Article |
Drosophila paramyosin is important for myoblast fusion and essential for myofibril formation
2 Molecular Biology Institute, San Diego State University, San Diego, CA 92182
3 Department of Genetics, University of Cambridge, Cambridge, CB2 3EH, UK
Address correspondence to Sanford I. Bernstein, Dept. of Biology, San Diego State University, 5500 Campanile Dr., Life Sciences 371, San Diego, CA 92182-4614. Tel.: (619) 594-5629. Fax: (619) 594-5676. E-mail: sbernst{at}sunstroke.sdsu.edu
![]() |
Abstract |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Key Words: contractile protein; cytoskeleton; contraction; muscle; thick filament
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The mechanism by which thick filaments attain precise regularity in striated muscle remains unknown. Although previous studies demonstrated that myosin possesses self-assembly ability (Huxley, 1963), myosin filaments formed in vitro lack important features of in vivo thick filaments. Accumulating evidence suggests that the assembly of myosin into thick filaments of distinct lengths, diameters, and flexural rigidities requires the presence of other proteins (Ziegler et al., 1996). In vertebrate muscles, several additional proteins are associated with thick filaments: C-protein, H-protein (mammals) or 86-kD protein (birds), M-protein, myomesin, M-creatine kinase, skelemin, adenosine monophosphate deaminase, and titin (Epstein and Fischman, 1991; Barral and Epstein, 1999). Analogues of some of these proteins exist in invertebrates. For instance, in Drosophila melanogaster, three members of the titin family have been identified: projectin (Ayme-Southgate et al., 1991), kettin (Hakeda et al., 2000; Kulke et al., 2001), and D-titin (Machado and Andrew, 2000; Zhang et al., 2000).
Some proteins are unique to thick filaments of invertebrate striated muscles. For instance, D. melanogaster possesses paramyosin (Vinós et al., 1991; Becker et al., 1992; Maroto et al., 1995), miniparamyosin (Becker et al., 1992; Maroto et al., 1995, 1996), myosin rod protein (Standiford et al., 1997), and flightin (Vigoreaux et al., 1993; Reedy et al., 2000). Caenorhabditis elegans has paramyosin (Mackenzie and Epstein, 1980; Kagawa et al., 1989) and -, ß-, and
-filagenin (Liu et al., 1998, 2000). The diversity of thick filament components may account for the highly variable lengths and diameters of muscle thick filaments from different species.
Paramyosin and myosin are the most abundant invertebrate thick filament proteins. Paramyosin is present in all invertebrate muscles studied (Maroto et al., 1995). This protein is a rodlike molecule with high -helical content in its long central domain. This domain is flanked by short nonhelical NH2- and COOH-terminal regions. Two paramyosin monomers can dimerize into a coiled coil. Analysis of paramyosin and myosin heavy chain rod sequences revealed a remarkable pattern of alternating concentrations of charge associated with a 28-residue repeat (Cohen and Parry, 1998). Interactions between these segments of opposite charge are thought to play a major role in the assembly of both of these proteins into the thick filaments (McLachlan and Karn, 1982; Kagawa et al., 1989).
Drosophila paramyosin is a protein comprised of 879 amino acid residues with a molecular mass of 105 kD. The central 823 residues form an
helix; this is flanked by nonhelical domains of 32 NH2-terminal and 24 COOH-terminal residues (Becker et al., 1992; Maroto et al., 1995). By using an alternative promoter and alternative RNA splicing, the paramyosin gene produces a transcript encoding miniparamyosin. Miniparamyosin shares its COOH-terminal region with paramyosin and has a unique NH2-terminal domain of 114 amino acids (Becker et al., 1992; Maroto et al., 1995). Paramyosin is present in both embryonic and adult muscles. However, miniparamyosin is only present in adult musculature (Maroto et al., 1996).
Paramyosin is thought to facilitate thick filament assembly. Mutant analysis in C. elegans shows that thick filament length and diameter are affected by paramyosin content (Mackenzie and Epstein, 1980). Based on biochemical, genetic, and structural studies of C. elegans thick filaments, Epstein et al. (1995) proposed a thick filament structure model in C. elegans. In this model, paramyosin, together with the filagenins, forms the tubular thick filament core in which seven paramyosin subfilaments are interconnected by an internal sleeve of filagenins. Each paramyosin subfilament contains four strands of paramyosin coiled coils throughout its length. The motor protein myosin attaches on the surface of the core, yielding the functional thick filament (Epstein et al., 1995; Müller et al., 2001). Notably, no homologues of C. elegans filagenins have been identified in D. melanogaster and miniparamyosin and flightin do not exist in C. elegans. Thus, the process of thick filament assembly in different organisms and the molecular mechanism involved in the formation of different types of myofibrils in each organism remain to be clarified.
