Myosin Heavy Chain Expression in Respiratory Muscles of the Rat William A. LaFramboise, Jon F. Watchko, Beverly S. Brozanski, Monica J. Daood, and Robert D. Guthrie Department of Pediatrics, Magee-Womens Hospital, University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania
Myosin heavy chain (MHC) isoforms of hind limb adult rat muscles and muscles with a range of respiratory activities were analyzed by a sodium dodecyl sulfate polyacrylamide gel electrophoresis technique that allowed electrophoretic separation of the three fast and one slow MHC isoform found in typical rat muscle. Costal and crural diaphragm muscle samples expressed a mixture of MHC~/S1OW' MHC 2A , and MHC 2X but little MHC 2B • In contrast, MHC 2B was the dominant MHC isoform in the genioglossus, intercostal, and three abdominal muscles, all of which exhibited minimal expression of MHC~/S1OW' The amount of MHC 2X (relative to total MHC composition) was similar in the diaphragm, genioglossus, and transversus abdominis muscles, while considerably less was detected in the rectus abdominis and external oblique muscles. These results indicate that MHC,x is broadly and variably distributed among respiratory muscles. Furthermore, these data suggest that a large portion of 2X fibers (containing MHC 2X ) , which cannot be detected by standard histochemical analysis, may be present in the genioglossus and transversus abdominis muscles as has been demonstrated for the diaphragm muscle. We speculate that an association exists between the level of MHC,x expression and frequency of respiratory recruitment.
Myosin comprises nearly the total composition of sarcomeric muscle thick filaments and its primary structure is composed of two a-helical myosin heavy chains (MHC) containing both the "rod" and "globular head" domains (1). Multiple MHC carboxy-terminal or rod domains aggregate to form the backbone structure of thick filaments. The MHC globular head domain contains the actin-binding site required for attachment to thin filaments (2) and the ATPase enzyme activity that provides energy for cross-bridge cycling (3). The S-2 "hinge" region, a portion of the rod, contains variable sequences that are associated with alterations in shortening velocity (4). Thus, fundamental structural and contractile properties of sarcomeric muscle fibers are determined by characteristics of their thick filaments and their MHC composition. Differences among myofibers regarding morphologic, enzymatic, and contractile characteristics have frequently been used to classify various fiber types (5). Recent data indicate that differences among these fiber types are associated with
(Received in original form May 2, 1991 and in revised form August 12, 1991) Address correspondence to: Robert D. Guthrie, M.D., Deparment of Pediattics, Magee-WomensHospital, 300 Halket Street, Pittsburgh, PA 152133180. Dr. LaFramboise's current address: Department of Biochemistry, J405, Health Sciences Building, SJ-70, University of Washington, Seattle, WA 98195. Abbreviations: N,N'-bis-methylene acrylarnide, Bis; myosin heavy chain, MHC; sodium dodecyl sulfate, SDS. Am. J. Respir. Cell Mol. BioI. Vol. 6. pp. 335-339, 1992
variations in MHC gene expression. Reiser and colleagues (6) have demonstrated a correlation of shortening velocity with MHC composition, whereas Schiaffino and associates (7) have been able to discriminate histochemically defined fiber classes using antibodies specific to MHC composition. Consequently, determination of MHC phenotype provides a structural basis for explaining the histochemical and functional differences among whole muscles and individual muscle fibers. Although identification of MHC isoforms has become increasingly important, the highly conserved nature of the MHC gene and protein family makes delineation of individual members difficult (8). Investigators rely primarily on differences in noncoding regions to discriminate among the genes of this multigene family (9). However, these differences do not extend to the MHC protein level where molecular weights are comparable and antigenicity is well conserved. We have recently utilized an electrophoretic and immunochemical assay to separate and identify the various MHC isoforms found in typical rat skeletal muscle (MHC~/'low, MHC 2A , MHC 2B• MHC 2X ) (10). Of particular interest was the finding that the primary respiratory muscle, the diaphragm muscle, contained an abundance of the MHC 2X isoform, whereas hind limb muscles expressed less of this MHC (11, 12). The purpose of the present study was to assay the MHC composition of various rat muscles known to exhibit respiratory activity and to determine whether MHC 2X is commonly expressed among these muscles.
