was estimated for these conditions (numerical aperture and excitation wavelengths). Thus our sampling rates were at least twofold greater than the Nyquist rate (115 nm).

Image analysis. Data were analyzed from 8-bit/pixel RGB false-color images. Alexa 488 labeling was displayed on the red channel, and Texas red signal was shown on the green channel. To estimate the degree of SERCA2a/-actinin colocalization, we conducted a standard particle analysis with ImageJ 1.31 software (NIH). Background fluorescence of each image (pixels with fluorescence intensity below a threshold value of 25%) was subtracted. Only particles larger than 3 pixels and smaller than 1,000 pixels were considered. Distribution of the sampled particles was arrayed in squared matrixes (Microsoft Excel 2002; Redmond, WA), and the interparticle distance was measured. The distance to the nearest neighboring particle was determined for each particle and plotted in a frequency distribution histogram. The extent of colocalization was estimated from the corresponding cumulative probability histogram. The statistical significance (P) of the developmental differences of SERCA2a/-actinin colocalization and the maximal difference (D) were determined using the Kolmogorov-Smirnov test.

Intracellular [Ca²⁺] measurements. Single myocytes were incubated in 20 µM fluo-4 AM (Molecular Probes) at RT. Adequate fluorescence levels were obtained with loading periods of 30 min for AD cells and 45 min for NB cells. Signs of cell damage (i.e., contracture, cytoplasmic granules, or membrane blebs) appeared when longer incubation times were used. The incubation period was ended by rinsing the cells with dye-free normal Tyrode solution at RT for 15 min. Loaded myocytes were then placed on the stage of an inverted laser scanning confocal microscope (LSM 410; Carl Zeiss) equipped with a x40 oil-immersion objective lens (Plan-Neofluar, numerical aperture = 1.3; Zeiss). Fluo-4 was excited with the 488-nm line of an argon laser. Emitted fluorescence was collected through a 515-nm long-pass emission filter. Fluo-4 images were recorded in line-scan mode with 512 pixels per line at 250 Hz. Action potentials (APs) were triggered by field stimulation with the use of platinum electrodes embedded in the experimental chamber. All AP-stimulated intracellular Ca²⁺ transients were recorded in normal Tyrode solution at RT. Only myocytes that exhibited vigorous contractility in response to field stimulation were used in this study. Ca²⁺ transient image files were analyzed with ImageJ 1.3 software (NIH).

SR microsomes preparation. SR microsomes were obtained from NB (1 day old) and AD (200 g) rat ventricles as previously described (33). Briefly, rats were anesthetized with intraperitoneal injection of pentobarbital sodium (10 mg/g body wt). Hearts were quickly removed by thoracotomy and rinsed in Ca²⁺-free Tyrode solution. After the atria and all visible vascular tissues were trimmed off, the ventricles were immersed in a solution containing protease inhibitors (in µM: 0.1 aprotinin, 500 benzamidine, 1 leupeptin, 1 pepstatin A, and 200 PMSF) at 4°C. These protease inhibitors were used throughout the entire microsome preparation procedure. The ventricles were homogenized (3 times for 30 s) with a Polytron homogenizer at low speed in ice-cold saline solution (in mM: 154 NaCl and 10 Trizma maleate, pH 6.8). The resulting homogenate was then centrifuged for 20 min at 4,000 g. The 4,000-g supernatant was filtered through cheesecloth and spun again for 20 min at 8,000 g. The 8,000-g supernatant was spun in an ultracentrifuge at 100,000 g for 40 min. The 100,000-g pellet, which contained SR microsomes, was resuspended in a storage solution (in mM: 154 NaCl, 10 Trizma maleate, and 300 sucrose, pH 6.8), flash frozen, and stored at –80°C until used. The protein concentration of all NB and AD SR microsome preparations was determined using the Lowry method.

Ryanodine binding. Ryanodine binding was quantified in SR microsome preparations from NB (1 day old) and AD hearts and from AD skeletal muscle. The AD skeletal muscle was used as a positive control. The binding assays were carried out by incubating the SR microsomes (1 mg of total protein) in 150 µl of reaction mixture containing 100 mM KCl, 20 mM HEPES, and 60 nM [³H]ryanodine (56.9 Ci/mmol). In addition, specific ryanodine and dihydropyridine binding was measured in NB and AD ventricle (whole tissue) homogenate. The protein concentration in the whole tissue homogenate was 10 mg/ml. In these assays, the 150-µl reaction mixture contained either 1 M KCl, 100 µM free Ca²⁺, 20 mM HEPES, and 0–100 nM [³H]ryanodine or 25 mM Tris, 10 mM Na-HEPES, 1 mM EDTA, 1.1 MgCl₂, and 15 nM [³H]PN200-110. All the binding assays were conducted in the presence of protease inhibitors (0.1 mg/ml leupeptin and 0.1 mg/ml aprotinin). Nonspecific binding was measured using either 37 µM cold ryanodine or 16.6 µM cold nifedipine. The incubation lasted for 90 min and was carried out at 37°C. To define the Ca²⁺ dependence of ryanodine binding, we repeated the assay at different free [Ca²⁺] (10^–7 to 10^–3 M). The cardiac Ca²⁺ dependence data were fitted with Eq. 1,

(1)

where B_max corresponds to the maximal number of binding sites, K_a represents half-maximal activation, and n_H is the Hill coefficient. The skeletal Ca²⁺ dependence data were fitted with Eq. 2,

(2)

where K_i represents half-maximal inactivation and n_H is the Hill coefficient of the inactivation component of the equation.

