© 1997 American Heart Association, Inc.
Articles |
Dissociated Spatial Patterning of Gap Junctions and Cell Adhesion Junctions During Postnatal Differentiation of Ventricular Myocardium
the Laboratory of Eukaryotic Molecular Genetics (B.D.A., A.I.M.), National Institute for Medical Research, London, England; the Department of Anatomy and Developmental Biology (L.U.R.K., K.W.), University College London; the Department of Cardiac Medicine (N.J.S., S.R.), National Heart and Lung Institute, Royal Brompton Hospital, London; and the Department of Cell Biology and Anatomy (R.P.T., R.G.G.), Cardiovascular Developmental Biology Center, Medical University of South Carolina, Charleston. E-mail Robert Gourdie@musc.edu
Correspondence to Dr R.G. Gourdie, Department of Cell Biology and Anatomy, Cardiovascular Developmental Biology Center, Medical University of South Carolina, 171 Ashley Ave, Charleston, SC 29425.
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Abstract |
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Nonuniformity in the spatial patterning of gap junctions between heart muscle cells is now recognized as an important determinant of electromechanical function in working myocardium. Breakdown of the normal geometry of electrical intercellular connectivity in diseased myocardium correlates with reentry, arrhythmia, and conduction disturbance. The developmental mechanism(s) that determines this precise spatial order in gap junction organization in normal myocardium is at present unknown. To examine this question, we have used immunoelectron and immunoconfocal microscopy to analyze the spatial distributions of gap junctional (connexin43), desmosomal (desmoplakin), and adherens junctional (N-cadherin) components during maturation of rodent and canine left ventricular myocardium. In rats, a striking divergence in the distribution of gap junctions and cell adhesion junctions emerged within the first 20 days of postnatal life. It was found that although gap junctions initially demonstrated dispersed distributions across myocyte cell membranes, desmosomes and adherens junctions showed more rapid polarization toward cell termini (ie, nascent intercalated disks) after birth. Over subsequent postnatal development (20 to 90 postnatal days), gap junctions became progressively concentrated in these cell adhesion junction–rich zones of membrane. Quantitative analyses of this process in a series of rats aged 15 embryonic and 1, 5, 10, 20, 40, 70, and 90 postnatal days indicated that significantly higher levels (P<.01) of N-cadherin and desmoplakin than of connexin43 were immunolocalized to cell termini by as early as postnatal day 5. Although all three junctions types showed increasing polarization to myocyte termini with development, variation between junctions remained significant (P<.05) at all times points between 5 and 70 postnatal days. Only at 90 postnatal days, when the animals were nearly full grown, did the proportions of gap junction, desmosome, and adherens junction at intercalated disks become statistically similar (P>.05). Examination of myocardium from 1- and 3-month-old canines revealed that related differential changes to the spatiotemporal distribution of intercellular junctions occurred during postnatal maturation of the dog heart, suggesting that the process was not rodent specific. It is concluded that this progressive change in the organization and pattern of association between gap junctions and cell adhesion junctions is likely to be an important factor in maturation of electromechanical function within the mammalian heart.
Key Words: heart • development • gap junction • intercellular adhesion junction • anisotropy
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Introduction |
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Gap junctions are sievelike aggregates of intercellular channels that permit the regulated flow of ions, metabolites, and electrical current between cells.1 Working muscle cells in the mammalian heart are connected together by large numbers of gap junctions, and these multiple sites of coupling provide the physical basis for propagation of depolarizing membrane potential from one cardiac muscle cell to another.2 3 4 As such, the gap junction is fundamental to coordinating the synchronous pumping action of the myocardium. It has been increasingly recognized that the spatial distribution of gap junctions is likely to be a key determinant of electromechanical function in the heart. Applied and theoretical electrophysiological investigations have provided evidence for the importance of nonuniformities in gap junction distribution in patterning the spread of electrical excitation in developing, mature, and diseased myocardial tissues.5 6 7 8 Morphological studies have correlated extensive remodeling of gap junction distribution with potential arrhythmogenic foci in ischemic heart disease9 10 11 12 and in hypertrophic myocardiopathy.13 14 During growth of the heart, progressive changes in the distribution of intercellular junctions15 16 17 18 19 20 have been correlated with the development of uniform electrical anisotropy,8 a characteristic of the rapid and orderly propagation of action potential through adult cardiac muscle. Although it is certain that the geometry of connectivity between myocytes is an important determinant of cardiac electrophysiological properties, the generation of this spatial order remains incompletely understood.