Materials and Methods Muscle samples were obtained from four adult rats (SpragueDawley; 90 to 158 days of age) after administration of a le-
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thal dose of sodium pentobarbital (60 mg/kg). The soleus and tibialis anterior muscles, which are activated in the maintenance of posture and during locomotion, were removed from the hind limb. Other muscles were selected that exhibited varying degrees of respiratory activation. These included the costal diaphragm, crural diaphragm, midthoracic intercostals, external abdominal oblique, transversus abdominis, and rectus abdominis muscles, which were removed from the thoracoabdominal region, and the genioglossus muscle, which was removed from the hypopharynx. Samples were minced with iris scissors and extracted on ice for 40 min in 4 vol of buffer (300 mM NaCl, 100 mM NaH 2PO., 50 mM Na 2HPO., 1 mM MgCl" 10 mM Na.P 20 " 10 mM EDTA) at pH 6.5 as previously described (13). Extracts were centrifuged at 13,000 x g for 30 min at 4 C (Model 235C microfuge; Fisher Scientific, Fairlawn, NJ), and the supernatants recovered and diluted in 9 vol of 1 mM EDTA and 0.1% {j-mercaptoethanol (vol/vol). The diluted extracts were stored overnight at 4 0 C to allow precipitation of myosin filaments. The filament-containing solution was subsequently centrifuged at 13,000 x g for 30 min at4° C (Model 235C; Fisher Scientific) to form a pellet. The pellet was resuspended (1:1) in 0.5 M NaCl and 10 mM NaH 2PO. buffer then diluted 1:100 in sodium dodecyl sulfate (SOS) buffer composed of 62.5 mM Tris-HCI, 2 % (wt/vol) SOS, 10% (vol/vol) glycerol, 5 % (vol/vol) {j-mercaptoethanol, and 0.001 % (wt/vol) bromophenol blue at pH 6.8 (14). The samples were boiled for 2 to 3 min and stored at -80 0 C. The qualitative and quantitative distribution of MHC isoforms among study muscles as determined by electrophoresis was not altered by the use of protease inhibitors (leupeptin, pepstatin, phenylmethylsulfonyl fluoride) during the myosin extraction and filament precipitation steps, which was consistent with the work of Tsika and co-workers (15). Separation of MHC isoforms was accomplished using a previously described electrophoresis protocol (10). Briefly, electrophoresis was carried out on slabs (18 em X 16 em X 0.75 rnm) consisting of an 11.5-cm separating gel and a 4.5em stacking gel. Both gels were prepared from a 30% acrylamide stock solution containing 28.5 % (wt/vol) acrylamide and 1.5% (wt/vol) N,N'-bis-methylene acrylamide (Bis). Separating gels were poured at a T = 5% (T = total concentration of acrylamide + Bis) and C = 5 % (C = percentage of total monomer due to Bis) while stacking gels ofT = 3%, C = 5% were utilized. Separating gels polymerized 2 h at room temperature before addition of the stacking gel. Final concentrations of Tris-HCl in both stacking and separating gels were as previously described (14); however, glycerol (30% vol/vol) was added to the separating gel (16, 17). Polymerization was activated by addition of 0.03% (wt/vol) ammonium persulfate and 0.10% (vol/vol) tetramethylethylenediamine (TEMED). Electrophoresis was performed with Tris-glycine running buffer at pH = 8.3 (14) in a vertical slab gel apparatus (SE 600; Hoefer Scientific Instruments). One- to three-microliter volumes of myosin extract (500 to 1,500 ng) were loaded on the gels. After the plates were secured in the electophoresis unit, the power supply (EC400; E-C Apparatus) and the cooling unit (Endocal RBC-3; Neslab) were turned on. Electrophoresis was carried out for 22 to 24 h at constant 120 V. The temperature of the buffer was maintained at 150 C for the duration of the procedure. 0
Separating gels were stained with silver (18), and the location and density of bands determined with a transmittance densitometer (GS 365; Hoefer Scientific Instruments). A preliminary study compared software packages provided to calculate area under the peaks determined by the densitometer (GS 365 Electrophoresis Data Reduction System 1986; Metragraphics Softward Corp., Scotts Valley, CA). The "automatic" analysis generated a Gaussian distribution from the plotted peaks, and area determinations were derived from this distribution. The "manual" mode required the operator to define the region to be integrated from the original output of the densitometer. Both analyses yielded comparable results except when multiple, closely positioned bands were present. In those cases, the manual mode gave more accurate results compared with planimetric analysis performed by hand. For consistency, the data presented in this study were obtained utilizing the manual analysis. The identity of the MHC bands of the costal diaphragm and hind limb muscles has been previously determined using 5% SOS polyacrylamide gel electrophoresis and irnmunoblotting techniques (10, 12). Co-migration of the other respiratory and appendicular MHC isoforms with those of the costal diaphragm was used to identify the pattern of MHC expression in these muscles. Intrasample variability was typically 1 to 3% for a given MHC isoform, with the greatest difference encountered being 5.4 % in the expression of MHC 2A between two separate aliquots of a costal diaphragm sample with a reciprocal 5 % difference in the expression of MHC~/"ow, Interanimal variability as reflected by the coefficient variation for a given MHC isoform was typically in the range of 11 to 28%.
Results The electrophoretic technique used in this study allowed separation of the four MHC isoforms previously identified in typical adultrat skeletal muscles (Figure 1) (10, 11, 19), and their order of mobility was shown to be MHC~/"ow > MHC 2B > MHC 2X > MHC 2A (12). Separation of four distinct MHC isoforms in the tibialis anterior muscle provided a control for the sieving properties of each gel. Gels that did not resolve each of the four MHC isoforms were not analyzed. No additional MHC bands were detected during coelectrophoresis of samples of the various muscles under analysis. All of the respiratory muscles analyzed in this study contained the three fast and one slow MHC isoform in varying distribution. Costal and crural diaphragm samples contained approximately 25 % MHC~/'IO'" more than double the percentage of MHCw'l~ expressed in the other respiratory muscles (Figure 2). The three abdominal respiratory muscles, i.e., the transversus abdominis, external oblique, and rectus abdominis, expressed nearly equivalent low amounts of MHCw'l~ as did the intercostal muscles (Figure 2). The genioglossus muscle was notable in that it contained barely detectable amounts of MHC wS10W (2 ± 3%) (Figure 2). Among the hind limb muscles, the soleus muscle contained 90 ± 5 % MHC~/'IOW, with the balance comprised of MHC 2A (Figures 1 and 2), whereas the tibialis anterior muscle con; tained only 6 ± 4% MHC~/'IOW' with 94% of the myosin composition distributed among the three fast MHC isoforms (Figures 1 and 2). In contrast to the relatively high level of MHC~lsIOW ex-
LaFramboise, Watchko, Brozanski et al.: Myosin Heavy Chain Expression in Respiratory Muscles
100
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Figure 1. Separation of myosin heavy chain (MHC) isoforms by 5 % sodium dodecyl sulfate polyacrylamide gel electrophoresis. sol = soleus; ta = tibialis anterior; cos = costal diaphragm; cru = crural diaphragm; int = intercostal; rec = rectus abdominis; tran = transversus abdominis; eo = external abdominal oblique; gg = genioglossus. The migration of MHCs in order of electrophoretic mobility is MHC~/SIOW (slow) > MHC 28 (2B) > MHC2X (2X) > MHC 2A (2A).
