Centre for Research in Neuroscience, Montreal General Hospital Research Institute; and Department of Neurology and Neurosurgery and Department of Biology, McGill University, Montreal, Quebec H3G 1A4, Canada
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ABSTRACT |
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Buss, Robert R. and
Pierre Drapeau.
Physiological Properties of Zebrafish Embryonic Red and White
Muscle Fibers During Early Development.
J. Neurophysiol. 84: 1545-1557, 2000.
The zebrafish is a model
organism for studies of vertebrate muscle differentiation and
development. However, an understanding of fish muscle physiology during
this period is limited. We examined the membrane, contractile,
electrical coupling, and synaptic properties of embryonic red (ER) and
white (EW) muscle fibers in developing zebrafish from 1 to 5 days
postfertilization. Resting membrane potentials were 73 mV in 1 day ER
and
78 mV in 1 day EW muscle and depolarized 17 and 7 mV,
respectively, by 5 days. Neither fiber type exhibited action
potentials. Current-voltage relationships were linear in EW fibers and
day 1 ER fibers but were outwardly rectifying in some ER fibers at 3 to
5 days. Both ER and EW fibers were contractile at all ages examined (1 to 5 days) and could follow trains of electrical stimulation of up to
30 Hz without fatiguing for up to 5 min. Synaptic activity consisting
of miniature endplate potentials (mEPPs) was observed at the earliest
ages examined (1.2-1.4 days) in both ER and EW fibers. Synaptic
activity increased in frequency, and mEPP amplitudes were larger by 5 days. Miniature EPP rise times and half-widths decreased in ER fibers by 5 days, while EW fiber mEPPs showed fast kinetics as early as
1.2-1.4 days. ER and EW muscle fibers showed extensive dye coupling
but not heterologous (red-white) coupling. Dye coupling decreased by 3 days yet remained at 5 days. Somites were electrically coupling, and
this allowed filtered synaptic potentials to spread from myotome to
myotome. It is concluded that at early developmental stages the
physiological properties of ER and EW muscle are similar but not
identical and are optimized to the patterns of swimming observed at
these stages.
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INTRODUCTION |
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Two types of muscle fibers are
easily recognized in most fishes (Greer-Walker and Pull
1975) by their characteristic red and white coloration in fresh
specimens (Bone 1978
; Johnston 1981
). A
typical teleost myomere, zebrafish included, has a superficial band of
red muscle fibers that runs parallel to the rostral-caudal axis of the
body and deeper layers of white muscle that run at an oblique angle to
this axis (Alexander 1969
; Van Raamsdonk et al.
1979
). Microscopic, immunological, and histochemical
examination reveals a greater diversity of fish muscle fiber types. The
myotome of the adult zebrafish is composed of five distinguishable
layers of muscle cells: 1) the superficial adult red layer,
2) the intermediate pink layer, 3) the deep white
layer, 4) the scattered dorsal and ventral fiber layer, and
5) the red muscle rim layer (Van Raamsdonk et al.
1978
, 1980
, 1982a
,
1987
; Waterman 1969
). At one extreme, superficial red fibers have small diameters, contain many mitochondria, have a rich blood supply, and have metabolic specializations, making
them suitable for sustained activity but not rapid contractions. At the
other extreme, each of these characteristics is the opposite for the
deep white muscle fibers, suggesting a rapidly fatiguing muscle capable
of fast and powerful contractions. How these different muscle fibers
develop and their precise functional roles in the embryo are less well understood.
Morphological development of muscle fibers has been studied in
considerable detail in the zebrafish, although the physiological roles
of different muscle fibers have not been directly examined. A
functional characterization of muscle development is important to
understand motor control during normal development and to assess dysfunction in the numerous interesting locomotor mutants that have
been isolated in the first large genetic screens of vertebrate development (Granato et al. 1996).
At a morphological level, a simpler pattern of muscle fibers is
observed in embryonic and young larval zebrafish. Embryonic white (EW)
fibers (lateral presomitic cells) form mediolateral to the notochord
and do not migrate (Du et al. 1997). When embryonic myotomal segments first form, they are block-shaped and composed entirely of EW fibers running parallel to the notochord. Within a few
hours the EW fibers take on their characteristic oblique chevron-shaped
orientations (Van Raamsdonk et al. 1974
). Embryonic red
(ER) fibers (adaxial cells) later form at the midline next to the
notochord and then migrate to the surface of the muscle (Du et
al. 1997
); their differentiation is regulated by the
Hedgehog and TGF-
gene families
(Blagden et al. 1997
; Currie and Ingham 1996
; Du et al. 1997
). A thin band of muscle
consisting of approximately 30 ER fibers per segment blankets these
deep fibers and retains their parallel orientation. An anatomical and
histochemical division into these two fiber types at early
developmental stages is present in a diversity of fish species
(Batty 1984
; Forstner et al. 1983
; Matsuoka and Iwai 1984
; O'Connell 1981
;
Proctor et al. 1980
; Stoiber and Sänger
1996
; Veggetti et al. 1993
), including a number
of cyprinid species related to zebrafish (El-Fiky et al.
1987
; Stoiber and Sänger 1996
). During the
first week of development, the ER and EW fibers are the only muscle
types present. El-Fiky and Wieser (1988)
suggest the ER
fibers are the main organs of gas exchange prior to gill development
and that the deeper muscle layers are involved in locomotion. In later
larval development the ER fibers divide to ultimately form the red
muscle rim fibers (Van Raamsdonk et al. 1979
,
1982b
; Waterman 1969
).
