Nature Neuroscience
- 9, 1388 - 1396 (2006)
Published online: 15 October 2006; | doi:10.1038/nn1793
Activation of a presynaptic glutamate transporter regulates synaptic transmission through electrical signalingMargaret Lin Veruki1, 2, Svein Harald Mørkve1, 2 & Espen Hartveit11 University of Bergen, Department of Biomedicine, Jonas Lies vei 91, N-5009 Bergen, Norway. 2 These authors contributed equally to this work.
Correspondence should be addressed to Espen Hartveit espen.hartveit@biomed.uib.no Whereas glutamate transporters in glial cells and postsynaptic neurons contribute significantly to re-uptake of synaptically released transmitter, the functional role of presynaptic glutamate transporters is poorly understood. Here, we used electrophysiological recording to examine the functional properties of a presynaptic glutamate transporter in rat retinal rod bipolar cells and its role in regulating glutamatergic synaptic transmission between rod bipolar cells and amacrine cells. Release of glutamate activated the presynaptic transporter with a time course that suggested a perisynaptic localization. The transporter was also activated by spillover of glutamate from neighboring rod bipolar cells. By recording from pairs of rod bipolar cells and AII amacrine cells, we demonstrate that activation of the transporter-associated anion current hyperpolarizes the presynaptic terminal and thereby inhibits synaptic transmission by suppressing transmitter release. Given the evidence for presynaptic glutamate transporters, similar mechanisms could be of general importance for transmission in the nervous system.Following release from a synaptic terminal, glutamate binds to pre- and postsynaptic receptors before it is rapidly cleared from the synaptic cleft by diffusion and taken up by specific, high-affinity glutamate transporters1. Glutamate transporters are located in glial cells2 and in some postsynaptic neurons3, but there is also evidence for a presynaptic localization4,
5,
6,
7,
8,
9,
10,
11,
12. The function of such presynaptic neuronal transporters is poorly understood. They could contribute to maintaining the presynaptic transmitter pool1, to the clearance of glutamate7 or to the regulation of synaptic transmission by acting as a negative feedback signal on presynaptic release9,
10. Whereas all glutamate transporters (excitatory amino-acid transporters, EAAT, 1–5) are electrogenic13, transport also generates a non–stoichiometrically coupled anion current14,
15. It has been proposed that the transporters with the most prominent anion current (EAAT4 and EAAT5)14,
15,
16 are strong candidates for regulating presynaptic function14,
16, possibly by regulating transmitter release.
In the retina, there is evidence for presynaptic glutamate transporters in photoreceptors5,
7,
10,
11,
12 and bipolar cells7,
9. We investigated the functional properties of a glutamate transporter in axon terminals of rat retinal rod bipolar cells and examined its role in excitatory synaptic transmission between these cells and postsynaptic AII amacrine cells. By recording simultaneously from pairs of synaptically connected cells, we demonstrate a role for the presynaptic transporter-associated anion current in regulating synaptic transmission by hyperpolarization or shunting inhibition of presynaptic axon terminals.
Results
A glutamate transporter at rod bipolar cell axon terminals
We determined the expression of a glutamate transporter in rod bipolar cells by voltage-clamp recording in retinal slices, with the recording pipette positioned at the cell body or the axon terminal (ref. 17 and Fig. 1). Pulses of glutamate from a puffer pipette evoked an inward current (Fig. 1b; peak amplitude 19.9  3.1 pA (mean s.e.m.); n = 7) that was reversibly blocked (to 3 1% of control; n = 3; Fig. 1b) by DL-threo- -benzyloxyaspartic acid (TBOA), a nonselective, nontransported blocker of glutamate transporters18. With symmetrical chloride (chloride equilibrium potential (ECl)  3 mV), the reversal potential (Erev) of the glutamate-evoked response was 0.1 2.2 mV (n = 3) and the I-V curve displayed inward rectification (ref. 9 and Fig. 1c,d). The Erev followed changes in ECl (Erev was -51 mV for ECl = -43 mV; n = 2; data not shown).
 | | Figure 1. Localization of a glutamate transporter at axon terminal of rod bipolar cells. | | |  | (a) Videomicrograph of Lucifer yellow–filled cell. Scale bar, 10  m. (b) Left, current activated in rod bipolar cell by glutamate ('Glu', 100 M, 1.5 s) from puffer pipette. Right, TBOA blocked the response. Duration of drug application indicated by horizontal bar above current trace. Whole-cell recordings with blockers of GABAA, GABAC, glycine and ionotropic glutamate receptors. (c) Current-voltage (I-V) graph of glutamate-evoked response. Glutamate (1 mM, 500 ms) applied at membrane potentials from -80 mV to +60 mV, (10 mV steps). Each trace, average of three trials. (d) Averaged I-V relationship of glutamate-evoked response (peak current; mean s.e.m.; n = 3). Data points normalized to response at -80 mV. (e) Spatial sensitivity profile to pressure-applied glutamate. Left, photomicrograph of Lucifer yellow–filled cell (puffer pipette in lower right corner). Right, responses to glutamate (100 M, 5 ms). Vertical position of each trace corresponds to that of puffer pipette. Co2+ replaced Ca2+ in extracellular solution. Scale bar, 10 m. (f) Top, current activated in outside-out axon terminal patch by glutamate (100 M, 500 ms) from puffer pipette. Bottom, TBOA blocked the response. (g,h) I-V properties (g) and averaged I-V relationship (h) of glutamate (1 mM, 100 ms, ultrafast application) response in outside-out axon terminal patches. Data in g show responses at -80, -60 and -40 mV. Data in h represent peak current (mean s.e.m.; n = 3–12 patches per data point). Data points normalized to response at -80 mV. Intracellular solutions: D (CsCl; b–d, ECl  3 mV; see Methods for details); E (KSCN; e–h).
