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NEUROSCIENCE |
1 Laboratory of Cellular and Molecular Physiopathology of the Retina, National Institute for Health and Medical Research (INSERM Unité 592), Université Pierre et Marie Curie-Paris6, Paris, France
2 IGBMC, 1 rue Laurent Fries BP10142, 67404 Illkirch cedex, France
3 Fondation Ophtalmologique A. de Rothschild, Paris, France
4 Centre National d'Ophtalmologie des quinze-vingts, Paris, France
5 School of Biomedical Sciences, University of Newcastle, New South Wales, Australia
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Abstract |
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(Received 29 July 2006;
accepted after revision 7 September 2006;
first published online 14 September 2006)
Corresponding author M. J. Roux : IGBMC-ICS, 1 rue Laurent Fries BP10142, 67404 Illkirch cedex, France. Email: mjroux{at}igbmc.u-strasbg.fr
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Introduction |
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To determine whether glutamate transporters may play similar function in the mammalian retina, EAAT5 was located in mouse retinal sections and responses to glutamate were recorded in mouse bipolar cells. EAAT5 was expressed in the synaptic terminals of photoreceptors (cones and rods) and rod bipolar cells. A glutamate transporter current reversing at ECl was recorded from rod bipolar cells, with a major contribution of the axon terminals. It could be elicited by triggering glutamate release through depolarization of the recorded cell, suggesting that EAAT5 acts as a presynaptic receptor in mouse rod bipolar cells. The charge carried by this current was 1.5 time larger than the one carried by the reciprocal inhibition received from amacrine cells, indicating that the feedback mediated by presynaptic EAAT5 plays a major role in controlling the output from rod bipolar cells.
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Methods |
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Mice strains used in this study were either of the C57BL/6J or Balb/c ByJ or crosses between them, purchased (pure strains) from Charles River (Lyon, France) or bred (crosses) at the Mouse Clinical Institute animal house (Illkirch, France). Procedures involving animals and their care were conducted in agreement with the French Ministry of Agriculture and the European Community Council Directive no. 86/609/EEC, OJL 358, 18 December 1986. Adult (9–19 weeks) mice were killed by cervical dislocation. The eyes were enucleated and immediately put in ice-cold bicarbonate-buffered saline (BBS), composed of (mM): NaCl 126, KCl 2.5, CaCl2 2.4, MgCl2 1.2, NaH2PO4 1.2, NaHCO3 18, glucose 11 previously bubbled with 95% O2–5% CO2. The cornea, lens and vitreous humor were removed. The retina was detached from the pigmented epithelium and embedded in agarose (1.5%) prepared in phosphate-buffered saline (PBS) (0.1 M; pH 7.4) kept at 42°C. After agarose solidification on ice, the retina was cut in 150 or 200 µm thick slices using a Leica VT1000S vibratome (Leica, Wetzlar, Germany). The slices were kept at room temperature in bubbled BBS for at least half an hour before recordings.
