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1Department of Molecular, Cellular and Developmental Biology, and 2Cell and Molecular Biology Graduate Program, University of Michigan, Ann Arbor, Michigan 48109
Submitted 23 April 2004; accepted in final form 18 June 2004
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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; Haffter et al. 1996). Genetic analysis has also been applied to study various aspects of behavior and behavioral development in zebrafish (Baier 2000; Granato et al. 1996). In fact, locomotory behaviors such as the escape response and swimming have been well characterized in fish.
The escape response in zebrafish consists of one or more very fast coils immediately after a touch (Eaton and Farley 1975; Saint-Amant and Drapeau 1998). The escape response initially appears at 21 h postfertilization (hpf) and requires the hindbrain (Saint-Amant and Drapeau 1998). After 28 hpf, embryos respond to touch with a bout of swimming with 
50 and 100% of embryos swimming after a tail touch at, respectively, 29 and 36 hpf. As with the early response, swimming after touch requires the hindbrain but not necessarily the more anterior brain regions.
The fact that the early behaviors require only the spinal cord and hindbrain suggests that the neural circuits responsible for these behaviors are localized to these regions of the CNS. Because the early hindbrain and spinal cord contain a relatively small number of neurons, many of which are well-characterized (Bernhardt et al. 1990; Eisen et al. 1986; Fetcho and Faber 1988; Kimmel et al. 1982; Kuwada et al. 1990; Mendelson 1985), these neural circuits are likely to be relatively simple ones amenable to neurophysiological analysis. In fact, zebrafish embryos can be analyzed with electrophysiological methods from the moment that neurons become functional (Buss and Drapeau 2000; Ribera and Nusslein-Volhard 1998; Saint-Amant and Drapeau 2000), and using them on fish, including zebrafish, has elucidated many elements of the neural circuits for the escape response (Faber and Korn 1975; Hatta and Korn 1999; Oda et al. 1995, 1998; Ritter et al. 2001; Takahashi et al. 2002).
The fact that early zebrafish behaviors and their neural circuits are reasonably well characterized and that mutations in these behaviors have been generated suggest that the neurophysiological analysis of these mutants will be useful for understanding the genetic basis for behavior. One such mutation, shocked, was identified by the lack of swimming after touch at 48 hpf (Granato et al. 1996). Normally 48 hpf zebrafish respond to a mechanosensory stimulus with an escape response consisting of a massive contraction of the trunk muscles on one side followed by swimming. sho larvae have normal looking muscles and respond with a tail flip but fail to initiate swimming in response to touch. Here we report that sho embryos are defective in their escape response as well as swimming. Furthermore, using electrophysiological and pharmacological methods we show that motor processing within in CNS is perturbed in sho embryos, suggesting that the sho gene product is required within the CNS for normal network function.
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METHODS |
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Zebrafish were bred and raised according to established procedures (Westerfield 1993), which meet the guidelines set forth by the University of Michigan animal care and use protocols. The developmental staging of the embryos was determined by counting somites as described in Kimmel et al. (1995). The mutant line sho was provided by Dr. Michael Granato at the University of Pennsylvania.
Behavioral observation and video capturing
Embryonic and larval behaviors were observed using a Leica stereo dissection microscope. Mechanosensory stimuli were delivered with a thin tungsten wire probe. Videos were captured using a Panasonic wvbp-330 CCD camera and a Scion LG-3 frame grabber (Scion, Frederick, MD) and analyzed with National Institutes of Health Image on a Mac G4.