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We measured the spatial sensitivity of the response by systematically shifting the position of the puffer pipette between the dendrites in the outer plexiform layer and the axon terminal in the inner part of the inner plexiform layer (Fig. 1e). Responses with the largest amplitudes and shortest onset and rise times were observed when the puffer pipette was located closest to the axon terminal in the inner plexiform layer (Fig. 1e; n = 4), suggesting a high concentration of glutamate transporter molecules in this region7,
9.
To verify the location of a transporter at rod bipolar axon terminals, we isolated outside-out patches from axon terminals in the slice. At a holding potential of -80 mV (ECl 3 mV), the application of glutamate evoked a small inward current (average 0.7 0.2 pA; n = 4; data not shown). When intracellular chloride (Cl-) was replaced by thiocyanate (SCN-), which has a higher permeability in the associated anion channel of glutamate transporters19, the glutamate-evoked response increased to 3.9 1.1 pA (Fig. 1f; n = 3). Patch responses to glutamate were blocked completely and reversibly by TBOA (Fig. 1f; n = 3). The response displayed strong inward rectification and no reversal at holding potentials up to +50 mV (Fig. 1g,h; n = 12 patches). These results suggest that rat rod bipolar cells express a glutamate transporter in the axon terminal region.
Fast kinetics of glutamate transporters at axon terminals
We studied the kinetics of the transporter with ultrafast application of glutamate to outside-out patches from rod bipolar axon terminals. With brief pulses of glutamate, the response rose rapidly to a peak (8.8 1.2 pA; n = 11), with a 20–80% rise time of 439 23 s (Fig. 2a), similar to that measured for AMPA receptors in patches from AII amacrine cells20. The latency to onset of the response, measured from the time of solution exchange (Methods), was 330 s. The time course of decay was best fitted with a double-exponential function ( fast = 12.9 2.1 ms; 76 3% amplitude contribution; slow = 54 5 ms; Fig. 2a).
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With longer pulses of glutamate (100 ms or 500 ms long), the 20–80% rise time (431 44 s; n = 10) and peak amplitude (8.0 1.4 pA) were similar to the corresponding values obtained for brief pulses. After the peak, the response decayed rapidly to a plateau (Fig. 2b). The decay was best fitted with a double-exponential function (fast = 4.0 0.8 ms; 82 3% amplitude contribution; slow = 203 28 ms; n = 10). The ratio of peak amplitude to steady-state amplitude was 1.55 0.05 (n = 10). The deactivation phase after termination of the glutamate pulse was well fitted with a double-exponential function (n = 10; Fig. 2b) with fast = 17 6 ms (70 6% amplitude contribution) and slow = 91 17 ms.
We estimated concentration-response properties by applying pulses of glutamate to outside-out patches (Fig. 2c,d). For the peak response, the effective concentration for half-maximum response (EC50) was 31.9 M and the Hill coefficient was 1.0. For the steady-state response, EC50 was 16.9 M and the Hill coefficient was 1.1. Thus, 50% of the maximum activity was reached at glutamate concentrations that were only moderately higher than estimates for the extracellular concentration in the CNS (0.2–7 M; ref. 21). The response kinetics depended strongly on the concentration of glutamate. At lower concentrations (1–10 M), the response rose relatively slowly to a plateau without inactivation, whereas at higher concentrations (100 M to 10 mM), the response rose rapidly to a peak and then decayed to a steady-state plateau (inactivation; Fig. 2c,e).
Synaptic release of glutamate activates the transporter
To determine whether synaptically released glutamate from a rod bipolar cell terminal can activate glutamate transporters on that same cell, we took advantage of the observation that exocytosis and glutamate release in bipolar cells undergo strong paired-pulse depression9,
22,
23,
24, which does not depend on a depression of Ca2+ influx. To measure glutamate release, we applied a paired-pulse stimulus to a rod bipolar cell that we recorded simultaneously with a synaptically connected AII amacrine cell. Both depolarizing pulses evoked an inward current in the rod bipolar cell, but the amplitude was larger for the first response (Fig. 3a). In the AII cell, the first stimulus evoked a large inward current (mediated by AMPA receptors20,
25), whereas the second stimulus evoked a much smaller response (Fig. 3a), reflecting reduced release of glutamate23,
24. For the rod bipolar cells, the difference current between the first and second responses (Fig. 3a, inset) had an onset of 4.1 0.7 ms, a 20–80% rise time of 4.4 0.8 ms (similar to that observed in ref. 9) and a peak amplitude of 17.6 4.1 pA (n = 4). The excitatory postsynaptic current (EPSC; evoked by the first stimulus) in the AII amacrine cell had an onset of 1.7 0.1 ms and a 20–80% rise time of 594 67 s (similar to the results of ref. 25). Both the onset and the rise time were significantly longer for the rod bipolar cell difference current than for the AII amacrine EPSC (Fig. 3a, inset; P < 0.05; paired t-test). When difference currents were recorded directly from rod bipolar axon terminals (Fig. 3b), the latency to onset was 2.9 0.3 ms, the 20–80% rise time was 3.6 0.5 ms and the peak amplitude was 12.3 2.4 pA (n = 7), not significantly different from the values for difference currents measured at the soma (P > 0.12). The difference current was reversibly blocked by TBOA (Fig. 3c), suggesting that it primarily corresponded to a glutamate transporter–mediated current. Replacing Ca2+ in the bath solution with Co2+ blocked both the presynaptic, voltage-gated Ca2+ current and the associated glutamate transporter–mediated current (see below).
