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Department of Biological Sciences, University of Alberta, Edmonton, Alberta T6G 2E9, Canada
Submitted 27 June 2003; accepted in final form 12 October 2003
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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-benzyloxyaspartate(200 µM) resulted in a significant increase in the frequency of mEPCs. These results suggest the presence of AMPA and NMDA receptors in association with motoneuron axons of larval zebrafish. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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; Katz 1971). Since this classical description of the NMJ, neurotransmitters other than ACh have been located in motoneurons and at the NMJ (Malomouzh et al. 2003; Pinard et al. 2002; Waerhaug and Ottersen 1993). One neurotransmitter that may be released from the motoneuron terminals along with ACh is glutamate (Barthelemy-Requin et al. 2000; Berger et al. 1995; Malomouzh et al. 2003; Waerhaug and Ottersen 1993). Despite the lack of knowledge surrounding the physiological actions of glutamate at the NMJ, there is a growing body of literature that suggests its presence there (Chen et al. 1998; Fu et al. 1995; Grozdanovic and Gossrau 1998; Liou et al. 1996; Pinard et al. 2002).
Glutamate is the predominant excitatory neurotransmitter in the vertebrate CNS. Glutamate-mediated transmission is regulated by ionotropic and metabotropic glutamate receptors. Ionotropic glutamate receptors (iGluRs) consist of three main groups: N-methyl-D-aspartate receptors (NMDARs), 
-amino-3-hydroxy-5-methyl-4-isoxazole-4-propionic acid receptors (AMPARs), and kainic acid receptors (KARs). NMDARs require glycine as a coagonist, are Mg2+ sensitive and are permeable to K+, Na+, and Ca2+. They are composed of a combination of two NMDAR subunits 1 (NR1) and two NMDAR subunits 2A-D (NR2A-D) (Hollmann and Heinemann 1994), and in some cases, may also contain NR3 subunits (Nishi et al. 2001; Perez-Otano et al. 2001). KARs and AMPARs are often grouped together as one receptor type (non-NMDARs) due to their similar pharmacological and kinetic properties. The majority of AMPA and KA receptors are monovalent cation channels that conduct Na+ and K+, although, some subunit compositions of KA and AMPA receptors are also permeable to Ca2+ (Hollmann and Heinemann 1994). All three receptor types can be found pre- or postsynaptically in the CNS (MacDermott et al. 1999), where they have been reported to be involved in synaptic plasticity phenomena such as long-term potentiation or depression (LTP and LTD) as well as a number of other cellular mechanisms (for review, see Grant and O'Dell 2001; Luscher et al. 2000).
The zebrafish (Danio rerio) has recently gained increasing popularity for developmental studies due to the ease of genetic manipulation, the transparent nature of embryos and larvae, prolific reproduction, and rapid development. Although some developmental changes continue to occur into early adulthood (van Raamsdonk et al. 1983), the motor network is in place by 
30 h postfertilization (hpf) (Drapeau et al. 2002). The body musculature of embryos and young larvae consists of superficial slow red fibers and medial fast white fibers (van Raamsdonk et al. 1978; Waterman 1969). A single layer of red fibers runs parallel to the body while multiple layers of white fibers lie underneath the red fibers at an oblique angle, thereby allowing for visual distinction between the two muscle types (van Raamsdonk et al. 1982). There are two classes of motoneurons within the spinal cord (primary and secondary) that have different functional properties (Liu and Westerfield 1988; Westerfield et al. 1986) and different patterns of innervation (de Graaf et al. 1990; van Raamsdonk et al. 1983). In each body hemi-segment, there are three primary motoneurons and 20–25 secondary motoneurons (Myers et al. 1986; Westerfield et al. 1986). White muscle fibers are electrically coupled (Buss and Drapeau 2000) and polyinnervated by one primary and two to three secondary motoneurons, while red fibers are only innervated by secondary motoneurons (de Graaf et al. 1990; Liu and Westerfield 1988).
