Immunofluorescent studies for myosin, actin, tropomyosin and α-actinin in cultured cardiomyopathic hamster heart cells

DEVELOPMENTAL

BIOLOGY

97,

338-348

(1983)

Immunofluorescent Studies for Myosin, Actin, Tropomyosin and a-Actinin in Cultured Cardiomyopathic Hamster Heart Cells LARRY F. Department Received

of Anatomy, September

LEMANSKI’

AND

University

ZENG-HONG

of Wisconsin,

24, 1982; accepted

in revised

Madison,

form

Tu Wisconsin January

53706

10, 1983

Primary cultures of cardiac myocytes from newborn normal and genetically cardiomyopathic (strain UM-X7.1) hamsters were analyzed by electron microscopy and immunofluorescent staining for myosin, actin, tropomyosin, and cY-actinin. Antibody staining of these contractile proteins demonstrates that both normal and cardiomyopathic (CM) myocytes contain prominent myoflbrils after 3 days in culture, although the CM myofibrils are disarrayed and not aligned as those in normal cells. The disarray becomes even more pronounced in CM cells after 5 days in culture. The immunofluorescent staining patterns of individual myofibrils h normal and CM cells were similar for myosin, actin, and tropomyosin. However, cY-actinin staining reveals that the CM myofibrils have abnormally wide and irregularly shaped Z bands. Electron microscopy confirms the irregular Z-band appearance as well as the myofibril disarray. Thus, CM cardiomyocytes clearly show an aberrant pattern of myofibril structure and organization in culture.

INTRODUCTION

while these similarities are well documented, the etiologies are poorly understood. Additional characteristics of cardiomyopathic hamster hearts include abnormalities in synthesis of actin and myosin (Bester and Wieland, 1974), in electrocardiology (Laird, 1974), in sarcoplasmic reticulum adenosine triphosphatase and calcium binding (Dhalla et al., 1975; Wrogemann et al, 1974; Louis and Irving, 1974; Singh et ah, 1975), in oxidative phosphorylation and calcium transport in mitochondria (Wrogemann et al., 1975; Thakar et al., 1973; Kako and Heggtveit, 1975), in various electrolytes (Lossnitzer and Bajusz, 1974), in water content (Kidd et al, 1981), and in L-carnitine levels (Paulson et al, 1981). Jasmin and Bajusz (1975) showed that administration of dibenamine, an adrenergic blocker that improves microcirculation, lowers the heart lesion in juvenile and adult animals. Thus, it appears that at least some of the above-mentioned abnormalities are secondary effects of hypoxic conditions resulting from decreased blood flow to the heart muscle in advanced juvenile and adult animals. To date, very little investigative attention has been directed toward analyzing the heart myocytes of young cardiomyopathic hamsters. The major problem with studies on advanced stages of the cardiomyopathy is that many secondary effects probably result from the heart failure. In order to understand the primary defect(s) of the cardiomyopathy it seems essential to study young animals before the experimental results become “clouded” by the cascading effects of secondary abnormalities. In addition, it would obviously be desirable to examine normal and cardiomyopathic (CM) cardiomyo-

Hornburger et al. (1962) first reported the discovery of an autosomal recessive condition in an inbred strain (BIO 14.6) of Syrian hamster which resulted in a dystrophy-like disease of the skeletal muscle, heart failure, and premature death of the animal. Several additional strains have since been derived from the original BIO 14.6 hamsters and one, UM-X7.1, has the advantage of showing good consistency in the temporal onset of disease symptoms (Bajusz, 1969,1973). This dystrophy affects the heart more severely than skeletal muscle and homozygous recessive animals die of heart failure after only one-third to one-half the normal life span. In recent years, numerous publications have appeared on cardiomyopathic hamsters. Most of these papers are descriptive morphological, physiological, or biochemical studies dealing with late juvenile or adult animals (Bajusz and Rona, 1972). Such studies are important in that they have indicated a close similarity between the pathology of the hamster cardiomyopathy and that of human heart disease. For example, the disorientation of myofibrils is a rather classical trait of hypertrophic obstructive cardiomyopathy in humans (Heggtveit and Nadkarni, 1971; Ferrans et aZ., 1972), yet virtually nothing is known about the cell level mechanisms by which directional derangement of the myofibrils takes place; it is possible that the mechanism(s) may turn out to be very similar to that causing the derangement of myofibrils in the cardiomyopathic hamster hearts. Thus, i To whom all correspondence should be addressed: Department Anatomy, University of Wisconsin, Madison, Wise. 53706.

