Synaptic Activation of Presynaptic Glutamate Transporter Currents in Nerve Terminals

Exocytosis is followed by an inward current in bipolar cell terminals

Whole-cell patch-clamp recordings were made from visually identified bipolar cell terminals in slices of goldfish retina (Fig. 1a,b). Because glutamate transporter currents have been identified in the dendrites of teleost bipolar cells (Grant and Dowling, 1995), somatic and dendritic currents were eliminated by recording only from terminals that had been isolated by severing of the axon during the slicing procedure. Isolated terminals also have the advantage of a simple electrotonic structure (spherical) (Mennerick et al., 1997), which reduces artifacts in time-resolved membrane capacitance measurements and enables fast voltage-clamping (Gillis, 1995). Isolated terminals were identified from the current response to a -10 mV step (Fig. 1c,d) (see Materials and Methods). Previously, isolated bipolar cell terminals dissociated acutely from goldfish retina have been used to study neuro-transmitter release by measurement of depolarization-evoked changes in membrane capacitance (von Gersdorff and Matthews, 1994). They exhibit pronounced paired-pulse depression of exocytosis (Mennerick and Matthews, 1996; Burrone and Lagnado, 2000) and glutamate release (von Gersdorff et al., 1998), which is not a result of a depression of Ca²⁺ influx. We therefore used differences in the membrane currents evoked by the first and second stimuli of a paired-pulse protocol to identify currents associated with glutamate release from bipolar cell terminals.

Experiments were performed with NO₃^- as the major intracellular anion to augment currents through transporter-associated anion channels (Wadiche et al., 1995). The extracellular solution contained picrotoxin (50 μm) and strychnine (1 μm) to block inhibitory currents evoked by GABA and glycine released from amacrine cells. From a holding potential (V_h) of -60 mV, two 1 msec depolarizations to 0 mV were delivered with an interpulse interval of 100 msec. The first depolarization evoked a Ca²⁺ current (I_Ca) with mean peak amplitude of -195 ± 30 pA (n = 10) (Fig. 2a₁) and an increase in membrane capacitance (ΔCm) of 15 ± 3 fF (Fig. 2a₂). This corresponds to the fusion of ∼600 vesicles over 55 ribbon-type active zones, or ∼11 vesicles per active zone (von Gersdorff et al., 1996). The mean amplitude of I_Ca evoked by the second depolarization was similar to that evoked by the first pulse (103 ± 1%, Fig. 2a₁), but the second pulse ΔCm was only 9 ± 3% of the first pulse ΔCm (Fig. 2a₂). After repolarization to V_h, I_Ca deactivated almost immediately (the tail current decays with a time constant of 90 μsec at -60 mV) (Mennerick and Matthews, 1998). However, a large inward current (I_trans) was observed after the first depolarization (Fig. 2a₃). I_trans peaked 2.8 ± 0.2 msec after the onset of the depolarizing pulse, had an average peak amplitude of -54.9 ± 7.3 pA, and decayed monoexponentially with a mean time constant of 8.5 ± 0.6 msec (n = 10). After the second depolarization, the maximum amplitude of I_trans was 19 ± 1% of that measured after the first pulse (Fig. 2a₃). Addition of ionotropic glutamate receptor antagonists (10 μm NBQX, 100 μm dl-AP-5) did not inhibit I_trans (n = 12, data not shown).

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Figure 2.

Depolarization-evoked exocytosis is associated with activation of I_trans in bipolar cell terminals but not in the calyx of Held. a, Bipolar cell terminal response to a pair of 1 msec depolarizations from -60 to 0 mV with CsNO₃ intracellular solution. a₁, Membrane current evoked by a 1 kHz voltage sine wave (used to measure membrane capacitance), with depolarization-evoked I_Ca during intervals. a₂, Membrane capacitance (C_m), showing an increase after the first depolarization but no additional increase after the second depolarization. Baseline C_m for this terminal was 4.3 pF. a₃, Currents evoked by the first and second depolarizations superimposed on an expanded time scale (first, black; second, gray). Note I_trans after the first but not the second depolarization. I_trans decayed with a time constant of 9.6 msec. b, P9 rat calyx of Held response to a pair of 1 msec depolarizations from -80 to 0 mV with CsNO₃ intracellular solution. I_Ca and ΔC_m were much larger than for bipolar cell terminals, but I_trans was not observed. Baseline C_m for this calyx was 21.2 pF.