Planar lipid bilayers. Single SR Ca²⁺ release channels were reconstituted into artificial planar lipid bilayers by fusing SR microsomes as previously described (33). Briefly, planar bilayers were formed across a 150-µm-diameter aperture in a Delrin partition. Bilayer-forming solution contained a mixture of phosphatidylethanolamine and phosphatidylcholine (7:3, 50 mg/ml decane; Avanti Polar Lipids, Pelham, AL). SR microsomes were added to one side of the bilayer (cis, cytoplasmic) (51). Membrane potential was controlled in the cis compartment by using a conventional patch-clamp amplifier (Axopatch 200B; Axon Instruments, Burlingame, CA), while the other side of the bilayer (trans) was connected to the ground of the amplifier. The current signal was digitized at a rate of 10 kHz with a 12-bit analog-to-digital/digital-to-analog converter (Digidata 1200; Axon Instruments), filtered with a Bessel filter at 2 kHz, and stored for later analysis. The recording solutions contained 20 mM HEPES-Tris (pH 7.4), 10 µM added Ca²⁺, and the charge carrier. The charge carrier was either Cs⁺ [50/250 mM (cis/trans) CsCH₃SO₃] or Ca²⁺ [2–30 mM trans Ca(CH₃SO₃)₂]. Free [Ca²⁺] in the cis chamber was adjusted to various levels (0.1 µM to 1 mM) and was buffered with either EGTA (see Fig. 8) or HEDTA (see Fig. 9). Free [Ca²⁺] in the recording solutions was measured off-line with custom-made Ca²⁺ electrodes (1).

View larger version (24K): [in this window] [in a new window]   Fig. 8. Stationary [Ca2+] dependence of NB and AD RyR channel activity. A: stationary single-channel activity from NB (left) and AD (right) RyR recorded at a holding potential of 0 mV, with 50/250 (cis/trans) mM CsCH3SO3, in the presence of the indicated cytoplasmic (cis) [Ca2+]. Zero-current level is indicated by dotted lines. Openings are shown as downward deflections. B: steady-state [Ca2+] dependence of normalized (to the maximal) Po from NB (n = 4; diamonds) and AD (n = 5; circles) RyRs. Smooth curves were generated using Eq. 4.

View larger version (24K): [in this window] [in a new window]   Fig. 9. Modulation of NB and AD RyR channel activity: ATP and caffeine activation. A: unitary current traces show activation of a single NB RyR by sequential addition of different concentrations (in mM) of ATP and caffeine to the cis side. Channel activity was recorded in the presence of 50/250 (cis/trans) mM CsCH3SO3 and 0.01 µM cytoplasmic (cis) Ca2+. Zero-current level (closed state, C) is indicated by the dotted line. Openings are shown as downward deflections. Bar values show pooled data from 4 individual reconstitutions in which the same sequential activation by ATP and caffeine was induced. Washout condition was equivalent to control (pCa = 7). Caf, caffeine. B: activation of a single AD RyR by sequential addition of different concentrations of ATP and caffeine to the cis side. Bar values show pooled data from 5 individual reconstitutions in which the same sequential activation by ATP and caffeine was induced. Washout condition was equivalent to control (pCa = 7).

Data acquisition, unitary current measurements, statistical analysis, and data processing were performed using commercially available software packages (pCLAMP version 8.0; Axon Instruments, Union City, CA; Microsoft Excel 2002; and Microcal Origin version 7.0; Northampton, MA). Gating kinetics analysis was conducted with custom-made routines written in the LabView version 7.0 programming language (National Instruments, Austin, TX). Averaged data points represent means ± SE. Statistical significance was determined with two-sample paired t-tests. The data presented were obtained from 60 isolated myocytes (10 AD and 50 NB), 9 SR microsome preparations (2 AD and 7 NB), and 97 bilayer incorporations (52 AD and 45 NB). Preliminary versions of this work were previously presented at Annual Biophysical Society Meetings.

	RESULTS

TOP ABSTRACT METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
SERCA2a shows sarcomeric distribution in NB ventricular myocytes. In NB ventricular myocytes, a poorly developed SR has been associated with the absence of a well-formed T-tubular system. For this reason, we examined the level of maturity and structural organization of the SR in relation to the sarcomeric arrangement of the cytoskeleton in NB rat. To this end, SERCA2a and -actinin were immunostained in double-labeling experiments conducted on acutely isolated myocytes from NB (2 days old) and AD rat hearts. Our results indicate that in NB myocytes, the fluorescence signal arising from SERCA2a (Fig. 1A, left) was already strong and well defined, consistent with a high level of SR organization. In addition, this arrangement exhibited a distribution oriented transversely to the longitudinal axis of the cell. Regular spacing of 2 µm between fluorescent bands was evident, corresponding well with the sarcomeric length observed with -actinin staining in the same NB myocyte (Fig. 1A, middle) and in AD myocytes (Fig. 1B, middle). The colocalized pixels (Fig. 1A and B, right, in white) indicate that SERCA2a and -actinin are closely associated in a sarcomeric arrangement. The high degree of colocalization indicates that this transversely striated pattern most likely arises from the Z lines and the junctional SR. The exclusion of SERCA2a and -actinin from the nuclear region of the cell further indicates the sarcomeric distribution of the SR in NB myocytes. In AD myocytes, a clear correlation between SR distribution (Fig. 1B, left) and the sarcomere (Fig. 1B, middle) was also evident as a transversely striated pattern. However, a second pattern of axially oriented stripes was evident only in AD. Because this pattern did not colocalize with -actinin, it could be attributed to the longitudinal free SR. Quantification of the distance between fluorescent particles revealed a significant reduction in colocalization with development (Fig. 1C). This could further indicate the emergence of a more prominent longitudinal SR in AD.