0012-1606/83 Copyright All rights

$3.00

0 1983 by Academic Press. Inc. of reproduction in any form reserved.

of

338

LEMANSKI

AND Tu

Card&myopathic

cytes under identical growth conditions (i.e., using cell culture methods). In earlier reports (Bajusz, 1969) aberrant myofibril structure and organization were observed in in vivo heart cells of cardiomyopathic animals 30 days of age or older; it has been suggested that this abnormality may result from hypoxia due to heart failure in vivo (Jasmin and Bajusz, 1975). In the present study, to assess whether the abnormal myofibril organization is a primary defect or a secondary effect of in vivo development, we prepared primary cultures of ventricular heart cells from newborn normal and CM hamsters. Cardiomyocyte myofibril organization was analyzed by ultrastructural and immunofluorescent methods after 3 and 5 days in culture. The studies clearly demonstrate that an aberrant pattern of myofibril organization and Z-band formation is present in cultured cells derived from CM hamsters. Normal cells cultured under identical conditions do not show these abnormalities. Thus, our results strongly suggest that the aberrant myofibrils in CM heart cells result not from secondary effects of in vivo development, but, more likely, from primary cell-level effects of the genetic mutation. MATERIALS

AND

METHODS

Animals. Normal Syrian hamsters were obtained from the Research Animals Resources Center at the University of Wisconsin, Madison, and were bred randomly. Genetically cardiomyopathic Syrian hamsters were obtained from our breeding colony, UM-X7.1/UW subline, derived from breeding stock provided by Dr. Jasmin of the University of Montreal, and inbred according to his protocols. The normal and CM animals were housed under identical conditions in the same room at 23°C on a light cycle of 12 hr light-12 hr dark. They were fed Purina Lab Chow and water ad libitum with supplements of a hamster seed mixture and lettuce. Tissue culture. Hearts of 3-day-old normal and CM hamsters were used for culture. The animals were killed by cervical dislocation and the hearts immediately removed using sterile techniques. The extirpated hearts were washed several times in cold Hanks’ solution to remove residual blood and the atria were removed and discarded. The ventricles were minced into small pieces (1 mm3) and washed again in Hanks’ solution. The pieces were incubated with agitation in Hanks’ solution containing 0.08% trypsin (1:250, Difco) and 0.02% collagenase (132 U/mg, GIBCO) at 35°C for 10 min. This was repeated four times. The first supernatant was discarded, then the three additional supernatants were diluted twofold with cold Eagle’s MEM, collected by lowspeed centrifugation, and washed two additional times in 100% Eagle’s MEM. The resulting pellets were re-

