Invertebrate Muscles: Muscle Specific Genes and

Transcription

Physiol Rev 85: 1001–1060, 2005;
doi:10.1152/physrev.00019.2004.
Invertebrate Muscles: Muscle Specific Genes and Proteins
SCOTT L. HOOPER AND JEFFREY B. THUMA
Neuroscience Program, Department of Biological Sciences, Ohio University, Athens, Ohio
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
B. Thick filament proteins
C. ␣-Actinin and other Z line proteins
D. Ca2⫹ binding proteins and their targets
E. Giant sarcomere-associated proteins
F. Miscellaneous other proteins
G. Summary
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Hooper, Scott L., and Jeffrey B. Thuma. Invertebrate Muscles: Muscle Specific Genes and Proteins. Physiol Rev
85: 1001–1060, 2005; doi:10.1152/physrev.00019.2004.—This is the first of a projected series of canonic reviews
covering all invertebrate muscle literature prior to 2005 and covers muscle genes and proteins except those involved
in excitation-contraction coupling (e.g., the ryanodine receptor) and those forming ligand- and voltage-dependent
channels. Two themes are of primary importance. The first is the evolutionary antiquity of muscle proteins. Actin,
myosin, and tropomyosin (at least, the presence of other muscle proteins in these organisms has not been examined)
exist in muscle-like cells in Radiata, and almost all muscle proteins are present across Bilateria, implying that the
first Bilaterian had a complete, or near-complete, complement of present-day muscle proteins. The second is the
extraordinary diversity of protein isoforms and genetic mechanisms for producing them. This rich diversity suggests
that studying invertebrate muscle proteins and genes can be usefully applied to resolve phylogenetic relationships
and to understand protein assembly coevolution. Fully achieving these goals, however, will require examination of
a much broader range of species than has been heretofore performed.
I. INTRODUCTION
A. Why Study Invertebrate Muscles, Genes,
and Proteins?
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
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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
0031-9333/05 $18.00 Copyright © 2005 the American Physiological Society
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FIG. 1. Phylogeny of Animalia (except Ecdysozoa, see Figs. 2 and 3). This phylogeny and those in Figs. 2 and 3 are from Tree of Life
(http://tolweb.org/tree/phylogeny.html; Coordinator, David Maddison, Univ. of Arizona), the University of California Museum of Paleontology
Phylogeny exhibit (http://www.ucmp.berkeley.edu/exhibit/phylogeny.html), and Systema Naturae 2000 [SJ Brands (comp.) 1989 –2002. Systema
Naturae 2000. Amsterdam, The Netherlands; http://sn2000.taxonomy.nl/]. In all three figures, gray lines represent unresolved relationships, and
differing horizontal line lengths (e.g., Platyzoa) are simply for figure composition, and in particular do not indicate evolutionary distance.
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.
C. Scope of Review and Literature Database
A comprehensive review of invertebrate muscle is
unavailable. However, our invertebrate muscle database
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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
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INVERTEBRATE MUSCLE GENES AND PROTEINS
FIG.
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2. Phylogeny of Ecdysozoa (except Insecta, see Fig. 3).
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.
II. REVIEW OF VERTEBRATE MUSCLE
SPECIFIC PROTEINS
Due to the number of references in this review, this
section is only sparsely referenced; References 6 and 128
are excellent reviews of vertebrate muscle. All muscles
contain thick and thin filaments. Thick filaments are composed of myosin. Myosin is composed of three pairs of
proteins: the heavy chain and the essential and regulatory
light chains (Fig. 4). The tails of the heavy chains form an
␣-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
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(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 wellorganized 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.
All vertebrate striated muscle sarcomeres are very
similar (Fig. 5). Two Z lines, composed largely of ␣-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
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FIG. 3. Phylogeny of Insecta. Insects in red have asynchronous muscles, insects in green do not, and insects in black have not been examined
for this characteristic.
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
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FIG. 4. Myosin is composed of three paired molecules: the heavy
chain and the essential and regulatory light chains. Part of the heavy
chains form a coiled coil tail; the remainder of the heavy chains and the
two light chains form two globular heads. HMM, heavy meromyosin; S1,
S2, two HMM subfragments; LMM, light meromyosin (see text). [Modified from Rayment and Holden (972).]
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FIG. 5. Thick and thin filament arrangement in vertebrate striated muscle
sarcomere. Also shown are the sarcomere associated proteins ␣-actinin, titin,
myomesin, M-protein, and myosin binding proteins C and H. Red, actin; blue,
myosin; yellow, ␣-actinin; triangle, titin
serine/threonine protein kinase activity;
ellipse and circle in M line, myomesin
and M-protein, respectively.
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 ␮m 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.
III. INVERTEBRATE MUSCLE PROTEINS
We have attempted to order this review hierarchically. However, reviews, papers describing large numbers
of mutants with a common phenotype (e.g., flightlessness), and treatments describing the global advantages of
particular organisms do not nicely fit into this hierarchy
because they cover a large number of topics. Rather than
cite them repeatedly below, we instead summarize these
papers here.
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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,
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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).
A. Thin Filament Proteins
1. Actin
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
TABLE
1.
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
Actin and myosin information for relatively little studied groups
Group/Protein
Pterobranchia actin
Annelida actin
Gastropod actin
Species
Muscle Versus
Cytoplasmic
Isoforms?
Identification Method
Muscle or
Developmental Stage
Specific Expression?
Reference Nos.
Saccoglossus kowalevskii
Glycera, Nereis, species
unreported
Helobdella triserialis
Two cDNAs
Protein purification
Unknown
Unknown
Unknown
Unknown
127
658
One cDNA, apparent
family
Unknown
1269
Aplysia californica
Aplysia californica
One cDNA, 3–5 gene
family
One cDNA
Cloning of 5⬘-flanking
and part of coding
region of one gene
Many tissues
including
muscle, but not
segmental
ganglia
Unknown
cDNA clone
muscle specific
Unknown
Unknown
Unknown
Unknown
401
246
Unknown
Unknown
643
366
Unknown
Unknown
658
Immunohistochemistry
Three genes
Unknown
Two genes muscle
specific
1011
1308
Protein purification or
immunohistochemistry
Expressed sequence tag
Unknown
Unknown
Differentially
expressed in
different muscles
Unknown
Unknown
Unknown
Zebra mussel
Haliotis rufescens
Lophophorata, Brachiopoda
actin
Chaetognatha actin
Glottidea, unreported
species
Sagitta friderici
Paraspadella gotoi
Chelicerata actin
Various
Cephalochordata myosin
Amphioxus (notochord)
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247, 658,
1021, 1204
1124
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 proPhysiol Rev • VOL
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teins (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 ob-
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served 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 transacting 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
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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
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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 Å 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 latePhysiol Rev • VOL
1009
ralis) 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,
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1010
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-tomyosin 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
Physiol Rev • VOL
(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
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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.
Muscle actin is part of a highly organized ensemble of
proteins that might be expected to coevolve. A more
fruitful approach therefore may be to build phylogenies
1011
for multiple muscle proteins (e.g., actin, troponin, myosin
light and heavy chains) and superimpose the trees to
arrive at the ensemble’s most likely phylogeny. When this
is done (Fig. 6D), an ancestral set of muscle proteins gives
rise to two branches, one of which leads to the vertebrate
smooth muscle ensemble and the arthropod and vertebrate cytoplasmic ensembles. The other branch gives rise
to urochordate smooth muscle and all striated muscles.
Note that this phylogeny agrees with those determined by
other methods (Figs. 1–3); in particular, in each branch all
vertebrates are more closely related to each other than
they are to nonvertebrates, and urochordates are more
closely related to vertebrates than they are to arthropods.
2. Tropomyosin
Vertebrate muscle tropomyosin is a dimer of tropomyosin molecules arranged in an in-parallel, in-register
coiled-coil. There are typically two isoforms, and tropo-
FIG. 6. Various molecular geneticbased phylogenies. A–C are based on actin alone. D is based on superimposition
of troponin C, myosin essential and regulatory light chains, myosin heavy chain,
actin, and (for the vertebrates) muscle
regulatory factor trees. [A modified from
Carlini et al. (161); B modified from
White and Crother (1280); C modified
from White and Crother (1281); D modified from Oota and Saitou (903).]
Physiol Rev • VOL
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1012
myosin 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
Physiol Rev • VOL
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–
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506, 661– 663, 668 – 672, 726, 752, 806, 970, 978 –983, 1032,
1045, 1073, 1166, 1172, 1173, 1277, 1278, 1285, 1295, 1296,
1312).
3. Troponin
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)
Physiol Rev • VOL
1013
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).
4. Calponin/caldesmon
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
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1014
identified in Eisenia (1017) and mollusks (76, 100, 225,
1012).
5. C. elegans unc-87
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 ␣-actinin,
and has the same actin binding sequence as calponin (361,
607). When unc-87 mutants are expressed in animals with
decreased myosin heavy chain activity, the age-dependent
disorganization is less, suggesting that it arises from contraction force. unc-87 is therefore not believed to be
required for sarcomere formation, but instead to serve a
structural role (360).
B. Thick Filament Proteins
1. Myosin heavy chain
There are at least 13 classes of myosin; all muscle
(and some nonmuscle) myosins belong to class II (61, 104,
172, 200, 374, 433, 1069, 1070, 1198, 1302). We cover here
only muscle myosins. References 26, 66, 400, 776, and
1134 review vertebrate and invertebrate muscle and nonmuscle myosins, and Reference 42 reviews vertebrate
(primarily) and invertebrate domain structure. Reference
275 reviews myosin in C. elegans. We also do not review
here how myosin aggregates to form the thick filament, as
these data are in the second review, but do note that
individual scallop (and presumably other invertebrate)
combined myosin heavy and light chain molecules have
the structure shown in Figure 4 (i.e., possess a long rod
and have two globular heads separated by a cleft and
connected to the rod by flexible necks) (1222).
References 172, 200, 374, and 433 examine myosin
heavy chain motor domain phylogenetic relationships.
These works are primarily concerned with the relationships among the different myosin classes, and in particular contain muscle myosins from too few invertebrate
species to shed light on invertebrate phylogenetic relationships. The available data suggest two important (from
the point of view of this review) myosin II branchings.
First, vertebrate smooth muscle myosin and some invertebrate nonmuscle (but still class II) myosins early split
from the myosins leading to vertebrate striated muscle
and all invertebrate muscle myosins. A second split resulted in two branches, one of which contains all striated
(both skeletal and cardiac) vertebrate and invertebrate
myosins and the other of which contains nematode muscle myosin (which has a special type of striation called
obliquely striated, see third review). However, given the
paucity of invertebrate species contained in this work and
Physiol Rev • VOL
in particular the lack of smooth invertebrate muscles in it,
these groupings must be considered preliminary.
A) CNIDARIA. A cDNA clone of a striated muscle specific
myosin heavy chain has been sequenced and is more
similar to striated muscle heavy chain isoforms than to
smooth or nonmuscle isoforms in either vertebrates or
invertebrates. Because cnidarians also have smooth muscle, at least one other muscle isoform is presumably
present. Southern blotting suggests this/these isoform(s)
would be from other genes, not alternative splicing (5,
1061).