At a physiological level, the first movements (17 h)
(Saint-Amant and Drapeau 1998) are observed shortly
after neuromuscular innervation commences (Liu and Westerfield
1992
), which is when muscle contractile properties emerge
(Van Raamsdonk et al. 1977
). Muscle pioneers are the
first fibers contacted by primary motoneurons, the first to form
functional synapses, and the first to contract (Melancon et al.
1997
). Secondary motoneurons begin to innervate muscle at ~26
h (Myers et al. 1986
), and swimming is first observed soon after at 27 h (Saint-Amant and Drapeau 1998
).
At this period, all fibers have metabolic properties similar to adult
white muscle, i.e., they are fatigable, and it is not until near the
fifth day that the superficial fibers begin to acquire the metabolic
characteristics of adult red muscle (Van Raamsdonk et al.
1978
). However, even at this early age, both fibers have
distinct red and white myofibrillar properties (Blagden et al.
1997
), but their functional roles are unknown. A rapidly
fatigable muscle may be one reason why brief periods of burst swimming,
followed by extended periods of rest (Van Raamsdonk et al.
1974
), are only observed at early larval stages. Rapidly
fatiguing swimming in early larval life appears to be a consequence of
muscle metabolic properties in a variety of cyprinids (El-Fiky
et al. 1987
) and whitefish (Forstner et al.
1983
).
The goal of this study is to compare the physiological properties of, and synaptic inputs to, developing ER and EW muscle fibers of the zebrafish. Developmental changes in the properties of these muscles are examined from when swimming behavior is first manifested (day 1) to when zebrafish actively swim to capture prey (day 5).
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METHODS |
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Preparation
Experiments were performed on zebrafish (Danio rerio)
embryos and larvae of the Longfin strain raised at 28.5°C and
obtained from a breeding colony maintained according to
Westerfield (1993). Results are taken from recordings
made on 84 muscle fibers. All procedures were carried out in compliance
with the guidelines stipulated by the Canadian Council for Animal Care
and McGill University. Zebrafish were anesthetized in 0.02% tricaine
(MS-222) dissolved in physiological extracellular Evans
(1979)
solution consisting of (in mM) 134 NaCl, 2.9 KCl, 2.1 CaCl2, 1.2 MgCl2, 10 HEPES,
and 10 glucose (Drapeau et al. 1999
), osmolarity
adjusted to ~290 mOsm, pH 7.8. The animal was then pinned through the
notochord to a silicone elastomer (Sylgard)-lined dish and the skin
overlying the axial musculature removed with a glass pipette and fine
forceps. The preparation was moved to the recording setup and then
continuously perfused with a tricaine-free Evans solution containing 15 µM D-tubocurarine or 1 µM tetrodotoxin (TTX) to
paralyze the animals. In dye coupling measurements (n = 25) a high magnesium and low calcium extracellular Evans solution (in
mM: 123 NaCl, 2.9 KCl, 0.7 CaCl2, 10 MgCl2, 10 HEPES, and 10 glucose, osmolarity
adjusted to ~290 mOsm, pH 7.8) was used instead of a paralyzing
agent. Although these fibers were not used for electrophysiological
measurements, robust spontaneous synaptic activity was observed in all
25 cells. Recordings made in this solution were very stable due to an
absence of muscle contractions, and for this reason this solution was used during paired recordings (n = 5).
Whole cell recordings
Standard whole cell recordings (Hamill et al.
1981) were performed on the superficial ER fibers and the first
two layers of EW fibers (van Raamsdonk et al. 1982b
). ER
and EW fibers correspond to the adaxial and lateral presomitic cells of
Devoto et al. (1996)
and Du et al.
(1997)
. All physiological measurements were performed on dorsal
and ventral fibers located one to two segments rostral or caudal to the
anus. A patch pipette controlled by a micromanipulator was used to
tease off one or two overlying ER fibers to expose the underlying EW
fibers. Muscle fibers were visualized with Hoffman modulation optics
(×40 water immersion objective). Experiments were performed at room
temperature (22°C). Patch-clamp electrodes were pulled from
thin-walled Kimax-51 borosilicate glass and were filled with either a
K-gluconate (for physiological measurements), Cs-gluconate (for paired
recordings), or CsCl (for dye coupling measurements) solution to yield
electrodes with resistances of 1.5-3 M
as described previously
(Drapeau et al. 1999
). The K-gluconate solution was
composed of (in mM) 116 D-gluconic acid-potassium salt, 16 KCl, 2 MgCl2, 10 HEPES, 10 EGTA, and 4 Na2ATP, osmolarity adjusted to 290 mOsm, pH 7.2. K-gluconate and KCl were replaced with Cs-gluconate and CsCl in the
Cs-gluconate solution or with CsCl in the Cs-chloride solution. The
liquid junction potential was
5 mV for K-gluconate and Cs-gluconate
electrodes and
2 mV for CsCl electrodes when measured in Evans
solution, and records were corrected for this potential. Recordings
were made with an Axoclamp-2A amplifier (Axon Instruments) in bridge
mode and were low-pass filtered at 10 kHz and digitized at 20-30 kHz.
