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NEUROSCIENCE |
1 Department of Biological Sciences
2 Centre for Neuroscience, Biological Sciences Building, University of Alberta, Edmonton, Alberta, Canada, T6G 2E9
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Abstract |
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 Top Abstract IntroductionMethodsResultsDiscussionReferences |
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with depolarization was found to be greater at 48 hpf than at 33 hpf. AMPARs in 33 hpf fish had a conductance of 
9 pS, and in older fish 15 pS. Finally, the desensitization blocker, cyclothiazide, increased by 4-fold in 48 hpf preparations, but only 1.5-fold in 33 hpf fish. These results are consistent with the hypothesis that the major mechanism underlying the developmental speeding in AMPAR kinetics in zebrafish CNS is a switch in receptor subunits. To our knowledge this is the first study to suggest that AMPARs change subunits during development in fish.
(Received 6 February 2007;
accepted after revision 3 April 2007;
first published online 5 April 2007)
Corresponding author: Declan W Ali, Department of Biological Sciences and Centre for Neuroscience, Biological Sciences Building, University of Alberta, Edmonton, Alberta, Canada, T6G 2E9. Email: declan.ali{at}ualberta.ca
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Introduction |
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TopAbstractIntroductionMethodsResultsDiscussionReferences |
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The strength of synaptic transmission at excitatory synapses depends, in part, upon receptor properties such as channel conductance, open/close rates and burst behaviour (Ambros-Ingerson & Lynch, 1993; Benke et al. 1998; Lin et al. 2002). The amplitude and time course of the synaptic currents that are mediated by AMPARs are key players of synaptic information efficacy and cell excitability, and changes in the nature of these currents will impact upon cell activity. Developmental changes in the properties of synaptic currents have been observed at excitatory synapses (Sakmann & Brenner, 1978; Hestrin, 1992; Wall et al. 2002). However, information about developmental changes in AMPAR-mediated mEPSCs is somewhat contradictory. For example, in hippocampal pyramidal cells, AMPA receptor-mediated currents show an increase in decay times with maturation (Seifert et al. 2000), whereas other studies report a developmental decrease in decay time constants (Taschenberger & von Gersdorff, 2000; Iwasaki & Takahashi, 2001; Joshi & Wang, 2002; Wall et al. 2002; Yamashita et al. 2003; Rumpel et al. 2004). The time course of synaptic currents depends upon factors such as receptor affinity (Jones et al. 1998), desensitization (Jones & Westbrook, 1996), glutamate dynamics in the synaptic cleft (e.g. release, clearance) and synaptic morphology (Cathala et al. 2005). The two main mechanisms that are thought to primarily underlie the developmental speeding of AMPA mEPSCs are (1) changes in receptor subunit composition (Lawrence & Trussell, 2000; Kumar et al. 2002); and (2) changes in synaptic morphology (Cathala et al. 2005). It is unknown if a developmental change in AMPAR kinetics occurs in zebrafish, and if so, what the underlying mechanism might be.
Mauthner cells (M-cells) are the largest neurons in the teleost's CNS and play key roles in escape behaviours (Kimmel et al. 1981; Metcalfe et al. 1990). The earliest synapses on M-cells are established by around 24 h postfertilization (hpf) (Kimmel et al. 1990), with functional inputs occurring around 26–27 hpf (Grunwald et al. 1988; Ali et al. 2000b). Some characteristics of GluRs on developing M-cells have already been investigated (Ali et al. 2000a), but the developmental profile of AMPAR properties is still to be determined. The zebrafish is an excellent preparation to study synaptic development, because the wide variety of locomotor mutants that are available and the recent sequencing of the genome allow a thorough genetic investigation into synaptic maturation. Furthermore, because Mauthner cells are involved in simple locomotor behaviours, such as the C-start reflex, the development of excitatory synapses associated with these cells can be related to behaviours in developing fish (Liu & Fetcho, 1999; Fetcho, 2006).
Therefore, we have undertaken this study to examine the developmental properties of AMPAR-mediated currents in vivo, on zebrafish M-cells. We show that there is indeed a developmental change in the AMPAR-mEPSC kinetics and that this change appears to be due to subunit switching rather than a change in synaptic morphology.
