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Neuroscience Program, Department of Biological Sciences, Ohio University, Athens, Ohio
ABSTRACT I. INTRODUCTION A. Why Study Invertebrate Muscles, Genes, and Proteins? B. Invertebrate and Vertebrate Phylogeny C. Scope of Review and Literature Database II. REVIEW OF VERTEBRATE MUSCLE SPECIFIC PROTEINS III. INVERTEBRATE MUSCLE PROTEINS A. Thin Filament Proteins 1. Actin 2. Tropomyosin 3. Troponin 4. Calponin/caldesmon 5. C. elegans unc-87 B. Thick Filament Proteins 1. Myosin heavy chain 2. Catchin/myorod 3. Myosin light chains 4. Myosin light-chain phosphorylation 5. Paramyosin/miniparamyosin 6. Filagenins 7. C. elegans unc-45 8. Myonin 9. Flightin 10. Zeelin C. {alpha}-Actinin and Other Z Line Proteins D. Ca2+ Binding Proteins and Their Targets E. Giant Sarcomere-Associated Proteins 1. Crayfish connectin/kettin, C. elegans kettin, and Drosophila titin/kettin 2. C. elegans titin 3. Unidentified very large sarcomere proteins 4. Projectin/twitchin 5. C. elegans unc-89 F. Miscellaneous Other Proteins 1. Nebulin 2. Nesprin 3. Amphiphysin 4. Various muscle cell attachment proteins 5. Dystrophin-related proteins 6. Intermediate filaments 7. The 29-kDa ascidian protein G. Summary ACKNOWLEDGMENTS REFERENCES
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I. INTRODUCTION |
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Although pure research has its defenders, science is generally justified by perceived human benefit. Two arguments suggest that studying invertebrate muscle genes and proteins can reveal generally applicable principles that could benefit humans. First, the last common ancestor of vertebrates and invertebrates had muscle, and most vertebrate muscle genes and proteins have invertebrate homologs. The experimental advantages of invertebrate preparations often allow these genes and proteins to be investigated more easily, or at a greater level of detail, than is possible in vertebrates. Many human diseases result from errors in muscle protein structure, and thus invertebrate studies have the possibility of improving human health. Second, invertebrate muscle genes and proteins show great variation. Despite this variety, in all cases the proteins must functionally interact correctly. Comparative studies therefore provide a rich arena in which to investigate the relationship between protein assembly structure and activity, again an area with clear relevance to human well-being.
B. Invertebrate and Vertebrate Phylogeny
Figures 1–3 show a contemporary, molecular biology-based, tree of life (4 and sources listed in the legend to Fig. 1). Three things are of particular importance. First, Cnidaria (corals, jellyfish) are separate and equal to Bilatera. All Bilatera are thus equally distant from all Cnidaria. Second (as has been long known), echinoderms, tunicates, and amphioxus are more closely related to vertebrates than they are to other invertebrates. Third, Ecdysozoa, Lophotrochozoa, and Deuterostomia, each of which contains invertebrates, are presently coequal branches. This tripartite split will presumably be eventually resolved into two bipartite branchings, but whether the Ecdysozoa and Lophotrochozoa, or one of them and the Deuterostomia, will end up being most closely related is as yet unclear. This issue has profound implications for comparative research since, depending on the ultimate resolution of bilaterian relationships, it may be that lobsters are more closely related to humans than they are to leeches. Although research on muscle genes may help resolve this issue, it is so far insufficient to do so. We have therefore organized the data presented here in simple concordance with the relationships shown in Figures 1–3.
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A comprehensive review of invertebrate muscle is unavailable. However, our invertebrate muscle database contains over 6,700 references, and this massive literature cannot be covered in a single review. We therefore intend to produce a series of canonic reviews covering all journal articles (due to their limited availability, books and book chapters are not included) on invertebrate muscle written before 2005. To that end, the field has been divided into subsets, of which this review covers the first. Subsequent reviews will cover 1) thick filament, thin filament, and sarcomere structure; the molecular basis of contraction and its regulation; asynchronous muscle and catch; 2) muscle and synaptic ultrastructure and excitation/contraction coupling; 3) voltage- and ligand-gated ionotropic channels; 4) metabotropic channels (modulation); and 5) integrative properties and production of behavior. Even with this broad net, some boundaries had to be drawn. In particular, papers dealing with metabolic pathways are generally not included, and no attempt to cover molting and muscle proteases, regeneration, or muscle development has been made (papers that identify regulatory regions in muscle protein genes are included, but papers further "upstream" are not). The database is available at http://crab-lab.zool.ohiou.edu/invert. Every effort will be made to maintain this site for at least 10 years from publication date.
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II. REVIEW OF VERTEBRATE MUSCLE SPECIFIC PROTEINS |
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-helical coiled-coil tail. The other end of each heavy chain and one essential and one regulatory chain form one of the combined molecule's two heads, each of which contains an ATPase activity and can independently bind to the thin (actin) filament. Trypsin severs the myosin tail, resulting in light meromyosin, which contains only tail (rod) sequences and heavy meromyosin (HMM), which contains part of the tail and the two head regions (30, 42, 525, 1135, 1138). Further digestion of HMM results in the S1 and S2 fragments, S1 consisting of the heads and S2 of the HMM tail portion. Thin filaments are a double helix of polymerized actin monomers. Tropomyosin and troponin are two thin filament-associated proteins involved in contraction regulation in striated muscle (muscles with well-organized sarcomeres, Fig. 5). The other type of vertebrate muscle, smooth muscle, does not have well-organized sarcomeres. Vertebrate smooth muscle contraction is regulated both by myosin light-chain phosphorylation by myosin light-chain kinase and a thin filament-based regulatory system based on the actin-binding proteins calponin and caldesmon.
