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Am J Physiol Heart Circ Physiol 279: H202-H209, 2000;
0363-6135/00 $5.00
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Vol. 279, Issue 1, H202-H209, July 2000

Colocalization of dihydropyridine and ryanodine receptors in neonate rabbit heart using confocal microscopy

Franklin Sedarat1, Liqun Xu1, Edwin D. W. Moore2, and Glen F. Tibbits1

1 Cardiac Membrane Research Laboratory, Simon Fraser University, Burnaby, British Columbia V5A 1S6; and 2 Department of Physiology, University of British Columbia, Vancouver, British Columbia V6T 1Z3, Canada


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Because of undeveloped T tubules and sparse sarcoplasmic reticulum, Ca2+-induced Ca2+ release (CICR) may not be the major mechanism providing contractile Ca2+ in the neonatal heart. Spatial association of dihydropyridine receptors (DHPRs) and ryanodine receptors (RyRs), a key factor for CICR, was examined in isolated neonatal rabbit ventricular myocytes aged 3-20 days by double-labeling immunofluorescence and confocal microscopy. We found a significant increase (P < 0.0005) in the degree of colocalization of DHPR and RyR during development. The number of voxels containing DHPR that also contained RyR in the 3-day-old group (62 ± 1.8%) was significantly lower than in the other age groups (76 ± 1.3 in 6-day old, 75 ± 1.2 in 10-day old, and 79 ± 0.9% in 20-day old). The number of voxels containing RyR that also contained DHPR was significantly higher in the 20-day-old group (17 ± 0.5%) compared with the other age groups (10 ± 0.7 in 3-day old, 11 ± 0.6 in 6-day old, and 11 ± 0.5% in 10-day old). During this period, the pattern of colocalization changed from mostly peripheral to mostly internal couplings. Our results provide a structural basis for the diminished prominence of CICR in neonatal heart.

calcium; dyadic coupling; excitation-contraction coupling


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

EXCITATION-CONTRACTION COUPLING (E-C coupling) in adult cardiac myocytes requires Ca2+ influx through sarcolemmal L-type Ca2+ channel or dihydropyridine receptor (DHPR), followed by Ca2+-induced Ca2+ release (CICR) from the sarcoplasmic reticulum Ca2+-release channel or ryanodine receptor (RyR) (5, 6). A major structural specialization in adult cardiac myocytes is dyadic couplings formed between sarcoplasmic reticulum (SR) and either the external or T-tubular sarcolemma (SL) (12, 21). The close apposition of the SR and SL defines a functionally restricted space that acts like an incomplete barrier to diffusion underneath the sarcolemmal membrane. In this space, which has been referred to as fuzzy space, RyRs are located very close (<20 nm) to the SL and T-tubule membranes and thus are exposed to a high local intracellular calcium concentration ([Ca2+]i) whenever neighboring DHPRs open (17). The macroscopic behavior of CICR depends critically on the spatial relationship of the DHPR and RyR in dyadic couplings, as well as on SL and SR Ca2+-channel kinetics (22).

During mammalian heart development, the morphology undergoes significant changes as does the mechanism of E-C coupling (7). The SR volume in neonates is less than in the adult, and both the amount of SR and SR Ca2+ uptake per gram of muscle increase with age (19). Most newborn mammalian cardiomyocytes do not develop T tubules until 8-10 days of age (11). With the lack of a developed T-tubular system in neonatal heart, the spatial relationship of DHPR to RyR may be different from that in the adult heart. Because the proximity of DHPR and RyR is a key factor for CICR, neonatal myocardium may not rely on CICR and the triggered release of Ca2+ from SR to provide contractile Ca2+. It has been shown, for example, that Ca2+-channel blockers have little effect on tension generation in neonatal cardiac muscle (16). This suggests that the Ca2+ current may not directly contribute Ca2+ for contraction or for CICR in neonatal cells. Alternatively, reverse-mode Na+/Ca2+ exchange has been suggested as a major source of Ca2+ influx for contraction in neonatal myocytes (3). The cardiac SL Na+/Ca2+ exchanger is abundant and functionally well developed in the late fetal/early newborn rabbit heart, and the density appears to decline postnatally (1). Given that the colocalization of DHPR and RyR is an essential element in E-C coupling in adult heart, it is of interest to determine the developmental changes in the geometric arrangement of DHPRs and RyRs in neonatal heart. In the present study we report the distribution pattern and degree of colocalization of DHPR and RyR in rabbit myocardial cells during ontogeny.


    MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Animals. Male or female neonatal New Zealand White rabbits were used in four age groups: 1- to 3-day old, 6- to 7-day old, 10- to 11-day old, and 20- to 21-day old. For each age group, 5 hearts and 10 cells/heart (50 cells/age group) were studied.

Antibody characterization. Mouse monoclonal anti-RyR antibodies (IgG1, clone C3-33, 1 mg/ml) were purchased from Affinity Bioreagents. Specificity of this antibody in rabbit cardiac muscle was determined with the use of Western blots as previously described (2). Rabbit polyclonal anti-DHPR antibody (CNC1, 0.469 mg/ml) was used; the specificity of this antibody has been previously described (10). CNC1 specifically binds to the class C alpha 1-subunit of the L-type Ca2+ channel.

Cell isolation procedure. The method of cell isolation was adapted from Mitra and Morad (18). All of the solutions were prepared with double-deionized water and filtered with a 0.2-µm filter. The solutions were aerated with 100% O2 before and during the isolation. Neonatal rabbits were anesthetized and heparinized by an intraperitoneal injection of pentobarbital sodium (60 mg/kg body wt) and heparin (2,700 USP/kg body wt). The heart was excised and kept in cold (4°C) dissection solution (in mM: 126 NaCl, 4.4 KCl, 5 MgCl2, 24 HEPES, 22 glucose, 20 taurine, 5 creatine, 5 sodium pyruvate, and 1 NaH2PO4, pH 7.4 at 37°C) for ~1 min to arrest the heart. After aortic cannulation, the coronary arteries were washed out by 5-10 ml of cold modified Kraftbruhe (KB) solution (in mM: 10 taurine, 70 glutamic acid, 25 KCl, 10 KH2PO4, and 22 dextrose, pH 7.3 at 22°C), which was nominally free of Ca2+ and Na+. The heart was then mounted on a Langendorff apparatus. The heart was retrogradely perfused with prewarmed (37°C) and oxygenated KB solution for 4 min. The heart was then digested with a collagenase (0.5 mg/ml, collagenase type II; Worthington) solution for 5-8 min at a flow rate of 1-4.7 ml/min at 34-35°C (enzyme concentration, flow rate, and enzyme digestion time depend on animal age; Table 1). This was followed by perfusion with an EGTA (0.5 mM) containing KB solution for 5 min to wash out the digestive enzymes. The digested heart was transferred to a petri dish containing 5 ml EGTA-KB solution, and the ventricles were dissected from the heart. The ventricles were gently teased with forceps to disperse individual myocytes. The cell suspension was filtered through a coarse nylon mesh (200 µm) to remove tissue chunks.

                              
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  		Table 1.   Protocols for cell isolation in different age groups

Indirect immunofluorescence labeling. Double labeling was performed on isolated rabbit myocytes with anti-RyR and anti-DHPR primary antibodies. Isolated myocytes were initially fixed with 2% buffered paraformaldehyde for 10 min. The fixed cells were quenched for aldehyde groups in 0.75% glycine buffer for 10 min. The cells were then permeabilized with Triton X-100 (0.1%) for 10 min. After being washed with PBS for 10 min, the cells were incubated with nonconjugated goat anti-rabbit IgG (Molecular Probes, Eugene, OR; 2 mg/ml) to block possible cross-reactions between the secondary anti-rabbit antibody and rabbit myocardial cells. The antibody dilution was 1:100 (2-h incubation time in 1.5 ml microcentrifuge tubes with gentle agitation). Excess antibody was washed off with the use of antibody wash solution [0.05% Triton X-100 in SSC (150.7 mM sodium chloride and 17.5 mM sodium citrate)] for 10 min and then PBS for another 10 min. Subsequently, the cells were incubated overnight with primary antibodies in antibody buffer solution (2% goat serum, 1% BSA, 0.05% Triton X-100, and 3 mM NaN3 in SSC). The antibody buffer solution contained goat serum and BSA to block nonspecific binding sites. Antibody dilution for primary antibodies was 1:80 to 1:100 for the polyclonal anti-DHPR and 1:100 to 1:200 for the monoclonal anti-RyR. Excess primary antibodies were washed off with the use of antibody wash solution (2 × 10 min) and then PBS (10 min). The cells were then incubated with Alexa-conjugated secondary antibodies (Molecular Probes; 2 mg/ml), diluted 1:100, for 2 h. Highly cross-adsorbed Alexa488 goat anti-rabbit IgG (H+L) and Alexa594 goat anti-mouse IgG (H+L) were used. After application of labeled secondary antibodies, the test tubes were wrapped in aluminum foil to prevent light exposure. The cells were then washed twice with the antibody wash solution (10 min) and once with PBS (10 min). In each step, solutions were added to disposable centrifuge tubes containing a suspension of isolated myocardial cells, and the tubes were gently agitated. At the end of each step, the tubes were centrifuged for 5 min at 5,000-10,000 rpm (depending on the size of the cells), and then the pelleted cells were used for the next step of the experiment. The cells were rinsed in a 90% glycerol-PBS mixture containing 2.5% 1,4-diazabicyclo-[2.2.2] octane (DABCO) and then mounted on a slide. In control experiments, single staining (to ensure the functionality of each primary antibody) and single staining with reverse secondary antibody were performed. To determine nonspecific binding, staining control experiments with secondary antibody without primary antibody were also performed.

