The pathway of myofibrillogenesis determines the interrelationship between myosin and paramyosin synthesis in Caenorhabditis elegans
Department of Cell Biology Box 3011, Duke University Medical
School, Durham, NC 27710, USA
* Present address: Department of Biology, University of North Carolina
Asheville, Asheville, NC 28804, USA
Author for correspondence (e-mail:
f.schachat{at}cellbio.duke.edu)
Accepted 17 March 2003
![]() |
Summary |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Key words: myosin B, myosin A, paramyosin, Caenorhabditis elegans, mutant, thick filament, myofibrillogenesis
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Brenner's initial description of C. elegans as a genetic system
(Brenner, 1974) revealed more
than 25 genetic complementation groups that affected C. elegans
movement. These included the genes for the two major proteins of body-wall
muscle thick filaments, paramyosin (unc-15) and the myosin B heavy
chain (unc-54) (Epstein et al.,
1974
; Waterston et al.,
1977
). Since then, screens for motility defects, suppressor
mutations and embryonic lethals have increased the number of genes implicated
in muscle assembly, organization and function several fold
(Francis and Waterston, 1991
;
Waterston, 1989a
;
Williams and Waterston, 1994
;
Zengel and Epstein, 1980a
).
Mutants in genes encoding major and minor structural components of the
myofilament lattice, as well as proteins that direct the organization and
assembly of C. elegans muscle, have been characterized and mapped
(Barral and Epstein, 1999
;
Benian et al., 1989
,
1999
;
Moerman and Fire, 1997
;
Ono and Benian, 1998
). The
existence of a diverse collection of motility-defective mutants, coupled with
the sequencing of the C. elegans genome
(Hodgkin et al., 1995
) and
extensive ultrastructural analysis of its striated muscle, led us to initiate
investigations on the interrelationship between myosin and paramyosin
synthesis and the underlying structural integrity of C. elegans thick
myofilaments.
C. elegans body-wall muscle thick filaments form bipolar
structures consisting of a core of paramyosin and filagenins
(Epstein et al., 1985;
Liu et al., 1998
;
Miller et al., 1986
;
Muller et al., 2001
) that
serves as a scaffold for the binding of two myosins: A and B. These myosins
are homodimeric with respect to their heavy chains (Schachat et al.,
1977
,
1978
) and are differentially
distributed on thick filaments. Myosin A represents approximately 20% of the
body-wall muscle myosin and is restricted to the central bipolar H-zone of
thick filaments, while myosin B composes approximately 80% of the body-wall
muscle myosin and is localized to the polar arms
(Miller et al., 1983
). Mutants
in the paramyosin and myosin B heavy chain genes impair movement and exhibit
aberrant thick filaments and A-band organization, although electron
microscopic observations show that the thin filament lattice is well-defined
and appears to be essentially intact
(Bejsovec and Anderson, 1988
;
Epstein et al., 1974
;
Mackenzie et al., 1978
;
MacLeod et al., 1977b
;
Waterston et al., 1977
).
Mutations in the myosin A gene (myo-3) result in embryonic lethality,
probably because myosin A is required for early events in thick filament
initiation and muscle cell elongation
(Waterston, 1989b
). In
addition, recent studies have provided insight into the interactions between
paramyosin and myosin that may direct assembly of the central and distal
regions of thick filaments. In particular, Muller et al.
(2001
) demonstrated that there
are significant segmental differences in both protein composition and
structure within the paramyosin-containing thick filament core, and Hoppe and
Waterston (2000
) characterized
a 322-amino-acid region within the rod portion of myosin A that is required
for its interaction with paramyosin.
Because molecular interactions among the components of the thick filament
are likely to dictate important steps in the assembly pathway, we investigated
the effects of eliminating the interactions between myosin B and paramyosin.
Previously, few studies of C. elegans have addressed the question of
whether mutations in one thick filament protein directly affect the expression
of others. Garcea et al.
(1978) found that a small
C-terminal deletion in myosin B does not affect the synthesis of myosin A,
even though it results in improperly assembled thick filaments and partial
paralysis; and paramyosin mutants did not appear to affect myosin accumulation
(Waterston et al., 1977
). But
whether other mutations, particularly those that eliminate the expression of
myosin B or paramyosin, have similarly benign effects on the synthesis of the
contractile proteins they interact with remains an open question.