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-actinin, define the sarcomere edges. The thin filaments attach to each Z line and extend toward the center of the sarcomere. The region adjacent to each Z line containing only thin filaments is the I band. One-half of each I band therefore belongs to one sarcomere and the other half to the adjacent sarcomere. The thick filaments are located at the center of the sarcomere. The region of the sarcomere with only thick filaments is the H band, and the region defined by their extent is the A band. At the very center of the sarcomere there is often also a line (due to the presence of additional proteins) called the M line. Sarcomeres consisting of only thick and thin filaments and Z lines would be inherently unstable. To appreciate this, consider a muscle fiber stimulated to contract while maintained in an isometric (constant length) condition. If the thick filaments were exactly centered in the sarcomere, equal numbers of myosin heads would engage the thin filaments on the two sides of the M line, the thick filament would feel equal force in both the right and left directions, and the thick filaments would therefore remain centered in the sarcomere. However, if a thick filament was even slightly uncentered, it would experience greater force in one direction and would therefore slide in that direction. The force imbalance on the thick filament would now be even greater, and it would thus continue to slide until it reached the Z line. Solving this difficulty requires a mechanism that develops a centering force if the thick filaments become uncentered. For example, if the thick filaments were attached to the Z lines by springs, any thick filament movement away from the center would decrease force in the shortened spring, and increase force in the stretched spring, which would recenter the thick filament.
Electron microscopic evidence of filaments linking the thick filaments to the Z line (and which could thus function as springs) was early obtained in both vertebrates and invertebrates (781). However, these filaments are not composed of polymerized smaller subunits but are instead enormous single proteins, and it took almost 20 years to characterize them chemically. Intriguingly, many of them contain large numbers of immunoglobulin and fibronectin III repeats. The largest of these proteins is titin (3 MDa, 
30,000 amino acids, Fig. 5). Titin can be 1 µm in length, and single titin molecules connect the M and Z lines. Titin's NH2 terminus extends through the Z line in close association with the thin filament. At the A/I junction it leaves the thin filament and joins the thick filament, with which it runs until reaching the M line, where it overlaps the titin filaments from the other half-sarcomere. The M line-associated proteins M-protein, myomesin, and skelemin (which is generated from the myomesin gene by alternative splicing) are other members of this family, as are the A band myosin binding proteins C and H (also called C-protein and 86-kDa protein) (Fig. 5). These proteins are much smaller than titin (a few hundred kiloDaltons), and unlike titin lack a serine/threonine kinase activity. Myosin light-chain kinase and telokin are two other members of this protein family, found in smooth muscle.
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III. INVERTEBRATE MUSCLE PROTEINS |
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For a recent general review of sarcomere structure and proteins (primarily vertebrate, but includes some invertebrate work), see Reference 182; Reference 933 reviews all aspects of both vertebrate and invertebrate muscle. Reference 1136 reviews the early history of muscle protein isolation and analysis. Reference 188 is a dated, primarily vertebrate, review of calcium binding proteins, troponin C, and myosin light chains. References 112, 276, 330–332, 451, 1223; 17, 276, 1261; and 437 review, respectively, Drosophila, Caenorhabditis elegans, and amphioxus muscle. References 236, 237, 440–443, 571, 815, 869 and 43, 378, 673, 987, 1266, 1282, respectively, describe phenotypic mutant derivations in Drosophila and C. elegans. Reference 333 reviews basic methods in Drosophila muscle biology. Gene expression profiles that included muscle specific proteins have been performed in jellyfish (Cyanea capillata) tentacle (1307); oyster (Crassostrea gigas) mantle (799); Mytilus galloprovincialis (1210); the platyhelminths Clonorchis sinensis (652) and Schistosoma japonicum (291, 1214, 1215); the nematodes Brugia malayi (118, 119), C. elegans (719, 769, 774, 798, 1259), Globodera species (951), Haemonchus contortus (436), Meloidogyne incognita (773), Onchocerca volvulus (697), and Strongyloides stercoralis (798) [for nematodes, multiple expressed sequence tag databases are now available on-line (926, 927, 1293)]; the mites Psoroptes ovis (555) and Boophilus microplus (220); and amphioxus notochord (1124) and Ciona intestinalis embryo (1048, 1049), larva (632, 1048), and adult (173, 1048), and cDNA clones covering nearly 85% of C. intestinalis mRNA species are available (1050). References 1060 and 281, 814, 1159 show two-dimensional electrophoresis profiles of, respectively, C. elegans and Drosophila proteins. References 814 and 1159 show that anatomically different Drosophila muscles (fibrillar vs. tubular) have different protein compositions. Also uncategorized are papers describing muscle post mortem changes, food-related properties, and calorimetric measurements of muscle proteins (this list is not comprehensive) (18, 19, 65, 170, 267, 269, 308, 367–372, 462, 463, 467, 493, 518–520, 542, 543, 546, 565, 588, 589, 620, 687, 701, 753, 764, 765, 770, 807, 808, 839, 840, 856, 865, 878, 899, 900, 907, 908, 910–923, 1043, 1044, 1068, 1093, 1191, 1192, 1199, 1200, 1300, 1313–1316).
Actin and myosin heavy chain have been extensively studied, and these sections are therefore subordered by phylogenetic group. For some groups not all the articles associated with it are explicitly covered in the text (e.g., descriptions of isolation techniques). For completeness, in these cases all references dealing with that group are listed immediately after the relevant subtitle. References about purification of myosin as an oligomer (i.e., heavy and light chains together) are listed in this manner in the myosin heavy chain section. Table 1 provides actin and myosin heavy chain data for groups for which only limited information is available (for actin, Pterobranchia, Annelida, Gastropoda, Brachiopoda, Chaetognatha, Chelicerata; for myosin heavy chain, Cephalochordata, Urochordata, Annelida, Chaetognatha, Chelicerata).
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Mammals have six (two striated muscle, two smooth muscle, and two cytoplasmic) (1203) and teleost fish have nine (1211) actin isoforms. References 303 and 418 are general (vertebrate and invertebrate) reviews of actin molecular genetics, References 557 and 1025 review vertebrate and invertebrate actin isoforms, and Reference 499 reviews ascidian actin. Reference 238 shows that muscle actins isolated from a variety of invertebrates all have the same molecular weight and coelectrofocus with the 
-form of vertebrate smooth muscle actin, but are immunologically distinct from each another.