B) MOLLUSCA. For bivalves, see References 67, 163–165,
405, 452, 453, 496, 567, 594, 597, 599, 622, 738, 740, 862,
864, 873, 874, 941, 1101, 1246); gastropods, References 32,
52; cephalopods, References 563, 580, 768. Myosin heavy
chain cDNAs or genes have been sequenced in scallops
Argopecten (also known as Aequipecten) (873, 874), Patinopecten (405), Pecten (496), and Placopectin (941) and
squid Loligo (768). Scallop (Placopectin, Argopectin) has
three or four isoforms alternatively spliced from a single
gene and expressed in a muscle-specific fashion (874,
941). Mollusk striated and catch (a special muscle type
that maintains contraction with very little energy expenditure, see second review) muscle myosin heavy chains
differ almost exclusively in only one region (surface loop
1). The two isoforms have different ATPase activities
(941) and ADP affinities and dissociation rates (622). The
sequence responsible for light-chain positioning on the
heavy chain has been identified in the scallop (452). Regulatory light-chain kinase phosphorylates scallop myosin
heavy chain (1101), and Mytilus myosin contains a tightly
bound endogenous kinase (163). Which residues were
phosphorylated in these works is unknown, but phosphorylation of specific serines in Mytilus tailpiece increases myosin solubility and favors molecular folding
(164, 165). Fish light chains bind to scallop heavy chain
(567, 1101). Squid has at least two isoforms, alternatively
spliced from one gene (768).
C) PLATYHELMINTHS. See References 14, 387, 572, 904,
1028, and 1276 for platyhelminth data. Planaria have two
muscle myosin heavy chain genes expressed in nonoverlapping sets of muscle (572, 904). Although all planarian
muscles are nonstriated, planaria heavy chain myosins
most closely resemble the striated muscle myosin heavy
chains of other organisms (572). Phylogenetic analysis
using myosin heavy chain genes suggests that the platyhelminths are polyphyletic and that two platyhelminth
groups, the Acoela and Nemertodermatida, are the earliest extant bilaterians (1028). Myosin may be useful in
platyhelminth vaccine development (13, 108, 114, 241,
254, 268, 282, 509, 682, 849, 935, 1103–1105, 1338 –1340).
D) NEMATODA. See References 24, 94, 250, 251, 275, 278,
280, 302, 402, 445– 447, 497, 541, 693, 711, 715–717, 792,
793, 812, 844, 855, 898, 956, 1053, 1161, 1260, 1265, 1274,
1275, 1279, and 1329 for nematode data. C. elegans has
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four myosin heavy chain isoforms (MHC A, B, C, D),
coded by widely separated genes (myo-3, unc 54, myo-2,
myo-1, respectively), some on different chromosomes
(10, 251, 280, 541, 715, 717, 793, 1053). MHC A and B are
used only in body wall muscle and MCH C and D only in
pharyngeal muscle (24, 280, 711). The molecular basis of
this differential expression is beginning to be investigated
(497, 898). MHC A is present only in the center of the thick
filament and MHC B only at the ends (792), possibly
because MHC A’s more hydrophobic rod surface binds
better to paramyosin (which forms the core of nematode
thick filaments, see second review) (447). Depending on
the mutation, MHC B mutants (280, 716, 717) are paralyzed with a reduced number of thick filaments composed
entirely of MCH A (94, 250, 278) or very slowly moving
with normally assembled thick filaments (250, 812). MHC
B overexpression disrupts muscle structure (302). MHC A
mutations are embryonic lethal and contain no thick filaments, presumably because MHC A is necessary for the
initial steps of thick filament formation (1260). Two regions of the MHC rod sufficient to allow thick filament
initiation have been identified (445, 447), as has a region
for interaction with paramyosin (446). The terminal, nonhelical tailpiece of MHC A is not required for thick filament formation, although the thick filaments are disorganized relative to control animals (445). MHC B, but not
MHC A, synthesis and accumulation is reduced in mutants
lacking paramyosin (1279), and MHC A overexpression
can partially compensate for MHC B and paramyosin
mutations (754). Myosin heavy chain B pre-RNAs have
been used to examine intron splicing in C. elegans (881).
Partial or complete myosin heavy chain cDNAs have been
sequenced in Schistosoma mansoni (855), Toxocara canis (877), and Onchocerca volvulus (1275), and a myosin
heavy chain gene has been sequenced in Brugia malayi
(1274). The Onchocerca and Brugia data are for body wall
(putative MHC B homolog) myosins. Myosin may be useful in nematode vaccine development (877, 961, 962).
E) CRUSTACEA. See References 55, 202, 203, 439, 570, 638,
684, 721, 783, 834, 1016, 1040, 1090 –1092, and 1346 for
crustacean data. Lobster has at least two myosin heavy
chain isoforms (684, 783), and crayfish three to four (638,
1040). Heavy chain isoform expression differs in fast versus slow muscles in crayfish (638, 1040) and lobster (203,
684, 783) (although Ref. 834 reported no difference) when
the muscles are examined in bulk, but individual fibers
show a continuum of myosin heavy chain isoforms (782).
ATPase activity differences between slow and fast muscles are due solely to heavy chain isoform differences
(684). Lobster abdominal myosin is more susceptible to
tryptic proteolysis and is cleaved at different locations
(also unlike vertebrate myosin, in a Ca2⫹ concentrationindependent manner), than is rabbit myosin (734, 1090,
1092, 1346). A specific myosin exists in the Antarctic
isopod Glyptonotus, presumably to maintain muscle funcPhysiol Rev • VOL
1015
tion at very low temperature (439), a conclusion supported by evidence that lobster does not express temperature-dependent myosin heavy chain isoforms (unlike, for
instance, fish) (721).
F) INSECTA. See References 89, 109 –111, 139, 180, 185,
223, 235, 347, 406, 424, 434, 435, 550, 591, 616 – 618, 692,
791, 816, 823, 879, 880, 1013, 1023, 1024, 1095, 1115, 1116,
1127–1129, 1132, 1162, 1253, 1272, 1302, 1336, and 1344 for
insect data. Drosophila has at least 15 myosin heavy chain
isoforms (all alternatively spliced from a single gene)
(109 –111, 185, 347, 406, 434, 435, 550, 1023, 1024, 1253);
many show tissue or developmental specificity (109, 180,
185, 347, 406, 550, 616, 816, 880, 1024, 1336; Refs. 823, 1132
are reviews). The molecular basis of expression specificity is beginning to be investigated (235, 347, 424, 434, 435,
1115, 1116). Exon variation in the heavy chain head region
suggests the region where light chain binding occurs is
important for myosin function (110). Exon variation in the
rod suggests that variation in the rod hinge is associated
with contraction rate changes (185). ATP-induced dissociation of actin from indirect flight muscle isoform thick
filaments is fast, similar to mammalian skeletal muscle
values (1095). However, comparing different isoforms
shows that they all have similar, fast, cross-bridge detachment rates (791) and unitary step (single cross-bridge
cycle) displacements (692). Variation in these parameters
thus does not explain isoform differences in thin filament
displacement velocity in Drosophila.
Multiple myosin heavy chain mutants have been isolated in Drosophila (180, 185, 616 – 618, 816, 819, 879, 880),
for some of which rescue and overexpression data are
available (223). Single amino acid mutations in the rod of
Drosophila indirect flight muscle myosin cause flightmuscle degeneration with age (even though myofibril assembly appears normal) (618), and mutations in the head
dramatically decrease thick filament number (617). Mutants that produce no myosin heavy chain produce relatively normal thin filament arrays (89, 180), and altering
actin-to-myosin ratios suggests that filament imbalances,
not lack of thick filaments per se, are responsible for
observed sarcomere structure defects (89). In support of
this, mutants that reduce thick filament number disproportionately affect flight and jump muscles (which have
highly organized sarcomeres) compared with intersegmental muscles (which have more variable myofilament
arrays) (879). Drosophila expressing only embryonic myosin heavy chain isoforms are viable but flightless with
normal-appearing muscles, and the flight muscles show
progressive deterioration (1272). Drosophila embryonic
myosin has lower actin sliding rates and basal ATPase
activity than adult indirect flight muscle myosin, and replacing Drosophila indirect flight muscle myosin with the
embryonic isoform slows muscle kinetics, which presumably underlies the flightlessness of such mutants (1127,
1129). The kinetic rate constants of the two isoforms
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1016
show different dependencies on the presence of phosphate, suggesting that the difference between them is that
in the embryonic form the rate-limiting step of the crossbridge cycle is ADP release, whereas in the indirect flight
muscle form the rate-limiting step is phosphate release
(during the cross-bridge cycle, myosin head rotation is
associated first with phosphate and then ADP release, see
second review) (1131). Substituting only the embryonic
S1 exon (one of four in this region that differ) decreases
ATPase activity, but does not affect actin sliding velocity,
flight muscle ultrastructure, or flight ability (1128). However, switching of indirect flight muscle exons into the
embryonic isoform (when the embryonic isoform is the
one being expressed in the flight muscles), or embryonic
exons into the flight muscle isoform, alters wingbeat frequency and increases or decreases, respectively, the muscle’s maximum power generation and optimal wingbeat
frequency for power generation (1130). Unlike Drosophila, two myosin heavy isoforms are present in fleshfly
(Phormia) asynchronous and locust synchronous flight
muscle (1344).
2. Catchin/myorod
An alternative splice of the myosin heavy chain gene
codes for catchin/myorod (1075–1077, 1299), a protein
found in smooth muscle thick filaments in a variety of
bivalves. The protein contains the COOH-terminal rod of
myosin heavy chain but has a unique NH2-terminal head
(1076, 1299) and, although most highly expressed in catch
muscles (1299), is not necessary for catch (1298). Despite
the identity of the tail portions, pure catchin/myorod polymerizes very differently than myosin or myosin rod.
Adding myosin heavy chain to myorod preparations dramatically alters myorod polymerization (1075). A similar
isoform, myosin rod protein, is encoded in Drosophila by
a gene internal to the myosin heavy chain gene. Myosin
rod protein is present in a variety of larval, embryonic,
and adult cardiac, visceral, and somatic (including direct
flight) muscles and localizes with myosin in the A band
(950, 1114). Myosin rod protein may be associated with
the relatively disordered thin and thick filament packing,
variable thin-to-thick filament ratio, and bent thin filaments observed in some direct flight muscles (1114). In
muscles expressing high levels of myosin rod protein,
thick filaments and sarcomeres still form in animals lacking myosin heavy chain, suggesting that sarcomere formation does not require actin-myosin heavy chain interaction (950).