Here, we report that a null mutation in myosin B has no discernible effect on paramyosin synthesis, while a null mutation in paramyosin specifically decreases myosin B synthesis. The non-reciprocal nature of these effects is consistent with an ordered pathway of thick filament assembly in which myosin B synthesis and incorporation into nascent thick filaments is dependent on the presence of a properly assembled, paramyosin-containing thick filament core.
![]() |
Materials and methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Radiolabeling and preparation of myofilament proteins
C. elegans were grown at 25°C on 8P NGM (nematode growth
media) plates (Schachat et al.,
1978) with a bacterial lawn of the E. coli strain OP50.
For accumulation studies, animals were harvested and transferred to NP NGM
plates and fed low specific activity (2.3x109 Bq
mmol1 l1) 35S-NA22-labeled
Escherichia coli for 72 h, as described previously
(Schachat et al., 1978
).
Homogenates were prepared using a French Press Cell at 54 500 kPa. Myofibrils
were prepared by low salt washes (50 mmol l1 NaCl, 10 mmol
l1 Tris, pH 7.4, 15 mmol l1
mercaptoethanol, 2 mmol l1 EGTA, 2 mmol l1
MgCl2, 0.1 mmol l1 diisopropyl fluorophosphate
and 10 µg ml1 each of the peptide protease inhibitors
leupeptin, antipain and chymostatin). Aliquots were saved from each step of
the myofibrillar preparation procedure. Previous studies had shown that
virtually all the soluble proteins are in the supernatant (G. White,
unpublished observations). The final pellet from a myofibrillar preparation
was diluted into sample buffer for electrophoresis. Serial dilutions of
samples were run on 8.0% sodium dodecyl sulfatepolyacrylamide
(SDSPAGE) gels using the Laemmli buffer system
(Laemmli, 1970
) to be certain
that exposures were in a linear range, and the resulting gels were dried and
exposed to Kodak X-ray film for 648 h. Digital images were captured
using an Adobe Photoshop-driven Howtek Scanmaster 3, and quantitative
densitometry was performed as described previously
(Thys et al., 1998
). Total
actin, which results primarily from unaffected tissue, was used as an internal
standard for normalization to obtain relative rates of accumulation and
synthesis. To estimate the fraction of actin that arises from non-muscle and
body-wall muscle, the actin-to-myosin protein ratio in squid transverse arm
muscle (1.8±0.1; N=3; Kier
and Schachat, 1992
), which exhibits thick filament dimensions and
sarcomere organization comparable to C. elegans
(van Leeuwen and Kier, 1977
;
Mackenzie and Epstein, 1981
),
was compared with the ratio in C. elegans wild-type 3rd instar larvae
(6.3±0.6; N=3). Calculation indicates that no more than 30% of
the total actin arises from C. elegans muscle. Subtracting the
contribution of pharyngeal muscle (20% of the total muscle), we estimate that
actin from sources other than body-wall muscle comprises at least 75% of the
total actin in wild-type C. elegans. Coupled with the electron
microscopic observations that a periodic and well-packed thin filament lattice
is present in CB1214 and unc-54 mutants
(Bejsovec and Anderson, 1988
;
Epstein et al., 1974
;
Waterston et al., 1977
,
1980
), we infer that total
actin does not vary significantly in the mutants studied and is a valid
standard for normalization.
Pulse labeling of nematodes
A mixed population of animals was harvested and treated with 1.25% NaOCl,
0.5 mol l1 NaOH for 10 min to produce eggs. Eggs were then
washed, plated and grown at 25°C. When animals entered larval stage 3, as
judged by examination of the developing gonad with Nomarski optics
(Sulston and Horvitz, 1977),
they were transferred to plates with high specific activity
(2.3x1010 Bq mmol1 l1)
35S-labeled bacteria for 2 h
(Schachat et al., 1977
;
Garcea et al., 1978
). Animals
were harvested, disrupted in the French Press Cell and analyzed by
SDSPAGE as previously described. For quantification, serial dilutions
of a minimum of three independent labeling experiments of each strain of
C. elegans were analyzed. Protein translation rates were normalized
to account for the different numbers of methionines and cysteines found in
each of the myofilament proteins analyzed.
