Skip to main content
NCBI home page
The Journal of Physiology logoLink to The Journal of Physiology
. 2010 Aug 31;588(Pt 20):3943–3956. doi: 10.1113/jphysiol.2010.191437

Fast glutamate uptake via EAAT2 shapes the cone-mediated light offset response in bipolar cells

Matthew J M Rowan 1, Harris Ripps 2,3, Wen Shen 1
PMCID: PMC3000584  PMID: 20807794

Abstract

Excitatory amino acid transporters (EAATs) are responsible for extracellular glutamate uptake within the retina, and are expressed by retinal neurons and Müller cells. Their role within glutamatergic synapses is not completely understood. In the salamander retina, five distinct EAAT-encoding genes have been cloned, making the amphibian retina an excellent system to study EAAT function. This study focused on sEAAT2, which is expressed in photoreceptor terminals and Off-bipolar cells in two isoforms, sEAAT2A and sEAAT2B. Using whole-cell patch-clamp recording, florescence imaging and antibody labelling methods, we systematically studied the functions of these two isoforms at the synapse between photoreceptors and bipolar cells, both in dark and with photic stimulation. Both sEAAT2A and sEAAT2B were sensitive to dihydrokainic acid (DHKA), a known EAAT2-specific inhibitor. Each isoform of sEAAT2 was found to play a role in tonic glutamate uptake at the cone synapse in darkness. Furthermore, presynaptic sEAAT2A strongly suppressed the rapid, transient glutamate signal from cones following light-offset. This was achieved by quickly binding exocytosed glutamate, which subsequently limited glutamate spillover to adjacent receptors at postsynaptic sites. Since the intensity and duration of photic stimulation determine the magnitude of these cone transient signals, we postulate that presynaptic cone EAATs contribute to the encoding of contrast sensitivity in cone vision.

Introduction

EAATs are a group of Na+- and K+-dependent membrane transporters. The molecular structures of EAATs are well conserved in mammalian and non-mammalian neurons and glial cells, and are expressed in the photoreceptors and bipolar cells of primate (Hanna & Calkins, 2007), mouse (Rauen et al. 2004) and salamander (Eliasof et al. 1998a). These transporters display two separate conductances when activated: a coupled Na+,H+,K+-dependent conductance which is necessary for the transporter to bind and translocate glutamate, and a stoichiometrically uncoupled Cl− conductance (Danbolt, 2001). The anionic conductance is indicative of EAAT uptake and has been used to assess EAAT activity (Arriza et al. 1997; Otis & Jahr, 1998). An EAAT-mediated Cl− conductance has been well documented within photoreceptors (Picaud et al. 1995; Grant & Werblin, 1996; Gaal et al. 1998). Although EAATs are present on salamander Müller cells, the glial cells in this species do not extend processes to the invaginations in cone terminals (Lasansky, 1973). Thus, EAATs in Müller cells perform less glutamate uptake at the salamander cone-bipolar cell synapse as compared with its activity in the inner retina (Brew & Attwell, 1987). This suggests that the EAATs localized within photoreceptor terminals are of major importance in removing synaptic glutamate within the outer plexiform layer (OPL). Pharmacological studies indicate that EAAT uptake can be blocked by highly specific, non-transportable antagonists, such as the EAAT2-specific inhibitor dihydrokainic acid (DHKA), and the broad EAAT inhibitor dl-threo-b-benzyloxyaspartic acid (TBOA).

This neuronal transporter plays a critical role in maintaining dark glutamate levels in the distal retina and also has been shown to slow the onset of light-evoked responses in horizontal cells (Roska et al. 1998; Veruki et al. 2006), indicating that EAAT2 controls tonic glutamate levels in the synaptic clefts of photoreceptors, which continuously release glutamate in the dark. A recent study suggests that accumulation of glutamatergic vesicles in cones during light stimulation causes a large, rapid exocytosis as light turns off (Jackman et al. 2009), followed by a large, transient spike in bipolar cells that receive cone inputs. The role EAATs play in encoding these transient glutamate signals in the distal retina is largely unknown.

The salamander retina is an ideal system in which to examine the role of EAAT2 in photoreceptor transmission, as salamander photoreceptors are readily accessible for electrophysiological study. Two forms of EAAT2 have been isolated and cloned from the salamander retina, designated sEAAT2A and sEAAT2B. sEAAT2A has been localized immunohistochemically to photoreceptor terminals and Müller cells within the OPL, while sEAAT2B is thought to be localized specifically in Off-bipolar cells. Importantly, both sEAAT2A and sEAAT2B have similar pharmacological properties, as both transporters are similarly inhibited with DHKA, with no significant difference in sensitivity in the micromolar range (Eliasof et al. 1998b).

In the present study we used whole-cell patch-clamp recording combined with fluorescence dye dialysis and antibody labelling methods to examine in detail the function of both EAAT2 isoforms at the glutamatergic synapse between photoreceptors and bipolar cells, both in the dark and in response to photic stimulation. We find that sEAAT2A and -2B are involved in maintaining a low level of tonic glutamate input to bipolar cells in darkness, and that fast presynaptic EAAT2 glutamate uptake acts to limit the light offset signal from cones. A similar mechanism might be of general importance within glutamatergic synapses in retinas of other species.

Methods

Retinal slice preparation

Larval tiger salamanders (Ambystoma tigrinum), purchased from Kons Scientific (Germantown, WI, USA) and Charles Sullivan (Nashville, TN, USA), were used in this study. The animals were kept in aquaria at 13°C under a 12 h dark–light cycle with continuous filtration. The retinas were collected from animals kept at least 6 h in the dark. Briefly, the animals were decapitated and double-pithed and the eyes were enucleated. All procedures were performed in accordance with the guidelines of the National Institutes of Health Guide for the Care and Use of Laboratory Animals and approved by the University's Animal Care Committee.