A) CNIDARIA. Two coral actin cDNA clones have been identified, one of which is expressed only in adults, the only stage with muscles. Although not verified by in situ hybridization, cnidaria may thus have a muscle specific actin. The putative muscle actin gene showed greatest homology to metazoan cytoplasmic actins (which metazoa are not clear from the article) (321). However, invertebrate muscle actins are typically most similar to vertebrate cytoplasmic actins (see below), and thus, depending on which metazoans the authors used for comparison, this similarity is not strong evidence that the adult actin gene is not a muscle gene. Hydra has three or more transcribed genes, but whether any are muscle specific is unknown (304).
B) CEPHALOCHORDATA. Cephalochordate (Branchiostoma only) actin gene number and expression are confusing. Work using antibodies specific for vertebrate smooth muscle, striated muscle, and cytoplasmic actins shows that B. lanceolatum has at least three actin isoforms, with the antismooth and antistriated antibodies staining separate sets of muscles and the anticytoplasmic antibody staining most other cells (note that Ref. 1204 is wrong in stating cephalochordates express only one muscle actin isoform). Despite early confusion on this point (373, 946, 1234), in these animals the notochord is an innervated muscle (310, 311, 511, 831, 1124, 1273). Interestingly, this tissue nonetheless stains with the anticytoplasmic antibody (285, 1096).
Cloning in B. lanceolatum identified a cytoplasmic and a muscle actin on the basis of "diagnostic" amino acids (see below) (127). However, no tests for tissue specific expression to verify these assignments were made. Cloning in B. floridae identified one cytoplasmic and two muscle actins on the basis of diagnostic amino acids (623). In situ hybridization showed that staining for one muscle actin was less strong in gill slit muscle and that the cytoplasmic form was weakly expressed in axial muscles in embryos, which suggests muscle and stage specific variation in actin gene expression. Cytoplasmic actin was present in notochord during early development, but none of the three forms was present in adult notochord. Cloning in B. belcheri identified only one cytoplasmic and one muscle actin gene (on the basis of diagnostic amino acids alone), but Southern blot analysis suggested the presence of "a number" of additional actin genes (624).
Expressed sequence tag analysis of a B. belcheri notochord cDNA library suggests that notochord has three actin genes, each of which codes for an identical actin that differs from the cytoplasmic and muscle clones already identified in this species (1124). The notochord, muscle, and cytoplasmic actin amino acid sequences are identical in the regions used to construct the oligonucleotide primers used to clone the muscle and cytoplasmic actins. It is thus unclear why the notochord genes were not identified in the earlier work (one possibility being a difference in codon usage, although codon variation is small for all amino acids in question, two of the seven amino acids in each primer have unique codons, and for the other five, only third nucleotide use varies). PCR analysis of notochord, muscle, and ovary libraries shows that one notochord actin gene is expressed only in notochord but some of the others are also expressed in muscle (1124). The most conservative interpretation of these data is that in Cephalochordata there are three actin groups (cytoplasmic, muscle, and notochord), each of which may contain more than one actin gene, but the number of these genes, and which tissues each is expressed in, is not clear.
C) UROCHORDATA. Urochordate (87, 173, 176, 428, 429, 500, 605, 623, 625–632, 858, 875, 1047, 1049, 1096, 1126, 1181–1183, 1204, 1280, 1281) actin genes have been studied almost exclusively in Phlebobranchia (Ciona intestinalis, Ascidia ceradotes) and Stolidobranchia (Styela clava, S. plicata, Halocynthia roretzi, Molgula oculata, M. occulta). Ciona has eight nonmuscle and six muscle actin genes (classified by diagnostic amino acid position, not verified by in situ hybridization) (note Ref. 1204 is wrong in stating urochordates express only one muscle actin). Three muscle actin genes encode identical proteins (173). All six muscle actins have amino acid positions diagnostic of vertebrate striated muscle actin. Ciona thus appears to have no counterpart to vertebrate smooth muscle actin, a conclusion supported by the failure of antismooth muscle antibodies to stain Ascidia muscles (1096). Gene expression profiles show that muscle actin is expressed in Ciona embryos (1049), larvae (632), and adults (173). Five muscle actins are expressed only in embryos and larvae, while the sixth is expressed solely in adults (173). In neither embryos nor larvae was an actin gene showing notochord specific expression mentioned (632, 1049).
Two-dimensional gel electrophoresis indicates that S. clava embryos, tadpoles, and adults contain three major and two minor actin isoforms. Two of the major isoforms are likely cytoplasmic and the third a muscle actin (1181). Cloning indicates four to seven muscle actin genes (according to diagnostic amino acid position). The four well-described genes encode identical proteins, but show different temporal and spatial expression patterns. In particular, one is expressed in a wide variety of tissues (including nonmuscle), but only in larva and younger animals, whereas another is expressed primarily in muscle cells in embryos but in nonmuscle tissue in adults. None of the genes has been shown to be expressed at high levels in adult muscle, which may suggest an adult-specific actin gene remains to be found. None of the clones stained the notochord (87). A muscle actin gene (by diagnostic amino acid position) that shows some amino acid variation (5.6%) from the S. clava muscle actins (and which might thus be the "missing" adult muscle actin gene) has been isolated from an adult S. plicata muscle cDNA library (605). In embryo and larva, this gene is expressed only in muscle cell lineages or functional muscle cells. Although the clone's origin shows the gene is expressed in adult muscle, whether it is expressed only in muscle is unknown (1182).
