Morphological analysis of contracting and quiescent adult rabbit cardiac myocytes in long-term culture.код для вставкиСкачать
THE ANATOMICAL RECORD 227:285-299 (1990) Morphological Analysis of Contracting and Quiescent Adult Rabbit Cardiac Myocytes in Long-Term CuIture MARLENE L. DECKER, DAVID G. SIMPSON, MONICA BEHNKE, MELISSA G . COOK, AND ROBERT S. DECKER Departments of Medicine (Cardiology) (M.L.D., M.B., M.G.C., R.S.D.), Cell Biology and Anatomy (D.G.S., R.S.D.) and Feinberg Cardiovascular Research Institute (R.S.D.), Northwestern University Medical School, Chicago, Illinois ABSTRACT Isolated rabbit ventricular cardiac myocytes adapt readily to primary culture. As the myocytes spread and flatten over the culture substratum, the myofibrillar apparatus retains a “rod-like” orientation. Development of contractile activity is crucial in the maintenance of the integrity of the myofibrillar apparatus during prolonged culture. Myocytes that fail to beat display morphological indications of atrophy; conversely, myocytes that commence beating show no such morphological signs of myofibrillar disorganization. The subcellular organization of other elements of the contractile apparatus, including the transverse tubular system and the sarcoplasmic reticulum, retain their structural relationship with the myofibrils in beating myocytes but not in quiescent cells. Cultured adult myocytes represent a n important model to investigate the influence of mechanical factors on the organization and maintenance of the adult cardiac phenotype. The isolation and culture of adult cardiac myocytes represents a n increasingly popular approach to investigate directly phenomena that regulate myocyte behavior under precisely defined conditions (Jacobson and Piper, 1986; Liebermann et al., 1987). Such a model system offers distinct advantages in analyzing the influence of neurohumeral and mechanical factors on myocyte structure and function which cannot be readily retrieved from perfused or intact hearts. Previous reports have demonstrated that the majority of freshly isolated calcium tolerant, adult r a t cardiac myocytes gradually rounded-up when placed in primary culture (Jacobson, 1977; Claycomb and Palazzo, 1980; Schwarzfeld and Jacobson, 1981; Moses and Claycomb, 1982b; Nag et al., 1983; Eppenberger et al., 1988; Piper et al., 1988). As the myocytes “rounded,” they lost many of their myotypic features, including elements of the intercalated disc, organized myofibrils, and their accompanying sarcoplasmic reticulum; the transversetubular system (T-tubules), and numerous mitochondria, many of which were apparently blebbed and released from the sarcolemmal surface (Moses and Claycomb, 1982b, 1984; Nag et al., 1983). Although many of these rounded myocytes subsequently attached to a substratum (Piper e t al., 19881, commenced beating (Bugaisky and Zak, 1989; Spahr et al., 19891, and reacquired a morphology that closely resembled the adult myocyte from which they were derived (Moses and Claycomb, 1982a, 1984; Nag et al., 19831, the question persisted-how favorably did these “redifferentiated” (Jacobson and Piper, 1986) myocytes compare with their in vivo counterparts (Jacobson and Piper, 1986; Bugaisky and Zak, 1989; Eppenberger et al., 1988)? Although considerable effort has been expended de0 1990 WILEY-LISS, INC. scribing the structural alterations that accompany the culture of “redifferentiating” r a t cardiac myocytes (Jacobson, 1977; Claycomb and Palazzo, 1980; Schwarzfeld and Jacobson, 1981; Nag e t al., 1983; Moses and Claycomb, 1982a,b, 1984, 1985; Jacobson and Piper e t al., 19861, few attempts have been made to compare and contrast such observations with myocytes isolated and cultured from other species (Cooper et al., 1986; Haddad e t al., 1988a), nor has the adaptation of the “redifferentiated” myocyte been compared to the “rapidly attached” heart cell (Jacobson and Piper, 1986). Two aspects of in vitro rat myocyte behavior required further comparison with myocytes cultured from other laboratory animals. First, most rat myocytes rounded-up during the first week of culture, resulting in disruption of the contractile apparatus (Nag et al., 1983; Moses and Claycomb, 1982a,b, 1984; Eppenberger et al., 1988; Piper et al., 1988; Bugaisky and Zak, 1989; Spahr et al., 1989), regardless of the nature of the culture environment (Piper et al., 1988). Second, as rat myocytes entered the second week of in vitro life, they commenced beating spontaneously (Bugaisky and Zak, 1989). Neither feline (Cooper et al., 1986; Decker et al., 1989a,b) nor rabbit (Haddad et al., 1988a) cardiac myocytes displayed these properties; rather, they maintained a rod-shaped configuration while spreading over the culture substratum and exhibited no contractile activity except when cultured a t high densities. Received June 30, 1989; accepted November 2, 1989. Address reprint requests to Marlene L. Decker, Department of Medicine (Cardiology), Searle 2-575, Northwestern University Medical School, 303 E Chicago Ave., Chicago, IL 60611. M.L. DECKER ET AL. 286 The principal objective of the present investigation was to describe the morphological changes that developed in the contractile apparatus when rabbit myocytes were maintained at a low density in a quiescent state and to compare and contrast these observations with those obtained from high density, synchronously beating myocyte preparations. Our observations were focused on interpreting the evolving changes in cytoarchitecture and the changing relationships between the organelles that constitute the contractile apparatus of the heart cell in quiescent and contractile myocytes. The principal goal of this investigation was to evaluate how mechanical factors influenced the maintenance of the adult phenotype in vitro. MATERIALS AND METHODS Isolation and Culture of Adult Rabbit Myocytes and/or transmission or scanning electron microscopy (Decker et al., 