3. Myosin light chains
Vertebrate myosin possesses two light chains, the
essential and regulatory, in equimolar amounts (one each
per myosin molecule, and hence two each per myosin
heavy chain dimer). All known invertebrate myosins simPhysiol Rev • VOL
ilarly contain two essential (also called SH, catalytic,
myosin light chain 1, and alkali light chain) and two
regulatory (also called myosin light chain 2) light chains,
both of which have an ellipsoidal shape (Fig. 4) in a
variety of species (1111). [Caution: in early scallop papers, the regulatory light chain is also called the EDTA
chain because EDTA exposure removes one or both (depending on concentration and temperature) regulatory
light chains (32, 169, 553, 596, 1139), presumably due to
the enhanced regulatory light chain binding to the heavy
chain that Ca2⫹ and Mg2⫹ induce (31, 1139). EDTA sensitivity is not present in many other species (553, 654),
and the term is not used in modern parlance, but could be
confusing when reading the older literature.]
Whether all invertebrate myosins contain equimolar
amounts of the two light chains, however, is not as clear.
Early work in the scallop suggested that there were two
moles of essential light chain but only one of regulatory
light chain per myosin heavy chain dimer (1139). More
recent work, however, shows that scallop myosin does
contain two moles of each light chain per dimer (553,
1294). The situation in Crustacea remains unclear. Lobster and crayfish myosin contains two light chain types, ␣
and ␤, each with multiple variants, but which type is
essential and which is regulatory is unknown (570, 834,
984). In some lobster muscle fibers, ␣-chain immunohistochemical staining is higher in the center of the thick
filaments, whereas ␤-chain staining is always uniformly
distributed (133). Although these data are suggestive, they
do not prove nonuniform chain distribution because they
could result from nonuniform masking of the ␣-chain in
different thick filament regions. No difference is seen by
two-dimensional gel electrophoresis in lobster light
chains from slow versus fast muscle fibers (684), although
different isoforms are present in the two muscle types in
crayfish (1040). In earthworms, different light chain isoforms are expressed in a muscle-specific fashion, but
which are regulatory and which are essential is unknown
(160). Ascaris has two light chains in an approximate 1:1
molar ratio, but which are regulatory and which are essential is unknown (844). In vertebrates the light chains
are associated with the myosin heads near the neck region (Fig. 4A). Scallop light chains are similarly situated,
and the regulatory and essential chains extensively interact (34, 217, 309, 399, 598, 1112, 1140, 1217, 1219, 1244,
1245, 1247, 1283, 1294). Bivalve light-chain levels show
characteristic changes in response to various pollutants
(1000).
Essential light chains have been investigated in amphioxus (438), ascidia (1148), annelid (249a), several bivalves (70, 190, 375, 496, 723, 739, 740, 862, 940, 1139,
1246), abalone (32), squid (595, 596, 1258), and Drosophila (181, 288 –290, 665, 1153, 1155, 1156). In amphioxus,
scallop, and Drosophila only one gene was found. Early
work with gel electrophoresis showed that essential light
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chains vary across bivalve species (740) but were identical in all muscle types in the one species (Spisula) in
which this question was examined (739). Later radioimmunoassay work using scallop antibodies showed that
the essential and regulatory light chains differ immunologically, that the antibodies did not cross-react with
other invertebrate essential light chains, and that striated
and smooth, but not heart, essential light chains were
immunologically identical (1246). Sequencing work
showed that several essential light-chain transcripts are
present in scallop (375), and identical isoforms are
present in striated and catch muscle (940) (cardiac muscle was not examined in this work). Essential light-chain
isoforms, generated by alternative splicing and protein
phosphorylation, are expressed in a muscle-specific fashion (indirect flight muscles vs. all others) in Drosophila
(288, 289, 1155, 1156, 1160). The splice patterns are maintained across several Drosophila species (665).
Regulatory light chains have been investigated in
amphioxus (1124); ascidia (1148); earthworm (228, 1072)
and the pogonophore Riftia (971); several bivalves (70,
375, 496, 592, 593, 597, 724, 739, 740, 800, 824, 862, 940,
1139, 1157, 1246), abalone (32), and squid (595, 596, 725);
C. elegans (227, 1029, 1030); Limulus (1248) and tarantula
(425); and Drosophila (924, 1153, 1155, 1156, 1178). Early
gel electrophoresis work showed that regulatory light
chains vary across bivalve species (740). Later radioimmunoassay work showed that scallop regulatory light
chain antibodies did not cross-react with other invertebrate regulatory light chains, and that striated and
smooth, but not heart, regulatory light chains are immunologically identical (1246). Bivalves (375, 940) and Drosophila (924, 1178) have one gene, and C. elegans has two
(227). Muscle, developmental stage, or gender specific
regulatory light-chain isoforms exist in earthworm (228),
C. elegans (1030), and Drosophila (1156, 1160, 1178).
Muscle specific isoforms are apparently not present in all
bivalves, as only one isoform is present in clam (Spisula
sachalinensis) adductor mantle, leg elevator, and heart
muscle (739), but two (phasic vs. catch muscle) in scallop
(592, 593, 824, 940). However, the different phasic and
catch muscle ATPase activities in bivalves are due to
differences in the myosin heavy chains, not the regulatory
light chains (940). Blocking scallop heavy chain thiol
groups blocks regulatory chain binding (594, 597, 896).
In C. elegans one gene is redundant, but deletion of
the other causes lethal pharyngeal muscle defects in hermaphrodites. This likely occurs because the first gene is X
linked, and its expression is downregulated (dosage compensated) in hermaphrodites (which have two X chromosomes). Consequently, without the second gene, in hermaphrodites insufficient regulatory light chain is produced in the pharynx (1030). The Drosophila isoforms are
believed to arise by posttranslational modification (1155,
1178), although several transcripts are expressed (924).
Physiol Rev • VOL
1017
Drosophila null mutation heterozygotes in the regulatory
light chain have reduced adult thoracic musculature,
somewhat disorganized indirect flight muscle, and altered
wing beat frequency spectrum or are flightless (464, 1251,
1252); changing a single amino acid from serine to alanine
reduces flight muscle power output (252). Drosophila
regulatory light-chain NH2-terminal deletions reduce indirect flight muscle calcium sensitivity, maximum amplitude of oscillatory work, sarcomere length, dynamic stiffness, and elastic modulus (476, 820). The stiffness and
elastic modulus changes suggest that in these muscles
regulatory light chain could link the thick and thin filaments.
4. Myosin light-chain phosphorylation
Expressed sequence tag analysis shows that a light
chain kinase is present in amphioxus notochord (1124). A
scallop cAMP-dependent kinase that phosphorylates only
one regulatory light chain isoform and a protein that
modulates the kinase have been identified (1098 –1100,
1102). A Limulus light chain specific kinase has been
purified (1071). The Drosophila light chain kinase gene
has been sequenced. Tissue- and stage-specific isoforms
are present; only some forms are Ca2⫹/calmodulin dependent (586, 1179). Reducing Drosophila regulatory lightchain phosphorylation reduces myosin ATPase activity
(1154) and, by reducing muscle stretch activation (a special property of the asynchronous muscles present in this
organism, see second review) (1110, 1180), indirect flight
muscle power output (252, 1110, 1180). Regulatory lightchain phosphorylation increases tarantula striated muscle
contraction; phosphorylation levels vary with Ca2⫹ concentration (425). Ascaris cuticle muscle regulatory light
chain is 48% phosphorylated in controls; GABA application reduces this phosphorylation to 18% (747). Phosphorylation of Limulus regulatory light chains in purified thick
filaments changes filament structure (674). Twitchin also
phosphorylates regulatory light chains (415). Light chain
phosphatases have been characterized in clam (1194) and
scallop; the scallop enzyme is Ca2⫹ dependent (474).
5. Paramyosin/miniparamyosin
We cover here only paramyosin genes and mutants
(see second review for structure of paramyosin containing thick filaments). Paramyosin/miniparamyosin have
been studied in cephalochordata (167, 437), echinodermata (876, 1195, 1256, 1284), annelids (1013, 1018, 1270,
1284), mollusks [Polyplacophora (786), bivalves (2, 56 –
60, 197, 198, 205–208, 270, 392, 397, 529, 547–549, 636, 641,
642, 677, 704, 738, 740, 766, 775, 786, 789, 795–797, 944,
991, 992, 1027, 1113, 1137, 1255–1257, 1270, 1284, 1290,
1291, 1311) (called tropomyosin A in Refs. 57, 60, 642, 795,
1027 and tropomyosin in Refs. 56, 58, 397, 547, 549, 641,
797), gastropods (401, 529, 786, 958, 1018, 1216, 1256,
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1018
1291), cephalopods (56, 561, 619, 762, 1197) (called tropomyosin A in Ref. 762 and tropomyosin in Ref. 619)],
brachiopods (1284), Chaetognatha (1011), platyhelminths
(91, 156, 298, 359, 387, 444, 478, 521, 635– 637, 830, 1056,
1087, 1207, 1284, 1305), nematodes [C. elegans (24, 277,
346, 402, 514, 710, 939, 1059, 1133, 1256, 1262, 1263),
Ascaris lumbricoides (1284), Brugia malayi (648, 1274),
O. volvulus (231, 688), Dirofilaria immitis (376, 688)],
Limulus (248, 270, 468, 675, 676) (called tropomyosin A in
Ref. 468), mites/ticks (297, 967, 1021), Crustacea (202,
270, 677, 834, 836, 852, 1016, 1040, 1256, 1284), Diptera (D.
melanogaster) (27, 90, 696, 743, 1013, 1018, 1231, 1232),
Coleoptera (Heliocopris japetus, Pachnoda ephippiata)
(145), Heteroptera (Lethocerus) (141, 145, 270, 676, 677),
and Orthoptera (Locusta migratoria, Schistocerca gregaria) (93, 590, 1284). References 270, 528, 677, 1013,
1018, 1256, and 1284 are reviews. Paramyosin is observed
in amphioxus notochord (311, 437), catch (56, 60, 270,
397, 448, 766, 1027, 1137), smooth (24, 401, 876, 1013,
1018, 1027, 1195, 1284), obliquely striated (24, 396, 561,
1018, 1262, 1284), and cross-striated (93, 145, 166, 202,
248, 270, 468, 561, 675, 676, 852, 944, 1013, 1016, 1018,
1021, 1040, 1056, 1231, 1284) muscle and thus is not a
marker of catch ability or muscle type (except for existing
only in invertebrates). Paramyosin expression varies in
mollusk cross, obliquely, and nonstriated (smooth) muscles, although which types express more is inconsistent
across species (561, 1018).
Paramyosins from two bivalves, a gastropod, and a
polyplacophore all have similar molecular weights, electrophoretic properties, solubility, and guanidine-HCl denaturation and pepsin and trypsin susceptibilities (786).
Paramyosins from two smooth and one striated molluscan muscles, and four arthropod (Limulus, Homarus,
barnacle, insect) striated muscles, have similar molecular
weights and are immunologically similar (270). Heteroptera and Coleoptera indirect flight muscle paramyosins
also have similar intrinsic sedimentation rates and circular dichromism values (145). However, this does not mean
that all paramyosins are identical; paramyosin from different species (248, 528, 529, 766, 876, 1256) and multiple
isoforms within single species (2, 202, 693, 834 – 836, 852,
1040, 1056, 1106, 1231, 1243) show differences in antigenic
structure, molecular weight, isoelectric focusing, phosphorylation ability, or amino acid composition. The two
to three paramyosin isoforms present in decapod crustacea show muscle specific expression (834 – 836, 852,
1040).
