Centre for Research in Neuroscience, McGill University and Montreal General Hospital Research Institute, Montreal, Quebec H3G 1A4, Canada
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ABSTRACT |
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Nguyen, Peter V.,
Laurent Aniksztejn,
Stefano Catarsi, and
Pierre Drapeau.
Maturation of neuromuscular transmission during early development in
zebrafish. We have examined the rapid development of synaptic
transmission at the neuromuscular junction (NMJ) in zebrafish embryos
and larvae by patch-clamp recording of spontaneous miniature endplate
currents (mEPCs) and single acetylcholine receptor (AChR) channels.
Embryonic (24-36 h) mEPCs recorded in vivo were small in amplitude
(<50 pA). The rate of mEPCs increased in larvae (3.5-fold increase
measured by 6 days), and these mEPCs were mostly of larger amplitude
(10-fold on average) with (5-fold) faster kinetics. Intracellular
labeling with Lucifer yellow indicated extensive coupling between
muscle cells in both embryos and larvae (
10 days). Blocking
acetylcholinesterase (AChE) with eserine had no effect on mEPC kinetics
in embryos at 1 day and only partially slowed (by ~1/2) the
decay rate in larvae at 6 days. In acutely dissociated muscle cells, we
observed the same two types of AChR with conductances of 45 and 60 pS
and with similar, brief (<0.5 ms) mean open times in both embryos and
larvae. We conclude that AChR properties are set early during
development at these early stages; functional maturation of the NMJ is
only partly shaped by expression of AChE and may also depend on
postsynaptic AChR clustering and presynaptic maturation.
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INTRODUCTION |
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Understanding the mechanism of synapse formation
during embryonic development is an important goal of neurobiology. Much
of what we know about the formation and maturation of synapses is based
on studies of the highly accessible peripheral neuromuscular junction
(NMJ). The cellular and molecular events taking place during embryonic
development of the NMJ have been characterized in considerable detail
in vitro and in vivo in a variety of vertebrate and mammalian species
(reviewed by Hall and Sanes 1993;
Jacobson 1991
). Ultimately, the cellular analysis of
functional development needs to be determined in vivo. Detailed studies
of neuromuscular synapse formation have been performed with cultured
cells, but only a few studies have examined the functional changes
occurring in vivo because of the difficulty of performing
electrophysiological recordings in early embryos.
Early studies (before the advent of high-resolution patch-clamp
techniques) reported developmental transitions in endplate potentials
in the embryonic rat (Bennett and Pettigrew 1974;
Dennis et al. 1981
; Diamond and Miledi
1962
) and Xenopus (Kullberg et al.
1977
). We chose to investigate the development of neuromuscular transmission in the zebrafish for a variety of reasons. First, this
animal offers the advantage of obtaining large numbers of externally
fertilized eggs at precise developmental stages that are easily
identified because of the optical clarity of the embryos. In
particular, the migrations and pathfinding of identified neurons such
as the spinal motoneurons have been described in detail in the embryo
and larva (Myers et al. 1986
; Westerfield et al.
1986
) and appear to be related to the onset of early
locomotor behaviors (Saint-Amant and Drapeau 1998
).
However, only limited physiological information is available concerning
the development and properties of neuromuscular synaptic transmission
(Grunwald et al. 1988
; Liu and Westerfield
1988
; Westerfield et al. 1986
). In particular, an analysis of the endplate currents (EPCs) and the underlying single
acetylcholine receptor (AChR) properties has not been done to date
during in vivo NMJ maturation in any preparation. Another interest of
the zebrafish is that recent genetic screens have identified mutations
affecting embryonic locomotor behaviors (Granato et al.
1996
), suggesting that the molecular bases of the development of these behaviors can be studied. However, an accurate description of
the mutant phenotypes requires the characterization of the functional
deficits before precise knowledge of the consequences of gene mutations
can be established. A functional definition of embryonic locomotion
requires an understanding of the mechanisms responsible for the
maturation of motoneurons and of neuromuscular synaptic transmission,
which together constitute "the final common pathway for all
behavioral acts" (Sherrington 1947
).