pression in the diaphragm versus other respiratory muscles, costal and crural samples contained the lowest amount of MHC 28 expressed among respiratory muscles (costal, 7 ± 1%; crural, 12 ± 6%) (Figure 2). In fact, MHC 28 was the dominant isoform found in each of the other respiratory muscles and was expressed at levels from 4 to 5 times the amount found in the diaphragm (Figure 2). The highest levels of MHC 28 expression were found in the three abdominal muscles (Figure 2). Again, expression ofthe MHC 28 isoform in the intercostal muscles was comparable to that of the abdominal muscles (intercostal, 50 ± 6 %; transverse abdominis, 51 ± 8%) (Figure 2). The costal and crural diaphragm samples contained the highest levels of MHC 2A expression among the respiratory muscles tested (Figure 2). The three abdominal muscles contained the lowest amounts of this isoform, with the genioglossus and intercostal muscles falling in between (Figure 2). Differences in the distribution of MHC 2X between the diaphragm and the other respiratory muscles were not as distinct as was the case for the other MHC isoforms (Figure 2). The diaphragm samples contained substantial proportions of MHC 2X (costal, 32 ± 12%; crural, 41 ± 5%), but this was also the case for the genioglossus (37 ± 9 %) and the transversus abdominis (35 ± 10%) muscles. The lowest level of MHC 2X expression was in the external oblique and rectus abdominis muscles at approximately 15% (Figure 2).
Discussion This is the first analysis of the MHC composition (including MHC~/SIOW, MHC 2A , MHC 28 , and MHC 2X ) in a group of muscles sharing the property of repetitive respiratory activation. All of these muscles from the rat expressed each of the four MHC isoforms common to the appendicular musculature. The most apparent differences among the muscles studied were in the almost reciprocal distribution of the 2A and 2B MHC isoforms. More specifically, the diaphragm muscle differed from the accessory respiratory muscles by the incorporation of substantial amounts of MHC~/slow, MHC 2A , and MHC 2X but little MHC 28 in its composition. In contrast, the accessory respiratory muscles expressed a predominance of MHC 28 , with low amounts of MHC~/slow and MHC 2A • These differences were particularly evident in the abdominal mus-
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Figure 2. Proportion of specific MHC isoform in relation to total isomyosins for each muscle (mean ± SD). cos = costal diaphragm; cru = crural diaphragm; int = intercostal; gg = genioglossus; tran = transversus abdominis; eo = external abdominal oblique; rec = rectus abdominis; sol = soleus; ta = tibialis anterior.
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cles, e.g., the rectus abdominis and the external oblique, in which two thirds of the MHC composition was MHC 2B • MHC 2X was also low in these two abdominal muscles, but the transversus abdominis, genioglossus, and intercostal muscles expressed substantial proportions of MHC 2X that matched or approached the levels present in diaphragm samples. Electrophoretic and immunochemical techniques have made it possible to identify 2X myofibers (fibers expressing MHC 2X ) as a subset of "fast" fibers distinct from the 2A and 2B population (7, 10). These fibers appeared as 2B fibers when standard histochemical techniques were used (11, 20, 21). The results of this study indicate that MHC 2X constitutes a significant portion of the MHC complement of accessory respiratory muscles as it does in the costal diaphragm (12). It has been established that MHC isoforms rarely coexisted in adult rat diaphragm fibers, a third of which were 2X fibers (12). Comparable segregation of MHC isoforms in the accessory respiratory muscles would yield similar large populations of 2X fibers in the genioglossus and transversus abdominis muscles. It is important to resolve MHC 2X from MHC 2B and 2X from 2B fibers in order to correlate data from histochemical and physiologic assays. For example, 2X fibers such as in the rat genioglossus and abdominal muscles typically stained as "fast" 2B fibers according to standard myosin ATPase histochemistry (11, 19, 21). Motor units composed of 2B fibers contracted rapidly and fatigued quickly, whereas 2X motor units had a lower shortening velocity and higher fatigue resistance (11, 22, 23). Consequently, a substantial population of 2X fibers in the genioglossus and transversus abdominis muscles would reduce the shortening velocity and susceptibility to fatigue compared with muscles dominated by 2B fibers. Conversely, muscles that contain predominantly MHC 2B , e.g., the external abdominal oblique, would be expected to be more fatigable than those with small proportions of MHC 2B , e.g., the costal diaphragm, as a recent preliminary report suggests (24). The inability to discriminate between 2X and 2B fibers may explain reports that various tongue and laryngeal muscles exhibited high resistance to fatigue despite the presence of large numbers of 2B muscle fibers (25, 26). Long-term, phasic