The sources of this variation are not completely
known [for very early work, some may arise from proteolytic degradation during extraction (1311)]. Paramyosin is
phosphorylatable (2, 197, 249, 1056, 1059, 1101, 1256,
1257), which could explain differences in antigenic structure, isoelectric focusing, and phosphorylation ability
(since paramyosins with different phosphorylation states
Physiol Rev • VOL
in vivo would vary in their ability to be further phosphorylated in vitro). The amount of phosphorylation
observed depends on extraction procedure (197). In
mollusks Ca2⫹-, cAMP-, and calmodulin-dependent kinases phosphorylate paramyosin (1036), neuromodulators alter phosphorylation ability (2), and phosphorylation alters myofibril ATPase activity (1257), suggesting
that paramyosin phosphorylation could be functionally
relevant.
Paramyosin isoforms could also arise from multiple
genes. Mytilus has two paramyosin genes, but whether
one is a separate miniparamyosin gene is unknown
(1255). Two genes may be present in the nematode Anisakis (939), and the platyhelminth Schistosoma has more
than one (444). Paramyosin cDNAs or genes have been
cloned in the cestode Taenia solium (1207) (from the T.
solium gene a mini-paramyosin cannot expressed); nematodes Brugia (648), Onchocerca (231, 688), and Dirofilaria (376, 688); tick Boophilus (297); and mite Blomia,
(967) but whether these are single copy is unclear.
C. elegans [unc-15 (514, 993, 1263)] has a single
paramyosin gene but two paramyosin isoforms, which are
differentially located in the thick filaments, one being
associated with the filament core and the other with the
myosin heavy chains (240, 693). Reference 514 asserts
that paramyosin expression differs in body wall and pharyngeal muscles, but we have been unable to find sources
showing this difference. Regardless, paramyosin is expressed in all C. elegans muscle (24). C. elegans paramyosin mutants show varying degrees of locomotion impairment, and in the most extreme phenotype are paralyzed
as adults (1263). The body wall muscles of these mutants
show aberrant myofilament lattice and thick filament
structure (277, 1263). Changing a single charge on the
paramyosin can disrupt thick filament assembly (346),
and paramyosin appears to be required to determine thick
filament length (710) [as is also the case in Limulus (468)
and locust (590)]. Many C. elegans paramyosin mutants
that disrupt body wall muscle function do not alter pharyngeal muscle function, although the pharyngeal thick
filaments are missing their normal paramyosin core
(1263). This difference is believed to be because pharyngeal muscles have less paramyosin, use different myosin
heavy chains than body wall muscles, and have only one
sarcomere (514, 1263). Paramyosin degradation is increased in C. elegans mutants lacking myosin heavy chain
B, although synthesis is unaffected (1279).
Drosophila (1232) has a single paramyosin gene that
is required for proper myoblast fusion, myofibril assembly, and muscle contraction (696). Paramyosin is much
more highly expressed in tubular than in flight muscle in
Drosophila (1231). Comparing paramyosin content of
solid- versus open-cored thick filaments (see second and
third reviews) across several insects shows that paramyo-
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sin is a larger percentage of filament mass in solid-cored
filaments (93).
Paramyosin is an important component in allergic
and immune reactions and may be useful in platyhelminth
and nematode vaccine development (3, 8, 13, 25, 83, 105–
108, 159, 171, 193, 201, 241, 245, 284, 307, 315, 344, 357,
358, 422, 495, 509, 521–523, 527, 583, 585, 635, 636, 644,
645, 648, 680, 681, 683, 685, 700, 726, 727, 777–780, 830,
846 – 848, 934 –936, 939, 965–969, 988 –990, 1078 –1080,
1117, 1165, 1172, 1173, 1187–1189, 1208, 1209, 1242, 1303–
1306, 1341). An important point for working on these
animals is that aldehyde fixation blocks muscle paramyosin staining in them (637).
Miniparamyosin is present in Drosophila (90, 744,
1013), shrimp (1016), and mite striated (1021) muscle and
Eisenia obliquely striated and smooth (1013) muscle.
Whole animal extracts show it to be present in annelid,
snail, mussel, and multiple arthropod species, but not C.
elegans, a result supported by analysis of the C. elegans
paramyosin gene (744). Drosophila mini-paramyosin is
encoded by the same gene as paramyosin, but its expression is transcriptionally regulated from a separate promoter (27, 90, 744). Drosophila miniparamyosin is expressed only in adults and has several isoforms (with
much greater molecular weight variation than those
known for paramyosin) that are expressed in a muscle
specific pattern (90, 743). The sequences that regulate
miniparamyosin/paramyosin expression are beginning to
be described (27). Miniparamyosin overexpression results
in nearly normal thick filaments and sarcomere length.
However, some defects are present in indirect flight muscle with some flight impairment, and these defects increase with age (28).
6. Filagenins
C. elegans thick filaments are associated with three
30-, 28-, and 20-kDa proteins (␣-, ␤-, and ␥-filagenin, respectively) (240), whose cDNAs have been sequenced
(694, 695). The proteins are believed to form the thick
filament core by cross-linking paramyosin subfilaments
(693). The ␣-form is located with myosin heavy chain A in
the middle of the thick filaments while the ␥-form is
located more laterally with myosin heavy chain B (695).
The ␣- and ␥-forms appear at different developmental
stages (695). The ␤-form is not present in pharyngeal
muscles (694).
Temperature-sensitive unc-45 mutants have reduced
body wall muscle thick filament number and are paralyzed as adults (279); homozygous, non-temperature-sensitive mutations result in arrested muscle development
and are lethal (1213). The gene has been sequenced and is
expressed in all C. elegans muscles, and only in muscle
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(1212). The protein associates with myosin heavy chain B
in the thick filament. In myosin heavy chain B null mutants unc-45 muscle localization is not seen (and thick
filament structure is near normal) (22). unc-45 binds
myosin heads (both scallop and C. elegans, although unfortunately myosin heavy chain A and B were not separately tested) (72). Temperature-sensitive mutations in a
region of unc-45 homologous to two fungal proteins involved in protein segregation result in (at the restrictive
temperature) myosin heavy chain A and B being scrambled in the thick filaments (as opposed to being located in
distinct thick filament regions as in wild type) (71). This
work suggests that unc-45 is a chaperone that helps
correctly integrate myosin heavy chain B into the thick
filament (693, 1318).
8. Myonin
A 230-kDa thick filament associated protein named
myonin that appears to regulate myosin heavy chain
ATPase activity has been identified in clam (1310).
9. Flightin
Flightin is a 20-kDa protein that has been found only
in Drosophila (1225, 1229), but efforts to identify it in
other species have not been reported. It is found only in
indirect flight muscles and is localized in the A band
except for the H-band region (Fig. 8) (1229). The protein
has multiple phosphorylation sites, and its phosphorylation state increases with age, particularly after eclosion
(1227). The single amino acid myosin gene mutations
noted above that cause age-dependent myofibril degeneration also fail to accumulate phosphorylated forms of
flightin (618). Mutations that block thick filament assembly block all flightin phosphorylation, whereas mutations
that block thin filament assembly alter flightin phosphorylation temporal sequence (1224). Flightin null mutants
are viable but flightless (976). Mutant sarcomeres appear
relatively normal in pupa and early adult but are ⬃25%
longer than normal. When the animals attempt to fly, the
indirect flight muscles become disrupted, and eventually
site-specific thick filament cleavage occurs. Flightin binds
to the myosin rod, and the flightin:myosin molar ratio is
between 1:1 and 1:2 (44).
10. Zeelin
Zeelin 1 and 2 have been isolated from Lethocerus
muscles (294, 1037). Zeelin 1 is present in flight and leg
muscles, and zeelin 2 is only in flight muscles. In flight
muscles the zeelins are present in discrete A-band regions, whereas in leg muscles the entire A band is labeled
(the identification in Ref. 1037 that these proteins were in
the Z disk was an artifact of fiber glycerination) (294).
Both proteins are associated with the thick filament, with
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zeelin 1 closer to the filament shaft than zeelin 2, and ⬃0.5
mol zeelin/ mol myosin (294). Zeelin function is unknown.
The small size and location of the zeelins are similar to
Drosophila flightins, but antibodies to zeelin do not recognize flightin (294).
protein) 43-kDa Drosophila Z-disk associated protein has
been identified, but its structure and function are unknown (998).
C. ␣-Actinin and Other Z Line Proteins
A wide variety of Ca2⫹ binding proteins (calmodulin,
Ca -dependent Ca2⫹ binding protein, troponin C, the B
subunit of phosphatase 2B, myosin light chains, parvalbumins) have evolved from a common ancestor with four
Ca2⫹ binding domains (see Refs. 574, 1144 for vertebrate
references, Refs. 610, 611, 818 for extensive summaries of
Ca2⫹ binding proteins and their evolutionary relationships, and Ref. 746 for a comparison of Ca2⫹ binding site
amino acid complements). These proteins are divided into
two types (209, 210). Proteins of the first (e.g., vertebrate
parvalbumin) are present in high quantity, have high Ca2⫹
affinity, have Ca2⫹ binding regions with a stable configuration, do not interact with known targets, are believed to
function as Ca2⫹ buffers and to help resequester Ca2⫹
after muscle contraction or to protect against deleterious
effects of high Ca2⫹ concentrations during prolonged contractions, and show relatively rapid evolutionary divergence (reviewed in Ref. 421). Proteins of the second are
present in lower quantity, have lower Ca2⫹ affinity, have
Ca2⫹ binding regions whose conformation changes upon
Ca2⫹ binding, are believed to be more directly involved in
excitation-contraction coupling (examples include troponin C and calmodulin), and are relatively well conserved.
We cover here only muscle specific Ca2⫹ binding proteins
(in particular, calmodulin, which is present in a wide
variety of tissues, is not included). Reference 348 reviews
Ca2⫹ binding proteins in general.
Amphioxus (Branchiostoma lanceolatum) has seven
sarcoplasmic Ca2⫹-binding proteins of the first type, all
arising by alternative splicing from one gene. One isoform
has been crystallized and its structure determined at 2.4-Å
resolution (194, 196). The two isoforms for which it has
been examined bind three Ca2⫹ (582, 1141, 1146, 1150).