View larger version (75K): [in this window] [in a new window]   Fig. 1. Developmental changes in sarcoplasmic reticulum (SR) distribution. Confocal images of immunostained myocytes show cellular distribution of SR Ca2+-ATPase (SERCA2a; pseudocolored green) and -actinin (pseudocolored red). A: SERCA2a (left) and -actinin (middle) distribution in a 2-day-old rat myocyte. SERCA2a/-actinin colocalization is shown (in white) in the merged image (right). B: SERCA2a (left) and -actinin (middle) staining in an adult (AD) rat myocyte. Areas of colocalization are shown (in white) in the merged image (right). C: bar graph (bottom) shows the distribution of nearest neighbor (interparticle) distances between SERCA2a and -actinin. A cumulative probability histogram (top) of colocalization indicates a significantly different distribution in AD and 2-day-old neonates (Komogorov-Smirnov test, D = 0.11 and P < 0.005).

View larger version (31K): [in this window] [in a new window]   Fig. 2. Characteristics of Ca2+ transients in acutely isolated newborn (NB) ventricle myocytes. A: inhomogeneous distribution of intracellular [Ca2+] in fluo-4-loaded NB myocytes. A color-coded fluorescence intensity (F/F0) image (top) shows a subsarcolemmal ring of elevated [Ca2+] during an action potential (AP)-induced Ca2+ transient in a 5-day-old NB myocyte. A surface plot of the corresponding image is shown at bottom. Trace at right shows transverse fluorescence intensity measured at the line marked with an asterisk. B: a transverse "x-t" line-scan image (top) was obtained during an AP-induced Ca2+ transient in a 4-day-old NB myocyte. Fluorescence intensity measured at the center (C; shaded curve) and subsarcolemma (SS; solid curve) is shown at bottom. C: developmental changes of Ca2+ transient kinetics. Averaged fluorescence values (10 traces) from AD and NB (5 days old) were normalized and overlapped. Time to peak in AD was 125.2 ms, whereas that in NB was 268 ms. Ca2+ transient decay () was fitted with a single-exponential function (blue curves) for both AD ( = 261.8 ms) and NB ( = 271.3 ms). D: pooled data illustrate developmental changes in the Ca2+ transient kinetics. *Statistically different (P < 0.05) from 1-day-old NB.

Because previous studies have demonstrated the existence of RyRs in NB cells, it is possible that at this age, RyR-mediated Ca²⁺ release makes an important contribution to the Ca²⁺ transient. To estimate this component, we recorded Ca²⁺ transients in AD and NB cells in the absence (control) and in the presence of a high dose (1 or 10 µM) of ryanodine (Fig. 3A). The ryanodine application had two different effects: reduction of the peak amplitude of the Ca²⁺ transient and elevation of the resting [Ca²⁺]. In AD cells, peak amplitude was reduced 70% and the resting Ca²⁺ level was substantially elevated. However, in 1-day-old NB, peak amplitude was reduced 13% and the resting Ca²⁺ level was only slightly elevated. This implies that the contribution of RyR-mediated Ca²⁺ release to the Ca²⁺ transient in NB cells is small. This could result from the absence of functional RyR-mediated SR Ca²⁺ stores in NB cells. To directly test this possibility, we applied caffeine to NB cells. Figure 3B shows results of a representative experiment conducted on a 2-day-old NB ventricular myocyte. Figure 3B, top, shows a longitudinal "x-t" line scan, and the trace (Fig. 3B, bottom) shows the corresponding averaged fluorescence intensity. To ensure adequate SR Ca²⁺ load, we first elicited AP-induced Ca²⁺ transients at a frequency of 0.5 Hz. After a 4-s quiescent period, 5 mM caffeine was rapidly applied to the cell. As a result, a transient increase of intracellular [Ca²⁺] was observed. This transient was slightly larger and slower compared with the electrically stimulated transients. Thus the minor contribution of RyR-mediated SR Ca²⁺ release to the electrically stimulated Ca²⁺ transient in NB cells cannot be attributed to the absence of functional RyR Ca²⁺ stores.

View larger version (34K): [in this window] [in a new window]   Fig. 3. Developmental changes in the SR contribution to the Ca2+ transient. A: AP-induced Ca2+ transients recorded before (control) and 15 min after addition of 1 and 10 µM ryanodine in AD and NB (2 day old), respectively. Ryanodine induced an increase in basal fluorescence level (indicated by dotted lines) of 50% in AD and 4.5% in NB and a reduction in transient amplitude of 70% in AD (n = 4) and 13% in NB (n = 6). B: RyR-mediated Ca2+ release in NB (n = 4) is illustrated with a line-scan image (top) and averaged fluorescence of Ca2+ transients (bottom) recorded in a NB (2 day old) myocyte. Ca2+ transients were elicited first with field stimulation, and after a quiescent period, a Ca2+ transient was induced by adding 5 mM caffeine to the bath solution.