The retinal slices were prepared in a dark room, under a dissecting microscope equipped with powered night-vision scopes (BE Meyer Co., Redmond, WA, USA), an infrared illuminator (850 nm), an infrared camera and a video monitor. Briefly, the retina was removed from an eyecup in Ringer solution and mounted on a piece of microfilter paper (Millipore, Billerica, MA, USA), with the ganglion cell layer downward. The filter paper with retina was vertically cut into 250 nm slices using a tissue slicer (Stoelting Co., Wood Dale, IL, USA). A single retinal slice was mounted in a recording chamber and superfused with oxygenated Ringer solution, consisting of (mm): NaCl (111), KCl (2.5), CaCl2 (1.8), MgCl2 (1.0), Hepes (5.0) and dextrose (10), pH 7.7. For recording Ca2+ channel currents from cones, 10 mm BaCl and 40 mm TEA were substituted for an equimolar amount of Na+. For recordings of capacitance and endogenous EAAT currents, 20 mm TEA was substituted for an equimolar concentration of Na+. The recording chamber was placed on an Olympus BX51WI microscope equipped with a CCD camera linked to a monitor.

Whole-cell patch-clamp recording

Whole-cell recordings were performed on photoreceptors and bipolar cells in dark-adapted retinal slices, using an EPC-10 amplifier and Pulse software (HEKA Instruments Inc., Bellmore, NY, USA). Patch electrodes (5–8 MΩ) were pulled with an MF-97 microelectrode puller (Sutter Instrument Co., Novato, CA, USA). All recordings were performed under 850 nm infrared illumination to avoid exposure to visible light. Two-second photic stimulation was performed with red LED illumination (660 nm peak emission) focused directly upon the retinal slice, controlled with a HEKA amplifier output. In all recordings, inhibitory inputs from glycine and GABA were blocked with 10 μm strychnine and 100 μm picrotoxin, respectively. For recording transporter currents in response to exogenous glutamate, electrodes were filled with (mm): KCl (40), potassium gluconate (60), MgCl2 (1), EGTA (5.0), Hepes (10), and ECl = −20 mV in photoreceptors or ECl = 0 mV within Off-bipolar cells (pH 7.3). For experiments on bipolar cell dark membrane currents and light responses, 5 mm KCl and 95 mm potassium gluconate were substituted for the calibration of ECl = −60 mV. To record current–voltage relationship curves from photoreceptors and bipolar cells, KCl and potassium gluconate were replaced with CsCl and CsF, respectively, to block large K+ conductances.

For capacitance recordings from cones, the internal solution included (mm): CsF (94), TEA (9.4), MgCl2 (1.9), MgATP (9.4), EGTA (5.0), Hepes (32.9). The phase angle setting was made with Pulse software based on the EPC-10+ amplifier circuitry, and verified using a model cell.

For recordings of fast endogenous EAAT currents, CsF was replaced with CsNO3 to enhance EAAT currents and the other solutions remained the same as for capacitance measurements. Data were analysed and plotted using Pulse (HEKA) and Igor (WaveMetrics, Inc., Lake Oswego, OR, USA) or Microsoft Excel software.

Photoreceptors and rod- or cone-dominant bipolar cells were identified by their morphology with Lucifer Yellow dialysis through electrodes, as well as with their light response patterns. A gravity-driven perfusion system was used to superfuse all external solutions. The perfusion tube was placed 3 mm away from the retinal slice and was manually controlled for delivering drugs during the experiments. All of the chemicals used in this study were purchased from Sigma and Tocris Bioscience (Ellisville, MO, USA).

Immunohistochemistry

Freshly enucleated eyes were fixed in 4% paraformaldehyde in Ringer solution for 30 min followed by dissection of the retina from the eyecup and then rinsed extensively in Ringer solution. The eyecups were dehydrated in graded sucrose solutions (10%, 15%, 20% and 30%), and following overnight immersion in 30% sucrose, they were embedded in OCT compound (Ted Pella, Redding, CA, USA), frozen, and sectioned at 14 μm thicknesses. Frozen sections were collected on slides, air dried, and stored at 80°C.

Immunohistochemistry was performed on retinal sections and flat-mount retinas that were removed from eyecups after fixation. Retinal sections and flat-mounts were rinsed with PBS containing 0.1% Tween and 0.3% Triton X-100 (PBST-T), and then treated with a blocking solution consisting of 10% normal goat or donkey serum and 0.1% sodium azide in PBST-T. Retinal sections and flat mounts were then incubated with the primary antibody mixture, either with 3% goat or donkey serum and 0.1% sodium azide in PBST-T for 2 h at room temperature or for 5 days at 4°C, respectively. Controls lacking the primary antibody were unstained. After numerous washes in PBS with 0.1% Tween (PBST) containing 0.1% sodium azide, immunoreactivity was tested after incubation with immunofluorescent secondary antibodies for 40 min at room temperature for retinal sections and overnight at 4°C for flat mounts in PBST-T containing 0.1% sodium azide. The retinas were subsequently rinsed with PBST, mounted with Vectorshield (Vector Laboratories, Burlingame, CA, USA) and viewed with a confocal laser scanning microscope (Zeiss, LMS 700). Images were acquired via 40× and 60× oil-immersion objectives, and then processed with Zen software (Trumbull, CT, USA).

Results

sEAAT2A in photoreceptor terminals

Antibody labelling has shown that sEAAT2A is present in photoreceptor terminals (Eliasof et al. 1998b). We used DHKA, a non-transported and EAAT2-selective antagonist, to separate sEAAT2A currents from total glutamate transporter currents in both rods and cones. DHKA is a reliable antagonist with Ki values between 24 μm and 79 μm for EAAT2, and has no significant effect on other EAATs near these concentrations (Eliasof et al. 1998a). Experiments were performed on dark-adapted retinal slices with an inhibitory cocktail consisting of 100 μm picrotoxin and 10 μm strychnine to block the effects of GABAergic and glycinergic inputs from the network. A high Cl− intracellular solution was used for photoreceptors to mimic natural conditions, in which the rod ECl is around −20 mV (Thoreson et al. 2002). Because glutamate uptake occurs most efficiently at lower voltages (Levy et al. 1998), photoreceptors in dark-adapted retinal slices were held at −50 mV, a potential negative to the ECl in the experiments. Glutamate at 1 mm was applied to activate glutamate transporters, which elicited large, sustained steady state inward currents in both rods and cones (Fig. 1). These inward currents are EAAT specific Cl− currents, as identified in previous studies (Grant & Werblin, 1996; Gaal et al. 1998). With the cells held at −50 mV, the amplitude of the glutamate-elicited currents at −50 mV was significantly larger in cones (131.21 ± 12.56 pA; mean ± s.e.m.; n = 7), as compared with −30.35 ± 3.73 pA (n = 7) in rods. DHKA at 100 μm reduced the inward currents in cones but had no effect on rod inward currents; residual DHKA-insensitive currents in cones and rods were completely blocked by 3 μm TBOA (Fig. 1A). Plots of the current–voltage relationship of glutamate transporter currents from cones and rods (Fig. 1B) showed a characteristically linear relationship at potentials negative to the ECl. Currents reversed near the ECl, −20 mV. In cones, 100 μm DHKA reduced the amplitude of glutamate-elicited currents over a broad voltage spectrum, but had no effect on the ECl reversal potential (Fig. 1B, left panel, n = 11).