Bipolar cell recordings
Slices were observed under infrared differential interference contrast (DIC) using a x63 objective and a Hamamatsu C8484 camera on a Leica DMLFS microscope. The preparation was continuously perfused at 
2 ml min–1 with bubbled BBS. Pipettes (6–8 M
) were pulled from GC150TF borosilicate glass capillaries (Harvard Apparatus) on a horizontal puller (DMZ Universal Puller, Zeitz Instrumente, Munich, Germany). Four different intracellular solutions were used. Two contained 10 mM EGTA-Na4, and either (mM) KCl 42, K-gluconate 98, MgCl2 1, EGTA-Na4 0.5, Hepes 5, ATP-Na2 5 (ECl
=
–28.9 mV, junction potential of 12.4 mV), or KCl 138, MgCl2 3, CaCl2 1, Hepes 10, ATP-Na2 3, GTP-Na3 0.5 (ECl
= 1.9 mV, junction potential of 3.9 mV). These solutions will be referred to as ECl
=
–29 mV and ECl
= 2 mV, respectively. The ECl
= 2 mV solution was used for most of the experiments presented here. Another solution containing 0.5 mM EGTA was otherwise identical to the ECl
= 2 mV solution. Finally, a Cs-based solution with very low EGTA (0.1 mM) was used to monitor reciprocal inhibition from amacrine cells. It contained (mM): CsCl 125, MgCl2 1, TEA-Cl 15, EGTA- Na4 0.1, glutamic acid 5, Hepes 10, ATP-Na2 3 and GTP-Na3 0.5. All solutions contained 10 µM of Alexa Fluor (488 or 594) hydrazide (Molecular Probes, Eugene, OR, USA), and pH was adjusted to 7.4 with NaOH or CsOH. Potentials were corrected post-recording for the calculated junction potential. For simplicity, potentials indicated in the text are rounded to integer values. All experiments were performed at room temperature (20–25°C). Data were acquired using a Multiclamp 700A amplifier, a Digidata 1322A interface and the pCLAMP9 software (Molecular Devices, Sunnyvale, CA, USA). Data were filtered prior to digitization at a frequency of 1/2 or 1/5th of the acquisition frequency, which was 200 Hz for puffed glutamate experiments and 10 kHz for depolarizing pulse experiments. Agonists and antagonists were applied either through bath application or locally with a puff pipette connected to a Picospritzer III (Parker Hannifin, Fairfield, NJ, USA). DL-Threo-ß-benzyloxyaspartate (DL-tBOA), 1,2,5,6-tetrahydropyridin-4-yl)methylphosphinic acid (TPMPA) and 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX) were obtained from Tocris (Ellisville, MI, USA); all others chemicals were obtained from Sigma-Aldrich (Lyon, France).
All values are indicated as means ± S.E.M.
Immunohistochemistry
Retinas from C57BL6/J mice were obtained from animals (8-weeks old) that had been killed by an overdose of sodium pentobarbital (200 mg kg–1, I.P.). Retinas were fixed by immersion with 4% paraformaldehyde in 0.1 M phosphate buffer, pH 7.4 for a duration varying between 5 min and 2 h. Vibratome sections (40 µm thick) were then cut and labelled using standard protocols, with a EAAT5 rabbit antiserum (dilution 1/5000) previously characterized (Pow & Barnett, 2000; Pow et al. 2000). Labelling was detected using biotinylated secondary antibodies, streptavidin–horseradish peroxidase complex, and revealed using diaminobenzidine as a chromogen. Immunolabelled sections were viewed using a Zeiss Axioskop.
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Results |
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In the mouse retina, EAAT5 immunolabelling was strongly dependent on the duration of fixation. For fixation times (between 5 and 20 min), cells in the inner nuclear layer with the characteristic shape of rod bipolar cells (RBCs) were labelled (Fig. 1A and B) from dendrites to axon terminals, with a stronger signal in the axon terminals (Fig. 1B). Double immunolabelling with the RBC specific marker PKC
confirmed the expression of EAAT5 in RBC (see accompanying online supplemental material, Supplemental Fig. 1). A diffuse staining was observed in the outer plexiform layer (OPL), which could correspond to RBC dendrites or photoreceptor terminals. For longer fixation times (2 h), the RBC labelling was strongly reduced, limited to the axon terminals (Fig. 1C). In contrast, the OPL staining was much stronger and could be attributed to photoreceptor terminals (Fig. 1C). The labelling pattern included small rounded terminals and larger, more heavily stained terminals in the inner part of the OPL, suggesting that both rod and cone terminals expressed EAAT5.