Like other vertebrates, neuromuscular transmission in zebrafish occurs through the action of ACh interacting with nAChRs (Liu and Westerfield 1992; Nguyen et al. 1999). The presence of glutamate at vertebrate NMJs has been well established (Berger et al. 1995; Chen et al. 1998; Fu et al. 1995; Grozdanovic and Gossrau 1998; Liou et al. 1996; Malomouzh et al. 2003; Pinard et al. 2002; Waerhaug and Ottersen 1993); however, there is some controversy over whether iGluRs are located on the postsynaptic or on the presynaptic membrane. It has also been suggested that a developmental switch from expression of iGluRs to metabotropic glutamate receptors occurs (Liou et al. 1996). Because the zebrafish is an excellent model to investigate neuromuscular transmission in vivo, we thought its use might help to resolve some of the controversies surrounding the presence of presynaptic iGluRs.
We investigated the presence of presynaptic iGluRs through immunohistochemistry, electophysiological, and pharmacological manipulation of the NMJ. Immunohistochemistry suggests the presence of NR2A subunits in association with the axons of primary motoneurons, whereas electrophysiology suggests a role for glutamate receptors in facilitation of ACh release during early development.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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All use of animals adhered to guidelines established by the University of Alberta and the Canadian Council of Animal Care. Adult fish (Carolina Biological, Burlington, NC) were spawned daily and fertilized eggs were collected and kept at 28.5°C. Prior to all protocols larval zebrafish were anesthetized in 0.02% MS-222.
Immunohistochemistry
Whole zebrafish larvae (4–6 days postfertilization, dpf) were fixed for 2 h in 2% paraformaldehyde and permeablized with 4% Triton X-100 for 30 min. Permeablized animals were incubated in primary antibody (zn-1, mouse anti-zebrafish 1:50, University of Oregon) to identify primary motoneurons, and anti-NR2A (Calbiochem, EMD Biosciences, San Diego, CA) for 48 h at 4°C on a rotary shaker. Fish were washed in phosphate-buffered saline for 3 h every 15 min on a shaker and then incubated in secondary antibody coupled to Alexa568 (goat anti-rabbit 1:1000, Molecular Probes, Eugene, OR) and Alexa488 (goat anti-mouse 1:1000, Molecular Probes) for 4 h at room temperature on a shaker. Fish were washed in phosphate-buffered saline for 3 h every 15 min, then mounted in glycerol for viewing. Imaging was done using a Hamamatsu C4742–95 CCD camera mounted on a Leica DMLFSA epifluorescence microscope with water-immersion x20 and x40 objectives. Image capture was done using Openlab software (Improvision, Lexington, MA).
Immunoprecipitation and immunoblotting
Adult zebrafish brains were rapidly dissected in ice-cold physiological saline, placed in sucrose buffer [containing (in mM) 320 sucrose, 0.01 EDTA, and 10 Tris-HCl, pH 7.4] containing protease inhibitors (3 mM PMSF, 40 µM leupeptin, 4 µM pepstatin A, and 0.4 mg/ml aprotinin) and immediately placed on dry ice. Whole brains were homogenized by hand with an eppendorf mortar and pestle and centrifuged at 1,000 g for 15 min. The supernatant was removed and the pellet was resuspended in sample buffer containing 2% SDS and protease inhibitors and boiled for 10 min to solubilize membrane proteins. Protein quantification was performed using the Lowry Protein Assay (Bio-Rad, Hercules, CA). For immunoprecipitation, the NR2A antibody (8 µg/ml; Calbiochem, EMD Biosciences, San Diego, CA) was added to the resuspended pellet (500 µg protein) and incubated overnight at 4°C, followed by 30 µl of protein A/G PLUS-agarose (Santa Cruz Biotechnology, Santa Cruz, CA) for 1 h at 4°C. Immunoprecipitates were washed, eluted, subjected to SDS-PAGE (8%), according to the method of Laemmli (1970), and transferred to nitrocellulose membranes using a semi-dry transfer apparatus (Bio-Rad, Hercules, CA). Membranes were blocked in blocking buffer (5% skim milk powder, 0.3% Tween-20 in TBS (TBS-T)) for 1 h at room temperature and incubated in anti-NR2A primary antibody (1:800) overnight at 4°C. Membranes were washed several times in TBS-T and incubated in HRP secondary antibody (1:200,000 goat anti rabbit IgG, Santa Cruz Biotechnology, Santa Cruz, CA) for 1 h at room temperature on a shaker. Signals were detected with enhanced chemiluminescence (SuperSignal West Femto, Pierce, Rockford, IL) and developed on X-ray film.
