1 Department of Anatomy and Neurobiology, Washington University Medical School,
St. Louis, MO 63110, USA
2 Department of Molecular and Cellular Biology, Harvard University, Cambridge,
MA 02138, USA
3 Biology I, University of Freiburg, 79104 Freiburg, Germany
Author for correspondence (e-mail:
sanesj{at}mcb.harvard.edu)
Accepted 12 August 2005
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SUMMARY |
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Key words: Acetylcholine receptor, Muscle, Neuromuscular junction, Zebrafish
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Introduction |
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Despite this impressive evidence, recent observations have raised questions
about the `synaptic organizer hypothesis' in general and the `agrin
hypothesis' (McMahan, 1990) in
particular. In mice genetically engineered to lack either DNA topoisomerase or
the HB9 transcription factor, the phrenic nerve fails to form and the
diaphragm muscle develops aneurally. Nonetheless, AChR clusters form on
myotubes in these muscles, and most reside in the central `end-plate' band
through which the nerve usually grows
(Yang et al., 2000
;
Yang et al., 2001
;
Lin et al., 2001
). Likewise,
AChR clusters form initially in agrin mutant mice, although they subsequently
disperse (Yang et al., 2001
;
Lin et al., 2001
;
Misgeld et al., 2005
).
At least three explanations can be envisioned for these results. One is that the aneural AChR clusters in vivo are akin to `hot spots' on cultured myotubes; growing axons use agrin to organize new postsynaptic sites and disperse ectopic clusters in vivo as they do in vitro. Second, clustering observed in topoisomerase, HB9 or agrin mutants might be a compensatory response to failed innervation rather than part of a normal developmental progression. Both of these explanations are consistent with the organizer and agrin hypotheses. Less consistent is a third alternative, that the initially aneural AChR clusters might be recognized by axons and incorporated into synapses. In this case, the role of agrin could be to stabilize clusters or to organize subsequent steps in synaptic maturation.
Distinguishing among these alternatives requires analysis of the earliest
steps in NMJ formation. Such analysis is difficult in mammals because live
imaging of muscles is infeasible at the embryonic stages when NMJs form, and
because muscles are innervated by numerous axons that form NMJs
asynchronously. We therefore used zebrafish embryos because their transparency
allows in vivo imaging and their myotomal muscles are initially innervated by
individual, identified axons. We show that, contrary to the predictions of the
organizer and agrin hypotheses, motor axons form some NMJs, particularly those
on early-born, slow adaxial muscle fibers
(Devoto et al., 1996;
Blagden et al., 1997
;
Barresi et al., 2000
;
Stickney et al., 2000
), by
incorporating pre-existing postsynaptic structures, and that axonal outgrowth
is guided by components associated with pre-existing clusters. Postsynaptic
differentiation is nonetheless nerve dependent, in that axons are crucial for
the maturation and maintenance of the postsynaptic apparatus and may be
required for the formation of AChR clusters on later-born, fast muscle
fibers.
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Materials and methods |
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Transgenic lines
To label motor axons, we generated transgenic lines in which the expression
of GFP or a membrane-associated (farnesylated) derivative of GFP (mGFP) was
regulated by genomic elements from the zebrafish hb9 gene. A PAC
clone containing the genomic hb9 locus (BUSMP706I09178Q2) was
isolated by screening the gridded PAC-library 706 (RZPD, Berlin, Germany) with
a radioactively labeled hb9 cDNA probe (AY445044). Restriction
analysis and Southern blotting revealed that the hb9-coding region,
together with 3 kb of 5' sequence, were present on a 5 kb
EcoRI-fragment. This fragment was subcloned, then the regulatory
sequences were amplified by PCR and inserted upstream of cDNAs encoding GFP or
mGFP (GAPGFP4) (Moriyoshi et al.,
1996). The plasmid DNAs were digested with EcoRI and
Acc651, purified, and injected into one-cell embryos (20 pg/embryo).
Resulting transgene-positive adults were mated, and embryos were inspected by
fluorescence microscopy, revealing germ-line transmission for seven hb9:GFP
and two hb9:mGFP founders. Motoneurons were labeled in all lines. For each
construct, the line with the strongest expression of GFP was selected for
study. Further description of the regulatory elements and of expression in
non-neural cells will be presented elsewhere (D.M., unpublished).
Histochemistry
For histological analysis, dechorionated embryos were fixed at 4°C for
4-6 hours with 4% paraformaldehyde, and rinsed in water. Fixed embryos were
then stained with znp-1 (Zebrafish International Resources Center, University
of Oregon, Eugene, OR), fluorophore-conjugated -bungarotoxin (BTX;
Molecular Probes) and anti-myosin (F59; Developmental Studies Hybridoma Bank,
University Iowa, Iowa City, IA), or fluorophore-conjugated phalloidin
(Molecular Probes), followed by appropriate fluorophore-conjugated secondary
antibodies. In some cases, embryos were stained live with BTX (see below)
before fixation. Animals older than 24 hpf were permeabilized by immersion in
acetone for 7 minutes at 20°C, after fixation and before
immunostaining.