Heart Cells

339

suspended in medium plus glutamine and, to enrich the cultures for myocytes, the dissociated heart cell suspensions were preincubated in an Erhlenmeyer flask for 60 min at 37°C. Most of the fibroblasts attached to the bottom of the flask during this period. The remaining unattached cells (now -90% myocytes) were diluted to an average density of 2 X lo5 dispersed cells/ml of medium. Two-milliliter aliquots of the myocyte-enriched cell suspensions were added to each 30-mm Corning tissue culture dish with a rat tail collagen-coated glass coverslip on the bottom. The culture medium consisted of Eagle’s MEM containing 15% fetal calf serum, penicillin (100 U/ml), streptomycin (0.1 mg/ml), and fungizone (0.25 pg/ml). The cultures were incubated at 3’7°C in an atmosphere of 5% CO%and 95% air. The culture medium was changed every second day. For the present study, 3- and 5-day cultures were examined. Antigen and antibody preparation. Antibodies raised against highly purified muscle protein antigens were used for the immunofluorescent studies. Myosin heavy chain, actin, and tropomyosin from chicken heart muscle were purified by preparative-slab SDS-polyacrylamide gel electrophoresis (Lazarides, 1975). n-Actinin prepared from porcine skeletal muscle (Go11 et al, 1972) was a generous gift from Dr. Darrel Go11 and Dr. Judith Schollmeyer. Antibodies against these proteins were prepared in young rabbits by methods detailed in earlier papers (Lemanski, 1979; Lemanski et aZ., 1980). Briefly, preimmune rabbits were screened carefully to eliminate nonspecific staining on the tissue samples to be used in the experiments. Antibodies to cy-actinin were obtained by giving an initial subcutaneous injection of 2 mg protein in Freund’s complete adjuvant followed by three subcutaneous booster injections of 2 mg of protein in Freund’s incomplete adjuvant at 2-week intervals. Antibodies to myosin heavy chain, actin, and tropomyosin (i.e., those purified by SDS-polyacrylamide gel electrophoresis) were prepared as above except that the relevant bands were cut from gels after slight Coomassie blue staining, homogenized in physiological saline, and injected into the rabbits without Freund’s adjuvant. The rabbits were bled from the lateral ear veins 8 days after the final injection and the -y-globulins obtained by ammonium sulfate fractionation. The antibodies were aliquoted and stored at -70°C until ready for use. Purity of antibody preparations was tested by double immunodiffusion and immunoelectrophoresis against purified,antigen and against crude heart muscle homogenates from chicken and hamster. Antibodies against myosin heavy chain, cY-actinin, and tropomyosin each gave single sharp precipitation lines indicative of high specificity (Lemanski et al, 1980). Our anti-a&in antibodies were nonprecipitating with purified actin or homogenates. The specificity of all anti-

340

BEVELOPMENTAL

BIOLOGY

bodies was tested further by staining isolated unfixed myoAbrila freshly prepared by methods detailed in our earlier paper (Lemanski, 1979); antimyosin specifically stained the A bands, anti-a-actinin the Z bands, and anti-a&in and anti-tropomyosin the I-band regions (Fig. I). Immunqfluoreacent stain&g. After 3 or 5 days in eulture, the coverslips on which heart cells grew were rinsed briefly in three changes of phosphate-buffered saline (PBS) and fixed in a periodate-lysine-paraformaldehyde solution (McLean and Nakane, 1974) for 30 min at room temperature. The coverslips were washed in two changes of PBS for 15 min, placed in 7% sucrose-PBS for 15 min, and transferred to 25% glycerol-PBS for 30 min. An indirect immunofluorescent method was used to stain the cultured cells. The primary antibodies in 25% glycerol-PBS were applied to the fixed cells for 60 min at 37OC.After several rinses in PBS-glycerol totaling 60 min, the cells were stained with FITC-labeled goat anti-rabbit IgG (Miles Laboratories) diluted 1:40 with 25% glycerol: PBS for 60 min at 37*C. The controls for the antibody staining included (a) incubation with the preimmune globulins from the same rabbit that

VOLUME

97,

1983

produced the apecific antibodies, (b) staining with an* tibodies absorbed with excess purified antigen, and (c) staining with second antibody only. There was very little background staining on these controls. The FITCstained cells were viewed with a Zeiss Universal light microscope equipped with epifluoreseent illumination using a xenon light source. Photographs were taken on 35”mm Kodak Plus-X film (ASA 125) at SO-setexposure times. Ekctrm microscow. For routine morphology, the cells were fixed by immersing the coverslips in a. glutaraldehyde-formaldehyde picric acid mixture, buffered to pH ‘7.4with 0,15 N phosphate buffer (Ito and Karnovsky, 1968) for l-2 hr at 23’C. After a brief buffer rinse,~ the cells were postfixed for 60 min at 0°C in 1.0% osmium tetroxide buffered to pH 7.4 with 0.15 M phosphate. The cells were dehydrated in graded ethanols and embedded by inverting Epon-filled Beem capsules over the areas of interest followed by polymerization at 60°C for 48 hr. The blocks were removed from the glass by alternating immersions in hot water and liquid nitrogen, thin sectioned with a diamond knife, mounted on bare copper grids, and doubly stained with lead citrate