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Methods |
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Wild-type Zebrafish (Danio rerio) embryos were raised at 28.5°C, and collected and staged as previously described (Kimmel et al. 1995; Westerfield, 2000). All procedures were carried out in compliance with the guidelines stipulated by the Canadian Council for Animal Care and the University of Alberta. Embryos and newly hatched larvae were anaesthetized in 0.02% tricaine (MS-222) (Sigma, St Louis, MO, USA) and dissected as described by Drapeau et al. (1999). Briefly, the entire hindbrain was exposed after removing the forebrain and rostral structures, but leaving the spinal cord intact.
Electrophysiology
The preparations were moved to the recording set-up and the chamber was continuously perfused at room temperature (20–24°C) with an aerated recording solution that contained 15 µM D-tubocurarine (Sigma) to paralyse the preparations, but lacked tricaine. The recording solution contained (Evans, 1979) (mM): 134 NaCl, 2.9 KCl, 2.1 CaCl2, 1.2 MgCl2, 10 Hepes, and 10 glucose, osmolarity adjusted to 290 mosmol l–1, pH 7.8. Whole-cell recordings were obtained from Mauthner neurons located in rhombomere 4. These cells were easily identified under Nomarski differential interference contrast (DIC) optics based on their morphology and location in the hindbrain (Fig. 1A; Eaton and Farley, 1973). Cells were filled with Lucifer Yellow (0.1%) to confirm their identity. Patch-clamp electrodes were pulled from borosilicate glass and when filled with a Cs-gluconate solution had resistances of 3.5–5 M
. The Cs-gluconate intracellular solution was composed of (mM) 115 Cs-gluconate, 15 CsCl, 2 MgCl2, 10 Hepes, 10 EGTA, and 4 Na2ATP, osmolarity adjusted to 280 mosmol l–1, pH 7.2.
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, and recordings in which the series resistance changed by at least 15% or membrane potentials were more positive than –45 mV were discarded. The liquid junction potential between the pipette and external solution was not corrected for. Analysis of mEPSCs
Synaptic activity was monitored using pClamp 8.1 software (Axon Instruments). Synaptic events were detected (using the template function for events >2.5 standard deviations above the basal noise) and analysed with Axograph 4.6 (Axon Instruments). The software detected all events that could be recognized visually. All events were then inspected visually, and those with uneven baselines or overlaying events (<5%) were discarded. The decay time course was analysed over the first 30 ms for pure AMPA events and was fitted with a sum of exponential curves. The presence of one or two exponential components during the decay of the mEPSCs was tested by comparing the sum of squared errors of the fits over the same decay ranges (Clements & Westbrook, 1991; Legendre, 1998). Averages of these events were always best fitted with one exponential curve. The rise times were defined as the time from 20% to 80% of the mEPSC amplitude. All data values are given as means ± S.E.M. Correlations were calculated with the use of a least-squares linear regression analysis. Significance was determined using one-way ANOVAs and Fisher LSD tests for normally distributed, equal variance data and paired t tests. Kruskal–Wallis ANOVA and Dunn's method of comparison were used for non-normal distributions. * denotes P < 0.05 and ** denotes P < 0.01.
Non-stationary fluctuation analysis
We performed non-stationary fluctuation analysis to estimate the single-channel current (i) and the available number of channels (N) (NSFA; Sigworth, 1980). The mEPSCs were aligned on their rise times, and events with obvious artifacts were manually discarded. Non-stationary analysis was performed on the deactivation phase of the responses. The averaged current and variance over time were computerized using Axograph 4.6 software after digital filtering of the traces at 2 kHz. Between 20 and 80 traces per patch were obtained for analysis.
The variance (
2) was plotted against the mean current (I) and the data points were fitted with the following parabolic function (Sigworth, 1980):
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| (1) |
b2 is the variance of the background noise. A linear fit to the initial part of the curve (10–15% of the response range) yields a slope factor that corresponds to the single-channel current (i) (Benke et al. 2001). All estimates for which the initial rising phase of the current-variance plot was well-fitted by a linear fit, produced similar values for conductance (
), as previously reported (Traynelis et al. 1993; Benke et al. 1998). The slope conductance value () for each channel was then calculated by the equation
=
i/(Vm
–
Erev), with Vm
=
–60 mV and Erev
= 0 mV. Chemicals
All drugs were dissolved in the aerated recording solution and applied by bath perfusion. Strychnine (5 µM), picrotoxin (100 µM), CNQX (10 µM), APV (50 µM), cyclothiazide (CTZ; 10 µM), kynurenic acid (KYN; 50 µM), SYM (10 µM) and GYKI (50 µM) were obtained from Sigma. Tetrodotoxin (TTX; 1 µM) and DL-TBOA (100 µM) were purchased from Tocris (Avonmouth, UK).