Figure 1. One millimolar glutamate-elicited EAAT currents in photoreceptors at holding voltage −50 mV, ECl = −20 mV.

Figure 1

Open in a new tab

A, steady state glutamate currents in cones (left) and rods (right) in control and with either 100 μm DHKA or 3 μm TBOA. B, typical current–voltage relationship of glutamate-elicited currents in cones (n = 11) and rods (n = 8). Cones are inhibited by DHKA (grey trace) over a wide voltage range, while rods were unaffected. C, average peak steady state glutamate currents (black bars) in cones and rods. Cones are significantly inhibited by DHKA (36.97 ± 5.65%; P < 0.0001; n = 7), whereas rods are unaffected. Both cones and rods are fully inhibited by TBOA (90.10 ± 6.57%, P < 0.0005, n = 7 and 95.83 ± 3.39%, P < 0.01, n = 7, respectively). Asterisk denotes statistically significant differences.

In cones, DHKA inhibited a significant portion (36.97 ± 5.65%) of the control glutamate currents at −50 mV (Fig. 1C left panel, P < 0.0001, n = 7), whereas 90.10 ± 6.57% of the remaining transporter current in cones was blocked with TBOA (Fig. 1C left panel, P < 0.0005, n = 7). The TBOA-sensitive glutamate current is probably mediated by at least one other subtype of sEAAT, most likely sEAAT5, previously detected in salamander photoreceptors (Eliasof et al. 1998b). Our results agree with those of a previous study (Eliasof & Werblin, 1993) of isolated cones. Although rods were unaffected by 100 μm DHKA under the same conditions (Fig. 1A and B, right panels), the vast majority of the overall glutamate-induced currents in rods were blocked by TBOA, which suppressed 95.83 ± 3.39% of the current recorded in the control group (P < 0.01, n = 8, Fig. 1C right panel). The data from rods is consistent with a previous study in rat, in which rod EAAT currents were completely inhibited with TBOA, but were unaffected by 200 μm DHKA (Hasegawa et al. 2006). Therefore, presynaptic sEAAT2A clearly plays a unique role within cone photoreceptors in the OPL of tiger salamander retina.

Evidence of sEAAT2B in Off-bipolar cells

The presence of EAAT2 in bipolar cells has been reported in primate, rat and salamander retina (Eliasof et al. 1998a; Rauen et al. 2004; Hanna & Calkins, 2007). A previous study depicted sEAAT2B as a neuronal-specific glutamate transporter in bipolar cells of the salamander retina (Eliasof et al. 1998b). To determine the specific locale of sEAAT2B, we used a salamander-specific antibody against sEAAT2B, developed by Dr Susan Amara. Confocal imaging of a retinal section labelled with sEAAT2B with and without background staining of Acridine Orange (AO), a cell nuclei marker, indicates that sEAAT2B is present in both bipolar cell dendrites and axon terminals within the distal layer of the inner plexiform layer (IPL) (Fig. 2). sEAAT2B labelling was present in displaced Off-bipolar cells, which have somas located in the outer nuclear layer (ONL) (Fig. 2A). Displaced Off-bipolar cells in salamander have been shown to receive predominant input from cones (Maple et al. 2005). Localization of sEAAT2B within the OPL was detected by single-plate confocal scanning at the photoreceptor terminal with sEAAT2B and SV2 double-labelling, a synaptic vesicle protein II (SV2) which labels photoreceptor terminals, in the flat-mounted retina. Figure 2B displays an example of sEAAT2B (red) and SV2 (green) labelling at the distal boundary of the OPL, and a superimposed image of both stains. Double-labelling indicated that anti-sEAAT2B and anti-SV2 labelled both bipolar cell dendrites and photoreceptor terminals, with no co-localization detected at the terminals of rods or cones (Fig. 2B, right panel). sEAAT2A and sEAAT2B are therefore separately expressed at the pre- and postsynaptic sites of the OPL, offering an opportunity to study the role of two EAAT2 variants at specific pre- and postsynaptic sites within the retina.

Figure 2. Confocal imaging of sEAAT2B labelling in retina.

Figure 2

Open in a new tab

A, sEAAT2B labelling (red) with and without background Acridine Orange (AO) (green) nuclear stain in a retinal section. sEAAT2B labels both regular and displaced Off-bipolar cells located in the INL and ONL, respectively. B, single scanning imaging from a double-labelled flat-mounted retina, in which SV2 and sEAAT2B are separately located at photoreceptor terminals and the postsynaptic dendrites, respectively, in the distal OPL.

sEAAT2B-mediated currents were investigated in Off-bipolar cells in dark-adapted retinal slices in whole-cell voltage-clamp mode. Lucifer Yellow was added to the electrode solution to depict the cell morphology after recording. Recordings were made with ECl = 0 mV. In the control, glutamate receptors in Off-bipolar cells were blocked by 100 μm 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX), a specific AMPA receptor inhibitor. PTX at 100 μm and 10 μm strychnine were applied to block inhibitory inputs. Glutamate at 1 mm with CNQX elicited an inward current in cells held at −60 mV. The CNQX-insensitive currents were blocked with 100 μm DHKA, indicating an EAAT2-specific transporter current (Fig. 3A). The current–voltage relationship of the DHKA-sensitive currents was linear and had a reversal potential near the ECl. (Fig. 3B, n = 6). Similar results were obtained in displaced Off-bipolar cells and non-displaced Off-bipolars (Fig. 3C). Displaced Off-bipolar cells displayed the morphological features of cone-dominated bipolar cells, as previously described (Maple et al. 2005). Non-displaced Off-bipolar cells normally had the morphology of rod–cone mixed cells, with the axon terminals located throughout the Off sublamina, as described previously (Pang et al. 2004). We analysed DHKA-sensitive currents from Off-bipolar cells with somas located in the ONL and inner nuclear layer (INL), and found that the average currents were 26.01 ± 3.48 pA (n = 8) and 28.74 ± 4.09 pA (n = 7), respectively (Fig. 3C). This indicates that sEAAT2B-mediated currents in Off-bipolar cells were significantly smaller than sEAAT2A-mediated currents in cones.