stimulation of rat muscles has been shown to induce the expression of MHC 2X , suggesting that this isoform may also be characteristic of muscles phasically activated in vivo such as the respiratory muscles (19,27). Although this study was not designed to test this proposition, it is interesting that the results were consistent with that hypothesis. The external oblique and rectus abdominis muscles are reported to be the least often recruited for ventilatory support of the accessory muscles of the anterolateral abdominal wall examined in our study (28, 29), and they displayed the lowest levels of MHC 2X expression. Higher levels of MHC 2X were present in the diaphragm and genioglossus muscles, which contract with each inspiration (30). Comparable levels of MHC 2X to the diaphragm and genioglossus were found in the transversus abdominis muscle-an abdominal muscle that is thought to be consistently activated during expiration (28). The intercostal samples obtained for this study involved both inspiratory and ex-
piratory regions but insufficient functional data are available to compare their activation patterns and MHC 2X levels. The rat MHC multigene family consists of members clustered on two chromosomes that undergo tissue-specific regulation (9, 31). The factors that coordinate expression of MHC isoforms within mature muscles have not been determined although changes occur at the protein level as a result of pharmacologic manipulations, adjustments in neural activation patterns, and alterations in work output (32-35). This study, in combination with other recent reports, emphasizes the importance of the MHC 2X isoform as a critical member of the rat MHC contractile protein family (7, 19). It should be noted that the MHC 2X isoform has been identified in the rat and mouse based on immunochemical and electrophoretic analyses established for these readily available laboratory animals. Further studies are required to determine whether this particular isoform or an analogous form exists in humans. Acknowledgments: The writers thank Maureen Davis for assistance in preparation of the manuscript. We thank Marcia Ontell, Ph.D., for her critical review of the manuscript and for her research advice and support. This research was supported by grants from the Richard King Mellon Foundation (to Dr. Guthrie), the Twenty Five Club of Magee-Womens Hospital (to Dr. Guthrie), a Research Grant Award from the American Lung Association (to Dr. Brozanski), and Grant HL-02491 from the National Heart, Blood and Lung Institute, National Institutes of Health (to Dr. Watchko).
References 1. Harrington, W. F., and M. E. Rodgers. 1985. Myosin. Annu. Rev. Bio-
chem. 53:35-73. 2. Chaussepied, P., and M. F. Morales. 1988. Modifying preselected sites on proteins: the stretch of residues 633-642 of the myosin heavy chain is part of the actin-binding site. Proc. Natl. Acad. Sci. USA 85: 7471-7475. 3. Maruta, S., T. Miyanishi, andG. Matsuda. 1989. LocalizationoftheATPbinding site in the 23kDa and 20kDa regions of the heavy chain of the skeletal muscle myosin head. Eur. J. Biochem. 184:213-221. 4. Collier, V. L., W. A. Kronert, P. T. O'Donnell, K. A. Edwards, and S. I. Bernstein. 1990. Alternative myosin hinge regions are utilized in a tissuespecific fashion that correlates with muscle contraction speed. Genes Dev. 4:885-895. 5. Burke, R. E., D. N. Levine, T. Tsairis, and F. E. Zajac. 1973. Physiological types and histochemical profiles in motor units of the cat gastrocnemius. J. Physiol. (Lond.) 234:723-748. 6. Reiser, P. J., R. L. Moss, G. C. Giulian, and M. L. Greaser. 1985. Shortening velocity in single filters from adult rabbit soleus muscles is correlated with myosin heavy chain composition. J. BioI. Chem. 260: 9077-9080. 7. Schiaffino, S., L. Gorza, S. Sartore et al. Three myosin heavy chain isoforms in type 2 skeletal muscle fibres. J. Muscle Res. Cell Motif. 10:197-205. 8. Miller, J. B., S. B. Teal, and F. E. Stockdale. 1989. Evolutionary conserved sequences of striated muscle myosin heavy chain isoforms. J. Bioi.
Chem.264:13122-13130. 9. Mahdavi, V., S. Izumo, and B. Nadal-Ginard. 1987. Developmental and hormonal regulation of sarcomeric myosin heavy chain gene family. eire.
Res. 60:804-814. 10. LaFramboise, W. A., M. 1. Daood, R. D. Guthrie, P. Moretti, S. Schiaffino, and M. Ontell. 1990. Electrophoretic separation and immunological identification of type 2X myosin heavy chain in rat skeletal muscle.
Biochem. Biophys. Acta 1035:109-112. II. Schiaffino, S., L. Gorza, and S. Ausoni. 1990. Muscle fiber types expressing different myosin heavy chain isoforms. Their functional properties and adaptive capacity. In The Dynamic State of Muscle Fibers. D. Pette, editor. De Gruyter, Berlin. 329-341. 12. LaFramboise, W. A., M. J. Daood, R. D. Guthrie et al. 1991. Emergence of the mature myosin phenotype in the rat diaphragm muscle. Dev. Bioi. 144: 1-15. 13. Butler-Browne, G. S., and R. G. Whalen. 1984. Myosin isozyme transitions occurring during the postnatal development of the rat soleus muscle.