Neville gel electrophoresis
To separate the two myosin heavy chain proteins A and B present in C.
elegans body-wall muscle, SDSPAGE gels were run as described by
Neville (1971). Myofilament
protein preparations, silver staining or autoradiography, and quantification
were performed as described previously
(Thys et al., 1998
).
RNA preparation and slot blot analysis
Total RNA was prepared from asynchronous populations of each strain
according to Austin and Kimble
(1989). RNA was loaded onto
nitrocellulose filters with duplicate loadings of 4 µg and 2 µg.
32P-labeled riboprobes were prepared from myosin B and myosin A
genomic fragments (Miller et al.,
1986
) and from an actin (act-3) probe
(Krause et al., 1989
) that
hybridizes with all C. elegans actin genes, which we subcloned into
pGEM3Z. All genomic fragments were kindly provided by Dr David Miller.
Hybridization was performed overnight at 55°C. Blots were washed to very
high stringency (Honda and Epstein,
1990
) and were exposed with intensifying screens at
80°C for 318 h. Quantitative densitometry was performed as
described above.
![]() |
Results |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
To determine whether a reciprocal reduction in myosin expression is observed in the absence of paramyosin, the paramyosin null mutant CB1214 was subjected to long-term pulse labeling as described above. Densitometric analysis showed that myosin accumulation was reduced by 28% compared with the wild type (Fig. 1A, lane c; Fig. 1B). These observations indicate that normal accumulation of myosin requires expression of wild-type paramyosin and the generation of properly assembled nascent thick filament cores.
Decreases in the rate of myosin synthesis account for the reduced
accumulation of myosin in the paramyosin and myosin B null mutants
To determine whether the reduced accumulation of myosin and paramyosin in
the myosin B and paramyosin null mutants results primarily from reduced
synthesis or from post-translational events such as increased proteolysis,
their rates of synthesis were compared to those of the wild type by pulse
labeling at high specific activity for 2 h. To ensure comparability of
short-term labeling in strains with different growth rates and to maximize the
incorporation of radiolabel, synchronous populations of eggs were generated
over a 3 h window and permitted to develop to larval stage 3, as assessed by
vulval development. After labeling, homogenates were prepared and analyzed by
SDSPAGE (Fig. 2A), and
the relative synthetic rates (normalized to actin) during the time interval
were measured by autoradiography and quantitative densitometry
(Fig. 2B). Comparison with the
wild type showed a decrease in the rate of myosin synthesis in the myosin B
null mutant CB190 (Fig. 2A,
lanes a and b), consistent with its inability to generate myosin B. By
contrast, the rate of paramyosin synthesis in CB190 was the same as in the
wild type, implying that the reduction in paramyosin accumulation in CB190
(Fig. 1) is a consequence of
increased degradation rather than impaired synthesis.
|
In the paramyosin null strain CB1214 (Fig. 2A, lane c), myosin synthesis was reduced by 33% (Fig. 2B). These data demonstrate that the 28% reduction in myosin accumulation measured in the paramyosin null mutant results from decreased myosin synthesis. Thus, the presence of wild-type paramyosin is essential for normal myosin synthesis.
Myosin B accumulation is differentially affected in the paramyosin
null mutant
The myosin present in C. elegans homogenates comes from several
sources. The body-wall muscle is the primary contributor, making up
approximately 75% of the total myosin, while the remainder arises primarily
from pharyngeal muscle. Because paramyosin mutants do not affect pharyngeal
muscle structure or function (Waterston et
al., 1977), we focused our attention on the two sarcomeric
myosins, A and B, that are expressed in C. elegans body-wall muscle.
These two myosins have non-overlapping functions, localize to different
regions of the thick filament and are involved in different stages of thick
filament assembly, and we sought to determine whether they were differentially
affected by the paramyosin mutations.
Using the Neville (1971)
gel electrophoresis system described by Karn et al.
(1985
) to resolve the
body-wall muscle myosin A and B heavy chains, we found that the paramyosin
null specifically interferes with accumulation of myosin B.