Figure 3. sEAAT2B-mediated currents in Off-bipolar cells.

Figure 3

Open in a new tab

A, with CNQX added to block AMPA receptors in Off-bipolar dendrites, glutamate elicits a steady inward current that is blocked by DHKA, indicating an sEAAT2B-mediated current. B, the current–voltage relationship curve of the DHKA-sensitive currents in Off-bipolar cells. C, morphology of displaced and regular (non-displaced) Off-bipolar cells, following Lucifer Yellow dialysis during whole-cell recoding, with statistical analysis of the sEAAT2B-mediated currents from each morphological type. EAAT2B currents were not significantly different among the subtypes of Off-bipolar cells.

Glutamate transporter currents were investigated from On-bipolar cells in the presence of the specific antagonists (RS)-α-cyclopropyl-4-phosphonophenylglycine (CPPG) and (+)-α-methyl-4-carboxyphenylglycine (MCPG), to block metabotropic glutamate receptors in On-bipolar dendrites. Glutamate at 1 mm elicited glutamate transporter currents in On-bipolar cells that were DHKA insensitive, but were TBOA sensitive (data not shown), indicating that On-bipolar cells lack sEAAT2B, but are likely to express other EAATs. Our data agree with immuno-antibody labelling (Eliasof et al. 1998a) that sEAAT2B is Off-bipolar cell specific in the salamander.

sEAAT2 maintains synaptic glutamate levels in dark

Rod and cone photoreceptors are constantly depolarized in dark, which allows for a continuous release of glutamate. Therefore, sEAAT-specific glutamate uptake most likely takes place at the photoreceptor terminal to prevent glutamate depletion at the presynaptic release site (Hasegawa et al. 2006). Accordingly, blocking sEAAT2-mediated glutamate reuptake in cones would cause an increase in glutamate levels in the OPL.

In general, bipolar cells in salamander retina receive mixed inputs from both rods and cones, although some bipolar cells receive a majority of input from either rods or cones, which are designated rod-dominant and cone-dominant bipolar cells, respectively (Pang et al. 2004). Rod- or cone-dominant neurons can be distinguished by the location of their axon terminals. In addition, rod-dominant bipolar cells display a slow recovery at light offset, resembling the light response of rods (Schnapf & Copenhagen, 1982), while cone-dominant bipolar cells inherit the features of the cone light response, characterized by a fast response at the offset of a light stimulus. These criteria were used to characterize the effect of DHKA on the different subtypes of bipolar cells. Cell morphology was revealed with Lucifer Yellow dialysis. Picrotoxin at 100 μm and 1 μm strychnine blocked inhibitory inputs, and 100 μm cyclothiazide (CTZ) was included to prevent AMPA receptor desensitization in bipolar cells. Dark currents were recorded from Off-bipolar cells that were voltage-clamped at −60 mV, near the ECl, to minimize EAAT currents. DHKA at 100 μm was then applied, which generated a steady inward current in Off-bipolar cells (Fig. 4Aa and b). Since DHKA inhibits sEAAT2 reuptake of glutamate at the synapse, the observed inward current in Off-bipolar cells was most likely glutamatergic, caused by glutamate accumulation in the synaptic cleft and subsequent activation of postsynaptic AMPA receptors. CNQX, which specifically blocks AMPA receptors located in Off-bipolar cell dendrites, was used to verify that the DHKA-elicited current was a consequence of increased synaptic glutamate in the OPL. Indeed, these inward currents were fully blocked by 100 μm CNQX (Fig. 4Aa and b). The average DHKA-induced AMPA currents were significantly larger in cone-dominant Off-bipolar cells (−32.76 ± 6.00 pA, n = 15), when compared with rod-dominant Off-bipolar cells (−8.373 ± 0.4 pA, n = 9) as illustrated in Fig. 4Ac (P < 0.01, n = 9). As DHKA blocks both sEAAT2A and sEAAT2B, the larger DHKA-induced current in cone-dominated Off-bipolar cells (Fig. 4Aa) is likely to be due to the fact that both sEAAT2A and sEAAT2B uptake was blocked at this synapse, while at rod-dominated Off-bipolar cell synapses (Fig. 4Ab), only EAAT2B uptake is blocked. This finding also suggests that presynaptic uptake via sEAAT2A is significantly stronger at the cone to bipolar cell synapse.

Figure 4. DHKA-elicited currents in rod- and cone-dominated bipolar cells in dark-adapted retinal slices at holding voltage −60 mV, ECl = −60 mV.

Figure 4

Open in a new tab

Aa, DHKA elicits an inward current, which can be blocked by CNQX, in cone-dominated Off-bipolar cells (n = 15). b, DHKA elicits a CNQX sensitive inward current in rod-dominant Off-bipolar cells (n = 9). c, comparison of the average absolute DHKA-elicited currents in cone- and rod-dominant Off-bipolar cells. DHKA elicited a significantly larger current in cone-dominant Off-bipolar cells (cone-dominant: −32.76 ± 6.00 pA; rod dominant: −8.373 ± 0.4 pA; P < 0.01). Ba, DHKA elicits an outward current, which can be blocked by CPPG, in cone-dominant On-bipolar cells (n = 4). b, DHKA failed to elicit a current in all rod-dominant On-bipolar cells (n = 5). c, comparison of the effects of DHKA in cone- and rod-dominant On-bipolar cells (cone-dominant: 16.79 ± 5.62; rod-dominant: 0.035 ± 0.18 pA; P < 0.05). Asterisk denotes statistically significant differences.