Dev. Bioi. 102:324-334. 14. Laemmli, U. K. 1970. Cleavage of structural proteins during the assembly
LaFramboise, Watchko, Brozanski et al.: Myosin Heavy Chain Expression in Respiratory Muscles
of the head of bacteriophage T4. Nature 227:680-685. 15. Tsika, R. W., R. E. Herrick, and K. M. Baldwin. 1987. Subunit composition of rodent isomyosins and their distribution in the hindlimb skeletal muscles. J. Appl. Physiol. 63:2101-2110. 16. Carraro, D., and C. Catani. 1983. A sensitive SDS-PAGE method separating myosin heavy chain isoforms of rat skeletal muscles reveals the heterogeneous nature of the embryonic myosin. Biochem. Biophys. Res. Commun. 116:793-802. 17. Danieli-Betto, D., E. Zerbato, and R. Betto. 1986. Type 1, 2A, and 2B myosin heavy chain electrophoretic analysis of rat muscle fibers. Biochem. Biophys. Res. Commun. 138:981-987. 18. Oakley, B. R., D. R. Kirsch, and N. R. Morris. 1980. A simplified ultrasensitive silver stain for detecting proteins in polyacrylamide gels. Anal. Biochem. 105:361-363. 19. Termin, A., R. S. Staron, and D. Pette. 1989. Myosin heavy chain isoforms in histochemically defined fiber types of rat muscle. Histochemistry 92:453-457. 20. Eddinger, T. J., and R. L. Moss. 1987. Mechanical properties of skinned single fibers of identified types from rat diaphragm. Am. J. Physiol. 253:C21O-C218. 21. Gorza, L. 1990. Identification of a novel type 2 fiber population in mammalian skeletal muscle by combined use of histochemical myosin ATPase and anti-myosin monoclonal antibodies. J. Histochem. Cytochem. 38: 257-265. 22. Close, R. I. 1972. Dynamic properties of mammalian skeletal muscles. Physiol. Rev. 52:129-197. 23. Schiaffino, S., S. Ausoni, L. Garza, L. Saggin, K. Gundersen, and T. Lomo. 1988. Myosin heavy chain isoforms and velocity of shortening of type 2 skeletal muscle fibers. Acta Physiol. Scand. 134:565-566. 24. Watchko, J. F., B. S. Brozanski, T. L. O'Day, R. D. Guthrie, and G. C. Sieck. 1991. In vitro contractile and endurance properties of the external abdominal oblique and costal diaphragm muscles during development. Am. Rev. Respir. Dis. 143(Suppl.):A561. (Abstr.)
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25. Hinrichsen, C., and A. Dulhunty. 1982. The contractile properties, histochemistry, ultrastructure and electrophysiology of the cricothyroid and posterior cricoarytenoid muscles in the rat. J. Muscle Res. Cell Motil. 3:169-190. 26. Van Lunteren, E" R. J. Salome, P. Manubay, G. S. Supinski, and T. E. Dick. 1990. Contractile and endurance properties of geniohyoid and diaphragm muscles. J. Appl. Physiol. 69:1991-1997. 27. Ausoni, S., L. Gorza, S. Schiaffino, K. Gundersen, and T. Lomo. 1990. Expression of myosin heavy chain isoforrns in stimulated fast and slow rat muscles. J. Neurosci. 10:153-160. 28. Gilmartin, J. J., V. Ninane, and A. DeTroyer. 1987. Abdominal muscle use during breathing in the anesthetized dog. Respir. Physiol. 70: 159-171. 29. DeTroyer, A., M. Estenne, V. Ninane, D. Van Gansbeke, and M. Gorini. 1990. Transversus abdominis muscle function in humans. J. Appl. Physiol. 68:1010-1016. 30. Sauerland, E. K., and S. P. Mitchell. 1975. Electromyographic activity of the intrinsic and extrinsic muscles of the human tongue. Tex. Rep. BioI. Med. 33:445-455. 31. Emerson, C. P., and S. I. Bernstein. 1987. Molecular genetics of myosin. Annu. Rev. Biochem. 56:695-726. 32. Carraro, D., L. Dalla-Libera, C. Catani, and D. Danieli-Betto. 1982. Chronic denervation of rat diaphragm: selective maintenance of adult fast myosin heavy chains. Muscle Nerve 5 :515-524. 33. Mabuchi, K., D. Szvetko, K. Pinter, and F. A. Sreter. 1982. Type 2B and 2A fiber transformation in intermittently stimulated rabbit muscles. Am. J. Physiol. 242: C373-C38I. 34. Gregory, P., R. B. Low, and W. S. Stirewalt. 1986. Changes in skeletalmuscle myosin isoenzymes with hypertrophy and exercise. Biochem. J. 238:55-63. 35. Izumo, S., B. Nadal-Ginard, and V. Mahdavi. 1986. All members of the MHC multigene family respond to thyroid hormone in a highly tissuespecific manner. Science 231:597-600.
Myosin heavy chain (MHC) isoforms of hind limb adult rat muscles and muscles with a range of respiratory activities were analyzed by a sodium dodecyl sulfate polyacrylamide gel electrophoresis technique that allowed electrophoretic separation of the three fast and one slow MHC isoform found in typical rat muscle. Costal and crural diaphragm muscle samples expressed a mixture of MHC~/S1OW' MHC 2A , and MHC 2X but little MHC 2B • In contrast, MHC 2B was the dominant MHC isoform in the genioglossus, intercostal, and three abdominal muscles, all of which exhibited minimal expression of MHC~/S1OW' The amount of MHC 2X (relative to total MHC composition) was similar in the diaphragm, genioglossus, and transversus abdominis muscles, while considerably less was detected in the rectus abdominis and external oblique muscles. These results indicate that MHC,x is broadly and variably distributed among respiratory muscles. Furthermore, these data suggest that a large portion of 2X fibers (containing MHC 2X ) , which cannot be detected by standard histochemical analysis, may be present in the genioglossus and transversus abdominis muscles as has been demonstrated for the diaphragm muscle. We speculate that an association exists between the level of MHC,x expression and frequency of respiratory recruitment.