View larger version (35K): [in this window] [in a new window]   Fig. 4. Developmental changes in specific [3H]ryanodine binding. A: nonlinear [3H]ryanodine binding (n = 3) to SR microsomes from NB (diamonds) and AD (circles). B: Scatchard analysis of the data shown in A. Ryanodine binding site density (Bmax) was 0.49 pmol/mg protein in NB and 1.35 pmol/mg protein in AD, and the apparent ryanodine binding affinity (Kd) was 32 ± 5 nM in NB and 25 ± 4 nM in AD. C: [Ca2+] dependence of specific [3H]ryanodine binding to SR microsomes obtained from NB heart (1 day old; diamonds), AD heart (circles), and AD skeletal muscle (triangles). Smooth curves were generated using Eq. 1 for heart and Eq. 2 for skeletal muscle. Bmax was 0.46, 0.78, and 0.92 pmol/mg protein for NB heart, AD heart, and AD skeletal muscle, respectively. Half-maximal activation (Ka) was 2.9, 2.0, and 3.4 µM for NB heart, AD heart, and AD skeletal muscle, respectively. Half-maximal inactivation (Ki) for skeletal muscle was 22 µM. The Hill coefficient (nH) was 2, 1.7, and 1.8 for NB heart, AD heart, and AD skeletal muscle, respectively. D: specific [3H]ryanodine (RyR) and [3H]nifedipine (dihydropyridine receptor, DHPR) binding density measured in crude homogenates from NB and AD rat ventricles (n = 5). Ryanodine binding density was 34.7 ± 0.7 and 134.5 ± 4 fmol/mg wet wt in NB and AD, respectively. *Statistically different (P < 0.05) from AD. Nifedipine binding density was 8.7 ± 1 and 20.5 ± 0.8 fmol/mg in NB and AD, respectively. Values for DHPR-to-RyR ratio (right axis) were obtained from the ratio of [3H]nifedipine/[3H]ryanodine binding to the whole heart homogenate. DHPR-to-RyR binding density ratio was 0.25 and 0.15 for NB and AD, respectively.

Reconstitution of single NB RyR Ca²⁺ release channels. Representative single RyR channel activity reconstituted from NB and AD SR microsomes is shown in Fig. 5. These recordings were obtained at –10 mV in a 0.01/10 mM (cis/trans) Ca²⁺ gradient. The NB and AD channels were spontaneously active in the presence of 10 µM Ca²⁺ (Fig. 5, top). In both cases, the corresponding total amplitude histograms indicate that the main current amplitude was 3 pA. In both channels, control gating was characterized by frequent brief openings with few events lasting more than a few milliseconds. The permeation and gating of both the NB and AD channels were clearly modified by the addition of ryanodine (Fig. 5, bottom). Ryanodine induced long-duration opening events to a subconductance level, which is the classic action of ryanodine on a single RyR channel (41). Thus these results positively identify these channels as RyR channels and, more importantly, confirm that the NB SR microsome preparations contain functional RyR channels. The next step was to determine whether the functional properties of single NB and AD RyR channels are similar.

View larger version (38K): [in this window] [in a new window]   Fig. 5. Ryanodine effect on NB and AD single RyR channel activity. Single-channel activity from NB (left) and AD (right) SR microsomes recorded under control conditions [10 µm CaCl2 (cis)/10 mM Ca-HEPES (trans), –10 mV of holding potential] (top) and 15 min after addition of 20 µM ryanodine to both sides of the bilayers (bottom). Opening events are shown as downward deflections. Zero-current level corresponds to the closed state (C) and is indicated by horizontal dotted lines. Total amplitude histograms constructed for each condition are shown beneath recordings. Gaussian functions fitted to total amplitude histograms show a mean amplitude in NB of 2.4 and 1.3 pA before and after ryanodine, respectively, and a mean amplitude in AD of 2.8 and 1.7 pA before and after ryanodine, respectively.

(3)

where F is Faraday’s constant, R is the gas constant, T is temperature, and [Cs⁺]_cis/[Ca²⁺]_trans equals 1. The calculated P_Ca/P_Cs values of the NB and AD RyR channels were similar with values of 10, indicating that both channels were 10-fold more selective for Ca²⁺. Thus the NB preparations not only contain functional RyR channels, but these channels also have permeation properties identical to those of their AD counterparts.

View larger version (39K): [in this window] [in a new window]   Fig. 6. Single-channel permeation properties of NB and AD RyR. A: Cs+ conductance. Single-channel activity from NB (left) and AD (middle) SR microsomes recorded in 50/250 (cis/trans) mM CsCH3SO3 and symmetrical 10 µM Ca2+ at the indicated membrane potentials. Opening events are shown as downward deflections. Zero-current level corresponds to the closed state (C) and is indicated by horizontal dotted lines. A current-voltage relationship (right) was constructed with pooled data from 4 (AD; circles) and 3 (NB; diamonds) experiments. Cs+ conductance values obtained by linear regression are 426 ± 35 and 413 ± 22 pS for NB and AD, respectively. B: Ca2+ conductance. Single-channel activity from NB (left) and AD (middle) SR microsomes recorded at indicated potentials. Unitary current was recorded in 0.01/30 (cis/trans) mM Ca(CH3SO3)2 and symmetrical 20 mM HEPES. A current-voltage relationship (right) was constructed with pooled data from 3 (AD; circles) and 4 (NB; diamonds) experiments. Ca2+ conductance values obtained by linear regression are 102 ± 15 and 98 ± 18 pS for NB and AD, respectively. C: Ca2+/Cs+ selectivity. Single RyR channel activity was recorded from NB and AD SR microsomes with a 2-s voltage ramp (from –50 to +70 mV, 0 mV holding potential). The trans solution contained 30 mM Ca(CH3SO3)2 and 20 mM HEPES (pH 7.4), whereas the cis solution contained 30 mM CsCH3SO3, 20 mM HEPES, and 10 µM CaCl2 (pH 7.4). Cation selectivity was estimated from the apparent reversal potential (ER, indicated by arrows). Zero-current level (closed state) is indicated by horizontal dotted lines. The permeability ratio (PCa/PCs), calculated using Eq. 3, has a value of 9.3 for NB and 10.6 for AD.