| | Figure 3. Synaptic release of glutamate activates the transporter in rod bipolar cells. | | |  | (a) Simultaneous, dual whole-cell recordings of synaptically connected rod bipolar cell (RB) and AII amacrine cell (AII). Diagram at left indicates recording configuration, both cells in voltage clamp (RB at left; AII at right). Top, paired-pulse stimulus applied to rod bipolar cell (interpulse interval 100 ms); middle and bottom, corresponding responses in RB and AII. Notice strong paired-pulse depression of response in AII. Each trace, average of three trials. Inset, overlaid and normalized traces showing response of AII cell to first stimulus and difference current of RB cell (response to first stimulus minus response to second stimulus). Notice difference in rise time. Intracellular solutions: B (CsCl; RB) and H (potassium gluconate; AII). GABAA, GABAC, glycine and NMDA receptors were blocked pharmacologically. (b) Top, overlaid responses of RB cell to first (1) and second (2) depolarization of paired-pulse stimulus (as in a); bottom, difference current (1 - 2). Inflection during rising phase of inward current (arrow) probably corresponds to transient inhibition of the Ca2+ current caused by synaptic cleft acidification48. (c) As in b (and for the same RB cell), but now in the presence of TBOA. Note that the difference current (1 - 2) was blocked by TBOA. In b and c, GABAA, GABAC and glycine receptors were blocked pharmacologically. Diagram at left indicates whole-cell voltage-clamp recording at axon terminal (b and c). Intracellular solution: D (CsCl; b,c).
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The fast rise time we measured in patches rules out the hypothesis that the slower rise of the presynaptic response to synaptically released glutamate is due to inherently slow kinetics of the glutamate transporter. The fast rise time of the EPSC in the AII amacrine cells rules out the hypothesis that slow, asynchronous release from the rod bipolar cell is responsible. Instead, our results suggest that the transporter proteins on axon terminals are, on average, located further from release sites than the postsynaptic glutamate receptors on AII cells. If we compare the rise times observed during local, synaptic activation with those in patches (Fig. 2e), a rough estimate suggests that, on average, the glutamate transporter molecules on axon terminals sense a peak glutamate concentration of approximately 100 M.
Spontaneous release of glutamate activates the transporter
In recordings from rod bipolar axon terminals with intracellular Cl- replaced by SCN-, we observed spontaneous, low-amplitude (5.3 0.3 pA; n = 13), rapidly rising and slowly decaying inward currents that were reversibly blocked by TBOA (n = 7; Fig. 4a,b). The 20–80% rise time was 1.5 0.1 ms, significantly longer than that for the patch responses (P < 0.0001; Fig. 4c). The decay phase was well fitted with a double-exponential function (Fig. 4b) with fast = 14.1 0.8 ms (82 3% amplitude contribution) and slow = 76.5 12.2 ms, not significantly different from the time constants measured with brief application of glutamate to outside-out patches (P > 0.12). We observed a gradual, time-dependent reduction in the frequency of events, from 1.3 0.1 Hz measured 1–3 min after establishing the whole-cell configuration to 0.4 0.1 Hz measured approximately 10 min later (n = 10; P < 0.0001; paired t-test). Depolarizing axon terminals from -60 mV to -48 mV increased the frequency of events from 1.1 0.2 Hz to 1.8 0.4 Hz (n = 6; P = 0.024; paired t-test). These results suggest that the observed events are quantal transporter currents corresponding to single-vesicle (or multivesicular26) release of glutamate from the recorded cell. Similar responses have been seen in tiger salamander cone photoreceptors, caused by the photoreceptors' own vesicular glutamate release10.
| | Figure 4. Spontaneous glutamate release activates the transporter in rod bipolar cells. | | |  | (a) Axon terminal whole-cell voltage-clamp (Vhold = -60 mV) recording (depicted at left) displaying rapidly activating, spontaneous inward currents (top), blocked by TBOA (bottom). Intracellular solution: E (KSCN). (b) Average waveform of rapidly activating, spontaneous inward currents, same axon terminal as in a (dotted line; n = 154 events); double-exponential fit to decay phase (continuous line). (c) Average waveform in b (GluT quantal; dotted line) overlaid with average waveform of response of outside-out patch to brief, ultrafast application of glutamate (1 mM; same as in Fig. 2a). (c–e) Waveforms normalized by peak amplitude and onsets aligned by eye. Same time scale used for c–e. Average waveform in b (GluT quantal; dotted line) was overlaid with average waveform of (d) spontaneous EPSCs (n = 233 events) recorded in an AII amacrine cell20 (AII spEPSC) and (e) spontaneous IPSCs (n = 61 events) from same axon terminal (RB spIPSC). (f) Simultaneous, dual recording of neighboring rod bipolar cells illustrate slowly activating, spontaneous inward currents occurring synchronously in both cells (control, recovery), reversibly blocked by TBOA. Diagram at left indicates recording configuration with both cells in voltage clamp (Vhold = -60 mV). TBOA also reduced holding current (not illustrated). Intracellular solution: A (CsCl). In a,b and f, GABAA, GABAC, glycine and ionotropic glutamate receptors were blocked pharmacologically (Fig. 1b).