H. roretzi adult body wall muscle contains two actins differing in isoelectric point (875). The gene(s) coding for these actins are not identified, but differ from those coding for larval muscle actin (625, 626, 628–630, 1047). H. roretzi has seven larval muscle [verified by in situ hybridization (625, 630)] actin genes. The genes are arranged in two clusters, one of which has five tandemly repeated genes in the same orientation and the other of which has two genes, arranged head to head on opposite DNA strands, that share a common interposed promoter (626, 628, 629). Two of the actins are identical, and all are very similar. Genes in each cluster have similar regulatory elements and are believed to be coordinately controlled (626, 628, 629, 1047). The sequences responsible for this regulation are beginning to be identified (428). When reporter genes with a H. roretzi 5'-upstream muscle actin gene (from the 5 gene cluster) flanking region are introduced into C. savignyi, reporter gene function is observed only in larval muscle cells, suggesting that larval muscle actin regulatory processes have been conserved in the two groups (429).
Two works in Molgula species show that changing actin gene expression can affect morphological development. The first (627, 631) involves M. oculata, which has typical tailed larvae, and M. occulta, which has tailless larvae. M. oculata has two actin muscle genes (classified by diagnostic amino acids, but verified in the larva by in situ hybridization), one expressed in larva and the other in adults. M. occulta has two actin genes orthologous to the M. oculata larva gene, but in situ hybridization with M. oculata-derived probes do not detect any muscle actin production in M. occulta. However, when M. occulta gene 5'-flanking regions are attached to a reporter gene, reporter gene product is observed in M. occulta embryo vestigial muscle cells. Thus both actin gene promoter functionality and proper spatial production of the trans-acting factors that activate the genes are present in M. occulta. The coding regions of the M. occulta genes, however, contain insertions, deletions, and codon substitutions that would result in their producing nonfunctional actin. M. occulta taillessness thus appears to be due, at least in part, not to changes muscle actin gene activation, but to changes in gene coding regions such that the activated genes produce no functional actin.
The second work involves precocious development in M. citrina larvae (1126). In most Molgula species, mesenchyme cells, believed to be adult muscle progenitors, remain undifferentiated in larvae, but in M. citrina they begin to differentiate during the larval stage. A M. citrina actin gene has been identified that is expressed in juveniles and adults (it is not known if it is exclusively expressed in muscle at these stages), but not in larva tail muscle, suggesting that it codes for an adult muscle actin. In situ hybridization shows that the gene is expressed in the precociously differentiating mesenchyme cells in larva and (at least) early after larval metamorphosis. Precocious development of adult features in this species is thus likely also associated with precocious expression of an adult muscle actin gene.
Larvacea muscle (classification confirmed by in situ hybridization) actin genes have been investigated only in Oikopleura longicauda (858). This work identified one actin gene expressed in larval and adult tail muscle (larvaceans retain their tails as adults), but not in adult heart. Undiscovered muscle actin genes thus presumably exist in this species.
D) ECHINODERMATA. Actin genes have been studied in echinoids (Stronglyocentrotus purpuratus, S. franciscanus, Lytechinus pictus, Heliocidadris erythrogramma, H. tuberculata) and asteroids (Pisaster ochraceus, Dermasterias imbricata) (199, 215, 218, 258, 265, 292, 293, 342, 508, 513, 568, 569, 602–604, 606, 650, 651, 909, 948, 1055, 1062, 1063, 1085, 1204, 1280, 1281). Echinoids have 6–10 cytoplasmic actins, but all possess only one muscle actin gene [early studies suggesting as many as 20 actin genes in Stronglyocentrotus and Lytechinus species apparently being mistaken (265, 508)] (215, 218, 265, 292, 293, 342, 568, 569, 650, 651, 909, 1055, 1085). In S. purpuratus, the muscle actin gene is expressed at high levels only in postpluteus muscle (1085). Dermasterias has eight actin genes, but which are cytoplasmic and which muscle is unknown (603). Pisaster has five actin genes, one believed to be cytoplasmic, two muscle specific, and two unspecified (identifications on the basis of cDNA library source tissue, not verified by in situ hybridization) (602–604, 606).
E) MOLLUSCA. For bivalves, see References 153, 230, 296, 556, 558, 658, 738–740, 859, 930, 1000, 1125, 1137, 1204, 1210, 1281, 1309; for gastropods, see Refs. 246, 366, 401, 643, 826, 1280, 1281; for cephalopods, see Refs. 161, 448, 595, 1106, 1204. Although one of the first techniques to isolate large quantities of invertebrate thin filaments was developed in bivalves (1137), and regulation of bivalve actomyosin has been extensively studied (see second review), relatively little is known about bivalve actin genes. Early electrophoresis work showed no difference in actin across a wide range of bivalve species (740) or between different tissues within one examined species (Spisula) (739). Actin cDNA clones have been isolated from scallop (Placopecten magellanicus) (930) and oyster (Crassostrea gigas) (153). The scallop actin gene is likely a muscle actin and appears to be the primary actin expressed in adductor muscle. Southern blot analysis suggests 12–15 actin genes in the genome. Scallop actin polymerizes more slowly than rabbit actin, and once polymerized, the cleft between actin subdomains 2 and 4 is larger in scallop than in rabbit actin (556, 558). The oyster cDNA was used to locate the gene and its upstream region, but nothing is known about temporal or tissue expression. Actin gene polymorphisms can be used to identify clam species muscle (to prevent fraudulent use of less desirable species in consumer products) (296) and to study interspecies hybridization (230). Bivalve actin levels change in characteristic ways in response to various pollutants (1000).
The information available about gastropod muscle actin is presented in Table 1. Cephalopods are only slightly better investigated. Southern blot analysis suggests that coleoid (all cephalopods but Nautilus) have at least three actin loci. Phylogenetic analysis suggests three actin isoforms, two of which were identified as muscle or cytoplasmic on the basis of being, respectively, "related to the mollusk muscle type" and "clustered among the other mollusk cytoplasmic" actins (161). However, almost all these sequences were obtained from GenBank, not from published work showing tissue specificity. As such, although the sequence comparisons showing three actin isoforms are likely valid, the identification of the isoforms as to type needs experimental confirmation. A capillary sodium dodecyl sulfate gel electrophoresis technique for actin separation in squid has been developed (1106).