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To test if EAAT5 could carry a detectable current in the mouse retina, whole-cell patch-clamp recordings were obtained from RBCs in adult mouse retinal slices. Recorded cells were chosen in the two outer rows of cell bodies of the inner nuclear layer. Cell identification was established post-recording, based on the morphology revealed by the Alexa dye included in the intracellular solution, according to the classification of Ghosh et al. (2004): only cells with an axon ending in the inner part of the IPL and terminating in a small group of fat varicosities were considered for further analysis (Fig. 2A). Puff-application (100 ms) of glutamate (1 mM) in the vicinity of the axon terminal evoked a transient and slowly decaying inward current (IGlu) (Fig. 2Ba). IGlu could not be attributed to a mechanical artefact due to the puff itself, as applications of BBS + 2 mM sucrose did not evoke any current (n
= 6, data not shown). In addition, fast inward postsynaptic currents were often seen on top of IGlu (Fig. 2Ba), as expected from the known inputs from both GABAergic and glycinergic amacrine cells on RBC terminals. These synaptic events were fully blocked by a cocktail of ionotropic glycine and GABA (A and C) receptor inhibitors (1 µM strychnine, 100 µM bicuculline methiodide and 50 µM TPMPA), leaving the slow phase unaffected (Fig. 2Bb, n
= 5). They were also blocked by a cocktail of glutamate receptor inhibitors (50 µM CNQX, 50 µM
DL-2-amino-5-phosphonovaleric acid (DL-AP5), 500 µM (RS)--methyl-4-carboxyphenylglycine (MCPG)), which had no effect on the slow phase (data not shown, n
= 5). This dual pharmacology was expected for inhibitory postsynaptic currents (IPSCs) as the amacrine cells are depolarized by glutamate. The absence of effect on the slow phase excluded that IGlu could be due to the direct activation of glutamate ionotropic receptors, to the indirect activation of GABA or glycine receptors, or to the modulation of a conductance via activation of metabotropic glutamate receptors.
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The slow phase of the glutamate response was further characterized on RBCs for which no spontaneous or glutamate-evoked synaptic currents were recorded. This kind of glutamate response was frequently encountered when the axon terminals were near the surface of the slice. In these cells, the mean peak amplitude of IGlu was –28.8 ± 4.1 pA (n = 15) at a holding potential of –64 mV (intracellular solution ECl = 2 mV), with individual values ranged from –13.3 to –61.6 pA. The glutamate transporter agonist D-aspartate (1 mM) also induced an inward current (–26.2 ± 5.1; n = 9). As expected from the results obtained from cells receiving synaptic inputs shown in Fig. 2, IGlu was blocked at 91.5 ± 1.5% (n = 13) by bath application of 50 µM DL-tBOA, confirming that the slow phase was due to the activation of glutamate transporters (Fig. 3A and Supplemental Fig. 2a). Dihydrokainate (500 µM) did not have any effect (1.3 ± 6.0% of current diminution, n = 6), indicating that the EAAT2 isoform was not contributing to IGlu (Fig. 3A and Supplemental Fig. 2b). IGlu was unaffected by a bath-applied cocktail of blockers: 1 µM strychnine, 100 µM bicuculline methiodide, 50 µM TPMPA, 50 µM CNQX and 50 µM DL-AP5 (100.1 ± 2.1% of current remaining, n = 9) or 50 µM CNQX and 50 µM DL-AP5 + 500 µM MCPG (97.9 ± 5.4% of current remaining, n = 14) (Fig. 3A and Supplemental Fig. 2c).
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Exogenous glutamate evokes a transporter current from the rod bipolar cell soma/dendrites
A contribution of a transporter current in the response to the glutamate released by photoreceptors has been reported in the ON cone or mixed input bipolar cells in the teleost retina (Grant & Dowling, 1995, 1996). Looking for a comparable contribution in mouse rod bipolar cell, we applied glutamate at the level of the outer plexiform layer above the recorded cell. Puffs of glutamate (5 ms, 1 mM) evoked a small inward current peaking at –4.6 ± 0.5 pA (n = 9) at –64 mV, which was blocked by 75 µM DL-tBOA (79.0 ± 2.6% of current diminution, n = 6 data not shown). When progressively increasing the puff duration from 5 to 50 ms, a secondary peak of larger maximal amplitude developed (Supplemental Fig. 3a), suggesting that longer puffs activated both dendritic/somatic and axonal transporters, the larger component coming from the axonal compartment. This was supported by the comparable time course of currents evoked by 100 ms glutamate puffs applied either above the dendritic arbor or the axon terminals of a given RBC (Supplemental Fig. 3a), and by the fact that some potential RBCs, whose axons were cut during the slicing procedure, presented similar responses to glutamate (n = 3). Due to its small amplitude and the difficulty to isolate it from the larger axonal component, this dendritic and/or somatic EAAT5 current was not studied further.