Myosin comprises nearly the total composition of sarcomeric muscle thick filaments and its primary structure is composed of two a-helical myosin heavy chains (MHC) containing both the "rod" and "globular head" domains (1). Multiple MHC carboxy-terminal or rod domains aggregate to form the backbone structure of thick filaments. The MHC globular head domain contains the actin-binding site required for attachment to thin filaments (2) and the ATPase enzyme activity that provides energy for cross-bridge cycling (3). The S-2 "hinge" region, a portion of the rod, contains variable sequences that are associated with alterations in shortening velocity (4). Thus, fundamental structural and contractile properties of sarcomeric muscle fibers are determined by characteristics of their thick filaments and their MHC composition. Differences among myofibers regarding morphologic, enzymatic, and contractile characteristics have frequently been used to classify various fiber types (5). Recent data indicate that differences among these fiber types are associated with
(Received in original form May 2, 1991 and in revised form August 12, 1991) Address correspondence to: Robert D. Guthrie, M.D., Deparment of Pediattics, Magee-WomensHospital, 300 Halket Street, Pittsburgh, PA 152133180. Dr. LaFramboise's current address: Department of Biochemistry, J405, Health Sciences Building, SJ-70, University of Washington, Seattle, WA 98195. Abbreviations: N,N'-bis-methylene acrylarnide, Bis; myosin heavy chain, MHC; sodium dodecyl sulfate, SDS. Am. J. Respir. Cell Mol. BioI. Vol. 6. pp. 335-339, 1992
variations in MHC gene expression. Reiser and colleagues (6) have demonstrated a correlation of shortening velocity with MHC composition, whereas Schiaffino and associates (7) have been able to discriminate histochemically defined fiber classes using antibodies specific to MHC composition. Consequently, determination of MHC phenotype provides a structural basis for explaining the histochemical and functional differences among whole muscles and individual muscle fibers. Although identification of MHC isoforms has become increasingly important, the highly conserved nature of the MHC gene and protein family makes delineation of individual members difficult (8). Investigators rely primarily on differences in noncoding regions to discriminate among the genes of this multigene family (9). However, these differences do not extend to the MHC protein level where molecular weights are comparable and antigenicity is well conserved. We have recently utilized an electrophoretic and immunochemical assay to separate and identify the various MHC isoforms found in typical rat skeletal muscle (MHC~/'low, MHC 2A , MHC 2B• MHC 2X ) (10). Of particular interest was the finding that the primary respiratory muscle, the diaphragm muscle, contained an abundance of the MHC 2X isoform, whereas hind limb muscles expressed less of this MHC (11, 12). The purpose of the present study was to assay the MHC composition of various rat muscles known to exhibit respiratory activity and to determine whether MHC 2X is commonly expressed among these muscles.
Materials and Methods Muscle samples were obtained from four adult rats (SpragueDawley; 90 to 158 days of age) after administration of a le-
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thal dose of sodium pentobarbital (60 mg/kg). The soleus and tibialis anterior muscles, which are activated in the maintenance of posture and during locomotion, were removed from the hind limb. Other muscles were selected that exhibited varying degrees of respiratory activation. These included the costal diaphragm, crural diaphragm, midthoracic intercostals, external abdominal oblique, transversus abdominis, and rectus abdominis muscles, which were removed from the thoracoabdominal region, and the genioglossus muscle, which was removed from the hypopharynx. Samples were minced with iris scissors and extracted on ice for 40 min in 4 vol of buffer (300 mM NaCl, 100 mM NaH 2PO., 50 mM Na 2HPO., 1 mM MgCl" 10 mM Na.P 20 " 10 mM EDTA) at pH 6.5 as previously described (13). Extracts were centrifuged at 13,000 x g for 30 min at 4 C (Model 235C microfuge; Fisher Scientific, Fairlawn, NJ), and the supernatants recovered and diluted in 9 vol of 1 mM EDTA and 0.1% {j-mercaptoethanol (vol/vol). The diluted extracts were stored overnight at 4 0 C to allow precipitation of myosin filaments. The filament-containing solution was subsequently centrifuged at 13,000 x g for 30 min at4° C (Model 235C; Fisher Scientific) to form a pellet. The pellet was resuspended (1:1) in 0.5 M NaCl and 10 mM NaH 2PO. buffer then diluted 1:100 in sodium dodecyl sulfate (SOS) buffer composed of 62.5 mM Tris-HCI, 2 % (wt/vol) SOS, 10% (vol/vol) glycerol, 5 % (vol/vol) {j-mercaptoethanol, and 0.001 % (wt/vol) bromophenol blue at pH 6.8 (14). The samples were boiled for 2 to 3 min and stored at -80 0 C. The qualitative and quantitative distribution of MHC isoforms among study muscles as determined by electrophoresis was not altered by the use of protease inhibitors (leupeptin, pepstatin, phenylmethylsulfonyl fluoride) during the myosin extraction and filament precipitation steps, which was consistent with the work of Tsika and co-workers (15). Separation of MHC isoforms was accomplished using a previously described electrophoresis protocol (10). Briefly, electrophoresis was carried out on slabs (18 em X 16 em X 0.75 rnm) consisting of an 11.5-cm separating gel and a 4.5em stacking gel. Both gels were prepared from a 30% acrylamide stock solution containing 28.5 % (wt/vol) acrylamide and 1.5% (wt/vol) N,N'-bis-methylene acrylamide (Bis). Separating gels were poured at a T = 5% (T = total concentration of acrylamide + Bis) and C = 5 % (C = percentage of total monomer due to Bis) while stacking gels ofT = 3%, C = 5% were utilized. Separating gels polymerized 2 h at room temperature before addition of the stacking gel. Final concentrations of Tris-HCl in both stacking and separating gels were as previously described (14); however, glycerol (30% vol/vol) was added to the separating gel (16, 17). Polymerization was activated by addition of 0.03% (wt/vol) ammonium persulfate and 0.10% (vol/vol) tetramethylethylenediamine (TEMED). Electrophoresis was performed with Tris-glycine running buffer at pH = 8.3 (14) in a vertical slab gel apparatus (SE 600; Hoefer Scientific Instruments). One- to three-microliter volumes of myosin extract (500 to 1,500 ng) were loaded on the gels. After the plates were secured in the electophoresis unit, the power supply (EC400; E-C Apparatus) and the cooling unit (Endocal RBC-3; Neslab) were turned on. Electrophoresis was carried out for 22 to 24 h at constant 120 V. The temperature of the buffer was maintained at 150 C for the duration of the procedure. 0