To examine the properties of developing neuromuscular synapses in the
zebrafish, we have combined whole muscle cell patch-clamp recordings of
synaptic activity in the embryo and larva in vivo and recordings of
single AChRs in acutely dissociated muscle fibers at these two early
developmental stages. By recording postsynaptic currents, we have been
able to relate the properties of synaptic transmission to single AChR
channels. Previous morphological studies have shown that the three
primary motoneurons on either side of each of the segmented spinal cord
somites extend axons that reach the muscle fibers along the middle of
the segmental myotomes and form tiny "en passant" synapses starting
at 17 h after fertilization (Liu and Westerfield 1992;
Myers et al. 1986
; Westerfield et al. 1986
), which is the time at which spontaneous contractions of the tail are first observed (Saint-Amant and Drapeau
1998
). These contractions eventually cease, and by 27 h
the embryos swim briefly in response to a touch (Eaton and
Farley 1973
; Kimmel et al. 1972
; Saint-Amant and Drapeau 1998
), the time at which the
more numerous secondary motoneurons innervate the muscles (Myers
et al. 1986
; Westerfield et al. 1986
). The
embryos hatch on the second day of development, and within a few days
the newborn larvae swim in short bursts at near-maximal rates
(Eaton and Farley 1973
; Kimmel et al.
1972
; Saint-Amant and Drapeau 1998
). We found it difficult to completely suppress (e.g., with TTX or high-Mg/low-Ca concentrations) the large-amplitude contractions, with the tail reaching the head of unrestrained embryos. It was thus not possible to
maintain patch-clamp recordings from the very fragile (unfused) muscle
cells in embryos <24 h old. We report here our observations with
embryos between 24 and 30 h, when swimming is first manifested (Saint-Amant and Drapeau 1998
), and contrast these with
the observations in 3- to 6-day larvae, when swimming has reached a
maximal rate. We show that the mEPCs increase greatly in frequency,
amplitude, and kinetics during this early period of maturation without
a change in AChR properties.
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METHODS |
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Embryonic and larval zebrafish preparations
Whole cell voltage-clamp recordings of mEPCs were performed on
living zebrafish embryos and larvae (ages 24 h to 6 days). All
procedures were carried out in compliance with the guidelines stipulated by the Canadian Council for Animal Care and McGill University. Intact animals were anesthetized by immersion in tricaine (0.1 mg/ml, Sigma) dissolved in a modified extracellular saline solution (Westerfield et al. 1986) consisting of (in mM)
116 NaCl, 2 KCl, 0.7 CaCl2, 10 MgCl2, 10 glucose, and 10 HEPES, adjusted to pH 7.2, 290 mosmol. The reduced
calcium and elevated magnesium concentrations were necessary to
attenuate muscle contractions during our experiments. In some
experiments, 1 µM TTX (Sigma) was added, and similar results were
obtained. Fine tungsten wires (0.001 in.) were used to securely pin
specimens through the notochord onto a Sylgard-lined petri dish. A
glass needle and fine forceps were used to expose the axial muculature
of a specimen by lifting and then carefully peeling its skin. After
this initial procedure, tricaine was washed out and replaced with
drug-free saline. The viability of the preparation was assessed by
looking for vigorous heart contractions and blood circulation.