Immunohistochemistry
Whole-mount antibody labeling was performed as described in Westerfield (1993). The following antibodies and dilution concentrations were used: F59 (1:10), which labels slow-twitch skeletal muscles (Devoto et al. 1996); anti-acetylated 
-tubulin (1:10,000), which labels axons by recognizing microtubules (Bernhardt et al. 1990); znp-1 (1:100), which labels primary motor axons (Westerfield 1993); zn-5 (1:500), which labels a subset of neurons including cerebellar neurons, hindbrain commissural neurons, branchiomotor neurons, and secondary spinal motor neurons (Chandrasekhar et al. 1997, 1999); zn-12 (1:2,000), which labels the RB peripheral axons (Trevarrow et al. 1990); 3A10 (1:100), which labels the Mauthner cells and commissural primary ascending interneurons (CoPA) neurons (Hatta 1992); anti-SV2 (1:100), which labels synaptic vesicles (Feany et al. 1992); and Mab35 (1:100), which labels nAChRs (Tzartos et al. 1981). The antibody staining was visualized using the Vectastain ABC Kit from Vector Laboratories (Burlingame, CA) and 0.15 mg/ml diaminobenzidine to produce a brown peroxidase reaction product. Molecular Probes (Eugene, OR) Alexa fluorescent anti-mouse and anti-rat antibodies were used as secondary antibodies for the double-labeling experiments. Fluorescent images were collected and processed with a Zeiss confocal microscope (Carl Zeiss, Thornwood, NY) and LSM 500 software. All chemicals were purchased from Sigma-Aldrich (St. Louis, MO) unless noted otherwise. All of the antibodies were purchased from the Developmental Studies Hybridoma Bank (Iowa City, IA), except F59, which was provided by Dr. Frank Stockdale at Stanford University, and anti-acetylated -tubulin from Sigma-Aldrich.
Calcium Green-1 imaging
Muscle cells from 1.5- to 2-day-old embryos were dissociated using a modified trypsin digestion protocol (Westerfield 1993) and cultured for 1 day. Intracellular calcium levels were visualized by adding Calcium Green-1 AM (Molecular Probes) to the bath solution 1–2 h before the experiment to allow sufficient uptake of the dye. The bath solution contained (in mM) 10 HEPES, 145 NaCl, 5 KCl, 1 NaH2PO4, 2 MgSO4, 10 glucose, and 2 CaCl2. Calcium transients were triggered by ejecting carbachol (1 mM, in bath solution) from a micropipette with its tip positioned approximately 100 µm from the muscle cell. The pressure (10 psi) and duration (100 ms) of the ejection were controlled by a Picospritzer II (Parker Hannifin, Fairfield, NJ). The calcium response was captured with a Scion frame grabber and analyzed with NIH Image on a Mac G4.
Dissection
The dissection protocols for in vivo patch clamping have been described previously (Buss and Drapeau 2000; Drapeau et al. 1999; Saint-Amant and Drapeau 2003). Briefly, a zebrafish embryo was anesthetized and paralyzed in recording solution containing (in mM) 134 NaCl, 2.9 KCl, 2.1 CaCl2, 1.2 MgCl2, 10 glucose, 10 HEPES, and 0.0075 d-tubocurarine, 290 mOsm, pH 7.8) that contained tricaine (0.02%). The embryo was immobilized by inserting tungsten wires (25 µm diam) through the notochord. For muscle recordings, target cells were exposed by peeling off the skin overlying several segments. When recording from neurons, the muscle mass was removed by treating embryos briefly with a dilute solution of collagenase (0.02%) followed by gentle suction through a broken pipette to expose the spinal cord.