In paired recordings an Axopatch 1D (Axon Instruments) was used as a
second amplifier. Cells were discarded if the resting membrane
potential was more depolarized than
50 mV. Measurements were made
3-5 min after obtaining the whole cell configuration to ensure cell dialysis.
Dye coupling between muscle fibers
All fibers reported in this study were filled with 0.2% fluorescent sulforhodamine B to examine the extent of dye coupling between muscle fibers, and electrophysiological measurements were taken from these same fibers. Images were captured with a Panasonic BP510 charge-coupled device (CCD) camera and a Scion Corporation LG3 frame grabber using Scion/NIH Image software. Whole cell configuration was maintained for 10-15 min in each fiber to allow complete equilibration of the dye. Dye coupling was quantified by counting the number of fluorescent fibers and the number of segments that contained fluorescent fibers.
Current-voltage relationships and membrane time constants
Recordings were made in D-tubocurarine to eliminate
CNS evoked muscle contractions. Current steps were manually incremented through an A-M Systems isolated pulse stimulator, captured using Axotape software (Axon Instruments), and measurements were made in
Axoscope 7 software (Axon Instruments). Current injections of 300-ms
duration were made once every 2-4 s. A small hyperpolarizing or
depolarizing current was injected and constantly adjusted to maintain a
membrane potential of 65 mV in 3- and 5-day animals. Some embryonic
muscle fibers were unstable at
65 mV, so all recordings were
performed at
75 mV, which is closer to the resting membrane potential
at this age. During recordings, the electrode response was constantly
monitored at high gain and high sweep speed on an analog oscilloscope
and the electrode balanced using the bridge circuitry. Electrode
resistance was generally low (3-5 M
) but was as high as 10 M
in
some recordings, and the electrode capacitance was <3 pF. Both the
electrode resistance and capacitance were an order of magnitude smaller
than those for the muscle membrane, resulting in small and fast
transients that were easily compensated and unlikely to contribute to
measurement errors.
Membrane time constants were determined by fitting the voltage response to a hyperpolarizing current injection (15- to 25-mV deflection) with a sum of exponential curves. The presence of one or two exponential components was tested by comparing the sum of squared errors of the fits.
Muscle contraction rates
In many muscle fibers, it was possible to inject sufficient
current in the center of the fiber to evoke a muscle contraction and
still maintain the whole cell configuration. Following voltage recordings, the current pulse duration was reduced to 15-20 ms and the
intensity increased to 2-7 nA, which was sufficient to evoke a muscle
contraction. A duration of 15-20 ms was chosen to mimic the cycle
duration of motoneuron output that must be occurring for the larvae to
swim at the observed tail beat frequencies of approximately 30-60 Hz
(Budick and O'Malley 1999; Eaton et al.
1977
; Saint-Amant and Drapeau 1998
). Briefer
durations of current were usually ineffective regardless of the
intensity of stimulation up to 10 nA. The contraction of the muscle was
observed on a video monitor and recorded on a VCR (Panasonic S-VHS)
along with stimulus parameters, which were recorded on Axotape for
later analysis. Inter-pulse interval was gradually decreased until the individual contractions fused together, which is referred to as the
tetanus fusion rate. Long (300 ms) depolarizing current injections were
used to determine the rheobasic contraction threshold, which was
recorded as the membrane potential at which the first twitch contraction was observed.
Analysis of miniature endplate potentials (mEPPs)
mEPPs were recorded in Evans solution containing TTX to block
CNS-evoked muscle contractions but not spontaneously occurring synaptic
vesicle release. A small hyperpolarizing or depolarizing current was
injected into the fiber to hold the membrane potential at 75 mV
(day 1) or
65 mV (days 3-5). Five-minute
recordings were made in Fetchex (Axon Instruments), and analysis was
performed off-line using Axograph 4.0 (Axon Instruments). Events were
detected using the template function set at 4-5 SD above baseline
noise. It was not possible to discriminate events <0.5 mV in amplitude due to the background noise even with Cs-gluconate pipettes. Small events were excluded followed by visual examination and deletion of
erroneous events. The remaining events were used to calculate peak
amplitudes, 20-80% rise times, half-widths, and mEPP frequencies.
Paired recordings
Paired recordings were performed in a high magnesium (10 mM) and low calcium (0.7 mM) solution, which abolished muscle contractions and allowed stable recordings with two electrodes. A Cs-gluconate intracellular solution was used to potentiate mEPPs and facilitate their detection. These fibers were not used for calculating any parameters listed in Tables 1 or 2. All paired recordings were made from longitudinally adjoining ER fibers of day 3 animals. Positive or negative current was ejected through the recording electrodes to equalize the membrane potential of each cell.
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The goal of this experiment was to determine whether the small and slow mEPPs occurred simultaneously with large and fast mEPPs in adjacent fibers, the small and slow events being due to filtering through intercellular junctions. Digitized data were examined by scrolling through 50-ms windows in Axoscope 7. Whenever an event was detected (>0.5 mV) in one fiber, a simultaneous corresponding event was searched for in the other fiber. This preliminary analysis revealed that events large enough to be detectable, i.e., >0.5 mV, always occurred simultaneously in both fibers with one mEPP appearing as a filtered and attenuated version of the other mEPP. Time-to-peak and peak amplitude measurements were then made by eye for 50 consecutive paired mEPPs. The peak amplitude and time-to-peak values of the larger mEPP was divided by the peak amplitude and time-to-peak value of the smaller mEPP and was normalized to 100% to express the extent of filtering between each pair of mEPPs.