Immunohistochemistry shows the proteins are localized
at the Z line (1202). One protein of the second type, Ca2⫹
vector protein, has been characterized in amphioxus and
binds two Ca2⫹ (62, 63, 209, 211, 574, 1124). The molecule’s three-dimensional solution structure, backbone dynamics, and Ca2⫹ binding properties have been studied in
great detail (62, 63, 1169 –1171). Its cDNA has been sequenced (1320), and although clearly a Ca2⫹ binding protein, no homolog has been found in invertebrates or vertebrates (1202). The protein is present in a wide variety of
tissues, but in highest concentration (50 –100 ␮M) in muscle (209, 412). Immunohistochemistry shows it to be
strongly localized at the Z lines and weakly localized at
the M lines, and it is more difficult to extract from muscle
than the sarcoplasmic Ca2⫹-binding proteins (1202). Ca2⫹
D. Ca2ⴙ Binding Proteins and Their Targets
2⫹
␣-Actinin has been isolated or identified by immunohistochemistry in annelid (Eisenia) obliquely striated and
smooth muscle (1010); bivalve (scallop) cross-striated,
obliquely striated, and smooth muscle (559, 1074); gastropod (Helix) cross-striated (heart) and smooth muscle
(1010); cephalopod (squid) obliquely striated muscle
(1196); chaetognath (Sagitta friderici) body wall muscle;
platyhelminth (Schistosoma mansoni) muscle of unidentified type (706); C. elegans obliquely striated muscle
(316), mite cross-striated muscle (1021); and Drosophila
(1010, 1035, 1228), honeybee (Apis mellifera) (1034), and
giant water bug (Lethocerus cordofanus) cross-striated
(146, 393) muscle. In insects, ␣-actinin staining was seen
throughout the Z line (1010). In the obliquely striated
muscles of earthworm, snail, and C. elegans, in which thin
filaments attach to peglike dense bodies analogous to
cross-striated Z lines (see third review), the staining was
limited to these formations (316, 1010). In earthworm and
snail smooth muscle, the staining was limited to discontinuous patches, suggesting that in these muscles the thin
filaments also have discontinuously distributed anchorage sites (1010).
Drosophila has a single ␣-actinin gene that gives rise
to three isoforms (338, 1008). Two are muscle specific.
One is present in larval and adult head and abdominal
muscles and the other in adult indirect flight muscles and
tubular jump and leg muscles (1008, 1035, 1228). Mutations can be fatal or cause severe muscle defects (327,
338, 441– 443). Rescue experiments show that 1) the indirect/jump/leg muscle isoform can replace the cytoplasmic and larval/head/abdominal forms and 2) increasing
␣-actinin’s length 15% does not affect its function (264).
␣-Actinin and kettin (see below) bind actin simultaneously and well (1205). A C. elegans ␣-actinin cDNA
clone has been isolated (73).
In C. elegans a small heat shock protein, HSP25, that
binds ␣-actinin and vinculin but not actin, is found in
muscle-dense bodies and M lines (253). Early immunohistochemistry in Drosophila and honeybee identified small
(150 –200 kDa) (some present in only very restricted sets
of muscles) and larger (400, 600 kDa) (of which the
600-kDa band showed some muscle specificity) non-actinin, non-projectin Z-disk proteins (1034, 1035, 1228). An
apparently novel (the 10 amino acid sequence against
which the monoclonal antibody used to identify it was
raised showed no close match to any known Drosophila
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vector protein forms a high-affinity, Ca2⫹-strengthened,
complex with Ca2⫹ vector target protein (209, 942). Ca2⫹
vector target protein is also present in both muscle and
nonmuscle tissues, but equally stains the Z and M lines
(1202). Ca2⫹ vector target protein does not appear to have
any enzymatic role, and its function is unknown. However, it has two immunoglobulin domains and thus could,
as do other immunoglobulin containing muscle proteins
(see sect. IIIE), contribute to sarcomere structure (1142).
cAMP-dependent protein kinase phosphorylates four
Ca2⫹ vector target protein sites, and phosphorylation decreases Ca2⫹ vector protein and Ca2⫹ vector target protein affinity (943).
Low-molecular-weight muscle-associated Ca2⫹ binding
proteins are present in several other invertebrates. Nereis diversicolor and Perinereis vancaurica tetradentata (annelid) sarcoplasmic Ca2⫹ binding proteins bind
three Ca2⫹, but that of Nereis virens only binds two.
The N. diversicolor and Perinereis proteins have been
sequenced (189, 213, 242, 577). The Nereis protein’s
three-dimensional structure (54, 195, 1230) and dynamics of Ca2⫹ and Mg2⫹ binding (274) have been determined, and the conformational changes associated with
Ca2⫹ binding are being intensively studied (178, 216,
266, 705, 953, 1094). Earthworm has three Ca2⫹ binding
protein isoforms that bind either two or three Ca2⫹, and
which are present in higher concentrations in fast relaxing muscles (consistent with their being involved in
Ca2⫹ buffering) (459, 460). The 20-kDa Ca2⫹ binding
proteins have been isolated from two scallops. The
Patinopecten protein has been sequenced and likely
binds two Ca2⫹ (191, 1143). Clams and oysters have
three Ca2⫹ binding protein isoforms (1141). Low (17
kDa)- and high (450 kDa)-molecular-weight Ca2⫹ binding proteins and a calsequestrin (the major Ca2⫹ binding protein in vertebrate skeletal and cardiac muscle)
homologs are present in the membrane fraction of
Mytilus anterior byssal retractor muscle (1301). A Ca2⫹
binding protein uniquely expressed in muscle that is
closely associated with the contractile machinery, and
which cross-reacts to antibodies to amphioxus sarcoplasmic Ca2⫹ binding protein, has been identified in
Aplysia, as has another present in both muscle and
neurons (932). A 20-kDa, muscle- and tegument-associated, Schistosoma Ca2⫹ binding protein has been isolated and sequenced (408, 409, 1118), and a gene identified that codes an 8-kDa Ca2⫹ binding protein of unknown location, but which is only expressed during the
animal’s free swimming stage (964). C. elegans calsequestrin is expressed only in body wall muscle. Its
function is unknown, but the sequences regulating its
expression are beginning to be identified (177).
Unlike other known invertebrate Ca2⫹ binding proteins, crayfish (102, 214, 498, 1287–1289), lobster (1289),
and shrimp (1145, 1147, 1289) sarcoplasmic Ca2⫹ binding
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proteins are dimers of two nonidentical proteins, each of
which binds three (for a total of 6 per dimer) Ca2⫹; ␣␣,
␣␤, and ␤␤ are all present. Ca2⫹ binding induces large
conformational changes. The amino acid sequences of the
crayfish (498) and shrimp proteins are known (1145,
1147), and the crayfish protein has been crystallized (612).
Additional Ca2⫹ binding proteins have been isolated from
crayfish and lobster muscle (420, 759), one of which has
sequence similarity to calcyphosine, a vertebrate nonmuscle Ca2⫹ binding protein (1051). Although early work
was unable to find sarcoplasmic Ca2⫹ binding proteins in
locust (212), 16- to 20-kDa Ca2⫹ binding proteins with
muscle specific (tubular vs. indirect) expression were
identified in Drosophila (46, 1160). The 23- to 24-kDa Ca2⫹
binding proteins were subsequently isolated from Drosophila and Calliphora (551, 560). The cDNA for the
Drosophila protein has been cloned and is expressed in
tubular but not indirect flight muscle (551). The gene is
not single copy (551), and multiple isoforms are observed
(560). A Drosophila calbindin homolog that is primarily
expressed in neurons but is present in a few muscles has
also been identified (986).
E. Giant Sarcomere-Associated Proteins
Like vertebrates, invertebrates also have large sarcomere proteins involved in centering the thick filaments in
the sarcomere and generating muscle stiffness. Invertebrate muscles, however, show a number of variabilities
not present in vertebrate sarcomeres (144, 323, 621). First,
invertebrate sarcomeres cover a very wide length range
(crayfish giant sarcomeres are ⬃8 ␮m at rest whereas
those of Drosophila indirect flight muscles are 3.3 ␮m).
Second, invertebrate muscles have very different abilities
to be stretched without damage (crayfish giant sarcomeres can stretch to 13 ␮m, Drosophila indirect flight
muscle sarcomeres to only ⬃3.5 ␮m). Third, invertebrate
muscles show a very wide passive stiffness range (a tension of 8 mN/mm2 stretches crayfish sarcomeres 4 ␮m,
but Drosophila sarcomeres only 0.2 ␮m).
Given these differences, invertebrates might be expected to have a wide variety of giant muscle proteins,
and indeed, a large number were early identified on the
basis of molecular weight, sarcomere position, and antibody reactivity (316, 455, 456, 544, 545, 639, 698, 730 –732,
757, 845, 850, 851, 884, 1014, 1016, 1021, 1033–1035, 1037,
1218, 1220, 1221, 1228, 1343). It is now clear that these
proteins belong to multiple isoform families of wide size
range, that gel electrophoresis molecular weight estimates for very large proteins can be highly inaccurate
(323), and that many of these proteins, even those from
different families, are sufficiently similar to immunologically cross-react. Gene sequencing in C. elegans and Drosophila has begun to provide order to this field. However,
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in older literature terms such as “mini-titin,” “titin/
twitchin,” “titinlike,” etc., are widespread, and what protein is being described is not always clear. Even worse,
vertebrate titin is also called connectin, and some authors
refer to all large (⬎2 MDa) invertebrate sarcomere proteins as connectin (756), even though sequence data show
that some of these large proteins are very different. Moreover, in an extremely unfortunate happenstance, a Drosophila cell adhesion molecule present on embryonic
muscles is called connectin (870 – 872). Readers must
therefore exercise caution in the older literature. Reviews
covering invertebrate sarcomere giant proteins (at least in
part) include References 95, 98, 142, 144, 544, 756, 758,
1184, 1185, 1190, 1343, 1345.
To provide order to these proteins, it is helpful to first
examine human skeletal muscle titin (Figs. 7A and 8) (182).
The protein’s first 80 kDa, with its multiple immunoglobulin
domain repeats, binds to the Z disk and possibly actin.
Additional immunoglobulin repeats (which can provide elasticity by unfolding under tension, Ref. 649) and an elastic
PEVK region occur next and span the adjacent half I band in
which the molecule moves toward the thick filaments. Next
is a large region of a repeated immunoglobulin/fibronectin
domain motif, which is associated with the thick filaments in the A band. The molecule ends with a 250-kDa
M-line region that contains a serine/threonine kinase
activity and multiple immunoglobulin repeats. Thus, in
addition to being huge (one basis for calling various
invertebrate giant proteins titin-like), titin also has regions in which only or primarily immunoglobulin repeats exist, areas expected to act as springs, a region
with a repeated immunoglobulin/fibronectin motif, and
a kinase (and hence a wide range of epitopes against
which antibodies can be raised). A large variety of
proteins could thus resemble one or another portion of
titin but still be distinctly different from one another.
Titin is believed to fulfill four functions (379, 755, 1186).
First, its springlike nature helps keep the thick filaments centered in the sarcomere. Second, this same
nature provides much of muscle passive resistance to
stretch and elasticity. Third, it may help determine
sarcomere length in development. Fourth, although its
FIG. 7. The motif structure of human titin and the major invertebrate giant sarcomere-associated proteins. Red rectangles, immunoglobulin
domains; green rectangles, fibronectin domains; blue rectangles, SEK domains; yellow rectangles, serine/threonine kinase activity; spring, PEVK
domain. SEK and PEVK domains are so named (single letter amino acid code) because these are the most prevalent amino acids in these regions.