Fig. 3 demonstrates clearly
that accumulation of myosin B is reduced by almost 60%, while myosin A
accumulation is not affected. Because myosin B composes approximately 50% of
the total myosin, the 60% reduction accounts fully for the observed reductions
in total myosin accumulation (28±4%) and synthesis (33±3%) in
CB1214 (Figs 1,
2).
|
Myosin B mRNA levels are reduced in the paramyosin null mutant
To confirm the implications of the paramyosin null pulse-labeling studies
and to determine whether the reduction in myosin B synthesis is a consequence
of a reduced rate of translation or a reduction of mRNA levels, the levels of
myosin A, myosin B and actin mRNAs were compared by quantitative slot blots.
Myosin B mRNA was reduced by 52% in CB1214 compared with N2 controls, while
myosin A mRNA levels were affected only slightly, if at all
(Fig. 4). Thus, the
steady-state mRNA levels indicate that the reduced rate of myosin B synthesis
in paramyosin mutants is a direct consequence of lower steady-state levels of
myosin B mRNA.
|
Myosin B accumulation is also differentially affected by paramyosin
missense mutations
To determine whether mutations that produce defective paramyosins also
alter myosin B accumulation, three paramyosin missense mutants, CB73, HE2000
and CB1215, were analyzed. These mutants, selected on the basis of motility
defects, all generate paramyosins with a single amino acid substitution that
results in aberrant thick filament structure
(Brenner, 1974;
Mackenzie and Epstein, 1980
;
Waterston et al., 1977
;
Zengel and Epstein, 1980a
).
Neville gels (Fig. 5) show that
they all exhibit reduced ratios of myosin B to total body-wall muscle myosin.
So, defects in paramyosin that affect the integrity of thick filaments and
motility, as well as the total absence of paramyosin, affect the expression of
myosin B. These observations demonstrate that proper accumulation of myosin B
depends on its interaction with paramyosin and/or the integrity of the
paramyosin core of nascent thick filaments.
|
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Studies on CB675, a C-terminal deletion mutant of myosin B, demonstrated
that a defect in myosin B can severely affect mobility and thick filament
structure, but it does not necessarily affect either the synthesis or
accumulation of myosin A (Garcea et al.,
1978). Synthetic and accumulation studies on the myosin B null
mutant CB190 presented here extend those observations, showing that the
synthesis of myosin A as well as paramyosin is unaffected by the absence of
myosin B. However, the absence of myosin B affects the accumulation of myosin
A and paramyosin differently. Myosin A accumulation, like its synthesis, is
independent of myosin B, but the absence of myosin B results in increased
paramyosin degradation. Thus, the synthesis of myosin A and paramyosin is not
dependent on interactions with myosin B; but myosin B increases the stability
of paramyosin. The increased stability of paramyosin in the presence of myosin
B is probably due to the physical interaction between myosin B and the
paramyosin core, which stabilizes the core and/or reduces its exposure to
proteases.
To determine whether the synthesis of myosin was similarly independent of
its native interactions with paramyosin, myosin expression was analyzed in the
paramyosin null mutant CB1214. The original studies of CB1214 demonstrated
that paramyosin is essential for proper thick filament assembly and A-band
formation (Epstein et al.,
1974); without paramyosin, thick filaments are significantly
shorter and thicker than those of the wild type (Mackenzie and Epstein,
1980
,
1981
). Here, we show that the
absence of paramyosin and/or the consequent disruption of thick filament
structure reduces the accumulation of body-wall muscle myosin by approximately
30% a decrease that is substantial even when compared with the 50%
reduction in myosin accumulation when myosin B is completely absent in
CB190.
The reduced accumulation of myosin B in the paramyosin null does not result
from either a generalized defect in body-wall muscle development, as occurs in
myosin A or perlecan (Waterston,
1989b; Zengel and Epstein,
1980b
), or an increase in the degradation of improperly assembled
myosin, as occurs in dominant-negative myosin B mutations [unc-54(d)
mutations] or in mutations of the myosin chaperonin
(Barral and Epstein, 1999
;
Barral et al., 2002
). Rather,
the parallel decreases in myosin B accumulation, total myosin synthesis and
steady-state myosin mRNA levels reported here demonstrate that the reduced
myosin accumulation in the absence of paramyosin is due to a decrease in
myosin B synthesis. Given the specific nature of the interaction between
myosin B and paramyosin, the most likely explanation for the decrease in
myosin B synthesis is that the efficient translation of myosin B is dependent
on a nascent thick filament structure with which it or its
mRNAribonuclear protein complex can interact.