View larger version (58K): [in this window] [in a new window]   Fig. 7. Modal gating kinetics of NB and AD RyR. Stationary single RyR channel activity from NB (A) and AD (B) recorded at a holding potential of 0 mV is shown at left. Ionic conditions in both cases were 0.01/30 (cis/trans) mM Ca(CH3SO3)2. Examples of high-open probability (Po) mode opening events are underlined with red lines, and low-Po mode events are underlined with dotted lines. Dwell-time dependency plots from NB and AD are shown at right. Dependency plots were constructed from at least 1 x 106 events by concatenating 10 min of continuous recordings from 4 different bilayers. The red areas in the contour surface correspond to positive shut-open dwell-time correlations, whereas the blue regions illustrate negative correlations.

(4)

The parameter P represents the maximal P_o, K_act corresponds to the activation midpoint, a is the activation coefficient, K_inact represents the inactivation midpoint, and i is the inactivation coefficient. The best fit was obtained for NB when K_act = 1.6 µM, a = 5.5, K_inact = 6.3 mM, and i = 2.2, and for AD when K_act = 3.2 µM, a = 2.3, K_inact = 5 mM, and i = 2.3. Thus the NB and AD RyR channel activity exhibited very similar cytosolic Ca²⁺ sensitivity.

Cytoplasmic ATP is another important endogenous activator of the RyR channel. Figure 9A shows its effect in NB and AD RyR channels in the presence of a low cytosolic [Ca²⁺]. The single RyR channel recordings shown are 10- and 5-s segments obtained from NB and AD, respectively, in each of the sequential (left to right) experimental conditions indicated. The bar values (n = 4) below each recording indicate the P_o in each experimental condition. In control conditions (100 nM cytosolic Ca²⁺), the NB and AD RyR channels exhibited a very low level of activity, consistent with the data presented in Fig. 8B. Additions of progressively larger cytosolic concentrations of ATP (from 0.4 to 2 mM) activated the channels to P_o values of 0.4. This effect was fully reversible when ATP was removed. Addition of caffeine substantially activated the channels as well. Thus there were no appreciable differences in the responses of NB and AD RyR channels to the ATP or caffeine challenges.

The inhibitory action of cytosolic Mg²⁺ also was defined for both the NB and AD RyR channels (Fig. 10). In this study, single RyR channel activity was recorded at 0 mV in the presence of 5 µM cytosolic Ca²⁺ with different cytosolic concentrations of Mg²⁺ (0.1 to 2 mM). Representative NB and AD single-channel recordings are shown (Fig. 10, left). Single RyR channel activity decreased as the cytosolic Mg²⁺ concentration was raised. The presence of cytosolic Mg²⁺ reduced both the P_o and unitary current of the channels. The reduction in current amplitude occurred because Mg²⁺, being permeable, can compete with charge carrier for occupancy of the pore (33). The reduction in P_o occurred because Mg²⁺ directly competes with Ca²⁺ for the activation site of the channel (28, 38). To test this interpretation, we repeated the Mg²⁺-induced inhibition experiments at a higher cytosolic [Ca²⁺] (100 µM). Under these conditions, cytosolic Mg²⁺ still reduced P_o, but higher concentrations were required. These data are summarized in dose-response plots (Fig. 10, right). These plots were constructed from pooled data from four experiments at each [Ca²⁺]. The P_o values were normalized to the maximal value obtained before Mg²⁺ addition. The smooth curves were generated using the following equation:

(5)

View larger version (39K): [in this window] [in a new window]   Fig. 10. Modulation of NB and AD RyR channel activity: Mg2+ inhibition. A: single-channel recordings (left) show the effect of different Mg2+ concentrations on NB RyR channel activity recorded at a holding potential of 0 mV in the presence of either 5 or 100 µM free Ca2+ on the cytoplasmic (cis) side of the channel and 50/250 (cis/trans) mM CsCH3SO3 as charge carrier. A dose-response curve (right) of pooled data from 4 different channel experiments shows the inhibitory effect of Mg2+ in the presence of 5 (diamonds) and 100 µM (circles) cytoplasmic (cis) Ca2+. Smooth curves were generated using Eq. 5. The best fit was obtained at 5 µM Ca2+ when Kd = 256 µM and nH = 2.5. The best fit was obtained at 100 µM Ca2+ when Kd = 1.6 mM and nH = 1.6. B: single-channel recordings (left) show the effect of different Mg2+ concentrations on AD RyR channel activity recorded at a holding potential of 0 mV in the presence of either 5 or 100 µM free Ca2+ on the cytoplasmic (cis) side of the channel. A dose-response curve (right) of pooled data from 3 different channel experiments shows the inhibitory effect of Mg2+ in the presence of 5 (diamonds) or 100 µM (circles) cytoplasmic (cis) Ca2+. The best fit was obtained at 5 µM Ca2+ when Kd = 180 µM and nH = 4.6. The best fit was obtained at 100 µM Ca2+ when Kd = 2.4 mM and nH = 3.3.