In all cases the NH2 terminus is to the left. The sequences in B are aligned at a conserved site (vertical double headed arrow) where a linker region
between two immunoglobulin domains is missing. Data are from References 97, 99, 144, 306, 323, 587, 708, 906, 1097, 1107.
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FIG. 8. Sarcomere location of the major
invertebrate giant muscle proteins. Top: a repeat of the vertebrate sarcomere shown in Fig.
5. Bottom: position of vertebrate titin and major
invertebrate giant proteins. Triangles represent
serine/threonine kinase position when known.
Line lengths are not proportional to protein molecular weight.
target(s) is unknown, titin’s kinase could regulate the
activity of other sarcomere proteins.
Except for amphioxus (455) and ascidia (845), in
which the proteins are presumably vertebrate titin, no
known invertebrate giant protein is long enough to
span an entire half sarcomere (566). Proteins in the 3
MDa range (similar to titin’s molecular weight) have
been identified by gel electrophoresis in some (but are
clearly absent from other) crayfish muscles (732) and in
the 5 MDa range in barnacle (730), but these lengths are
insufficient to span half the large (10 ␮m) sarcomeres
present in them. Very large (4-MDa) proteins that connect the Z line to the thick filaments are present in the
annelid Neanthes, but how far along them the proteins
extend is unknown (545). All other invertebrate giant
proteins are also too small to span a half sarcomere.
Present best knowledge is thus that invertebrate elasticity and sarcomere structure arise from proteins that
connect Z line and thick filament proteins, but which do
not extend all the way to the M line.
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1. Crayfish connectin/kettin, C. elegans kettin,
and Drosophila titin/kettin
Genes capable of producing 1) ⬃2-MDa proteins
(called connectin in crayfish and titin in Drosophila) with
an immunoglobulin repeat-rich region and springlike
PEVK regions but no protein kinase domain, and 2) ⬃550kDa proteins (called kettin in all species) consisting of
only immunoglobulin domains, have been completely or
partially sequenced in C. elegans (144, 391), Drosophila
(144, 587, 640, 708, 1337), Bombyx mori (584, 1123), and
crayfish (323). The reported Drosophila sequences show
some variation. The sequence predicted from the Drosophila genome project (144) lacks eight NH2-terminal
immunoglobulin repeats (left rectangle, below sequence)
present in the sequence of Reference 708, and has four
COOH-terminal fibronectin repeats (right rectangle) not
present in Reference 708. Drosophila and crayfish kettin
are produced by alternative splicing from the titin/connectin gene. C. elegans kettin is instead produced by its
own gene (see below for a large C. elegans sarcomere
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protein). In Bombyx, kettin is also produced by its own,
single-copy gene, and there are two titinlike genes. Given
the presence in C. elegans (see below) of another giant
sarcomere protein that is distantly, if at all, related to the
crayfish connectin/insect titin/C. elegans-Drosophilacrayfish kettin protein family, but which is nonetheless
named titin, the nomenclature in this field clearly needs
revision. Reference 144 suggests that the crayfish connectin/insect titin/C. elegans-Drosophila-crayfish kettin proteins be all renamed SLS proteins (from sallimus, the
gene that codes for Drosophila titin/kettin), with kettin
becoming just an SLS isoform.
Crayfish connectin runs from the Z line to the tips of
the thick filaments and stretches with the sarcomere to
lengths up to 3.5 ␮m (323). Crayfish connectin contains
both a PEVK region and a SEK region, which is also
elastic (322), and connectin proteolysis in skinned crayfish fibers abolishes resting tension (732). Antibodies to
an epitope on crayfish connectin that is not present in
Drosophila titin recognize a large protein in barnacle
ventral muscle (323), Calliphora larval muscle (323), and
beetle (Allomyrina), bumblebee (Bombus), and waterbug
(Lethocerus) leg muscle (323, 884). The beetle, bumblebee, and waterbug protein is recognized by antibodies to
vertebrate titin and is localized in the I band in beetle
(884). The crayfish connectin antibody also recognized a
3-MDa protein in waterbug, but not beetle or bumblebee,
flight muscle (884). A 540-kDa protein that localized to Z
lines, elongated upon muscle stretch, bound actin, and
was recognized by antibodies against vertebrate titin, and
a 500-kDa Lethocerus muscle protein was early isolated
from crayfish claw muscle (731). Given the cloning evidence for kettin in crayfish (323), this protein is presumably kettin.
C. elegans kettin has an immunological staining pattern consistent with dense body (equivalent to Z line)
expression, all muscles were stained (144, 587). Drosophila kettin is present in all larval body wall and visceral
muscles and in adult leg and flight muscles. Mutants of
what was believed to be the kettin gene have abnormal
muscles; homozygotes cannot hatch and heterozygotes
cannot fly (391). However, in light of titin and kettin
arising from one gene, it is now unclear if these effects are
due to a lack of kettin or titin. Kettin is located at the Z
line in flight muscle, with the NH2 terminus in the line and
the COOH terminus outside (140). Kettin binds to actin
filaments (1205) and in vitro promotes anti-parallel association of thin filaments (the same arrangement as that
present in vivo, see Fig. 5) (1205). Kettin’s immunoglobulin domains bind ␣-actinin and actin (640) and are consistent with a model in which each repeat and its flanking
region bind to one thin filament actin monomer, each
immunoglobulin/linker binding ⬃3 nm of the thin filament
(587, 1205). With this model, each sarcomere’s kettin
molecules would overlap by a small amount and extend
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some 60 nm outside the 80- to 120-nm Z-line width (587).
Kettin competes with tropomyosin for actin binding,
which could exclude tropomyosin from the Z line (1205).
Kettin’s COOH terminus can bind myosin; kettin could
thus link the Z line to the thick filaments (621) and is
therefore believed to play a major role in muscle stiffness
(140, 621). However, it also has large numbers of immunoglobulin domains, and single kettin molecules respond
to tension with steplike relaxations presumably stemming
from unfolding of individual immunoglobulin domains
(649). Kettin may therefore also play a role in muscle
elasticity.
Drosophila titin is required for myoblast fusion and
the formation of striated muscle sarcomeres (708, 1337).
Titin also plays an important role in muscle elasticity.
Although immunoglobulin/fibronectin unfolding presumably can play a role in this process, it is not its only
source, since muscle length changes in 2.3 nm quanta
when force is applied to indirect flight muscles under
conditions in which titin is the sole length-absorbing element. This distance is much less than that which would
occur from immunoglobulin or fibronectin domain unfolding and may correspond to single beta-sheet unfolding
(123). Multiple titin isoforms (in addition to kettin) are
expressed with some muscle specificity (144). Drosophila
titin is also present on condensed chromosomes and required for proper chromosome structure and function
(707, 708). A titin-like protein is present in Anopheles
(144).
2. C. elegans titin
The C. elegans titin gene produces a 2,200-kDa protein with multiple immunoglobulin and fibronectin repeats, an elastic region, and a kinase; a 1,200-kDa protein
with the immunoglobulin and fibronectin repeats and
elastic region, but no kinase; and a 301-kDa protein consisting of only the kinase and the COOH-terminal (primarily) immunoglobulin repeats (306). Immunohistochemistry shows that the 2,200- and 1,200-kDa isoforms are
expressed in all muscles except in the pharynx. The 2,200and 1,200-kDa proteins bind at their NH2 terminal to the
dense bodies (Z-line equivalents). The 2,200-kDa protein
may be long enough to reach slightly into the A band, and
thus its kinase could act on thick filament proteins. The
position of the 301-kDa protein is unknown. The gene is
not expressed until after C. elegans muscles are formed.
No C. elegans titin mutants are available, and thus the
functions of these proteins are unknown, as are the kinase’s targets. A survey of the C. elegans genome searching for immunoglobulin containing proteins identified
twitchin, unc-89, unc-52, and several titinlike sequences
(1167).
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3. Unidentified very large sarcomere proteins
Barnacle has 5,000- and 1,200-kDa proteins that
monoclonal antibodies to vertebrate titin recognize; the
5,000-kDa protein links the thick filament to the Z line
(730). This could be a crustacean equivalent to C. elegans
titin, but sequence data will be required to confirm this
relationship. Antibodies to vertebrate titin recognize a ⬎2
MDa scallop protein and label scallop thick filaments
along their entire length and most strongly at their tips
(1221). A similarly large titinlike (by antibody binding)
protein is present in squid (542). These data indicate that
mollusks contain a very large sarcomere protein, but its
relationship to other invertebrate muscle proteins is unknown.
4. Projectin/twitchin
A family of 600- to 1,000-kDa proteins with repeating
fibronectin/immunoglobulin repeats and a serine/threonine kinase domain exists in mollusks, C. elegans, crustaceans, and insects. The protein is called twitchin in C.
elegans, twitchin or twitchinlike in mollusks, and projectin in crustaceans and insects.
Twitchin (unc-22) was identified from C. elegans
mutants with a constant twitch and abnormal body wall
muscle sarcomeres (131, 678, 679, 809, 811, 813, 1003).
Twitchin consists of several immunoglobulin repeats,
multiple repeats of an immunoglobulin/fibronectin motif,
a serine/threonine kinase, and several immunoglobulin
repeats (96, 97, 314, 810). Up to six fibronectin/immunoglobulin repeats can be deleted without strong phenotypic effects (562). Bacterially expressed twitchin kinase
autophosphorylates (664). The 60 residues COOH terminal to the kinase core inhibit kinase activity (664) by
binding to the active site (intrasteric regulation) (457, 458,
578, 579). The Ca2⫹ binding protein S100A1 relieves the
inhibition and increases kinase activity 10-fold (413, 507).
Although several other Ca2⫹ binding proteins, including
calmodulin, bind to the autoinhibitory sequence (417),
they do not relieve the inhibition (664). Twitchin staining
is observed at both the A band and the dense body,
suggesting it links the thick filaments to the dense body
(767, 810). There are likely at least two twitchin genes, as
unc-22 mutants remove staining from body wall, anal, and
vulval muscles, but not pharynx (810). Mutations in unc54, which codes for myosin heavy chain B, can suppress
the unc-22 phenotype; the unc-54 mutants in isolation
have normal muscle structure, but the animals are stiff
and slow (812).
Striated and smooth scallop muscle and mussel
smooth catch muscle contains a 600 – 800 kDa (0.2 ␮m)
rod-shaped protein with a kinase domain (1220). Antibodies against the protein stain Limulus, Lethocerus (indirect flight), crayfish, and chicken muscle and bind to the
A/I junction in all except Lethocerus, in which they bind
Physiol Rev • VOL
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to the I band. Antibodies raised to the immunoglobulin
repeat or kinase domains of C. elegans twitchin recognize
the protein. Although both C. elegans twitchin and titin
have immunoglobulin repeats and a kinase, the protein’s
molecular weight suggests it is a projectin/twitchin.