In the working myocardium of adult mammalian ventricle, gap junctions colocalize in nonuniform distributions at myocyte boundaries with two other intercellular junction types, desmosomes and adherens junctions.21 This codistribution occurs at intercalated disks, zones of end-to-end electromechanical coupling between neighboring muscle cells. In functional distinction to the gap junction, desmosomes and adherens junctions act to fasten myocytes together mechanically and to provide anchorage sites for the cytoskeleton and myofibrillar contractile apparatus, respectively. Myocytes normally have multiple intercalated disks, forming irregular branchlike patterns of disk contact with, on average, 9 to 12 neighboring cells.4 10 11 The basis of the correlated distributions of gap junctions and cell adhesion junctions remains unclear. There is evidence for roles of cell adhesion molecules on the formation of gap junctions from studies in vitro of a variety in cell types.22 23 24 However, whether adhesive interactions between cells also determine and maintain nonuniformities in the spatial patterning of gap junctions is uncertain.
Although the morphogenesis of the rodent heart is complete at birth, the differentiation of neonatal working myocytes is not advanced.25 During postnatal growth, working myocytes undergo large hypertrophic increases in size accompanied by marked accretion of myofibrillar contractile apparatus, T tubules, glycogen, and mitochondria. This period is also associated with pronounced changes in gap junctional electrical connectivity between these cells.15 16 17 At birth, myocardial gap junctions are uniformly dispersed across myocyte membranes. Over postnatal growth, concomitant with a loss of lateral intercellular connections, there is a progressive accumulation of gap junctions in intercalated disks. In the rat, this process occurs over a relatively extended period of development and does not culminate until the animal is well past sexual maturity (40 to 90 postnatal days). Comparable results indicating a prolonged phase of reorganization of cardiac intercellular junctions have recently been reported for humans in early childhood18 19 20 and have been suggested to underlie alteration in the rate and anisotropy of conduction during postnatal development.8 19 At present, however, there is only limited data on the maturational changes in mechanical junctions that accompany differentiation of the intercalated disk. Such information is important, because it may provide insight into the mechanisms underpinning the patterning and maintenance of intercellular electrical connectivity in the normal and diseased heart. Therefore, the present study set out to determine, in detail, the sequential patterns of distribution and expression of gap junctional and cell adhesion junctional components during the differentiation of the intercalated disk in the developing rat and canine ventricle.
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Materials and Methods |
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Experimental Animals and Tissue Preparation
Rat embryos at 15 days after conception and rats at 1, 5, 10, 20, 40, 70, and 90 postnatal days were obtained from timed-pregnant Sprague-Dawley rats. For light microscopic analyses, three hearts from each of the stages studied were dissected out and processed immediately for methanol freeze substitution as follows: Specimens were immersed in liquid nitrogen–cooled isopentane for 1 minute, transferred to -80°C methanol for 7 days, and subsequently warmed in three 24-hour steps at -20°C, 0°C, and 20°C. After methanol freeze substitution, hearts were oriented, embedded in paraffin, and serially sectioned as detailed previously.16 Ventricular tissues for immunoelectron microscopy were processed as detailed in Green et al.26 Frozen sections (10 µm) were cut from paraformaldehyde-fixed (4%, 3 hours in PBS) blocks of ventricular myocardium from normal 1-month-old (2 animals) and 3-month-old (1 animal) dogs, kindly provided by Dr N. Sydney Moise (Cornell University, Ithaca, NY). All light and electron microscopic analyses of rat and dog tissues were confined to the left ventricular wall.