Results are presented as means ± SE, and significant denotes a significant relationship (P < 0.05) determined using the Student's t-test or Mann-Whitney rank sum test. Correlations were tested using the Pearson Product Moment Correlation, and a significant relation was noted when P < 0.05. Sulforhodamine B was purchased from Molecular Probes (Eugene, OR) and all other chemicals from Sigma Chemical (St. Louis, MO).
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RESULTS |
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To avoid the errors inherent with voltage-clamp recording from
electrically coupled muscle cells (e.g., Broadie 1999;
Nguyen et al. 1999
), experiments were performed in
current-clamp mode. Muscle fibers were examined at three time periods
of development (Kimmel et al. 1995
): in embryos
(day 1.2-1.5; referred to as day 1) near the
time swimming-like behavior is first observed (Saint-Amant and
Drapeau 1998
), in quiescent posthatching larvae (day
3.4-3.6; referred to as day 3) and in active
free-swimming larvae (day 4.9-5.8; referred to as day
5). As the appearance of the two types of muscle fibers did not
change over the brief (4 day) interval examined, we refer to them as ER
or EW muscle fibers. Resting membrane potentials recorded with
K-gluconate intracellular solutions (Table 1) were
73 ± 2 (SE) mV in ER and
78 ± 1 mV in EW fibers and
depolarized by 7 mV (EW) and 17 mV (ER) during development to the free
swimming larval stage (day 5). At all stages examined, the
resting membrane potentials of the EW fibers were more negative than
those of the ER fibers. By day 5 the difference in resting
membrane potential between EW and ER fibers was 15 mV.
Dye coupling
Dye coupling was observed in 69 of 74 muscle fibers examined in this study, the exceptions being 5 EW fibers at 3-5 days. No obvious differences in dye coupling were observed in rostral or caudal segments (±10 segments from the anus) or in dorsally or ventrally located fibers. Dye coupling in ER fibers spanned up to five segments (only 3 segments are shown in Fig. 1) at all stages, which included up to 38 labeled fibers at day 1 and 12 at days 3-5. The average number of dye-coupled segments decreased by 38%, and the number of fibers filled decreased by 73% (Table 1) during the transition from day 1 to day 3, and there was no further change by day 5. Fibers were dye-coupled in both dorso-ventral and rostro-caudal directions (Fig. 1, A2 and B2).
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Dye coupling was less extensive in the EW than in ER fibers at all corresponding stages examined (Fig. 1; Table 1) but showed a similar decrease in segmental coupling (by 39%) and decrease (by 70%) in the number of fibers coupled from day 1 to day 3, and, similar to the ER fibers, no further change in coupling was observed by day 5. Unlike ER fibers, coupling in the EW fibers was generally restricted (21 of 26 fibers) to the rostro-caudal axis in larvae where 1-4 segments could contain dye-filled fibers. Dorso-ventral coupling in addition to rostro-caudal coupling was observed in day 1 EW fibers where 3-13 fibers were found coupled over 3-5 segments at later stages. Coupling between EW fibers running in different oblique orientations (Fig. 1A3) was observed occasionally. However, coupling was never observed between ER and EW fibers.
Current-voltage (I-V) relationships
The I-V relationship of each type of muscle fiber
was examined to reveal any differences or changes in the membrane
properties during development of the ER and EW fibers. At day
1, ER fibers had mean input resistances of 31 ± 2 M
(Table 1) and showed no rectification, and the time course of the
voltage response was similar in both depolarizing and hyperpolarizing
directions (Fig. 2A1). The
mean input resistance of day 3 ER fibers increased to
74 ± 10 M
(Table 1) when hyperpolarizing currents were
injected. In some cells outward rectification was observed with
depolarizing currents, and the initial voltage response was steeper in
the depolarizing direction (Fig. 2A2). Interestingly, the
input resistance at day 5 decreased to 30 ± 5 M
, a
value similar to that observed at day 1. It is likely that
the initial increase in input resistance from day 1 to
day 3 was principally due to muscle fiber uncoupling, and
the decrease in input resistance observed from day 3 to
day 5 was due to insertion of voltage-gated ion channels
into the membrane. Similar to day 3, some day 5 fibers rectified and showed a steeper initial voltage response in the
depolarizing direction (Fig. 2A3; Table 1). Action
potentials were not observed in any ER fibers.
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Day 1 EW fibers had a mean input resistances of
33 ± 6 M (Table 1) and showed a similar voltage response in
both depolarizing and hyperpolarizing directions (Fig. 2B1).
Similar to ER fibers, the input resistance in day 3 EW
fibers was slightly higher (46 ± 8 M
) and then decreased to
27 ± 5 M
by day 5 (Table 1). Rectification was not
observed in any EW fibers. The initial voltage response was steeper in
the depolarizing direction than the hyperpolarizing direction in
day 3 EW fibers (Fig. 2B2), but this was not
apparent in day 5 EW fibers (Fig. 2B3). No action
potentials were observed in any EW fibers.