The stability of wild-type myosin B to proteolysis in the CB1214 background
is consistent with the studies of Bejsovec and Anderson
(1988), who reported that
wild-type myosin B was expressed at levels consistent with its gene dosage in
unc-54(d)/+ heterozygotes. The increased proteolytic susceptibility
of the dominant missense mutants of myosin B when compared with wild-type
myosin B and recessive mutants, such as CB675
(Garcea et al., 1978
),
probably reflects differences in their ability to interact productively with
the unc-45 myosin chaperonin
(Barral and Epstein, 1999
;
Barral et al., 2002
).
It is difficult to reconcile the observations on myosin and paramyosin null
mutations with a single mechanism for the regulation of contractile protein
synthesis. The finding that myosin A and paramyosin synthesis are independent
of the presence of myosin B is consistent with models in which the synthesis
of each of the thick filament proteins is entirely independent
(Saad et al., 1986). However,
the specific decrease in myosin B synthesis in the paramyosin null mutant is
inconsistent with such a mechanism. In the paramyosin null, myosin B synthesis
is clearly dependent on the presence of paramyosin. This observation is more
readily interpreted in terms of a modified `cotranslational assembly' model
(Isaacs and Fulton, 1987
), in
which synthesis of each thick filament protein is limited by its incorporation
into pre-existing thick filament assemblies.
The non-reciprocal effects of myosin B and paramyosin null mutations on
thick filament protein synthesis suggest that, if there is a single class of
explanation, it requires consideration of other aspects of the assembly
process. Our observations point directly to the pathway that Epstein,
Waterston and colleagues have presented for C. elegans thick filament
assembly. It holds that the central bipolar region containing myosin A forms
first, nucleating the polar elongation of the paramyosin core, which is then
followed by the addition of myosin B to the polar arms (Epstein et al.,
1985,
1986
;
Waterston, 1989b
). This
pathway, combined with the structural and expression defects of mutations, is
illustrated in Fig. 6. Our
studies, in conjunction with those of Garcea et al.
(1978
) and the observation
that mutations in myosin A result in an embryonic lethal phenotype
(Waterston, 1989b
), suggest
that the effects of myosin B and paramyosin null mutations on thick filament
protein synthesis are a consequence of this ordered assembly pathway. Myosin A
defects that prevent the generation of a bipolar central region of the
filament preclude all downstream steps in contractile protein synthesis and
myofilament assembly; the paramyosin null mutant impacts only the later
synthesis of myosin B, which interacts with a stable paramyosin core during
thick filament elongation; and the myosin B null mutant does not affect the
synthesis of either myosin A or paramyosin, because formation of the nascent
thick filament bipolar core precedes and is independent of interactions with
myosin B.
|
![]() |
Acknowledgments |
---|
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Austin, J. and Kimble, J. (1989). Transcript analysis of glp-1 and lin-12, homologous genes required for cell interactions during development of C. elegans.Cell 58,565 -571.[Medline]
Barral, J. M. and Epstein, H. F. (1999). Protein machines and self assembly in muscle organization. BioEssays 21,813 -823.[CrossRef][Medline]
Barral, J. M., Hutagalung, A. H., Brinker, A., Hartl, F. U. and
Epstein, H. F. (2002). Role of the myosin assembly protein
UNC-45 as a molecular chaperone for myosin. Science
295,669
-671.
Bejsovec, A. and Anderson, P. (1988). Myosin heavy-chain mutations that disrupt Caenorhabditis elegans thick filament assembly. Genes Dev. 2,1307 -1317.[Abstract]
Benian, G. M., Ayme-Southgate, A. and Tinley, T. L. (1999). The genetics and molecular biology of the titin/connectin-like proteins of invertebrates. Rev. Physiol. Biochem. Pharmacol. 138,235 -268.[Medline]
Benian, G. M., Kiff, J. E., Neckelmann, N., Moerman, D. G. and Waterston, R. H. (1989). Sequence of an unusually large protein implicated in regulation of myosin activity in C. elegans.Nature 342,45 -50.[CrossRef][Medline]
Brenner, S. (1974). The genetics of
Caenorhabditis elegans. Genetics
77, 71-94.