Electrophysiology
Anesthetized 4- to 6-day-old zebrafish larvae were pinned through the notochord to silicon elastomer (Sylgard)-lined recording dishes and perfused with extracellular saline containing (in mM) 134 NaCl, 2.9 KCl, 2.1CaCl2, 1.2 MgCl2, 10 HEPES, 10 glucose, and 0.001 tetrodotoxin (TTX, Tocris, Avonmouth, UK). A section of skin was removed to allow access to the axial muscles. After removal of the skin, red muscle fibers were gently removed with a micropipette. Red fibers run parallel to the body, whereas white fibers lie at an oblique angle, allowing for visual distinction between the two muscle types (van Raamsdonk et al. 1982). Whole cell patch-clamp recordings were performed on axial white fibers in voltage-clamp mode (Hamill et al. 1981). For investigation of (NMDA receptor properties, Mg2+-free extracellular solution was used. The Mg2+-free recording solution consisted of equimolar NaCl replacing MgCl2. The pipette solution contained (in mM) 130 CsCl, 2 MgCl2, 10 HEPES, 10 EGTA, 2 CaCl2, 4 Na2ATP, and 0.4 LiGTP. Polished pipettes were pulled from thin-walled glass (World Precision Instruments, Sarasota, FL) and had tip resistances of 1–2.5 M
, resulting in whole cell series resistance values of 1.5–5 M. Series resistances were monitored carefully during recordings and abandoned if a change of >15% occurred. All series resistances were compensated by 90–95%, and fibers were clamped at –65 mV throughout all recordings. Whole cell recordings were performed with an Axopatch 200B and captured with pClamp 8.1 software. Data were sampled at 100 kHz and low-pass filtered at 10 kHz. Recording analysis was performed using Axograph software.
After rupture of the membrane, recordings were allowed to stabilize prior to initiation of recording. After stabilization, the next 2 min of each recording was used as a measure of the baseline frequency of spontaneous release. Miniature endplate current (mEPC) frequency in the presence of agonists was determined at the visually identified peak frequency or after 6–6.5 min of agonist perfusion. Recordings of mEPCs were analyzed for any kinetic changes associated with agonist perfusion, including rise time, decay time, peak amplitude, proportions of small (<400 pA) and large (>400 pA) events (as determined through amplitude distribution analysis), and frequency of spontaneous release. Kainic acid, AMPA, NMDA, and glutamate (Sigma) were used as agonists. Vehicle controls were completed for 0.1% DMSO. Because of the high stringency of the recording conditions, we were able to maintain high-quality recordings for only 12–15 min, and it was difficult to achieve washout of the agonists in most of our experiments. However, in the few recordings that were held for 20 min, we were able to washout the agonists, and in these cases, we observed a reduction in the mEPC frequency back to approximately baseline levels.