Whole cell voltage-clamp recordings
Standard whole cell voltage-clamp recordings (Hamill et
al. 1981) were performed on dorsal and ventral muscle fibers
(epaxial and hypaxial muscles) (see Myers 1985
) in vivo
under direct visualization with the use of Hoffmann modulation optics
(×40 water immersion lens, Nikon Labophot microscope). All experiments
were performed at room temperature (22°C). Patch-clamp electrodes
were pulled from thin-walled, Kimax-51 borosilicate glass (3- to 5-M
resistance) and were filled with (in mM) 130 CsCl, 2 MgCl2,
10 HEPES, 10 EGTA, and 4 NaATP, pH 7.2, 290 mosmol. The series
resistance (5-10 M
) was compensated by 60-80%. Whole cell
currents were recorded with an Axopatch-1D amplifier (Axon
Instruments), filtered at 2 kHz (
3 dB) and digitized at 10 kHz,
allowing the resolution of events with rise times of >0.1 ms. Data
were acquired with pClamp 6.0 software (Axon Instruments) and were
analyzed off-line with Axograph 3.5 (Axon Instruments). Cells with
resting potentials less than
50 mV were discarded, and all recordings
were obtained from a holding potential of
60 mV. For some experiments
0.1 mM eserine (Sigma), an inhibitor of acetylcholinesterase (AChE),
was dissolved in the modified saline solution, and specimens were
preincubated in drug for
30 min before recording to ensure
penetration of the eserine. We performed statistical comparisons by
using a Kolmogorov-Smirnov test for mEPC data.
Dye coupling
In some experiments fluorescent Lucifer yellow (Li salt; Sigma)
was included (0.1%) in the patch pipette solution to determine the
extent of coupling between the muscle cells at different developmental stages. In some experiments we added 1-3 mM halothane, 1-octanol, or
1-heptanol (Sigma) to the extracellular solution and superfused the
preparations for 30 min before starting the recordings to uncouple
the cells with these gap junction blockers. The cells were photographed
with a still camera after removing the pipette at the end of the experiment.
Acute dissociation of muscle cells
To enhance recording of single AChRs, we acutely dissociated
muscle cells by treating skinned embryos (as described previously) for
5-10 min with 0.1% collagenase (Type XII, Sigma Chemical) dissolved
in normal extracellular solution (containing 1.3 mM CaCl2
and 1 mM MgCl2). The collagenase was washed off, and the cells were dissociated with a fine glass needle while incubating the
tissue fragments in a modified recording solution that was nominally
Mg- and Ca-free and that contained 2 mM EDTA. Dissociated cells were
transferred to tissue culture dishes coated with polylysine (Sigma) to
enhance cell adhesion and were kept in normal extracellular solution
for 2 h. No obvious changes in cell morphology were noticed during
these short time periods.
Single-channel recording
Electrodes of 5- to 10-M resistance were coated with dental
wax to ~50 µm of their tips, and the recording solution level was
kept low to reduce capacitive noise (<0.3 pA RMS). The electrodes were
filled with either recording solution containing 1.3 mM
CaCl2, 1 mM MgCl2, and 0.4 µM ACh for
cell-attached recordings or with KCl pipette solution (as for whole
cell recordings, but containing KCl instead of CsCl) for recording in
the outside-out configuration (Hamill et al. 1981
).
Recordings were filtered at 5 kHz (
3 dB) and digitized at 20 kHz.
Data were stored to computer disk or digital tape and were analyzed
off-line with pClamp 6 software (Axon Instruments). For some
experiments, cells were continuously superfused with recording solution
containing drugs (0.4 µM ACh, 10 µM D-tubocurarine;
Sigma). The membrane potential of the muscle fibers was determined in
the whole cell configuration and was used to correct the pipette
potential applied during recordings.
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RESULTS |
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Morphological features of embryonic and larval muscle fibers in vivo
As an initial step toward a physiological characterization of neuromuscular synaptic properties in living zebrafish embryos, we compared the morphological features of myotomes and muscle fibers in embryos and larvae at two different developmental stages. Between 24 and 30 h postfertilization, chevron-shaped myotomes (Fig. 1A) were clearly demarcated in the dorsal (top) and ventral regions of the zebrafish embryo. However, individual myocytes within each segmental myotome could not be clearly discerned at this stage. Ventral axial muscles were less prominent at this stage than were dorsal muscles. In sharp contrast larvae showed long, fused muscle fibers with discrete striations at 2-µm intervals (an isolated muscle fiber from a 3-day larva is shown more clearly in Fig. 4A2), and the demarcations between individual fibers were clearly evident (Fig. 1B). At this stage of development, skin peeling was easily performed, and the result was invariably a more pristine and intact preparation than with embryos.