Epstein, H. F., Miller, D. M., Ortiz, I. and Berliner, G. C. (1985). Myosin and paramyosin are organized about a newly identified core structure. J. Cell Biol. 100,904 -915.[Abstract]
Epstein, H. F., Ortiz, I. and Mackinnon, L. A. (1986). The alteration of myosin isoform compartmentation in specific mutants of Caenorhabditis elegans. J. Cell Biol. 103,985 -993.[Abstract]
Epstein, H. F., Waterston, R. H. and Brenner, S. (1974). A mutant affecting the heavy chain of myosin in Caenorhabditis elegans. J. Mol. Biol. 90,291 -300.[Medline]
Francis, R. and Waterston, R. H. (1991). Muscle cell attachment in Caenorhabditis elegans. J. Cell Biol. 114,465 -479.[Abstract]
Garcea, R. L., Schachat, F. H. and Epstein, H. F. (1978). Coordinate synthesis of two myosins in wild-type and mutant nematode muscles during larval development. Cell 15,421 -428.[Medline]
Hodgkin, J., Plasterk, R. H. and Waterston, R. H. (1995). The nematode Caenorhabditis elegans and its genome. Science 270,410 -414.[Abstract]
Honda, S. and Epstein, H. F. (1990). Modulation of muscle gene expression in Caenorhabditis elegans: differential levels of transcripts, mRNAs, and polypeptides for thick filament proteins during nematode development. Proc. Natl. Acad. Sci. USA 87,876 -880.[Abstract]
Hoppe, P. E. and Waterston, R. H. (2000). A
region of the myosin rod important for interaction with paramyosin in
Caenorhabditis elegans striated muscle.
Genetics 156,631
-643.
Isaacs, W. B. and Fulton, A. B. (1987). Cotranslational assembly of myosin heavy chain in developing cultured skeletal muscle. Proc. Natl. Acad. Sci. USA 84,6174 -6178.[Abstract]
Karn, J., Dibbs, N. and Miller, D. M. (1985). Cloning nematode myosin genes. In Cell and Muscle Motility VI (ed. J. Shays), pp. 185-238.New York: Plenum.
Kier, W. M. and Schachat, F. H. (1992). Biochemical comparison of fast- and slow-contracting squid muscle. J. Exp. Zool. 168,41 -56.
Krause, M., Wild, M., Rosenzweig, B. and Hirsh, D. (1989). Wild-type and mutant actin genes in Caenorhabditis elegans. J. Mol. Biol. 208,381 -392.[Medline]
Laemmli, U. K. (1970). Cleavage of structural proteins during assembly of the head of Bacteriophage T4. Nature 227,680 -685.[Medline]
Liu, F., Bauer, C. C., Ortiz, I., Cook, R. G., Schmid, M. F. and
Epstein, H. F. (1998). ß-Filagenin, a newly identified
protein coassembling with myosin and paramyosin in Caenorhabditis elegans.J. Cell Biol. 140,347
-353.
Mackenzie, J. M., Jr and Epstein, H. F. (1980). Paramyosin is necessary for determination of nematode thick filament length in vivo. Cell 22,747 -755.[CrossRef][Medline]
Mackenzie, J. M., Jr and Epstein, H. F. (1981). Electron microscopy of nematode thick filaments. J. Ultrastruct. Res. 76,277 -285.[Medline]
Mackenzie, J., Schachat, F. H. and Epstein, H. E. (1978). Immunocytochemical localization of two myosins within the same muscle cells in Caenorhabditis elegans. Cell 15,413 -419.[Medline]
MacLeod, A. R., Waterston, R. H. and Brenner, S. (1977a). An internal deletion mutant of a myosin heavy chain in Caenorhabditis elegans. Proc. Natl. Acad. Sci. USA 74,5336 -5340.[Abstract]
MacLeod, A. R., Waterston, R. H., Fishpool, R. M. and Brenner, S. (1977b). Identification of the structural gene for a myosin heavy-chain in Caenorhabditis elegans. J. Mol. Biol. 114,133 -140.[Medline]
Miller, D. M., III, Ortiz, I., Berliner, G. C. and Epstein, H. F. (1983). Differential localization of two myosins with nematode thick filaments. J. Cell Biol. 34,477 -490.