Electrophysiology and motor neuron recordings
The dissected embryo was under constant perfusion of recording solution (2–5 ml/min). The intracellular solution consisted of (in mM) 116 potassium gluconate, 16 KCl, 2 MgCl2, 10 HEPES, 10 EGTA, and 4 Na3ATP, 273 mOsm, pH 7.2. In experiments that require the removal of K+ conductance, KCl and K-gluconate were replaced with cesium chloride and Cs-gluconate, respectively, in the intracellular solution. A small amount of sulforhodamine (0.2%) was also included in the intracellular solution to facilitate the identification of patched cells. Patch pipettes were pulled from borosilicate silicate glass to yield electrodes with resistances of 6–10 M
for muscle and 10–30 M for neuron. The pipette junction potential has been measured to be 6.4 mV, and all potentials were corrected for this junction potential. Recordings were made with an Axopatch 200 amplifier (Axon Instruments, Union City, CA), low-pass filtered at 5 kHz and sampled at 10 kHz. Data were collected with Clampex 8.2 software (Axon Instruments) and analyzed with Clampfit 9.0 software (Axon Instruments). Fictive swimming was elicited by a sudden change in light intensity or by ejecting bath solution from a pipette with a 15- to 30-µm tip opening. The duration (10–30 ms) and pressure (20–50 psi) of the stimulation were regulated by a Picospritzer.
The primary motor neurons were identified by their position, size, and axonal projections to muscle. The input resistance, Ri, was determined by recording the changes in membrane potential in response to incremental current injections. Each stimulation lasted 300–500 ms and was repeated every 5 s. For the study of action potential properties, step-wise current injections were made near the threshold, and the spike generated from the smallest current stimulation was used for the analysis. For the analysis of current/voltage relationships, the voltage-dependent currents induced by incremental voltage steps were recorded. In these voltage-clamp recordings of the motor neurons, QX-314 (2 mM) was included in the intracellular solution to block the trains of unclamped action potentials that were invariably induced by the suprathreshold depolarizing voltage steps in the absence of QX-314. This also reveals other conductances normally masked by the voltage-dependent sodium currents. Clampfit 9.0 software was used for the analysis of the membrane properties and action potential properties of the motor neurons.
RB recordings
Similar to the motor neurons, we were able to obtain whole cell recordings of the RB neurons. These neurons were identified by their position, size, and axonal projections in the dorsolateral fasciculus (DLF) of the spinal cord. Action potential properties and membrane properties were studied as those in the motor neurons with the exception that QX-314 was not included in the intracellular solution for any recording. At suprathreshold voltage steps, only a single action potential was observed at the onset of the steps instead of the continuous trains of spikes observed in motor neurons.
Miniature endplate potential analysis
Tetrodotoxin (TTX, 1 mM) was added to recording solution lacking d-tubocurarine. Spontaneous release events were detected automatically using the template search function in Clampfit 9.0. Events with amplitudes <0.5 mV and other erroneous events were excluded after visual inspection.
Exogenous muscle stimulation in paralyzed animals
Embryos were prepared as described in the preceding text for physiology. A higher concentration of d-tubocurarine (15 µM) was used to block any CNS-triggered activity. A constant positive pressure was applied to a micropipette containing either caffeine (6 mM) or KCl (20 mM). The resulting stream was targeted to the exposed musculature to induce contractions, which were videotaped. Contractions were also triggered by exogenous depolarization of single muscle cells under whole cell configuration. Steps of depolarizing current (3–6 nA) at varying frequencies (10–40 Hz) were applied to muscle in the current-clamp mode. The resulting contractions were videotaped.
N-methyl-D-aspartate-induced activity
Current-clamp recordings from muscle cells or neurons were maintained for 25 min. During each recording, N-methyl-D-aspartate (NMDA, 10–100 µM) was introduced to the recording solution by perfusion at 5 min and continued for 10 min. At 15 min, TTX (1 µM) was introduced to the bath in addition to NMDA until the end of the recording. For each recording, three intervals were analyzed. Interval 1, from 0–5 min, was the predrug control. Interval 2, 10–15 min, was used to study the effect of NMDA on each cell. Interval 3, 20–25 min, was used to study the TTX-resistant response of the cell to NMDA.