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Differential activity profiles in living embryos and larvae
We compared the spontaneous neuromuscular synaptic activity
profiles of embryos (24-36 h) and larvae (3-6 days). We bathed the
preparations in a high-Mg/low-Ca solution to prevent muscle contractions and to isolate spontaneous (presumably quantal) events. The recordings were performed near the middle of the somite, where neuromuscular innervation occurs (Liu and Westerfield
1992). Similar levels of synaptic activity were observed in
preliminary experiments (not shown) in the presence of 1 µM TTX at a
normal Ca concentration compared with the levels observed in
high-Mg/low-Ca solution at both stages of development, indicating that
these were spontaneous, not evoked, events. In embryos, whole cell
voltage-clamp recordings of spontaneous miniature EPCs (mEPCs) revealed
a mean frequency of 26 ± 20 (SD) min
1
(n = 5; Fig. 1B shows selected traces with
several events each). In contrast, in 6-day larvae, the mean frequency
was significantly higher, 92 ± 38 min
1
(n = 5; P < 0.01). Hence the mean
frequency of spontaneous synaptic transmitter release was ~3.5-fold
higher in the 6-day larvae than in the more immature 1-day embryos. The
mEPCs were blocked at all stages in the presence of 10 µM
D-tubocurarine, as shown previously for the block of
neuromuscular transmission in larvae (Grunwald et al.
1988
).
mEPCs recorded in embryos and larvae show distinct kinetic characteristics
A casual inspection of mEPCs recorded in embryos (Fig. 1A) and larvae (Fig. 1B) revealed two primary characteristics. First, mEPCs recorded from embryos were invariably small in size, in contrast to the larger mEPCs measured from larvae. Second, the embryonic mEPCs were slower in their kinetics of onset and decay than those observed in larvae. In general, mEPCs with fast kinetics of onset and decay emerged at ~36-42 h of development (not shown), and the transition from slow to fast mEPC kinetics was largely completed by 3 days of development.
A more thorough analysis of these mEPCs showed that in embryos the mean amplitude was 12.3 ± 10.4 pA (n = 5 muscle cells, in different animals). The amplitudes measured in larvae were significantly greater, 62.3 ± 16.1 pA at 3 days and 122.2 ± 120.7 pA at 6 days (P < 0.005, n = 5 preparations; Fig. 2, A-E, and Table 1). The median amplitudes at 27 h and 6 days were 7 and 73 pA, respectively (Fig. 2F). In no case could the amplitude histograms be described by a single Gaussian distribution. In summary, spontaneous mEPCs recorded at these two developmental stages differed significantly in their mean amplitudes.
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We also observed significant age-dependent increases in the kinetics of onset and decay of mEPCs. The average mEPC rise times (10-90% maximum amplitude) in 24- to 36-h embryos versus 3- and 6-day larvae were 2.4 ± 2.1 ms versus 0.23 ± 0.04 ms and 0.49 ± 0.28 ms, respectively (P < 0.0001 for embryos compared with larvae, n = 5 for each age; Table 1). The medians of the rise time distributions for these two stages were 1.95 ms in embryos and 0.35 ms in 6 day larvae, respectively (Fig. 3A). In embryos, the mean decay time constant of mEPCs was 7.2 ± 2.3 ms (Table 1), which was significantly longer (P < 0.0001) than the mean decay time constant measured in larvae at 3 days (1.3 ± 0.4) and 6 days (2.1 ± 1.3 ms; Table 1). The median decay time constants for these two stages were 6.8 ms in embryos and 1.5 ms in 6-day larvae, respectively (Fig. 3B). Finally, there were clear biases in the distributions of mEPC sizes and their matching decay times at both stages (Fig. 3C). At 27 h, mEPCs were distributed such that the majority had decay time constants of >4 ms and amplitudes of <50 pA. In larvae by sharp contrast, mEPCs observed at 3 and 6 days had decay time constants that were mostly <2 ms, with relatively few events >4 ms in duration (Fig. 3C), and most of these mEPCs had amplitudes of >100 pA.