Miller, D. M., Stockdale, F. E. and Karn, J. (1986). Immunological identification of the genes encoding the four myosin heavy chain isoforms of Caenorhabditis elegans. Proc. Natl. Acad. Sci. USA 83,2305 -2309.[Abstract]
Moerman, D. G. and Fire, A. (1997). Muscle: structure, function, and development. In C. Elegans II (ed. D. L. Riddle, T. Blumenthal, B. J. Meyer and J. R. Priess), pp.417 -470.Cold Spring Harbor, New York: Cold Spring Harbor Press.
Muller, S. A., Haner, M., Ortiz, I., Aebi, U. and Epstein, H. F. (2001). STEM analysis of Caenorhabditis elegans muscle thick filaments: evidence for microdifferentiated substructures. J. Mol. Biol. 305,1035 -1044.[CrossRef][Medline]
Neville, D. M., Jr (1971). Molecular weight
determination of protein-dodecyl sulfate complexes by gel electrophoresis in a
discontinuous buffer system. J. Biol. Chem.
246,6328
-6334.
Ono, S. and Benian, G. M. (1998). Two
Caenorhabditis elegans actin depolymerizing factor/cofilin proteins,
encoded by the unc-60 gene, differentially regulate actin filament
dynamics. J. Biol. Chem.
273,3778
-3783.
Saad, A. D., Pardee, J. D. and Fischman, D. A. (1986). Dynamic exchange of myosin molecules between thick filaments. Proc. Natl. Acad. Sci. USA 83,9483 -9487.[Abstract]
Schachat, F., Garcea, R. L. and Epstein, H. F. (1978). Myosins exist as homodimers of heavy chains: demonstration with specific antibody purified by nematode mutant myosin affinity chromatography. Cell 10,405 -411.
Schachat, F. H., Harris, H. E. and Epstein, H. F. (1977). Two homogeneous myosin in body-wall muscle of Caenorhabditis elegans. Cell 10,721 -728.[CrossRef][Medline]
Sulston, J. E. and Horvitz, H. R. (1977). Post-embryonic cell lineages of the nematode, Caenorhabditis elegans.Dev. Biol. 56,110 -156.[Medline]
Thys, T. M., Blank, J. M. and Schachat, F. H.
(1998). Rostral-caudal variation in troponin T and parvalbumin
correlates with differences in relaxation rates of cod axial muscle.
J. Exp. Biol. 201,2993
-3001.
van Leeuwen, J. and Kier, W. M. (1977). Functional design of tentacles in squid: linking sarcomre ultrastructure to gross morphological dynamics. Phil. Trans. R. Soc. Lond. B 352,551 -571.
Waterston, R. H. (1989a). Molecular genetic approaches to the study of motility in Caenorhabditis elegans. Cell Motil. Cytoskeleton 14,136 -145.[Medline]
Waterston, R. H. (1989b). The minor myosin heavy chain, mhcA, of Caenorhabditis elegans is necessary for the initiation of thick filament assembly. EMBO J. 8,3429 -3436.[Abstract]
Waterston, R. H., Fishpool, R. M. and Brenner, S. (1977). Mutants affecting paramyosin in Caenorhabditis elegans. J. Mol. Biol. 117,679 -697.[Medline]
Waterston, R. H., Thomson, J. N. and Brenner, S. (1980). Mutants with altered muscle structure of Caenorhabditis elegans. Dev. Biol. 77,271 -302.[Medline]
Williams, B. D. and Waterston, R. H. (1994). Genes critical for muscle development and function in Caenorhabditis elegans identified through lethal mutations. J. Cell Biol. 124,475 -490.[Abstract]
Zengel, J. M. and Epstein, H. F. (1980a). Identification of genetic elements associated with muscle structure in the nematode Caenorhabditis elegans. Cell Motil. 1, 73-97.[Medline]
Zengel, J. M. and Epstein, H. F. (1980b). Mutants altering coordinate synthesis of specific myosins during nematode muscle development. Proc. Natl. Acad. Sci. USA 77,852 -856.[Abstract]