Changes in membrane capacitance were also used to measure depolarization-evoked release at the large terminals of the calyx of Held (Sun and Wu, 2001). Whole-cell recordings were made from visually identified calyces in slices of brainstem from P5 to P14 rats. With CsNO₃ intracellular solution, two 1 msec depolarizations (100 msec interval) from -80 to 0 mV were delivered. I_Ca evoked by the first depolarization had a mean amplitude of -1.4 ± 0.1 nA, and ΔCm was 78 ± 18 fF (n = 6), corresponding to the release of 600–900 synaptic vesicles (Taschenberger et al., 2002) from ∼600 conventional active zones (Sätzler et al., 2002). I_Ca evoked by the second depolarization was 92 ± 3% of the first pulse I_Ca, whereas second pulse ΔCm was only 23 ± 9% of the first pulse ΔCm (Fig. 2b₁,b₂). In contrast to bipolar cell terminals, there was no difference in the tail currents measured after the first and second depolarizations (n = 6, Fig. 2b₃).

During the course of whole-cell recordings from bipolar cell terminals, the amount of release per stimulus gradually runs down. If I_trans is activated by released transmitter, as suggested by the correlation between I_trans and ΔCm with paired pulses, a similar decrease in I_trans amplitude would be expected. Delivery of 1 msec depolarizations at 15 sec intervals for 5 min was associated with a gradual decline in ΔCm to 17 ± 4% of the initial value (n = 3) (Fig. 3a₂,b), whereas I_Ca was fairly constant during this period (94 ± 2%) (Fig. 3b). However, the peak amplitude of I_trans was found to decrease to the same extent as ΔCm (20 ± 3%), with a very similar time course (n = 3) (Fig. 3a₁,b). I_trans is therefore likely to be activated by released glutamate. The rate of decay of I_trans was independent of its amplitude, being 8.1 ± 0.6 msec for the first three stimuli and 7.7 ± 0.3 msec for the 13th to 15th stimuli, when the peak amplitude of I_trans had decreased to 22 ± 6% of its initial value (n = 3). However, the time to peak became shorter as the amplitude of I_trans decreased such that the current appeared to peak before the end of I_Ca (Fig. 3a₁).

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Figure 3.

Correlated run-down of ΔCm and I_trans amplitude in bipolar cell terminals. a₁, Currents and a₂, ΔCm evoked by 1 msec depolarizations from early (i) and late (ii) stages of a recording. I_trans and ΔCm decreased during the recording. Baseline Cm for this terminal was 3.9 pF. b, I_trans peak amplitude (measured 1.5–2.5 msec after depolarization), I_Ca amplitude, and ΔCm normalized to the first response and plotted against stimulus number. Stimulus interval was 15 sec. Mean ± SEM for three terminals.

Exogenous glutamate activates an inward current in bipolar cell terminals

The expression of a glutamate-activated current in bipolar cell terminals was investigated further by puff-application of l-glutamate (10 mm) to isolated terminals in the presence of a mixture of bath and puff-applied ionotropic glutamate, GABA, and glycine receptor antagonists (10 μm NBQX, 100 μm dl-AP-5, 50 μm picrotoxin, 1 μm strychnine) plus 1 mm CdCl₂ to block Ca²⁺-dependent synaptic transmission. In four experiments, the metabotropic glutamate receptor (mGluR) antagonists MCPG (500 μm; blocker of group I and II mGluRs) and CPPG (200 μm; blocker of group II and III mGluRs) were also included in the extracellular solution. In the presence of these antagonists and with CsNO₃ intracellular solution, l-glutamate evoked a large, slowly decaying inward current (I_glu) with a mean peak amplitude of -241 ± 69 pA (V_h = -60 mV; n = 6) (Fig. 4a). The current-to-voltage relationship of I_glu was investigated in two terminals by varying V_h between -90 and +90 mV at the peak of the glutamate response and subtracting the current induced by the same voltage protocol before glutamate application. I_glu exhibited an inwardly rectifying I–V curve (Fig. 4b). This relationship is similar to that reported for glutamate transporter currents evoked in photoreceptors and retinal glial cells (Sarantis et al., 1988; Tachibana and Kaneko, 1988; Schwartz and Tachibana, 1990; Barbour et al., 1991). The sensitivity of I_glu to glutamate transporter inhibitors was therefore examined. In the presence of the specific, nontransportable blocker TBOA (50 μm) (Shimamoto et al., 1998; Shigeri et al., 2001), I_glu was significantly inhibited (-17 ± 5 pA; n = 6; p < 0.05) (Fig. 4c). However, the nontransportable GLT1/EAAT2 subtype-specific antagonist DHK (200 μm) did not inhibit I_glu (-348 ± 101 pA; n = 4).