The effect of DHKA was also investigated in On-bipolar cells, which utilize metabotropic glutamate receptors on their dendrites. Activation of these metabotropic receptors closes cation permeable channels in the cells; therefore On-bipolar cells are hyperpolarized in dark by constant glutamate release from photoreceptors. DHKA-induced glutamatergic responses were recorded from cone- and rod-dominant On-bipolar cells held at −60 mV. DHKA at 100 μm increased glutamatergic inputs to cone-dominant On-bipolar cells, resulting in an outward current in the neurons, but did not affect rod-dominant On-bipolar cells (Fig. 4Ba and b). The outward currents in cone-dominant On-bipolar cells were blocked by 100 μm CPPG, a specific antagonist for metabotropic glutamate receptors in On-bipolar dendrites (Fig. 4B, n = 4). The average DHKA-induced metabotrophic currents were significantly larger in cone-dominant On-bipolar cells (16.79 ± 5.62 pA, n = 5) when compared with rod-dominant On-bipolar cells (0.035 ± 0.18 pA, n = 5), as shown in Fig. 4Bc (P < 0.01, n = 5). Results from On-bipolar cells suggest that presynaptic sEAAT2A plays an important role at the synapses of cone-dominant On-bipolar cells. These findings are in accordance with the results in Fig. 1, namely that sEAAT2 is a cone-pathway specific transporter.

Fast uptake via sEAAT2A shapes light-evoked cone signals to bipolar cells

As presynaptic sEAAT2A uptake was found to maintain lower tonic glutamate levels from cones in dark, we postulated that the transporter might also play a role in shaping light-evoked, transient glutamate release from cones to bipolar cells. Bipolar cells receiving cone inputs receive fast, transient potentials when presynaptic cones are depolarized (Cadetti et al. 2005, 2008), a phenomenon that naturally occurs at light offset. The ‘off-overshoot’ response is a cone-specific signal that encodes the intensity and duration of a light stimulus, and perhaps the motion of light across the visual field (Wu, 1988). The effect of sEAAT2A on the light-evoked, glutamatergic ‘off-overshoot’ was studied in bipolar cells with application of DHKA. Intracellular and extracellular solutions were kept as in Fig. 3, with ECl = −60 mV; controls included 100 μm picrotoxin, 10 μm strychnine and 100 μm CTZ. A 2 s light stimulus was used to evoke current responses in the neurons. The effect of 100 μm DHKA on light-evoked responses was tested on both cone- and rod-dominant bipolar cells. Typical light-evoked current responses in Off- and On-bipolar cells receiving inputs predominately from cones are depicted in Fig. 5 (black traces). Light stimulation produced an outward current in Off-bipolar cells and an inward current in On-bipolar cells. Cones rapidly release a store of vesicles at the offset of a light stimulus which in turn evokes a transient glutamatergic current in cone-dominated bipolar cells, as shown by the black traces in Fig. 5A and B. DHKA significantly increased the amplitude of the transient current at the light offset in both cone-dominant Off- and On-bipolar cells, although the increase was much larger in Off-bipolar cells (grey traces, Fig. 5A and B). In cone-dominant Off-bipolar cells, the overshoot was enhanced by DHKA from an average control amplitude of −12.88 ± 2.11 pA to −59.94 ± 5.15 pA (P < 0.01 n = 6, Fig. 5C). The average enhancement of the overshoot in On-bipolar cells was far less, i.e. from 14.07 ± 3.52 pA to 24.05 ± 5.89 pA with DHKA (P < 0.05, n = 5, Fig. 5B). In contrast, DHKA had no apparent influence on the light offset current response in cells receiving a majority of inputs from rods (Fig. 5D). Our results suggest that sEAAT2 rapidly binds extracellular glutamate, thereby limiting the amplitude of the ‘off-overshoot’ signal from cones to bipolar cells, most significantly within Off-bipolar cells.

Figure 5. The contribution of sEAAT2A to light responses in rod- and cone-dominated bipolar cells.

Figure 5

Open in a new tab

A, cone-dominated Off-bipolar cells display a transient inward current at the offset of a 2 s light stimulus. DHKA causes a large enhancement in the light-offset current. B, cone-dominated On-bipolar cells display a transient outward current at the offset of a 2 s light stimulus. DHKA enhances the light-offset response. C, enlarged light-offset currents from a cone-dominated Off-bipolar cell. D, rod-dominated Off-bipolar cells display a slow light-offset response; DHKA did not alter the light-offset response in the neurons.

Inhibition of sEAAT2A did not affect vesicular release from cones

Light-evoked offset responses in second-order neurons originate from presynaptic cones (Wu, 1988). In order to isolate a mechanism underlying the observed EAAT2-specific suppression of cone signals, it is necessary to determine whether EAAT2A inhibition directly increases the magnitude of glutamate release from cones. To test this, we recorded the effects of DHKA on the cone light response, the voltage-dependent Ca2+ channel currents, and cell capacitance changes following depolarization. Controls included 100 μm PTX and 10 μm strychnine to block inhibitory synaptic inputs.

The effect of 100 μm DHKA on light-evoked responses was recorded from cones in dark-adapted retinal slices, with the same internal solution used for Fig. 1. Cones were voltage clamped at −50 mV, slightly negative to the dark membrane potential of the neurons. A light stimulus closed cGMP gated channels in the cones, halting cation influx in dark and causing an outward current shift. At the termination of a light stimulus, a large inward current can be elicited in cones (Fig. 6A). DHKA at 100 μm had no effect on light-evoked responses in cones when a low Cl− intracellular solution (potassium gluconate) was used (n = 5), indicating that the inhibition of sEAAT2A had no direct effect on the light-evoked cone response. DHKA also had no effect on the cone dark membrane potential, in agreement with a previous study in salamander (Yang & Wu, 1997).

Figure 6. EAAT2A inhibition does not alter cone light responses or exocytosis.

Figure 6

Open in a new tab

A, cones display a characteristic outward current during a light response. DHKA has no significant effect on the light-evoked responses of cones, or on currents following the light response (n = 5). B, voltage gated Ca2+ currents were isolated in cones with external Ba2+ with a 25 ms voltage step to −10 mV. DHKA does not alter voltage-dependent Ca2+ channel currents in cones (n = 7). C, cellular capacitance measurements (Cm) were performed prior to and following a 100 ms depolarizing step from −70 mV to −10 mV to measure exocytosis. DHKA did not affect the depolarization-evoked capacitance changes (n = 5).