Some stained embryos were sectioned in a cryostat and counterstained with Neurotrace (Molecular Probes), but most were deyolked and mounted on slides in glycerol. Initial morphometric analysis was performed on a Zeiss Axioplan2 fluorescent microscope. Confocal images were acquired using 20x (NA0.7), 40x (NA1.15) and 60x (NA1.2) long-working distance water objectives, or a 60x (NA1.45) oil objective. Stacks of optical sections were collected in the z dimension. The step size, based on the calculated optimum for each objective, was between 0.25 and 0.5 µm. Subsequently, each stack was collapsed into a single image (maximum intensity or z-projection), and additionally rendered to provide views from multiple angles. Analysis was performed offline using the Metamorph Universal Imaging software.
Time-lapse imaging
Embryos were staged, dechorionated, and incubated at room temperature for
45 minutes in L-15 medium containing 5% DMSO and 2.5 µM BTX (Molecular
Probes). Embryos were then rinsed in L-15 and mounted on glass coverslips with
1% low-melting temperature agarose in embryo medium containing 0.003% PTU and
0.02% (w/v) tricaine (MS-222). A coverslip was inverted onto a central-well
organ culture dish (Falcon #35-3037) containing medium/PTU/tricaine, and the
dish was placed into a temperature-controlled stage (Cell Microtemp Systems;
28-30°C) and mounted on a confocal microscope. Stacks of optical sections
were collected in the z dimension at intervals of 2-5 minutes for a
period of 3-8 hours. Initial analysis used collapsed images, but for scoring
axon-myotube contacts, we viewed expanded stacks plane by plane. Step size and
processing were as described above.
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Results |
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Labeling primary motoneurons with GFP
To image CaP, we sought to label it with Green Fluorescent Protein (GFP).
Unfortunately, primary motoneurons are not labeled in previously described
transgenic zebrafish with strong neuron-specific expression of GFP
(Higashijima et al., 2000;
Park et al., 2000
;
Udvadia, 2001
). We therefore
generated new lines in which GFP was linked to regulatory elements of the
zebrafish hb9 gene (see Materials and methods). HB9 is a
transcription factor that is expressed in motoneurons and required for early
stages of their development (Jessell,
2000
). Primary motor axons were intensely labeled in HB9:GFP fish
(Fig. 1E). CaPs were labeled by
14 hpf, before their axons extended in the myotomes
(Fig. 1F). Between 14 and 24
hpf, GFP labeled CaP axons at least as effectively as a well-characterized
neuron-specific antibody, znp-1
(Trevarrow, 1990
)
(Fig. 1). We conclude that the
HB9:GFP line is suitable for following the earliest steps in NMJ
formation.
|
By 14 hpf (in rostral segments), the somata of primary motoneurons
were prominent in the ventral portion of the neural tube
(Fig. 1F), and the first
adaxial myotubes had formed in the myotomes (see below). Over the next hour,
CaP initiated axonogenesis by extending filopodia out of the neural tube (see
below); in parallel, additional myotubes began to form adjacent to the pioneer
cells. Following the filopodia, a broad foot-shaped axon extended, roughly
perpendicular to the long axis of the myotubes and midway between their ends
(Fig. 1G). Initially thick and
uniform in diameter, the axon became variable in diameter by 17 hpf
(Fig. 1H), and then broke into
large varicosities separated by thin segments (19 hpf,
Fig. 1I). Over the next few
hours, small varicosities formed between the larger ones (22 hpf,
Fig. 1J). The smallest
varicosities enlarged rapidly, so that varicosities were more (though not
completely) uniform in size and shape by 24 hpf
(Fig. 1K). This stereotyped
sequence of steps is summarized in Fig.
1L.
AChR clustering precedes axon-myotube contact
To ascertain the relationship of AChR clusters to growing motor axons, we
incubated live embryos with fluorophore-conjugated -bungarotoxin (BTX),
which binds specifically to AChRs, then visualized axons with anti-znp-1 or
GFP, as above. No clusters were detected at 14 hpf, before primary motoneurons
extended axons (Fig. 2A). By 16
hpf, however, AChR clusters were present on myotubes at the medial edge of the
myotome (Fig. 2B). As new
myotubes formed, AChRs clustered on them, in advance of the extending axons
(Fig. 2C). All AChR clusters on
rostral segments were covered by axons by 20 hpf
(Fig. 2D). Thus, AChRs cluster
on myotubes before they are innervated.