Transient transporter current following depolarization
To examine whether glutamate transporters could act as presynaptic receptors, calcium-dependent vesicular release of glutamate was evoked from the recorded RBC by short depolarizations. Paired-pulse stimulations were used to obtain additional information on the release properties of these RBC terminals (pairs of 2 ms voltage jumps from –84 to –24 mV, 100 ms apart). A transient inward current (Itrans) followed the fast capacitive current upon repolarization (Figs 4, 5 and 6). In addition, as for responses to glutamate puffs, IPSCs were frequently seen (Fig. 4).
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Itrans ran down quickly over time and could rarely be detected after 3 min following establishment of the whole-cell configuration (Fig. 5C), in a similar fashion to what has been reported from the large terminals of goldfish bipolar cells (Palmer et al. 2003). The run-down time courses were identical for the first and second pulses (data not shown). Run-down of Itrans was not due to inactivation/internalization of the transporters, as IGlu could still be evoked with comparable amplitude after complete disappearance of Itrans (data not shown). In fact, it may correspond to a depletion of the vesicular pool: following an apparent complete run-down after trains of 2 ms paired-pulses at 5 s intervals, Itrans could frequently be re-observed after a few minutes without depolarization.
To circumvent this limitation, depolarizations were applied quickly after the passage in whole-cell, to maximize the recording time for Itrans. In doing so, Itrans could be recorded in a large majority of the tested RBC (93%, 331 out of 355). On the contrary, Itrans was not observed in any of the 25 cone bipolar cells patched in this study. All the pharmacological tests were performed using puff applications in the vicinity of the RBC axon terminals. While the exact concentration of the applied drug reaching the terminal could not be known, this method allowed for a fast wash of the antagonist compatible with the observed run-down of Itrans. Figure 5A shows traces of the current evoked by paired depolarizations before (black trace), at the end of (dark grey trace), and 1 min after application of 75 µM DL-tBOA (light grey trace). To determine the DL-tBOA-sensitive fraction of Itrans, the zero current was estimated as being reached after more than 3 min in the whole-cell configuration. Puff applications of 75 µM DL-tBOA (between 100 ms and 2 s, dark grey trace) blocked Itrans at 97.6 ± 1.23% for the first peak, and at 94.8 ± 1.4% for the second peak (n = 9) (see percentage of remaining current in Fig. 4C). Itrans quickly recovered its initial amplitude (light grey trace). When glutamate (1 mM) was applied locally with a puff pipette, Itrans was completely suppressed following the depolarization (Fig. 5D, dark grey trace). Although the peak amplitude of Itrans was always smaller (65.2 ± 4.8%, n = 9) than the steady-state current evoked by the application of glutamate, it could be as large as 88% of the amplitude of the glutamate-induced current. These observations suggested that Itrans was produced by the glutamate transporter mediating IGlu characterized above. In addition, Itrans was not affected by glutamate receptor blockers (97.3 ± 0.9% remaining current in the presence of 50 µM CNQX, 50 µM DL-AP5 and 500 µM MCPG, n = 5).
The DL-tBOA-sensitive current was isolated by subtraction for kinetic analysis (Fig. 5B). When the cell was held at –84 mV, a 2 ms pulse to –24 mV evoked a current of –32.7 ± 3.5 pA (n = 11). Itrans peaked 5.6 ± 0.1 ms (n = 11) after the start of the depolarization, and decayed with a time constant of 10.2 ± 0.4 ms (n = 11). A second pulse 100 ms later evoked a smaller current (–14.5 ± 1.4 pA, n = 11). The amplitude ratio of the second pulse compared to the first was on average 0.46 ± 0.02 (n = 11). However, the kinetic parameters of the second pulse were similar to those of the first one, as it peaked 6.6 ± 0.3 ms (n = 11) after the start of the depolarization and decayed with a time constant of 9.1 ± 0.6 ms (n = 11).