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Figure 4.

Activation ofI_glu by exogenous glutamate in bipolar cell terminals but not the calyx of Held. a, Puff application of l-glutamate (10 mm) to a bipolar cell terminal at -60 mV with CsNO₃ intracellular solution. Control response in the presence of NBQX (10 μm), dl-AP-5 (100 μm), picrotoxin (50 μm), strychnine (1 μm), and CdCl₂ (1 mm). b, The I–V relationship of I_glu in one terminal. A protocol comprising brief incremental voltage steps from -90 mV was delivered both before glutamate application and at the peak of the response, and the I–V curve obtained from the difference in the evoked currents at each potential. This recording was made with the addition of the mGluR antagonists MCPG (500 μm) and CPPG (200 μm) and the GLT1 antagonist DHK (200 μm) to the extracellular solution. c, A glutamate response in the presence of the broad-spectrum glutamate transporter inhibitor TBOA (50 μm) and the average peak I_glu amplitudes for control (n = 6) and TBOA (50 μm; n = 6) recordings. Only the first response from each terminal used as substantial rundown was observed for subsequent responses (^*p < 0.05 compared with control). d, Electrophoretic application of l-glutamate (0.5 m) to the calyx of Held (P11) at -80 mV with KNO₃ intracellular solution. Left, Presynaptic action potential with “postpotential” marked by arrowhead, presumably arising from capacitively coupled activity in the postsynaptic neuron. Similarly, postsynaptic spikes evoked by l-glutamate application were observed in the presynaptic recording (middle). Postsynaptic activity was blocked by NBQX (5 μm), leaving no presynaptic l-glutamate-evoked current (right).

For comparison, l-glutamate (0.5 m) was applied electrophoretically to the calyx of Held (KNO₃ intracellular solution; V_h =-80 mV). Recordings were confirmed as presynaptic by the action potential waveform evoked by afferent fiber stimulation (Borst et al., 1995) (Fig. 4d). In the presence of NBQX (5 μm), an antagonist of the low-affinity AMPA-type glutamate receptor, no current was evoked by l-glutamate (Fig. 4d). In the absence of NBQX, high-frequency capacitively coupled spikes originating from action potential firing in the postsynaptic cell were recorded, confirming that l-glutamate was delivered to the synapse (n = 12) (Fig. 4d). Interestingly, using the same electrophoretic procedure in the presence of NBQX (5 μm) and d-AP-5 (50 μm), prominent anion currents were observed in surrounding glial cells (H. Taschenberger and H. von Gersdorff, unpublished observations).

I_trans has the pharmacological profile of a transporter current

Because the glutamate-evoked current in bipolar cell terminals was significantly inhibited by the glutamate transporter antagonist TBOA, the sensitivity of I_trans to transporter antagonists was investigated. TBOA (50 μm) or DHK (500 μm) was bath-applied in the presence of ionotropic glutamate receptor antagonists (10 μm NBQX, 100 μm dl-AP-5). I_trans was evoked by a pair of 1 msec depolarizations to 0 mV. In the presence of TBOA, I_trans was inhibited significantly (Fig. 5a). Current amplitude measured 2–3 msec after the depolarization was -3.8 ± 0.2 pA (n = 6), compared with a peak I_trans amplitude of -54.9 ± 7.3 pA under control conditions (n = 10; p < 0.05). The amount of glutamate release in the presence of TBOA, as measured by ΔCm, was not significantly different from control experiments (control, 15.3 ± 3.2 fF, n = 10; TBOA, 17.6 ± 3.9 fF, n = 6) (Fig. 5a). The block by TBOA of the current evoked by a 1 msec depolarization suggests that it is mediated solely by glutamate transporters, with no significant contribution from Ca²⁺-activated conductances. In contrast to TBOA, DHK (500 μm) had no inhibitory effect on either the amplitude of I_trans (-54.5 ± 13.8 pA; n = 7) (Fig. 5a) or the rate of decay (7.2 ± 0.9 msec; n = 7). The peak amplitude of I_trans was also normalized to baseline Cm to control for variability in terminal size, and no significant difference was found between DHK and control experiments (control, -14.6 ± 1.9 pA/pF, n = 10; DHK, -16.9 ± 4.7 pA/pF, n = 6) (Fig. 5b). All subsequent I_trans amplitudes will be expressed as current density.