Agonist specificity was investigated through antagonist application. Antagonists were bath applied for 2 min prior to initiation of agonist perfusion. The competitive antagonist D-(–)-2-amino-5-phosphonopentanoic acid (AP-5; 50 µM; Tocris, Avonmouth, UK) was used to block NMDA receptor activation (Ali et al. 2000). Nonspecific [6-cyano-7-nitroquinoxalene-2,3-dione (CNQX), 50 µM; Sigma] and specific non-NMDAR antagonists were used to investigate the contribution of AMPARs and KARs. (2S,4R)-4-methylglutamate (Sym 2081, Sym; 10 µM: Sigma, St Louis, MO) acts specifically on KARs to desensitize them and prevent their activation (Donevan et al. 1998; Zhou et al. 1997). 1-(4-aminophenyl)-4-methyl-7,8-methylenedioxy-5H-2,3-benzodiazepine (GYKI 52466, 50 µM, Sigma) was used as a specific antagonist for AMPARs (Cossart et al. 2002; Donevan and Rogawski 1993). To investigate potential effects of glutamate, the general glutamate transport blocker D-threo--benzyloxyaspartate (D-TBOA; 200 µM; Tocris, Avonmouth, UK) was used (Jabaudon et al. 1999; Waagepetersen et al. 2001). D-tubocurarine (10 µM; Sigma) was used to block nAChRs.
Local perfusion was used to directly apply agonist onto the NMJ. To detect the perfusate, domestic food coloring (5%) was added to the agonist solution. To ensure that the perfusate was removed quickly from the animal a bath flow rate of 3–4 ml/min was used. The perfusion pipette was placed at the horizontal midline of the larvae with bath flow away from the spinal cord. A cone-shaped flow of perfusate was clearly visible flowing ventrally away from the midline. No back-flow over the spinal cord was ever visible.
Statistical analysis
Due to some data sets being nonnormal, nonparametric analyses were used throughout. The Mann-Whitney U test was used to compare all treatments. Significance was assigned where P values were <0.05. All treatments were replicated four to eight times. SigmaStat software was used for all statistical analyses and data were plotted using SigmaPlot software.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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Whole-mount preparations of 4–6 dpf zebrafish reveal peripheral staining in the axial musculature with NR2A antibodies (Fig. 1, A1 and B1; n = 6). To identify which peripheral axons were expressing the NMDAR subunits, we performed double-labeling experiments using the NR2A antibody and the zn-1 antibody that specifically targets primary motoneurons in larval zebrafish (Myers et al. 1986). The NR2A staining appears to be associated with motoneurons, as shown by colocalization with zn-1 antibody staining (Fig. 1, A and B). Staining with anti-NR2A reveals what appears to be one common, ventral exit point from the spinal cord, after which axons project to the dorsal, ventral, and mid musculatures (Fig. 1, A1 and B1). The NR2A staining corresponds well with previously published images of primary motoneuron projections (Behra et al. 2002; Eisen and Melançon 2001; Myers et al. 1986; Westerfield et al. 1986). The three primary motoneurons, caudal (CaP), middle (MiP), and rostral (RoP), project to the ventral, dorsal, and mid axial musculature, respectively (Myers et al. 1986). NR2A staining colocalizes with zn-1 staining in dorsal, ventral and mid musculature, (Fig. 1, A3 and B3), suggesting that NMDARs containing the NR2A subunit are associated with the axons of primary motoneurons.
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; Zhou and Kimelberg 2001), it was possible that the observed staining was associated with Schwann cells along motoneurons. We therefore performed immunostains using anti-NR2A antibodies on animals at an age at which myelination of peripheral axons has not yet occurred (Brosamle and Halpern 2002). NR2A staining in 26 hpf embryos (Fig. 1C) reveals similar staining of projections as seen in older animals, indicating that NR2A subunits are associated with the motoneurons and not Schwann cells. Negative controls were performed by excluding primary antibody from tissue incubations. Incubation in secondary antibodies resulted in dull homogenous staining throughout (Fig. 1D). These controls support the specificity of the primary antibodies used. Antibody staining for other NR2 subunits was inconclusive. In addition we tested antibodies targeted to some AMPA and kainate receptor subunits (GluR1, 2/3, GluR5, 6, 7, and KA2), but the results were also inconclusive.