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RESULTS |
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sho mutants, which were originally identified because they failed to initiate swimming after touch at 48 hpf (Granato et al. 1996), are also deficient in the escape response. At 24–26 hpf, sho embryos failed to respond to touch, whereas wild-type embryos responded with an escape response (Fig. 1). Wild-type sibling embryos respond to head touch with one, two, or three vigorous contractions that reorient the embryos in, respectively, 14.6, 66.7, and 18.8% of cases (n = 48) with 98% of multiple contractions alternating from side to side. Tail touch lead to one, two, three, or four contractions that projected the embryos forward in 14.6, 61, 19.5, and 4.9% of cases, respectively (n = 41) with 100% of multiple contractions alternating from side to side. On the other hand, sho embryos fail to respond normally to both head touch and tail touch in all cases (n = 13) at 24–26 hpf. At 48 hpf, wild-type siblings swim away following touch (n = 20), whereas sho embryos respond with a few uncoordinated trunk contractions or not at all (n = 13; Fig. 1).
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Defective distribution or morphology of sensory cells could potentially account for the behavioral defects observed in sho mutants. However, the distribution and morphology of the trigeminal, RB, and posterior lateral line neurons were normal in sho embryos. The distribution of large, Islet-positive RB in the dorsal spinal cord and trigeminal neurons were comparable between mutants [3.2 ± 0.1 (SE) RB neurons per hemisegment, n = 6 embryos] and wild-type sibling (3.2 ± 0.1 RB neurons per hemisegment, n = 5 embryos) embryos at 23 hpf as were the peripheral axons of trigeminal, RB, and posterior lateral line neurons labeled with anti-acetylated--tubulin and MAb zn-12 at 27 hpf (n = 8 for mutant and n = 8 for wild-type sibling embryos; Fig. 2 ). Thus there appears to be no obvious anatomical defects of sensory neurons observable at the light level that can explain the aberrant behavioral responses of the mutant embryos.
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). However, our finding that the threshold for mechanosensory stimulation for responses in muscles at 48 hpf suggests that sensory transduction in RB neurons is not obviously disturbed in sho embryos.
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-tubulin to label the posterior lateral line nerve. In all cases (n = 19), the posterior lateral line nerve had been disrupted by the pinning and eliminated from segments in which the skin had been removed (not shown). Although we could not prove the posterior lateral line is entirely normal, the abnormal muscle response to sensory stimulation cannot be explained by a possible dysfunction in the lateral line due to its removal in our electrophysiological protocols. Muscles in sho embryos are normal
The axial muscles in sho embryos were examined to see if they contributed to the observed behavioral defects. First, we confirmed that the organization and structure of the axial muscles were normal when examined with differential interference contrast optics (not shown) (Granato et al. 1996) and that the expression and distribution of muscle myosin was normal in sho embryos (not shown). Second, the ability of sho muscles to contract was tested pharmacologically and physiologically. Muscles from 48 hpf sho embryos (n = 5) as in wild-type siblings (n = 5) contracted in response to focal application of caffeine (6 mM) from a micropipette (not shown). Because caffeine causes the release of Ca2+ from intracellular stores, the resulting sho muscle contractions suggest that the site of defect is not in the Ca2+-dependent contractile machinery of muscle. The ability of sho muscles to contract upon depolarization of their membrane was examined by focal application of 20 mM KCl. sho muscles (n = 10) as wild-type sibling muscles (n = 5) contracted after application of KCl (not shown). Furthermore, sho muscles (n = 4) as wild-type sibling muscles (n = 2) followed repetitive stimulation by direct injection of a pulse of depolarizing current at frequencies up to 35–40 Hz when the muscles reached tetanus. Thus it appears that the excitation-contraction coupling mechanism involving the sensing of membrane depolarization to Ca2+ release from the sarcoplasmic reticulum (SR) is functional in the mutants. To directly examine if Ca2+ is released properly from the SR of mutants, dissociated muscles were loaded with Calcium-Green-1 AM, a Ca2+ indicator dye, and analyzed after application of an acetylcholine agonist, carbachol, in vitro. As expected from the normal response of muscles to depolarization, sho muscles (n = 6) as wild-type sibling muscles (n = 5) responded to carbachol with an increase in intracellular Ca2+ (Fig. 3). These results suggest that sho muscles respond normally to the neurotransmitter and that the excitation/contraction mechanism is normal. Thus it appears that muscles in sho mutants are normal.