Myotomes form in a rostral-to-caudal progression. Although the sequence of steps in NMJ formation was generally similar among segments, the degree to which AChR clustering preceded axon extension varied systematically along the rostrocaudal axis (Fig. 2E-I). In rostral segments, AChR clusters were initially present on only the few myotubes directly ahead of the advancing axon (Fig. 2B,C), and axons eventually extended beyond the clusters (Fig. 2D,E,I), consistent with the possibility that axons secrete (or their filopodia contain) an AChR-clustering agent. By contrast, a complete band of AChR clusters formed prior to axon outgrowth in caudal segments of the trunk (Fig. 2F). This difference appears to reflect a position-dependent delay between motoneuron and myotube development. In rostral segments, the first myotubes formed as the CaP axon exited the neural tube; AChRs clustered shortly after myotubes form, and the clusters were rapidly innervated by the growing CaP axon. In more caudal segments, however, myotubes had formed, and AChRs clustered, across the full width of the myotome before the CaP axon exited the neural tube. This striking mismatch provides additional evidence that AChR clustering is initiated without axonal contact. Moreover, the presence of a full band of clusters in caudal segments does not support the possibility that postsynaptic differentiation is initiated by a mid-range cue emitted from growth cones as they extend.
Aneural AChR clusters are incorporated into NMJs
The observations presented so far suggested that axons incorporate
initially aneural AChR clusters into NMJs. Before accepting this conclusion,
however, we needed to consider two other possibilities. First, clusters that
were aneural at the time of observation might have been transiently innervated
at an earlier stage. Second, aneural clusters might have dispersed as axons
extended, and been replaced by new, innervated clusters. To assess these
alternatives, we imaged NMJ formation using HB9:GFP fish in which AChRs had
been labeled with BTX. To ensure that fine filopodial processes did not go
undetected, we generated an additional transgenic line that incorporated a
membrane-associated (farnesylated) derivative of GFP (HB9:mGFP). Patterns of
expression in HB9:mGFP fish were similar to those described above for HB9:GFP
fish, but the membrane-targeted derivative provided superior labeling of
filopodia (Fig. 3). Stacks of
optical sections were collected at intervals of 2-5 minutes for a period of
4-6 hours, beginning 16-18 hpf. Segments 7-12 were filmed, because more
rostral segments were occluded by yolk.
AChR clusters were present before any filopodia or other processes extended from GFP- or mGFP-labeled CaP axons. In the segment shown in Fig. 3A, for example, clusters were present from the initial view at 17 hpf, but no processes extended from CaP until nearly 19 hpf. (The time of initial CaP extension is later here than in Fig. 1 because it is a more caudal segment; see Fig. 2E.) At this point, a slender filopodium extended from the neuronal somata toward the myotome. Thereafter, numerous additional filopodia extended in multiple directions, many of which subsequently retracted. The filopodia were numerous (up to 20 per neuron at a single time), long (up to 100 µm from the axon shaft to the filopodial tip), and irregular in shape. They were highly dynamic, extending or retracting up to 65 µm in 2 minutes (the time between frames). AChR clusters were present for up to 2 hours prior to the extension of filopodia, and were observed in all 26 segments (from 16 fish) imaged starting at 16-17 hpf.
As filopodia and lamellopodia extended, some contacted previously aneural AChR clusters (Fig. 3A,B). A subset of these contacts remained stable for the remainder of the viewing period, during which time other filopodia extended, contacted additional AChR clusters, or remodeled. Because we collected images at 2-5 minute intervals, and we never saw initially distinct AChR clusters completely disperse in <15 minutes, we can rule out the possibility that aneural clusters were replaced by distinct, nerve-induced clusters between views. Thus, many AChR clusters form aneurally and at least some NMJs are formed by the incorporation of these initially aneural clusters.
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|
A second, related line of evidence for the recognition of cluster-associated components by axons was that processes contacting an AChR cluster frequently became stably associated with it. To quantify the selectivity of this stabilization, we analyzed all processes that extended from one CaP axon in each of seven fish. Each process was scored for whether it contacted a cluster of AChRs, and whether it persisted to the end of the imaging period. All of the processes (n=102) that failed to contact an AChR cluster retracted, whereas most of the processes (63/71) that contacted an AChR cluster remained associated with it (Fig. 4B).
|
The sequence shown in Fig. 3B illustrates several of the phenomena that appear to reflect the local guidance of axons. A filopodium initially contacted a cluster lying rostral to the central band, forming a contact that precisely matched the shape of the cluster. Following initial contact, filopodial activity became focused in its direction. Additional clusters were not encountered, however. Subsequently, the filopodium in contact with the AChR cluster retracted. Following this loss of axonal contact, multiple filopodia extended in all directions, a few of which contacted a proximal AChR cluster in the central band. Filopodia extending beyond this contacted cluster did encounter additional clusters, and the axon extended along the end-plate band. Thus, CaP processes became stably associated with AChR clusters, and the association preceded the reorientation of filopodial activity and axonal extension. Together, these observations suggest that axonal growth is influenced by components associated with AChR clusters.