Immunoprecipitation and immunoblotting
To determine the specificity of the NR2A antibodies in zebrafish, we immunoprecipitated membrane proteins from adult zebrafish brain to first concentrate the protein and then immunoblotted with anti-NR2A. We used adult brains to obtain tissue with high levels of NMDARs. Results clearly indicate a single protein band of 165–170 kDa (NR2A) that is detected by the anti-NR2A antibody (Fig. 1E; n = 7), confirming the specificity of the antibodies. Negative controls were performed in which the primary antibody (anti-NR2A) was omitted from the immunoprecipitation step. NR2A subunits were not detected in these control experiments. Positive controls were performed by running rat brain homogenate (Santa Cruz, Santa Cruz, CA) on SDS-PAGE. Taken together, these results suggest that anti-NR2A detects the NR2A subunits that appear to be associated with the axons of developing primary motoneurons.
Electrophysiology
LOCAL PERFUSION. We next decided to use electrophysiology to investigate the presence of axonal or presynaptic NMDARs at the NMJ and to confirm the immunohistochemical results. We hypothesized that activation of these peripheral NMDARs will likely depolarize the axons and will influence the release of ACh at the endplate. We therefore attempted to record mEPCs from muscle fibers in the presence of glutamate receptor agonists. This was done using a relatively high bath flowrate (3 ml/min), flowing from dorsal to ventral over the preparation. The agonist perfusion pipette was placed at or ventral to the horizontal midline of the fish, also perfusing from dorsal to ventral. Whole cell patch-clamp recordings were performed on white muscle ventral to the perfusion pipette in the presence of TTX (1 µM) to ensure that we recorded mEPCs. With this design, perfusion of only the ventral axial muscle is attained with no back-flow of perfusate over dorsal muscle and the spinal cord. Perfusates may dilute by as much as 10-fold over a 300- to 500-µm distance (Herreras et al. 1994), and we therefore used a relatively high concentration of NMDA to ensure activation of axonal or presynaptic receptors. Separate control experiments were performed whereby we perfused saline or 0.1% DMSO (vehicle) over the preparations for the duration of the recording. We recorded solely from white muscle because the immunohistochemistry suggested that NMDARs were associated with primary and not secondary motoneurons. Application of 500 µM NMDA caused a significant (P < 0.05) rise in the frequency of spontaneous ACh release that was 346.7 ± 145.2-fold (n = 4). This increase was blocked by bath application of 50 µM AP-5 (Fig. 2A; n = 4). Because NMDARs appeared to be present at the NMJ, we tested for the possible presence of AMPARs and KARs. Perfusion with KA (250 µM) induced responses that were significantly (P < 0.05) greater than saline controls by 165.5 ± 34.9-fold (n = 4) and that were prevented with the non-NMDAR antagonist CNQX (50 µM; Fig. 2B; n = 4). Graphical representations of raw traces (Fig. 2, C and D) illustrate the significant increase in mEPC frequency in the presence of KA and NMDA (P < 0.05). There was no increase in mEPC frequency in any of the control experiments (P > 0.05). The observed increase in mEPC frequency during local application of agonists directly onto the trunk musculature suggests that glutamate receptors are likely associated with the axons of motoneurons in the peripheral nervous system.
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; Zhou et al. 1997). Representative recordings illustrate the KA-induced increase in spontaneous ACh release compared with controls (Fig. 5A). Concentration of 10, 50, and 100 µM KA were tested. No significant response to 10 µM KA was observed; however, in the presence of 50 and 100 µM KA, significant increases (P < 0.05) in mEPC frequency were observed (Fig. 5B; n = 4–6). The more moderate increase (150.7 ± 56.9-fold; n = 5) in the presence of 50 µM KA was chosen for complete pharmacological investigation. We found that Sym 2081 (10 µM) had no effect on the KA-induced increase in mEPC frequency (Fig. 5, A and C; n = 5). In addition, Sym 2081 (10 µM) alone had no effect on the frequency of spontaneous release (P > 0.05; n = 4). To investigate whether KA was nonspecifically activating AMPA receptors, GYKI 52466 (50 µM) was applied in the presence of KA (50 µM). GYKI 52466 prevented the KA-induced increase in frequency of spontaneous ACh release (P < 0.05), suggesting that KA acts nonspecifically to activate peripheral AMPARs.
