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1 Division of Neuroscience, University of Edinburgh, 1 George Square, Edinburgh EH8 9JZ, UK2 Wellcome Laboratory for Molecular Pharmacology, Department of Pharmacology, University College London, Gower Street, London WC1E 6BT, UK
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(Received 3 March 2004;
accepted after revision 22 April 2004;
first published online 23 April 2004)
Corresponding author D. J. A. Wyllie: Division of Neuroscience, University of Edinburgh, 1 George Square, Edinburgh EH8 9JZ, UK. Email: dwyllie1{at}staffmail.ed.ac.uk
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Introduction |
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In a recent study, Brickley et al. (2003) concluded that incorporation of the NR2D subunit into heteromeric NMDA receptor complexes might restrict such receptors to extrasynaptic sites (see also Momiyama, 2000). Given that NR2D mRNA levels are at their highest early in development (Watanabe et al. 1992; Monyer et al. 1994), the activation of these receptors by glutamate may be important in mediating effects that are distinct from the ‘classical’ role played by ionotropic glutamate receptors in ‘fast’ synaptic transmission. Recombinant and native NR2D-containing NMDA receptors show slow deactivation times (time constant of decay of around 5000 ms) following brief applications of glutamate (Monyer et al. 1994; Vicini et al. 1998; Wyllie et al. 1998; Misra et al. 2000). However, it is unclear what mechanism determines the slow kinetics of NR2D-containing receptors.
The proposed topology of NMDA receptor subunits includes an extracellular amino terminus, three membrane spanning segments (M1, M3 and M4), one membrane-embedded inverted loop (M2) that forms the channel pore and an intracellular carboxy terminus (for a review see Dingledine et al. 1999). Two polypeptide segments, S1 (
150 residues preceding M1) and S2 (the extracellular loop between M3 and M4), are found in glutamate receptor subunits and show sequence homology with bacterial periplasmic amino acid binding proteins (Stern-Bach et al. 1994). Functional binding studies and structural data have shown that the S1/S2 segments form a ligand-binding pocket and confer ligand-specificity in AMPA/kainate receptors and in S1S2 fusion proteins (Kuusinen et al. 1995; Armstrong et al. 1998; Keinanen et al. 1998; Lampinen et al. 1998). A sketch of the proposed structure of ionotropic glutamate receptors is shown in Fig. 1.
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To address the importance of the length of time glutamate is bound to the receptor, a mutation analogous to the T671A mutation in the NR2A subunit was introduced into the NR2D subunit (T692A). This mutation should disrupt the binding of glutamate. The mutant NR2D subunit cRNA was coexpressed with wild-type NR1a cRNA in Xenopus laevis oocytes and the effect of the mutation on glutamate and glycine potency was examined under two-electrode voltage-clamp configuration. Single-channel properties of the receptor were analysed in outside-out membrane patches and concentration jump experiments were performed to examine the macroscopic current decay of NR1a/NR2D(T692A) receptors following a brief application of glutamate.
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Methods |
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The wild-type expression plasmids for the rat NR1a (exon 5-lacking) and NR2D NMDA receptor constructs were as in Kuner & Schoepfer (1996). The T692A mutation was introduced using a similar PCR strategy to that described in Anson et al. (1998). The mutation was confirmed by DNA sequencing.
Heterogous expression of NMDA receptors
cRNA was synthesized as run-off transcripts from linearized plasmid DNA using the Promega (Madison, WI, USA) RiboMAx RNA synthesis kit. Reactions were supplemented as previously described (Anson et al. 1998). NR1 and NR2D cRNAs were mixed in a nominal ratio of 1: 9 before injection. Xenopus laevis were killed, in accordance with current Home Office guidelines, by exposure to an overdose of anaesthetic. Oocytes were removed, defolliculated and injected with either NR1a/NR2D wild-type or NR1a/NR2D(T692A) cRNA mixtures. Oocytes were incubated at 19°C for 2–3 days in modified Barth's solution (mM): 88 NaCl, 1 KCl, 2.4 NaHCO3, 0.33 Ca(NO3)2, 0.82 MgSO4, 0.44 CaCl2, 15 Tris-Cl, adjusted to pH 7.4 with NaOH, supplemented with 50 IU ml–1 penicillin (BRL, Bethesda, MD, USA), 50 µg ml–1 streptomycin (BRL), and 30 µMD(–)-2-amino-5-phosphonopentanoic acid (APV) (Tocris Cookson, UK), followed by storage at 4°C until they were used for electrophysiological measurements (2–7 days after injection).