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The slower rise time of the presumed quantal transporter currents, compared to that observed for patch responses (Fig. 4c), suggests that even the transporter molecules closest to release sites are not as close as postsynaptic AMPA receptors on AII amacrine cells. This was supported by the observation that the rise time of quantal transporter currents was longer than the rise times of spontaneous EPSCs in AII amacrine cells20 (Fig. 4d). Spontaneous inhibitory postsynaptic currents (IPSCs), recorded in the same axon terminals, displayed 20–80% rise times that were also faster (425 17 s; n = 7) than those observed for the quantal transporter currents (Fig. 4e), suggesting that the rise times of the quantal transporter currents are not artificially prolonged because of high series resistance.
In continuous recordings from rod bipolar cells, we observed additional TBOA-sensitive currents, occurring at a frequency of 0.3 0.1 Hz with a peak amplitude of 7.2 0.7 pA (symmetrical chloride; n = 12; Fig. 4f). These currents were easily distinguished from presumed quantal transporter currents by their slower rise time (30.1 4.9 ms) and longer duration (half-width 137 15 ms). When recording from pairs of rod bipolar cells, we observed that these currents could occur synchronously in both cells (6 of 6 pairs; Fig. 4f). Both synchronously and asynchronously occurring currents were blocked by TBOA (n = 3 pairs; Fig. 4f) and could occur long after rundown of evoked transmitter release in whole-cell recordings (5–15 min). These results suggested that the currents were caused by spillover of glutamate following transient release from neighboring cells. Similar currents were observed in axon terminal recordings.
TBOA reduced the holding current of rod bipolar cells (7.6 0.7 pA; n = 10), suggesting that glutamate transporters might be tonically activated by extracellular glutamate in the slice. However, exposing outside-out patches from axon terminals to TBOA also reduced an inward holding current (1.0 0.2 pA; n = 3), suggesting that the transporter displays a substrate-independent anion 'leak' current16,
27,
28. Thus, for the TBOA-evoked reduction of holding current in the slice, we could not distinguish between substrate-dependent and substrate-independent mechanisms.
Spillover of glutamate activates the transporter
The results presented above suggest that glutamate released from rod bipolar cells escapes the synaptic cleft and activates transporters on neighboring rod bipolar cells. To test this directly, we recorded from pairs of rod bipolar cells. All recordings were performed from cells located below the surface layer. Depolarization of one cell to -45 mV or more positive potentials evoked an inward current ('lateral response') in the nonstimulated cell (Fig. 5a; 97 of 112 pairs). Lateral responses were typically evoked in each cell of a pair (Fig. 5a). The average center-to-center distance between cell bodies was 15.4 0.5 m (range 8.7–45.1 m; n = 97 pairs). For cell pairs without lateral responses, the average distance was 32.6 7.5 m (range 12.5–131 m; n = 15). Lateral responses were not observed when either cell had a severed axon (n = 3).
| | Figure 5. Spillover of glutamate between rod bipolar cells activates the transporter. | | |  | (a) Dual recording of neighboring rod bipolar cells. Left, recording configuration. Both cells in voltage clamp, depolarizing stimulus (to -20 mV; 100 ms) applied alternately to each cell. Right, responses when stimulus was applied to cell 1 (top) or to cell 2 (bottom). Notice lateral response (transient inward current) in nonstimulated cell. Transient current increase of stimulated cell (arrow in top trace for cell 1, bottom trace for cell 2) appeared when cell was repolarized to holding potential. At this point, voltage-gated Ca2+ current started deactivating, the driving force for chloride increased (ECl 0 mV) and GABAergic feedback29 and transporter-associated currents9 increased transiently. (b) I-V relationship of lateral response (peak current; mean s.e.m.; n = 6–7 cells per data point). (c) Lateral responses (bottom traces) with increasing depolarization (–50 mV to -25 mV; top trace) of stimulated cell. (d) Block by Co2+ (2.5 mM) of lateral response (and response in stimulated cell). Here and below, traces for stimulated (1) and nonstimulated (2) cell in black (control) and gray (test). (e–h) Bicuculline (e) TPMPA (f) and CNQX (g) did not block lateral response. In g: cells in perforated patch (amphotericin B), GABAergic feedback response in stimulated cell appeared as outward current during depolarization in control condition (Vm = -20 mV), suggesting ECl was more negative than -20 mV. Same scale bars for d–g. (h) TBOA blocked lateral response in nonstimulated cell and part of tail current in stimulated cell. Intracellular solutions: B (CsCl; a,c–f,h) and A (CsCl; b,g).
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Lateral responses displayed inward rectification (Fig. 5b) with Erev = -0.9 2.9 mV (ECl 3 mV; n = 5). With depolarization to -20 mV, the onset of the lateral response occurred at 7.9 0.5 ms, the 20–80% rise time was 7.2 0.7 ms and the average peak amplitude was 9.5 0.9 pA (n = 46). For each cell pair, the voltage threshold of the lateral response was similar to that of the reciprocal, self-inhibitory response caused by GABAergic feedback inhibition from postsynaptic amacrine cells25,
29 (–45 mV for 6 of 7 pairs; -50 mV for 1 of 7 pairs; Fig. 5c).