Antibodies
Gap Junctions
Two antibodies directed against gap junctional connexin43 (Cx43) were used. The specificity of the first reagent (HJ) for cardiac-type gap junctions has been extensively characterized by immunoblotting, ultrastructural, and immunohistochemical methods.26 27 28 29 The second anti-Cx43 antibody, designated Z43, is a commercially available mouse monoclonal antibody (Zymed Laboratories Inc). We and the manufacturer have confirmed antibody specificity by Western blotting, and immunolabeling of cardiac gap junctions by Z43 has been shown in earlier studies.29
Desmosomes
The anti-desmoplakin antibody, DP145, is a rabbit polyclonal antibody raised against a trpE bacterial fusion protein containing half of region B and all of region C of the desmoplakin C-terminus30 and gives immunofluorescent labeling patterns typical of desmosomes in foreskin epithelium.31 Specificity of DP145 was further verified by Western blotting (see Fig 1
) of a desmosome-enriched preparation from bovine muzzle.
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Adherens Junctions
The pancadherin antibody recognizing adherens junctions, designated NC, was raised against the conserved C-terminal 24–amino acid residues of N-cadherin and was generously supplied by Dr Benjamin Geiger of the Weizmann Institute, Israel. This antibody gives specific immunofluorescent labeling of adherens junctions in cardiac intercalated disks from a wide range of mammalian species.19 32
Immunoconfocal Microscopy and Immunoelectron Microscopy
The specificity of the antisera HJ, DP145, and NC for ultrastructurally defined gap junctions, desmosomes, and adherens junctions, respectively, was confirmed by immunogold labeling on ultrathin sections imaged by a Phillips EM301 electron microscope, as detailed in Green et al.26 For immunoconfocal microscopy, labeling by indirect immunofluorescence was carried out using protocols and controls described previously.16 28 29 A microwave-based antigen retrieval protocol14 was used before labeling with NC antibodies. Double labeling of intercellular junctional molecules and actin was achieved by treating Z43-, HJ-, DP145-, or NC-immunolabeled sections with actin-binding phalloidin conjugated to rhodamine. Double labeling for Cx43 gap junctions and cell adhesion junctions was carried out with mouse monoclonal Z43 in simultaneous incubations with either DP145 or NC rabbit polyclonal antibodies; multiple labeling protocols described previously were used.33 34 Imaging was done on Biorad MRC-500 (Bio-Rad Microscience) and Leica 4D TCS (Leica UK Limited) scanning laser confocal microscopes.
Image Analysis and Statistics
The PC-Image (Foster-Findlay Associates)–based methods used in the present study for quantitative analyses of junctional area from single confocal optical sections of immunolabeled tissue have been detailed in previous reports19 28 29 and validated by direct comparison to freeze-fracture electron microscopic morphometry.26 The proportion of immunofluorescence occurring at cell termini (ie, at intercalated disks) for each junction type was quantified on 12 optical sections from left ventricular myocardium for each developmental stage (ie, 3 ratsx4 images per rat). The method used was based on a protocol outlined by Peters et al.19 Briefly, random fields of longitudinally sectioned myocytes were analyzed to determine the proportion of label at intercellular abutments lying transverse to the long axis of the myocytes (ie, forming intercalated disks) relative to overall immunolabeling. For the purpose of this analysis, putative disks were defined as linear arrays of four or more fluorescent spots, orthogonal to the myocyte long axis. Phase-contrast microscopy was used to further confirm the colocalization of the linear arrays of fluorescent puncta with zones of end-to-end abutment between myocytes.