Whole-oocyte recordings
Responses to glutamate, aspartate, NMDA or glycine were measured with a two-electrode voltage-clamp amplifier (TEC05, npi electronics, Tamm, Germany or Geneclamp, Axon Instruments) at –60 mV using 0.5–2 M
electrodes filled with 3 M KCl. Oocytes were perfused in a modified Ca2+-free normal frog Ringer solution (NFR) solution supplemented by 1.8 mM BaCl2 as described by Anson et al. (1998). For glutamate, aspartate and NMDA dose–response measurements, the NFR was further supplemented with 20 µM glycine and for glycine dose–response measurements the NFR was supplemented with either 30 µM glutamate for wild-type receptors or 10 mM glutamate for mutant receptors.
Application of solutions was either controlled manually or by the computer program, CellWorks Lite (npi electronics). Data were filtered at 100 Hz, digitized at 300 Hz and reduced to a final sampling rate of 10 Hz (oversampling with equal weight averaging of 30 samples) before storage.
Data analysis for dose–response curves
Data analysis was performed using the program CVFit (see http://www.ucl.ac.uk/Pharmacology/dc.html). Dose–response curves were fitted individually for each oocyte with the Hill equation:
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Single-channel recordings
Before making patch-clamp recordings the vitelline membrane of each oocyte was removed in a hypotonic solution containing (mM): 200 sodium methylsulphate, 20 KCl, 1 MgCl, 10 Hepes (pH 7.4 with KOH). Outside-out patches were held at –100 mV and recordings were made in an external solution containing (mM): 125 NaCl, 3 KCl, 1.25 NaH2PO4, 20 Hepes, 0.85 CaCl2 plus 20 µM glycine (pH 7.4 with NaOH). Patch pipettes were made from thick-walled borosilicate glass (Clark Electromedical Instruments, Pangbourne, UK), and contained (mM): 141 potassium gluconate, 2.5 NaCl, 10 Hepes, 11 EGTA (pH 7.4, with KOH). After fire-polishing the tips, patch pipettes had resistances of 10–20 M. Single-channel activity of NR2D(T692A)-containing NMDA receptors was evoked by either glutamate (200 µM), aspartate (100 µM) or NMDA (100 µM) and recorded with an Axopatch-1D amplifier (Axon Instruments, Foster City, CA, USA) and recorded on digital audio tape (Biologic DTR 1205, Biologic Instruments, Claix, France).
Analysis
Single-channel currents were replayed from digital audio tape, filtered at 2 kHz and digitized at 20 kHz. The amplitudes of single channel currents and their durations were measured using the SCAN program and analysed using the EKDIST program (Colquhoun & Sigworth, 1995; available from http://www.ucl.ac.uk/Pharmacology/dc.html). A resolution of 40 µs was imposed on both openings and closings in each data record. Distributions of fitted amplitudes that had durations greater than 415 µs were fitted with a mixture of two Gaussian components using method of maximum likelihood. Distributions of open periods, shut times, burst lengths and total open time per burst were fitted with either mixtures of exponential components using the method of maximum likelihood. Bursts of single-channel openings were defined by a critical gap length (tcrit) that was calculated to give equal numbers of misclassified events, i.e. the number of gaps that were incorrectly assigned as being within a burst when they were in fact between bursts was equal to the number that were classified as being between bursts when they were in fact within bursts (see Magleby & Pallotta, 1983; Clapham & Neher, 1984). To examine the frequency of direct transitions between two conductance levels the durations of the open period before and after the direct transition needed to be longer than two filter rise times (332 µs) to be included in the analysis. A critical amplitude (A crit) was determined to separate the two conductance levels that produced equal numbers of misclassified events. This amplitude is denoted by the dashed line in the asymmetry plots. In such histograms the occurrence of a transition between two states (represented as the ith amplitude level followed by the (i+1)th amplitude, where i is an integer) is indicated by a single point.