Both lateral and self-inhibitory responses were blocked by replacing Ca2+ with Co2+ extracellularly (Fig. 5d). The lateral response was not blocked by antagonists of GABAA (bicuculline; n = 5; Fig. 5e), GABAC ((1,2,5,6-tetrahydropyridine-4-yl) methylphosphinic acid, TPMPA; n = 3; Fig. 5f), non-NMDA receptors (6-cyano-7-nitroquinoxaline-2,3-dione, CNQX; n = 4; Fig. 5g) or NMDA receptors (3-(R)-2-carboxypiperazin-4-propyl-1-phosphonic acid, CPP; n = 3; data not shown), suggesting that it did not depend on the same synaptic circuitry as the self-inhibitory response29. In contrast, TBOA completely and reversibly blocked the lateral response (n = 6; Fig. 5h). This suggested that the lateral response is mediated by activation of a glutamate transporter on the nonstimulated cell, following spillover of glutamate from the synaptic cleft.
Lateral responses were observed with intracellular solutions with strong (5 mM EGTA) or weak (0.1 mM EGTA) Ca2+-buffering capacity in the stimulated cell. In perforated-patch recordings, we observed lateral responses qualitatively similar to those observed with standard whole-cell recordings (n = 4), but the peak amplitude was reduced as a result of the reduced driving force for chloride (ECl < 0 mV; Fig. 5g).
To examine whether increased activity of glutamate transporters in retinal cells at physiological temperatures would prevent spillover of glutamate30,
31,
32 between rod bipolar axon terminals, we recorded from pairs of rod bipolar cells at 34 °C. In this condition, the peak amplitude of the lateral response was 12.2 1.9 pA (n = 13), not significantly different from that at room temperature (P = 0.06). Synchronous and asynchronous spontaneous transporter currents similar to those observed at room temperature (Fig. 4f) were also observed at 34 °C (data not shown).
Transporter activation inhibits synaptic transmission
A potential function of a presynaptic glutamate transporter is suppression of transmitter release through hyperpolarization or shunting inhibition of the axon terminal mediated by the non–stoichiometrically coupled anion current9,
10. To investigate whether the glutamate-evoked conductance increase is large enough to influence the membrane potential of a rod bipolar cell, we first applied glutamate to the axon terminals of rod bipolar cells recorded in the perforated-patch configuration (using gramicidin to avoid perturbing intracellular chloride). In the voltage-clamp condition, Erev was -64.4 1.3 mV (n = 6; Fig. 6a), suggesting that ECl is close to this value33,
34,
35. In the current-clamp condition (with the same cells), glutamate application evoked hyperpolarizations at membrane potentials positive to Erev (–13.7 1.8 mV at -40 mV) and depolarizations at membrane potentials negative to Erev (11.6 1.9 mV at -90 mV; Fig. 6a). To systematically investigate the impact of the magnitude and Erev of such conductances on the integrative properties of rod bipolar cells, we used dynamic-clamp recording from axon terminals. We injected a conductance waveform (peak conductance 400 pS) and measured the change in membrane potential while systematically varying Erev of the current. At a membrane potential of -60 mV, the injected conductance evoked an inhibitory postsynaptic potential (IPSP) of -5.1 0.5 mV when Erev was -70 mV and an excitatory postsynaptic potential (EPSP) of 5.3 1.0 mV when Erev was -50 mV (n = 3; Fig. 6b). We also examined the ability of the transporter-associated anion conductance to act as a shunting inhibition by combining injection of a transient excitatory conductance (Erev = 0 mV; peak conductance 400 pS) with that of a steady-state inhibitory conductance (Erev = Vm). At a membrane potential of -60 mV, the excitatory conductance evoked an EPSP of 25 mV in the absence of shunting inhibition (Fig. 6c). When we added the inhibitory conductance (Erev = -60 mV), the size of the EPSP was reduced (Fig. 6c). On average, the peak amplitude of the EPSP was reduced by 1–2 mV per 100 pS of shunting conductance (Fig. 6c; n = 3).
| | Figure 6. Action and interaction of excitatory and inhibitory conductances at rod bipolar cell axon terminals. | | |  | (a–c) Left, recording configuration. Co2+ replaced Ca2+ in extracellular solution. Between trials, membrane potential returned to -60 mV. (a) Response to glutamate (100 M; 120–200 ms) applied to rod bipolar cells recorded at axon terminal (perforated patch; gramicidin). Middle, I-V relationship (peak current; mean s.e.m.; n = 6). Data points normalized to current at -90 mV. Right, glutamate-evoked responses in one of the same cells (current clamp). Note hyperpolarization at Vm = -40 mV, no response at -65 mV, depolarization at -90 mV. Each trace, average of 6–13 trials. Recordings in the presence of blockers of GABAA, GABAC, glycine and ionotropic glutamate receptors. (b) Conductance (Ginj) injected at axon terminal (Vm = -60 mV) evoked EPSPs or IPSPs depending on Erev (–80 to -20 mV; 10 mV steps). Each trace, average of 10 trials. Conductance waveform from lateral response with peak current of 24 pA (midway between population average and maximum response observed), peak conductance 400 pS (Erev 0 mV). Right, relation between Erev and PSP peak amplitude (mean s.e.m.; n = 3). (c) Amplitude of axon terminal EPSP evoked by excitatory conductance (Gexc; Erev = 0 mV) injected at axon terminal (Vm = -60 mV) was reduced in proportion to magnitude of shunting inhibitory conductance (Gshunt; Erev = -60 mV). Each trace, average of 10 trials. Right, reduction of EPSP peak with increasing magnitude of Gshunt (mean s.e.m.; n = 3). Intracellular solutions: G (KCl; a) and F (potassium gluconate; b,c).