1988a,b). The organization of the myofibrillar apparatus was also evaluated by staining briefly fixed coverslips with rhodamine-conjugated phalloidin (Molecular Probes, Junction City, OR), which specifically binds to f-actin (Wulf et al., 1979) or with a monoclonal (CM-52) anti-myosin antibody developed by Clark et al. (1982). Coverslips were mounted with Aquamount for viewing with a Leitz Orthoplan fluorescence microscope equipped with fluorescein and rhodamine optics. The nature and distribution of attachment sites was assessed on live cells with interference reflection microscopy a s described previously (Decker et al., 1988a). Petri dishes were briefly rinsed in serum-free MEM and preserved in situ according to the transmission electron microscope (TEM) protocols briefly outlined below (Decker et al., 1988a). Routine preparations were fixed with 4% glutaraldehyde buffered in 0.1 M sodium cacodylate buffer (pH 7.4) for 2-4 hours, rinsed in three changes of 0.1 M cacodylate buffer plus 7.5% sucrose, postfixed in 2% osmium tetroxide for 1 hour, rinsed again and then en bloc stained in aqueous uranyl acetate (0.5%) to enhance membrane contrast. Other cultures were treated with 2% tannic acid or osmium ferrocyanide (0.8%)to stain the glycocalyx and intensify myofibrillar contrast or to stain the sarcoplasmic reticulum (Decker et al., 1988a). The preparations were dehydrated and then infiltrated with 50150 mixture of ethanol and Medcast (Ted Pella, Inc., Tustin, CA); the dishes were then drained of epoxy and a thin layer of fresh plastic added. At this juncture, Beem capsules were inserted into the epoxy and the plastic was polymerized for 18 hours at 55°C. The dishes were then removed and the capsules filled with resin and polymerized for a n additional 2 days a t 55°C. En face, transverse and sagittal thin sections (Decker et al., 1988a) were stained with uranyl acetate and lead citrate and viewed with a JEOL 100 CX electron microscope. Myocytes plated onto coverslips were employed to study changes in cell surface topography with the scanning electron microscope (SEMI. Coverslips were rinsed twice in serum-free medium and fixed in 4% glutaraldehyde for 1 hour a t room temperature. The cells were then rinsed in 0.1 M cacodylate buffer plus 7.5% sucrose, postfixed in buffered 1% OsO,, and stabilized with 1% tannic acid and 2.5% uranyl acetate. The samples were then dehydrated, critically point dried, sputter coated with gold, and viewed in a JEOL 35CF scanning electron microscope operating at 10 kV (Decker et al., 1988a). Ventricular cardiac myocytes were enzymatically isolated from 1.8 kg adult male New Zealand white rabbits by a protocol previously described in considerable detail (Haddad et al., 1988a). Rabbits were briefly heparinized (500 U/kg body wt.), anesthetized with sodium pentobarbital (30 mglkg body wt.), and the hearts removed and mounted onto a Langendorff perfusion apparatus in a sterile laminar flow hood. Hearts were perfused briefly with nominally Ca2+ free KrebsRinger bicarbonate (KRB) buffer containing 15 mM HEPES (pH 7.35) followed by KRB supplemented with 80 U/ml class I1 collagenase (Worthington Biochemical, Freehold, NJ), 0.5 mgiml testicular hyaluronidase (Sigma Chemical Co., St. Louis, MO), and 20 pM Ca2+ (Powell et al., 1980) a t 37°C in a 5% co,/95%02 atmosphere. When the heart became flaccid (about 30-45 min), it was removed, teased apart in KRB plus 1% bovine serum albumin (BSA, ICN Immunobiologics, Lisle, IL), filtered through a 550 pm nylon mesh to eliminate connective tissue debris and centrifuged (50g for 90 seconds) to remove rounded, non-viable cells. The pelleted myocytes were resuspended in KRB plus 2% BSA, re-centrifuged, and the pellet finally diluted with Eagle’s minimum essential medium (MEM) with Earle’s salts (Northwestern University Media Center) supplemented with 5% fetal calf serum and antibiotics. Myocytes were plated onto 35 mm petri dishes (Corning, NY) or coverslips previously coated with laminin (20 g/ml) at either a low (1x lo4 cellsldish) or high (1x 10 cells/dish) density. Proliferation of nonmyocytic cells was inhibited by adding 10 pM cytosine1-p- -D-arabinofuranoside (Sigma Chem. Co., St. Louis, MO) to the culture medium (Haddad e t al., 1988a). Cultures were incubated a t 37°C in a humidified atmosphere of 5%co,/95% air, and the culture medium was exchanged every other day. The quality of the cultures RESULTS was monitored by measuring adenosine triphosphate (ATP) and creatine phosphate (CrP) content, the reProperties of Freshly Attached Myocytes lease of creatine kinase (CK) and lactate dehydrogenase (LDH) enzyme activity into the culture medium Calcium tolerant myocytes adhere rapidly to a lamiand the ability of the myocytes to exclude trypan blue nin-coated substratum (Fig. 1).Such freshly prepared a s described previously (Haddad et al., 1988a). myocyte cultures exclude trypan blue, release little LDH activity into the culture medium, and possess Light and Electron Microscopy “normal” levels of CrP and ATP in the appropriate proCoverslips and petri dishes were selected after 1hour portion (Table 1). Adherent myocytes also appear to be and 1 , 4 , 7 , 1 4 ,and 28 days of culture and prepared for modestly loaded when compared to age-matched coundifferential interference (DIC), interference reflection terparts maintained unattached in suspension. Mea(IR), phase contrast (PC), fluorescence microscopy, surements of intrasarcomeric dimensions obtained P CONTRACTION AND MYOFIBRILLAR STRUCTURE 287 Figs. 1-6. The morphology of freshly attached (1 hr) cardiac myocytes is illustrated. Cells adhere randomly, but frequently appear in close contact (arrows) with one another (Fig. 1).Phase contrast (Fig. 2) and IRM (Fig. 3) images demonstrate the close contacts (arrows) that develop along the length of the basal myocyte surface (Fig. 3), running parallel to Z-lines (Fig. 2). Minute adhesion plaques (Figs. 2, 