Itrans dependence on Ca2+ channel current
When the voltage of 2 ms depolarizing pulses was varied, current could be detected from –50 mV upward (Fig. 6A–C). The peak amplitude of the first and second pulses as a function of the depolarization voltage could be fitted to a Boltzmann distribution, with half-potentials of –42.0 ± 1.1 mV and –43.4 ± 1.1 mV, respectively (n = 7). This voltage dependence of Itrans, similar to that of L-type voltage-gated Ca2+ channels reported from mouse bipolar cells (Berntson et al. 2003), suggested that Itrans was due to glutamate release from the recorded cell. While the weaker depolarizations induced responses of comparable amplitudes for the first and second pulses, with weak paired-pulse facilitation below –44 mV, a marked paired-pulse depression was observed from –40 mV upward.
As additional evidence that Itrans was subsequent to synaptic release, we tested its dependence upon Ca2+ currents. Puff application (10 s) of 100 µM Cd2+ next to the axon terminal strongly reduced Itrans (Fig. 6D, n = 4). In paired pulse experiments as described above, the first peak was reduced by 86.5 ± 2.3%, the second by 80.0 ± 3.9% (n = 4). Moreover, when recordings were performed in Ca2+-free BBS, no Itrans could be detected, while the cell did respond to puff applications of 1 mM D-aspartate, confirming that they possessed functional glutamate transporters (n = 3, data not shown). These experiments confirmed that Itrans was dependent on Ca2+ influx through voltage-activated Ca2+ channels.
Comparison of the charge carried by Itrans and reciprocal inhibition
When the intracellular solution contained 10 mM EGTA, the observed IPSCs mostly corresponded to spontaneous rather than reciprocal inhibition synaptic events. The event frequency was not increased in the 100 ms following depolarization compared to baseline, and IPSCs could still be observed after the run-down of glutamate release (data not shown). The ratio of the current time integrals of Itrans and of the mean of the IPSCs was 2.1 ± 0.4 (n = 10 cells) for depolarizations evoking maximal Itrans. To increase the probability of monitoring reciprocal inhibition, we first decreased the EGTA concentration in the pipette to 0.5 mM. This induced an increase in the average Itrans first and second peak amplitudes from –69.7 ± 6.6 pA and –43.1 ± 3.9 pA with 10 mM EGTA to –100.6 ± 5.9 pA and –66.0 ± 5.2 pA with 0.5 mM EGTA. Consequently, the charge carried by Itrans increased from –531 ± 71 pA ms (n = 10) to –2074 ± 408 pA ms (n = 6), while the charge carried by a mean IPSC did not change (–317 ± 55 pA ms for 10 mM EGTA, –330 ± 108 pA ms for 0.5 mM EGTA). Lowering the EGTA in the pipette also had a tendency to accelerate the Itrans run-down (Fig. 5C), but not significantly (P = 0.49, repeated measurements ANOVA, n = 9 and 10 cells). Reciprocal inhibition was still not seen following 2 ms depolarizations. Many studies in which reciprocal inhibition has been reported (Hartveit, 1999; Singer & Diamond, 2003; Vigh & von Gersdorff, 2005) used longer depolarizations and Cs+/TEA rather than K+-based internal solutions. We thus used a low EGTA (0.1 mM) Cs+/TEA-based intracellular solution, including 5 mM glutamate, to try to slow down the run-down of Itrans. While 2 ms depolarization did evoke reciprocal inhibition in only 1 cell out of 14, longer depolarizations (12–52 ms in 10 ms steps) progressively evoked more reciprocal IPSCs as well as a longer slow phase (Fig. 7). IPSCs were blocked by a cocktail of 1 µM strychnine + 50 µM TPMPA + 100 µM bicuculline methiodide (Fig. 8A and B) allowing to isolate the noisy slow phase. On the other hand, puff application of 75 µM DL-tBOA inhibited the slow phase but not the IPSCs. For 50 ms depolarizations, the time integral of Itrans (slow phase) was 1.40 ± 0.15 time larger than the charge carried by reciprocal IPSCs (mean of the charge from 4 sweeps prior to inhibitory cocktail application) (n = 4 cells).