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These results suggest that activation of the glutamate transporter in rod bipolar cell axon terminals can suppress transmitter release by counteracting stimulus-evoked depolarization. Testing this hypothesis directly, by blocking the transporter pharmacologically and examining the effect on the synaptic transmission between rod bipolar cells and AII amacrine cells, could be problematic, because TBOA evokes a steady-state desensitization of the AMPA receptors on AII amacrine cells (data not shown), presumably by increasing the extracellular concentration of glutamate. Instead, we measured the voltage response of rod bipolar cells to injection of an excitatory conductance waveform, first in the control condition and then in the presence of TBOA. The conductance waveform (Fig. 7a) mimicked a light-evoked, excitatory conductance measured in rod bipolar cells36. In the control condition (intact transporters), the EPSP displayed both a transient peak and a steady-state plateau (Fig. 7a). In the presence of TBOA (blocked transporters), the peak voltage excursion was 7.0 0.4 mV (n = 7) more depolarized than that in the control condition and the steady-state plateau was 4.2 0.6 mV more depolarized (Fig. 7a), presumably caused by the abolishment of the synaptic activation of the glutamate transporter and the consequent reduction of the inhibitory conductance (ECl = -62 mV; refs. 33,
34,
35). To examine whether these EPSPs could differentially influence the release of glutamate from rod bipolar cells, we used them as presynaptic voltage-clamp commands in paired recordings of synaptically connected rod bipolar cells and AII amacrine cells. The EPSP recorded in the presence of TBOA evoked an EPSC in the AII amacrine cell that rose faster to the peak (20–80% rise time 10.3 0.6 ms) than the EPSC evoked by the EPSP recorded in the control condition (20–80% rise time 17.3 0.3 ms; n = 3; P = 0.002; paired t-test; Fig. 7b).
| | Figure 7. Activation of glutamate transporter in rod bipolar cell axon terminals suppresses synaptic transmission. | | |  | (a) Voltage responses of rod bipolar cell (RB) to injection of excitatory conductance (Gexc; Erev = 0 mV; 10–90% rise and decay time 90 ms; 500 pS amplitude) in control (black) and with TBOA (red). Each trace, average of four trials. Left, current-clamp configuration. Right, rising phase of voltage responses at higher time resolution. GABAA, GABAC and glycine receptors blocked pharmacologically. (b) EPSCs in AII amacrine cell, evoked by voltage clamping the presynaptic rod bipolar cell with EPSP waveforms recorded in a (black, control; red, TBOA). Each trace, average of nine trials. Left, voltage-clamp configuration, command waveforms (Vcom) applied to rod bipolar cell. Notice shorter rise time of AII amacrine EPSC when voltage clamping rod bipolar cell with EPSP waveform recorded with TBOA. Right, initial phase of EPSCs at higher time resolution. (c) Increased EPSP amplitude in rod bipolar cell when glutamate release was reduced by paired-pulse stimulus. Left, recording configuration, rod bipolar cell in current clamp (perforated-patch; gramicidin), AII amacrine cell in voltage clamp. Rod bipolar cell injected with paired-pulse excitatory conductance waveform (Gexc; Erev = 0 mV; 10–90% rise and decay time 8 ms; 600 pS amplitude; 50 ms interpulse interval). Each trace, average of six trials. EPSCs of AII amacrine cell verified reduced release of glutamate after second compared to first stimulus. In b and c, GABAA, GABAC, glycine and NMDA receptors were blocked pharmacologically. Intracellular solutions: F (potassium gluconate) in a; C (CsCH3SO3; RB) and H (potassium gluconate; AII) in b; F (potassium gluconate; RB) and H (potassium gluconate; AII) in c.
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The effect of blocking the presynaptic glutamate transporter on the synaptic transmission between rod bipolar and AII amacrine cells could be due to either a steady-state or a transient inhibitory conductance, or a combination of both. To detect a functional role for a transient increase of transmitter release, we injected a paired-pulse excitatory conductance waveform into rod bipolar cells recorded simultaneously with postsynaptic AII amacrine cells. By recording the AII amacrine EPSCs evoked by the first and the second conductance stimulus, we verified the presence of paired-pulse depression of release (Fig. 7c). Notably, the voltage response of the rod bipolar cell to the second conductance stimulus, which evoked less glutamate release, reached a larger peak amplitude (5.3 0.5 mV more depolarized; P < 0.0001; paired t-test; n = 5 with conventional whole-cell recording (ECl = -62 mV); n = 4 with gramicidin perforated-patch recording) than the voltage response to the first conductance stimulus, and sometimes developed a longer-lasting depolarizing plateau (Fig. 7c). These results suggested that the second depolarization of the rod bipolar cell evoked a smaller transporter-mediated inhibitory conductance than the first depolarization, corresponding to a reduction in the amount of glutamate released.