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Results |
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(33 hpf), 253 ± 48 M (48 hpf) and 87 ± 19 M (72 hpf), while the resting membrane potentials were –54 ± 1 mV (33 hpf), –58 ± 2 mV (48 hpf) and –61 ± 2 mV (72 hpf). Isolation of non-NMDA-mediated mEPSCs
We identified M-cells in vivo based upon the location of their large cell bodies adjacent to the otic vesicles in the hindbrain. Cells were filled with Lucifer yellow to confirm their identity (Fig. 1A).
Miniature EPSCs, unlike evoked currents, allow accuracy in determining the properties of postsynaptic receptors because their decay kinetics in particular can be highly influenced by developmental changes in the synchrony of vesicular release or release probability (Silver et al. 1996; Wall et al. 2002; Cathala et al. 2005). Therefore, we focused our analysis on mEPSCs to determine the developmental changes in the properties of postsynaptic receptors.
A high level of synaptic activity was observed when we recorded from the M-cells in normal physiological saline (Fig. 1B). The M-cells were clamped at a holding potential of –60 mV, where all inhibitory currents were small and inward. To differentiate inward, excitatory currents from inhibitory currents, blockers for glycine and GABA receptors (5 µM strychnine and 100 µM picrotoxin, respectively) were added to the bath during all recordings. In these initial experiments, 50 µM APV was added to the bath to block all NMDA receptor activity (Fig. 1C), but we found that isolated non-NMDA mEPSCs were identical in all aspects when recorded in the presence or absence of APV. Therefore we omitted APV for all further recordings at –60 mV, but included it when recording at potentials more positive than –60 mV. Miniature EPSCs were recorded in the presence of 1 µM TTX (Fig. 1C). The remaining mEPSCs were completely blocked by the non-NMDA antagonist CNQX (10 µM; Fig. 1C). Individual mEPSCs from a recording were averaged, and all analyses were performed on averaged mEPSCs (Fig. 1D).
Developmental profile of non-NMDA mEPSCs
Because it is difficult to properly clamp cells with large and extensive dendrites, whole-cell recordings are often subject to the problem of inadequate space clamp (Rall, 1969). Accurate control of dendritic potentials from a point voltage source is difficult, and synaptic events that are generated at more distal locations may be filtered leading to attenuation and widening. Thus, these events will appear to have longer rise times, smaller peak amplitudes and longer decays (Rall, 1969). Therefore, the correlation between the rise time and peak amplitude of events at 33 hpf, 48 hpf and 72 hpf (Fig. 2A) were examined for all recordings to determine if cells were properly space clamped. The lack of a correlation at all ages (r = 0.02 at 33 hpf; r < 0.01 at 48 hpf; r = 0.04 at 72 hpf), suggests that the M-cells remained electrically compact over the ages examined, and that space-clamping errors were not a major issue in our study. The mEPSC amplitude distribution at all ages remained unimodal (Fig. 2B), with the average peak amplitude increasing significantly between 33 hpf and 48 hpf (33 hpf: 23.9 ± 2.0, n = 10; 48 hpf: 30.0 ± 1.5, n = 9 and 72 hpf: 29.4 ± 0.9, n = 8; P < 0.05) (Fig. 2B). Amplitude histograms from representative experiments are shown in Fig. 2C. The tight, normal distributions suggest that there was very little or no variation in quantal size.
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mEPSCs are mediated solely by AMPARs
To determine if the mEPSCs were mediated by AMPA or kainate receptors, we examined the effect of the selective AMPAR antagonist (GYKI; 50 µM), and the selective kainite-desensitizing agent SYM (10 µM) on mEPSCs. We found that GYKI completely blocked all mEPSCs and that SYM had no effect on either the mEPSC frequency, amplitude, rise time or the decay time constant (P > 0.05; see Fig. 1 in online Supplemental material). These results suggest that the mEPSCs were due solely to the activation of AMPA receptors and lacked a kainate component at all ages.
The marked change in postsynaptic currents during development depends on a number of factors (Edmonds et al. 1995; Conti & Weinberg, 1999), most notably a change in synaptic morphology (Cathala et al. 2005) or a switch in receptor subunits (Kumar et al. 2002; Wall et al. 2002). Therefore, we next investigated the mechanism underlying the developmental changes observed in the AMPAR-mediated mEPSC kinetics between 33 hpf and 48 hpf.