To investigate the function of D. melanogaster paramyosin, we used a genetic approach. We functionally disabled the paramyosin gene by mobilizing a P element in its promoter region. We observed that homozygous paramyosin mutants die as late embryos and that myofibril assembly is disrupted. Surprisingly, we found that paramyosin is also required for myoblast fusion. In the absence of paramyosin, myoblast fusion is sometimes blocked, resulting in the absence of some muscle fibers. We rescued the homozygous paramyosin mutant to adulthood using a paramyosin transgene, thereby proving that defects observed in myoblast fusion and myofibril assembly arise specifically from the absence of paramyosin. Antibody localization confirmed that paramyosin is present in myoblasts before fusion and is localized in discrete foci at the contact sites of fusing myoblasts. Our results demonstrate that paramyosin functions as a cytoplasmic protein in early embryonic development and is important for myoblast fusion before its assembly into thick filaments.
![]() |
Results |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
We used a paramyosin-specific antibody to determine paramyosin expression levels in the lines that could be rescued by the paramyosin transgene. We identified one strongly hypomorphic mutant in which the paramyosin expression level is <1% (Fig. 2 A). We refer to this line as prm1. Homozygous prm1 mutants die at the late embryo stage and are rescued to adulthood by the pm transgene.
|
The mutation in prm1 is paramyosin-specific
Because paramyosin and miniparamyosin are encoded at the same locus (Fig. 1), we wondered if the mutation of the paramyosin promoter region affects miniparamyosin expression. Expression of paramyosin and miniparamyosin in pm-transgenerescued flies was analyzed by Western blotting using paramyosin and miniparamyosin antibodies. Results showed that both paramyosin and miniparamyosin expression in rescued flies are normal (Fig. 2 B). Because the transgene used in rescue does not contain the miniparamyosin-specific exon, all the miniparamyosin expressed in rescued adult flies must be encoded by the endogenous miniparamyosin locus. This indicates that miniparamyosin expression is not affected by the deletion in the paramyosin promoter region caused by imprecise P element excision.
The deletion in prm1 uncovers a portion of the 284-bp CG13306, an open reading frame for which no cDNAs have been reported (Fig. 1). Deletion of CG13306 is not the cause of the mutant phenotypes we document because homozygous prm1 mutants could be rescued with the pm transgene (see Results), which is truncated within the CG13306 open reading frame.
Mutation of the paramyosin gene severely affects muscle development
Homozygous prm1 embryos display grossly normal morphology including normal segmentation and epidermal denticle belts (unpublished data), but they fail to hatch at late embryonic stage 17. Compared with normal embryos, manually hatched late stage 17 prm1 homozygous embryos are flat and motionless, suggesting possible muscle defects.
We investigated the muscle development of homozygous prm1 embryos by staining muscle fibers of late stage 16 embryos with muscle myosin heavy chain antibody (Fig. 3). At this stage, the process of body wall muscle development is complete and each of the abdominal hemisegments (A2-A7) has 30 muscles (Bate, 1993). Each of these muscles is unique in terms of its position, size, sites of attachment, and patterns of innervation. Every muscle has a full complement of nuclei, which remains unchanged until the end of larval life (Bate, 1990, 1993). In prm1 mutant embryos, the regular muscle pattern is disrupted. At stage 16, over 95% of the mutant embryos have obviously detectable muscle losses and aberrantly shaped muscle fibers. Compared with wild-type counterparts, these aberrant fibers are shorter or thinner and have fewer nuclei (unpublished data). Muscle fiber absence in mutant embryos usually occurs in groups. It can take place at any position in the embryo, in all three major muscle subtypes: dorsal, lateral, and ventral. In embryos in which muscle development is severely disrupted, we observed an absence of several muscle groups in different segments of the same embryo. However, muscles that successfully developed are located in normal positions. We never observed cross-segment fibers or duplicated fibers. The mutant phenotype of muscle fiber losses in prm1 is confirmed in homozygotes for the deficiency Df(3L)hi22ki, which covers the paramyosin gene (unpublished data). The muscle fiber phenotype in the deficiency line is slightly worse than in prm1, reflecting leaky expression of paramyosin in prm1.
|
|
|
|
|
|
Rescue of prm1 mutant phenotypes with the paramyosin transgene
To verify that the failure of myoblast fusion and aberrant myofibril assembly of somatic embryo body wall muscles in prm1 mutant embryos are indeed caused by the paramyosin mutation, we analyzed the muscle development and myofibril assembly of rescued embryos. The phenotypes observed in prm1 mutant embryos are restored to normal in rescued organisms. Rescued embryos have a normal embryonic body wall muscle pattern (Fig. 3, AC). Each missing muscle fiber or group of fibers is restored and no duplicated or aberrant muscles are observed. The embryo body wall muscles of rescued embryos have a regular striation pattern of sarcomeres (Fig. 7).