All statistical analyses and tests were carried out using Minitab software (Minitab Inc). First, overall variation between junction molecules (JMs) was analyzed within each of the seven time points. Before ANOVA, graphical analysis (using the Minitab %Normplot macro) of raw data and ANOVA residuals indicated that the data showed no significant deviations (P>.05) from normality. Tests of homogeneity of variance (using the Minitab %Vartest macro) revealed no significant differences (P>.05) between variances within the 1-, 5-, 10-, 20-, 40-, and 90-day time points, as assessed by confidence intervals for factor standard deviations and Bartlett's test of homogeneity of variance. Arcsin transformation (arcsin 
Y/100)35 of the 70-day data enabled variance within this time point to assume homogeneity (P>.05).
Initially, the factors used in the model were as follows: JM, RAT, and JMxRAT.
However, because the F values for RAT and JMxRAT never reached significance (P>.05) within any of the seven time points, these factors were relegated to the error term. The final model used for ANOVA was simply JM.
In the Table, these analyses of overall variation in gap junction, desmosome, and adherens junction distribution within time points are referred to as ANOVA set A. Before assessment of confidence levels for JM variation, the P values for each of the time points were adjusted conservatively according to Bonferroni.35
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Next, we assessed variation of JM in the three pairings of JMs possible within each time point (ie, Cx43 versus DP, Cx43 versus NC, and DP versus NC). Again, each data set was tested for homogeneity of variance. Of the 21 data sets thus analyzed, 19 showed homogeneity of variance (P>.05), and ANOVAs were carried out for these on untransformed data. Arcsin transformation significantly improved homogeneity of variance in the remaining two data sets corresponding to the 70-day time point Cx43 versus DP (to P>.05) and Cx43 versus NC (to P>.01) comparisons. These 21 pairwise comparisons are referred to as ANOVA set B in the Table
. Before assessment of confidence levels for JM variation, the P values for the three hypotheses tested within each time point were adjusted according to Bonferroni.35
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Results |
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In adult ventricular myocardium, electrical and mechanical intercellular junctions show correlated distributions at intercalated disks (Figs 1
and 2). In the present study, our main aim was to characterize the differentiation of this arrangement, in particular, to determine the spatial patterning of gap junctions and cell adhesion junctions during disk formation. As a first step, immunoelectron microscopy was used to verify that gap junctions, desmosomes, and adherens junctions between working ventricular myocytes were specifically and independently discriminated from one another by HJ/Z43 (anti-Cx43), DP145 (anti–desmoplakin-1), and NC (anti–N-cadherin), respectively.
Fig 1 illustrates that accumulations of immunogold particles are confined to the appropriate junction type for each of the antisera without significant background labeling. Furthermore, there is no cross-reaction of the antisera for inappropriate junctional types or significant association of gold label with nonjunctional plasma membranes. Consistent with previous studies of rats26 and humans,19 extensive searching of immunogold-labeled ultrathin sections revealed that all gap junctions identifiable between ventricular myocytes were labeled by anti-Cx43 antibodies. Although two other gap junction proteins, connexin40 and connexin45, have been identified in rat cardiac tissues,29 36 labeling of these does not contribute to establishing the spatial distribution of gap junctions in rat working myocardium. Connexin40 is not present in the working ventricular myocardium of the postnatal and adult rat,29 36 37 38 and connexin45, which is present only at extremely low levels in the ventricular myocardium, colocalizes with Cx43-containing gap junctions.36
After establishing the specificity of the antisera, we examined the immunolabeling patterns of the three intercellular junctions over the postnatal period. In Fig 2, the distributions of immunolabeled gap junctions (Cx43), desmosomes (desmoplakin), and adherens junctions (N-cadherin) are compared at matching locations on adjacent wax sections of rat ventricle at 1, 20, and 90 postnatal days. The ventricular myocytes are sectioned in longitudinal orientation, and myofibril arrays are delineated by double labeling with actin-binding rhodamine-phalloidin. At 1 postnatal day, immunolabeling of all junction types appeared uniformly distributed, occurring in relatively dispersed patterns across the membranes of ventricular myocytes (Fig 2a through 2c). At 20 postnatal days, however, a striking difference in the distribution of gap junctions and cell adhesion junctions is apparent (Fig 2d through 2f). Whereas Cx43-positive gap junctions show uniform concentrations at both zones of side-to-side and end-on (ie, intercalated disklike) contact between myocytes, desmoplakin-positive desmosomes and N-cadherin–positive adherens junctions are most prominently localized at cell termini. However, this difference in distribution is transient (by 90 postnatal days, immunolabeling of all three junction types is concentrated at intercalated disks; Fig 2g through 2i).