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Because sensory processing and muscle function were not obviously abnormal in sho embryos, outgrowth by motor axons and development of the neuromuscular junction were examined in the mutants. Outgrowth by the presynaptic primary motor axons labeled with diI or MAb znp-1 in sho embryos (n = 10) was normal as was the pattern of neuromuscular junctions (Fig. 4). Simultaneous labeling for both the presynaptic synaptic vesicle antigen, SV-2, and the postsynaptic nicotinic acetylcholine receptors showed that presynaptic and postsynaptic elements were juxtaposed in the axial muscles at the light level and that the pattern of neuromuscular junctions was indistinguishable between mutant (n = 10) and wild-type siblings (n = 10).
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). Both slow and fast twitch muscles contract during swimming (Buss and Drapeau 2002). Slow twitch muscle recordings from partially curarized 48 hpf wild-type sibling embryos (n = 11) showed a long burst (1994 ± 288 ms) of rhythmic depolarizations in response to mechanosensory stimulation of the tail or visual stimulation (light dimming; Fig. 5 ). The frequency of the rhythmic depolarizations (28.6 ± 1.0 Hz) corresponded to the frequency of swimming of 33 Hz observed at 48 hpf (Buss and Drapeau 2001). In 48 hpf sho embryos (n = 11), a much shorter (124 ± 53 ms), irregular depolarization was recorded after mechanosensory or visual stimulation. Thus the slow twitch muscle recordings correlate with the behavioral response at 48 hpf and suggest that there is a defect in the CNS output to the muscles and/or the function of the neuromuscular junction in sho embryos.
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The nature of the CNS defect in sho embryos was investigated by a combination of anatomical and physiological methods. The overall patterning of neurons and axonal tracts analyzed with anti-acetylated--tubulin, MAb zn-5, and MAb 3A10 antibodies that label all or subsets of neurons were normal in sho embryos (not shown). The distribution and development of many spinal and hindbrain neurons such as the Mauthner cell, the command neuron for the escape response, and the CoPA and ventral longitudinal descending interneurons (VeLD) spinal interneurons showed no obvious defects.
Electrophysiological analysis of primary motor neurons demonstrated that the response of primary motor neurons to sensory stimulation was aberrant in sho embryos (Fig. 7). Normally a long burst of rhythmic action potentials initiated from a sustained plateau-like depolarization is recorded in response to touch in wild-type sibling motor neurons (2,360 ± 41 ms, n = 6) at 48 hpf. The frequency of action potentials in the bursts (28.5 + 1.1 Hz) corresponded with the frequency of the swimming of 33 Hz at 48 hpf (Buss and Drapeau 2001). In sho embryos, however, the initial depolarization is not sustained, and only a short burst of action potentials is evident after stimulation (217 ± 53 ms, n = 5). The aberrant response of motor neurons in the mutants correlates with the abnormal response at the neuromuscular junction and suggests that a major component of the defect in sho embryos is due to a defect in the CNS.
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). Comparable afterdischarges can be initiated in both sho and wild-type sibling motor neurons after brief depolarizations from a patch electrode filled with a solution in which K+ was replaced by cesium (Fig. 9). These afterdischarges are blocked by extracellular cobalt (1 mM) in both sho (n = 4) and wild-type sibling (n = 6) motor neurons. This suggests that the Ca2+-dependent currents that underlie the afterdischarge are not compromised by the mutation. Fifth, direct hyperpolarization of the motor neuron during fictive swimming should terminate a plateau potential if it is dependent on the intrinsic properties of motor neurons. On the other hand, if the sustained depolarization requires synaptic input, then hyperpolarizing the motor neurons during a fictive swimming episode should not terminate the depolarization. We found that hyperpolarizations did not terminate mechanosensory-induced depolarization in wild-type motor neurons (n = 4; Fig. 10 ), suggesting that the sustained depolarizations during fictive swimming are not due to intrinsic properties of motor neurons. Thus it appears that the membrane properties of motor neurons are normal and that synaptic input to motor neurons is defective in the mutants.