F) PLATYHELMINTHS. Cestoda is represented by Taenia solium (15, 155), Diphyllobothrium dendriticum (1237–1241), and Echinococcus granulosus (229). Taenia has seven actin isoforms (15). Two genes, with identical coding sequences, have been cloned (155). Diphyllobothrium has five actin genes (1239, 1241), three muscle specific (1237). Two actin genes have been isolated from Echinococcus, and Southern blotting indicates as many as eight (229). Trematoda is represented by Schistosoma mansoni (1, 234, 706, 901, 1342) and Fasciola hepatica (1119). Two-dimensional gel electrophoresis identifies seven Schistosoma actin isoforms, and two actin genes have been cloned. Fasciola has three actin isoforms, one of which is specific to tegumental spines. Turbellaria is represented by Dugesia lugubris, which has at least two major actin isoforms, one muscle specific (928, 929).
G) NEMATODA. All information comes from Caenorhabditis elegans and Onchocerca volvulus (300, 608, 609, 646, 718, 902, 1052, 1204, 1235, 1264, 1280, 1281). In C. elegans four actin genes have been identified, two of which produce identical proteins and none which differs by more than three amino acids (300, 609). Three of the genes are clustered (10, 300). All four genes are transcribed, and three acquire a 22-nucleotide leader sequence via RNA trans-splicing (608). Genetic evidence suggests that two of the genes are involved in muscle thin filaments (646, 1264). However, the actins are so similar that they migrate as a single species by isoelectric focusing (1052), no isoform-specific antibodies have been generated, and thus other evidence of tissue specific isoform expression is lacking. One reference (902) states that all four genes code for muscle specific actins (and refers to a personal communication about a putative cytoplasmic actin gene), but on what basis is unclear. Expression of total actin varies during development (718). C. elegans actin can be efficiently extracted at high purity (902). Some biochemical differences between C. elegans and rabbit actin have been identified (902), and the structure of C. elegans actin resolved at 1.75 
resolution (1235). There is suggestive evidence that C. elegans actin folding may be chaperoned (667). O. volvulus has four actin genes that can be divided into two classes on the basis of EcoR1 digestions. The genes are arranged in two clusters, each of which contains one copy of each class (1328). Nothing is known about tissue or stage specific expression.
H) CRUSTACEA. Crustacean (202, 526, 720, 783, 826, 1206, 1280, 1281) actin genes have been relatively little studied. Artemia (species unreported) has 8–10 genes and 4 isoforms, one muscle specific (709, 905). cDNA clones have been isolated from the shrimp Marsupenaeus japonicus and crayfish Procumbarus clarkii, but nothing is known about their expression (526, 720). Crab (Gecarcinus lateralis) has seven or eight actin genes, and immunocytochemistry suggests some tissue and stage specific expression (1206). In lobster (Homarus americanus), different muscles contain different amounts of total actin (783).
I) INSECTA. The references for insecta are as follows: Diptera: Drosophila (16, 20, 64, 77, 89, 92, 111, 115, 129, 130, 204, 222, 259–262, 334, 336, 339, 340, 350, 419, 430, 431, 449, 512, 536, 540, 634, 647, 658, 702, 703, 722, 733, 826, 829, 868, 897, 925, 974, 1038, 1039, 1041, 1057, 1108, 1109, 1175, 1176, 1280, 1281, 1347), Aedes aegypti (466), Mayetiola destructor (1086), Dacus dorsalis (also known as Bactrocera dorsalis) (411), Phormia regina (591); Lepidoptera: Bombyx (826–829); Coleoptera: Heliocopris japetus (139); Heteroptera: Lethocerus cordofanus (139, 146, 658, 1037). D. melanogaster has six actin genes that are widely dispersed throughout the genome and produce three major mRNA size classes (339). The genes are similarly dispersed in other Drosophila species (702). Gene coding sequences, but not intron positions, are highly conserved (334). Four (cytogenetic positions 57B, 79B, 87E, and 88F) of these genes code muscle actin (64, 111, 340, 430, 733, 826, 1041, 1175), at least some of whose expression is muscle or developmental stage specific (64, 77, 111, 204, 340, 430, 536, 634, 868, 1041, 1175, 1176). In particular, Act88F is expressed primarily in indirect flight muscles (64, 340, 430, 722) [although it is coexpressed with other muscle actin genes, and its absence causes behavioral defects, in a small number of other muscles (868)], Act79B is primarily expressed in "tubular" muscles (an anatomically specific muscle type, see third review) (64, 204, 340, 883) [an early report (1347) that Act79B is the larval muscle actin apparently being in error], and Act57B and Act87E are expressed in embryonic and larval muscle (340, 1175) and a variety of adult nontubular, nonindirect flight muscles (340). Similar data are obtained from D. virilis (703) and, for Act88F, in D. simulans (92), except that in D. virilis gene coexpression occurs in more muscles (although this difference may stem from enhanced sensitivity of modern techniques). Regulatory regions of the Act57B, Act79B, and Act88F genes have been identified in D. melanogaster (204, 350, 431), and the Act88F gene promoter has been used to drive green fluorescent protein expression (11).
The different actins are functionally nonequivalent (336), and (although they differ by only 15 amino acids) mammalian cytoplasmic actin cannot substitute for Act88F (129). Drosophila indirect flight muscle actin requires posttranslational modification for normal polymerization (419, 722, 1057) (which presumably underlies the indirect flight muscle "actin III" reported in Ref. 449; see also Refs. 430, 647) and is the only known actin to have an unacetylated, free NH2 terminus (1057).