	DISCUSSION

TOP ABSTRACT METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
In this study, two novel contributions were presented. First, the developmental changes of the intracellular Ca²⁺ transient, particularly of the RyR-mediated fraction, were defined in acutely isolated ventricular myocytes from the NB rat heart. The originality of this part of the study is that most previous works focusing on this topic have been conducted either in different species [rabbit (18)] or with different experimental preparations [multicellular (12, 49), primary cultures (21, 31), and fetal cells (46)]. A second unique aspect is the multidisciplinary experimental approach used in this study that includes scanning confocal microscopy of the SR cellular distribution and RyR-mediated Ca²⁺ release, radiolabeled ryanodine and dihydropyridine (DHP) binding, and single RyR channel recording to directly define, for the first time, the functional profile of NB RyR. Our results indicate that the RyR-mediated CICR contribution to the AP-triggered intracellular Ca²⁺ transient in NB cells is negligible. This is interesting because the NB cells have functional intracellular Ca²⁺ stores that contain working RyR channels. Furthermore, our single-channel studies revealed that these NB RyR channels have similar permeation, gating kinetics, and regulatory properties compared with AD RyR channels. Therefore, our results indicate that other mechanisms must exist to explain why in NB cells these channels do not contribute more substantially to the AP-triggered intracellular Ca²⁺ transient. Possible answers are discussed below.

It is possible that RyR channel density in NB cells is just too low to make a substantial contribution to the AP-triggered intracellular Ca²⁺ transient. One way to evaluate this possibility is to compare RyR density and the fraction of RyR-mediated CICR in NB and AD cells. The RyR-mediated CICR contribution in AD cells represents 70% of the AP-triggered Ca²⁺ transient (see Fig. 3). In NB cells, however, it represents only 13% of the AP-triggered Ca²⁺ transient. If RyR contribution is directly proportional to the RyR density, the expected RyR density in NB should be 18% of that present in the AD [according to a simple linear relationship of (70% CICR/100% RyR density) in AD = (13% CICR/18% RyR density) in NB]. However, ryanodine binding data from Fig. 4A indicate that the actual RyR protein density in SR microsomes from NB cells is 36%, a value twofold larger than the simplistic prediction. Several factors should be considered, however, before any conclusion is drawn. Our binding data from SR microsomal preparations involved normalization to total SR protein. Because this total may also change during development, RyR and DHPR protein density was determined in crude homogenates of NB and AD ventricular tissue (Fig. 4D). Our results in crude homogenate were normalized to milligrams of wet tissue. If we assume a total protein concentration of 109 mg/ml cell volume and a ventricular density of 1.06 g/ml (3), our results would indicate a density of ryanodine binding sites of 337 and 1,306 fmol/mg protein in NB and AD, respectively. Similarly, the density of DHP binding sites would correspond to 84 and 190 fmol/mg protein in NB and AD, respectively. Our values correspond well with those reported previously for RyR and DHPR density (14, 23, 35, 39). In addition, our value for the RyR-to-DHPR ratio in AD tissue of 6.4 also agrees with the values of 6.9 and 7.3 in rat ventricle reported by Wibo et al. (53) and Bers and Stiffel (4), respectively. Together, our binding data showed that the RyR protein density in NB was 25% of that present in the adult. Therefore, our results indicate that some RyRs do not effectively contribute to the AP-triggered Ca²⁺ transient.

This idea was recently addressed by Sedarat et al. (45), who evaluated the level of coimmunolocalization of DHPRs and RyRs in NB rabbit heart myocytes of different ages. Their results indicate that in 3-day-old myocytes, the fraction of RyRs colocalized with DHPRs is no larger than 10%. Thus our observation that RyR-mediated CICR represents 13% of the AP-triggered Ca²⁺ transient is consistent with that fraction. Although in the AD rat, only 37% of RyRs colocalize with DHPRs (44), the large ryanodine-sensitive component of the Ca²⁺ transient indicates that in the mature heart, even the uncoupled RyRs are efficiently recruited by CICR during an AP. In NB, however, only a fraction of RyRs appears to participate in the E-C coupling. This raises an interesting question: What is the functional role of the rest of the RyRs in NB cells? The "other" RyRs simply may be in the pipeline for future insertion into the developing E-C coupling apparatus, or they may be mediating intracellular Ca²⁺ signaling unrelated to E-C coupling.

Thus the lower RyR channel density in NB cells likely is not the only reason for the small RyR-mediated CICR contribution to the AP-triggered intracellular Ca²⁺ transient. The arguments presented above and published previously have assumed that the RyR channels in NB and AD cells are functionally equivalent entities. The validity of this assumption was tested in our study.

To our knowledge, this study is the first in which NB RyR functional properties have been defined at the single-channel level. Unitary conductance (in Cs⁺ and Ca²⁺), ion selectivity (P_Cs/P_Ca), and stationary modal gating kinetics, as well as the channel's sensitivity to exogenous (caffeine and ryanodine) and endogenous (Ca²⁺, ATP, and Mg²⁺) ligands, were carefully defined in this work. No substantial differences were detected during development. The steady-state cytosolic Ca²⁺ sensitivity of the NB RyR channel was nearly identical to that found in AD and corresponded to the RyR type 2 isoform (7, 8, 29). Thus the single-channel results presented clearly validate the assumption that the RyR channels in NB and AD cells are functionally equivalent entities and are most likely arising from the same molecular isoform (i.e., type 2). Consequently, changes in single-channel properties cannot be proposed as a mechanism to explain the developmental differences in RyR contribution to the AP-triggered intracellular Ca²⁺ transient.