As mentioned at the beginning of Results, the paramyosin transgene rescues the embryonic lethality of prm1. Homozygous prm1 survives to adulthood in the presence of the paramyosin transgene. Indirect flight muscles from rescued adult prm1 flies have normal myofibril structure (Fig. 9). In longitudinal sections, myofibrils have regular Z discs and M lines and sarcomere length is unchanged compared with wild type (Fig. 9, A and B). In transverse sections, thick and thin filaments are packed in a hexagonal manner and all the thick filaments have hollow centers (Fig. 9, CF). This suggests that paramyosin expressed from the transgene is correctly assembled into thick filaments and that this further restored the assembly of myofibrils during myofibrillogenesis.
|
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The role of paramyosin in myoblast fusion
The events surrounding myoblast fusion in D. melanogaster have been studied extensively (Paululat et al., 1999; Taylor, 2002). Fusion always takes place between founders and fusion-competent myoblasts. An individual founder fuses with fusion-competent myoblasts to form muscle precursors with two or three nuclei. These precursors enlarge by recruiting and fusing with additional fusion-competent myoblasts to form multinucleate myotubes.
The most apparent mesodermal defect in embryos mutant for the paramyosin gene is random loss of muscle fibers or muscle fibers in aberrant shapes (Fig. 3). However, the defect in muscle development is not caused by reduction in myoblast number in early development (Fig. 4). Unfused myoblasts are present in the locations of missing muscles before being degraded (Fig. 4 F), suggesting a role of paramyosin in the progression of cells from myoblasts to myotubes.
The structural features of paramyosin and its cytoplasmic location seem inconsistent with a role in cell adhesion, instead suggesting paramyosin functions as a cytoskeleton protein in early embryonic stages before myofibril formation. Paramyosin molecules form rodlike coiled-coil -helical dimers that lack domains reminiscent of cell adhesion molecules. Furthermore, paramyosin is localized beneath the cell membrane of mesodermal cells and epidermal cells (Fig. 6). Paramyosin might attach the cytoskeleton to membrane adhesion sites, akin to the role of coiled-coil intermediate filaments in other organisms. This function might arise due to the lack of cytoplasmic intermediate filament proteins in Drosophila (Goldstein and Gunawardena, 2000).
As is the case for paramyosin, there are several other cytoplasmic proteins that are important for Drosophila myoblast fusion (Paululat et al., 1999; Taylor, 2002). These include the myofibrillar protein D-titin (Zhang et al., 2000), Blown fuse (Doberstein et al., 1997), and Myoblast city (Rushton et al., 1995; Doberstein et al., 1997; Erickson et al., 1997), found in both founders and fusion-competent myoblasts, as well as Rolling pebbles (Rols)/Antisocial (Ants) present only in founders (Chen and Olson, 2001; Menon and Chia, 2001). It is suggested that Rols/Ants functions as an intracellular adaptor protein that relays signals from the immunoglobulin family membrane protein Dumbfounded (Ruiz-Gómez et al., 2000) to the cytoskeleton during myoblast fusion (Chen and Olson, 2001; Menon and Chia, 2001).
The presence of the myofibrillar protein D-titin in developing myoblasts implicates cytoskeletal components in the process of myoblast fusion (Zhang et al., 2000). Defects in myoblast fusion similar to those in prm1 occur in Drosophila D-titin mutants. Compared with wild-type muscles, embryo body wall muscles of D-titin mutants are smaller and thinner. Occasionally, a few missing muscles are observed in severe mutant alleles (Zhang et al., 2000). The motor protein nonmuscle myosin II is also important for muscle fiber formation in Drosophila embryos (Bloor and Kiehart, 2001). In zipper mutants of nonmuscle myosin II, some ventral muscles are deleted. The involvement of D-titin and possible involvement of nonmuscle myosin II in myoblast fusion support the hypothesis that contractile elements play a role in this process. Later, we discuss a model in which paramyosin serves as part of nonmuscle myosin minifilaments that interact with the actin cytoskeleton to regulate cortical cytoskeleton dynamics and promote myoblast fusion.