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The functional defect was further characterized by analyzing the synaptic input to the motor neurons in sho embryos. Because one major deficit is the lack of a sustained depolarization in the mutant motor neurons (see preceding text), the source of the sustained depolarization was examined more closely. Glutamatergic synaptic transmission via NMDA receptors is a good candidate for the generation of sustained depolarizations because they are activated by glutamate only at depolarized membrane potentials and therefore could be responsible for sustaining the depolarization. In fact, in the lamprey, CNS plateau potentials are NMDA dependent (Di Prisco et al. 1997). Indeed APV (50 µM), a NMDA receptor inhibitor, blocked the sustained depolarization in wild-type motor neurons normally induced by mechanical stimulation so that only a short burst similar to that seen in sho motor neurons was evoked (Fig. 11; before APV 1993 ± 494 ms, n = 4; following APV, 248 ± 27 ms, n = 4). Furthermore, application of NMDA (10–100 µM) to embryos induced wild-type sibling motor neurons to depolarize slowly (25 µM NMDA depolarized motor neurons by 5–13 mV, n = 4) and exhibit episodes of plateau-like depolarization, in addition to the slow depolarization, that gave rise to bursts of action potentials. These transient plateau-like depolarizations induced by NMDA could be studied without action potentials in the motor neurons by intracellular application of 2 µM QX-314 from the patch pipette (Fig. 12).
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DISCUSSION |
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The CNS physiology defects exhibited by sho mutants could be due to morphological abnormalities in neurons or to neurophysiological defects within the CNS. Previous analysis of zebrafish behavioral mutants has identified a number of mutations that exhibit aberrant physiology and morphology in the nervous system. space cadet, unplugged, and diwanka embryos exhibit defects in the morphology of specific neurons (Lorent et al. 2001; Zeller and Granato 1999; Zhang and Granato 2000) and abnormal behavior (Granato et al. 1996). On the other hand, touch-insensitive mutants such as macho, alligator, and steifftier exhibit aberrant voltage-sensitive Na+ currents in mechanosensory neurons that result in abnormal behavioral responses (Ribera and Nusslein-Volhard 1998). Mutations have also been identified that affect the function of the neuromuscular junction. One such mutation, twitch once, is a loss-of-function mutation of the gene encoding for rapsyn (Ono et al. 2002).
Our behavioral, morphological, and physiological analyses suggest that the sho gene encodes a product that is required for proper morphological and/or physiological development within the CNS. Although our analysis was far from comprehensive, so far we have not detected any morphological defects. The overall pattern of neurons and axons within the CNS appears normal, and morphological development of early, identifiable neurons such as the RB sensory neurons, CoPA and VeLD spinal interneurons, Mauthner cell, and primary motor neurons appears normal. The apparent normal morphology of several different neurons rules out the possibility that sho encodes for a factor necessary for process outgrowth by all neurons. It is still very possible, however, that a morphological abnormality at the ultrastructural level or in the many neurons not yet examined closely could be the basis for the CNS defect.