Distinct AChR clustering patterns on slow and fast muscle fibers
Myotomal muscles contain two types of muscle fiber that differ in
developmental origin and molecular composition. The first-born fibers, called
adaxial, form from myoblasts near the notochord. They contain slow myosin, and
initially form a monolayer on the medial surface of the somite. Most of the
adaxial fibers then migrate radially to the lateral edge of the myotome. The
few adaxial fibers nearest the horizontal midline (2-6 per segment) do not
migrate and remain near the notochord; they have been called `muscle
pioneers'. As migration occurs, myoblasts of a second population (lateral
presomitic cells) fuse in the space between the notochord and the adaxial
cells, giving rise to fast muscle fibers
(Devoto et al., 1996;
Blagden et al., 1997
;
Barresi et al., 2000
;
Stickney et al., 2000
). These
events, like the other aspects of neuromuscular development described above,
occur in a rostrocaudal sequence (Fig.
5A'-D').
|
After forming NMJs on muscle pioneers, the CaP axon extended past them. CaP remained on the medial surface of the myotome, and thereby contacted myoblasts of the later-born population, which were fusing to form fast muscle fibers (Fig. 5E,G; see also Fig. 2D). New AChR clusters formed beneath the advancing axon, usually a few myotubes behind the tip of the growth cone (Fig. 5F,G; see also Fig. 2I). Clusters on the fast fibers were generally smaller than those on pioneers (Fig. 5G). Thus, AChR clusters formed on adaxial cells before CaP had contacted them but formed on fast fibers following axonal contact.
Once it reached the distal (ventral) edge of the myotome, CaP had contacted
all fibers on the medial surface, and had initiated the formation of NMJs on
them. Observations reported here are restricted to these NMJs. Subsequently,
during the second day of embryogenesis, CaP, and the secondary motor axons
that followed it, branched and turned to innervate fibers deeper in the
myotome (Beattie, 2000).
Formation of precise synaptic appositions
Apposition of the CaP axon to AChR clusters was initially imprecise, in
that some clusters were only partially covered by axonal processes, and some
axonal segments extended beyond the borders of clusters (e.g.
Fig. 2D-I,
Fig. 6A,B). Apposition between
axonal varicosities and AChR clusters became precise by 22-24 hpf in rostral
segments. At this time, each axonal varicosity was centered over a myotube,
each myotube bore a single AChR cluster, and each axonal varicosity was
precisely aligned with the cluster (Fig.
6C-E).
Morphometric analysis of confocal images revealed five processes that contributed to the matching of axonal varicosities to AChR clusters. First, CaP axons became apposed to AChR clusters in the center of myotubes as they extended (Fig. 6F, dotted line). Second, aneural clusters lateral to the central endplate band (asterisks in Fig. 2B-D) were lost after the initial cluster had been contacted (Fig. 6F, solid line). Third, clusters on adaxial fibers other than the pioneers were lost as those fibers migrated laterally and became displaced from CaP (Figs 2E). Fourth, protrusions from the main axonal shaft that were not in contact with clusters were lost (Fig. 2C, Fig. 6A). Finally, clusters were often initially larger than even the largest varicosities that overlay them, but they decreased in size over a 3-5 hour period to match the size of the varicosities (Fig. 6G). The fact that matching occurred by shrinkage of the AChR cluster rather than by expansion of the varicosity suggests that the nerve terminal influences postsynaptic morphology.
Axons remodel and stabilize AChR clusters
Although AChR clusters form aneurally, axons might affect their fate. An
opportunity to test this idea was provided by a fortuitous observation on
HB9:mGFP transgenic fish. As expected, motor axons were unlabeled in
approximately one-quarter of the offspring from matings between HB9:mGFP
heterozygotes, and were labeled in approximately three-quarters; fluorescence
was clearly brighter in approximately one-third of the mGFP-positive fish than
it was in the remaining two-thirds, indicating that the brighter third were
homozygous. Primary motor axons, including the CaP axons, failed to extend in
70% of the segments of such putative homozygotes (called
HB9:mGFP+/+; Fig.
7A). Time-lapse imaging of HB9:mGFP+/+ fish from 14-22
hpf failed to reveal any axonal or filopodial extension in segments that were
still axon-free at 22 hpf, whereas normal axon extension occurred in adjacent
segments (data not shown). Thus, axon-free myotomal segments were aneural (not
innervated) rather than being transiently innervated (denervated). The failure
of extension reflected a toxic effect of high levels of mGFP rather than a
lost function of a gene interrupted by transgene insertion, because
attenuating mGFP expression by injecting a morpholino oligonucleotide
complementary to sequences in the transgene restored axon formation in
HB9:mGFP+/+ fish (data not shown). Thus, overexpression of mGFP in
HB9:mGFP+/+ fish led to cell-autonomous blockade of axon
outgrowth.
|
First, between 19 and 22 hpf, the number of clusters increased >2-fold in segments that were not innervated but changed little in innervated segments (Fig. 7F). The lack of a net change in innervated segments reflects a balance between the formation of NMJs on newly formed myotubes, and the loss of aneural clusters from innervated myotubes (`side clusters', Fig. 6F) and from adaxial myotubes migrating laterally. Staining with anti-myosin or the actin-binding toxin phalloidin revealed no difference in myotube shape or number between innervated segments and segments that were not innervated (data not shown). Accordingly, the presence of supernumerary clusters in segments that were not innervated suggests that the dispersal of aneural clusters in innervated myotubes requires the presence of an axon.