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Discussion |
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In the mammalian retina, EAAT5 localization has been reported in the rat (Pow & Barnett, 2000; Pow et al. 2000), cat (Fyk-Kolodziej et al. 2004), rabbit and macaque retina (Pow et al. 2000). Rod photoreceptor terminals were labelled in all species, while expression in cone terminals was only confirmed in the cat retina, excluded in the macaque retina and remained uncertain in the rat and rabbit retina. RBC terminals as well as some cone bipolar cells were labelled in the adult rat retina (Pow & Barnett, 2000), while EAAT5 was not reported in rabbit, macaque and cat bipolar cells (Fyk-Kolodziej et al. 2004). In the cat retina, some amacrine and ganglion cells were labelled in addition to photoreceptors. We report here that in the mouse retina, EAAT5 is expressed in terminals of cone and rod photoreceptors. Cone pedicles were more strongly stained than rod spherules, a striking difference with the macaque retina. EAAT5 is also expressed in the axon and axon terminals of RBCs, most of the labelling in the INL and IPL colocalizing with the RBC-specific marker PKC.
Glutamate transporters as postsynaptic receptors in the mammalian retina
Vertebrate photoreceptors continuously release glutamate in the dark and hyperpolarize in response to light, resulting in a decrease in glutamate release. This translates into the hyperpolarization of OFF bipolar cells via the closure of AMPA or kainate receptors (DeVries, 2000) and into the depolarization of ON bipolar cells via activation of mGluR6 metabotropic receptors (Slaughter & Miller, 1981; Nawy & Copenhagen, 1987). An additional glutamate-sensitive conductance has been reported on the dendrites of lower vertebrate ON bipolar cells (Saito et al. 1979; Kondo & Toyoda, 1980; Nawy & Copenhagen, 1987) and attributed to a glutamate transporter (Grant & Dowling, 1995, 1996; Wong et al. 2005). Previous studies of glutamate responses of mammalian bipolar cells did not report the existence of glutamate-dependent dendritic current besides the one controlled by mGluR6 (de la Villa et al. 1995; Euler et al. 1996). Hasegawa et al. (2006) recently reported that glutamate transporters did not contribute to the RBC response following rod depolarization. Thus the DL-tBOA-sensitive current of small amplitude that we observed following puff applications of glutamate on mouse RBC dendrites/soma most probably originates from non-synaptic EAAT5. It may rather arise from EAAT5 located on the RBC soma, as suggested by the immunolocalization data (Fig. 1 and Supplemental Fig. 1).
Glutamate transporters as presynaptic receptors in the mammalian retina
A presynaptic current with the characteristics of a transporter-associated current has been observed in lower vertebrate and mammalian photoreceptors (Sarantis et al. 1988; Tachibana & Kaneko, 1988; Picaud et al. 1995b; Hasegawa et al. 2006) and in goldfish bipolar cells (Palmer et al. 2003). Both EAAT2 and EAAT5 have been localized in photoreceptor terminals and in bipolar cells of the salamander (Eliasof et al. 1998), goldfish (Vandenbranden et al. 2000) and mammalian retina (Rauen et al. 1996; Pow & Barnett, 2000; Pow et al. 2000; Fyk-Kolodziej et al. 2004). However, EAAT2 has a small Cl– conductance and is sensitive to dihydrokainate, while the currents reported here and from goldfish bipolar cell terminals were not (Palmer et al. 2003). Hence EAAT5, which is the glutamate transporter with the largest Cl– conductance, is the most likely candidate to account for the currents reported in non-mammalian species and in the present study.
In mouse RBC, short depolarizations evoke a transient current that can be blocked either by the glutamate transporter inhibitor DL-tBOA (75 µM), or by the Ca2+ channel blocker Cd2+ (100 µM) or by removal of external Ca2+. As expected from the release properties of bipolar cells from rats (Singer & Diamond, 2006) or lower vertebrates (von Gersdorff & Matthews, 1997; von Gersdorff et al. 1998; Palmer et al. 2003), a strong paired pulse depression was observed. The EAAT5 current may have been missed in previous studies for various reasons, the most obvious one being its fast rundown in whole-cell configuration. Classically, recordings in whole-cell mode are not performed right away after breaking into the cell, to allow enough time to achieve an equilibrium in the composition between the pipette medium and the intracellular compartment, and a proper compensation for the membrane capacitance and access resistance. In addition, responses to glutamate were often envisaged from a dendritic point of view, glutamate being puffed in the vicinity of dendrites rather than on axon terminals. When considering feedback IPSCs in RBC terminals, this current if still present in the initial recordings could have been masked by long depolarizing pulses as in Hartveit (1999) and by K+ channel tail current evoked by stronger depolarization when the repolarizing potential is far from EK.