Alternatively, it is possible that the signaling environment around the RyR channels in the NB cells will affect their contribution to the AP-triggered intracellular Ca²⁺ transient. In AD cells, most RyR channels are associated to DHPRs, whereas in NB cells, because of the absence of T tubules, the RyRs are either coupled to the sarcolemma in the periphery or uncoupled, forming corbular SR (45). For this reason, the Ca²⁺ transient reflects primarily the Ca²⁺ influx across the sarcolemma. Our results (Fig. 2, A and B) are consistent with this view; that is, Ca²⁺ signal starts in the cell's periphery and then propagates to the center, where it activates uncoupled RyRs less effectively because of slower kinetics (2, 13). This pattern of AP-triggered Ca²⁺ entry also occurs in AD atrial myocytes (5, 20, 26) and Purkinje cells (9), where a well-developed T-tubular system is absent.

Other mechanisms by which the local signaling environment could screen or attenuate the RyR function in NB involve cross talk with inositol 1,4,5-trisphosphate receptors [Ins(1,4,5)P₃Rs]. This interaction could be favored because of the relatively large Ins(1,4,5)P₃R density found in the immature heart (40). In this regard, it has been reported in cultured cells from NB rat that Ins(1,4,5)P₃R-mediated Ca²⁺ release can reduce the contribution of RyRs to the intracellular Ca²⁺ signaling due to Ca²⁺ depletion of the SR (21a). Although our work did not directly address this question, the sizeable caffeine-sensitive Ca²⁺ release observed in NB myocytes (Fig. 2B) makes this mechanism unlikely.

In addition, it must be considered that all of the structures that form the dyad (i.e., T-tube membrane, junctional space, and junctional SR) contain proteins that can modulate directly or indirectly the RyR channel activity. Developmental changes of some of these proteins that have been reported to occur in the rat may affect RyR's behavior. Although some proteins reduce their concentration during development (like calreticulin; Ref. 27), the common pattern is to increase. Examples of these proteins include DHPR (53), junctin (52), triadin (48), and calsequestrin (27). Other luminal SR proteins (i.e., histidine-rich Ca²⁺ binding protein) also may interact differently with the RyR during development (24). Whether or not all these proteins are responsible for the RyR behavior in NB, and to what extent, are questions that remain unanswered.

In conclusion, our study shows that there is a marked difference between AD and NB cells in the contribution of RyR-mediated CICR to the AP-triggered intracellular Ca²⁺ transients. The CICR component in NB is very small and cannot be attributed to the absence of functional intracellular Ca²⁺ stores or to differential single RyR channel properties. Instead, our results suggest that the small contribution of CICR to the AP-triggered Ca²⁺ transients in NB cells arise, at least partially, from the lower density of RyRs. Other contributing factors not investigated in the present study may include the cellular RyR distribution in NB cells and the relative distance between these Ca²⁺ release channels and the sarcolemmal Ca²⁺ trigger.

	GRANTS

TOP ABSTRACT METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
This work was supported by National Heart, Lung, and Blood Institute Grants HL-57832 (to M. Fill) and HL-62571 (to R. Mejía-Alvarez) and by American Heart Association Grants 0130142N (to J. A. Copello) and 9950382N (to R. Mejía-Alvarez).

	ACKNOWLEDGMENTS

 
We thank Dr. Rafael A. Rosales and Dr. Carlos Villalba-Galea for writing the Kernel analysis routines used in this work for the gating kinetics analysis.

	FOOTNOTES

 

Address for reprint requests and other correspondence: R. Mejía-Alvarez, Dept. of Physiology, Loyola Univ. Chicago, 2160 S. First Ave., Maywood IL 60153 (E-mail: rmejia{at}lumc.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

	REFERENCES

TOP ABSTRACT METHODS RESULTS DISCUSSION GRANTS REFERENCES

 