Second, between 22 hpf and 24 hpf, the number of clusters increased in innervated segments, reflecting the formation of additional myotubes, but decreased by >80% in segments that were not innervated (Fig. 7C,D,F). By 26 hpf, most of the segments that lacked axons were completely devoid of AChR clusters (data not shown). Thus, although AChR clusters form aneurally, their maintenance requires innervation.
Third, between 22 hpf and 24hpf, AChR cluster size decreased less in segments that were not innervated than in innervated segments or in control fish (Figs 7G). This result supports the hypothesis proposed above, that the generation of a perfect apposition between pre- and postsynaptic specializations involves axon-dependent remodeling of the AChR cluster to match the outline of the axonal varicosity.
Taken together, these results suggest that innervation is essential for the remodeling and maintenance of AChR clusters. Observations at a later stage provide evidence against an alternative interpretation, that a lack of innervation leads to defects in the ability of the myotube to participate in synaptogenesis. Although primary axon outgrowth was defective in HB9:mGFP+/+ fish, secondary axons grew into most segments, including those that were initially not innervated. The secondary axons formed NMJs in which terminal varicosities were apposed to AChR clusters (Fig. 7E); in some cases, these fish survived for at least 2 additional days. This result indicates that segments that were not innervated remained competent to form and maintain AChR clusters and NMJs.
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Discussion |
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First, AChR clusters form on myotubes that have not yet been contacted by
axons, and at least some NMJs form by the incorporation of these pre-existing
AChR clusters. This view differs from the classical one, that growing axons
organize AChR clusters at sites of contact with myotubes, and disperse aneural
clusters that they do not contact (Anderson
and Cohen, 1978; Frank and
Fischbach, 1979
;
Ziskind-Conhaim et al., 1984
;
McMahan, 1990
;
Sanes and Lichtman, 1999
). It
is consistent with more recent studies showing that AChR clusters form in
aneural mouse muscles (Yang et al.,
2000
; Yang et al.,
2001
; Lin et al.,
2001
) but, as explained in the Introduction, those studies were
also consistent with the conventional view.
Second, although motor axons are not required for the formation of AChR
clusters, they affect postsynaptic development in multiple ways. (1) Clusters
remodel after they are contacted by axons, leading to the precise apposition
of pre- and postsynaptic specializations
(Fig. 6A-C). (2) Aneural
clusters are lost soon after NMJs form
(Fig. 6F). (3) All AChR
clusters are eventually lost if axons fail to extend
(Fig. 7). These observations
support the idea that axons both disperse aneural and maintain innervated
clusters, and thus suggest that nerves provide both stabilization factors that
act locally and dispersal signals that act at a distance. A crucial local
stabilization factor is likely to be z-agrin; indeed, defects in agrin mutant
mice (Burgess et al., 1999;
Yang et al., 2001
;
Lin et al., 2001
) can be
explained by a loss of this stabilizing influence
(Misgeld et al., 2005
), even
though they were initially interpreted as supporting the idea that agrin acts
as a clustering agent in vivo, as it does in vitro
(McMahan, 1990
;
Ruegg et al., 1992
;
Ferns et al., 1992
;
Gesemann et al., 1995
). Recent
studies of mutant mice suggest that the neurotransmitter, acetylcholine, is a
crucial dispersal signal in vivo (Misgeld
et al., 2005
; Lin et al.,
2005
), although there is also evidence that locally presented
factors, such as agrin, can promote AChR cluster dispersal at a distance in
vitro (Madhavan et al., 2003).
Third, a single motor axon can form synapses in two distinct ways: by the
incorporation of initially aneural AChR clusters or by the organization of
clusters following contact with a myotube. For CaP, it appears that these
modes correspond to distinct fiber types, slow/pioneer and fast, respectively
(Fig. 5). The correspondence
might reflect an intrinsic, qualitative difference in synaptogenic ability
between the two fiber types, but it is also possible that it results only from
the different times at which the two populations are born. Adaxial/pioneer
fibers form considerably before they are contacted by CaP, whereas fast fibers
fuse as CaP contacts them. If AChR clustering can occur aneurally a few hours
following myotube formation, the developmental sequence would lead to the
formation of CaP-encountering clusters on muscle pioneers but not on fast
myotubes. Thus, consistent with results in mammals
(Misgeld et al., 2005),
axon-derived factors such as agrin might promote AChR clustering on some
myotubes but serve to prevent dispersal on others.