5) also appear at the distal ends (large arrowhead) of the cells. Such cells display a dimpled cell surface (bracketed arrows) (Fig. 4)with small blebs (arrowheads) emerging between the costameres (Fig. 5). Thin-sections (Fig. 6) reveal that these blebs contain “normal” mitochondria (m); also illustrated are the internalized elements (arrows) of the intercalated disc. Sarcomeres are preserved in a modestly contracted state. Z-lines; I-band; A-band. Fig. 1, X 450; Figs. 2,3, X 1,200; Fig. 4, X 840; Fig. 5, X 5075; Fig. 6, X 12,000. from suspended and attached cells reveals that a significant reduction (6.4*0.1%)in Z-2 widths is apparent in adherent myocytes when both preparations are preserved identically. The rapid development of minute attachment plaques (Figs. 2 , 5 ) at the distal ends of the myocyte and the appearance of close contacts that parallel the Z-lines (Figs. 2 , 3 ) coincide with apparent tension development as evidenced by a reduction in I-band 288 M.L. DECKER ET AL. TABLE 1. Changes in high energy phosphate content and cell viability during primary culture of adult rabbit cardiac myocytes LDH Preparation Intact Heart Freshly isolated myocytes 1-Hr myocyte culture 1-Day myocyte culture 7-Day myocyte culture 14-Day myocyte culture Non-Beating 28-Day myocyte culture Non-Beating 14-Day myocyte culture Beating 28-Day myocyte culture Beating Trypan Blue %Rounds3 ND 15.6 2 2.4 4.9 ? 0.4 1.7 t 0.3 1.4 ? 0.2 Exclusion’ ND 74.2 t 4.9 95.3 t 0.4 98.7 t 0.6 98.1 ? 0.5 1.6 2 0.2 1.5 t 0.1 98.4 t 0.4 7.9 t 1.1 25.6 t 2.0 1.5 k 0.3 1.5 t 0.2 98.7 2 0.4 3.4 t 0.2 40.2 t 4.5 26.6 2 1.2 1.5 t 0.2 2.5 t 0.4 95.4 t 0.2 4.8 42.7 t 2.4 29.4 2 1.9 1.5 k 0.3 2.1 t 0.4 96.3 t 0.4 6.4 t 1.5 CrP ATP CrPIATP 45.5 2 1.8 34.6 k 4.1 47.3 3.8 45.7 t 3.1 46.8 t 4.1 28.4 5 1.3 20.7 t 3.0 29.7 k 1.7 34.4 5 1.3 28.1 t 1.9 1.6 t 0.1 1.7 t 0.4 1.6 t 0.1 1.3 2 0.1 1.7 2 0.1 39.1 t 3.5 24.7 2.2 38.7 2.5 * 2 2 Release’ ND 34.2 t 6.3 5.1 2 0.7 5.7 t 0.5 6.4 2 0.8 2 2.1 ‘Values are means ? S.E. of 10-15 replicate cultures that are exposed to 10 uM Ara-C or the mean ? S.E. of five whole hearts or myocyte preparations. Adenosine triphosphate (ATP) and creatine phosphate (CrP) are expressed in nanomoles per mg. protein. Lactate dehydrogenase (LDH) activity is expressed as % of total LDH released into the culture medium after a 2 hr. incubation in fresh medium. % of cells that do not stain with trypan blue after a 2 hr. incubation in fresh medium. ND Not determined ‘Values are means ? S.E. for 5 cultures exposed to lOuM Ara-C. Values represent mean number of round cells as % of total counted cells for 5 cultures. width and, therefore, in sarcomere length (Fig. 6). SEM images also depict the existence of randomly scattered protrusions on the sarcolemmal surface that develop between adjacent costameres as myocytes adhere to the substratum (Figs. 4, 5 ) . Thin sections reveal that mitochondria present in such blebs appear indistinguishable from those that intermingle among neighboring myofibrils (Fig. 6). Morphology of Cultured Myocytes After attachment to laminin, rod-shaped myocytes gradually spread into a highly flattened configuration. Although this dramatic morphological transformation accompanies their adaptation into primary culture, neither ATP or CrP levels nor the ratio of these highenergy metabolites display any significant reduction (Table 1). Moreover, the release of CK and LDH activity is negligible after 1day and greater than 95% of the cultured myocytes exclude trypan blue during the course of culture. Furthermore, unlike rat preparations that display significant cell rounding, regardless of the extracellular matrix molecules that the cells are cultured on, fewer than 10% of the rabbit myocytes round during prolonged culture (Table 1). Day 1 quiescent myocyte cultures Within 24 hours of plating, the distal ends of the adherent myocytes display well-developed adhesion plaques when observed with interference reflection microscopy (IRM) and SEM (Figs. 9-11). IRM further reveals the continued presence of close contacts along the length of the myocyte, which are oriented perpendicular to the long axis of the cell (Fig. 9). The terminal edges of these distal plaques further demonstrate close apposition with the substratum and the contour of the cell surface suggests the development of subsarcolemma1 actin stress fibers (Fig. 11). Thin sections of this region reveal that many of the Z-lines are disrupted and that resident myofibrils lose their classical sarcomeric organization (Fig. 12). Proximal to the adhesion sites (Fig. 12), the structural organization of the myocyte closely resembles that of the freshly isolated cell with the myofibrillar apparatus retaining its registry and T-tubules assuming their normal morphological relationships. Fluorescence microscopy further illustrates that the organization of the myofibrillar apparatus, as visualized with rhodamine phalloidin or monoclonal anti-myosin antibodies, appears normal (Figs. 7, 8). The mean length of these sarcomeres measures 1.78k0.05 pm ( n = 351, similar to values obtained 24 hours earlier, implying that the heart cells remain “loaded.” Lastly, at sites where adjacent myocytes are closely apposed to one another, no evidence could be garnered for the reassembly of specialized intercellular junctions at this time (Fig. 12). Day 7 quiescent myocyte cultures Although rabbit myocytes retain their rod-shape through the first week of culture, the distal ends of the cells transform into fan-shaped plaques (Fig. 13-16) that can be easily identified as adhesion plaques by interference reflection microscopy (Fig. 15).Such zones of contact disclose the presence of close contacts; focal contacts of the variety that characterize cardiac fibroblasts are not a prominent feature in the contact zones of quiescent myocytes. Lateral cell processes, likewise, develop and spread over the substratum at this time (Fig. 16). By day 7 in