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Coupling between muscle cells
One possible explanation for the presence of smaller and slower
events in the embryonic muscle cells is if these are extensively coupled in the embryo and the coupling is lost at the larval stage, as
described for Xenopus embryos (Armstrong et al.
1983) where the coupling is lost within 2 days of formation of
the NMJ. Accordingly, the embryonic cells would be more affected by
cable (or syncytial) filtering. We examined this by including the
small, gap junction-permeable fluorescent dye Lucifer yellow in the
patch pipette. As shown in Fig. 4,
B and D, many other muscle cells were labeled
with Lucifer yellow, indicating extensive coupling in both embryos (Fig. 4, A and B) and larvae (Fig. 4,
C and D). Larval muscle cells remained coupled
for
10 days, and only in older larvae (>14 days) did Lucifer yellow
not pass into other cells (not shown), indicating a rather delayed
uncoupling. However, the muscle fibers were too large at this stage for
whole cell recordings of mEPCs.
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In other experiments (not shown) we attempted to uncouple muscle cells
in younger larvae and embryos by incubating these in the presence of
1-3 mM of the common gap junction uncouplers halothane, 1-octanol, or
1-heptanol (de Roos et al. 1996; Niggli et al.
1989
). Halothane and 1-octanol failed to uncouple the cells;
although 1-heptanol prevented coupling, it also eliminated mEPCs even
at late stages (2-3 wk) when coupling was no longer present,
suggesting a toxic effect, e.g., such as blockage of AChRs
(Murrell et al. 1991
). We were therefore unable to
directly test the effects of coupling on mEPC properties and conclude
that extensive coupling exists at all the stages we could record from.
The small fraction (<5%) of low amplitude (<100 pA), slow (>4 ms) mEPCs observed in larvae (Fig. 3) could represent immature events similar to those observed in embryos or perhaps represent a minor population of poorly clamped events in the larger, larval muscle fibers, although we recorded from the middle of the fibers, presumably close to where they are innervated. That the mEPCs at 6 days were somewhat slower on average than at 3 days (Table 1) and had more of the very slow events (>4 ms) suggests that there was a small but gradual decrease in the effectiveness of the space clamp. This is consistent with an increase in muscle fiber size during this developmental period, and we were unable to effectively voltage clamp muscle cells in older larvae. We conclude that the mEPCs recorded in 24- to 27-h embryos were, on average, ~10-fold smaller in size and 3- to 4-fold slower in kinetics of onset and decay than were mEPCs measured in 6-day larvae.
Differential effects of AChE inhibition on mEPC decay times
What might be the molecular bases of the developmental difference in mEPC decay time constants? We addressed this question by first examining the effects of acute exposure of axial muscles to an inhibitor of AChE. We reasoned that blocking AChE, which hydrolyzes and inactivates the neuromuscular transmitter, should prolong the decay time constants of mEPCs because of decreased breakdown of and consequent prolonged exposure to ACh in the synaptic cleft. We found that, for embryos, 0.1 mM eserine had no significant effect on the mean decay time constant (6.0 ± 1.9 ms vs. 7.2 ± 2.3 ms in controls; P > 0.2, see Fig. 3B and Table 2). A similar lack of effect was observed at 1 mM eserine (not shown). However, we observed pronounced effects of eserine in larvae as mEPCs showed twofold longer decay time constants in the presence of the drug (Fig. 3B and Table 2). Nonetheless, it is noteworthy that the decay time constants measured in larvae in eserine (mean = 3.8 ms) did not match the embryonic control (drug-free) decay time constant (mean = 7.2 ms). Thus the activity of AChE in the synaptic cleft has only a partial role in determining the decay rates of mEPCs recorded in zebrafish axial muscles, and the transition from slow to fast mEPC decay rates cannot be attributed entirely to changes in AChE activity during development.
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Similar AChR channels in embryonic and larval muscle cells
We hypothesized that the observed developmental alterations in
mEPC kinetics may also arise from changes in the properties of AChR
channels during development, such as the expression of different
subunits or subtypes. To characterize the properties of AChRs, we
recorded single-channel activity from acutely dissociated muscle cells.