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Figure 5.

Sensitivity of I_trans in bipolar cell terminals to glutamate transporter inhibitors. a, Superimposition of a pair of currents (first, black; second, gray) evoked by 1 msec depolarizations to 0 mV (100 msec interval) in the presence of TBOA (50 μm) or DHK (500 μm). I_trans was observed with DHK but not with TBOA. Bottom, ΔCm evoked by the first and second depolarizations, delivered during intervals in Cm trace, for these responses. b, Mean peak I_trans amplitude normalized to terminal size for control (n = 10), TBOA (50 μm; n = 6), and DHK (500 μm; n = 7) responses. ^*p < 0.05 compared with control. c, Superimposition of the membrane currents evoked by pairs of 25 msec depolarizations to -30 mV (100 msec interval; first, black; second, gray). I_trans was observed during the first depolarization under control conditions and in the presence of DHK (500 μm) but not in TBOA (50 μm) or THA (200 μm).

Because bipolar cells generally respond to stimulation with graded and sustained membrane potential changes, the activation of I_trans was investigated during longer depolarizations to a more physiologically relevant membrane potential. Release was evoked by a pair of 25 msec pulses (100 msec interval) from -60 to -30 mV. Paired-pulse depression of release for 25 msec depolarizations was difficult to measure with CsNO₃ intracellular solution because of the slow decay of a Ca²⁺-dependent anion conductance after the pulse. However, with Cs-gluconate intracellular solution, ΔCm evoked by the second pulse was on average only 13 ± 4% of that evoked by the first pulse (n = 7, data not shown). Under control conditions, I_trans was observed in addition to I_Ca during the first but not the second depolarization (n = 8) (Fig. 5c). The additional first pulse inward current peaked within 5 msec of the start of the depolarization and decayed with a mean time constant of 8.7 msec during the remainder of the pulse. Interestingly, it was found from ΔCm measurements that the rate of release during a 25 msec pulse decreases rapidly after the first millisecond, such that ∼87% of the total release occurs within the first 5 msec. I_trans was evoked in the presence of DHK (500 μm; n = 4) (Fig. 5c) but was not observed in the presence of TBOA (50 μm; n = 6) (Fig. 5c) or the substrate glutamate transporter inhibitor THA (200–300 μm; n = 7) (Fig. 5c).

Homologs of EAAT2, 3, and 5 have been localized to retinal bipolar cells by staining with specific antibodies (Rauen and Kanner, 1994; Schultz and Stell, 1996; Eliasof et al., 1998; Reye et al., 2002). Of these, GLT1/EAAT2 (Rauen et al., 1996; Vandenbranden et al., 2000) and EAAT5 (Pow and Barnett, 2000) have been localized specifically to bipolar cell terminals. Given the insensitivity of I_trans to GLT1-specific DHK, the synaptically activated transporter current in bipolar cell terminals is likely to be mediated by a homolog of EAAT5.

I_trans is an anion current

Transporter currents can be produced by both electrogenic uptake and an associated anion conductance. The anion conductance is more permeable to NO₃^- than Cl^- and is impermeable to gluconate^- (Wadiche et al., 1995). Intracellular CsNO₃ was therefore replaced with Cs-gluconate to isolate electrogenic current. The ΔCm evoked by a 1 msec depolarization with Csgluconate intracellular solution (13 ± 2 fF; n = 10) was comparable with that with CsNO₃ solution, but I_trans was never observed (n = 10) (Fig. 6a), indicating a requirement for permeant anions. Consistent with this, I_trans was evoked by a 1 msec depolarization with CsCl intracellular solution (Fig. 6a). The current had a peak amplitude of -7.7 ± 0.8 pA/pF (n = 4), smaller than CsNO₃ recordings (-14.6 ± 1.9 pA/pF), but the rate of decay of I_trans was similar (7.6 ± 1.2 msec; n = 4) (Fig. 6b).

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Figure 6.