culture, the sarcolemmal surface of the myocyte has acquired a relatively smooth texture with only faint impressions of the subjacent myofibrillar apparatus apparent in SEM images (Fig. 16) and only a few sarcolemmal blebs of the variety encountered at day 1are visible. Significant alterations in the structure and organization of the contractile apparatus are apparent in myocytes maintained for 7 days in vitro. Fluorescent distribution of myofibrillar proteins reveals that contractile elements are restricted primarily to the cylindrical portion of the myocyte where their registry is disrupted (Figs. 13, 14). Furthermore, such myocytes CONTRACTION AND MYOFIBRILLAR STRUCTURE Figs. 7-1 2. The changes in myocyte structure after one day in vitro. Actin (Fig. 7) and myosin (Fig. 8 ) staining reveals a well ordered rnyofibrillar apparatus (M) which is confirmed in thin-sections (Fig. 12). IRM and SEM reveal that distal attachment plaques (arrowheads, Figs. 9,10,11) enlarge and close contacts are extensive (arrow, Fig. 9) along the length of the rnyocyte. In the plaque regions (Fig. 12), myofibrillar order is disrupted (arrows) although thick and thin fila- 289 ments do project into the plaque and can be observed just below the cell surface in SEM profiles (arrows, Fig. 11). The cell surface is also decorated with a few blebs (arrows) similar to those seen earlier (Fig. 10). Areas of close contact also develop between adjacent cells (arrowheads) but no junctions are apparent. (Fig. 12). Figs, 7, $, x 650; Fig. 9, x 1,200; Fig. 10, x 975; Fig. 11, X 3,300; Fig. 12, X 10,600. display sarcomeres whose Z-lines range from being the appearance of cytoplasmic “wedges” that develop partially intact to almost nonexistent (Figs. 17, 18). amongst the contractile elements. Such regions exhibit Myofibrillar register is progressively interrupted by parallel arrays of rough endoplasmic reticulum and nu- 290 M.L. DECKER ET AL. Figs. 13-20. Fluorescent, IRM, SEM and thin-section profiles of myocytes cultured for 7 days. In such myocytes (Fig. 16) both actin (Fig. 13) and myosin (Fig. 14) lose their normal registry in more proximal portions of the cell. Large fan-shaped distal adhesion plaques are well developed (Figs. 15, 16).Close contact zones (arrows) are only apparent in the periphery of the plaque while contact sites along the length of the myocyte (arrowhead) appear to be receeding and are being replaced by punctate focal contacts (Fig. 15). E n face (Fig. 17, 18) and transverse (Figs. 19, 20) sections obtained from the rod-shaped portion (Figs. 17,181; the rod-plaque junction (Fig. 19) and the plaque area, proper (Fig. 20) from myocytes like the one illus- trated in Fig. 16, reveal major changes in myofibril (M) structure and T-tubular organization. Some 2-lines are disrupted (open arrowhead) while others are nearly intact (arrowhead) (Figs. 17, 18). T-tubules retain their association with Z-lines and junctional sarcoplasmic reticulum (Fig. 181, but in the peripheral plaques, sarcolemmal invaginations (arrows) are apparent but not closely associated with sarcoplasmic reticulum or myofibrils (Fig. 19, 20). Parallel arrays of rough endoplasmic retriculum (RER) also appear at 7 days as the cytoplasmic compartment increases in size (Fig. 17). c, caveolae. Figs. 13, 14, x 600; Fig. 15, x 1,500; Fig. 16, x 1,500; Fig. 17, x 31,000; Fig. 18, x 25,000; Fig. 19, x 20,000; Fig. 20, x 25,300. CONTRACTION AND MYOFIBRILLAR STRUCTURE 291 merous polysomal profiles that are not normally en- display numerous caveolae (Fig. 24). Peripheral T-tucountered in adult myocytes (Figs, 17). bules also enlarge dramatically (Fig. 23) and become As the myocyte spreads, total reliance on frontal or decorated with caveolae (Figs. 24,25). These T-tubules en face images provides insufficient morphological in- are frequently found associated with elements of juncformation on the evolving subcellular reorganization; tional sarcoplasmic reticulum (Figs. 25, 26) and therefore, sagittal and transverse thin sections are em- aligned with myofibrillar Z-lines (Fig. 25). T-tubules ployed to clarify cell-substrate interactions; to evaluate and their neighboring junctional sarcoplasmic reticuthe structure of the myocyte processes, and, in partic- lum are also frequently observed but not always closely ular, to evaluate myofibrillar integrity. In sagittal sec- associated with neighboring myofibrils (Fig. 26). Nontions taken from the distal portions of a myocyte in the junctional sarcoplasmic reticulum retains its plexiform region of a n attachment plaque (Fig. 161, myofibrils morphology in these contracting heart cells (Fig. 23, rather abruptly lose their sarcomeric organization and inset). are reduced to arrays composed of thick and thin filaments, which in most instances, lack well defined ZQuiescent Myocytes material (Fig. 20). In areas of transition where the adNon-beating myocytes cultured for 2 weeks are hesion plaque is contiguous with the cylindrical spread extensively and are characterized by a n overt portion of the myocyte (Fig. 16), myofibrils display a disruption of the contractile apparatus when compared significant degree of disorder (Fig. 19). The sarco- to paired contracting myocyte preparations (See Figs. lemma reveal few caveolae on the apical and basal sur- 22, 28). Myofibrillar register is almost completely abfaces (Figs. 19, 20), unlike their in vivo counterparts. sent in these cells, with the remaining f-actin and myT-tubules retain their normal configuration and asso- osin assuming a variety of unusual configurations ciation with intact myofibrils (Fig. 18); however, in (Figs. 28, 29). Perhaps the most striking alteration in spread zones “T-tubule-like” invaginations develop on the contractile apparatus is a pronounced condensation the basolateral surface of the sarcolemma (Figs. 19, of actin that frequently develops in the perinuclear re20). T-tubules can be positively identified by the