Concentration jump experiments
Fast concentration jumps were achieved by the rapid switching, across the tip of a patch pipette, of two solutions flowing from either side of a piece of theta glass. The control solution was supplemented with 20 µM glycine, and the test solution contained 10 mM glutamate (+ 20 µM glycine). Outside-out patches, held at –100 mV, were exposed to glutamate for 100 ms, and jumps were made every 10 s. Measurement of liquid junction potentials at the end of experiments indicated that the exchange of control and test solutions occurred within 300 µs. The macroscopic agonist-evoked currents were filtered at 2 kHz and sampled at 10 kHz onto a computer hard disk via a CED 1401 Plus interface (CED Instruments, Cambridge, UK) using the computer program CJUMP. Individual jumps were averaged and the resulting mean current fitted (least squares) with a single exponential curve using the CJFIT program (available from http://www.ucl.ac.uk/Pharmacology/dc.html).
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Results |
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Figure 1A shows a partial amino acid sequence alignment of bacterial amino acid binding proteins and mammalian glutamate receptor subunits. Within the S2 region of NMDA NR2 receptor subunits and AMPA/kainate receptor this threonine residue is conserved. Figure 1B shows a cartoon of the topology of the NR2D subunit and the location of the S1 and S2 regions that form a ‘clamshell’ structure to bind glutamate and the location of the T692 residue in the NR2D subunit is highlighted.
NR1a/NR2D(T692A) receptor-channels show a reduction in glutamate potency
Figure 2 shows individual responses recorded under two-electrode voltage clamp to increasing concentrations of glutamate (in the presence of 20 µM glycine) from an oocyte expressing wild-type NR1a/NR2D NMDA receptor-channels (Fig. 2A) and from an oocyte expressing NR2D subunits carrying the T692A point mutation (Fig. 2B). It is apparent from these panels that considerably higher concentrations of glutamate are required to evoke responses in the oocyte expressing the mutant receptor-channel. However currents evoked by glutamate in oocytes expressing NR1a/NR2D(T692A) receptors (Imax= 290 ± 48 nA, n= 9) were slightly greater than those evoked by glutamate in oocytes expressing wild-type NR1a/NR2D receptors (Imax= 153 ± 34 nA, n= 5). The fact that we did not observe any reduction in the maximal current in oocytes injected with a similar amount of cRNA coding for mutated NR2D receptor compared to those injected with the cRNA for the wild-type subunit indicates that this mutation is unlikely to affect, to any great extent, the gating of the channel. A series of concentration–response curves were generated from oocytes expressing either NR1a/NR2D and NR1a/NR2D(T692A) receptor-channels. The pooled data are shown in Fig. 2C and illustrate clearly that the NR2D(T692A) mutation shifts the glutamate dose–response curve far to the right compared to the wild-type. The glutamate potency for the NR1a/NR2D(T692A) receptor-channels (glutamate EC50= 703 ± 43 µM, n= 9) is reduced by 1562-fold compared to wild-type (glutamate EC50= 0.45 ± 0.04 µM, n= 5). Despite the shift in glutamate potency there is little change in the Hill slope for the concentration–response curve of the mutant receptors when compared to wild-type receptor-channels (1.50 ± 0.13 and 1.31 ± 0.15, respectively). To characterize further the effects of the threonine to alanine mutation in the NR2D receptor subunit we examined the potency of two other NMDA receptors agonists on both wild-type NR2D- and NR2D(T692A)-containing receptor-channels. For both NMDA and aspartate there is a clear rightward shift in potency for responses recorded from receptor-channels carrying the point mutation. Overall the EC50 and Hill slope (nH) values for these curves are: for aspartate (Fig. 2D), EC50(WT)= 3.3 ± 0.3 µM, nH= 1.13 ± 0.1; EC50(T692A)= 191 ± 20 µM, nH= 1.29 ± 0.1; and for NMDA (Fig. 2E), EC50(WT)= 3.7 ± 0.2 µM, nH= 1.37 ± 0.1; EC50(T692A)= 117 ± 7 µM, nH= 1.58 ± 0.1 (n= 5–9 for each curve). Thus the mutation causes an approximately 1600-fold shift in glutamate potency but more modest shifts in aspartate and NMDA potencies of approximately 60-fold and 30-fold, respectively. In contrast to its effects on glutamate, NMDA and aspartate potencies, the mutation has little effect on glycine potency (Fig. 2F), with EC50(WT)= 0.08 ± 0.01 µM and EC50(T692A)= 0.11 ± 0.01 µM (n= 4). This is in agreement with our previous demonstration that a series of mutations in the S1 and S2 ligand binding domains of the NR2A NMDA receptor subunit have little effect on glycine potency (Anson et al. 1998).