Ion selectivity of I_trans. a, Currents evoked by a pair of 1 msec depolarizations to 0 mV, superimposed (first, black; second, gray) with (bottom) ΔCm evoked by the first and second depolarizations, delivered during intervals in Cm trace. I_trans was not observed with Cs gluconate but was evoked with either KCl or CsCl intracellular solution. Also shown is a recording with CsNO₃ intracellular solution, which contained 10 mm l-glutamate. b, Comparison of mean I_trans amplitude and τ_decay with KCl (n = 4), CsCl (n = 4), CsNO₃ (n = 10), or CsNO₃ plus l-glutamate (n = 4) intracellular solution. ^*p < 0.05.

Glutamate uptake is coupled to the countertransport of intracellular K⁺ (Zerangue and Kavanaugh, 1996; Levy et al., 1998), which was normally replaced in our recordings by Cs⁺. It was found previously that glutamate-evoked anion currents in cells expressing the GLT1 subtype of glutamate transporter are significantly reduced by substitution of K⁺ with Cs⁺ (Bergles et al., 2002), and intracellular Cs⁺ was shown to decrease transporter currents in isolated Müller cells by one-third (Barbour et al., 1991). Recordings of I_trans in bipolar cell terminals were therefore also made with a KCl-based intracellular solution (Fig. 6a). The peak amplitude of I_trans evoked by a 1 msec depolarization was -6.9 ± 1.2 pA/pF (n = 4) (Fig. 6b), not significantly different from CsCl recordings. The rate of decay of I_trans was also unaffected by K⁺ (8.8 ± 1.0 msec; n = 49) (Fig. 6b).

Under physiological conditions, glutamate transporters are working against a high cytoplasmic glutamate concentration, which will be lost during whole-cell recording. To determine whether this affected the size or kinetics of I_trans, experiments were also performed with a CsNO₃-based intracellular solution that contained 10 mm l-glutamate. This did not change significantly either the peak amplitude (-18.6 ± 5.2 pA/pF; n = 4) or rate of decay (7.1 ± 0.5 msec; n = 4) of I_trans evoked by a 1 msec depolarization (Fig. 6a,b).

I_trans is augmented in low extracellular pH buffer

After exocytosis, the synaptic cleft is transiently acidified because of the low pH of synaptic vesicles (Krishtal et al., 1987; Miesenböck et al., 1998). This may affect transporter function, because protons are cotransported during glutamate uptake (Zerangue and Kavanaugh, 1996). The magnitude of synaptic cleft acidification can be modulated by exogenous pH buffers (DeVries, 2001); therefore, the effect of extracellular HEPES on the transporter current was investigated. The concentration of HEPES in the extracellular solution was changed from 12 to 3 or 48 mm by substitution with NaCl, and the pH was set to 7.45 with NaOH solution. The final Na⁺ concentration ranged from 105 mm (with 48 mm HEPES) to 130 mm (with 3 mm HEPES). This difference in Na⁺ concentration should not greatly affect the current, because both are near saturation for the effect of Na⁺ on transporter function. We confirmed this by using a recent model of glutamate transporter function (Bergles et al., 2002), which showed that this difference in Na⁺ concentration affected the transporter current by a maximum of 4–9% (using 100 μm to 100 mm l-glutamate pulses; C. E. Jahr, personal communication). I_trans was evoked by a pair of 1 msec depolarizations to 0 mV with CsNO₃ intracellular solution. When the extracellular HEPES concentration was increased from 12 to 48 mm, neither the amplitude nor the decay of I_trans was altered significantly (-17.3 ± 1.2 pA/pF; 7.1 ± 0.4 msec; n = 3) (Fig. 7a,b). However, lowering the HEPES concentration from 12 to 3 mm caused a 230% increase in the amplitude of I_trans to -48.2 ± 8.0 pA/pF (n = 5; p < 0.05) (Fig. 7a,b), with no change in the rate of decay (8.5 ± 0.6 msec) (Fig. 7b). There was no significant difference in I_Ca or ΔCm evoked by 1 msec depolarizations between recordings with 3 and 48 mm extracellular HEPES (Fig. 7a), indicating that the increase in I_trans was not a result of increased glutamate release.

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Figure 7.

The effect of extracellular HEPES concentration on I_trans. a, Currents evoked by a pair of 1 msec depolarizations to 0 mV, superimposed (first, black; second, gray) in the presence of 3 or 48 mm extracellular HEPES. I_trans was greatly potentiated in low HEPES. Bottom, ΔCm evoked by the first and second depolarizations, delivered during intervals in Cm trace. b, Comparison of mean I_trans amplitude and τ_decay with 3 (n = 5), 12 (n = 10), and 48 (n = 3) mm extracellular HEPES. ^*p < 0.05.