The role of paramyosin in thick filament formation and myofibril assembly
Accumulating evidence suggests that the assembly of muscle myosin into thick filaments requires the presence of other proteins (Barral and Epstein, 1999). Biochemical, cell biological, and genetic studies in invertebrates support this hypothesis. In D. melanogaster, paramyosin, miniparamyosin, myosin rod protein, and flightin are all present in the A band region of some muscle sarcomeres (Vigoreaux et al., 1993; Maroto et al., 1996; Standiford et al., 1997). Flightin knockout flies show increased thick filament length in indirect flight muscles, suggesting that flightin regulates thick filament assembly (Reedy et al., 2000). In C. elegans, the thick filament protein UNC-45 acts as a myosin chaperone (Barral et al., 2002) and unc-45 mutations disrupt thick filament assembly (Barral et al., 1998). C. elegans' thick filament cores contain paramyosin and three filagenins (Liu et al., 1998). Mutation of paramyosin in C. elegans greatly reduces the length of thick filaments and disrupts the distribution of myosin isoforms (Mackenzie and Epstein, 1980; Epstein et al., 1986).
Consistent with the C. elegans studies, we found that the body wall muscles of homozygous prm1 embryos have a marked reduction in the number of observable thick filaments, resulting in areas containing only thin filaments. Two factors might contribute to the formation of thick filaments in this line. The paramyosin from leaky expression of prm1 may interact with other unidentified thick filament core proteins, nucleating myosin molecules into these abnormal thick filaments. Alternatively, myosin may directly interact with these unidentified proteins to form abnormal thick filaments. We conclude that paramyosin is important for the production of an adequate number of morphologically normal thick filaments.
Another specific characteristic of prm1 is that the striated pattern of embryonic body wall muscles is disrupted. Although thin filaments assemble in the absence of paramyosin, they could not organize into regular sarcomeric patterns. The localization of paramyosin in thick filaments does not support a role as a scaffold in organizing myofibrils. Thus, abnormal interaction of thin and thick filaments might be the direct cause of this phenotype. Mutations in other Drosophila muscle contractile proteins also disrupt sarcomere organization, including actin (Sparrow et al., 1991), troponin-T (Fyrberg et al., 1990), troponin-I (Beall and Fyrberg, 1991), myosin heavy chain (O'Donnell and Bernstein, 1988; Beall et al., 1989), and -actinin (Roulier et al., 1992). This indicates that correct interaction between thin and thick filaments is required for myofibril formation.
A model of paramyosin function in Drosophila muscle differentiation
The formation of myofibrils has been extensively studied in embryonic vertebrate cardiac and skeletal muscle cells and several models have been proposed. In cultured cardiomyocytes, Sanger and colleagues (Dabiri et al., 1997) suggested that premyofibrils, characterized by banded patterns of -actininrich Z-bodies and nonmuscle myosin IIB, form at the edges of spreading cardiomyocytes and develop into mature myofibrils. During the transition from premyofibrils to myofibrils, there is an exchange of nonmuscle myosin IIB filaments for muscle myosin II filaments and a growth and fusion of Z-bodies into Z-bands. Evidence from developing chicken embryo hearts (Ehler et al., 1999) did not substantiate the presence of premyofibrils in vivo. An alternative model, based on observations of cultured cardiomyocytes, proposes that spatially separate complexes of actin filaments and Z-bands (IZI brushes) and groups of myosin thick filaments assemble independently of one another; they become spliced together by titin filaments, and then inserted at the ends of fully formed myofibrils (Ojima et al., 1999). Although these models differ in detailed steps of myofibril formation, they agree that myofibrillogenesis starts with membrane association and that thick and thin filaments form independently. According to these previous studies and our evidence of paramyosin's dual roles in myoblasts and myofibrils, we propose a model of paramyosin in Drosophila muscle differentiation.
The transition of paramyosin from a cytoplasmic location to myofibrils can be divided into several steps in our model. First, before myoblast fusion, paramyosin binds to nonmuscle myosin filaments. These paramyosin-nucleated nonmuscle myosin filaments interact with filamentous actin in the cytoskeleton of myoblasts to regulate cell shape change and membrane penetration during cell fusion. Adaptor proteins, such as Rols/Ants, help keep these paramyosin nonmuscle myosin filaments in discrete foci at this stage. Second, after myoblast fusion, nonmuscle myosin is replaced by muscle myosin. Paramyosin, together with other thick filament core proteins, nucleate muscle myosin filaments into muscle thick filament precursors. Third, actin filaments and titin of IZI brushes incorporate precursor thick filaments into A-bands of nascent myofibrils, as in cultured cardiomyocytes (Dabiri et al., 1997; Holtzer et al., 1997). Filaments may be first aligned out-of-register, and then transported into position. Finally, both thin and thick filaments elongate and myofibril diameter increases by peripheral addition of myofilaments.
Overall, our work revealed a role of paramyosin in thick filament formation and myofibrillogenesis. We also discovered that paramyosin has an unexpected function in myoblast fusion. Antibody localization and analysis of contractile protein mutants available in the Drosophila system will permit testing of our model of paramyosin function in muscle differentiation.