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Representative traces are shown from recordings done in the presence of 1 mM glutamate, saline controls, and uptake inhibitors (Fig. 6A). In the presence of 1 mM glutamate, no significant (P > 0.05) change in frequency of spontaneous release was observed (Fig. 6; n = 5). However, in the presence of the general glutamate uptake inhibitor D-TBOA (200 µM) with 1 mM glutamate, significant (P < 0.05) increases in mEPC frequency were observed (Fig. 6; n = 6). During application of 200 µM TBOA alone, no change in mEPC frequency was observed (Fig. 6; n = 4).
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Analysis of recordings revealed the presence of multiple types of mEPCs (Fig. 8A). Of these pools of events, it is likely that large amplitude events with fast rise times are mEPCs from mature synapses on the muscle being recorded from. A second type of event is of a smaller amplitude (<400 pA) and displays relatively long rise times (Fig. 8A). It is probable that these events occur on cells other than the ones being recorded from and are highly filtered due to electrical coupling (Buss and Drapeau 2000; Nguyen et al. 1999). The third type of mEPCs have small amplitudes and fast rise times and may result from activation of immature synapses or simply from synapses with fewer numbers of nAChRs. It has previously been reported that two distinct pools of mEPCs can be recorded at the zebrafish NMJ (Nguyen et al. 1999). Nguyen et al. (1999) also found slow and fast rise time events with small amplitudes, although these were not reported as separate pools. Amplitude distributions clearly illustrate a bimodal distribution of mEPCs (Fig. 8, B and C) that is not altered in the presence of NMDA or AMPA. It is evident that there is a pool of events that has a mean amplitude of 200 pA with the division between event pools 400 pA (Fig. 8, B and C). The large (>400 pA)-amplitude events have a normal distribution with a mean value of 1,422 ± 110 pA (n = 7 experiments) in the presence of saline that is almost identical to the average amplitude of mEPCs in the presence of NMDA (1,433 ± 139 pA; n = 8 experiments). The average amplitude of large events is 1,455 ± 34 and 1,340 ± 140 pA in the presence of 0.1% DMSO and AMPA respectively (n = 6 experiments). Neither of these pairs of values are significantly different. Rise times versus amplitude plots clearly illustrate the different pools of mEPCs (Fig. 8, D and E). Small-amplitude events range in rise times from <0.1 to 0.4 ms, whereas larger-amplitude events have fast rise times between 0.05 and 0.09 ms.
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), peak amplitude, and proportion of all events that were large (>400 pA). These values were then compared with the average of all events occurring during agonist application, and the comparisons for large events are shown as ratios in Fig. 9, whereas the comparisons for small events are shown as ratios in Fig. 10. Analysis of agonist application data revealed no significant difference in the data for rise time, decay time, peak amplitude, or proportion of large (>400 pA)-amplitude events in the presence of NMDA, AMPA, KA, or glutamate (Fig. 9). The lack of difference in any of the above properties indicates that glutamate receptor agonists do not act postynaptically but rather on receptors that are located at some point presynaptic to the NMJ. This may be either at the level of the motoneuron axons or at the presynaptic terminal or both. These results also suggest that because the proportion of large events does not change in the presence of the iGluR agonists, then neither does the proportion of small events because these are reciprocal values.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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; Fu et al. 1995; Grozdanovic and Gossrau 1998; Liou et al. 1996; Malomouzh et al. 2003; Waerhaug and Ottersen 1993). Many of these reports have focused primarily on NMDARs; however, we also suggest the presence of non-NMDARs at the NMJ. Our use of specific non-NMDAR antagonists provides support for the presence of AMPARs but not KARs. When taken together, our results strongly suggest the presence of iGluRs in association with motoneuron axons outside the CNS. Furthermore, the abolishment of all mEPCs in the presence of curare suggests that iGluRs are not associated with muscle fibers. iGluRs at the NMJ
Data presented here suggest the presence of NMDARs in association with the axons of motoneurons and possibly also with presynaptic structures. The precise location of the iGluRs has yet to be thoroughly mapped out. Staining in 26 hpf embryos was similar to what was present at 4–6 dpf, indicating that NR2A subunits are associated with motoneuron axons and not glia. Although glial cell precursors are present in embryonic zebrafish (Gilmour et al. 2002), and may be associated with axon pathfinding and motoneuron differentiation (Gilmour et al. 2002; Park et al. 2002), it has been shown that myelination does not occur until 4 dpf (Brosamle and Halpern 2002). Therefore we have interpreted the positive NR2A immunostaining in the peripheral nervous system of 26 hpf embryos to indicate that the NMDARs are associated with motoneurons and not glial cells.