Discussion
We investigated the functional properties of a glutamate transporter located on presynaptic axon terminals of rod bipolar cells in rat retina. Activation of the transporter was detected as an increase in the transporter-associated non–stoichiometrically coupled anion current. When the transporter was studied in outside-out patches, we found very fast activation kinetics, similar to those reported for glutamate transporters in other systems27,
28,
37,
38 and to AMPA receptors on AII amacrine cells20 postsynaptic to rod bipolar cells. When activated by synaptically released glutamate, however, the activation kinetics were significantly slower. A comparison of the concentration dependence of the glutamate-transporter kinetics (in patches) with the time course of synaptically evoked responses, suggests that the glutamate transient at the presynaptic transporter reaches a lower concentration than that attained in the synaptic cleft27,
37. This suggests that most of the transporter molecules are located further from release sites than postsynaptic AMPA receptors are, most probably perisynaptically. We detected spontaneous activation of the presynaptic transporter, both by release of single transmitter vesicles from the same cell and by spillover after depolarization-evoked release from neighboring rod bipolar cells. From paired recordings of rod bipolar cells, we directly observed glutamate spillover with activation of the presynaptic transporter. This is consistent with accumulating evidence from several regions of the CNS, suggesting that glutamate escapes the synapse from which it is released, diffuses to neighboring synapses and activates transporters and receptors32,
39,
40. Specifically, there is evidence that diffusion of glutamate out of the synaptic cleft can activate glutamate transporters on nearby astrocytes in the hippocampus27 and Bergmann glial cells in the cerebellum37. It might be speculated that the observed spillover of glutamate between rod bipolar axon terminals is due to abolished activity of glial and/or neuronal glutamate transporters in the slice preparation. However, TBOA strongly reduced the amplitude of spontaneous EPSCs in AII amacrine cells and this can be counteracted by coapplication of cyclothiazide (unpublished data), suggesting that TBOA resulted in desensitization of the AMPA receptors on AII amacrine cells. This suggests that glutamate transporters are active in the slice preparation. In addition, recordings were performed from cells located below the surface layer, reducing, but not eliminating, the possibility that damage to superficial Müller glial cells (see below) explains the observed spillover. Spillover was also present at an elevated, physiological temperature, at which transporters are known to operate at higher rates31,
32. This cross-talk between neighboring synapses might be typical for synaptic microcircuits with little glial 'wrapping' of neuronal processes. Our results suggest that the microenvironment of the synaptic cleft between a rod bipolar cell and its postsynaptic partners is not compartmentalized or protected from the general extracellular environment in the inner plexiform layer9. In several areas of the CNS, similar spillover is limited by glial cells with glutamate transporters40. Whereas the high apparent affinity of the rod bipolar axon terminal glutamate transporter is likely to be important for the presynaptic response to spillover40, it is possible that spillover of glutamate can transiently desensitize, but not activate, the postsynaptic AMPA receptors on AII amacrine cells20,
41. It will be interesting to see whether similar spillover is observed if paired recordings of neighboring rod bipolar cells can be performed in an intact preparation.
Immunocytochemical investigations have detected the presence of all currently identified glutamate transporters (EAAT1–5) in the mammalian retina42,
43,
44,
45,
46, but only EAAT5 has been clearly detected in axon terminals of adult rod bipolar cells43. Whereas little is known about the functional properties of EAAT5 from heterologous expression studies, the functional properties revealed in our study are generally consistent with the presence of this transporter16. In retina, EAAT1 in Müller glial cells mediates the bulk of glutamate uptake30, and the potential contribution of the presynaptic rod bipolar glutamate transporter to transmitter clearance at the output synapses of rod bipolar cells is unknown.
The magnitude of the transporter-associated anion conductance activated by glutamate was sufficient to influence the membrane potential at the axon terminal of rod bipolar cells. In perforated-patch recordings with unperturbed ECl, Erev of the glutamate-evoked current was -64 mV, and glutamate evoked hyperpolarizations or depolarizations depending on the value of the membrane potential. Using dynamic-clamp electrophysiology, injection of realistic conductance waveforms at a membrane potential of -60 mV evoked IPSPs or EPSPs depending on Erev of the injected conductance. Furthermore, injection of a tonic conductance with Erev = Vm (= -60 mV) resulted in the shunting of EPSPs evoked by an excitatory conductance waveform. Notably, injection of an excitatory conductance into rod bipolar cells resulted in a larger EPSP when glutamate transporters were blocked compared to the control condition. When this EPSP was used as a voltage command in rod bipolar cells, the resulting EPSC in AII amacrine cells displayed a faster rise time compared to that in the control. Although the effect of blocking the glutamate transporter pharmacologically could be explained by a mechanism based on tonic activation of the presynaptic transporter, we used a paired-pulse stimulus to demonstrate that transient, stimulus-dependent activation of the transporter can influence the membrane potential of a rod bipolar cell. The inhibition exerted by activation of the glutamate transporter is one of three potential negative feedback mechanisms9 acting on the presynaptic axon terminals of rod bipolar cells. The other two involve GABAergic feedback from amacrine cells25,
29,
47 and suppression of voltage-gated Ca2+ current by protons released during exocytosis of synaptic vesicles48. Further work is required to determine the relative magnitude and functional importance of each of these mechanisms for synaptic output from rod bipolar cells, as well as the modulation of and potential interactions between these mechanisms.
Our results suggest that glutamate released from rod bipolar cell axon terminals acts locally on the terminal from which it is released, as well as on neighboring terminals by spillover, and exerts a negative feedback control on transmitter release by activating the transporter-associated nonstoichiometric anion conductance. The size of the diffusion domain49 around a rod bipolar axon terminal is unknown, but our evidence suggests that release from a single cell will influence that cell and its immediate neighbors. Similarly, the size of the corresponding confluence domains49—the number of cells that influence the extracellular glutamate concentration at a specific point—is also unknown. The transporter-associated anion conductance can influence the membrane potential of an axon terminal by both hyperpolarization and shunting inhibition. The evidence for glutamate transporters in other presynaptic terminals in the CNS suggests that similar mechanisms of regulating transmitter release could be of general importance.