I_trans has a large single-channel current

The permeability of transporter-associated anion channels is known to differ between transporter subtypes, with EAAT4 and 5 displaying a much larger anion current than EAAT1 or 2 (Fairman et al., 1995; Arriza et al., 1997; Eliasof et al., 1998). The Cl^- conductance of the anion channel associated with EAAT1 was found to be at the subfemtosiemens level (Wadiche and Kavanaugh, 1998), whereas a value of 0.7 pS was obtained for glutamate transporters in photoreceptors (Picaud et al., 1995b; Larsson et al., 1996). To investigate the anion conductance of glutamate transporters in bipolar cell terminals, an estimate of single-channel current was obtained by noise analysis of the transporter currents. Peak-scaled nonstationary noise analysis was applied to the decay of I_trans using a method described previously for postsynaptic currents (Traynelis et al., 1993; Benke et al., 1998). I_trans was evoked by 1 msec depolarizations to 0 mV with CsNO₃ intracellular solution. Ca²⁺-activated conductances are unlikely to contribute significant noise to the decay of I_trans, because current variance after the pulse in the presence of TBOA (1.9 ± 0.3 pA²) was not significantly greater than baseline variance (1.4 ± 0.4 pA²; n = 6). For each terminal, a plot of mean current variance against amplitude gave estimates for single-channel current (i) and the average number of channels open at the peak of the current (N) (Fig. 8a). At a V_h of -60 mV, an average value of -1.1 ± 0.1 pA was obtained for i (n = 7, Fig. 8b). As a control, I_trans was recorded in a different set of terminals at -90 mV and yielded a value for i of -1.5 ± 0.2 pA (n = 8) (Fig. 8b), consistent with the increased driving force at this membrane potential. There was considerable variability in the peak amplitude of I_trans between recordings. For a constant V_h, larger macroscopic currents were associated with an increase in N but not with an increase in i (Fig. 8c). From the single-channel current values obtained at -60 and -90 mV, the slope conductance between these potentials is 13.3 pS.

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Figure 8.

Peak-scaled nonstationary noise analysis of the decay of I_trans evoked by 1 msec depolarizations to 0 mV and nonstationary noise analysis of the decay of I_glu. a, Variance (σ²) versus amplitude (I) plot for I_trans in one terminal and fit of the data by σ² = iI - I²/N + b₁ to yield estimates for i (single-channel current) and N (average number of channels open at peak of response). For this terminal, i was -1.4 pA and N was 45. b, Values of i obtained for individual terminals at a holding potential (V_h) of -60 or -90 mV (closed/open circles) and mean ± SEM (gray squares). c, Plot of i (left) and N (right) against mean peak amplitude of I_trans for each terminal (closed circles, -60 mV; open circles, -90 mV). d, Variance versus amplitude plot for I_glu in one terminal and fit of the data by σ² = iI - I²/N + b₁. For this response, i was -0.5 pA and N was 1486. e, Values of i obtained from I_glu for individual terminals under control conditions and in the presence of DHK (200 μm).

An estimate of i was also obtained by noise analysis of the currents evoked by application of exogenous glutamate (I_glu) to bipolar cell terminals. A current variance/amplitude plot for the decay of each response (Fig. 8d) yielded a mean value for i of -0.51 ± 0.02 pA at -60 mV (n = 5 terminals) (Fig. 8e). The lower single-channel current value obtained by noise analysis of I_glu could result from the activation of different populations of transporters by the two techniques. For example, application of exogenous glutamate could also activate GLT1-type transporters, which have been identified at a distance from synaptic ribbons in goldfish bipolar cell terminals (Vandenbranden et al., 2000) and have a much smaller associated anion conductance than EAAT5 (Eliasof et al., 1998). However, noise analysis of I_glu evoked in the presence of DHK (200 μm; GLT1 Km = 8–53 μm) (Levy et al., 1998; Chen et al., 2002) gave a single-channel current value of -0.44 ± 0.04 pA (n = 3 terminals) (Fig. 8e), which was not significantly different from control recordings. GLT1-like transporters are therefore unlikely to contribute greatly to I_glu.