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To more precisely determine (1) when the difference in gap junction and cell adhesion junction distribution first emerges and (2) when the adult configuration is reached, a quantitative analysis of the proportion of junctional fluorescence located at zones of transverse abutment between myocytes (ie, cell termini/intercalated disks) was carried out (Table and Fig 3). At 15 embryonic days, clearly defined cell termini are few in number and difficult to discern on the basis of histology. However, between 1 and 5 postnatal days, as ventricular myocytes elongate and begin to develop well-aligned myofibrillar contractile apparatus, zones of end-to-end or terminal abutment between myocytes (ie, early disklike zones) are readily discriminated from regions of lateral apposition. Fig 3 indicates that variation in relative levels of Cx43, desmoplakin, and N-cadherin at myocyte termini show complex nonlinear patterns of change over postnatal life. Proportional levels of cell termini–associated desmosomes (red squares) and adherens junctions (green triangles) share similar curvilinear trends over postnatal development and show their most rapid increase between 1 and 20 postnatal days. This is in contrast to the proportional levels of gap junctions (blue circles) at cell termini, which demonstrate slower increases between 1 and 20 days. Compared with gap junctions, significantly higher levels (P<.01) of desmosomes and adherens junctions are localized to cell termini from as early as 5 postnatal days (Table). This difference between the two cell adhesion junctions and gap junctions is maintained at above the 5% level of confidence (P<.05) for time points up to 40 postnatal days. Only at 90 postnatal days, when postnatal growth is nearly complete, do the proportions of all three junction types at intercalated disks become statistically similar (P>.05).
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Finally, in order to confirm that this process was not restricted to rodents, studies were carried out on ventricular myocardium from dogs. As with the young rat, the distribution pattern of gap junctions was found to diverge from both that of desmosomes and adherens junctions in 1- and 3-month-old dogs. This is illustrated for the 3-month-old dogs by double labeling in Fig 4a, in which localization of desmosomes (red) to cell termini is contrasted with more uniformly spread distributions of gap junctions (green). Comparison of panels a and b of Fig 4 indicates that the progression of the process in a dog at 3 months (Fig 4a) is similar to that of a 20-day-old rat (Fig 4b).
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Discussion |
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The present study demonstrates a transient but striking divergence in the distributions of gap junctions and cell adhesion junctions during postnatal maturation of rat and dog ventricular myocytes. Although desmosomes and adherens junctions rapidly establish polarized distributions at myocyte termini after birth, gap junctions initially retain dispersed distributions across the membranes of myocytes. Over postnatal growth, the distribution of gap junction distribution becomes progressively more correlated with cell adhesion junction–rich zones of membrane, culminating in the formation of intercalated disks characteristic of adult ventricular myocardium.
In adult ventricle, gap junctions show precise three-dimensional patterns of colocalization with cell adhesion junctions at intercalated disks, zones of end-to-end electromechanical abutment between myocytes.2 3 4 12 19 21 28 A number of workers have now shown that this pattern of gap junctional organization is not present in newborn mammals but differentiates over postnatal life in association with progressive decreases in side-to-side intermyocyte connections.15 16 17 18 19 20 It has been proposed that this process is vital to the emergence of uniform anisotropic conduction of action potential, an electrophysiological characteristic of mature myocardial tissues, and essential for rapid and efficient depolarization of cardiac muscle.8 19 A previous study of humans during childhood revealed progressive reorganization of gap junctions and adherens junctions from dispersed to disk configurations, although a transient dissociation between the two junction types was not detected.19 The present study has demonstrated the dissociation in two evolutionarily distinct families; thus, this apparent difference is unlikely to be due to a species variation but is more probably attributable to the inherent constraints in following the time course in the available human surgical specimens. In practice, ethical considerations make an optimal temporal sequence of human specimens impossible to obtain, and this limits the ability to resolve transient events during the time course. Necessarily, in a study by Peters et al,19 comparison of gap and adherens junction distribution during postnatal development of human heart was confined to a number of individuals with ages ranging from 9 weeks and 7 years. It is conceivable, then, that the perinatal divergence in different intercellular junction types we describe in rats and dogs may have not been resolved in this previous study of humans. In the present study, with the benefit of the temporal precision attainable in animal studies, we were able to resolve changes in spatial organization using a comprehensive time series in which three individuals were sampled at each of eight time points over the 90 days of postnatal growth.