Methods
Review of experimental design.
The procedure for killing the rats was approved by the Norwegian Animal Research Authority.
Electrophysiology.
Albino rats (4–7 weeks postnatal) were deeply anesthetized with halothane in oxygen and killed by cervical dislocation. A detailed account of the general methods, including preparation of retinal slices, has been previously published20. The extracellular perfusing solution (bubbled with 95% O2, 5% CO2) had the following composition: 125 mM NaCl, 25 mM NaHCO3, 2.5 mM KCl, 2.5 mM CaCl2, 1 mM MgCl2 and 10 mM glucose (pH 7.4). Except where otherwise stated, all experiments were done at room temperature (22–25 °C). A series of intracellular solutions were used; in the figures, these are referred to by a capital letter (A–H) and by their main constituent (CsCl in solutions A, B and D; CsCH3SO3 in C; KSCN in E; potassium gluconate in F and H; and KCl in G; details in the Supplementary Methods online). Lucifer yellow was added to all intracellular solutions (1 mg ml-1) and the pH was adjusted to 7.3 with CsOH or KOH as appropriate.
Drugs were either added directly to the extracellular solution or were locally applied by pressure from a patch pipette or a multibarrelled pipette complex. The concentrations of drugs were as follows (supplied by Tocris Bioscience, unless otherwise noted): 10 M bicuculline methchloride (to block GABAA receptors), 10 M CNQX or 6,7-dinitroquinoxaline-2,3-dione (DNQX) (to block non-NMDA-type glutamate receptors), 20–30 M CPP (to block NMDA-type glutamate receptors), 100 M glutamate, 1 M strychnine (Sigma; to block glycine receptors), 50 M TPMPA (to block GABAC receptors), 0.3 M tetrodotoxin (TTX) and 50 M TBOA. All AII amacrine cell recordings were done with TTX in the bath.
Ultrafast drug application from a theta-tube pipette was performed as previously described20. Drugs were dissolved in HEPES-buffered solution containing 145 mM NaCl, 2.5 mM KCl, 2.5 mM CaCl2, 1 mM MgCl2, 5 mM hemisodium-HEPES and 10 mM glucose; pH was adjusted to 7.4 with HCl. Under optimal conditions, the 20–80% rise time of the solution exchange ranged from 100 s to 220 s. Data from experiments with 20–80% rise times > 300 s were omitted.
Recordings were made with an EPC9-dual or an EPC10-triple amplifier (HEKA Elektronik). Cells and patches were generally held at a membrane potential of -60 mV. The digital sampling interval was 10–250 s and the signal was low-pass filtered with a corner frequency (–3 dB) that was one-fifth to one-third the inverse of the sampling interval. The average series resistance (Rs) was 17 1 M (n = 66) for rod bipolar cell body recordings, 106 5 M (n = 29) for rod bipolar axon terminal recordings and 29 2 M (n = 16) for AII amacrine cell recordings. The Rs for rod bipolar axon terminal recordings is likely to have been overestimated owing to incomplete resolution of the very rapid capacitive current decay (L. Oltedal, S.H.M., M.L.V. and E.H., unpublished data). The average Rs was 99 6 M (n = 18) for perforated-patch recordings. Conductance injection experiments were done as detailed in Supplementary Methods.
Data analysis.
Data were analyzed with FitMaster, PulseFit and PulseTools (HEKA Elektronik), Igor Pro (WaveMetrics) and AxoGraph (Molecular Devices). Spontaneous, transient currents were detected with a threshold of -3 pA (MiniAnalysis; Synaptosoft) and verified by eye. The number of events analyzed for each axon terminal recording ranged from 43 to 169. The decay time course of averaged responses was estimated by curve fitting with single- or double-exponential functions. For measurement of Erev values, data points of I-V relationships were typically fitted by fifth- to seventh-order polynomial functions. Concentration-response data were fitted with a Hill-type equation. Each patch was tested with two to four different concentrations of glutamate. Responses were normalized to the response to 1 mM glutamate. To control for rundown, measurements of responses to different concentrations of glutamate were interleaved with measurements of responses to 1 mM glutamate. Patches with more than 30% reduction in the response to 1 mM glutamate were excluded from the analysis. Onset of a response was measured as the time from start of the stimulus to 5% of the peak response amplitude.
Data are presented as mean s.e.m. (n = number of cells, cell pairs or patches) and percentages are presented as percentage of control. Statistical analyses were performed using Student's two-tailed t-test (unpaired, unless otherwise stated) and differences were considered statistically significant at the P < 0.05 level. For illustration purposes, capacitive transients were truncated and most records of raw data were low-pass filtered (digital nonlagging Gaussian filter; -3 dB at 0.5–2 kHz). Unless otherwise noted, traces in figures represent individual traces.
Note: Supplementary information is available on the Nature Neuroscience website.
Received 28 June 2006; Accepted 26 September 2006; Published online: 15 October 2006.
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Acknowledgments
Financial support from the Norwegian Research Council (NFR 155397/310 and 161217/V40), the Meltzer fund (University of Bergen) and the Faculty of Medicine at the University of Bergen (fellowships for M.L.V. and S.H.M.) is gratefully acknowledged.
Competing interests statement:
The authors declare that they have no competing financial interests. |