Wheat germ agglutinin. Wheat germ agglutinin (WGA) was used to visualize SL and T-tubular patterns. Nonpermeabilized isolated myocytes were incubated with WGA (100 µg/ml) coupled to tetramethylrhodamine isothiocyanate (TRITC; Sigma) for 30 min. The cells were rinsed three times with PBS and then mounted on a slide.

Microscopy. Samples were examined with the use of a Zeiss LSM 410 laser scanning confocal microscope. An Ar-Kr 488/568 laser provided the excitation light beam, and a neutral density filter (T 0.01) was used to uniformly attenuate the intensity of laser light. The excitation light (488 and 568 nm) passed through a dual-band dichroic beam splitter (FT 488/568) that allowed the capture of images in both green and red channels simultaneously. The green and red emissions were separated by a dichroic splitter (FT 560) and filtered (515- to 540-nm band-pass filter for green and >610-nm long-pass filter for red emission). The Z interval was adjusted to 0.25 µm, and the number of sections was adjusted to 50-70 planes depending on the diameter of the cells. Two three-dimensional (3-D) images were acquired from the two different emission wavelengths representing DHPR and RyR. To determine the amount of colocalization of DHPR and RyR, images were imported into Optimas 5.2 image processing software. With the use of Analytical Language for Images, macros were written to analyze the 3-D data sets. A threshold was applied to the images to exclude ~99% of the signal found in the control images. The two 3-D images were compared voxel by voxel to determine the degree of colocalization. To get the best possible resolution, we optimized the performance of the optical system. A highly corrected objective (Zeiss Plan-Apochromat ×63, numerical aperture 1.40 oil), standard Zeiss immersion oil, and coverslips of 0.17-mm thickness were used. To reduce spherical aberration due to the refractive index (eta ) mismatch, mounting medium was prepared with glycerol (eta  = 1.47) to make it similar to that of immersion oil with eta  = 1.518 (96% similarity). Because there is lower resolution when recording optical sections in focal planes away from the coverslip, the slides were left upside down after slide preparation; this allowed the myocardial cells to attach to the coverslip. Despite the use of a plan objective, just a small part of the field of view nearest the optical axis was imaged to reduce the effect of off-axis aberrations. A plan-apochromat objective was used to eliminate chromatic aberrations. It should be noted that even in confocal microscopy, to some extent images may suffer from degradation because of out-of-focus light contributing to in-focus areas and also from anisotropy in the imaging properties (i.e., inferior axial resolution compared with the lateral resolution). In this study, the best compromise between signal level and spatial resolution was found by setting the confocal pinhole to the diameter of the Airy disk. By application of the Nyquist theorem, it was determined that a pixel diameter of ~100 nm (in reference to the specimen) was needed to properly sample the data in the xy-plane. By adjustment of the zoom setting in the confocal microscope, a pixel size equal to 100 nm was obtained.

Materials. All chemicals used were purchased from Sigma Chemicals unless specified otherwise.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Immunofluorescence. Immunoblots of crude membrane extracts from ventricular tissue of neonatal rabbits indicated that the MA3-916 anti-RyR antibody binds specifically to an antigen of the predicted relative molecular mass (~565 kDa). Control experiments demonstrated that there is no cross-reactivity between the two sets of primary and secondary antibodies, as shown in Fig. 1.