We tested other NR2 antibodies and antibodies against some AMPA and KA receptor subunits, but the results were inconclusive, so we focused primarily on NR2A subunits. NR2A subunits in zebrafish appear to run at a slightly different molecular weight (165–170 kDa) compared with rat NR2A, which is 180 kDa. This likely reflects subtle differences between the zebrafish and rat isoforms.
Variations between NR2 subunits are a major source of diversity in NMDARs. One such difference is the voltage dependence of the Mg2+ block (Hollmann and Heinemann 1994; Kirson et al. 1999). Other differences in NR2 subunits include responses to both glycine (Regalado et al. 2001) and glutamate (Chen et al. 2001) and mechanisms of modulation (Krupp et al. 1996). It has also been suggested that variations in surface expression pattern occur between subunits. Barria and Malinow (2002) suggest that NR2A subunits require ligand binding to be inserted into the membrane, whereas NR2B subunits do not. This may provide a mechanism that allows for selective surface expression of axonal NMDARs. NR2A subunits also have shorter decay times (Cathala et al. 2000), which may be important for faster on and off responses to changes in ligand availability. Also, NR2A are more permeable to Ca2+, which has been shown to be functionally important when NMDARs are the only Ca2+-permeable receptor at a synapse (Lei and McBain 2002). Any of the preceding subunit-specific characteristics could be important for the selective expression of NR2A subunits at the NMJ.
Recently, the presence of glutamate at the NMJ has been suggested in mammalian preparations (Barthelemy-Requin et al. 2000; Waerhaug and Ottersen 1993). There is also evidence that glutamate and ACh can be coreleased from vertebrate synaptosomes (Vyas and Bradford 1987). It does, however, seem most likely that glial cells surrounding the NMJ are involved in glutamate uptake and release. Although many types of glutamate transporters are probably present at the NMJ, previously published literature suggests that glial transporters are primarily responsible for glutamate clearance (Bergles and Jahr 1998; Danbolt 2001). The lack of effect in the presence of glutamate alone suggests that glutamate is either being taken up or being metabolized (Danbolt 2001; Matthews et al. 2000). This strongly suggests that glutamate is being removed from the site of action at a rate that prevents activation of iGluRs. The lack of effect of TBOA in the absence of applied glutamate suggests that little glutamate is released spontaneously. It is possible that endogenously released glutamate is capable of activating the receptors under normal swimming conditions when motoneurons are firing at frequencies 
30 Hz. Rapid removal of glutamate reflects the tight regulation of this neurotransmitter at the synaptic cleft to prevent excitotoxicity and diffusion (Asztely et al. 1997).
It is possible that glutamate is released from glial cells surrounding the NMJ. Although this has not been reported to occur at the NMJ, glial cells of the CNS are thought to have vesicular machinery (Calegari et al. 1999) and are able to release glutamate in a nonpathological, vesicular manner (Araque et al. 2000; Pasti et al. 2001). This release could be mediated through a feedback mechanism involving signaling molecules like NO or perhaps through muscarinic AChRs, which are known to be present on glial cells surrounding the NMJ (Robitaille et al. 1997).