FIG. 1. Corresponding phase-contrast and immunofluorescsnt micrographs of unfixed hamster skeletal muscle myoflbrile prepared by homogenization of glycerol extracted muscle in standard salt solution (0.01 M KCl, 0.006 M MgCl,, 0.006 M EGTA, 0.006 M phosphate buffer, pH 7.0). An indirect staining method was used; apacific primary antibodies prepared in rabbits were applied to myofibrils dried on glass slides followed by FITC-labeled goat anti-rabbit IgG. Arraws indicate corresponding areas on the paired micrographs. (a) Antimyosin specifically stains the A bands, X8100; (b) anti-actin the I-band regions, X2620; (c) anti-tropomyosin the I-band regions, X2620; (d) anti-aactinin the Z banda, X2620.

LEF~ANSKI AND Tu

Car&myopathic

Heart

Cells

FIG. 2. Typical normal oardiomyocyte cultured 3 days and stained for myosin by an indirect immunofluorescent method. The A bands of the myofibrils are stained specifically and reveal that most of the myofibrils are arranged in parallel arrays, X1340. FIGS. 3-5. Cardiomyopathic (CM) hamster heart myocytes cultured 3 days and stained for myosin. The A-band staining of individual myofibrils is similar to that in normal cells; however, in CM cells, the myofibrils are variably disoriented with respect to each other, X1340.

and uranyl acetate. Sections were viewed on Hitachi HU-11DS or Jeol 106CX electron microscopes at 75 or 60 kV. RESULTS

Three- and five-day primary cultures derived from the heart ventricles of both normal and CM newborn hamsters contain a mixture of cardiac myocytes and nonmuscle cells (mostly fibroblasts). After 3 days in culture, approximately 75% of the cells were myocytes, while after 5 days 50% were myocytes. This presumably resulted from the higher mitotic index of fibroblasts, since most of the nonmusele cells had been removed by our preincubation step as described under the Materials

and Methods section. In spite of this heterogeneity, the muscle cells could be distinguished easily from the nonmuscle cells in the cultures. The myocytes are irregular in shape and display phase-dense cytoplasm with numerous cross-striated myofibrils. They beat spontaneously at 59-120 contractions per minute. The fibroblasts appear similar in shape to the cardiomyocytes, but have a phase-lucent cytoplasm. Furthermore, they do not contain cross-banded myofibrils nor do they contract. When cultured cardiomyocytes are stained with FITClabeled antibodies against the contractile proteins, intense fluorescence is apparent in the myofibrils of both normal and CM cells, Antimyosin antibodies specifically stain the A bands of organized myofibrils in the cul-

342

DEVELOPMENTALBIOLOGY

VOLUME

97,1983

FIG. 6. Typical normal cardiomyocyte cultured for 5 days and stained for myosin by an indirect immunofluorescent have become more numerous in the cells and continue to show parallel arrangements with sarcomeres in register, FIGS. 7-9. Cardiomyopathic (CM) hamster heart myocytes cultured 5 days and stained for myosin. The staining appears similar to that seen in normal control heart cells. Myofibril disarray is obvious in the CM cardiomyocytes X1250; X680; X1100.