Separating gels were stained with silver (18), and the location and density of bands determined with a transmittance densitometer (GS 365; Hoefer Scientific Instruments). A preliminary study compared software packages provided to calculate area under the peaks determined by the densitometer (GS 365 Electrophoresis Data Reduction System 1986; Metragraphics Softward Corp., Scotts Valley, CA). The "automatic" analysis generated a Gaussian distribution from the plotted peaks, and area determinations were derived from this distribution. The "manual" mode required the operator to define the region to be integrated from the original output of the densitometer. Both analyses yielded comparable results except when multiple, closely positioned bands were present. In those cases, the manual mode gave more accurate results compared with planimetric analysis performed by hand. For consistency, the data presented in this study were obtained utilizing the manual analysis. The identity of the MHC bands of the costal diaphragm and hind limb muscles has been previously determined using 5% SOS polyacrylamide gel electrophoresis and irnmunoblotting techniques (10, 12). Co-migration of the other respiratory and appendicular MHC isoforms with those of the costal diaphragm was used to identify the pattern of MHC expression in these muscles. Intrasample variability was typically 1 to 3% for a given MHC isoform, with the greatest difference encountered being 5.4 % in the expression of MHC 2A between two separate aliquots of a costal diaphragm sample with a reciprocal 5 % difference in the expression of MHC~/"ow, Interanimal variability as reflected by the coefficient variation for a given MHC isoform was typically in the range of 11 to 28%.
Results The electrophoretic technique used in this study allowed separation of the four MHC isoforms previously identified in typical adultrat skeletal muscles (Figure 1) (10, 11, 19), and their order of mobility was shown to be MHC~/"ow > MHC 2B > MHC 2X > MHC 2A (12). Separation of four distinct MHC isoforms in the tibialis anterior muscle provided a control for the sieving properties of each gel. Gels that did not resolve each of the four MHC isoforms were not analyzed. No additional MHC bands were detected during coelectrophoresis of samples of the various muscles under analysis. All of the respiratory muscles analyzed in this study contained the three fast and one slow MHC isoform in varying distribution. Costal and crural diaphragm samples contained approximately 25 % MHC~/'IO'" more than double the percentage of MHCw'l~ expressed in the other respiratory muscles (Figure 2). The three abdominal respiratory muscles, i.e., the transversus abdominis, external oblique, and rectus abdominis, expressed nearly equivalent low amounts of MHCw'l~ as did the intercostal muscles (Figure 2). The genioglossus muscle was notable in that it contained barely detectable amounts of MHC wS10W (2 ± 3%) (Figure 2). Among the hind limb muscles, the soleus muscle contained 90 ± 5 % MHC~/'IOW, with the balance comprised of MHC 2A (Figures 1 and 2), whereas the tibialis anterior muscle con; tained only 6 ± 4% MHC~/'IOW' with 94% of the myosin composition distributed among the three fast MHC isoforms (Figures 1 and 2). In contrast to the relatively high level of MHC~lsIOW ex-
LaFramboise, Watchko, Brozanski et al.: Myosin Heavy Chain Expression in Respiratory Muscles
100
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Figure 1. Separation of myosin heavy chain (MHC) isoforms by 5 % sodium dodecyl sulfate polyacrylamide gel electrophoresis. sol = soleus; ta = tibialis anterior; cos = costal diaphragm; cru = crural diaphragm; int = intercostal; rec = rectus abdominis; tran = transversus abdominis; eo = external abdominal oblique; gg = genioglossus. The migration of MHCs in order of electrophoretic mobility is MHC~/SIOW (slow) > MHC 28 (2B) > MHC2X (2X) > MHC 2A (2A).
pression in the diaphragm versus other respiratory muscles, costal and crural samples contained the lowest amount of MHC 28 expressed among respiratory muscles (costal, 7 ± 1%; crural, 12 ± 6%) (Figure 2). In fact, MHC 28 was the dominant isoform found in each of the other respiratory muscles and was expressed at levels from 4 to 5 times the amount found in the diaphragm (Figure 2). The highest levels of MHC 28 expression were found in the three abdominal muscles (Figure 2). Again, expression ofthe MHC 28 isoform in the intercostal muscles was comparable to that of the abdominal muscles (intercostal, 50 ± 6 %; transverse abdominis, 51 ± 8%) (Figure 2). The costal and crural diaphragm samples contained the highest levels of MHC 2A expression among the respiratory muscles tested (Figure 2). The three abdominal muscles contained the lowest amounts of this isoform, with the genioglossus and intercostal muscles falling in between (Figure 2). Differences in the distribution of MHC 2X between the diaphragm and the other respiratory muscles were not as distinct as was the case for the other MHC isoforms (Figure 2). The diaphragm samples contained substantial proportions of MHC 2X (costal, 32 ± 12%; crural, 41 ± 5%), but this was also the case for the genioglossus (37 ± 9 %) and the transversus abdominis (35 ± 10%) muscles. The lowest level of MHC 2X expression was in the external oblique and rectus abdominis muscles at approximately 15% (Figure 2).