Membrane time constants
Membrane voltage responses to 300-ms hyperpolarizing current
injections were fit with the sum of two exponential curves with membrane time constants fast and
slow. At all ages and in both ER and EW
fibers,
fast mean values ranged from 0.5 to
4.4 ms, and
slow means ranged from 16 to 76 ms
(Table 1). At day 1, EW fibers had a faster
fast (0.5 ± 0.2 ms) than that of ER
fibers (3.2 ± 0.8 ms). The
fast exceeded
the membrane time constant and may reflect an active conductance, or
the presence of unfused myoblasts as observed previously (Nguyen
et al. 1999
). The
slow was very
similar (ER = 71 ± 3 ms vs. EW = 76 ± 4 ms; Table
1). However,
slow contributed to 75% of the
response amplitude in ER fibers, whereas in EW fibers only 21% of the
response amplitude was due to
slow. This
difference might be explained by the more extensive dye coupling
observed in ER fibers. At day 3,
slow decreased similarly in EW and ER fibers
to 40 ± 9 and 39 ± 11 ms. The contribution of
slow to the peak amplitude changed little (21 to 19%) in EW fibers, while it decreased from 75 to 36% in ER fibers.
These changes in membrane time constants closely parallel the changes
in dye coupling observed at these same ages. Accordingly,
fast likely corresponded to the membrane
response of the fiber recorded from and perhaps immediately adjacent
fibers (immediate compartment), whereas
slow
was the response of more distant coupled muscle fibers (remote
compartment). However, changes in fiber size and channel composition,
which were not examined in this study, are also likely to influence the
changes in time constants observed. By day 5,
slow changed little in EW fibers (36 ± 7 vs. 40 ± 9 ms) and only contributed to 12 ± 1% of the
amplitude of the voltage response. At this age
slow contributed to 49% of the amplitude of
the voltage response in ER fibers, which by this time had developed a
significantly faster
slow (16 ± 3 vs. 36 ± 7 ms) than day 5 EW fibers.
Contraction thresholds and maximal contraction rates
During the I-V recordings (300-ms current pulses) the
membrane depolarization required for muscle to twitch was noted. The voltage threshold for muscle contraction changed little in day 1-5 ER fibers [means varied from 42 ± 4 mV to
48 ± 7 mV (Table 1)]. In contrast, EW fibers did not contract in
response to long depolarizing current that depolarized the membrane to
35 mV. Larger current injections that depolarized the membrane well
above 0 mV resulted in robust muscle twitches and the loss of whole cell configuration. It was apparent that EW muscle did not respond well
to focal stimulation, which was not unexpected in these unexcitable and
multiply innervated muscle fibers.
Following I-V recordings, stimulus parameters were adjusted
to resemble more closely the currents the muscle fibers would presumably receive during swimming at maximally observed contraction rates. The electrode seal was lost in several fibers once muscle contractions started, but recordings were relatively stable in others.
The local (point) current necessary to evoke a muscle contraction
depolarized the membrane well above 0 mV and out of physiological
range, providing further evidence for the requirement for distributed,
multi-terminal innervation to evoke muscle contraction. Of the
successful recordings, ER and EW fibers at all stages were capable of
following trains of 15-20 ms stimulation delivered at frequencies up
to 30 Hz (observed in a day 5 ER fiber), which predicts a
sustainable upper limit of 30 Hz for alternating tail beat. Thirty
hertz is below the maximum tail beat frequencies (~50 Hz)
(Budick and O'Malley 1999) reported in larval
zebrafish. This discrepancy is likely because our study measured
sustained muscle contraction rates and cannot rule out the possibility
that more rapid muscle contraction rates, which fatigue within a few muscle contractions, are possible. The average muscle tetanus fusion
rates (Table 1) were fastest at day 5 and appear slower at
day 1, which correlates well with the gradual increase in
swimming tail beat frequency observed in developing zebrafish
(Budick and O'Malley 1999
; Eaton et al.
1977
; Saint-Amant and Drapeau 1998
). Stimulation
was generally delivered for 2-5 min, during which time muscle fatigue
was not observed.
mEPPs
Figure 3, A-D, shows representative examples of mEPPs recorded in day 1 and day 5 muscle fibers. Readily apparent are the higher frequency of events at day 5 versus day 1 and the prominence of large amplitude events at day 5 compared with smaller event amplitudes at day 1. Representative mEPPs are shown on an expanded time scale in Fig. 3, Ai-Di. At day 1 mean mEPP amplitudes were similar in ER (0.8 ± 0.1 mV) and EW (0.87 ± 0.04 mV) and then increased ~3 and ~5 times, respectively, by day 5 (Table 2). The mEPP time courses were highly similar in day 5 ER and EW fibers and in day 1 EW fibers, but the time courses were distinctly slower in day 1 ER fibers. Mean rise times and half-widths were 4 and 6 times faster, respectively, in day 1 EW fibers than they were in ER fibers (0.58 ± 0.06 ms vs. 2.3 ± 0.2 ms rise times and 3.8 ± 0.1 ms vs. 24 ± 2 ms half-widths; Table 2). In contrast, by day 5 these values were comparable (0.33 ± 0.05 ms vs. 0.39 ± 0.04 ms rise times and 3.5 ± 0.2 ms vs. 3.6 ± 0.2 ms half-widths; Table 2). The frequency of detectable mEPPs increased in both ER and EW fibers from day 1 to day 5, but this increase was greatest in EW fibers where it increased 10 times compared with ER fibers where the frequency increased only 1.5 times. Specifically, day 1 ER fibers received a higher frequency of mEPPs (0.38 ± 0.04 Hz) than EW fibers (0.10 ± 0.02 Hz), but this difference was reversed by day 5 when ER fibers received mEPPs at 0.57 ± 0.11 Hz compared with frequencies of 1.00 ± 0.24 Hz observed in EW fibers. Differences in passive membrane properties did not account for slow events in day 1 ER fibers. Even though day 1 ER fibers had slower time constants than day 1 EW fibers, these values (as well as the resistances) were more similar to those of both types of fibers at day 5 (Table 2). Therefore the kinetics of mEPPs recorded in day 1 ER fibers were fundamentally slower than those of day 1 EW fibers and of both fiber types at later stages.