A shift in voltage-gated Ca2+ channel currents following EAAT2A inhibition might alter the amount of vesicular release from cones. To test if an EAAT2A-directed modulation of cone voltage-gated Ca2+ channels occurs, 100 μm DHKA was applied while recording Ca2+ channel currents in cones. Ba2+ and TEA were added to the external solution, with TEA and CsCl-containing electrode solution to isolate Ca2+ channel currents. Maximal sustained inward Ca2+ channel currents were elicited from cones by a single voltage-step from the holding potential to −10 mV, and were recorded for 25 ms (Fig. 6B). DHKA had an insignificant effect on the voltage gated Ca2+ channel currents in cones (n = 7), suggesting that the Ca2+-dependent glutamate release in cones is unaltered after EAAT2A inhibition.

To provide further evidence that EAAT2A activity did not modulate vesicular glutamate release, we recorded capacitance (Cm) changes following a depolarizing step. Transient increases in cell capacitance are correlated with the binding of vesicles at the plasma membrane. If EAAT2A inhibition changes any exocytotic mechanism, or somehow modifies the readily releasable pool of vesicles, a change in Cm following a depolarization would be apparent. Cells were voltage-clamped at −70 mV, and were subjected to a 600 Hz sine wave of ± 30 mV applied around the holding potential. HEKA software ‘Lock-in’ mode was utilized to measure the capacitance (Cm), access conductance (Gs) and membrane conductance (Gm) for each sine wave during the recordings. Gs and Gm did not significantly change during Cm recordings, indicating a stable control. Thirty-second intervals were permitted between depolarizing steps, which allows cone intracellular Ca2+ levels to return to prestimulus levels between pulses (Innocenti & Heidelberger, 2008). Cones had a total cell capacitance ranging from 18.3 to 24.8 pF in dark conditions. Depolarizing steps of 75 ms from −60 mV to −10 mV elicited a 202.4 ± 24.3 fF (n = 5) peak capacitance increase, within the range of values previously described in salamander cones in the slice preparation (Rabl et al. 2005). Assuming the fusion of a single vesicle at the cone synaptic terminal causes a capacitance increase of ∼0.056 fF (Thoreson et al. 2004), then the increase we observed in response to the 75 ms depolarizing step reflects the fusion of about 3614 vesicles undergoing exocytosis. DHKA at 100 μm did not affect the depolarization-evoked capacitance changes (Fig. 6C, n = 5). Taken together, the light-offset enhancement observed in bipolar cells after EAAT2A inhibition is probably not due to an enhanced presynaptic glutamate release, and EAAT2A activity therefore does not modulate the output of cones.

Comparison of EAAT2-mediated uptake at presynaptic cones and postsynaptic bipolar dendrites following fast glutamate release

Our results show that sEAAT2 plays a significant role in the encoding of fast cone signals in the OPL. Uncovering the distinct actions of the cone EAAT2A and bipolar cell EAAT2B is necessary, and offers a unique perspective into the physiological importance of presynaptic and postsynaptic transporters at the same synapse. Two separate experiments were devised to study the natural activity of EAAT2 in cones and bipolar cells following fast glutamate exocytosis. Both experiments utilized intracellular CsNO3 in the pipette solution (Fig. 7A and B) to amplify transporter kinetics following vesicular release from cones, as the NO3− ion is more readily conducted by the transporter after binding glutamate. TEA was included in the bath to reduce membrane conductance as in capacitance measurements, and the majority of cellular capacitance was compensated during these experiments. Picrotoxin at 100 μm and 10 μm strychnine were perfused constantly. All experiments were performed in dark conditions.

Figure 7. Presynaptic EAAT2A rapidly removes glutamate released from cones.

Figure 7

Open in a new tab

A, with intracellular NO3− added in the pipette to enhance EAAT currents, an inward current was elicited in cones following a 2 ms depolarizing step. A significant portion of the current was sensitive to DHKA. B, displaced Off-bipolar cells with truncated axons were chosen to study the EAAT2B activation following fast exocytosis in cones at the light offset. With intracellular NO3−, light responses were recorded from the Off-bipolar cells. Light offset responses were almost completely blocked with CNQX, indicting that transient glutamate release from cones has a negligible effect on EAAT2B in the dendrites of the Off-bipolar cells.

Cones were held at −70 mV and subjected to a short, 2 ms depolarizing step to −10 mV in order to measure evoked EAAT currents following vesicular release, as previously demonstrated in retinal bipolar cells of various species (Palmer et al. 2003, Wersinger et al. 2006). A 15 s interval was allowed following each depolarization. Large, transient inward currents were immediately evoked following the short depolarizing step (Fig. 7A). This inward tail current was partially inhibited with 100 μm DHKA, accounting for 35.3 ± 7.8% of the observed peak current following vesicular release (n = 7). This DHKA-sensitive transporter current indicated that cone specific EAAT2A rapidly binds extracellular glutamate following release. In addition, the DHKA-insensitive inward current following depolarization was completely eliminated with 50 μm TBOA (data not shown). TBOA is likely to also inhibit the EAAT5 subtype in cones, as suggested earlier. Our results suggest that in the presynaptic cone EAAT2A plays a significant role in limiting light-offset signals, through an immediate buffering of synaptic glutamate following release. The contribution of both transporters together is considered in the Discussion.

The activity of postsynaptic EAAT2B in Off-bipolar cells was studied in cells with truncated axons, to remove any possible transporter-associated currents that are likely to exist within the terminal. Only cone-dominant, displaced Off-bipolar cells were used in this experiment, which were readily accessible within the ONL. Immediately after breaking in, cells held at −60 mV were subjected to a 2 s light stimulus. Cells receiving strong cone inputs were selected (Fig. 7B, black trace) and 100 μm CNQX was then applied to inhibit AMPA inputs. The NO3− containing intracellular solution was allowed to diffuse for 1 min before resuming light responses. If EAAT2B was significantly activated following light offset, an observable DHKA-sensitive inward current should exist after the light cessation. We did not observe a significant DHKA sensitive current from the Off-bipolar cells with truncated axon (Fig. 7B, grey trace) and the results were repeated in four cells, indicating that dendritic sEAAT2B uptake following large glutamate release is significantly less effective as compared with presynaptic uptake via sEAAT2A. However, baseline currents were shifted inward slightly after the application of DHKA, suggesting that dendritic EAAT2B might play a lesser role in controlling the concentration of glutamate in the dark.