Immunohistochemistry, immunoelectron microscopy,
and partial amino acid and cDNA sequencing indicate that
Aplysia californica has a twitchin-related protein that colocalizes with contractile filaments (416, 822, 954). Like C.
elegans twitchin, Aplysia twitchin autophosphorylates (416,
822), has an autoinhibitory sequence that binds Ca2⫹/calmodulin without activating the kinase (137, 413, 414, 416,
417), and is activated (in this case, ⬎1,000-fold) by Ca2⫹/
S100A1 binding (413, 414, 417). However, Aplysia twitchin
phosphorylates regulatory myosin light chains well (415),
whereas C. elegans twitchin only poorly phosphorylates
peptides containing the expected nematode regulatory myosin light-chain phosphorylation site (98, 664). C. elegans
and Aplysia twitchin also differ with respect to Ca2⫹/calmodulin affinity, sensitivity to naphthalene sulfonamide inhibitors, and the effect of Zn2⫹ on Ca2⫹/S100 activation
(required in Aplysia, inhibitory in C. elegans) (98, 414, 417).
Studies on neuropeptide modulation of Aplysia muscle relaxation indicate that cAMP-dependent protein kinase phosphorylates Aplysia twitchin, suggesting that twitchin may
regulate muscle relaxation (954), consistent with unc-22’s
phenotypic effects (809). Aplysia twitchin’s sarcomere location is unknown. Phosphorylation of a Mytilus twitchin-like
protein is involved in catch (see second review) (151, 152,
324, 326, 1088, 1089, 1298).
Early work with monoclonal antibodies against
Lethocerus muscle proteins revealed an 800-kDa protein
(639). A Drosophila protein, projectin, that is believed to
be the homolog of this Lethocerus protein, has been sequenced (47– 49, 232, 329, 1107). Projectin is very similar
to C. elegans twitchin (49), and projectin antibodies
cross-react with C. elegans twitchin (1228). Several isoforms, all of which retain the kinase and which show
muscle specific expression (1082, 1228), arise by alternative splicing (48, 232, 1107). The bent locus codes Drosophila projectin. Homozygous embryos do not hatch,
and in the most severe allele no muscle contraction ever
occurs (48, 329). Heterozygote muscles show reduced
stretch activation (821, 1226). Projectin, paramyosin, and
myosin aggregates form different diameter fibers depending on projectin’s phosphorylation state (287). In locust,
projectin (along with paramyosin) may help determine
thick filament length (590). Drosophila projectin binds to
thin filaments (1271), and locust projectin blocks the
formation in vitro of paracrystals from solutions of actin
monomers, instead promoting thin filament formation
(949). Locust projectin autophosphorylates (287, 1271)
and phosphorylates 30-, 100-, 165-, and 200-kDa proteins
(but not myosin light chain) in vitro (287, 1271). The
30-kDa protein may be troponin I, and troponin phosphor-
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ylation by projectin increases troponin’s Ca2⫹ sensitivity
(1271). Actomyosin, actin filaments, and myosin inhibit
projectin autophosphorylation, and calmodulin stimulates
it (287). Projectin phosphorylation of other proteins, however, is unaffected by actomyosin, actin filaments, myosin, calmodulin, Ca2⫹, Ca2⫹ and calmodulin, or cAMP
(287, 1271). Calmodulin-dependent protein kinase II
(CaMKII) binds to projectin, which increases CaMKII activity twofold (286).
In synchronous muscle, projectin is located in the A
band similar to twitchin (47, 639, 1035, 1228). In locust,
the molecule extends into the I band, and the molecule’s
COOH terminus, with the kinase domain, is located in the
A band (1082). In asynchronous muscle, it is instead
located in the I band and forms filaments connecting the
Z line and thick filament (47, 639, 1033, 1035, 1228). Neither myosin nor actin is required for projectin’s initial
localization to the Z line during sarcomere assembly, but
in the absence of actin projectin’s Z line localization is
later lost (45). Proteins believed to be projectin have been
isolated from locust, bee, beetle, and Lethocerus (142,
884, 1034), and staining with an antibody likely to have
been raised against Drosophila projectin stains (as expected) Drosophila cross-striated muscle and annelid
(Eisenia) obliquely striated and smooth muscle (1014).
A large sarcomere protein with immunological similarity to vertebrate titin and honeybee, beetle, and Drosophila projectin that contains a serine/threonine kinase
and autophosphorylates was early isolated from crayfish
(456, 745). This protein, crayfish projectin, has been
cloned, and flexor and closer muscles express different
isoforms (906).
The unc-89 mutants have disorganized muscle structure and lack M lines (95, 1266). The unc-89 gene has
been cloned and is believed to be a homolog of vertebrate
obscurin (98, 99, 1097, 1122). The gene produces four
isoforms that are differentially expressed in different
muscles. Two of the isoforms contain extensive immunoglobulin repeats. One of these isoforms also contains two
protein kinase domains, although only one is believed to
be functional. The other two isoforms are short versions
containing only the tandem protein kinase domains. All
the isoforms are localized in the center of the A band. The
structure of a region of the molecule that may be involved
in protein-protein interactions has been determined in
detail (120, 121). unc-98, a protein containing four Zn
finger motifs, is also located at the M line in C. elegans
(787). A 400-kDa M-line protein has been identified in
Lethocerus (142, 639), but without sequence data the
relationship of this protein and the unc-89 product is
unknown.
Physiol Rev • VOL
F. Miscellaneous Other Proteins
1. Nebulin
Nebulin-like proteins are present in obliquely striated
Eisenia muscle (1015) and cross-striated muscle in Antarctic mussel shrimp (1016), Antarctic mite Halacarellus
(1021), lancelet Branchiostoma (cephalochordate) (313),
and hagfish skeletal and heart muscle (312). Nebulin-sized
proteins are present in echinoderms, an annelid, mollusks, crustaceans, and insects (698). In Eisenia and Halacarellus, anti-nebulin staining occurs along the entire
sarcomere, whereas in Branchiostoma staining is only in
the I band.
2. Nesprin
Vertebrate nesprins are giant (1 MDa) actin binding
(and nuclear) proteins with extensive spectrinlike repeats
(1335). Spectrin-related Drosophila proteins of 200 –300
kDa molecular mass, located at muscle adhesion sites in
early development and later at the Z line and which were
originally believed to be possible dystrophin orthologs,
were early identified (1007, 1233). It now appears these
proteins arise from a nesprin gene ortholog that could
produce isoforms as large as 11,720 amino acids (1335). A
similar gene has been identified in C. elegans (1335).
3. Amphiphysin
Vertebrates have two amphiphysins, one involved in
endocytosis and the other in striated muscle t tubules.
Drosophila has a single amphiphysin gene whose product
is involved only in muscle, as mutants have normal synaptic transmission but severely disorganized t tubule/sarcoplasmic reticulum systems (973, 1332) and defective
membrane protein integration in the postsynaptic muscle
membrane (761, 1327, 1332).
4. Various muscle cell attachment proteins
To transmit force to effectors, the thin filaments of
the terminal sarcomeres must attach to the plasma membrane, and thence to tendons or their equivalent. This
attachment is accomplished by a protein complex including integrin, talin, vinculin, and filamen. Several of these
proteins, prominently integrin, also control the complex’s
development. This complex is not fully understood in
vertebrates and is less so in invertebrates. We here therefore simply list a number of proteins that attach thin
filaments to the muscle cell membrane, or muscle cells to
tendons or body wall structures, including primarily references showing that they serve a structural role. These
proteins include connectin (Drosophila) (963), DIM-1 (C.
elegans) (1001), integrin and integrin-associated proteins
[jellyfish (975), C. elegans (349, 432, 450, 691, 712, 787,
1004 –1006, 1054), Drosophila (122, 124, 147, 183, 263, 666,
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713, 854, 955, 1292, 1297, 1331), kakapo (Drosophila)
(380, 1120), myotactin (C. elegans) (454), spondin (Drosophila) (1201), talin (C. elegans) (219, 825), jellyfish
(975), Drosophila (136)], and vinculin (C. elegans) (74,
75). References 117, 125, 132, 134, 135, 149, 154, 389, 390,
465, 633, 947, 1002, 1329, 1330 are reviews or papers
describing muscle adhesion-deficient mutants or antibodies against cell adhesion molecules. Analysis of talin homologs from yeast, slime mold, and nematode reveals a
conserved protein domain believed to be specialized for
thin filament binding (772). Early monoclonal antibody
work identified three to five C. elegans muscle attachment
proteins (317); which, if any, of the above proteins correspond to these antibodies is unknown.
5. Dystrophin-related proteins
Mutations in the human dystrophin gene cause Duchenne muscular dystrophy. The dystrophin complex may
also help transmit sarcomere force to the plasma membrane, or help maintain membrane integrity under the
stress of sarcomere contraction (1031). The human dystrophin glycoprotein complex consists of dystrophin, dystrobrevin, distroglycan, sarcoglycan, sarcospan, syntrophin, and nitric oxide synthase (168). Given this close
association, all these genes will be considered together
here. The sea urchin dystrophin gene has been identified
and shares with vertebrates a complex structure with
multiple promoters and gene products (853, 1250). Immunohistochemistry and Western blot analysis show that
leech (Pondtobdella) has dystrophin, dystroglycan, sarcoglycan, syntrophin, and sarcospan homologs (1020, 1022).
C. elegans (113, 352, 382, 384) and Drosophila (243, 377,
853) have dystrophin, dystrobrevin, dystroglycan, sarcoglycan, and syntrophin (but not sarcospan) homologs.
The C. elegans and Drosophila dystrophin genes again
have multiple promoters and gene products (113, 853). C.
elegans muscle has a nitric oxide synthase binding protein
that can partially suppress dystrophin mutation (354).
The C. elegans proteins are also likely a complex, as
dystrophin (113), dystrobrevin (352, 355), dystroglycan
(384), sarcoglycan (384), and syntrophin (382, 384) mutants all have the same phenotype: hyperactivity, head
bending during forward locomotion, a tendency to hypercontract, and ACh hypersensitivity. Further genetic evidence of interaction is provided by data that increased
dystrobrevin expression delays muscle degeneration and
locomotor defects in dystrophin mutants (353). In vitro
binding assays give direct evidence of dystrophin, dystrobrevin, and syntrophin interaction and show that dystrophin/distrobrevin binding requires the second coiled-coil
dystrobrevin domain (351, 383). The basis of the mutant
phenotype is not well understood, but acetylcholinesterase activity is decreased (by an unknown mechanism) in
dystrophin mutants, which may explain the ACh hyperPhysiol Rev • VOL
1027
sensitivity (356). Ca2⫹ channel activity is important for
phenotype expression, as increased Ca2⫹ channel activity
increases, and reduced Ca2⫹ channel activity decreases,
muscle degeneration in dystrophin null mutants (742).
References 168, 186, 226, 999, 1066, and 1067 are reviews
of C. elegans and Drosophila as model systems for studying Duchenne muscular dystrophy.
6. Intermediate filaments
In vertebrates another protein that may help transmit
force from the sarcomere to the cell membrane, or help
maintain muscle cell integrity in the face of the stress
involved, is the intermediate filament protein desmin. In
mammals desmin is one of a large number of intermediate
filament types (others include keratin, neurofilament protein, and nuclear lamins) that can be ordered into five or
six classes on the basis of sequence homology (9, 319 are
primarily vertebrate reviews). Invertebrate intermediate
filaments are much less well understood, and their relationship to the mammalian classes remains unclear (256,
345, 530, 994, 996, 997, 1249, 1267, 1268). We cover here
only work directly involving invertebrate muscle intermediate filaments.