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The properties of mEPPs were further examined by comparing the distributions of event amplitudes, rise times, and half-widths. Representative mEPP distributions from a day 1 ER and EW fiber are shown in Fig. 4. The amplitude distributions were very similar in the day 1 ER and EW fibers and appeared to be normally distributed, but the rise time and half-width distributions are clearly wider and displaced to the right in the ER fibers. Scatter plots of event rise times versus amplitude and half-widths (Fig. 5) do not reveal more than one population of events and further illustrate the difference between mEPPs in day 1 ER and EW fibers.
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mEPP distributions appeared normally distributed and were similar in day 5 ER and EW fibers. Thus amplitude, rise time, and half-width distributions are only shown for EW fibers (Fig. 6). Unlike day 1 fibers, some day 5 fibers displayed one population of mEPP amplitude, rise time, and half-width distributions (Fig. 6A), while others displayed two distributions (Fig. 6B). This was also apparent in scatter plots of rise times versus amplitudes (Fig. 7, A1 and B1) or half-widths (Fig. 7, A2 and B2). The distribution common to all fibers was composed of events with large amplitudes (>1-2 mV), fast rise times (<0.2-0.4 ms), and short half-widths (<4-5 ms; Fig. 7A). In the majority of cells (3 of 5 ER; 3 of 4 EW) a second population of events with smaller amplitudes (<1-2 mV), slower rise times (>0.2-0.4 ms), and longer half-widths (>4-5 ms) were observed (Fig. 7B). Closer examination of the fibers where this second population of small and slow events was observed revealed a lower baseline noise in these fibers. Thus two populations of events may be present in all day 5 fibers, but baseline noise prevents the detection of these smaller and slower events. No clear differences could be detected in the amplitude distributions of day 5 ER and EW fibers, and mean mEPP amplitudes were not significantly different in EW (3.7 ± 0.84 mV) and ER (2.3 ± 0.43 mV) fibers.
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Evidence for the filtered propagation of mEPPs between coupled muscle fibers
A simple explanation for the two types of events is that the population of large and fast mEPPs originated from synapses located on the cell recorded from, whereas the population of smaller and slower mEPPs would represent mEPPs originating in electrically coupled neighboring fibers that were recorded as filtered mEPPs. To test this possibility, simultaneous recordings were made from adjacent cells, and the presence of a large, fast event in one cell occurring simultaneously with a small, slow event in the other fiber was investigated. A total of five paired recordings were performed on adjacent ER fibers in day 3 larvae. A pair of dye-coupled fibers is shown in Fig. 8, A and B, and the corresponding paired recording is shown in Fig. 8C. Apparent is the presence of a large and fast event recorded form one electrode paired with a smaller and slower event recorded with the second electrode (Fig. 8, C and D). The larger and faster event was observed with equal occurrence in either of the electrodes. Fifty consecutive mEPPs were observed in each of the five paired recordings and time-to-peak and peak amplitude measured. In every instance, a larger and faster event occurred simultaneously with a smaller and slower event. On average the amplitude of filtered events was half (45 ± 7%) as small and the time-to-peak was twice (2.0 ± 0.1) as slow. In addition, a trend observed (r = 0.64) was that the events with the least attenuated amplitudes also showed the smallest increase in time-to-peak. These were events where the lowest amplitude was small to begin with (e.g., Fig. 8, C and D, 3 and 5). Conversely, events with the greatest amplitude attenuation showed the greatest increase in time-to-peak. These events resembled 2 of Fig. 8, C and D, where the large amplitude event was big to begin with. Thus the electrical coupling acted as a low-pass filter, strongly attenuating the synaptic events from immediately adjacent fibers while only weakly attenuating synaptic events from more distant cells.
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DISCUSSION |
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This is the first study examining the physiological development of
red and white fish muscle. At all stages examined, the resting membrane
potential of the ER fibers was more depolarized than that of the EW
fibers. A more depolarized resting potential in fish red muscle has
been reported in a variety of fishes (Andersen et al.
1963; Hidaka and Toida 1969a
; Stanfield
1972
; Takeuchi 1959
;
Teräväinen 1971
). The absolute membrane
potential of the EW muscle at the latest stage examined (day
5) was
71 mV, which contrasts with a mean resting potential of
82 mV in zebrafish adult white muscle (Westerfield et al.
1986
).
The input resistance of adult red muscle is generally lower than that
of white muscle (Alnaes et al. 1964; Hidaka and
Toida 1969a
; Klein and Prosser 1985
;
Nicolaysen 1976a
,b
; Stanfield 1972
; Teräväinen 1971
), while ER and EW fiber
resistances were similar at the same stages. The red and white muscle
of adult fish shows rectification in some species of ray finned fishes
but not in the silver carp (Eugene and Barets 1982
;
Hagiwara and Takahashi 1967
; Hidaka and Toida
1969a
), cartilaginous (Hagiwara and Takahashi 1967
), or jawless fishes (Alnaes et al. 1964
).