Glutamate transporter currents are observed in bipolar cell terminals

Our results demonstrate the activation of a previously unreported presynaptic current by very brief depolarization of isolated bipolar cell terminals in retinal slices. Several lines of evidence identify this as a glutamate transporter-associated current. First, current amplitude correlated with the amount of glutamate released, as measured by ΔCm, during both paired-pulse depression and rundown of release. Second, the current was not observed with gluconate as the major intracellular anion and was larger with NO₃^- than Cl^-. It is unlikely to reflect activation of the Ca²⁺-dependent Cl^- current expressed in bipolar cell terminals (Okada et al., 1995), because the amplitude of this current correlates directly with Ca²⁺ influx (Burrone et al., 2002) and is therefore not depressed by the paired-pulse protocol. Third, it was inhibited by the glutamate transporter antagonists TBOA and THA. In addition, a TBOA-sensitive current was evoked in bipolar cell terminals by application of exogenous glutamate in the presence of antagonists of Ca²⁺ channels, glutamate, GABA and glycine ionotropic receptors, and mGluRs. The pharmacology and large single-channel anion current of the transporter suggest that it is the EAAT5 subtype, which has been found to be strongly immunolocalized to rat bipolar cell terminals (Pow and Barnett, 2000).

In photoreceptors, glutamate uptake currents have also been found to arise in the synaptic terminal (Sarantis et al., 1988; Eliasof and Werblin, 1993; Picaud et al., 1995a; Grant and Werblin, 1996). Both photoreceptors and bipolar cells release glutamate from ribbon-type synapses. The expression of presynaptic glutamate transporters may therefore be related to the high capacity and rates of release attainable at these synapses (Mennerick and Matthews, 1996; von Gersdorff et al., 1996). Glutamate uptake has been reported to modulate postsynaptic responses at both the photoreceptor-to-bipolar cell (Vandenbranden et al., 1996; Gaal et al., 1998) and bipolar cell-to-ganglion cell synapses (Higgs and Lukasiewicz, 1999; Matsui et al., 1999; Chen and Diamond, 2002). It is likely that at least part of this uptake occurs via presynaptic transporters.

Glutamate transporter currents are absent from the calyx of Held

Glutamate transporters contribute to shaping the time course of postsynaptic currents and limit the effects of glutamate spillover at various conventional active zone synapses (Barbour et al., 1994; Tong and Jahr, 1994; Takahashi et al., 1995; Otis et al., 1996; Overstreet et al., 1999; Diamond, 2001; Arnth-Jensen et al., 2002), including the calyx of Held (Taschenberger and von Gersdorff, 2001). The major neuronal glutamate transporter (EAAC1/EAAT3) has been localized to presynaptic terminals in some brain regions (Rothstein et al., 1994; Furuta et al., 1997a; He et al., 2000; Chen et al., 2002) but is absent from others (Conti et al., 1998). In the present study, no presynaptic transporter current was identified in the calyx of Held. Recordings were made from terminals during the maturation phase of the synapse (P5–P14). Because the expression of glutamate transporters is regulated developmentally (Furuta et al., 1997b), it remains possible that presynaptic transporters may be present in adult animals. Our results, together with an absence of evoked glutamate transporter current in postsynaptic MNTB neurons (Taschenberger and von Gersdorff, unpublished observations), suggest that glutamate transporters may be expressed exclusively in glial cells at the calyx of Held synapse.

Transporter currents in bipolar cell terminals have fast kinetics

The transporter current evoked by a 1 msec stimulation peaked 2.8 msec after the onset of the depolarizing pulse and decayed with a time constant of 8.5 msec. Postsynaptic transporter-associated anion currents evoked by synaptically released glutamate in cerebellar Purkinje cells were reported to have much slower kinetics (Otis et al., 1997; Auger and Attwell, 2000). This may result from transporter localization at a distance from release sites because brief glutamate application to excised membrane patches evoked currents with faster kinetics (decay time constant of 7.8 msec for 1 msec application) (Otis et al., 1997). It is unknown how the decay time of glutamate transporter currents in bipolar cell terminals in situ is affected by the rate of glutamate diffusion or the possible presence of postsynaptic transporters with fast kinetics. However, the relatively fast time course of the recorded current suggests that glutamate clearance from the vicinity of the transporters is fairly rapid. Interestingly, the rate of decay of I_trans was unaffected by GLT1 antagonists. It is also unclear how the time course of the anion current is related to the kinetics of glutamate uptake by a transporter. It has been suggested that the anion current is activated on glutamate binding and deactivates as the transporter moves into subsequent conformational states (Otis and Jahr, 1998). The anion and electrogenic currents were found to have a similar time course for EAAT2 (Otis and Kavanaugh, 2000), but the anion current exhibited slower kinetics than the electrogenic current for GLT1 (Bergles et al., 2002). Similarly, the transporter-associated anion current activated by synaptically released glutamate in Purkinje cells was reported to have a much slower time course than the electrogenic current (Auger and Attwell, 2000). In bipolar cell terminals, electrogenic uptake currents were not detected in the absence of permeant anions, and it was therefore difficult to determine the relationship between the recorded current and glutamate transport.