Discussion This is the first analysis of the MHC composition (including MHC~/SIOW, MHC 2A , MHC 28 , and MHC 2X ) in a group of muscles sharing the property of repetitive respiratory activation. All of these muscles from the rat expressed each of the four MHC isoforms common to the appendicular musculature. The most apparent differences among the muscles studied were in the almost reciprocal distribution of the 2A and 2B MHC isoforms. More specifically, the diaphragm muscle differed from the accessory respiratory muscles by the incorporation of substantial amounts of MHC~/slow, MHC 2A , and MHC 2X but little MHC 28 in its composition. In contrast, the accessory respiratory muscles expressed a predominance of MHC 28 , with low amounts of MHC~/slow and MHC 2A • These differences were particularly evident in the abdominal mus-
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Figure 2. Proportion of specific MHC isoform in relation to total isomyosins for each muscle (mean ± SD). cos = costal diaphragm; cru = crural diaphragm; int = intercostal; gg = genioglossus; tran = transversus abdominis; eo = external abdominal oblique; rec = rectus abdominis; sol = soleus; ta = tibialis anterior.
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cles, e.g., the rectus abdominis and the external oblique, in which two thirds of the MHC composition was MHC 2B • MHC 2X was also low in these two abdominal muscles, but the transversus abdominis, genioglossus, and intercostal muscles expressed substantial proportions of MHC 2X that matched or approached the levels present in diaphragm samples. Electrophoretic and immunochemical techniques have made it possible to identify 2X myofibers (fibers expressing MHC 2X ) as a subset of "fast" fibers distinct from the 2A and 2B population (7, 10). These fibers appeared as 2B fibers when standard histochemical techniques were used (11, 20, 21). The results of this study indicate that MHC 2X constitutes a significant portion of the MHC complement of accessory respiratory muscles as it does in the costal diaphragm (12). It has been established that MHC isoforms rarely coexisted in adult rat diaphragm fibers, a third of which were 2X fibers (12). Comparable segregation of MHC isoforms in the accessory respiratory muscles would yield similar large populations of 2X fibers in the genioglossus and transversus abdominis muscles. It is important to resolve MHC 2X from MHC 2B and 2X from 2B fibers in order to correlate data from histochemical and physiologic assays. For example, 2X fibers such as in the rat genioglossus and abdominal muscles typically stained as "fast" 2B fibers according to standard myosin ATPase histochemistry (11, 19, 21). Motor units composed of 2B fibers contracted rapidly and fatigued quickly, whereas 2X motor units had a lower shortening velocity and higher fatigue resistance (11, 22, 23). Consequently, a substantial population of 2X fibers in the genioglossus and transversus abdominis muscles would reduce the shortening velocity and susceptibility to fatigue compared with muscles dominated by 2B fibers. Conversely, muscles that contain predominantly MHC 2B , e.g., the external abdominal oblique, would be expected to be more fatigable than those with small proportions of MHC 2B , e.g., the costal diaphragm, as a recent preliminary report suggests (24). The inability to discriminate between 2X and 2B fibers may explain reports that various tongue and laryngeal muscles exhibited high resistance to fatigue despite the presence of large numbers of 2B muscle fibers (25, 26). Long-term, phasic stimulation of rat muscles has been shown to induce the expression of MHC 2X , suggesting that this isoform may also be characteristic of muscles phasically activated in vivo such as the respiratory muscles (19,27). Although this study was not designed to test this proposition, it is interesting that the results were consistent with that hypothesis. The external oblique and rectus abdominis muscles are reported to be the least often recruited for ventilatory support of the accessory muscles of the anterolateral abdominal wall examined in our study (28, 29), and they displayed the lowest levels of MHC 2X expression. Higher levels of MHC 2X were present in the diaphragm and genioglossus muscles, which contract with each inspiration (30). Comparable levels of MHC 2X to the diaphragm and genioglossus were found in the transversus abdominis muscle-an abdominal muscle that is thought to be consistently activated during expiration (28). The intercostal samples obtained for this study involved both inspiratory and ex-
piratory regions but insufficient functional data are available to compare their activation patterns and MHC 2X levels. The rat MHC multigene family consists of members clustered on two chromosomes that undergo tissue-specific regulation (9, 31). The factors that coordinate expression of MHC isoforms within mature muscles have not been determined although changes occur at the protein level as a result of pharmacologic manipulations, adjustments in neural activation patterns, and alterations in work output (32-35). This study, in combination with other recent reports, emphasizes the importance of the MHC 2X isoform as a critical member of the rat MHC contractile protein family (7, 19). It should be noted that the MHC 2X isoform has been identified in the rat and mouse based on immunochemical and electrophoretic analyses established for these readily available laboratory animals. Further studies are required to determine whether this particular isoform or an analogous form exists in humans. Acknowledgments: The writers thank Maureen Davis for assistance in preparation of the manuscript. We thank Marcia Ontell, Ph.D., for her critical review of the manuscript and for her research advice and support. This research was supported by grants from the Richard King Mellon Foundation (to Dr. Guthrie), the Twenty Five Club of Magee-Womens Hospital (to Dr. Guthrie), a Research Grant Award from the American Lung Association (to Dr. Brozanski), and Grant HL-02491 from the National Heart, Blood and Lung Institute, National Institutes of Health (to Dr. Watchko).
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