The I-V curves of day 5 EW muscle were linear,
while some day 3 and 5 ER muscle showed outward
rectification. The more extensive coupling in ER fibers may have
contributed to the rectification as injected current could have spread
to other cells.
Action potentials were not observed in the ER or EW muscle fibers.
However, the absence of action potentials does not rule out the
existence of voltage-gated channels, which is suggested by the presence
of a faster rising membrane response to positive, compared with
negative, current injections in both ER and EW fibers (Fig. 2). The
white fibers of adult fish generate action potentials that may or may
not be overshooting, whereas adult red fibers in a wide variety of
fishes are incapable of generating action potentials (Andersen
et al. 1963; Eugene and Barets 1982
;
Gainer 1967
; Hidaka and Toida 1969a
;
Hudson 1969
; Takeuchi 1959
;
Teräväinen 1971
), including adult zebrafish
(Westerfield et al. 1986
). Although both fiber types had
distinct characteristics at day 1 and day 5, their resting potentials, input resistances and excitatory properties
were not comparable to those reported in other adult fish, suggesting
that the complement of ion channels and pumps is not fully expressed at
these stages.
Intercellular coupling between developing muscle cells has been
reported in mammals, birds, amphibia, and fish and is lost with
developmental maturation (Dennis 1981). Intercellular
junctions have been observed in ultrastructure studies of embryonic
zebrafish muscle (Waterman 1969
) and in rainbow trout
(Nag and Nursall 1972
). There was a clear trend toward
fiber uncoupling with age in this study (Fig. 1), although fibers were
not fully uncoupled at the latest stages examined. It cannot be assumed
that all fibers are fully uncoupled in adult zebrafish because in other
fish muscle fibers remain coupled into adulthood (e.g., the lamprey)
(Teräväinen 1971
).
If we assume that the extent of detectable fiber coupling was limited
by diffusion barriers, then it appears that the cytoplasm of all
myotomal muscle (ER and EW) is connected at these developmental stages.
The absence of coupling between ER and EW fibers could restrict
diffusion of intracellular factors involved in muscle differentiation
and growth to these distinct fiber types. No evidence was found to
suggest electrical coupling between motor axons and muscle
(Dennis 1981) as dye did not pass from muscle into
motoneuron axons and the bursting patterns of action potentials
observed in motoneuron recordings (Drapeau et al. 1999
)
were not observed as patterns of depolarization in curarized muscle.
With the exception of the lamprey (Teräväinen
1971), the white muscle fibers of the typical teleost will
contract in response to sub-threshold synaptic stimuli (Bone
1964
; Hagiwara and Takahashi 1967
; Hidaka
and Toida 1969a
). The maximal contraction rates (mean rates of
23-27 Hz, Table 1) of both the ER and EW fibers changed little during
day 3 to day 5, although contraction rates may
have been slower at day 1. These contraction rates are
within the range of contraction rates observed in other fish myotomal
muscle that range from 5 to 60 Hz (Altringham and Johnston
1988
; Johnston 1980
). The maximal contraction
rates observed in the ER and EW muscle are consistent with swimming
cycle periods observed behaviorally (Budick and O'Malley
1999
; Eaton et al. 1977
; Saint-Amant and Drapeau 1998
) and show that neural excitation is optimized to match the contractile apparatus. This finding shows that ER muscle could be co-active with EW muscle at all swimming speeds at the stages
examined. However, this would contrast with the situation in adult
zebrafish muscle where red muscle is believed to be active at slow
swimming speeds and white muscle is recruited during short bursts of
rapid swimming (Liu and Westerfield 1988
).
There have been a limited number of studies examining synaptic
properties in fish muscle. Comparable mEPP amplitudes were observed in
ER and EW fibers at the same stages (~1 mV at day 1 and
2-4 mV at day 5, Table 2). mEPP amplitudes are generally <2.5 mV in silver carp, lamprey, and hagfish (Alnaes et al.
1964; Balezina and Gulyaev 1985
; Hidaka
and Toida 1969b
), while events of 7 mV were found in the fast
contracting sonic muscle of the toadfish (Gainer and Klancher
1965
; Skoglund 1959
). Synaptic events recorded
in fish muscle have rapid decay kinetics, with current half-widths or
time constants no more than 2 ms in duration and mEPP half-widths
ranging from 5 to 10 ms (Alnaes et al. 1964
; Balezina and Gulyaev 1985
; Gainer and Klancher
1965
; Hidaka and Toida 1969b
; Macdonald
1983
; Macdonald and Montgomery 1986
;
Miledi and Reiser 1983
). The decay kinetics of day
5 zebrafish mEPPs were at least as fast having mean half-widths of
<4 ms (Table 2) due to mEPCs with decay time constants of <2 ms
(Legendre et al. 2000
; Nguyen et al.
1999
) and were comparable to the mEPPs recorded in 2-day-old
zebrafish myotomal muscle shown in Fig. 3 of Felsenfeld et al.
(1990)
. Rapid mEPC kinetics are consistent with the fast
contraction rates during swimming as the zebrafish is one of the
fastest swimming teleosts of its size (Plaut 2000
).