As shown in Fig.
5A1, myocytes from
1-day-old embryos were small, spherical cells of 10-20 µm in
diameter. Muscle cells isolated from larvae were long, fused
trapezoidal cells of 20-30 µm in diameter and 100 µm in
length (Fig. 5A2). Regular striations were
evident at 2-µm intervals. These dissociated cells thus appeared to
retain the morphology observed in vivo (Fig. 1). We isolated muscle
cells from 3-day-old larvae because cells from older specimens were
more difficult to dissociate, and the developmental changes in mEPC
properties were largely complete by this time. Unlike mammalian muscle
fibers (Brehm and Kullberg 1987
), there was no thickening of the en passant NMJ that could be discerned in early zebrafish muscle cells.
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Cell-attached recordings with 0.4 µM ACh in the pipette from muscle cells at both embryonic and larval stages revealed two types of channels (Fig. 5, B1 and B2). Amplitude histograms of channel openings in the experiment illustrated in Fig. 5 were best fit by two Gaussian distributions (Fig. 5, C1 and C2) with predominantly larger amplitude events. On average the channels had slope conductances (Fig. 6A, estimated over a range of holding potentials) of 45 pS (45.3 ± 4.4 pS, n = 28) and 60 pS (60.1 ± 2.6 pS, n = 27) and mean open times of <1 ms, as described subsequently (based on all recordings presented in Table 3). These events were likely due to the presence of two distinct channels (rather than subconductance states) as patches containing one or the other type of channel were observed in approximately one-half (15/28) of the patches. Furthermore, when two channels were present, either one could predominate, although the 60-pS channel was seen more frequently. Occasionally other conductance levels were observed (usually of ~20, 35, and >100 pS), but unlike the 45- and 60-pS channels these proved to be insensitive to D-tubocurarine.
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We characterized the properties of the channels in dissociated myocytes from embryos and larvae to ascertain whether their properties changed between these stages of development. As shown in Fig. 6A, ~45-pS and (more frequently) ~60-pS conductances were observed in embryonic myocytes and had reversal potentials near 0 mV. Similar conductances were observed in larval muscle cells as well (Table 3; Student's t-test, P > 0.05 for the difference between each type of conductance in each "type of recording"). Single exponential distributions were normally observed for the open and closed time histograms (Fig. 6, B1 and B2) with means of <1 ms and of a few hundred milliseconds, respectively (Table 3). Only occasionally (4/55 channels analyzed) were second, longer open (~5 ms) or closed times (~400 ms) detected (as seen for a small number of events in Fig. 6, B1, B2, and C1). Rather than reflecting distinct but rarely observed second states, we believe that these longer-lasting events were more likely due to missed events during unresolved bursts. We conclude that these channels shared similar kinetic properties in embryos and larvae despite their different conductances.
To ascertain that these were indeed AChR channels, the sensitivity of
the channels to application of ACh (0.4 µM) and
D-tubocurarine (10 µM) was examined in outside-out
patches because synaptic currents were found to be abolished at this
concentration of D-tubocurarine, which was shown previously
to suppress neuromuscular transmission in the zebrafish
(Grunwald et al. 1988). As shown in Fig.
7A, recordings from
outside-out patches revealed channels with properties indistinguishable
from those observed in cell-attached patches (Table 3) in that both 45- and 60-pS conductances with brief (<1 ms) openings were observed. A
very low, spontaneous level of activity was observed in the
absence of applied ACh (Control), and as shown in Fig. 7, A
(ACh) and B (representative of 3 such experiments), the
probability of channel opening increased rapidly on application of ACh.
Channel activity was completely suppressed by subsequent application of
10 µM D-tubocurarine in the presence of ACh [Fig. 7,
A (ACh + d-Tc) and B], confirming that these
channels were indeed AChR channels. Furthermore, in 12 recordings from cell-attached patches with D-tubocurarine in the pipette
solution, no channel activity was observed. We conclude from these
results that the same two types of AChR are present throughout the
developmental stages examined.