Multiple mutants of Drosophila indirect flight muscle actin that alter muscle force production, despite in some cases assembling into seemingly normal thin filaments, have been obtained (16, 20, 222, 260–262, 430, 815, 897, 974, 1038, 1108, 1109), as have mutants that disrupt myofibril structure (16, 222, 260, 536, 540, 722, 1109). In a study comparing several Act88F mutants, in almost all cases protein stability was similar (261). In one mutant in which muscle force generation is altered but thin filament structure appears normal, the mutation site is distant from the myosin binding site, and actin mutations can therefore have long-range effects on force generation (262). Experiments measuring the effect of actin mutation on profilin, ATP, and DNase I binding showed similar distant effects for some mutants (259). Other mutants identified Glu93 as part of a secondary myosin binding site (974), electrostatic charge on actin domain two as critical for thin filament regulation by tropomyosin (115), and the binding site of Clostridridium toxins (512). Mutants that produce no actin still produce relatively normal thick filament arrays, and experiments in which the actin-to-myosin ratio is altered suggest that filament imbalances, not lack of thin filaments per se, are responsible for the observed defects (89). Act88F epitope tagging on the COOH terminus results in flightlessness and disordered indirect flight muscle sarcomeres, but NH2 terminus tagging gives relatively normal sarcomeres and partially restores flight ability (130). Many [but not all (430, 540)] actin mutations induce heat shock protein production (260, 430, 431, 897, 925). The molecular basis of this induction is unknown but is independent of myofibril degeneration (430, 540, 1038, 1039).
Asynchronous muscle (a special type of muscle in which muscle contractions are not synchronized with motor neuron activity, see second review) in all species examined in Nepomorpha, and some species in Diptera, contain a ubiquitinated actin, arthrin (64, 93, 138, 1058). Arthrin and the tropomyosin/troponin complex are in equimolar concentration, suggesting that they may colocalize on the thin filament. This suggestion has not been verified, but it is known that the ubiquination site is on the opposite side of the thin filament from where tropomyosin binds (341). Arthrin's function is unknown, as arthrin activates myosin ATPase, is regulated by troponin/tropomyosin in the same manner as actin, and its presence does not alter actomyosin kinetics (138, 1058). Arthrin is not required for asynchronous muscle, as mutant Drosophila in which actin ubiquination cannot occur still fly (1058), and arthrin has not been found in any Hymenoptera species with asynchronous muscle, and is absent from some Cicadellidae (a family of Cicadelloidea), Diptera, Coleoptera, and Heteroptera species, even though all animals in these groups have asynchronous muscle (Fig. 3) (937, 1058). Arthrin evolved independently at least twice, although in all cases the ubiquination site (Lys-118) is identical (148, 1058).
Actin genes from three other Diptera (A. aegypti, M. destructor, D. dorsalis) believed to code for muscle actin (on the basis of nucleotide and amino acid similarity to D. melanogaster genes) have been investigated (411, 466, 832, 1086, 1236). Southern blot analysis suggests Aedes contains at least five actin-related sequences (466) with differential expression in different muscles (832, 1236). Four muscle genes (identity confirmed by hybridization of gene specific clones to RNA extracted from different tissues) have been cloned in D. dorsalis (411). Developmental stage specific expression and differential expression in different muscles are present. The 3'- and 5'-flanking regions of these genes show very little sequence homology both among themselves and to other known actin genes. In Lepidoptera (B. mori) three actin genes have been identified, one cytoplasmic and two muscle, one of which is expressed only in adult muscle and the other in both larval and adult muscles (827, 828).
J) ACTIN AS A PHYLOGENETIC CHARACTER. Much of the above work investigates phylogenetic relationships (92, 127, 161, 218, 285, 292, 300, 334, 366, 500, 568, 605, 623, 624, 626, 627, 826, 829, 858, 903, 1085, 1204, 1239, 1269). Until recently, the generally accepted conclusion was that all invertebrate muscle actins are most closely related to vertebrate cytoplasmic actins, with insect muscle actins diverging very early from those of other invertebrates. Unfortunately, much of this work may be seriously flawed. First, much of it is based on diagnostic amino acids (623), a small number of differing amino acids that were early and successfully used to classify mammalian actins, and then (uncritically) applied to invertebrate actins. This approach has been strongly criticized because it 1) treats the other amino acids as carrying no phylogenetic information and 2) does not analyze gene evolution as a dynamic process of "descent with modification" (i.e., it is insufficient to just compare differences between two extant species; instead, enough information from multiple species must be obtained to infer the history of changes that resulted in the present differences) (1281).
Second, actin gene duplications and conversions (including between muscle and cytoplasmic forms), which complicate phylogenetic comparisons, have occurred in many lineages (218, 258, 624, 1280). Third, Drosophila actin genes show pronounced codon bias, which could bias phylogeny construction (410). Fourth, actin may not be particularly suitable as a phylogenetic marker. In most organisms actin is a relatively large proportion of total protein. Growth rate changes might therefore disproportionately affect actin evolution, and even closely related invertebrate species can grow at very different rates. Indeed, the actin gene duplications noted above may have arisen specifically to allow rapid growth in some lineages. Furthermore, different actin isoforms presumably at least partly reflect a need for muscles with different functional properties, a need that in many cases may depend on organism life-style. Closely related invertebrate species with different life-styles could thus have experienced higher rates of actin evolution, again complicating phylogenetic analysis.
Fifth, actin-based phylogenies often have untenable relationships (Fig. 6). For instance, in phylogeny A, sea urchin and ascidia cytoplasmic actin are more closely related to mollusk cytoplasmic actin than to pufferfish cytoplasmic actin, even though sea urchin, ascidia, and pufferfish belong to Deuterostoma and mollusks to Lophotrochozoa (Fig. 1). Similarly, in phylogeny B, sea urchin cytoplasmic actin is more closely related to Cnidaria actin than to either ascidia or sea star cytoplasmic actin, even though Cnidaria belongs to Radiata and the other species to Bilateria. Again, in phylogeny C, sea star cytoplasmic actin is more closely related to urochordate cytoplasmic actin than it is to that of another echinoderm, sea urchin.