Effect of glutamate uptake in shaping the mEPSC
The glutamate transient in the synaptic cleft changes shape due to alterations in synaptic morphology that might occur during development (Lee & Sheng, 2000). These changes have been shown to lead to faster glutamate clearance from the cleft (Otis et al. 1996; Diamond & Jahr, 1997), resulting in a shorter mEPSC decay time constant. Therefore it was possible that the developmental decrease in that we had noted was due to faster clearance of glutamate from the cleft. To test this, we recorded AMPAR-mEPSCs in the presence and absence of the glutamate transporter blocker DL-TBOA. Our rationale was that if the faster events in older animals were due to the appearance of glutamate transporters, then DL-TBOA should block their activity and widen the mEPSC in older animals (48 hpf) compared with younger ones (33 hpf). However, we found that DL-TBOA did not affect the mEPSC frequency, amplitude, rise time and decay time constant at any age (P > 0.05; see Fig. 2 in online Supplemental material). The lack of effect of DL-TBOA indicates that glutamate uptake does not play a major role in shaping the AMPAR-mESPC in zebrafish.
Additionally, developmental changes in AMPAR-mEPSC kinetics might be due to an increased concentration of glutamate in the synaptic cleft. The glutamate receptor antagonist kynurenic acid (KYN) may be used to detect changes in the glutamate concentration in the synaptic cleft because the effect of KYN would be weaker if the glutamate concentration in the synaptic cleft is higher (Diamond & Jahr, 1997; Wadiche & Jahr, 2001; Yamashita et al. 2003). Application of KYN resulted in a reduction in the mEPSC peak amplitude (Fig. 4C) at both 33 hpf and 48 hpf as expected; however, the reductions were to similar extents (Fig. 4D; P > 0.05) suggesting that the glutamate concentration in the synaptic cleft does not change significantly during development.
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Voltage dependence of AMPAR-mediated mEPSCs
Because the I–V relationship of AMPARs is known to be subunit dependent (Geiger et al. 1995; Sah & Lopez De Armentia, 2003), we examined the voltage dependence of the mEPSCs by varying the membrane holding potential in steps of 20 mV increments from –80 mV to +80 mV. In order to analyse the current–voltage (I–V) relationship quantitatively, we averaged well-isolated AMPAR-mEPSCs occurring during 30–60 s time periods (n = 4–9) at each holding potential, and plotted the peak amplitude of each averaged mEPSC against the holding potential (Fig. 5A and B). We found that the I–V relationship at 33 hpf was linear (Fig. 5A), while at 48 hpf, it displayed outward rectification. The I–V relationship at 72 hpf also exhibited outward rectification (data not shown), similar to that at 48 hpf. Even though our outward rectification I–V relationship is similar to that observed in cells in the nucleus magnocellularis (Otis et al. 1995), outward rectification is rare for AMPA receptors and most cells containing AMPAR receptors show a linear or inwardly rectifying I–V relationship.
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Because our data suggested that the receptors at the two ages may indeed be different, we determined the single-channel conductance of the AMPARs at 33 hpf and 48 hpf. In order to do this, we performed non-stationary fluctuation analysis (NSFA) on AMPAR-mEPSCs. Estimates of AMPAR unitary conductance () and numbers of active channels (N) can be obtained from analyses of miniature synaptic events, which examine the variance of fluctuations in the decay phase of synaptic events (Traynelis et al. 1993; Benke et al. 1998). This analysis is based on the assumption that the opening and closing of individual channels are independent events. Thus, the event-to-event fluctuations in synaptic currents will be smallest at the peak and end portion of the event, and largest during the decay phase, as determined by single-channel conductance and the number of functional channels. Variance–current plots were constructed at both ages, and estimates of and N were derived from fitted functions (see Methods). We found a significant difference in between the two ages (33 hpf: 9.6 ± 0.52 pS and 48 hpf: 15.2 ± 1.17 pS; n
= 6; P
= 0.002; Fig. 7), suggesting that the AMPARs at 33 hpf and at 48 hpf are truly different. There was no significant change in the number of active channels during development. Specifically, we found that at 33 hpf the number of active channels was 28 ± 2 and at 48 hpf, it was 33 ± 5 (n
= 5 at 33 hpf and n
= 6 at 48 hpf; P
= 0.38).