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Single-channel amplitudes and transition asymmetry
Figure 3 compares the single-channel events evoked by glutamate and recorded at a holding potential of –100 mV from an outside-out membrane patch excised from either an oocyte expressing wild-type NR1a/NR2D channels (Fig. 3A) or one expressing NR1a/NR2D(T692A) channels (Fig. 3B). It is clear that the point mutation does not affect the amplitude of these events when compared to wild-type. NR1a/NR2D(T692A) channels displayed two conductance levels that are typical of NR2D-containing NMDA receptors. Amplitude histograms from two recordings are shown in Fig. 3C and D and each is fitted with a mixture of two Gaussian components. Overall the mean amplitudes (and relative areas) for the larger (main) conductance level (recorded at –100 mV) were 4.05 ± 0.19 pA (61 ± 4%) and 3.97 ± 0.10 pA (69 ± 3%) for wild-type (n= 4) and mutant (n= 6) receptor-channels, respectively. The mean values of the lower (sub) conductance level were 2.24 ± 0.07 and 2.03 ± 0.07 pA for wild-type and mutant receptor-channels, respectively. The values for the amplitudes and areas for each component in the distributions of wild-type and mutant receptor-channels are not significantly different (P > 0.05 in all tests). The values we report here for the amplitude of the main and subconductance levels are greater than those we reported in our previous study of recombinant NR1a/NR2D NMDA receptor-channels, also expressed in oocytes (see Wyllie et al. 1996). However, in our earlier study the internal recording solution was Na+-based (rather than K+-based, as used in the present study) and this may account for the differences we have observed.
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We also examined the properties of NR1a/NR2D(T692A) channels when activated by either aspartate (100 µM) or NMDA (100 µM) (n= 3 patches for each agonist) (Fig. 4). The amplitudes of NR1a/NR2D(T692A) single-channel currents evoked by aspartate or NMDA were not significantly different from those produced by glutamate and on average were 3.96 ± 0.27 pA (relative area 62 ± 2%) and 2.02 ± 0.17 pA (relative area 38 ± 2%) for aspartate-activated events and 3.98 ± 0.36 pA (60 ± 4%) and 1.99 ± 0.25 pA (40 ± 4%) for NMDA-activated single-channel currents. Typical amplitude histograms for aspartate- and NMDA-evoked single-channel currents are illustrated in Fig. 4A and B. As was the case for glutamate, aspartate and NMDA, each gave rise to events that displayed temporal asymmetry in direct transitions (for examples see Fig. 4C and D). The mean values for the percentage of transitions starting in the main conductance level and going to the subconductance level were 64 ± 0.3% for aspartate-evoked events and 60 ± 3% for NMDA-evoked events. These values are not statistically different from those obtained from recordings made with glutamate as the agonist.
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Table 1 summarizes the mean values for open periods, shut times and burst lengths for glutamate-evoked NR1a/NR2D(T692A) channel activity.