![]() |
Materials and methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Screening for late embryo motility defects
Embryos were collected for 1 h and aged to 22 h at 25°C, by which stage most wild-type embryos (including TM6C homozygotes) had hatched (Broadie and Bate, 1993). Unhatched embryos were dechorionated and observed for spontaneous peristaltic larval movement and tested for movement in response to touching by forceps. Mutant lines whose development appeared normal, but which were defective for larval movement, were rescreened to confirm the phenotype.
Genetic screen to isolate new paramyosin mutants
To isolate new alleles of the paramyosin gene by imprecise P element excision, individual male yw; prm106-5/TM2, Ubx Sb flies were mated to virgin female w; Sp/CyO; 23, Sb/TM2, Ubx flies, which produce transposase. The white+ gene in the P element was used as a marker for P element mobilization. Individual male F1 yw; prm106-5/
23, Sb flies with blotchy eyes were selected and then mated to yw; MKRS, Sb/TM3, y+ Ser virgin female flies. Sibling white-eyed F2 yw; prm106-5/TM3, y+ Ser flies from each cross were mated to each other to make a stable stock for each mutant line. Lines in which yw; prm106-5/prm106-5 flies could be recovered were probably progeny of flies with a precise P element excision and were discarded. Homozygous lethal lines likely contain new mutant alleles of the paramyosin gene and were kept for further analysis.
To identify paramyosin mutants, a paramyosin transgene P[w+, PM] (referred to as pm in the text) (Mardahl-Dumesnil, 1998) was crossed into these mutant backgrounds. Mutants that could be rescued to adulthood were considered to be paramyosin-specific mutants. Paramyosin-specific mutants, which are embryonic lethal, were used for paramyosin expression analysis. We used mouth hook color as a marker to differentiate homozygous mutant embryos from heterozygous embryos. Homozygous mutant embryos have yellow mouth hooks because they lack the balancer chromosome, which carries a black mouth hook marker (y+). Paramyosin expression levels were analyzed by Western blotting using antiparamyosin antibody (Maroto et al., 1995).
Protein electrophoresis and Western blotting
Protein electrophoresis of dechorinated embryos or dissected adult upper thoraces and Western blotting with antiparamyosin and antiminiparamyosin antibodies were performed as described by Maroto et al. (1995). Antibody detection was performed using the SuperSignal system (Pierce Chemical Co.). The relative amounts of proteins were determined by scanning densitometry of bands on films (Expression 636 scanner [Epson]; NIH Image 1.61).
DNA analysis
The insertion site of the P element in line prm1065 was determined by inverse PCR and cycle sequencing, essentially as described by the Berkeley Drosophila Genome Project (http://www.fruitfly.org/p_disrupt/inverse_pcr.html). To determine the extent of the deletion resulting from imprecise P element excision, genomic DNA was extracted from prm1 embryos. 100 ng of DNA were used as template for PCR with the Roche Expand Long Template PCR system (Roche Biosciences). Primers used were: forward primer, 5'GTTTTTAGTTCTCGGTTCTTTTTGTTGGAC3'; and reverse primer, 5'AAAAAGTTGCCGATTGCCACAAAGGCCACG3'. PCR products were cloned and sequenced.
Immunohistochemistry
Antibody staining of whole-mount embryos and of dissected embryos for confocal microscopy was performed as described (Bate, 1990; Rushton et al., 1995). Antibodies used are: antimuscle myosin and antinonmuscle myosin II antibodies (Kiehart and Feghali, 1986); antiparamyosin and antiminiparamyosin antibodies (Maroto et al., 1995); anti-DMEF2 (Lilly et al., 1995); mouse monoclonal antiß-galactosidase antibody (Promega); Cy5-labeled goat antirabbit Ig (Amersham Biosciences); and goat antimouse Ig Alexa Fluor 488 conjugate (Molecular Probes). FITC-phalloidin was purchased from Molecular Probes.
Electron microscopy
Electron microscopy of adult indirect flight muscles and embryos was done following previously published procedures (O'Donnell and Bernstein, 1988).
![]() |
Acknowledgments |
---|
This research was supported by an American Heart Association, Western States Affiliate predoctoral fellowship to H. Liu, grants MCB9604546 (National Science Foundation) and AR43396 (National Institutes of Health) to S.I. Bernstein, and grant 048476 (Wellcome Trust) to C.J. O'Kane.