Several of our results are compatible with a defect in a NMDA-sensitive process within the CNS of sho embryos. First, application of APV onto wild-type motor neurons phenocopies the mutation. Second, NMDA initiates fictive swimming activity in wild-type but not sho motor neurons. These results also suggest that NMDA is critical for the generation of sustained depolarization during swimming in zebrafish as it is in other organisms (Brownstone et al. 1994; Di Prisco et al. 1997; Schmidt et al. 1998). In fact, several of the key synapses in the Xenopus tadpole swim circuit are known to be glutamatergic (Roberts 2000). In Xenopus, the excitation of two classes of excitatory interneurons in the dorsal spinal cord by RB neurons is primarily mediated by AMPA receptors. These dorsal interneurons in turn activate, via AMPA and NMDA receptors, the interneurons that constitute the spinal pattern generator (Li et al. 2003, 2004). Additionally, the descending interneurons that are thought to provide excitatory synaptic drive for the pattern generator are mediated by NMDA and kainate receptors (Dale and Roberts 1984; Roberts and Alford 1986). Although the swim circuit is not well characterized in zebrafish compared with Xenopus, the multipolar commissural descending interneurons (MCoD) interneurons are active during swimming and likely to be glutamatergic (Higashijima et al. 2003; Ritter et al. 2001) and could potentially be a source of synaptic drive for the swimming pattern generator.
One possibility is that synaptic transmission mediated via NMDA receptors is defective in mutants. sho embryos could be defective in release of glutamate at glutamatergic synapses. However, the fact that exogenous NMDA initiates swimming in wild-type embryos but does not rescue sho embryos suggests that a defect over and above potential defects in glutamate release within the CNS must exist in mutants. Could the defect be one in the NMDA receptor itself? The comparable responses of sho and wild-type motor neurons to direct application of NMDA in the absence of synaptic transmission argue against this point. However, it remains possible that the voltage dependency of NMDA receptors may be defective in mutants.
Another possibility is that the membrane properties of neurons that are activated by NMDA action are aberrant in sho embryos. These could be neurons that directly or indirectly (polysynaptically) respond to NMDA. Analysis of the membrane properties of RB and motor neurons in mutants suggest that at least some of the passive membrane and voltage-gated properties are normal. However, the membrane properties of interneurons that receive sensory information that are presumptive analogues to the dorsal interneurons in Xenopus (Li et al. 2003, 2004; Roberts 2000) and/or those neurons that make up the swimming pattern generator could be defective in sho embryos. In lamprey and Xenopus, spinal neurons exhibit intrinsic, voltage-dependent oscillations of the membrane potential that act in concert with NMDA-mediated activation to mediate swimming (Aiken et al. 2003; Prime et al. 1999; Reith and Sillar 1998; Wallen and Grillner 1987). Motor neurons are thought to be part of the pattern generator in organisms such as Xenopus. At present, however, the status of potential intrinsic oscillatory mechanisms in zebrafish and whether they might be compromised in sho embryos is unclear. Thus sho is not likely to encode for a ubiquitous membrane property protein or a protein that regulates the synthesis, stability, or localization of such a protein found in many neurons including RB neurons but could encode for one expressed in a subset of CNS neurons that are normally activate during swimming. Furthermore, because the escape response and swimming are aberrant in sho embryos, the molecular factor is likely to be common to the neural circuits for both behaviors (Fetcho and Faber 1988; Grillner 2003; O'Malley et al. 1996). Given the similarities of how locomotion is regulated from mammals to lamprey, it would not be surprising if the sho product played similar roles in the generation of synaptic drive to motor neurons in a wide variety of organisms (Grillner 2003; Kiehn and Butt 2003). Ultimately definitive elucidation of the function of the sho product will require the molecular identification of the sho gene.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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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.
Address for reprint requests and other correspondence: J. Y. Kuwada, Dept. of Molecular, Cellular and Developmental Biology, University of Michigan, Ann Arbor, MI 48109-1048 (E-mail: kuwada{at}umich.edu).
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) with a transient elevation of their cytosolic Ca2+ levels. A and C are before carbachol, and B and D after. E (wild-type) and F (sho) are graphs of the relative fluorescence intensities of two representative cells.
), wild-type sibling muscle responded with a burst of rhythmic depolarizations that likely represented swimming. B: sho muscles responded with a short, nonrhythmic depolarization, which correlates with the uncoordinated behavior seen in the mutants.