Although the primary motor system of zebrafish is unique in some respects
(Beattie, 2000), several
observations suggest that our conclusions may be applicable to the more
commonly studied NMJs of birds and mammals. (1) Some aneural AChR clusters are
present in rodent and chick muscles, in the vicinity of axons
(Lin et al., 2001
;
Smith and Slater, 1983
), and,
in one case, in advance of axonal contact
(Dahm and Landmesser, 1991
).
(2) AChR clusters extend beyond nerve terminals at early stages in the
formation of the mouse NMJ
(Matthews-Bellinger and Salpeter,
1983
). (3) AChR clusters shrink transiently perinatally in rodent
muscle before growth resumes postnatally
(Bevan and Steinbach,
1977
).
Our work is based on the pioneering studies of Eisen, Westerfield and
colleagues (Eisen and Meyers,
1986; Meyers et al., 1986,
Westerfield et al., 1986
;
Fashena and Westerfield, 1999
;
Westerfield et al., 1990
;
Sepich et al., 1998; Liu and Westerfield,
1992
). In general, our results are consistent with theirs, but
they differ in one respect: they did not observe aneural AChR clusters prior
to axon outgrowth or in segments rendered aneural by the ablation of primary
motoneurons (Liu and Westerfield,
1992
). We believe this difference is due to the improved
sensitivity of fluorescence detection methods and the improved resolution of
confocal imaging. Aneural AChR clusters are smaller and dimmer than those at
mature NMJs, and thus might have been missed previously. Moreover, clusters on
aneural segments are transient (Fig.
7), and many of the observations in those previous studies were
made after they would have disappeared.
In summary, we have provided evidence that some embryonic NMJs are formed
by the incorporation of initially aneural postsynaptic specializations. This
scenario differs from the widely accepted one in which axons emit signals that
organize the differentiation of postsynaptic sites beneath nerve terminals,
and components organized by the axon then promote presynaptic differentiation.
It is important to note, however, that the two scenarios are not mutually
exclusive. Indeed, both appear to operate within a single myotome. It is
possible that some differences among muscles in patterns of synaptogenesis and
responsiveness to agrin (Pun et al.,
2002) result from the predominant use of one mechanism or the
other. Moreover, the roles we document for axons in remodeling postsynaptic
sites once they are formed are common to both scenarios. Finally, most models
for synaptogenesis in the central nervous system are based on the idea that
postsynaptic differentiation occurs subsequent and consequent to axonal
contact (Li and Sheng, 2003
).
However, neurons, like myotubes, can organize postsynaptic specializations in
the absence of input (Rao et al.,
1998
). It will be interesting to learn whether some central
synapses form by incorporation of initially aneural postsynaptic sites, using
mechanisms analogous to those described here for the NMJ.
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ACKNOWLEDGMENTS |
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Footnotes |
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REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Akimenko, M. A., Johnson, S. L., Westerfield, M. and Ekker,
M. (1995). Differential induction of four msx homeobox genes
during fin development and regeneration in zebrafish.
Development 121,347
-357.
Anderson M. J. and Cohen M. W. (1978). Nerve-induced and spontaneous redistribution of acetyl choline receptors on cultured muscle cells. J. Physiol. 268,757 -773.
Barresi, M. J., Stickney, H. L. and Devoto, S. H.
(2000). The zebrafish slow-muscle-omitted gene product is
required for Hedgehog signal transduction and the development of slow muscle
identity. Development
127,2189
-2199.
Beattie, C. E. (2000). Control of motor axon guidance in the zebrafish embryo. Brain Res. Bull. 53,489 -500.[CrossRef][Medline]
Bevan S. and Steinbach, J. H. (1977). The distribution of alpha-bungarotoxin binding sites of mammalian skeletal muscle developing in vivo. J. Physiol. 267,195 -213.[Medline]
Blagden, C. S., Currie, P. D., Ingham, P. W. and Hughes, S.
M. (1997). Notochord induction of zebrafish slow muscle
mediated by Sonic hedgehog. Genes Dev.
11,2163
-2175.
Burgess, R. W., Nguyen, Q. T., Son, Y. J., Lichtman, J. W. and Sanes, J. R. (1999). Alternatively spliced isoforms of nerve- and muscle-derived agrin: their roles at the neuromuscular junction. Neuron 1,33 -44.[CrossRef]
Cohen, I., Rimer, M., Lomo, T. and McMahan, U. J. (1997). Agrin-induced postsynaptic-like apparatus in skeletal muscle fibers in vivo. Mol. Cell Neurosci. 9, 237-253.[CrossRef][Medline]
Cohen, S. A. and Fischbach, G. D. (1973). Regulation of muscle acetylcholine sensitivity by muscle activity in cell culture. Science 94,76 -78.
Dahm, L. M. and Landmesser, L. T. (1991). The regulation of synaptogenesis during normal development and following activity blockade. J. Neurosci. 11,238 -255.[Abstract]
Devoto, S. H., Melancon, E., Eisen, J. S. and Westerfield,
M. (1996). Identification of separate slow and fast muscle
precursor cells in vivo, prior to somite formation.