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  		Fig. 1.   A: 3-dimensional confocal image of distribution of dihydropyridine receptor (DHPR) in a 20-day-old rabbit myocyte. Data were introduced by superposition (max-z projection) of a series of optical sections from front to back surfaces of cell. B: control experiment related to myocytes from same rabbit as in A. Cell was prepared with use of double-labeling procedure outlined in MATERIALS AND METHODS except that primary antibodies were omitted. Confocal microscope settings for images in A and B were identical. Absence of any specific signal on control image indicates that blocking of possible cross-reactions between secondary anti-rabbit antibody and rabbit myocardial cells was complete. C: same image as in B, but intensity and contrast of pixels in image were adjusted to make it possible to see pixels related to nonspecific binding of secondary anti-rabbit antibody. In calculation of colocalization degree, a threshold that excluded ~99% of signal found in image in B was applied to image in A to exclude any fluorescent signal related to nonspecific binding. Same procedure was applied to image of distribution of ryanodine receptor (RyR; not shown).

WGA. WGA staining patterns before and after T-tubular formation are illustrated in Fig. 2, A and B. T tubules were not observed in 3- and 6-day-old rabbit myocytes. In these age groups, WGA stained the SL, and a boundary around the periphery of the cell was observed. Although the cells were not permeabilized, a few internal spots were observed. This suggests that fixation of myocardial cells may induce some pores in the SL. T tubules were first observed in myocardial cells from 10-day-old rabbits. At this age, the T tubules were observed as small invaginations, whereas in more mature myocytes, WGA labeling was distributed throughout the entire cell, indicating a more developed T-tubular system. Different degrees of T-tubular development were observed in different cells obtained from the same animal. A nonuniform appearance of T-tubular development in the cells acquired from the same animal, particularly in myocytes from 10-day-old rabbits, indicated that in a given heart myocardial cells can be in different stages of development.


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  		Fig. 2.   Wheat germ agglutinin (WGA) staining pattern in myocardial cells before and after T-tubular formation. A: 6-day-old rabbit myocyte labeled with TRITC-WGA. Note fluorescent labeling as a boundary around cell (no T tubules). Arrows, staining related to internal organelles (refer to RESULTS); n, nuclear shadow. B: nearly fully developed T tubules in a myocardial cell isolated from 20-day-old rabbit heart.

RyR, DHPR, and colocalization staining patterns. Figure 3 is composed of representative confocal images acquired from isolated myocytes in each of the four age groups. Each panel shows RyR (red), DHPR (green), and colocalization (yellow) staining patterns in one focal plane close to the center of the cell.


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  		Fig. 3.   RyR staining pattern (left, pseudocolored red), DHPR staining pattern (middle, pseudocolored green), and colocalization staining pattern (right, colocalized pixels are pseudocolored yellow) in myocardial cells isolated from 3- (A), 6- (B), 10- (C), and 20-day-old (D) rabbit hearts. Nos. of voxels containing DHPR were 8,778, 11,563, 25,192, and 75,799 in A, B, C, and D, respectively. Nos. of voxels containing RyR were 91,612, 86,177, 185,275, and 339,733 in A, B, C, and D, respectively. Nos. of voxels containing both DHPR and RyR were 5,677, 8,609, 18,949, and 59,623 in A, B, C, and D, respectively. Note that above nos. were calculated from whole image stack, but just 1 focal plane from each image stack is shown in each panel. Refer to RESULTS for description of staining patterns.

Staining pattern of RyR was similar in all age groups and included striations, spaced at regular intervals of ~2 µm, that clearly indicated the existence of cytoplasmic arrays of RyR (red pixels in Fig. 3, A-D). This pattern is consistent with that of the Z lines in adult myocardial cells but was observed in even the youngest animals at a time when the T tubules had not yet formed.