pres- gions of most myocytes and the appearance of minute ence of their accompanying junctional sarcoplasmic re- myofibrils which approach the dimensions of stress fiticulum. Non-junctional sarcoplasmic reticulum con- bers (Fig. 28). Some of these fibers possess myosin, but tinues to exhibit its “honeycomb” pattern when found the vast majority of this immunofluorescently detectin association with a n “intact” myofibril, but in regions able protein exists in a rather diffuse pattern throughwhere orderly myofibrillar structure is lost, the sarco- out the cytoplasm of the myocyte (Fig. 29). Relatively plasmic reticulum is transformed into a branching ar- “normal” myofibrils displaying uniformly spaced, Zray typical of smooth endoplasmic reticulum which in- lines (Fig. 27) are often observed in electron microtermingles with rough endoplasmic reticulum (Fig. graphs along with abnormal myofibrils with irregu17). larly spaced and distorted Z-lines (Figs. 27, 30). The abnormal myofibrils reveal few, if any, myosin thick Day 14 myocyte cultures filaments amongst the actin thin filaments (Fig. 30). By days 10-12 in culture, a few densely plated cul- The honeycomb organization of the sarcoplasmic retictures of myocytes develop spontaneous contractile ac- ulum is also radically altered, with it assuming a mortivity. The cultures commence beating synchronously phology reminiscent of tubular smooth endoplasmic reduring this interval with individual myocytes contract- ticulum in areas devoid of intact myofibrils, but ing a t a rate of 150 beats per minute. Such cultures displaying disorganized myofibrillar elements (Fig. retain their contractile properties for well over a 32). The T-tubules in such regions exhibit a complex month. The acquisition of contractile function is accom- branching pattern, but they do remain closely associpanied by no change in high energy phosphate metab- ated with elements of the junctional sarcoplasmic reolism or loss of viability (Table 1);conversely, other ticulum (Fig. 32). That portion of the sarcoplasmic paired myocyte cultures where a significant number of reticulum associated with the remaining intact myomyocytes have detached remain quiescent and fail to fibrils retains its plexiform morphology (Fig. 31). beat even when maintained indefinitely. Day 28 myocyte cultures Contractile Myocytes Contractile rnyocytes. Synchronously contracting 4A beating myocyte can be immediately distinguished from a non-beating cell by the organization of its myo- week-old cells display a linearly ordered myofibrillar apfibrillar apparatus. Furthermore, such cells retain a paratus (Figs. 33,34); moreover, ultrastructural images of rod-like configuration not unlike that seen in 7-day-old the myofibrils reveal no obvious differences in their orgamyocytes except that the fan-like adhesion plaque is nization or structure when compared to the myofibrils transformed into well developed, branching processes present in 14-day-old beating cells (cf. Fig. 23 with Fig. (Fig. 21). The contractile apparatus retains its integ- 34). These myofibrils do, however, project well into the rity in the central core of the cell and myofibrils can be processes of these flattened myocytes (Fig. 33). Adjacent observed penetrating into such processes (Fig. 22). The cells in such preparations display normal intercalated myofibrils of these beating cells display no evidence of discs (Fig. 34) in which preferential attachment of myothe “disrupted” Z-lines (Fig. 23, inset) that are such a fibrils can be demonstrated. All such myocytes are binuprominent feature of 7-day old quiescent myocytes cleate (Fig. 341, and no evidence for karyokinesis is de(Figs. 17, 18). The basal sarcolemmal invaginations tectable in rabbit myocyte cultures. The only other present a t 7 days appear to develop into large dilated notable structural changes apparent in the beating myocavitations of the basal cell surface (Figs. 23,241 which cytes are an apparent increase in polyribosome and rough 292 M.L. DECKER ET AL. Figs. 21-26. are SEM, fluorescent and TEM images of contractile myocytes cultured for 14 days. Beating cells spread extensively but still retain a central rod-shaped segment (Fig. 21). Numerous phalloidin-positive myofibrils are apparent and many project into cell processes (Fig. 22). E n face images reveal myofibrils (M) with intact 2-lines (Figs. 23, inset; 25); T-tubules (T) appear to develop as basolateral invaginations (arrowheads) of the sarcolemma (Figs. 23, 24). The glycocalyx of these T-tubules (arrowheads) stains with tannic acid demonstrating their continuity with the cell surface (Figs. 23, 25); such T-tubules are associated with junctional components of the sarcoplasmic reticulum (arrows) (Figs. 25, 26). The sarcoplasmic reticulum (SR) retains its plexiform pattern around myofibrils (Fig. 23, inset). Saggital sections also show apical and basal caveolae (c) (Fig. 24) and basal T-tubules (TI; coated vesicles (cv) are also visible (Fig. 24). D, desmosome. Fig. 21, x 600; Fig. 22, X 750; Fig. 23, X 10,000; Fig. 23 inset, x 10,000. Fig. 24, x 25,700; Fig. 25, x 35,500; Fig. 26, x 62,500. CONTRACTION AND MYOFIBRILLAR STRUCTURE Figs. 27-32. depict the organization of the contractile apparatus in quiescent myocytes cultured for 14 days. Myofibrils (M) are only rarely well developed (arrow) when viewed in en face sections (Fig. 27). Phalloidin- (Fig. 28) and myosin- (Fig. 29) stained cells illustrate a profound disruption of myofibrils with only a few small fibrils clearly displaying both proteins (arrows). Much of the actin condenses (Fig. 28) while myosin is distributed in a diffuse-reticular pattern (Fig. 29). Many of the myofibrils (Fig. 30) reveal few thick filaments 293 and the Z-lines are distorted (arrows). Other myofibrils appear normal and display a plexiform sarcoplasmic reticulum (*) encircling these myofibrils (Fig. 31). Myofibrillar breakdown is associated with a disruption in sarcoplasmic reticulum (SR) order, but T-tubules still maintain their close association with junctional elements of the sarcoplasmic reticulum (arrow) (Fig. 32). ID, intercalated disc, c, caveolae. Fig. 27, x 4,500; Figs. 28, 29, x 500; Fig. 30, x 21,600; Fig. 31, x 34,000; Fig. 32, x 31,000. 