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DISCUSSION |
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We have observed spontaneous mEPCs in zebrafish embryos between 24 and 36 h, with slow rise times of ~2 ms and decay times of ~7
ms. The mEPC amplitudes were 10-80 pA (mean 12 pA). Primary motoneurons start innervating the muscles at ~17 h postfertilization, the same time at which spontaneous tail contractions are first observed
(Myers et al. 1986; Westerfield et al.
1986
). This is also the earliest time at which clusters of
AChRs have been detected in living embryos by staining with fluorescent
-bungarotoxin (Liu and Westerfield 1992
). Thus AChR
clustering appears to occur within a few hours of contact between the
motoneurons and muscle cells. After hatching, when larvae are able to
swim in fast bursts (Eaton and Farley 1973
;
Kimmel et al. 1972
; Saint-Amant and Drapeau 1998
), severalfold larger and faster synaptic currents were
observed (rise times of <0.5 ms and decay times of <2 ms), suggesting
an increased efficiency of transmission at maturing endplates. The neuronal sources of these mEPCs cannot be unambiguously defined because
of polyneuronal innervation. The caudal primary motoneuron and numerous
secondary motoneurons innervate the lateral ventral muscle fibers,
whereas the middle primary motoneuron and secondary motoneurons
innervate the dorsal muscles (Myers 1985
;
Westerfield et al. 1986
). This polyneuronal innervation
may account for the non-Gaussian distribution of mEPCs recorded at the
stages we examined if different presynaptic endings activated synaptic
responses of different amplitudes. Muscle cells were extensively
coupled in both embryos and larvae for
2 wk, in contrast to the
coupling between Xenopus embryonic muscle cells that is lost
after ~2 days (Armstrong et al. 1983
). This indicates
that the trunk contractions during early larval swimming are mediated
by a synsytium of coupled muscle fibers that may help minimize
disparities in the extent of NMJ maturation by enhancing the overall
performance of the muscle.
Two types of AChR channels were detected in muscle cells acutely
dissociated from embryos, 45- and 60-pS conductance channels with mean
open times of <1 ms. In the dissociated muscle cells, we could not
distinguish between synaptic and extrasynaptic AChRs. However, in
numerous other preparations, extrasynaptic AChRs appear to have
properties similar to synaptically localized AChRs and provide direct
information about the latter (Brehm and Kullberg 1987;
reviewed by Edmonds et al. 1995
; Schuetze and
Role 1987
). The same two types of AChR channels were observed
in larvae, ruling out an obvious transition in channel properties
during maturation at these early stages. However, more subtle changes,
e.g., in modal gating of the channels (Naranjo and Brehm
1993
) or in affinity for ACh, may not have been detected in our
experiments. In many other preparations, a switch from slow- to
fast-channel kinetics is observed during muscle maturation (reviewed by
Jacobson 1991
; Schuetze and Role 1987
). A
later postnatal switch in bovine (Mishina et al. 1986
)
and murine (Witzemann et al. 1996
) AChR subunit
composition has been shown to underlie the kinetic transitions in these
species. Thus in most preparations it appears that postsynaptic events determine the early maturation of the NMJ during the first few days or
weeks of development. In contrast, two types of AChR subtype with fast
kinetics have also been observed in mice (Sheperd and Brehm
1997
), and chick muscle has only one type of AChR with a mean
open time that remains long (Schuetze 1980
). Similar to
these latter species, the AChR openings that we observed in zebrafish appear to remain constant (albeit brief) throughout the early developmental period that we examined, suggesting that the channel properties may be expressed from the onset of embryogenesis. As we did not examine the properties of mEPCs and AChRs in mature animals,
we cannot exclude a later change in receptor properties. However it
seems unlikely that the synaptic currents can get much faster than
those in larvae as these are already among the fastest cholinergic
responses to be reported and are close to the limit of detection (and effectiveness).