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Vertebrate muscle tropomyosin is a dimer of tropomyosin molecules arranged in an in-parallel, in-register coiled-coil. There are typically two isoforms, and tropomyosin can form as a hetero- or homodimer of either. Nonmuscle tropomyosin isoforms are also widely present. Multiple tropomyosin isoforms are generally present in invertebrates, and most work has concentrated on distinguishing muscle and nonmuscle forms. Whether invertebrate tropomyosin is a dimer, and if so, a hetero- or homodimer, is less investigated. However, equilibrium sedimentation work in crayfish, oyster, abalone, and blowfly indicate that the tropomyosins self-associate (1291), and optical rotary dispersion and hydrodynamic measurements suggest blowfly tropomyosin is 100% -helical and in a two-stranded configuration (591). Given these data, invertebrate tropomyosin is presumably also a coiled-coil dimer. Muscle-type tropomyosin is in some cases expressed in nonmuscle tissues, and in some work below these tissues were used as tropomyosin source material. Reference 653 is a general (vertebrate and invertebrate) review covering muscle and nonmuscle tropomyosins. In the older literature, paramyosin is sometimes called tropomyosin A or even tropomyosin (for references, see sect. IIIB5 and second review); in this work tropomyosin is called tropomyosin B. In two very early articles, we are unable to determine, for the invertebrate portions, whether tropomyosin or paramyosin was being studied (938, 1174).
Tropomyosin has been identified by expressed sequence tagging, immunohistochemistry, protein isolation, or cloning in Cnidaria (53, 320, 385, 386, 699); amphioxus (1124); Ascidia (785, 1183); sea urchin (485, 486, 490–492, 1177); annelids (221, 658); mollusks: bivalves (179, 404, 469, 472, 481, 482, 484, 488, 494, 524, 658, 672, 738–740, 795, 931, 1152, 1254, 1291), gastropods (401, 480, 1291), cephalopods (448, 477, 483, 595, 806, 1197); brachiopods (658); Chaetognatha (1011); platyhelminths (283, 706, 1277, 1295); C. elegans (21, 515–517, 718) and other nematodes (39, 318, 388, 461, 501, 842); Chelicerata: mite (1021, 1285), horseshoe crabs (658, 659, 802, 804, 805), scorpion (805); Crustacea: lobster (202, 487, 670, 790, 803, 805, 837, 1042), shrimp (233, 668, 803, 805, 960, 981, 982, 1073, 1285), crayfish (103, 801, 803, 805, 1291), crab (669, 803, 805), hermit crab (803, 805), isopod (803, 805); and various insects (41, 84–86, 139, 146, 394, 395, 537–539, 591, 614, 658, 805, 1291).
Sea anemone has five isoforms (320) of unknown origin, but some tissue specificity is observed (320). Jellyfish has two tropomyosin genes (Ref. 53 is mistaken in stating only a single gene exists). One is expressed only in striated muscle (385). One tropomyosin gene has been found in Hydra (699), but is not expressed in epitheliomuscular cells. Unidentified epitheliomuscular cell specific genes may thus exist. Expressed sequence tag analysis shows that tropomyosin is expressed in Amphioxus notochord (1124). Striated and smooth muscle in ascidia has an identical tropomyosin, coded by a single gene and more similar to vertebrate striated than vertebrate smooth muscle tropomyosin. This common expression may be due to ascidian smooth muscle being troponin regulated (785). The isoform is expressed only at low levels in larval tail muscle, so undiscovered tropomyosin genes may exist. Sea urchin has at least two isoforms, antigenically identified as muscle or nonmuscle types (492). The muscle type binds actin (1177) and is located in muscle (486). Earthworm muscle possesses two isoforms of unknown origin that combine as a heterodimer (221). Bivalves and gastropods have as many as six isoforms (401, 404, 482, 489, 740), some expressed in a muscle specific manner (488, 489, 494, 931), and multiple tropomyosin genes (404, 494, 931). NH2-terminal blocking is important for head to tail polymerization, actin binding, and Ca2+ regulation in Akazara scallop tropomyosin (473). Bivalve tropomyosin levels change in characteristic ways in response to various pollutants (1000).
Platyhelminths have multiple isoforms, with some tissue-specific expression and likely multiple genes (283, 1277). C. elegans has one tropomyosin gene. Four isoforms arise by alternative splicing and show developmental stage (718) specific expression and are differentially expressed in different muscles (21, 515, 516). Limulus has four to six isoforms that are not expressed in a tissue-specific manner (804). Scorpion has four muscle isoforms that show some muscle specificity (805). Crustaceans have three or four muscle isoforms. One is uniquely expressed in cardiac muscle, and the others show considerable but not perfect muscle specific expression (487, 801, 803, 805, 837, 960, 1042). Two of the isoforms may arise by alternative splicing (837). Beetles and centipedes have three muscle tropomyosins, which show some muscle specificity (805). Locust has multiple isoforms with muscle specific expression. A cDNA clone has been sequenced, and two mRNAs are present, but whether they are from different genes is unknown (614). Drosophila has two muscle tropomyosin genes. Each produces four or five tissue or developmentally specific isoforms (including nonmuscle forms) by a combination of alternative splicing and multiple promoters (84–86, 381, 394, 395, 537–539, 1064) (two of these isoforms are the flight muscle specific troponin H, see below). Considerable evidence has been obtained in Drosophila on the regions of the tropomyosin gene that regulates its expression (686, 689, 690, 788, 1064).
Drosophila (538, 613, 817, 1164) and C. elegans (21, 516, 1282) tropomyosin mutations alter muscle structure and function. Some Drosophila mutants can be rescued by reproviding correct tropomyosin sequences (337, 1163, 1164). Despite the partial tissue specificity noted above, in one case substitution of different isoforms can also rescue tropomyosin mutants (794).
Tropomyosin is an important component of immune and allergic reactions (7, 25, 29, 33, 37, 38, 40, 41, 50, 51, 150, 157, 158, 179, 233, 255, 295, 362, 403, 479–484, 501–506, 661–663, 668–672, 726, 752, 806, 970, 978–983, 1032, 1045, 1073, 1166, 1172, 1173, 1277, 1278, 1285, 1295, 1296, 1312).