Submitted: 30 August 2002
Revised: 3 February 2003
Accepted: 4 February 2003
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Arredondo, J.J., R.M. Ferreres, M. Maroto, R.M. Cripps, R. Marco, S.I. Bernstein, and M. Cervera. 2001. Control of Drosophila paramyosin/miniparamyosin gene expression. Differential regulatory mechanisms for muscle-specific transcription. J. Biol. Chem. 276:82788287.
Ayme-Southgate, A., J. Vigoreaux, G. Benian, and M.L. Pardue. 1991. Drosophila has a twitchin/titin-related gene that appears to encode projectin. Proc. Natl. Acad. Sci. USA. 88:79737977.[Abstract]
Barral, J.M., and H.F. Epstein. 1999. Protein machines and self assembly in muscle organization. Bioessays. 21:813823.[CrossRef][Medline]
Barral, J.M., C.C. Bauer, I. Ortiz, and H.F. Epstein. 1998. Unc-45 mutations in Caenorhabditis elegans implicate a CRO1/She4p-like domain in myosin assembly. J. Cell Biol. 143:12151225.
Barral, J.M., A.H. Hutagalung, A. Brinker, F.U. Hartl, and H.F. Epstein. 2002. Role of the myosin assembly protein UNC-45 as a molecular chaperone for myosin. Science. 295:669671.
Bate, M. 1990. The embryonic development of larval muscles in Drosophila. Development. 110:791804.[Abstract]
Bate, M. 1993. The mesoderm and its derivatives. The Development of Drosophila melanogaster. Vol. II. M. Bate and A. Martinez-Arias, editors. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. 10131090.
Beall, C.J., and E. Fyrberg. 1991. Muscle abnormalities in Drosophila melanogaster heldup mutants are caused by missing or aberrant troponin-I isoforms. J. Cell Biol. 114:941951.[Abstract]
Beall, C.J., M.A. Sepanski, and E.A. Fyrberg. 1989. Genetic dissection of Drosophila myofibril formation: effects of actin and myosin heavy chain null alleles. Genes Dev. 3:131140.[Abstract]
Becker, K.D., P.T. O'Donnell, J.M. Heitz, M. Vito, and S.I. Bernstein. 1992. Analysis of Drosophila paramyosin: identification of a novel isoform which is restricted to a subset of adult muscles. J. Cell Biol. 116:669681.[Abstract]
Bloor, J.W., and D.P. Kiehart. 2001. zipper nonmuscle myosin-II functions downstream of PS2 integrin in Drosophila myogenesis and is necessary for myofibril formation. Dev. Biol. 239:215228.[CrossRef][Medline]
Bour, B.A., M. Chakravarti, J.M. West, and S.M. Abmayr. 2000. Drosophila SNS, a member of the immunoglobulin superfamily that is essential for myoblast fusion. Genes Dev. 14:14981511.
Broadie, K.S., and M. Bate. 1993. Development of the embryonic neuromuscular synapse of Drosophila melanogaster. J. Neurosci. 13:144166.[Abstract]
Campos-Ortega, J.A., and V. Hartenstein. 1985. The embryonic development of Drosophila melanogaster. Springer-Verlag, Berlin. 9-84.
Chen, E.H., and E.N. Olson. 2001. Antisocial, an intracellular adaptor protein, is required for myoblast fusion in Drosophila. Dev. Cell. 1:705715.[Medline]
Cohen, C., and D.A. Parry. 1998. A conserved C-terminal assembly region in paramyosin and myosin rods. J. Struct. Biol. 122:180187.[CrossRef][Medline]
Dabiri, G.A., K.K. Turnacioglu, J.M. Sanger, and J.W. Sanger. 1997. Myofibrillogenesis visualized in living embryonic cardiomyocytes. Proc. Natl. Acad. Sci. USA. 94:94939498.
Deak, P., M.M. Omar, R.D. Saunders, M. Pal, O. Komonyi, J. Szidonya, P. Maroy, Y. Zhang, M. Ashburner, P. Benos, et al. 1997. P-element insertion alleles of essential genes on the third chromosome of Drosophila melanogaster: correlation of physical and cytogenetic maps in chromosomal region 86E-87F. Genetics. 147:16971722.
Doberstein, S.K., R.D. Fetter, A.Y. Mehta, and C.S. Goodman. 1997. Genetic analysis of myoblast fusion: blown fuse is required for progression beyond the prefusion complex. J. Cell Biol. 136:12491261.
Ehler, E., B.M. Rothen, S.P. Hammerle, M. Komiyama, and J.C. Perriard. 1999. Myofibrillogenesis in the developing chicken heart: assembly of Z-disk, M-line and the thick filaments. J. Cell Sci. 112:15291539.