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4 ranging between 53 and 557 ms and 5 ranging between 161 and 2548 ms. Given this variability, we made the assumption that these two components represented shut times that occurred between separate activations (bursts) of channel activity. This is in contrast to our findings for wild-type NR1a/NR2D channels where only the last component of the shut time distribution was considered to represent shut times between separate activations. In these shut time distributions the time constant of the fifth component varied only 2-fold (140–290 ms; see Wyllie et al. 1998). The fact that two shut time components contained within the NR1a/NR2D(T692A) distributions are considered to be between bursts is analogous with the observation we made with NR1a/NR2A channels carrying the homologous T671A point mutation, where we concluded that the latter two components represented gaps between separate activations of these receptor-channels (see Anson et al. 2000). A tcrit was calculated as the gap most likely to separate the shut times contained within the third and fourth components of each distribution and thus allow us to identify individual activations (bursts) of channel openings. The mean tcrit calculated by the method of misclassifying equal numbers of events from each component of the distribution (Clapham & Neher, 1984) was 20.8 ± 4.4 ms. In the distribution shown in Fig. 5B the tcrit is 23 ms. Burst lengths. Following identification of an appropriate tcrit, distributions of burst lengths were constructed. The distribution resulting from imposing a tcrit of 23 ms on the data illustrated in Fig. 5A and B is shown in Fig. 5C. Burst length distributions were best fitted with a mixture of three exponential components. Distributions of wild-type NR1a/NR2D individual channel activations show extremely long-lasting bursts of activity – the slowest two components in such distributions have time constants of around 1400 and 5000 ms (see Wyllie et al. 1998). Such long-lasting bursts are missing in NR1a/NR2D(T692A) distributions where the longest component has a mean of 66.7 ms. Thus the T692A mutation results in an approximately 80-fold reduction in the time constant of the slowest component contained within burst length distributions. Analysis of distributions of total open time per burst indicated that these were best described by a mixture of three exponential components and had an overall mean of 3.60 ± 1.03 ms which gave a probability that a channel was open during a burst of 0.074 ± 0.03. This open probability is not significantly different from that observed for wild-type NR1a/NR2D channels of 0.04 ± 0.02 (P > 0.05) (see Wyllie et al. 1998). The top three traces in Fig. 5D show three bursts of NR1a/NR2D(T692A) channel openings while the lower trace in this panel is an example of a wild-type NR1a/NR2D burst but on a 100-fold slower time base.
Deactivation of macroscopic currents mediated by NR1a/NR2D(T692A) channels
Assuming we have correctly identified individual activations of NR1a/NR2D(T692A) channels we would predict that the deactivation of NR1a/NR2D(T692A) receptor-mediated currents would be considerably faster than those seen for wild-type channels, which decay with a time constant of between 3 and 5 s (see Monyer et al. 1994; Vicini et al. 1998; Wyllie et al. 1998; Misra et al. 2000).
Rapid applications of glutamate (10 mM) were applied for 100 ms to outside-out patches, held at –100 mV, and excised from oocytes expressing NR1a/NR2D(T692A) receptor-channels. We used a 100 ms application to ensure that we recorded a plateau to the current, as if the T692A mutation affects the dissociation rate of glutamate, briefer applications may not result in an equilibrium of agonist occupancy of the receptor being achieved. Figure 6A shows three individual sweeps recorded from a single experiment in which individual channel openings can be distinguished. Figure 6B shows the average of 75 jumps where the decay of the current following the removal of glutamate is fitted with a single exponential component with a time constant of 79 ms. The dashed line in Fig. 6B shows the comparable decay of wild-type NR1a/NR2D receptors. On average the deactivation of the NR1a/NR2D(T692A) channels could be described by a single exponential with a time constant of 72.6 ± 19.3 ms (n= 7). This is approximately 60-fold faster than the deactivation of wild-type NR1a/NR2D receptor-channels (Wyllie et al. 1998) and this difference is illustrated in Fig. 6C where the deactivation of the mutant and wild-type NR2D-containing NMDA receptors have been superimposed.
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Discussion |
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A number of structure–function studies have identified the residues in the S1 and S2 domains of the glutamate receptor subunits that control glutamate binding (Kuusinen et al. 1995; Paas et al. 1996; Laube et al. 1997; Anson et al. 1998, 2000; Wo et al. 1999). These findings have correlated well with evidence obtained from the analysis of the crystal structure of the GluR2 S1S2 protein fragment (Armstrong et al. 1998; Armstrong & Gouaux, 2000). Previously, we have reported the effects of mutating a threonine residue in the S2 domain of the NR2A subunit, which results in the disruption of glutamate binding to this receptor subunit and alters the kinetic behaviour of single-channel currents (Anson et al. 1998, 2000).