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Lastly, we examined the effect of cyclothiazide (CTZ) on AMPA mEPSCs. CTZ suppresses the desensitization of AMPA receptors (Partin et al. 1993; Wong & Mayer, 1993), but does so to a greater degree on the flip splice variants and on heteromeric receptors (Partin et al. 1994, 1995; Cotton & Partin, 2000). Therefore, differential effects of CTZ imply the presence of different subunits, which may underlie the speeding of the AMPAR kinetics in older animals. We used a low concentration (10 µM) of CTZ to discern between flip and flop forms of the subunits (Partin et al. 1994; Cathala et al. 2005). We found that CTZ had no effect on the mEPSC frequency, amplitude or rise time (P > 0.05 Fig. 8C); however, the decay time at both ages was significantly increased (33 hpf, P
= 0.011; 48 hpf P
= 0.012; paired t tests). The decay time constant in 33 hpf controls was 0.8 ± 0.07 ms and increased to 1.2 ± 0.1 ms (n
= 3) in the presence of CTZ, and in 48 hpf controls was 0.5 ± 0.03 ms and increased to 2.1 ± 0.2 ms (n
= 4) in the presence of CTZ. Therefore, CTZ showed a stronger potentiation of the decay time constant in the older animals (a 4-fold increase in at 48 hpf) compared with the younger ones (1.5-fold increase in at 33 hpf; Fig. 8D; P
= 0.025, ANOVA). These observations suggest that AMPARs at 33 hpf are composed of subunits that are different from those at 48 hpf.
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Discussion |
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TopAbstractIntroductionMethodsResultsDiscussionReferences |
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Here, we provide several lines of evidence that are consistent with a switch in AMPAR subunits. First, application of DL-TBOA and KYN did not alter the kinetics of the mEPSC, suggesting that glutamate uptake or the glutamate concentration in the synaptic cleft did not change during development. Second, the decay time course of AMPAR mEPSCs decreases between 33 hpf and 48 hpf. Third, in 48 hpf fish, the decay time constant is affected by voltage to a much larger degree than it is at 33 hpf. Fourth, CTZ increases the decay time constant of AMPAR mEPSCs 4-fold in 48 hpf fish, but only 1.5-fold at 33 hpf, and fifth, the main single-channel conductance of AMPAR at 33 hpf is 9 pS, but 15 pS at 48 hpf. Taken together, these data suggest that the population of AMPARs at 33 hpf is likely to be functionally different from that at 48 hpf.
Our results show that the time course of AMPAR mEPSCs became faster during development. The decay kinetics of AMPAR-EPSCs can be influenced by a number of factors, including the composition of the postsynaptic receptors (Edmonds et al. 1995; Conti & Weinberg, 1999), the rate at which they desensitize (Jones & Westbrook, 1996), the dynamics of glutamate in the synaptic cleft (e.g. release, clearance), and changes in synaptic structure and the surrounding neuropil (Cathala et al. 2005). Our findings are similar to a number of other studies, which show a developmental speeding of AMPA mEPSC kinetics in preparations such as the rat cochlear nucleus (Brenowitz & Trussell, 2001), cerebellar mossy fibre granule cells (Wall et al. 2002), median nucleus trapezoid bodies (Taschenberger & von Gersdorff, 2000) and mammalian neocortical pyramidal neurons (Kumar et al. 2002). These changes are thought to occur due to alterations in the composition of the AMPA receptor (Edmonds et al. 1995; Conti & Weinberg, 1999; Kumar et al. 2002). Similarly, a switch in receptor subunit composition is thought to underlie the developmental speeding of the decay time course in glycinergic and NMDA receptor-mediated synaptic currents (Hestrin, 1992; Takahashi et al. 1996; Singer et al. 1998). However, a recent study has shown that the speeding of AMPA EPSC kinetics in mouse cerebellar granule cells occurs via a different mechanism: maturation of synaptic morphology rather than subunit alterations (Cathala et al. 2005). Our findings suggest that in zebrafish M-cells, subunit switching is likely to be the main determinant of the accelerated decay of AMPAR-EPSCs.
The decay time constant increased with depolarization at all ages, but this occurred to a greater extent in older animals. A similar voltage-dependent effect on the AMPA mEPSC decay kinetics has been reported (Otis et al. 1996; Glowatzki & Fuchs, 2002; Morkve et al. 2002); however, it is not known which subunit(s) might be responsible for this property (Morkve et al. 2002). We observed a significant difference in the percentage increase in the decay time constant with depolarization between 33 hpf and 48 hpf animals, suggesting that the receptor gating properties contributing to the EPSC decay kinetics are different at those ages. Since receptor gating properties are dependent on the receptor's subunit composition, it is likely that the AMPARs in M-cells undergo a developmental switch in their subunit composition.