tured cells. Normal cardiomyocytes after 3 days in culture contain well-formed myofibrils that are, for the most part, oriented parallel to each other and to the longitudinal axes of the cells (Fig. 2). By contrast, antimyosin staining shows that the myofibrils in 3-daycultured CM cells are disoriented with respect to each other and fail to form the parallel arrays as in normal cells (Figs. 3-5). After 5 days in culture, the normal cells have increased in size and contain larger and more numerous myofibrils which continue to be oriented in parallel arrays (Fig. 6). The myofibril disarray in CM cells after 5 days has become even more pronounced and many myofibrils appear to form complex entanglements within the sarcoplasm (Figs. 7-9). In spite of the

method. The myofibrils X1100. of individual myofibrils after 5 days in culture,

disoriented arrangements, the antimyosin staining for individual myofibrils appears similar in normal and CM myocytes. Anti-actin and anti-tropomyosin antibodies specifically stain the I bands of the myofibrils as well as the lateral portions of the A bands, presumably where the thin filaments interdigitate with the thick filaments (Figs. 10-13). As with antimyosin-stained preparations, myofibril disarray in 3- and 5-day cultures of CM cardiomyocytes is readily apparent. Also, no obvious differences between normal and CM cells are noted in the staining of individual myofibrils. Immunofluorescent staining for ff-actinin in normal and CM cardiomyocytes again reveals myofibril disar-

LEMANSKI

AND

Tu

Cardiomyopathic

FIG. 10. Antiactin-stained CM cardiomyocytes after 3 days in culture. The actin similar to that for normal ones, ~666. FIG. 11. Immunofluorescent staining pattern for tropomyosin in CM cardiomyocyte of myofibrils is evident in CM myocytes, the I-band staining of individual myofibrils FIGS. 12, 13. Antitropomyosin stained CM cardiomyocytes after 5 days in culture. become even more extensive than at 3 days in culture, X784.

ray in 3- and 5-day CM cultures (Figs. 14-17). The 5day CM cultures further show an abnormal staining pattern for cY-actinin in individual myofibrils. The Zband staining appears wider and somewhat more diffuse than normal and, in many areas, the Z bands have an irregular “zig-zag” appearance. On occasion, a-actinin staining appears to extend across an entire sarcomere length from one Z band to another. In addition, some of the CM cardiomyocytes contain collections of cu-actinin-positive “spots” suggestive of Z bodies, usually in their peripheral cytoplasm. The normal cells show distinct anti-a-actinin staining of the myofibrillar Zbands which in most cells are registered by 5 days in

Heart

Cells

staining for individual

myofibrils in CM cells appears

after 3 days in culture. Although aberrant organization is the same as in normal cells, X1078. Disarray of myofibrils is obvious in CM cells and has

culture. The staining in normal cells is suggestive of thin distinct Z bands; Z body-like distributions are rarely seen. Electron microscopic observations of cultured normal and CM cardiomyocytes corroborate the immunofluorescent studies. Normal heart cells contain well-organized myofibrils that are in parallel arrangements with their distinct Z bands, usually in register (Fig. 18). The CM cardiomyocytes show aberrant disarrayed myofibrils which often criss-cross each other and frequently a single Z band may have more than one myofibril insertion (Fig. 19). The Z bands appear wider and more diffuse than normal in some areas. In addition, Z bodies

DEVELOPMENTAL BIOLOGY

344

VOLUME 97, 1983

FIG. 14. Anti-o-actinin staining pattern of normal heart cell after 5 days in culture reveals that the specifically stained Z bands are thin and many are registered, X1100. FIGS. 15-17. AntibcY-actinin staining of CM heart cells cultured for 6 days. The disoriented myofibrils show irregular zig-zag Z lines in some areas. Also, what appear to be 2 bodies and Z plaques are evident in the CM cells, X800.

and Z plaques with associated thin filaments have accumulated in the peripheral cytoplasm of many CM cells (Fig. 20). DISCUSSION