Epstein, H.F., and D.A. Fischman. 1991. Molecular analysis of protein assembly in muscle development. Science. 251:10391044.[Medline]
Epstein, H.F., B.J. Aronow, and H.E. Harris. 1976. Myosin-paramyosin cofilaments: enzymatic interactions with F-actin. Proc. Natl. Acad. Sci. USA. 73:30153019.[Abstract]
Epstein, H.F., I. Ortiz, and L.A. Mackinnon. 1986. The alteration of myosin isoform compartmentation in specific mutants of Caenorhabditis elegans. J. Cell Biol. 103:985993.[Abstract]
Epstein, H.F., G.Y. Lu, P.R. Deitiker, I. Oritz, and M.F. Schmid. 1995. Preliminary three-dimensional model for nematode thick filament core. J. Struct. Biol. 115:163174.[CrossRef][Medline]
Erickson, M.R., B.J. Galletta, and S.M. Abmayr. 1997. Drosophila myoblast city encodes a conserved protein that is essential for myoblast fusion, dorsal closure, and cytoskeletal organization. J. Cell Biol. 138:589603.
Fyrberg, E., C.C. Fyrberg, C. Beall, and D.L. Saville. 1990. Drosophila melanogaster troponin-T mutations engender three distinct syndromes of myofibrillar abnormalities. J. Mol. Biol. 216:657675.[Medline]
Goldstein, L.S., and S. Gunawardena. 2000. Flying through the Drosophila cytoskeletal genome. J. Cell Biol. 150:F63F68.[Medline]
Hakeda, S., S. Endo, and K. Saigo. 2000. Requirements of Kettin, a giant muscle protein highly conserved in overall structure in evolution, for normal muscle function, viability, and flight activity of Drosophila. J. Cell Biol. 148:101114.
Holtzer, H., T. Hijikata, Z.X. Lin, Z.Q. Zhang, S. Holtzer, F. Protasi, C. Franzini-Armstrong, and H.L. Sweeney. 1997. Independent assembly of 1.6 microns long bipolar MHC filaments and I-Z-I bodies. Cell Struct. Funct. 22:8393.[Medline]
Huxley, H.T. 1963. Electron microscope studies on the structure of natural and synthetic protein filaments from striated muscle. J. Mol. Biol. 7:281308.
Kagawa, H., K. Gengyo, A.D. McLachlan, S. Brenner, and J. Karn. 1989. Paramyosin gene (unc-15) of Caenorhabditis elegans. Molecular cloning, nucleotide sequence and models for thick filament structure. J. Mol. Biol. 207:311333.[Medline]
Kiehart, D.P., and R. Feghali. 1986. Cytoplasmic myosin from Drosophila melanogaster. J. Cell Biol. 103:15171525.[Abstract]
Kulke, M., C. Neagoe, B. Kolmerer, A. Minajeva, H. Hinssen, B. Bullard, and W.A. Linke. 2001. Kettin, a major source of myofibrillar stiffness in Drosophila indirect flight muscle. J. Cell Biol. 154:10451057.
Lilly, B., B. Zhao, G. Ranganayakulu, B.M. Paterson, R.A. Schulz, and E.N. Olson. 1995. Requirement of MADS domain transcription factor D-MEF2 for muscle formation in Drosophila. Science. 267:688693.[Medline]
Liu, F., C.C. Bauer, I. Ortiz, R.G. Cook, M.F. Schmid, and H.F. Epstein. 1998. ß-Filagenin, a newly identified protein coassembling with myosin and paramyosin in Caenorhabditis elegans. J. Cell Biol. 140:347353.
Liu, F., I. Ortiz, A. Hutagalung, C.C. Bauer, R.G. Cook, and H.F. Epstein. 2000. Differential assembly of - and
-filagenins into thick filaments in Caenorhabditis elegans. J. Cell Sci. 113:40014012.
Machado, C., and D.J. Andrew. 2000. D-Titin: a giant protein with dual roles in chromosomes and muscles. J. Cell Biol. 151:639652.
Mackenzie, J.M., Jr., and H.F. Epstein. 1980. Paramyosin is necessary for determination of nematode thick filament length in vivo. Cell. 22:747755.[CrossRef][Medline]
Mardahl-Dumesnil, M. 1998. The role of myosin heavy chain, enolase, and paramyosin in muscle assembly and function in Drosophila melanogaster. Ph.D. thesis. San Diego State University, San Diego. 167 pp.
Maroto, M., J.J. Arredondo, M. San Roman, R. Marco, and M. Cervera. 1995. Analysis of the paramyosin/miniparamyosin gene. Miniparamyosin is an independently transcribed, distinct paramyosin isoform, widely distributed in invertebrates. J. Biol. Chem. 270:43754382.