In the vast majority of teleosts, including the zebrafish, red and
white muscle receive numerous distributed endings from a plexus of
nerves spread over the surface of the muscle fibers (Altringham and Johnston 1981; Barets
1961
; Bone 1964
; Gainer 1969
;
Gainer and Klancher 1965
; Hudson 1969
;
Ogata 1988
; Shenk and Davidson 1966
;
Takeuchi 1959
; Westerfield et al.
1986
), which contrasts with the terminal innervation of white
muscle fibers in non- and basal-teleost groups (Best and Bone
1973
; Bone and Ono 1982
; Ono
1983
). The function of this multiterminal innervation may be to
allow the presynaptic motoneuron action potential to reach several
points of the muscle fiber faster and with greater efficiency than
would be possible by active or passive conduction along the muscle
fiber if activated by a single terminal junction (Shenk and
Davidson 1966
). Short-duration (<10 ms) current injected through the patch electrode was ineffective at eliciting muscle contraction, and in EW fibers long duration (300 ms) current injections did not evoke a muscle twitch until the membrane potential approached or overshot 0 mV, which is a seemingly unphysiological potential for
the membrane to reach.
In the zebrafish, the biophysical properties of individual
acetylcholine receptor molecules at the neuromuscular junctions are
also optimized to maximize current spread. At these junctions, the
endplate current is generated on reversal from open channel block
following removal of acetylcholine (Legendre et al.
2000). This results in a delayed closing of the acetylcholine
receptors and consequently a transient, rebound synaptic current. The
net consequence is a larger charge distributed over a broader time course, presumably enhancing the distributed depolarization leading to
muscle contraction.
Distinct changes were observed in the properties of mEPPs over the
stages examined. mEPP amplitudes increased in ER and EW fibers,
suggesting an increase in acetylcholine receptor density postsynaptically or an increase in the acetylcholine content of a
transmitter quantum or both. The increase in mEPP frequencies could be
due to an increase in the number of synaptic sites and/or probability
of vesicle release. Liu and Westerfield (1992) reported an increase in the number and size of acetylcholine receptor clusters during synaptogenesis. mEPP kinetics were similar in day 1 EW fibers and day 5 EW and ER fibers but were distinctly
slower in the day 1 ER fibers. Nguyen et al.
(1999)
reported a similar developmental increase in mEPC
kinetics as observed in ER fibers in this study, thus making it likely
that the superficial ER fibers were the fibers examined at the
embryonic stages. The large difference in synaptic kinetics and
frequency of occurrence in ER and EW fibers at the embryonic stages
could be due to immaturity of the ER fibers or differential innervation
by ER and EW muscle by different populations of motoneurons or both.
In adult zebrafish, populations of motoneurons with dorsal and ventral
positions in the spinal cord have been found to innervate white and red
muscle, respectively, with high specificity (Myers 1985;
Van Asselt et al. 1993
; Van Raamsdonk et al.
1983
). The dorsal population of motoneurons undoubtedly include
the primary motoneurons described by Myers et al.
(1986)
, which are the first motoneurons to exit the spinal cord
and form synapses on muscle fibers. In the adult zebrafish there is a
greater diversity of fiber types and most secondary motoneurons
innervate more than one class of muscle, e.g., white and intermediate
(De Graaf et al. 1990
). It is possible that the
outgrowing secondary motoneurons are forming synapses on ER fibers as
well as EW fibers in this period. We suggest that the primary
motoneurons of Myers et al. (1986)
only innervate EW
muscle fibers, and having started synaptogenesis at 17 h
(Liu and Westerfield 1992
), have undergone upward of
10 h of synaptic development by the time of recording. We
therefore hypothesize that both the ER and EW fibers are innervated by
the population of ventral secondary motoneurons (Myers et al.
1986
), which do not exit the spinal cord until ~26 h.
Recordings made on ER fibers could potentially be at the initial period
of synaptogenesis when synaptic currents are expected to have slow
kinetics (Kullberg and Owens 1986
; Kullberg et
al. 1977
; Nguyen et al. 1999
; Schuetze and Role 1987
).
Paired recordings performed on day 3 ER fibers clearly
demonstrate that small amplitude mEPPs with slow kinetics do not all arise as events with distinct kinetics but are due to low-pass filtering through an electrically coupled syncytium of muscle fibers.
All small and slow mEPPs are not necessarily filtered mEPPs, and we
would expect that an unknown fraction of these events (including
undetected events <0.5 mV in amplitude) would arise from newly formed
synapses. Electrical coupling between adjacent muscle fibers
undoubtedly places a limit on the possible movements of the zebrafish
embryo and larva. During swimming, propulsion is generated by
alternating, neural-mediated waves of contractions progressing down the
body of the fish (Lindsey 1978). Electrical coupling
would spread (limited by low-pass filtering) the neurally evoked mEPPs
to adjacent myotomes, depolarizing the membrane closer to contraction
threshold. However, the electrical coupling would limit the degree of
curvature in the body of the larvae that swim with shallow eel-like
movements (unpublished observations), an energetically efficient means
of movement for small larval fish (Batty 1984
). Thus the
physiological properties of larval zebrafish muscle complement the most
energetically favorable form of swimming in this developing fish.
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ACKNOWLEDGMENTS |
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This work was funded by the National Sciences and Engineering Research Council and the Medical Research Council of Canada. R. R. Buss holds an MRC doctoral research award.
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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 (E-mail: mcpd{at}musica.mcgill.ca).
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 16 February 2000; accepted in final form 31 May 2000.
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REFERENCES |
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