The present immunofluorescent and ultrastructural studies demonstrate that CM cardiomyocytes have an abnormal pattern of myofibril organization when grown in culture. The myofibrils in CM cardiomyocytes are disoriented with respect to each other and fail to align in parallel arrays as in normal control cells. Wada et a&Z.(1976) reported that disorganized myofibrils are present in portions of cultured CM heart cells (Strain 310 14.6) as revealed by electron microscopy of thin

sections. In the present study, our use of immunofluorescent staining of the contractile proteins allowed us to evaluate the distributions of myofibrils in whole cells and thus unequivocally confirm an extensively disarrayed myofibril arrangement in CM cardiomyocytes. The immunofluorescent staining patterns for myosin, actin, and tropomyoein in individual myofibrils of CM hamster cells were similar to those seen in normal cells; however, anti-a-actinin staining was abnormal in CM cardiomyocytee. The Z lines appeared wider, more diffuse, and irregular. Also, more 2 bodies were present in CM cells, These observations were confirmed by the ultrastructural studies. Such results could be interpreted to indicate that the CM cells are delayed in their differentiation when compared to normal, a possibility

LEMANSKI AND Tu

Cardimyoputhic

Hearl

cd.3

346

after 5 daysin culture. The myefibrils(arrows)showa parallelalignmentrelative FICI. 18.Electronmicrographof normalcardiomyocpte to eachother andto the longitudinalaxis of the cell, Nu, nucleus,X8466.

Wada et ul. (19%) suggested for cultured CM heart cells in I310 14.6 hamsters. However, that the disarray in CM cells becomes more pronounced as time in culture progresses would seem to argue against such a conclusion. Furthermore, the cultured CM cardiomyocytes formed myoflbrils and began contracting at the same time as normal cells. In addition, the Z-bodies and 2 body-thin filament complexes increased with longer culture times indicating that this does not represent a temporal delay in differentiation. That our results differ from Wada and co-workers could be due to strain differences of the cardiomyopathy (Wada et al. used 310 14.6 and we used UM-X’I.l/UW), We believe that a more likely alternative, however, is that in the preliminary report by Wada et al. only electron microscopy was used to accessmyofibril formation and there obviously would have been some diflkulty in observing on thin sections the overall

pattern of myofibrils that were so extensively disarrayed. Our own use of immunofluorescent methods overcame this limitation by allowing us to view the entire complement of myofibrils in a given cell. Thus, results of the present study would not support the hypothesis that myofibril formation itself is temporally delayed in cultured CM cardiomyocytes, but suggests rather that myoflbril formation in CM cells temporally parallels that in normal ones. Recent studies on newborn in v&o hearts of normal and CM hamsters (strain-UM-X7.1) using morphometric analyses at the ultrastructural level would support this conclusion (Kidd et aL, 1981). The myofibrils in cultured CM cells have failed to align properly, however. Why there are myoflbril abnormalities in CM cells remains a moot question. Preliminary SDS-polyacryl-

346

DEVELOPMENTALBIOLOGY

VOLUME 97,1983

LEMANSKI

AND Tu

Cardiemyopathic

amide gel electrophoresis experiments on in vivo embryonic hearts have been suggestive of protein differences between normal and CM hearts. One of the proteins which appeared to be reduced (not detectable on Coomassie blue-stained gels) in mutant hearts when compared to normal ones has a molecular weight of -50,000 daltons (interestingly within the molecular weight range for desmin and/or vimentin) (Nakanishi and Lemanski, unpublished observation). Differences in a-actinin (100,000 daltons) did not appear to be significant by electrophoretic techniques in spite of the abnormal immunofluorescent staining pattern for this protein in CM cells. Whether there might be some relationship between the aberrant myofibril orientation, the abnormal a-actinin staining pattern, and the abnormal electrophoresis pattern in CM hearts will require further study. The first published work suggesting that filaments of “intermediate size” are responsible for aligning muscle myofibrils into register appeared in our earlier papers describing the ultrastructure of developing axolotl (salamander) heart cells (Lemanski, 1972, 1973). In more recent studies, Gard and Lazarides (1980), using immunofluorescent methods, have suggested that intermediate filaments composed of desmin and vimentin are involved in the alignment of myofibrils in cultured chicken skeletal myotubes as well. Thus, if the electrophoretically “missing” band (-50,000) on gels of embryonic cardiomyopathic hamster heart turns out to be desmin or a desmin-like protein, then the implications as to why myofibrils are improperly aligned in CM cells could prove most interesting! Whatever future experiments yield along these lines, the present study makes it clear that myofibril formation is aberrant in CM hamster heart cells in culture and suggests that this condition in in vivo hearts is not caused solely by hypoxic conditions resulting from decreased coronary microcirculation as has been suggested in previous reports. We believe that a more likely possibility might be that the observed disarray is a primary defect of the mutation (or at least a defect that is expressed very early in development). This presumably would result in a decreased contraction efficiency for the CM heart cells which, in turn, could lead to hypertrophy, heart failure, and eventually to many other reported abnormalities at advanced stages of the disease. It is clear that further critical studies will be required to fully understand the genetically based cardiomyopathy in UM-X7.1/UW Syrian golden hamsters. FIG. other. FIG. and Z