The function of EAAT2A at the cone to bipolar cell synapse

The function of EAAT2A in the regulation of cone to bipolar cell signalling was studied by recording light-evoked glutamate currents in bipolar cells in response to photic stimuli with different durations. Since glutamate vesicles are readily replenished at cone terminals when light hyperpolarizes the cones, the longer the hyperpolarization, the more vesicles are accumulated in the cone terminals. Therefore, an increase in light duration would be expected to increase glutamate vesicle release in cones at the end of the light stimulation. Using stimuli of 400 ms, 700 ms, 1 s and 4 s durations, we were able to demonstrate this variation in glutamate release from cones (Fig. 8) in the postsynaptic current recordings from cone-dominated Off-bipolar cells. Figure 8A (dark trace) shows typical current responses as the duration of light stimulation was increased. With application of 100 μm DHKA to block EAAT2A, the control current amplitudes and duration were enhanced. This enhancement was more pronounced at 1 s and 4 s of light stimulation compared to the 400 ms light stimulus (Fig. 8A, grey traces). These results suggest that glutamate uptake via EAAT2A is maximally facilitated in response to longer light exposures that produce a greater glutamate release at the termination of the photic stimulus. Similar results were also observed with increase of light intensity. Using 2 s light pulses at three levels of light intensity showed that the greater the intensity, the greater the inward current, and the greater its enhancement by 100 μm DHKA at light offset (data not shown).

Figure 8. The role of EAAT2A in modulating glutamate release.

Figure 8

Open in a new tab

A, light responses recorded from an Off-bipolar cell stimulated with flash durations of 400 ms, 700 ms, 1 s and 4 s. DHKA enhances the amplitudes and durations of light responses when the flash duration exceeded 400 ms. B, glutamate transporter currents were recorded from a cone that was held at the resting dark potential of −35 mV, and were hyperpolarized to −70 mV for durations of 400 ms, 700 ms, 2 s and 4 s. This was followed by a brief depolarization to −10 mV at each duration to evoke glutamate release. The glutamate uptake currents were elicited by each depolarization pulse and were increased progressively with longer hyperpolarization times. C, hypothetical model depicting the role of sEAAT2 at the cone to Off- bipolar cell synapse of the dark-adapted retina. In control conditions, glutamate released stimulates AMPA receptors on Off-bipolar cells and excess glutamate is taken up into the terminals by the sEAAT2A transporter. Blocking transporter activity with DHKA produces accumulation of glutamate in the synaptic cleft, which binds to more AMPA receptors and greatly enhances light offset response.

To confirm that glutamate uptake in cones is facilitated following strong glutamate release, we recorded glutamate transporter currents using the same protocol as described in Fig. 7A, i.e. the cone was held at −35 mV to mimic the dark membrane potential, and was hyperpolarized to −70 mV before triggering a short 50 mV depolarization pulse to −10 mV. In addition, the duration of the hyperpolarizing steps was varied from 400 ms to 4 s to match the time course of light stimulation used in Fig. 8A. Moreover, the patch pipette again delivered NO3− intracellularly to enhance the transport current, so that the inward transporter currents were readily observed at the end of the depolarization pulse (Fig. 8B). As predicted, transporter currents increased with increasing the durations of the hyperpolarization step. These transporter currents were again DHKA sensitive (data not shown).

Figure 8C depicts a possible mechanism for sEAAT2 in modulating the transient glutamatergic synapses of cones-to-bipolar cells. In this model, sEAAT2A-mediated uptake into cone terminals rapidly buffers glutamate levels in the OPL, thus limiting glutamate diffusion after transient glutamate release in the dark-adapted retina. Blockage of the presynaptic transporter allows for glutamate spillover to adjacent areas in the synaptic cleft, resulting in a larger synaptic input to bipolar cells after light offset-evoked transient glutamate release. In addition, sEAAT2B may serve as a postsynaptic transporter in Off-bipolar cell dendrites, thereby contributing to glutamate uptake in the cone-to-Off-bipolar cell synapse. Our model also suggests that sEAAT2A might be in closer proximity to glutamate release sites in cone terminals than previously suggested.

Discussion

Possible function of the presynaptic glutamate transporter at cone synapses

Photoreceptors are unique in that they are able to transiently release glutamate in response to changing levels of light and dark. The rapid burst of transmitter release from cones at light offset has been reported as due to a buildup of vesicles on the synaptic ribbon in light. These stored vesicles undergo a rapid exocytosis at light offset (Jackson et al. 2009), causing the characteristic ‘off-overshoot’ signal observed from cone dominant On- and Off-bipolar cells as well as from ganglion cells. Our study demonstrates that EAAT2 is involved in encoding these light offset signals in the distal retina. We find that EAAT2 inhibition enhances the amplitude of cone-mediated transient responses in bipolar cells. However, it is noteworthy that DHKA has no direct influence on Ca2+ channel currents or exocytosis-associated capacitance changes in cones, indicating that the signal increase in bipolar cells following sEAAT2A inhibition is due to an increase of glutamate input in bipolar cells, without a shift in vesicular output from cones. We postulate that inhibition of sEAAT2A causes excessive glutamate spillover, thereby activating more AMPA receptors in bipolar dendrites after rapid release from cones. Rapid reuptake of glutamate in cones at light offset appears necessary for cone vision to retain its high sensitivity to changing levels of illumination. Our results show that at the levels of illumination used in this study, presynaptic sEAAT2A strongly reduces cone signal transmission to bipolar cells. Further study is necessary to determine the role of EAAT2A in shaping cone signals over a larger range of contrast illumination.