Transporter currents in bipolar cell terminals are potentiated by low pH

The amplitude of bipolar cell terminal transporter currents was increased significantly by reducing the concentration of extracellular pH buffer, which will lead to an increase in synaptic cleft acidification after exocytosis (DeVries, 2001). The effect of extracellular protons on glutamate transporter function is likely to be complex. Consistent with the cotransport of protons and glutamate, alkalization of the extracellular medium has been shown to reduce steady-state transporter currents evoked by glutamate (Zerangue and Kavanaugh, 1996; Watzke et al., 2000), although glutamate transport can also be inhibited by acidification (Billups and Attwell, 1996). Synaptically activated transporter currents in the cerebellum were unaffected by elevated pH, although steady-state currents were significantly depressed in the same cells (Auger and Attwell, 2000). The transporter-associated anion conductance may also be modulated directly by pH (Watzke et al., 2000), and there is evidence for an additional proton current associated with EAAT4, which is potentiated by arachidonic acid (Fairman et al., 1998; Tzingounis et al., 1998). The exact role of protons in transporter function therefore remains uncertain and additional work is necessary to determine how bipolar cell transporter currents are potentiated by low pH.

Possible function of glutamate transporters in bipolar cell terminals

Assuming a single-channel current of -1.1 pA and a 1:1 transporter-to-anion channel ratio, we estimate that the glutamate released by a 1 msec depolarization activates, on average, ∼50 transporters (-55/-1.1 pA). This is a surprisingly small number, considering the associated ΔCm (15 fF), which indicates the release of ∼600 vesicles. Given the large number of active zones per terminal (∼55) (von Gersdorff et al., 1996), it suggests that transporters are not expressed at high density in the immediate vicinity of release sites. It also suggests a role for presynaptic transporters with high anion permeability in rapid signaling mechanisms. Bipolar cell terminals have a low endogenous Cl^- concentration (∼23 mm), resulting in outward current through Cl^- channels at physiological membrane potentials (Billups and Attwell, 2002). Assuming a Cl^- reversal potential of -60 mV and a single-channel conductance of 9 pS (estimated from the amplitude of I_trans with CsCl or KCl intracellular solution), an outward current of ∼14 pA would result from the activation of 50 transporters at -30 mV. This would equate to a membrane hyperpolarization of 7 mV with 0.5 GΩ input resistance (Protti et al., 2000). This is likely to be an underestimate, because physiological pH buffering conditions are more similar to 3 mm than 12 mm extracellular HEPES (M. J. Palmer and H. von Gersdorff, unpublished observations). Future studies will aim to determine the function of this Cl^- current in bipolar cell terminals. In photoreceptors, it was found that activation of a transporter-associated Cl^- current by exogenous glutamate reduced K⁺-evoked depolarizations (Picaud et al., 1995a), and in bipolar cell dendrites, a similar glutamate-activated Cl^- current has been suggested to contribute to light responses (Grant and Dowling, 1995).

Finally, we note that although the number of transporters activated by a brief depolarization seems to be small, these results do not preclude the existence of a large population of extrasynaptic glutamate transporters. From the amplitude of the largest glutamate-evoked transporter current (-551 pA) and the single-channel current obtained from this terminal (-0.48 pA), >1000 EAAT5-type transporters may be present on bipolar cell terminals. Thus, in addition to being located strategically close to release sites for putative signaling purposes, transporters located extrasynaptically may serve a more traditional role in glutamate uptake. Presynaptic transporters may be necessary for glutamate clearance at this synapse, because Müller cells do not contact bipolar cell terminals directly (Pow and Barnett, 1999). We conclude that glutamate transporters in bipolar cell terminals may function to signal a fast hyperpolarization after phasic release and contribute to the reuptake process after tonic release from this ribbon-type synapse.