The depolarization-induced current was always smaller than the glutamate-evoked steady-state current, but still could be as large as 88% of its amplitude, indicating that most presynaptic transporters can be activated by the glutamate released following a single depolarization. This contrasts with what has been reported in the goldfish RBC, in which only 5% of the EAAT5-like glutamate transporters were activated by comparable depolarizations (Palmer et al. 2003).
It should be noted that in vertebrates, all reported transporters acting as presynaptic receptors were located at retinal glutamatergic ribbon synapses. Equivalent synapses exist in other sensory structures relying on graded potential neurons (e.g. auditory and vestibular hair cells) for which the localization of glutamate transporters has not been studied as thoroughly as in the retina. A fast and graded feedback directly linked to the transmitter release as the one offered by EAAT5 may be a common feature to these synapses, which have a higher rate of vesicular release than conventional synapses.
Role(s) of presynaptic EAAT5 in RBC
In mouse RBC axon terminals, ECl is close to –60 mV (Varela et al. 2005). Thus, EAAT5 will physiologically act as an inhibitory presynaptic receptor, opposing the terminal depolarization and providing a faster feedback than the one mediated by reciprocal synapses with amacrine cells. In conditions in which reciprocal inhibition could be monitored, the slower time course of the EAAT5 current makes it a larger charge carrier than inhibitory GABA and glycinergic inputs to the RBC terminals. It should be noted that, in those conditions, the glutamate release and clearance may have been affected. First, the Ca2+-buffer used was weaker than the endogenous buffer, considered to be equivalent to 0.4 mM BAPTA or 2 mM EGTA (Burrone et al. 2002). Second, as K+ was replaced by Cs+ in the intracellular solution, the capacity of presynaptic EAAT5 to clear the glutamate from the synaptic cleft should be reduced as intracellular K+ is required for proper glutamate transport. Both aspects should prolong Itrans and accentuate the amount of reciprocal inhibition received from amacrine cells, especially if EAAT5 is a key element in this clearance at RBC terminals, as recently reported for rod photoreceptors (Hasegawa et al. 2006). With K+-based solutions and higher Ca2+ buffering capacities, especially for short depolarizations, the EAAT5 current should have an even more preponderant role in the feedback received by the RBC terminals.
Both GABA receptors and glutamate transporters can be modulated by PKC, which is highly expressed in RBCs (Greferath et al. 1990). While GABA receptors are down-regulated by PKC in rod bipolar cells (Feigenspan & Bormann, 1994; Gillette & Dacheux, 1996), PKC is expected to up-regulate glutamate transporters (Casado et al. 1993). Thus the exact balance between RBC auto-inhibition via EAAT5 and feedback from amacrine cells may vary depending on retinal activity.
The current-mediated action of EAAT5 can be strengthened by the fact that the charge carrier is Cl–. Indeed, L-type Ca2+ channels in salamander rod photoreceptors can be modulated through a mechanism independent of G protein and phosphorylation (Kourennyi & Barnes, 2000), which may correspond to variations in the intracellular Cl– concentration, a lower [Cl–]i inhibiting Ca2+ entry (Thoreson & Stella, 2000; Thoreson et al. 2003). Following this line, Ca2+ channel inhibition by glutamate transporter activation reported in these cells can be explained through the EAAT5-mediated Cl– efflux (Rabl et al. 2003). A similar mechanism can be considered in RBCs, which also express L-type Ca2+ channels on their axon terminals (de la Villa et al. 1998). The relative amplitude of Itrans compared to IGlu suggests that the transporters are localized close to the release site, and thus close to the Ca2+ channels responsible for the Ca2+ entry controlling vesicular release. Thus, the impact of the Cl– flux mediated by EAAT5 may be larger on Ca2+ channel activity than on the global Cl– concentration in the terminal.
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, n
= 9). Data were normalized to the slope of the linear regression for each cell. The cells considered in A–C did not receive spontaneous GABAergic and/or glycinergic inputs to facilitate determination of the response peak.