One of our most compelling findings was the differential effect of CTZ at 48 hpf compared with 33 hpf. We were careful to use low concentrations of CTZ in order to ensure specificity (Partin et al. 1994; Cathala et al. 2005) on AMPARs. The dramatic increase in at 48 hpf, but not at 33 hpf, suggests that the alternatively spliced, flip variant may be present in greater quantities in older animals. CTZ blocks desensitization of AMPARs and preferentially has a stronger effect on the flip forms of the subunits (Partin et al. 1994). Thus the differential effects of CTZ at 48 hpf and 33 hpf argue in favour of the flop form associated with M-cells early in development, with the emerging onset of the flip version sometime later. This contrasts with a recent study showing that the flip form of GluR2 is the predominant form in the CNS at 30 hpf (Lin et al. 2006). Interestingly, while both forms increase in expression after 30 hpf, the flip variant is expressed in greater amounts (Lin et al. 2006). Finally, since CTZ is thought to have a greater degree of potentiation on homomeric receptors (Cotton & Partin, 2000), our results indicate that the AMPARs at 33 hpf may be heteromeric, while those at 48 hpf are likely to be homomeric. However, it should be noted that the differential effect of CTZ at the different ages may also be due to unknown developmental changes occurring within the synapse. For instance, if the receptors desensitize differently due to differential phosphorylation mechanisms, then the responses to CTZ may be different between the two ages.
We found that in older animals (48 hpf to 72 hpf), the AMPARs exhibited outward rectification, while at 33 hpf the relationship was linear. This was a curious finding because most AMPA receptors exhibit either a linear or inward rectification (Verdoorn et al. 1991; Koh et al. 1995; Lui & Cull-Candy 2000). However, some GluR2-containing receptors do display outward rectification in the edited form (Verdoorn et al. 1991; Morkve et al. 2002). The fact that we detected only a linear I–V relation in the young animals, and only outward rectification in older fish, supports our view that the AMPA receptors are different between the two ages. Site-directed mutagenesis of single amino acid residues of AMPAR subunits may cause outward rectification (Dingledine et al. 1992). Our findings suggest that the amino acid identity of AMPARs in zebrafish is most certainly different from that in mammals, and may underlie some of the interesting properties that we have seen.
The small but significant developmental increase in mEPSC amplitude could be due to a number of factors, including an increase in (1) the number of postsynaptic receptors; (2) single-channel conductance; (3) quantal content; or (4) a change in synaptic morphology. AMPARs on 33 hpf M-cells exhibited a smaller conductance (9 pS) than those at 48 hpf (15 pS). Our findings are similar to those observed in other systems where values of 8–10 pS and 15–20 pS have been found (Morkve et al. 2002; Momiyama et al. 2003; Bannister et al. 2005; Cathala et al. 2005). The conductance of AMPARs is critically dependent upon subunit composition (Swanson et al. 1997), suggesting that the different conductances we have seen at 33 hpf and 48 hpf are due to separate populations at these two ages. What might be the physiological significance of a subunit switch? Current theory suggests that a speeding in AMPAR kinetics occurs in order to ensure temporal precision and increased efficiency of information transfer (London et al. 2002). In zebrafish, the switch in subunits occurs between 33 h and 48 h, immediately prior to hatching at 48–52 hpf. The perceived advantage would be increased efficiency in the escape response that is necessary for survival.
In summary, we have characterized a number of developmental changes in AMPAR-mediated synaptic currents in Mauthner neurons. Our findings are similar to those of previous reports that show a developmental change in the composition of AMPARs (Bellingham et al. 1998; Taschenberger & von Gersdorff, 2000; Kumar et al. 2002; Wall et al. 2002). While our results suggest that a subunit switch occurs, they are not conclusive, and it is possible that unknown developmental mechanisms may account for a number of the changes that we have seen. Our findings indicate that the AMPARs at the ages examined contain at least one GluR2 subunit, and that the overall receptor subunit assembly probably changes between 33 hpf and 48 hpf. All of these changes occur between 33 hpf and 48 hpf, which is a critical period for synapse formation (Kimmel et al. 1981), sensory onset (Eaton & Farley, 1973) and morphological maturation of the M-cell (Kimmel et al. 1981).
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Footnotes |
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TopAbstractIntroductionMethodsResultsDiscussionReferences |
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