19. Electron micrograph The Z bands (Z) appear 20. Electron micrograph plaques (P) are evident

Heart

347

Cells

The‘authors gratefully acknowledge Ms. Ghousia Nizamuddin for assistance in preparing the illustrations and Ms. Sue Leonard for secretarial assistance. This work was supported by NIH Grant I-IL22550 and, in part, by a Grant-in-Aid from the American Heart Association. The work was done during the tenure of an American Heart Association Established Investigatorship Award to L.F.L. Zeng-Hong Tu is a Visiting Scholar at the University of Wisconsin from the Shanghai Institute of Materia Medica, Chinese Academy of Sciences, People’s Republic of China.

REFERENCES

BAJUSZ, E. (1969). Hereditary

cardiomyopathy: A new disease model. Heart J. ‘77, 686-696. BAJUSZ, E. (1973). A disease model of hereditary cardiomyopathy: Its usefulness and limitations. Rec. Advan. Studies Card Struet. MetaboL 2, 291-292. BAJUSZ, E., and RONA, G. (1972). Rec. Advan Studies Card Stmcct. MetaboL 1,225-324. BESTER, A., and WIELAND, G. (1974) The synthesis of myofibrillar and soluble proteins in cell-free systems and in intact cultured muscle cells from newborn polymyopathic hamsters. J. Mol. Cell. CardioZ. 7, 325-344. DHALLA, N., SINGH, A., LEE, S., ANAND, M., BERNATSKY, A., and JASMIN, G. (1975). Defective membrane systems in dystrophic skeletal muscle of the UM-X7.1 strain of genetically myopathic hamster. Clin. Sci. Mol. Med. 49, 359-368. FERRANS, V., MORROW, A., and ROBERTS, W. (1972). Myocardial ultrastructure in idiopathic hypertrophic subaortic stenosis. C&ulation 45, 769-792. GARD, D. L., and LAZARIDES, E. (1980). The synthesis and distribution of desmin and vimentin during myogenesis in vitro. Cell 19. 263275. GOLL, D. E., SUZUKI, A., CAMBELL, J., and HOLMES, G. R. (1972). Studies on purified a-actinin. J. Mol. BioL 67, 469-488. HEGGTVEIT, H., and NADKARNI, B. (1971). Ultrastructural pathology of the myocardium. Methods Achiev. Exp. PathoL 5,474-518. HOMBURGER, F., BAKER, J., NIXON, C., and WILGRAM, G. (1962). New hereditary disease of Syrian hamster. Primary, generalized polymyopathy and cardiac necrosis. Arch Intern Med 110,660-662. ITO, S., and KARNOVSKY, M. (1968). Formaldehyde-glutaraldehyde fixative containing trinitro compounds. J. Cell BioL 39, (2, Pt. Z), 168a (Abstr.). JASMIN, G., and BAJUSZ, E. (1975). Prevention of myocardial degeneration in hamsters with hereditary cardiomyopathy. Rec. Advan Studies Card Struct. MetabuL 6, 219-229. KAKO, K., and HEGGTVEIT, H. (1975). Metabolic changes in the myocardium of hamsters with hereditary muscular dystrophy. Rec. Advan Studies Card. Struct. MetaboL 6, 269-274. KIDD, P., JONES, A., LEMANSKI, L., RUDOLPH, A., and ALLEN, L. (1981). Histological and electron microscope-stereological study of the myocardium of newborn genetically cardiomyopathic hamsters. J. Ultrastruct. Res. 76, 107-119. LAIRD, C. (1974). Developmental Electrocardiography in Inbred Perinatal Hamsters. Advan. CardioL 13,250-269. LAZARIDES, E. (1975). Tropomyosin antibody: The specific localization of tropomyosin in nonmuscle cells. J. Cell BioL 65, 549-561. Amer.