At the low concentration (0.4 µM) of ACh used here, only single AChR
channel openings of <0.5 ms in duration were observed. At higher
concentrations, bursts or clusters of openings have been observed in
other preparations (reviewed by Edmonds et al. 1995;
Lingle et al. 1992
), and their durations appear to
determine the time course of synaptic events (e.g., at
Xenopus NMJs) (Kullberg and Owens 1986
).
Accordingly, we would expect several reopenings of AChRs at mature
synapses, given the mean synaptic decay time constant of ~2 ms that
we observed. It is interesting to note that in Xenopus
(Kullberg and Owens 1986
) and garter snakes
(Dionne and Parsons 1978
, 1981
) the mEPC decay rates are
related to contraction velocity. Because zebrafish larvae and adults
show a high frequency (20-40 Hz) of contractions during swimming
(Liu and Westerfield 1988
; Saint-Amant and
Drapeau 1998
) and have faster and more pronounced startle
contractions (Kimmel et al. 1972
; Saint-Amant and
Drapeau 1998
), we propose that the physiological properties of
the NMJ (brief AChR channel openings and fast mEPCs) have been
optimized for these high rates of muscle contraction.
The immature synapses recorded in embryos had time constants of
decay that were threefold longer than those recorded in larvae, suggesting prolonged channel reopenings during ACh release. The AChE
inhibitor eserine caused only a twofold slowing of larval mEPCs and did
not affect the time course of embryonic mEPCs, suggesting that AChE
expression appears to be an important determinant for the maturation of
synaptic transmission at NMJs but is not the only limit to mEPC decay.
The limited contribution of AChE to shaping mEPCs may have contributed
to the non-Gaussian amplitude histrograms observed both in embryos and
larvae if the prolonged presence of ACh in the synaptic cleft produced
broader and more variant patterns of AChR activation. AChR clusters
increase in number and size early during synaptogenesis (Liu and
Westerfield 1992). Together with an increase in AChR density
within clusters this could account for both the increase in mEPC
amplitude and kinetics. Alternatively, presynaptic differentiation may
determine the development of mature endplate properties in the
zebrafish. It has been shown that the growth cones of
Xenopus motoneurons in culture release ACh before contacting
a muscle cell (Hume et al. 1983
; Young and Poo
1983
). Contact enhances the rate of release of ACh (Sun
and Poo 1987
; Xie and Poo 1986
), and the
kinetics and frequency of mEPCs increase with time in culture
(Evers et al. 1989
). These findings with cultured cells
suggest that an extended diffusion barrier may exist at newly formed
zebrafish NMJs in vivo and may result in delayed and prolonged
activation of AChRs. Presynaptic contact with muscle cells could
culminate in tight, restricted apposition of the motor nerve terminal
to the muscle cell, and a higher density of AChE would accelerate the
mEPC decay rate to that observed by 3 days. Regardless of the
structural changes occurring during early development of the NMJ, our
results indicate that the same AChRs are present in the embryo and
larva and that AChE expression is not the only limit to maturation of
the mEPC.
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ACKNOWLEDGMENTS |
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This work was supported by the following salary awards: a Medical Research Council Centennial Fellowship to P. V. Nguyen, a Fonds de la Recherche en Santé du Québec (FRSQ)-Institute National de la Santé et de la Recherche Médicale Visiting Fellow to L. Aniksztejn, and an FRSQ Senior Research Scholarship to P. Drapeau. This work was also supported by a grant from the National Sciences and Engineering Research Council of Canada to P. Drapeau.
Present address: P. V. Nguyen, Dept. of Physiology and Division of Neurosciences, University of Alberta, Faculty of Medicine, Edmonton, Alberta T6G 2H7, Canada; L. Aniksztejn, INSERM U29, Hopital de Port-Royal, 123 Blvd. De Port Royal, 75014 Paris, France.
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FOOTNOTES |
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Address for reprint requests: P. Drapeau, Dept. of Neurology, Montreal General Hospital, 1650 Cedar Ave., Montreal, Quebec H3G 1A4, Canada.
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 7 December 1998; accepted in final form 24 February 1999.
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REFERENCES |
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