Development 122,3371
-3380.
Eisen, J. S., Meyers, P. Z. and Westerfield, M. (1986). Pathway selection by growth cones of identified motorneurons in live zebrafish embryos. Nature 320,269 -271.[CrossRef][Medline]
Fashena, D. and Westerfield, M. (1999). Secondary motoneuron axons localize DM-GRASP on their fasciculated segments. J. Comp. Neurol. 406,415 -424.[CrossRef][Medline]
Ferns, M., Hoch, W., Campanelli, J. T., Rupp, F., Hall, Z. W. and Scheller, R. H. (1992). RNA splicing regulates agrin-mediated acetylcholine receptor clustering activity on cultured myotubes. Neuron 8,1079 -1086.[CrossRef][Medline]
Frank, E. and Fischbach, G. D. (1979). Early events in neuromuscular junction formation in vitro: induction of acetylcholine receptor clusters in the postsynaptic membrane and morphology of newly formed synapses. J. Cell Biol. 83,143 -158.[Abstract]
Gautam, M., Noakes, P. G., Moscoso, L., Rupp, F., Scheller, R. H., Merlie, J. P. and Sanes, J. R. (1996). Defective neuromuscular synaptogenesis in agrin-deficient mutant mice. Cell 85,525 -535.[CrossRef][Medline]
Gesemann, M., Denzer, A. J. and Ruegg, M. A. (1995). Acetylcholine receptor-aggregating activity of agrin isoforms and mapping of the active site. J. Cell Biol. 128,625 -636.[Abstract]
Grinnell, A. D. (1995). Dynamics of
nerve-muscle interaction in developing and mature neuromuscular junctions.
Physiol. Rev. 75,789
-834.
Higashijima, S., Hotta, Y. and Okamoto, H.
(2000). Visualization of cranial motor neurons in live transgenic
zebrafish expressing green fluorescent protein under the control of the
islet-1 promoter/enhancer. J. Neurosci.
20,206
-218.
Jessell, T. M. (2000). Neuronal specification in the spinal cord: inductive signals and transcriptional codes. Nat. Rev. Genet. 1,20 -29.[CrossRef][Medline]
Jones, G., Meier, T., Lichtsteiner, M., Witzemann, V., Sakmann,
B. and Brenner, H. R. (1997). Induction by agrin of ectopic
and functional postsynaptic-like membrane in innervated muscle.
Proc. Natl. Acad. Sci. USA
94,2654
-2659.
Kimmel, C. (1995). Stages of embryonic development of the zebrafish. Dev. Dyn. 203,253 -310.[Medline]
Li, Z, and Sheng, M. (2003). Some assembly required: the development of neuronal synapses. Nat. Rev. Mol. Cell. Biol. 4,833 -841.[CrossRef][Medline]
Lin, W., Burgess, R. W., Dominguez, B., Pfaff, S. L., Sanes, J. R. and Lee, K. F. (2001). Distinct roles of nerve and muscle in postsynaptic differentiation of the neuromuscular synapse. Nature 410,1057 -1064.[CrossRef][Medline]
Lin, W., Dominguez, B., Yang, J., Aryal, P., Brandon, E. P., Gage, F. H. and Lee, K. F. (2005). Neurotransmitter acetylcholine negatively regulates neuromuscular synapse formation by a Cdk5-dependent mechanism. Neuron 46,569 -579.[CrossRef][Medline]
Liu, D. W. and Westerfield, M. (1992). Clustering of muscle acetylcholine receptors requires motoneurons in live embryos, but not in cell culture. J. Neurosci. 12,1859 -1866.[Abstract]
Madhavan, R. and Peng, H. B. (2003). A synaptic balancing act: local and global signaling in the clustering of ACh receptors at vertebrate neuromuscular junctions. J. Neurocytol. 32,685 -696.[CrossRef][Medline]
Matthews-Bellinger, J. A. and Salpeter, M. M. (1983). Fine structural distribution of acetylcholine receptors at developing mouse neuromuscular junctions. J. Neurosci. 3,644 -657.[Abstract]
McMahan, U. J. (1990). The agrin hypothesis. Cold Spring Harb. Symp. Quant. Biol. 55,407 -418.[Medline]
Misgeld, T., Burgess, R. W., Lewis, R. M., Cunningham, J. M., Lichtman, J. W. and Sanes, J. R. (2002). Roles of neurotransmitter in synapse formation: development of neuromuscular junctions lacking choline acetyltransferase. Neuron 36,635 -648.[CrossRef][Medline]
Misgeld, T., Kummer, T. T., Lichtman, J. W. and Sanes, J. R.
(2005). Agrin promotes synaptic differentiation by counteracting
an inhibitory effect of neurotransmitter. Proc. Natl. Acad. Sci.
USA 102,11088
-11093.