In the present study we have mutated the homologous residue in the NR2D subunit. This threonine residue is conserved throughout the ionotropic glutamate receptor subunit family in subunits that bind glutamate (see Fig. 1A), implying that this residue plays a pivotal role in glutamate binding, not only among NMDA NR2 receptor subunits but also within the AMPA and kainate receptor families. Indeed, X-ray crystallographic studies have confirmed the role of the threonine residue at position 655 (the homologous residue in the GluR2 subunit) in agonist binding of the 
-carboxyl group of glutamate to the GluR2 S1S2 protein fragment (Armstrong & Gouaux, 2000). It is intriguing to note that in both NR1 and NR3 subunits this threonine residue is absent and indeed in the NR3 subunit it is substituted with an alanine (Ciabarra et al. 1995; Sucher et al. 1995; Chatterton et al. 2002; see alignment in Fig. 1A) – thus, in part, this may explain the insensitivity of the NR3 subunits to activation by glutamate.
The reduction in agonist potency is consistent with a change in binding
The NR2D(T692A) mutation reduced the glutamate potency of NR1a/NR2D(T692A) receptors by three orders of magnitude, with little change in Hill slope compared to NR1a/NR2D wild-type receptors. The shift of around 1600-fold in glutamate potency is more than that seen with aspartate (approximately 60-fold shift) and NMDA (approximately 30-fold shift). As the EC50 for any agonist acting on a receptor is dependent on all the rate constants in the reaction scheme that describes its activation, it might be anticipated that we should have observed a change in the EC50 for the co-agonist, glycine, since the T692A mutation has, most likely, increased the dissociation rate constant for glutamate. However, the concentration–response curves for glycine were generated under conditions where NR2 receptor subunits were fully liganded by the saturating concentrations of glutamate used in these particular experiments (likewise, glutamate, aspartate and NMDA concentration–response curves were generated in the presence of saturating concentrations of glycine). Thus, the fact that we do not see a change in EC50 for glycine in these experiments is evidence against the idea that the T692A mutation affects the rate constants describing the binding of glycine to the NR1 receptor subunit.
Clearly a major difference between the agonists glutamate, aspartate and NMDA is in the length of the carbon chain backbone of the side chain of these ligands. The fact that the potencies of aspartate and NMDA are reduced to a lesser extent may suggest that the hydrogen bonding of these agonists to this threonine residue can be compensated for in a manner that is not achievable when glutamate occupies the binding site. Until a crystal structure for an NMDA NR2 binding site is elucidated it remains to be seen as to how various ligands orientate in the binding site and indeed whether different NMDA receptor agonists induce different degrees of domain closure, as has been reported to be the case when full and partial agonists occupy the ligand binding site of the GluR2 S1S2 protein fragment (Armstrong & Gouaux, 2000; Jin & Gouaux, 2003; Jin et al. 2003).
A reduction in glutamate potency could be explained not only by a change in binding but by also a change in gating (Colquhoun, 1998). However, such a large reduction in glutamate potency with little change in maximal response and Hill slope cannot be explained by a change in gating alone. If a change in gating had caused a 1600-fold shift in glutamate EC50, we would expect the maximal response to be reduced by a factor of about 2.5 million (see Colquhoun, 1998). This has not been observed here given that the maximal responses recorded from oocytes injected with similar amounts of cRNA were similar and is consistent with the data obtained from receptors containing the homologous mutation (T671A) in the NR2A subunit (Anson et al. 1998). Indeed wild-type NR2D-containing NMDA receptors have an extremely low probability of being open during an activation and are in fact closed for around 96% of the activation (see Wyllie et al. 1998). This observation implies that the ‘efficacy’ of glutamate at NR2D-containing receptors is extremely low. Consistent with the idea that gating is little affected by this mutation is our observation that the single-channel conductance of NR2D(T692A)-containing receptors is not different from wild-type. In as much as transitions between the main conductance and subconductance levels can be defined as a ‘gating’, the NR2D(T692A) mutation did not alter the temporal asymmetry of transition frequency between these two conductance levels when the channels were activated by any of the agonists used in this study. We did, however, observe a significant decrease in the mean value of the time constant of the third component of open period distributions of NR1a/NR2D(T692A) events when compared to wild-type. This reduction accounts for the difference in the overall mean value for open periods of the mutant (0.92 ms) and wild-type channels (1.50 ms; see Wyllie et al. 1998). Although this small decrease in open period duration cannot possibly account for the large change in glutamate potency observed, it may imply that coupling of the binding site to the activation gate of the channel has been disrupted in such a way as to increase the rate of channel closure from this open state. Given our data, we suggest that the most likely explanation for the differences in the potency shifts seen with glutamate, aspartate and NMDA is that they are attributable to the way these agonists interact with the binding site rather than any effect on gating properties of the channel.