The pattern of distribution of DHPR was different in the different age groups. In young animals, before the development of T tubules, fluorescence associated with the DHPRs was seen as discrete spots only in the periphery of the cell, most likely in the SL (green pixels in Fig. 3, A and B). After the formation of T tubules, however, fluorescence associated with DHPRs could be seen both at the periphery of the cell and in the cell interior (green pixels in Fig. 3, C and D). There was a close correlation between the age at which the T tubules began to form and the time at which DHPRs began to be seen in the cell interior. In 10-day-old rabbits, fluorescence associated with DHPRs was closer to the periphery than the center of the cell, and this parallels the pattern of T tubules at this age, which are detected as only small invaginations from the surface. In 20-day-old rabbits, however, DHPRs appeared to be distributed in a pattern similar to that of RyR: regularly spaced transverse bands (and at this age the T tubules have nearly fully developed). In pilot studies, live myocardial cells were labeled, by suspending them in a buffer that resembled the intracellular environment, and skinned with saponin. Because this approach did not use Triton X-100 and there were no significant differences in the staining pattern of DHPR, we believe there were either no or a negligible number of epitopes removed by the mild Triton X-100 treatment used in the present study.

In superimposed images, a voxel containing both DHPR and RyR is pseudocolored yellow. The pattern of distribution of colocalized DHPR and RyR was strikingly similar to the pattern of distribution of DHPR in every age group. In young animals, there were discrete points of colocalization in the periphery before T-tubular formation (yellow pixels in Fig. 3, A and B). In older animals, there was an increase in the number of colocalized voxels in the interior of the cell, and, finally, the colocalized voxels appeared as regularly spaced arrays in animals in which the T tubules had nearly fully developed (yellow pixels in Fig. 3, C and D). In 3- and 6-day-old rabbits, the sarcolemmal region was demarcated by discrete yellow spots. Many of the colocalized voxels were in register with the regularly spaced transverse bands of RyR in the cytoplasm. This suggests that DHPRs may be clustered in the SL in close association with subsarcolemmal junctional SR to make peripheral couplings. At the same time, a substantial amount of RyR-specific staining was observed in the interior of the cell that was not in close association with DHPRs and probably represents corbular SR. In 10-day-old rabbits, colocalization was detected in both the periphery and in discrete spots within the cell. This indicates both peripheral and internal couplings, suggesting that DHPRs are appearing inside the developing T tubules and are making internal couplings with SR elements. In the 20-day-old group, yellow voxels were most likely to appear in transversely oriented bands along the entire length of the cell. Figure 4 shows cross-sectional images of myocardial cells before and after T-tubular formation.


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  		Fig. 4.   A: cross sections of a 6-day-old rabbit myocardial cell. Cell has been rotated about x- and y-axes. Top: DHPR (green), RyR (red), and colocalization (yellow) staining patterns. Bottom: only voxels containing DHPR. It is apparent from cross sections that DHPR and also coincident voxels are located almost exclusively on cell surface at this age. Note elongation of distribution patterns along the z-axis because of lower resolution in z (axial resolution) compared with lateral resolution. B: cross (yz) sections of a 20-day-old rabbit myocyte. Colocalized voxels (yellow, top) and DHPR voxels (green, bottom) are distributed both on cell surface (arrowheads) and in interior.

Degree of colocalization. With the use of Optimas software, the degree of colocalization was calculated in the different age groups. The results are shown in Table 2.

                              
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  		Table 2.   Degree of colocalization in different age groups

For both DHPR and RyR, the degree of colocalization increased with age. Even in the youngest group, >60% of DHPRs were colocalized, but the colocalized voxels were restricted to the periphery. At the same time, 90% of RyRs were not colocalized. A lower degree of colocalization for RyR compared with DHPR was observed in all age groups because of a higher number of voxels containing signal specific for RyR compared with the number of voxels containing signal specific for DHPR. The number of voxels containing RyR was almost five- to seven-fold greater than the number of voxels containing DHPR. In the older age groups, the degree of colocalization increased. In 20-day-old animals, almost 80% of DHPRs and 17% of RyRs were colocalized. At this age, although >80% of RyRs were not colocalized, the pattern of colocalization was totally changed compared with younger animals. In these more mature cells, colocalized voxels were detected along the entire width of the myocardial cell. To quantify peripheral vs. internal couplings, the ratios of pixels containing peripheral couplings to the total number of coincident pixels were calculated in different age groups. In each age group, 20 cells were randomly selected. In each cell, images from one plane in the middle of the stack were chosen. In these image planes, numbers of coincident pixels located at the border of the cells were measured as peripheral couplings. As shown in Fig. 5, the percentages of peripheral couplings in the myocytes from 3-, 6-, 10-, and 20-day-old rabbits were 96.8 ± 1.0, 88.1 ± 1.8, 40.5 ± 3.5, and 24.8 ± 2.5, respectively.