Investigation of non-NMDARs suggests the presence of AMPARs, but not KARs, at the NMJ. This is suggested through the use of specific AMPAR and KAR antagonists (GYKI 52466 and Sym 2801 respectively). Because EC50 values for activation of AMPARs by KA have been reported to be between 56 and 150 µM (Jayaraman 1998; Lambolez et al. 1991; Lazzaro et al. 2002; Nakanishi et al. 1990; Patneau et al. 1993), the concentrations of KA that we used could most likely activate both AMPA and KARs. Prevention of the KA-induced increase in mEPC frequency by GYKI 52466 supports indiscriminant non-NMDAR activation by KA.
Although antibody staining suggests that only primary motoneurons express NMDARs, both small and large mEPCs increase in frequency during iGluR agonist application. One reason for this may be that both large and small mEPCs are due to spontaneous release from primary motoneurons. Another is that small mEPCs may be due to the presence of immature synapses along different branches of primary motoneurons. It is also possible that secondary motoneurons express NR2 subunits other than NR2A, and therefore were not detected. Whatever the cause, both small and large mEPCs increase in frequency during iGluR agonist applications, with the relative proportion of each remaining the same.
Role of iGluRs at the NMJ
Glutamate is the major excitatory neurotransmitter in the vertebrate CNS. During development, glutamate and its receptors have many important roles in axonal outgrowth (Metzger et al. 1998), learning and memory, (Atwood and Wojtowicz 1999) and synaptic strengthening (Gu et al. 1996). Previously reported work on glutamate receptors at the NMJ has suggested that Ca2+ may play roles in developmental changes in the presynaptic terminal (Fu et al. 1995). Aside from this, iGluRs may also alter muscle contraction (Koyuncuoglu et al. 1998) or aid in strengthening postsynaptic components of the NMJ. It is known that pruning of motoneurons is activity dependent and that postsynaptic AChR clustering is also dependent on the presence of the presynaptic motoneuron (Luo et al. 2002; Sanes and Lichtman 1999). If activation of iGluRs increases ACh release and subsequent activity at the developing NMJ, then they could play a role in both pre- and postsynaptic modifications.
Ionotropic glutamate receptors at the NMJ may be replaced with mGluRs at a later developmental stage (Liou et al. 1996). This may reflect a developmental change in requirements for glutamate at the NMJ. Initial excitatory actions of glutamate switching to inhibitory ones later in life may occur to strengthen the synapse early and then switch to a modulatory role. Strengthening, or other glutamate-mediated actions, could occur through a Ca2+-mediated mechanism or via another pathway such as NO signaling (Grozdanovic and Gossrau 1998; Malomouzh et al. 2003; Pinard et al. 2002).
Taken together, our results suggest the presence of iGluRs at the larval zebrafish NMJ. Electrophysiology and kinetic analysis of mEPCs suggest that iGluRs do not directly modulate nAChR kinetics. However, it is possible that increased Ca2+ in both the pre- and postsynaptic terminals could have a role in the development of the NMJ. Both pre- and postsynaptic components require Ca2+-dependent changes after motoneuron contact that could be facilitated by iGluRs. This does, however, remain an initial characterization of iGluRs at zebrafish NMJs. We are attempting to determine the precise location of the NMDARs, and the physiological function of these receptors at NMJs. In addition, we are investigating whether or not iGluRs are also associated with the NMJs on red muscle fibers. In the process of achieving these aims, it may be important to determine what other types of presynaptic receptors are present at the zebrafish NMJ. As different subunits have different functional properties, knowledge of their presence may aid in resolving the role of iGluRs. This will add to our understanding of NMJ development and modulation and perhaps synaptogenesis in general.
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ACKNOWLEDGMENTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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This work was supported by grants from National Science and Engineering Research Council (NSERC) and Canadian Foundation for Innovation (CFI) (to D. W. Ali), an NSERC studentship (to K. J. Todd), and NSERC and Alberta Heritage Foundation for Medical Research Summer studentships (to C.A.B. Slatter).
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FOOTNOTES |
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Address for reprint requests and other correspondence: D. W. Ali, CW405 Biological Sciences Bldg., Dept. of Biological Sciences, University of Alberta, Edmonton, Alberta T6G 2E9, Canada (E-mail: declan.ali{at}ualberta.ca).
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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