Vertebrate troponin contains three subunits: troponins T, I, and C. Ascidia (273, 882, 1183); Glycera, Lumbricus, and Nereis (annelids) (658); scallop (363, 861, 885, 888–891) and squid (477, 595, 892, 1081, 1193); nematode (461, 564); Limulus (658, 659); shrimp (895, 960), lobster (202, 790, 852, 860, 985, 1081), and crayfish (103); and Drosophila (139, 143) and Lethocerus (658) troponin also contains three subunits (although with considerable molecular weight variation). Troponin T is present in cross and obliquely striated Eisenia (earthworm) muscle (1012, 1019), Helix heart (1012), mite (1021), and Antarctic mussel shrimp (1016); troponin C in amphioxus (1149), sandworm (Perinereis) (1326), mussel (1081), and barnacle (12, 35, 192); troponin I in amphioxus (1124); and "troponin" (unidentified subunit) in Chaetognatha (Sagitta frederici) (1011). A fourth troponin, troponin H (called a tropomyosin isoform, not troponin H, by some authors), which is a fusion product of tropomyosin and a hydrophobic proline-rich sequence, is alternatively spliced from one of the tropomyosin genes, at least partly replaces troponin I, and exists in all indirect (asynchronous) and some direct (synchronous) flight muscles (143, 395, 537, 539, 937, 977). In mollusks, high (relative to vertebrates, but physiological for the organisms in question) monovalent ion or Mg2+ levels are required for troponin and tropomyosin to remain bound to the thin filament (363, 655). Lack of recognition of this need presumably is the reason that early studies found troponin was not present (552, 658), or was present at only low levels (660), in molluscan muscle.
Lobster actin:tropomyosin:troponin (with the troponin consisting of equimolar amounts of troponin T, I, and C) (790), and scallop actin:tropomyosin:[troponin I:troponin C], are present in molar ratios of 7:1:1 (657), as are Limulus actin:tropomyosin:[troponin T:troponin C], but in this species twice the amount of troponin I may be present (659). Shrimp troponin C, I, and T are present in a 1:1:1 molar ratio (895). Scallop troponin I and C are associated with the I band (657). Troponin is absent from surf clam foot muscle (175). Reference 656 compares the amino acid composition of troponin C in rabbit and a variety of invertebrates.
Cloning work has been performed for ascidia (184, 714, 1321, 1323), scallop (893, 1158), tick (1317), crayfish (575), and Drosophila (68, 88) troponin I; Amphioxus (1149), ascidia (1144, 1322), sandworm (1326), scallop (863, 886, 887, 894, 1121, 1325), squid (892), nematode (1168), Limulus (573), lobster (343), crayfish (576), barnacle (192), fire ant (945), and Drosophila (328, 423) troponin C; ascidia (272), C. elegans (833), Drosophila (101, 143, 335), and scallop (470, 471) troponin T; and Drosophila troponin H (395, 537, 539). Troponin-based phylogenies agree with Figures 1–3 (184, 187, 471, 1149, 1320). Multiple isoforms, some developmental stage or muscle specific, are present in ascidia (271, 272, 714, 882, 1321, 1323); scallop (888, 1325); C. elegans (843); shrimp, crayfish, and lobster (202, 343, 581, 783, 790, 834–836, 838, 852, 860, 895, 960, 985, 1083, 1286); barnacle (35, 192); Lethocerus (959), Anopheles (959), dragonfly (305, 735–737); and Drosophila (68, 69, 101, 257, 328, 335, 395, 423, 538, 539, 959). Stage-specific isoform changes are associated with increased Ca2+ sensitivity and altered twitch contraction kinetics in aging dragonflies (305, 736), and expression of the asynchronous muscle isoform with flight ability in bee (257).
Halocynthia (ascidia) has one adult (which produces two isoforms by alternative splicing) and at least three larval troponin I genes; sequences regulating gene expression have been identified (1321, 1323). Ciona, alternatively, has only one troponin I gene (which again produces two isoforms), and no homologs to the Halocynthia larval genes (714, 1324). Only one troponin I gene has been identified in Drosophila (68, 69, 88, 244); sequences regulating gene expression have been identified (741, 760). Ascidia, amphioxus, sandworm, and scallop have one troponin C gene (1319, 1322, 1325, 1326), C. elegans and barnacle two (192, 1168), lobster three (343), and Drosophila five (328, 423). The fourth intron of invertebrate troponin C genes shows great variability, suggesting that the ancestral gene may not have possessed it (1325, 1326). Ascidia has two troponin T genes (271, 272) and Drosophila and dragonfly one (101, 735). Troponin I, C, T, or H mutants can disrupt muscle function or development (68, 69, 88, 335, 516, 538, 771, 833, 866, 1168), although some polymorphism is tolerated (224). Drosophila troponin I mutants can be suppressed by tropomyosin (841) or myosin heavy chain (299, 615, 866, 867) mutations or troponin I second site mutations (952).
Calponin-like proteins have been identified in Eisenia (1017); Mytilus (325); Helix (1012); Echinococcus [called myophilin (748–751)] and Schistosoma (510, 1306); and Onchocerca (where it is highly immunogenic) (475), but not in crustacean or Drosophila muscle (1012, 1017). Calponin-like cDNAs in a nematode (Meloidogyne incognita) (162), Echinococcus (751), and Schistosoma (1306) have been sequenced. Echinococcus has only one gene, but multiple isoforms are expressed due to posttranslational modification, including phosphorylation by protein kinase C (750). In some Schistosoma species, several copies are present, and multiple isoforms are expressed (1306). Caldesmon-like proteins have been identified in Eisenia (1017) and mollusks (76, 100, 225, 1012).
The unc-87 mutants have almost normal embryonic muscles that become severely disorganized as the animals mature (1266). unc-87 codes for a 40-kDa protein that is located in the I band, bundles actin filaments (but not monomers) in the absence of tropomyosin or