Several studies present contrasting interpretations concerning the relationships between intercellular junctions during their assembly.22 23 24 39 40 41 Although the present work does not directly address the question of assembly of these junctions from their molecular components, it is relevant to the reorganization and maintenance of myocyte intercellular junctions subsequent to assembly. One explanation of our results is that over postnatal growth, gap junctions located at intercalated disks are preferentially retained over those at side-to-side (ie, nondisk) zones of contact. At disks, high concentrations of desmosomes and adherens junctions provide sites of stabilized sarcolemma, potentially favorable for the preservation of gap junctions located either close to or within the body of the disk. Nondisk-localized gap junctions, distant from the stabilizing influence of cell adhesion junctions, may be selectively internalized or degraded because of an increasing vulnerability to shearing forces generated by the contraction of neighboring myocytes. This process would explain the conspicuous increase in frequency of endocytosed gap junctions that has been described in the neonatal mammalian ventricle by a number of different workers.19 42 43 The possibilities also exist that gap junctions diffuse within the plane of the membrane and are trapped when they enter the stabilized zones at the disks or that their relocalization to these sites is mechanically driven. Whatever the mechanism underpinning this process, extrapolation of the trends illustrated in Fig 3 indicates that it may not necessarily cease at the end of maturational growth. Indeed, it may continue throughout the lifetime of the animal, albeit at a slower rate. Confirmation of this is clinically relevant, as increases in anisotropy in the aging myocardium have been linked to the propensity of the older heart for arrhythmias and disturbances of conduction.6 Understanding developmental interrelationships between electrical and mechanical junctions may provide insight into other cardiac pathologies, including those associated with localized disruptions to gap junctional distribution, such as at infarct border zones9 10 11 12 or regions of myofiber disarray.14 Indeed, a recent study of zones of myofiber disarray in adults with hypertrophic cardiomyopathy reported abnormal dissociation of gap junctional and desmosomal contacts,14 highly reminiscent of the transient divergence that we report here in gap junction and cell adhesion junction distribution during development of the normal heart.
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Acknowledgments |
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Dr Gourdie is a Basil O'Connor Scholar of the March of Dimes Birth Defects Foundation and gratefully acknowledges support for this project from the American Heart Association (South Carolina Affiliate) and British Heart Foundation (FS92005). Drs Thompson and Gourdie are supported by National Institutes of Health project grant HL-50582. Drs Angst and Magee are supported by British Heart Foundation project grant PG/94088; Dr Severs, by British Heart Foundation project grant PG/93136. Work at the National Institute for Medical Research was also supported by the Medical Research Council (UK). The comments on this manuscript by Drs Roger Markwald, Takashi Mikawa, Wanda Litchenberg, and Leonard Eisenberg are acknowledged with gratitude. We thank Dr Thomas Trusk and Joshua Spruill for helping to prepare figures and Patricia Germroth for assistance with immunohistology. We are grateful to Dr Stewart Denslow for his statistical consultation. The provision of canine heart samples by Dr N. Sydney Moise (Cornell University, Ithaca, NY) is acknowledged with gratitude.
Received December 19, 1995; accepted October 18, 1996.
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