Maroto, M., J. Arredondo, D. Goulding, R. Marco, B. Bullard, and M. Cervera. 1996. Drosophila paramyosin/miniparamyosin gene products show a large diversity in quantity, localization, and isoform pattern: a possible role in muscle maturation and function. J. Cell Biol. 134:8192.[Abstract]
McLachlan, A.D., and J. Karn. 1982. Periodic charge distributions in the myosin rod amino acid sequence match cross-bridge spacings in muscle. Nature. 299:226231.[CrossRef][Medline]
Menon, S.D., and W. Chia. 2001. Drosophila rolling pebbles: a multidomain protein required for myoblast fusion that recruits D-Titin in response to the myoblast attractant Dumbfounded. Dev. Cell. 1:691703.[Medline]
Müller, S.A., M. Häner, I. Ortiz, U. Aebi, and H.F. Epstein. 2001. STEM analysis of Caenorhabditis elegans muscle thick filaments: evidence for microdifferentiated substructures. J. Mol. Biol. 305:10351044.[CrossRef][Medline]
Nose, A., T. Isshiki, and M. Takeichi. 1998. Regional specification of muscle progenitors in Drosophila: the role of the msh homeobox gene. Development. 125:215223.
O'Donnell, P.T., and S.I. Bernstein. 1988. Molecular and ultrastructural defects in a Drosophila myosin heavy chain mutant: differential effects on muscle function produced by similar thick filament abnormalities. J. Cell Biol. 107:26012612.[Abstract]
Obinata, T. 1993. Contractile proteins and myofibrillogenesis. Int. Rev. Cytol. 143:153189.[Medline]
Ojima, K., Z.X. Lin, Z.Q. Zhang, T. Hijikata, S. Holtzer, S. Labeit, H.L. Sweeney, and H. Holtzer. 1999. Initiation and maturation of I-Z-I bodies in the growth tips of transfected myotubes. J. Cell Sci. 112:41014112.
Paululat, A., A. Holz, and R. Renkawitz-Pohl. 1999. Essential genes for myoblast fusion in Drosophila embryogenesis. Mech. Dev. 83:1726.[CrossRef][Medline]
Reedy, M.C., B. Bullard, and J.O. Vigoreaux. 2000. Flightin is essential for thick filament assembly and sarcomere stability in Drosophila flight muscles. J. Cell Biol. 151:14831500.
Roulier, E.M., C. Fyrberg, and E. Fyrberg. 1992. Perturbations of Drosophila -actinin cause muscle paralysis, weakness, and atrophy but do not confer obvious nonmuscle phenotypes. J. Cell Biol. 116:911922.[Abstract]
Ruiz-Gómez, M., N. Coutts, A. Price, M.V. Taylor, and M. Bate. 2000. Drosophila dumbfounded: a myoblast attractant essential for fusion. Cell. 102:189198.[Medline]
Rushton, E., R. Drysdale, S.M. Abmayr, A.M. Michelson, and M. Bate. 1995. Mutations in a novel gene, myoblast city, provide evidence in support of the founder cell hypothesis for Drosophila muscle development. Development. 121:19791988.
Sparrow, J., M. Reedy, E. Ball, V. Kyrtatas, J. Molloy, J. Durston, E. Hennessey, and D. White. 1991. Functional and ultrastructural effects of a missense mutation in the indirect flight muscle-specific actin gene of Drosophila melanogaster. J. Mol. Biol. 222:963982.[CrossRef][Medline]
Standiford, D.M., M.B. Davis, K. Miedema, C. Franzini-Armstrong, and C.P. Emerson, Jr. 1997. Myosin rod protein: a novel thick filament component of Drosophila muscle. J. Mol. Biol. 265:4055.[CrossRef][Medline]
Taylor, M.V. 2002. Muscle differentiation: how two cells become one. Curr. Biol. 12:R224R228.[CrossRef][Medline]
Vigoreaux, J.O., J.D. Saide, K. Valgeirsdottir, and M.L. Pardue. 1993. Flightin, a novel myofibrillar protein of Drosophila stretch-activated muscles. J. Cell Biol. 121:587598.[Abstract]
Vinós, J., A. Domingo, R. Marco, and M. Cervera. 1991. Identification and characterization of Drosophila melanogaster paramyosin. J. Mol. Biol. 220:687700.[Medline]
Zhang, Y., D. Featherstone, W. Davis, E. Rushton, and K. Broadie. 2000. Drosophila D-titin is required for myoblast fusion and skeletal muscle striation. J. Cell Sci. 113:31033115.
Ziegler, C., K. Jurk, B. Weitkamp, B. Kölsch, and B. Beinbrech. 1996. In vitro interactions of proteins from insect myosin filaments. Biophysics. 41:7987.
Related Article