Early work showed surprising variation with respect
to the presence of intermediate filaments in invertebrate
muscles, even between closely related species. For instance, large amounts of intermediate filaments were
present in all muscle types (smooth, irregularly, and obliquely striated) in Ascaris, located throughout the sarcomere and around dense bodies, but not in C. elegans
muscle (80 – 82). A wide survey of major phylogenetic
groups showed that intermediate filaments were also
present in Acanthocephala, Echiura, and Chaetognatha
muscle, but absent from the other examined invertebrate
groups, including Urochordata and Cephalochordata (78,
81). It was subsequently shown that some invertebrate
intermediate filament proteins lack an epitope present in
all vertebrate intermediate filament proteins, and thus
that lack of muscle staining in some groups may be due to
the lack of this epitope rather than a lack of intermediate
filaments (995). Particularly in light of later work showing
muscle intermediate filaments are present in some of the
above “negative” groups (see below), the absence assignments made in this early work should be viewed with
caution.
Molecular genetic work has shown that Styela (a
urochordate) has two intermediate filament type genes,
one of which is expressed in smooth body wall muscle
and shows sequence similarity to desmin (997). Similarly,
one of the five intermediate filament protein genes in
Ciona (another urochordate) is expressed in muscle (535,
1249). Despite some confusion (994), it is now also clear
that in Branchiostoma (Cephalochordata) 1 of the 10
known (530) intermediate filament proteins is present in
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1028
muscle (997). With respect to nematodes, however, the
early data have held, with muscles from species from
multiple genera (Acanthocheilonema, Anisakis, Ascaris,
Dirifilaria, Onchocerca, Toxocara, Trichinella) containing intermediate filament proteins or showing staining
(116, 1046, 1065, 1333, 1334), but none of the four intermediate filament proteins (of a total of 11) in C. elegans
for which it has been investigated being present in muscle
(although the absence of some does cause displaced muscles and paralysis) (364, 398, 531–534). Staining is also
present in the muscles of several platyhelminths (Echinococcus, Schistosoma, Taenia) (1046).
Intermediate filaments composed of a protein immunologically similar to desmin are also present in several,
but not all, annelids (79, 81, 239). An intermediate filament
protein gene identified using a tissue sample containing
body muscle and attached epidermis has been sequenced
in Lumbricus terristris, but it is unclear if this gene
codes the above protein (126). Proteins that form 2- to
4-nm filaments, but which are not similar to desmin or
vimentin by two-dimensional gel electrophoresis or immunological cross-reactivity, are present in sea urchin
muscle (957). Arthropods appear to lack all intermediate
filaments (except nuclear lamins) (81), an absence confirmed by the lack of intermediate filament genes in the
Drosophila genome (365, 1026).
7. The 29-kDa ascidian protein
A body wall protein from Halocynthia roretzi of
unknown function that copurifies with thin filaments is
localized near the plasma membrane, has some similarity
to heat shock proteins and ␣-cystallin, and has been isolated and sequenced (1084, 1151).
G. Summary
The most striking impression from the above is the
large number and variety of different proteins, and different isoforms of individual proteins, present across invertebrates. This variety provides two great opportunities.
First, because these proteins interact and thus must have
evolved together, comparison of concerted changes in
their sequences should provide powerful evidence with
respect to evolutionary relationships. In addition to the
studies on actin mentioned earlier, this work has been
begun with sequence comparisons of myosin regulatory
and essential light chain and troponin C (187); troponin C,
myosin heavy chain, myosin regulatory and essential light
chain, and actin (903); vertebrate, C. elegans, and Drosophila titin, twitchin, projectin, and unc-89 (427, 554);
and vertebrate, ascidian, and Drosophila troponin I (407).
The first work is in agreement with the phyla and lower
assignments in Figures 1–3 but does not resolve the question of invertebrate monophyly. The second indicates that
Physiol Rev • VOL
skeletal and cardiac muscle type tissues evolved before
the vertebrate/arthropod split, but vertebrate smooth
muscle appears to have arisen independently of other
muscle types, and arthropod striated, urochordate
smooth, and vertebrate muscle (except for smooth) share
a common ancestry. The third suggests that titin/kettin,
twitchin, projectin, and myosin light chain kinases form a
related group of which unc-89 is not a member, but it was
performed with too few invertebrate species to provide
insight into invertebrate phylogenetic relationships. The
fourth shows that a single troponin I gene (from which
multiple isoforms are produced by alternative splicing),
such as is present in ascidia and Drosophila, is the ancestral condition, and the gene duplications leading to the
three troponin I genes present in vertebrates occurred
after the ascidian/vertebrate split. Although these early
observations are intriguing, much further work is necessary for the phylogenetic opportunities present here to be
fully realized. Notable lacks are investigation of muscle
proteins in Cnidaria, in which so far only actin, myosin,
and tropomyosin have been shown to exist, and detailed
study of giant muscle proteins in Lophotrochozoa.
The second opportunity this variety provides is in
understanding protein ensembles. Despite their variety,
the different sets of proteins present in the various invertebrates must successfully perform relatively similar
functions (form thin and thick filaments and sarcomeres,
contract and relax on demand). Investigating how the
multiplicity of forms noted above continue to do so, and
the relationship between these forms and differences in
muscle structure and function, will presumably provide
great insight into fundamental questions of protein interaction and function that can be successfully applied to
other systems. The utility of this approach is well demonstrated by a recent analysis of myosin heavy and light
chains, which showed that head, neck, and tail domains of
the heavy chains, and the essential and regulatory light
chains, show very similar phylogenetic trees (600). This
work is beginning in Drosophila, in which large numbers
of myofibril mutants are available (332, 815, 869, 1223).
Comparison of phenotypes of single gene heterozygotes
to mutants pairwise heterozygous in various muscle component genes has been used to investigate gene product
interactions and how critical correct regulation of gene
product amount is in myofibril development (89, 331, 441,
1223), and null mutants of actin and myosin have been
used to examine the contributions of thin and thick filaments to sarcomere assembly (89). Similar genetic opportunities are available in C. elegans, as is also the ability to
use antisense RNA to inhibit expressions of specific genes
(301). The increasing availability of cloned genes in ascidia should allow similar work to proceed in this “higher”
invertebrate (174). The development of robotic approaches for simultaneously identifying the components
85 • JULY 2005 •
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of multiprotein mixtures (proteomes) should dramatically
enhance these efforts (36).
It is critical to point out, however, that to date most
work has been performed in a small number of “model”
systems (Drosophila, C. elegans, Bivalvia, decapod crustacea), organisms heavily studied due to their disease or
economic importance (parasitic platyhelminths and nematodes), or organisms relatively closely related to vertebrates (Cephalochordata, Urochordata, Echinodermata).
Although this bias was understandable in the early days in
which the challenge was defining the sarcomeric muscle
protein complement, and protein and gene analysis and
sequencing tools were relatively primitive, neither justification exists today. These concerns about relatively small
numbers of studied species are deepened by the observation that Drosophila and C. elegans have apparently undergone extensive gene loss and rapid sequence divergence relative to the common ancestral metazoan (601).
Full achievement of the insights about invertebrate phylogeny and protein structure and function that the multiplicity of invertebrate muscle proteins allows can only be
achieved by overcoming this species parochialism and
examining a much larger range of the enormous number
of invertebrate species that exist.
This work would not have been possible without, and we express our
deep gratitude to, C. A. Barcroft, R. P. Harrison, R. R. Heck III, K. H.
Hobbs, S. McKean, B. Revill, J. Thuma, and R. Thuma for help in creating
and maintaining the reference database and to the Ohio University
library system and OHIOLINK electronic journal center for providing
several thousand journal article photocopies and PDFs.
This work was supported by grants from the National Institutes of
Health and the University of Cologne, Cologne, Germany (to S. L.
Hooper).
Address for reprint requests and other correspondence: S. L. Hooper,
Neuroscience Program, Dept. of Biological Sciences, Irvine Hall, Ohio
University, Athens, OH 45701 (E-mail: [email protected]).
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85 • JULY 2005 •
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Scott L. Hooper and Jeffrey B. Thuma
Physiol Rev 85:1001-1060, 2005. doi:10.1152/physrev.00019.2004
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on the following topics:
Oncology .. Protein Assembly
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Physiology .. Tropomyosin
Physiology .. Excitation-Contraction Coupling
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Physiol Rev 85: 1417, 2005;
doi:10.1152/physrev.00027.2005.
CORRIGENDA
Volume 85, July 2005
Pages 1001-1060: Hooper, Scott L., and Jeffrey B. Thuma. “Invertebrate Muscles:
Muscle Specific Genes and Proteins” (http://physrev.physiology.org/cgi/content/
full/85/3/1001). On p. 1006, we regret that Table 1 was published without the last
four entries. The complete Table 1 is listed here.
TABLE
1.
Actin and myosin information for relatively little studied groups
Group/Protein
Pterobranchia actin
Annelida actin
Gastropod actin
Species
Identification Method
Muscle Versus
Cytoplasmic
Isoforms?
Reference Nos.
Saccoglossus kowalevskii
Glycera, Nereis, species
unreported
Helobdella triserialis
Two cDNAs
Unknown
Unknown
Unknown
Unknown
127
658
One cDNA, apparent
family
Unknown
1269
Aplysia californica
Aplysia californica
One cDNA, 3–5 gene
family
One cDNA
Cloning of 5⬘-flanking
and part of coding
region of one gene
Many tissues
including
muscle, but not
segmental
ganglia
Unknown
cDNA clone
muscle specific
Unknown
Unknown
Unknown
Unknown
401
246
Unknown
Unknown
643
366
Unknown
Unknown
658
Three genes
Unknown
Two genes muscle
specific
1011
1308
Expressed sequence tag
One cDNA
Unknown
Unknown
Differentially
expressed in
different muscles
Unknown
Zebra mussel
Haliotis rufescens
Lophophorata, Brachiopoda
actin
Chaetognatha actin
Glottidea, unreported
species
Sagitta friderici
Paraspadella gotoi
Chelicerata actin
Various
Cephalochordata myosin
Urochordata myosin
Amphioxus (notochord)
Ascidia
Annelida myosin
Earthworm, leech
Next column
Chaetognatha myosin
Sagitta friderici
Unknown
Chelicerata myosin
Various
Unknown
www.prv.org
Muscle or
Developmental Stage
Specific Expression?
Unknown
Next column
Unknown
Only in
differentiating
embryonic muscle
cells
Two isoforms in
muscle specific
fashion
Present in body wall
muscle, specificity
unknown
Unknown
0031-9333/05 $18.00 Copyright © 2005 the American Physiological Society
247, 658,
1021, 1204
1124
23, 728, 729,
784, 857
160, 1009,
1013
1011
247, 426,
1021
1417

Invertebrate Muscles: Muscle Specific Genes and

Transcription

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