294 M.L. DECKER ET AL. Figs. 33 and 34.The distribution of myofibrils in 28-day cultured contractile myocytes. Transverse sections reveal that myofibrils (MI penetrate into the peripheral processes of two interdigitating cells (Fig. 33). En face images (Fig. 34) of the perinuclear (N) region of the cell depict well ordered myofibrils (M). Lysosomes (L) are also prevalent in such cells. ID, intercalated disc. Fig. 33, x 15,000; Fig. 34, x 8,000. endoplasmic reticulum content and the presence of numerous lysosomal residual bodies (Fig. 34). Quiescent myocytes. Myofibrils, such as they are, become distributed in a complex branching pattern throughout the myoplasm with many projecting into the spread processes of the flattened, non-beating myocytes (Fig. 35). At this juncture much of the condensed f-actin deposits present in 14-day-oldmyocytes (Fig. 28) are replaced by a lacy network of minute myofibrils. Unlike stress fibers, which are difficult to visualize in the non-beating cells, these minute myofibrils disclose a regular periodicity (Fig. 35). Myosin distribution remains essentially unchanged from that observed in 14-day-old quiescent myocytes (See Fig. 291, with much of the protein present in the cytoplasm and not associated with the minute myofibrils. Such structures lack well-defined Z-lines, display little ev- CONTRACTION AND MYOFIBRILLAR STRUCTURE Figs. 35-37. A disorganized contractile apparatus in 28-day old quiescent cultured myocytes. Phalloidin-positive cells reveal virtually no myofibrillar registry, rather branching actin filaments displaying a regular periodicity (arrowheads) are apparent (Fig. 35). TEM show these structures (arrows) to be myofibrils (Fig. 36). T-tubules are oc- 295 casionally encountered; the sarcoplasmic reticulum (SR) exhibits a plexiform morphology when observed with myofibrils (M) but most often appears in a reticular pattern (arrows; Fig. 37) in non-beating cells. m, mitochondria. Fig. 35, X 2,000; Fig. 36, x 3,500; Fig. 37, x 33,100. 296 M.L. DECKER ET AL. idence of their previous orderly register, and, in most instances, reveal a paucity of thick filaments (Fig. 36). Ttubules and the sarcoplasmic reticulum remain evident in non-beating myocytes in spite of the marked reorganization in the contractile apparatus. T-tubules and their adjacent junctional sarcoplasmic reticulum and the honeycomb morphology characteristic of non-junctional sarcoplasmic reticulum can be observed in association with the modified myofibrils (Fig. 37). However, the sarcoplasmic reticulum assumes a reticular pattern in regions where myofibrillar disruption is apparent (Fig. 37). As in contractile cells, numerous polyribosomes are readily apparent in the sarcoplasm (Fig. 37). DISCUSSION Previous results from this laboratory demonstrate that quiescent cultures of adult rabbit cardiac myocytes can be established on laminin-coated surfaces that are suitable for combined biochemical and morphological study (Haddad et al., 1988a,b; Decker et al., 1988a,b). Both beating and non-beating myocyte preparations maintain high CrP/ATP ratios, release negligible amounts of cytoplasmic enzyme activity, and exclude trypan blue, testifying to the consistent quality of such cultures (Table 1).Adult rabbit myocytes retain their rod-shaped morphology and gradually flatten without passing through a n intervening round phase (Haddad et al., 1988a; Decker et al., 1988a,b) and the accompanying disruption of contractile elements that is characteristic of most cultured adult rat myocytes (Moses and Claycomb, 1982b; Nag et al., 1983; Jacobson and Piper, 1986; Piper et al., 1988; Eppenberger et al., 1988); nevertheless, a more subtle form of subcellular remodeling is evident a s adult rabbit myocytes adapt to culture. If freshly isolated myocytes are plated at high density (1x lo6 cellsi35 mm petri dish), then approximately 20% of the cultures commence beating synchronously between day 10 and 12 of culture. Although we can only speculate on the mechanism responsible for the initiation of contraction, only cultures that have re-established cell-cell contact and assembled a n intercalated disc at this juncture (Decker et al., 1989a) beat synchronously. Since approximately 50% of the adult rabbit heart cells detach from the culture vessel during the first week of culture (Haddad et al., 1988a1, it is conceivable that in high-density cultures some surviving pacemaker cells may establish contact with neighboring ventricular myocytes provoking contractile automaticity (Meier et al., 1986; Jacobson et al., 1988b). Conversely, in low-density cultures i t is likely that solitary rabbit ventricular cells normally remain quiescent because they can maintain high transmembrane resting potentials (Powell et al., 1980; Bkaily et al., 1984; Meier et al., 1986). The present investigation describes the subcellular reorganization that accompanies the adaptation of adult rabbit myocytes to in vitro life and clearly demonstrates that myocytes which fail to develop contractile function display “myofibrillar atrophy” and ultimately reconfigure those remaining structural components of the contractile apparatus to assume a pattern vaguely reminiscent of embryonic myofibrils (Fischman, 1967). Those myocytes that commence beating retain many of the morphological features which characterize their in vivo progenitors. The progressive disruption of the contractile apparabus represents the major structural alteration that evolves in the quiescent myocyte during the first 2 weeks of culture. During the first week of culture, fluorescent images of phalloidin- and anti-myosinlabeled myocytes illustrate some loss