1. Baudet S, Hove-Madsen L, and Bers DM. How to make and use calcium-specific mini- and microelectrodes. Methods Cell Biol 40: 93–113, 1994.[ISI][Medline]
2. Bers D. Excitation-Contraction Coupling and Cardiac Contractile Force. Developments in Cardiovascular Medicine. Dordrecht, The Netherlands: Kluwer Academic, 1993, vol. 122.
3. Bers D. Excitation-Contraction Coupling and Cardiac Contractile Force. Developments in Cardiovascular Medicine (2nd ed.). Dordrecht, The Netherlands: Kluwer Academic, 2001, p. 41–42, 2001.
4. Bers DM and Stiffel VM. Ratio of ryanodine to dihydropyridine receptors in cardiac and skeletal muscle and implications for E-C coupling. Am J Physiol Cell Physiol 264: C1587–C1593, 1993.[Abstract/Free Full Text]
5. Brette F and Orchard C. T-tubule function in mammalian cardiac myocytes. Circ Res 92: 1182–1192, 2003.[Abstract/Free Full Text]
6. Brillantes AM, Bezprozvannaya S, and Marks AR. Developmental and tissue-specific regulation of rabbit skeletal and cardiac muscle calcium channels involved in excitation-contraction coupling. Circ Res 75: 503–510, 1994.[Abstract/Free Full Text]
7. Chu A, Fill M, Stefani E, and Entman ML. Cytoplasmic Ca²⁺ does not inhibit the cardiac muscle sarcoplasmic reticulum ryanodine receptor Ca²⁺ channel, although Ca²⁺-induced Ca²⁺ inactivation of Ca²⁺ release is observed in native vesicles. J Membr Biol 135: 49–59, 1993.[ISI][Medline]
8. Copello JA, Barg S, Onoue H, and Fleishcer S. Heterogeneity of Ca²⁺ gating of skeletal muscle and cardiac ryanodine receptors. Biophys J 73: 141–156, 1997.[Abstract/Free Full Text]
9. Cordeiro JM, Spitzer KW, Giles WR, Ershler PE, Cannell MB, and Bridge JHB. Location of the initiation site of calcium transients and sparks in rabbit heart Purkinje cells. J Physiol 531: 301–314, 2001.[Abstract/Free Full Text]
10. Cukierman S, Yellen G, and Miller C. The K⁺ channel of sarcoplasmic reticulum. A new look at Cs⁺ block. Biophys J 48: 477–484, 1985.[Abstract/Free Full Text]
11. Endo M, Tanaka M, and Ogawa Y. Calcium induced release of calcium from the sarcoplasmic reticulum of skinned skeletal muscle fibres. Nature 228: 34–36, 1970.[CrossRef][Medline]
12. Escobar AL, Ribeiro-Costa R, Villalba-Galea C, Zoghbi ME, Pérez CG, and Mejía-Alvarez R. Developmental changes of intracellular Ca²⁺ transients in beating rat hearts. Am J Physiol Heart Circ Physiol 286: H971–H978, 2004.[Abstract/Free Full Text]
13. Fabiato A. Time and calcium dependence of activation and inactivation of calcium-induced release of calcium from the sarcoplasmic reticulum of a skinned canine cardiac Purkinje cell. J Gen Physiol 85: 247–289, 1985.[Abstract/Free Full Text]
14. Fitzgerald M, Neylon CB, Marks AR, and Woodcock EA. Reduced ryanodine receptor content in isolated neonatal cardiomyocytes compared with the intact tissue. J Mol Cell Cardiol 26: 1261–1265, 1994.[CrossRef][ISI][Medline]
15. Fruen BR, Bardy JM, Byrem TM, Strasburg GM, and Louis CF. Differential Ca²⁺ sensitivity of skeletal and cardiac muscle ryanodine receptors in the presence of calmodulin. Am J Physiol Cell Physiol 279: C724–C733, 2000.[Abstract/Free Full Text]
16. Goldstein MA and Traeger L. Ultraestructural changes in postnatal development of the cardiac myocytes. In: The Developing Heart, edited by Legato MJ. Boston, MA: Nijhoff, 1985, p. 1–20.
17. Gorza L, Vettore S, Tessaro A, Sorrentino V, and Vitadello M. Regional and age-related differences in mRNA composition of intracellular Ca²⁺-release channels of rat cardiac myocytes. J Mol Cell Cardiol 29: 1023–1036, 1997.[CrossRef][ISI][Medline]
18. Haddock PS, Coetzee WA, Cho E, Porter L, Katoh H, Bers DM, Jafri KS, and Artman M. Subcellular [Ca²⁺]_i gradients during excitation-contraction coupling in newborn rabbit ventricular myocytes. Circ Res 85: 415–427, 1999.[Abstract/Free Full Text]
19. Hamilton SL, Mejía-Alvarez R, Fill M, Hawkes MJ, Brush KL, Schilling WP, and Stefani E. [³H]PN200-110 and [³H]ryanodine binding and reconstitution of ion channel activity with skeletal muscle membranes. Anal Biochem 183: 31–41, 1989.[CrossRef][ISI][Medline]
20. Hüser J, Lipsius SL, and Blatter LA. Calcium gradients during excitation-contraction coupling in cat atrial myocytes. J Physiol 494: 641–651, 1996.[ISI][Medline]
21. Husse B and Wussling M. Developmental changes of calcium transients and contractility during the cultivation of rat neonatal cardiomyocytes. Mol Cell Biochem 164: 13–21, 1996.[CrossRef]
21. Jaconi M, Bony C, Richards SM, Terzic A, Arnaudeau S, Vassort G, and Puceat M. Inositol 1,4,5-trisphosphate directs Ca²⁺ flow between mitochondria and the endoplasmic/sarcoplasmic reticulum: a role in regulating cardiac autonomic Ca²⁺ spiking. Mol Biol Cell 11: 1845–1858, 2000.[Abstract/Free Full Text]
22. Kazazoglou T, Schmid A, Renaud JF, and Lazdunski K. Ontogenic appearance of Ca²⁺ channels characterized as binding sites for nitrendipine during development of nervous, skeletal and cardiac muscle systems in the rat. FEBS Lett 164: 75–79, 1983.[CrossRef][ISI][Medline]
23. Kim DH, Mkparu F, Kim CR, and Caroll RF. Alteration of Ca²⁺ release channel function in sarcoplasmic reticulum of pressure-overload-induced hypertrophic rat heart. J Mol Cell Cardiol 26: 1505–1512, 1994.[CrossRef][ISI][Medline]
24. Kim E, Shin DW, Hong CS, Jeong D, Kim DH, and Park WJ. Increased Ca²⁺ storage capacity in the sarcoplasmic reticulum by overexpression of HRC (histidine-rich Ca²⁺ binding protein). Biochem Biophys Res Commun 300: 19