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  		Fig. 5.   Percentages of peripheral couplings in different age groups representing ratio of colocalized pixels located in the periphery of cell to total no. of colocalized pixels. Values are means ± SE; n = 20 for each age group.

Statistical analysis. With the use of SPSS statistical analysis software, one-way ANOVA was performed to examine the equality of the means for the amount of colocalization in the different age groups.

A multiple-comparisons test indicated that the DHPR colocalization differs significantly between the first age group (1- to 3-day old) and all other groups (P < 0.0005). For RyR, the amount of colocalization in the oldest group (20-day old) differs significantly from all other groups (P < 0.0005).


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Immunofluorescent localization of DHPR and RyR. This study examined the subcellular distribution of SL L-type Ca2+ channels and SR Ca2+-release channels in neonate rabbit ventricular myocytes aged 3-20 days.

RyR staining pattern. We observed that the staining pattern of RyR was in transverse bands at regularly spaced intervals of ~2 µm. This is consistent with the RyR pattern in adult rabbit myocardial cells, as reported by Carl et al. (2). Jorgensen et al. (14) detected a similar RyR-specific pattern of transversely oriented rows of fluorescent foci in rat papillary myofibers. We found well-organized arrays of RyRs in the cytoplasm of myocytes even in 3-day-old rabbits at a time when the T tubules had not yet formed. This parallels the report of Carl et al. (2) of RyR staining pattern in adult rabbit atrial cardiac myocytes, which also lack a T-tubular system. The RyR pattern was similar in all age groups. Parameswaran et al. (20) studied the distribution of the RyR in postnatal developing porcine cardiac muscle (at 3, 5, 10, and 20 days) and reported no difference in immunofluorescence labeling of RyR in these age groups compared with adult porcine cardiac myocytes.

RyRs were partly colocalized and partly noncolocalized with SL or T-tubular DHPRs. Because the prominent structural difference between junctional and corbular SR is that junctional SR is connected to either T tubules or to SL via "feet" structures, whereas corbular SR is not (13), we conclude that RyRs in rabbit myocardial cells are located in both junctional and corbular SR. This is in agreement with the immunoelectron microscopy studies by Jorgensen et al. (14), which reported that RyRs were localized to junctional and corbular SR. Another consideration is whether or not the RyR staining pattern is different in junctional and corbular SR. In our images, the RyR staining pattern was identical in junctional and corbular SR. This parallels the study of the structure of corbular SR in rabbit cardiac muscle by freeze fracture. Dolber and Sommer (4) reported that the processes on the surface of corbular SR had all the anatomical features of junctional processes of junctional SR.

DHPR staining pattern. Before T-tubular formation, we observed the pattern of DHPR staining as discrete spots limited to the periphery of the cell. This pattern is consistent with DHPR staining pattern in rabbit atrial cells that lack a T-tubular system, as reported by Carl et al. (2). In 10-day-old rabbits, DHPRs were observed as discrete spots in the interior of the cell as well as in the periphery. In 20-day-old rabbits, DHPRs were distributed mostly in transverse bands similar to RyR staining pattern. The DHPR staining pattern in the 20-day-old rabbit is similar to that in adult rabbit, as reported by other investigators (2).

An interesting observation in our study was the presence of some intensive green fluorescent signals around the nuclear area in some of the 3- and 6-day-old rabbit myocytes. This was somewhat unexpected in immature cells that are essentially devoid of T tubules, and, as a result, the fluorescence associated with DHPR should be restricted to cell periphery. These spots were never observed in control experiments, which indicates that they are not related to nonspecific binding. This fluorescent staining is most likely related to alpha 1-subunits located in cytoplasmic organelles, either in the process of posttranslational assembly and packaging for the delivery to the SL or in the process of degradation.

Colocalization of DHPR and RyR. There are two major findings in this study related to colocalization of DHPR and RyR during the period of ontogeny: 1) the degree of colocalization of DHPR and RyR increases with age during development, and 2) there is a transition from couplings restricted to the periphery to mostly internal couplings along the entire length of the cell. It is conceivable that by increasing the number of couplings and the distribution of these couplings over the entire cell, the role of inward Ca2+ current (ICa), CICR, and SR Ca2+ release increases in the contraction of developing myocardial cells.