of myofibrillar order that may be correlated with the development of minor discontinuities within the Z-lines of some sarcomeres. A similar pattern of Z-line dissolution is apparent in the myofibrils of quiescent rat myocytes cultured for similar lengths of time (Jacobson and Piper, 1986; Schwartz et al., 1985); however, nonbeating feline myocytes are reported not to disclose such features (Cooper e t al., 1986). Since the development of such Z-line abnormalities is frequently associated with disuse atrophy in skeletal muscle (Jacobson and Piper, 1986), the present observations imply that non-beating, but adherent rabbit and rat cardiac myocytes display myofibrillar atrophy. Although myocytes isolated from each of these species would appear to be “loaded’ (Cooper et al., 1986) to the same degree (i.e., attached to a laminin- or fetal calf serum-treated substrata), the feline contractile apparatus apparently responds more favorably to this “in vitro load” than do its r a t or rabbit counterparts, disclosing little morphological sign of atrophic myofibrils for nearly 2 weeks (Cooper et al., 1986). The mechanism(s) that might mediate the disruption of myofibrillar organization are only now being investigated, but several experiments clearly indicate that the synthesis of myofibrillar proteins is almost completely suppressed in non-beating rabbit myocytes (Haddad et al., 198813; Decker et al., 1989b). Since the disappearance of intact myofibrils appears to proceed relatively rapidly, the degradation of contractile proteins may not be altered significantly in quiescent myocytes (Decker et al., 1989b). Future experiments designed to measure the rate of actin and myosin degradation will be required to verify this hypothesis. The present observations support the contention that synthesis of contractile and, perhaps, cytoskeletal proteins a s well (Simpson et al., 1988) may be regulated to a large extent by mechanical factors and that the myofibrillar proteins are degraded andlor disassembled in the absence of a “significant” in vitro load (Cooper et al., 1986; Mann et al., 1989) and/or spontaneous contractility (McDermott and Morgan, 1989). The disruption of the contractile apparatus proceeds unabated a s non-beating myocytes continue to spread and flatten a s culture is prolonged. At 2 weeks, fluorescence (Figs. 28, 29) and thin-section images (Figs. 27, 30) reveal a n apparent disintegration of myofibrillar elements with few “intact” myofibrils recognizable in most myocytes. These myofibrils, such as they are, reveal a paucity of thick filaments and aberrant “Zlines” (Fig. 30). At this juncture, myosin assumes primarily a diffuse perinuclear distribution with only a modest amount of co-localization with actin-positive fibrils (Figs. 28, 29). Ultimately, the myofibrillar apparatus becomes reordered into a n interdigitating pattern of “minute-myofibrils” (Fig. 35) that appear to replace the normal, repeating registry of the mature contractile apparatus. This structural reorganization is accompanied by a marked decline in the myofibrillar CONTRACTION AND MYOFIBRILLAR STRUCTURE volume density in these quiescent myocytes (Decker et al., 1989a)-whether such myocytes retain a contractile potential awaits further investigation. The fate of the T-tubule system and the organization of the sarcoplasmic reticulum also seems linked with changes in the organization of the contractile apparatus in non-beating myocytes. Through the first week of culture, little change could be documented in either the structure or the distribution of T-tubules or the sarcoplasmic reticulum. The former retained its close association with Z-lines and junctional sarcoplasmic reticulum and the latter encircled myofibrils, displaying its classical plexiform morphology (Fawcett and McNutt, 1969; Forbes and Speralakis, 1977). However, as these quiescent myocytes gradually spread and lose their myofibrils, the T-tubules are less frequently encountered, and the spatial relationships that T-tubules and the sarcoplasmic reticulum normally enjoy with myofibrils are lost in areas where myofibrillar disruption is apparent. T-tubules are no longer closely associated with Z-lines in many instances (Moses and Claycomb, 1982a) and the plexiform morphology that typifies the non-junctional sarcoplasmic reticulum is transformed into a tubulo-vesicular configuration somewhat reminiscent of smooth endoplasmic reticulum seen in steroid secreting cells (Fig. 32). One structural association remains constant throughout this period of myofibrillar atrophy, and that is the continued presence of junctional sarcoplasmic reticulum closely apposed to T-tubules (Fig. 32). Such results imply that in the absence of mechanical activity, the structural relationships between the myofibrillar apparatus and T-tubules and the sarcoplasmic reticulum are lost. Whether these “atrophied” myocytes retain the potential to contract if stimulated appropriately will be the focus of future experiments. It should be emphasized, however, that factors other than contractility may modify the organization of the T-tubule system and the sarcoplasmic reticulum. For example, the extensive cell spreading that transpires in vitro (Claycomb and Palazzo, 1980; Haddad et al., 1988a; Jacobson et al., 1988a,b; Piper et al., 1988) provokes a dramatic change in the surface to volume ratio of these cells. Since the depolarization of the sarcolemma and its T-tubule extensions promotes the synchronous release of calcium ions from the sarcoplasmic reticulum and other sources, the increase in surfacelvolume ratio that accompanies myoc t e spreading may effectively reduce the distance Ca’+ ions must diffuse to elicit excitation-contraction coupling in cultured heart cells (Langer et al., 1979, 1987), thereby obviating the need for a n extensively developed T-system and wellordered sarcoplasmic reticulum. Delcarpio and associates (Delcarpio et al., 1986) also