of a portion of CM cardiomyocyte after 5 days in culture. The myofibrils are disoriented with respect to each irregular in shape. Mi, mitochondria; MW, membrane whorl, X2300. of a portion of CM cardiomyocyte after 5 days in culture. In this peripheral portion of the cell, Z bodies (Z) and probably account for the a-actinin positive “spots” observed in immunofluorescent preparations, X44,340.

348

DEVELOPMENTAL BIOLOGY

LEMANSKI, L. F. (1972). Z-Bands in developing heart muscle of the Mexican salamander, Ambystoma mexicanum. Proc. Electron Mieroscop. Soc Amer. 30,24-25. LEMANSKI, L. F. (1973). Heart development in the Mexican salamander, Ambystwna mexicanum II. Ultrastructure. Amer. J. Anat 136, 487-526.

LEMANSKI, L. F. (1979). Role of tropomyosin in actin filament formation in embryonic salamander heart cells. J. Cell Biol. 82, 227238.

LEMANSKI, L. F., FULDNER, R. A., and PAULSON, D. J. (1980). Immunofluorescence studies for myosin, cY-actinin and tropomyosin in developing hearts of normal and cardiac lethal mutant axolotls, Ambystoma mexicanum. J. EmbryoL Exp. Morphol. 55,1-15. LOSSNITZER, K., and BAJUSZ, E. (1974). Water and electrolyte alterations during the life course of the BIO 14.6 Syrian golden hamster. A disease model of a hereditary cardiomyopathy. J. MoL Cell Cardial. 6, 163-177. LOUIS, C., and IRVING, I. (1974). Protein components of sarcoplasmic reticulum membranes from different animal species. Rio&m. Bi@ phya A& 365,193-202. MCLEAN, I. W., and NAKANE, P. K. (1974). Periodate-lysine-paraform-

VOLUME 97, 1983

aldehyde fixative. A new fixative for immunoelectron microscopy. J. Histochem. Cytochem. 22,1077-1083. PAULSON, D. J., TRIPP, M. E., LEMANSKI, L. F., and SHUG, A. L. (1981). L-Carnitine deficiency in the Syrian hamster and rat. Fed Proc. 40, 1589. SINGH, N., DHALLA, N., MCNAMARA, D., BAJUSZ, E., and JASMIN, G. (1975). Membrane alternation in failing hearts of cardiomyopathic hamsters. Rec. Advan Studies Card. Struct. MetaboL 6, 259-268. THAKAR, J., WROGEMANN, K., and BLANCHAER, M. (1973). Effect of ruthenium red on oxidative phosphorylation and the calcium and magnesium content of skeletal muscle mitochondria of normal and BIO 14.6 dystrophic hamsters. Rio&m. Biophys. Acta 314, 8-14. WADA, A., YONEDA, H., SHIBATA, H., INVI, Y., FUSHIMA, H., TAKEMURA, K., and ONISHI, S. (1976). Tissue-cultured heart cells from the cardiomyopathic hamster. J. MoL Cell Cardiol 8, 619-626. WROGEMANN, K., BLANCHAER, M., THAKAR, J., and MEZON, B. (1975). On the role of mitochondria in the hereditary cardiomyopathy of the Syrian hamster. Rec. Advan Studies Card, Struct. MetaboL 6, 231-241.

WROGEMANN, K., JACOBSON,B., and BLANCHAER, M. (1974). Sarcolemma1 ATPase activities in normal and BIO 14.6 dystrophic hamster skeletal muscle. Canad J. Rio&em. 52(I), 500-506.