Before T-tubular formation, we found that codistribution of DHPR and RyR was restricted to the peripheral SL. Other studies in chick myocardium (23) with no T tubules and also in rabbit atrial cells (2) revealed dyadic couplings between DHPR and RyR restricted to the peripheral SL. We observed more internal dyadic couplings in the older animals, which paralleled the development of the T-tubular network. In the 20-day-old animals, couplings were most likely to appear along the entire width of the cell. This is consistent with the distribution of DHPR and RyR couplings in adult rabbit ventricular cells, as reported by other researchers (2). In a recent study, localized SR Ca2+-release events (Ca2+ sparks) were reported to occur predominantly at the cell periphery of newborn (1-14 days) rabbit myocytes in contrast to adults, in which sparks occurred across the entire width of the cell (9). In all age groups, we observed many of the RyRs as non-colocalized with DHPRs. Even in 20-day-old rabbits, only 17% of RyRs were colocalized. In one study by Moore (EDW Moore, unpublished data) in adult rat myocardial cells, ~30% of RyRs were reported as colocalized with DHPR. It would be worthwhile to determine the role of these "uncoupled" RyRs in corbular SR and whether they are activated in the absence of a close association with the DHPR.

In this study, even in the youngest group, fluorescence related to DHPR was detected in clusters mostly in the regions of the SL that are in register with cytoplasmic arrays of RyR, which we refer to as morphological cross talk. This is in contrast to Na+/Ca2+ exchanger (NCX) immunolocalization studies, which have shown that antibodies against NCX label SL almost uniformly (3, 15). Although this morphological cross talk may suggest that the two proteins are in close association with each other, it does not necessarily mean that they are close enough to have functional cross talk. It should be noted that measurement of colocalization degree is always at the limits of the optical system. It has been postulated that the L-type Ca2+ channel should be within ~20 nm of the RyR to trigger Ca2+ release from the SR (16). Even under ideal conditions, the resolution of confocal microscopy is several times this distance. Changes in E-C coupling associated with cardiac hypertrophy investigated by Gomez et al. (8) suggest that functional cross talk is very sensitive to the geometric arrangement of DHPRs and RyRs in the dyad (8). With the use of a perforated patch-clamp technique, we studied inactivation kinetics of the L-type Ca2+ channel before and after application of 5 mM caffeine from myocytes of rabbits of the same age groups as used in the present study (24). These data, which show only the inactivation kinetics of the 20-day-old group to be affected by caffeine, are consistent with the notion that DHPR and RyR functional coupling is only observed in the oldest group.

Our data along with data from other investigators suggest that dyadic couplings in rabbit ventricular myocytes undergo both structural and functional changes during the first 20 days of life. In immature myocytes, elements of dyadic couplings (DHPR and RyR) are less coupled, either morphologically or functionally. Because cross signaling of DHPR and RyR is the cornerstone of CICR, and CICR is widely accepted as the fundamental component of E-C coupling in mature heart, there should be another alternative mechanism in the immature myocardium to provide the Ca2+ required for contraction. Because the NCX is abundantly localized in the SL and demonstrates high activity in fetal and newborn hearts, reverse-mode Na+/Ca2+ exchange may be a major pathway for transmembrane influx of Ca2+ (3). We speculate that in the rabbit myocytes at some time during ontogeny, most likely in the first month after birth, the transition to adult E-C coupling mechanism occurs, and the dependence on the NCX to provide contractile Ca2+ decreases. The characterization of the control systems that integrate these changes remains to be achieved.


    ACKNOWLEDGEMENTS

We thank the generous support of the Heart and Stroke Foundation of British Columbia and Yukon (to G. F. Tibbits) that made these studies possible. Rabbit polyclonal anti-DHPR antibody was generously provided by Dr. William A. Catterall at the Department of Pharmacology, University of Washington, Seattle, WA.


    FOOTNOTES

Address for reprint requests and other correspondence: G. F. Tibbits, Cardiac Membrane Research Laboratory, Simon Fraser Univ., Burnaby, BC V5A 1S6, Canada (E-mail tibbits{at}sfu.ca).

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. §1734 solely to indicate this fact.

Received 4 August 1999; accepted in final form 29 December 1999.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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Am J Physiol Heart Circ Physiol 279(1):H202-H209
0363-6135/00 $5.00 Copyright © 2000 the American Physiological Society


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