Single-channel and deactivation properties
Perhaps the most striking kinetic feature of NR1a/NR2D channel activations is their very long-lasting duration coupled with the fact that the channel remains closed for most of the activation (Wyllie et al. 1998). The slowest component in the burst length distribution is around 5000 ms, which is to be expected given the fact that deactivation of channel activity following brief exposure to glutamate of both native and recombinant NR2D-containing NMDA receptors takes several seconds (for example see Monyer et al. 1994; Vicini et al. 1998; Wyllie et al. 1998; Misra et al. 2000). In order to identify NR1a/NR2D(T692A) bursts of opening we chose to use a critical gap length that separated the third and fourth components contained within the shut time distributions. The first three components in such distributions showed the least variability, suggesting that they were not dependent on agonist concentration. This, together with the fact that deactivation rates of NR1a/NR2D(T692A) currents are considerably more rapid that those seen with wild-type, makes it unlikely that a different value of tcrit would correctly identify individual activations. The slowest component contained within distributions of NR1a/NR2D(T692A) burst lengths has a duration approximately 100 times less than the slowest component in corresponding distributions of wild-type burst lengths. From the values shown in Table 1 it can be seen that the overall probability of the channel being open during a burst is still low (< 0.1).
Deactivation times of macroscopic responses from NR1a/NR2D(T692A) receptors, after brief applications of glutamate, were around 60 times less than wild-type NR1a/NR2D receptors. Similarly an increase in the rate of decay was observed for NR1a/NR2A(T671A) receptors. Given that the deactivation of the macroscopic current is determined by the underlying kinetic behaviour of the single-channel events this increased deactivation rate is entirely expected. Indeed if one uses the model for NMDA receptor activation proposed by Lester & Jahr (1992) and increases the rate of dissociation by a factor of 1600 then the predicted change in the deactivation rate of a simulated macroscopic current is around 10-fold. This is not surprising, as the duration of individual activations will depend not only on the dissociation rate of the agonist but also on the duration and the number of openings (as well as the durations of shut times) contained within an activation. Therefore, just as changes in the EC50 are difficult to ascribe to alterations in either binding or gating parameters, the effect of altering the dissociation rate constant is difficult to quantify without knowledge of a kinetic scheme that adequately describes receptor activation. Recently, variations of the kinetic scheme originally proposed by Lester & Jahr (1992) describing the activation of recombinant NMDA receptors have been published (see Banke & Traynelis, 2003; Popescu & Auerbach, 2003). Neither of these newer schemes can account for the complex activation structure of NR2D-containing NMDA receptors and it remains to be seen whether an ‘all encompassing’ scheme, in which only the rate constants describing the entry to and exit from different states vary, can describe the heterogeneity of NMDA channel activations seen with different NR2 combinations. If this can be achieved then it may be easier to identify how alterations in the dissociation rate constant influence the observed EC50 for agonist action and more importantly the nature of individual channel activations. Nonetheless the channel properties we have described for NR1a/NR2D(T692A) receptors are entirely consistent with the notion that this mutation primarily affects the ability of glutamate to remain bound to the receptor.
NR2D-containing NMDA receptors appear to play an unusual and perhaps special functional role for ionotropic glutamate receptors due to their apparent exclusion from synapses (see for example Momiyama, 2000; Brickley et al. 2003) and their very slow deactivation kinetics. While our results do not address specifically the role played by such receptors they do indicate that the long-lasting activations most likely are a consequence of glutamate remaining bound to this receptor for the duration of the activation rather than initiating a series of conformational changes in the protein from which the channel can open and close in the absence of bound ligand.
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Footnotes |
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References |
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) or NR1a/NR2D(T692A) channels (•) and fitted with the Hill equation as described in the Methods. F, concentration–response curves evoked by increasing concentrations of glycine in the presence of a saturating concentration of glutamate (10 µM for wild-type or 10 mM for mutant) for currents recorded from oocytes expressing either wild-type NR1a/NR2D channels (