report that extensively spread adult rat myocytes contain fewer T-tubules than freshly isolated cells (Decker et al., 1989a). Nevertheless, when rabbit myocytes are cultured a t high density, some of the preparations commence beating synchronously between day 10 and 12 of culture. Such contractile activity promotes the retention of myofibrils in myocytes examined at 14 and 28 days of culture; moreover, no indication of Z-line fragmentation can be demonstrated in these contracting myocytes (Fig. 23). The morphological relationships between Ttubules, the sarcoplasmic reticulum, and myofibrils 297 also appear to be stabilized by beating. T-tubules are frequently encountered adjacent to myofibrillar Z-lines or a s branching baso-lateral invaginations of the sarcolemmal surface similar to those reported by other investigators (Moses and Claycomb, 1982a, 1985; Langer et al., 1987). The sarcoplasmic reticulum retains its honeycomb configuration with little evidence of reorganization similar to t h a t observed in the nonbeating myocytes. Clearly, contractile activity represents the principal trophic influence that maintains the organization of the myofibrillar apparatus and its associated membranous components in cultured adult rabbit myocytes. The results of the present investigation demonstrate that a unique subcellular remodeling of adult cardiac myocytes attends long-term culture and that in the absence of contractile function, myocytes display morphological signs of marked myofibrillar atrophy. Several investigations support the argument that the mechanical load placed upon heart cells may well be of primary importance in the maintenance of the structural and functional properties of cardiac myocytes. For example, transection of the chordae tendinae of feline right ventricular papillary muscle induces rapid and progressive atrophy of the cardiac myocytes (Cooper and Tomanek, 1982). This atrophic response is characterized by a significant reduction in myofibrillar volume density, a concomitant decline in both actin and myosin content, and a depression in the force generation of the unloaded papillary muscle (Thompson et al., 1984; Kent et al., 1985). A novel feature of this model is that the unloaded papillary muscle continues to contract in synchrony with the right ventricle (Cooper and Tomanek, 1982), implying that changes in load and not beating per se, control the composition and organization of the contractile apparatus. When the papillary muscle is reloaded, it quickly regains its normal structure and function (Thompson et al., 1984). The morphological dissolution of papillary myofibrils (Tomanek and Cooper, 1981) associated with the atrophy in this model closely resembles the events described in the present study where the focal loss of Z-line material and disruption of myofibrillar organization is accompanied by a major increase in the myocytic cytoplasmic ground substance. Recent observations further illustrate that cultured beating neonatal myocytes grow larger when attached to a substratum than corresponding unattached beating cells (Marino et al., 1987). Moreover, attached contractile myocytes apparently retain their myofibrillar organization, whereas unattached cells display a poorly ordered contractile apparatus. Such results further implicate “load” as the principal factor that regulates the composition and structure of the contractile apparatus in the isolated heart cell. In this regard, Cooper’s recent results also bear importantly on the issue of “load” (Cooper et al., 1986). Non-beating feline adult myocytes attached to laminin and cultured in serum-free conditions reveal only minor reductions in contractile protein content and minimal myofibrillar disruption when maintained in vitro for 2 weeks. Cooper et al. (1986) suggests that attachment, itself, induces a load significant enough to inhibit atrophy. The results in the present study, however, demonstrate that rabbit myocytes attached to a laminin substratum clearly exhibit atrophy. Further- M.L. DECKER ET AL. 298 more, the present observations suggest that attachment is not sufficient, in and of itself, to prevent the atrophy and reorganization of the contractile apparatus in cultured rabbit myocytes. Two caveats must, however, be mentioned in light of the present observations. Cooper’s study is conducted in the absence of serum, and the present investigation employs this supplement, which is recognized to enhance cell spreading, but also ensures long-term survival of cultured cardiac myocytes (Borg and Terracio, 1988; Jacobson et al., 1988a; Piper et al., 1988). Nevertheless, under the present culture conditions, significant evidence of myofibrillar atrophy [i.e., Z-line disruption and loss of contractile units (Decker et al., 1989a)l precedes significant cell spreading; therefore, disruption of the contractile apparatus does not appear to be closely linked with cell spreading per se. Secondly, Cooper’s group (Cooper et al., 1986) employed significantly higher concentrations of laminin for cell attachment than those used in the present study (Haddad et al., 1988a); whether these differences influence myocyte spreading or the composition of the contractile apparatus awaits further study. The results of these investigations suggest that attachment, alone, is insufficient to maintain a “normal” myofibrillar apparatus and that a n isometric load created in the beating myocyte appears to be a primary stimulus for supporting a fully developed contractile apparatus. ACKNOWLEDGMENTS The present study was supported by public health service grants HL 33616 and HL 19648 and the Feinberg Cardiovascular Research Institute of Northwestern University Medical School. The authors wish to thank Dr. William A. Clark for providing our laboratory a sample of the CM-52 monoclonal anti-myosin antibody. LITERATURE CITED Bkaily, G., N. Sperelakis, and J. 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