Moriyoshi, K., Richards, L. J., Akazawa, C., O'Leary D. D. and Nakanishi, S. (1996). Labeling neural cells using adenoviral gene transfer of membrane-targeted GFP. Neuron 16,255 -260.[CrossRef][Medline]
Myers P. Z., Eisen, J. S. and Westerfield, M. (1986). Development and axonal outgrowth of identified motoneurons in the zebrafish. J. Neurosci. 6,2278 -2289.[Abstract]
Niell, C. M., Meyer, M. P. and Smith, S. J. (2004). In vivo imaging of synapse formation on a growing dendritic arbor. Nat. Neurosci. 7, 254-260.[CrossRef][Medline]
Park, H. C., Kim, C. H., Bae. Y. K., Yeo, S. Y., Kim, S. H., Hong, S. K., Shin, J., Yoo, K. W., Hibi, M., Hirano, T. et al. (2000). Analysis of upstream elements in the HuC promoter leads to the establishment of transgenic zebrafish with fluorescent neurons. Dev. Biol. 227,279 -293.[CrossRef][Medline]
Pun, S., Sigrist, M., Santos, A. F., Ruegg, M. A., Sanes, J. R., Jessell, T. M., Arber, S. and Caroni, P. (2002). An intrinsic distinction in neuromuscular junction assembly and maintenance in different skeletal muscles. Neuron 34,357 -370.[CrossRef][Medline]
Rao, A., Kim, E., Sheng, M. and Craig, A. M.
(1998). Heterogeneity in the molecular composition of excitatory
postsynaptic sites during development of hippocampal neurons in culture.
J. Neurosci. 18,1217
-1229.
Ruegg, M. A., Tsim, K. W., Horton, S. E., Kroger, S., Escher, G., Gensch, E. M. and McMahan, U. J. (1992). The agrin gene codes for a family of basal lamina proteins that differ in function and distribution. Neuron 4,691 -699.[CrossRef]
Sanes, J. R. and Lichtman, J. W. (1999). Development of the vertebrate neuromuscular junction. Annu. Rev. Neurosci. 22,389 -442.[CrossRef][Medline]
Sanes, J. R. and Lichtman, J. W. (2001). Induction, assembly, maturation and maintenance of a post-synaptic apparatus. Nat. Rev. Neurosci. 11,791 -805.
Sanes, J. R., Feldman, D. H., Cheney, J. M. and Lawrence, J. C. (1984). Brain extract induces synaptic characteristics in the basal lamina of cultured myotubes. J. Neurosci. 2, 464-473.
Smith, M. A. and Slater, C. R. (1983). Spatial distribution of acetylcholine receptors at developing chick neuromuscular junctions. J. Neurocytol. 12,993 -1005.[CrossRef][Medline]
Stickney, H. L., Barresi, M. J. and Devoto, S. H. (2000). Somite development in zebrafish. Dev. Dyn. 219,287 -303.[CrossRef][Medline]
Sytkowski, A. J., Vogel, Z. and Nirenberg, M. W. (1973). Development of acetylcholine receptor clusters on cultured muscle cells. Proc. Natl. Acad. Sci. USA 1, 270-274.
Trevarrow, B., Marks, D. and Kimmel, C. B. (1990). Organization of hindbrain segments in the zebrafish embryo. Neuron 4,669 -679.[CrossRef][Medline]
Udvadia, A. J., Koster, R. W. and Skene, J. H.
(2001). GAP-43 promoter elements in transgenic zebrafish reveal a
difference in signals for axon growth during CNS development and regeneration.
Development 128,1175
-1182.
Walsh, M. K. and Lichtman, J. W. (2003). In vivo time-lapse imaging of synaptic takeover associated with naturally occurring synapse elimination. Neuron 37, 67-73.[CrossRef][Medline]
Westerfield, M., McMurray, J. V. and Eisen, J. S. (1986). Identified motoneurons and their innervation of axial muscles in the zebrafish. J. Neurosci. 6,2267 -2277.[Abstract]
Westerfield, M., Liu, D. W., Kimmel, C. B. and Walker, C. (1990). Pathfinding and synapse formation in a zebrafish mutant lacking functional acetylcholine receptors. Neuron 4, 867-874.[CrossRef][Medline]
Yang, X., Li, W., Prescott, E. D., Burden, S. J. and Wang, J.
C. (2000). DNA topoisomerase II beta and neural development.
Science 287,131
-134.
Yang, X., Arber, S., William, C., Yasuto Tanabe, L. L., Jessell, T. M., Birchmeier, C. and Burden, S. J. (2001). Patterning of muscle acetylcholine receptor gene expression in the absence of motor innervation. Neuron 30,399 -410.[CrossRef][Medline]
Ziskind-Conhaim, L., Geffen, I. and Hall, Z. W. (1984). Redistribution of acetylcholine receptors on developing rat myotubes. J. Neurosci. 9,2346 -2349.