Fast reuptake of glutamate by presynaptic sEAAT2A may be critical in preventing glutamate depletion within cone terminals. Since glutamate is continuously released in darkness, transmitter reuptake into cone terminals would serve to prevent synaptic depression in photoreceptors and prevent over-activation of postsynaptic receptors. Previous studies indicate that DHKA-sensitive transporters balance glutamate levels internally and externally in cones, and therefore help retain a steady tonic glutamate synapse in dark (Picaud et al. 1995; Gaal et al. 1998). Our results agree with these previous studies that DHKA-sensitive transporters are responsible for maintaining tonic glutamate levels in the OPL in dark; we suggest that EAAT2A activity is most important with respect to limiting excess glutamate buildup within the synaptic cleft.

Different types of EAATs control glutamate uptake in rod and cone terminals

It is generally accepted that EAAT uptake is tightly coupled with the associated glutamate transporter current, including the large, thermodynamically uncoupled anion current (Arriza et al. 1997; Otis & Jahr, 1998; Palmer et al. 2003). This Cl− conductance is variable among EAAT subtypes; for example, sEAAT5 has a larger Cl− conductance compared to sEAAT2. Previous studies have revealed that sEAAT5 is also present in photoreceptors (Eliasof et al. 1998), suggesting that glutamate uptake in salamander cones is performed by both sEAAT5 and sEAAT2A subtypes. As DHKA specifically blocks sEAAT2A, we could pharmacologically separate the currents of these two subtypes. We report that sEAAT2A mediates ∼37% of the total glutamate transporter current. Clearly, sEAAT2A does not elicit the most significant anion current. Although sEAAT2A is less permeable to Cl−, its contribution to glutamate clearance is highly significant, suggesting that despite its smaller Cl− conductance, the kinetics and binding affinity of EAATs are independent of the magnitude of their uncoupled Cl− conductance. Our finding also lends credence to the idea that different EAAT subtypes may play alternative roles in modulating synaptic activity.

Our study indicates that glutamate transporter currents are not blocked by DHKA in rods, suggesting that glutamate uptake in salamander rods is mediated by EAATs other than sEAAT2. Therefore, sEAAT2A appears to be cone specific, with respect to photoreceptor signalling. To understand why different subtypes of EAATs engage in glutamate uptake in rod and cone synapses, the functional differences between these two distinct types of photoreceptors should be considered. Cones have a significantly faster releasable pool of vesicles (Cadetti et al. 2008). We suggest that EAAT2A allows the cone terminal to provide a more efficient mechanism with which to buffer glutamate output, thereby suppressing over-activation of the postsynaptic cells and increasing contrast sensitivity in the cone system.

Although a previous immunocytochemical study indicates that sEAAT2A might also be present in Müller cells (Eliasof et al. 1998b), DHKA does not inhibit Müller cells (Barbour et al. 1991). Moreover, the processes of salamander Müller cells do not enter the invaginations of the cone terminal (Lasansky, 1973). Therefore, it appears that EAAT2A in Müller cells does not participate in DHKA-sensitive regulation of cone to bipolar cell synapses, especially after transient release. We agree with the perception from previous literature (Gaal et al. 1998; Furness et al. 2008) that glutamate uptake by Müller and glial cells is most likely to be a slow and inefficient process in regulating synaptic transmission. Vandenbranden et al. (2000) have previously reported EAAT2 localization far from the synaptic ribbon in goldfish (Carassius auratus) cones. We must note that in their study, tissues were fixed after a 3 h period of light adaptation. The localization of EAAT2 may be in closer proximity to synaptic ribbons in cones of the dark-adapted retina, as is implied in our study.

The postsynaptic transporter might be less important in control of cone synaptic gain

Molecular cloning studies have shown that EAAT2 exists in the salamander retina in two isoforms, sEAAT2A and sEAAT2B (Eliasof et al. 1998a), that are located at different pre- and postsynaptic sites in the OPL. Our results from axon-truncated bipolar cells suggest that postsynaptic sEAAT2B uptake in the OPL plays a less significant role compared with its presynaptic counterpart in cones. In addition, Fig. 4Ac and Bc depict statistical differences of glutamatergic currents in cone-dominant and rod-dominant Off-bipolar cells in the presence of DHKA. Since salamander rods appear to lack sEAAT2A, inhibition of sEAAT2A via DHKA causes more glutamate accumulation in the synaptic clefts of cone-to-Off-bipolar cells than in rod-to-Off bipolar cells. The small glutamatergic currents in rod-dominated Off-bipolar cells could be due either to the blockage of sEAAT2B by DHKA in the Off-bipolar cells dendrites, or to a lower glutamate receptor density in the dendrites of rod-dominated Off-bipolar cells. Since sEAAT2B elicits a smaller uptake-mediated Cl− conductance, we speculate that glutamate uptake via sEAA2B in Off-bipolar cell dendrites might cause a shift in membrane potential toward ECl. Although the dendritic EAAT2B current is small, sEAAT2B could theoretically act to counter the effect of glutamate receptors, as the ECl is likely to be negative to the glutamate reversal potential in Off-bipolar dendrites.

In further agreement with previous studies, our results also denote sEAAT2B as an excellent marker for Off-bipolar cells in the salamander retina. Immunocytochemical results show that sEAAT2B labels the entire Off-bipolar cell, including dendrites and axon terminals (Fig. 2). Although a strong antibody labelling of sEAAT2B is observed by immunocytochemistry, our electrophysiological results indicate that the transporter has a smaller overall Cl− conductance (Fig. 3) with respect to sEAAT2A in cones. Indeed, a previous study (Eliasof et al. 1998b) reported that no significant differences were observed between the overall conductances of sEAAT2A and sEAAT2B, which raises the possibility that most labelling in Off-bipolar cells represents an inactive form of sEAAT2B in the cytosol, rather than the active form expressed in the plasma membrane of cells.

Although sEAAT2B is the only neuronal-specific transporter found in salamander Off-bipolar cells, the function of sEAAT2B has yet to be determined. Nevertheless, there is a strong possibility that sEAAT2B acts as a presynaptic transporter in bipolar cell terminals, in much the same way that the sEAAT2A isoform acts presynaptically in cone terminals. Indeed, transient EAAT currents can be elicited from Off-bipolar cells following fast depolarizations (our unpublished observation). Therefore, sEAAT2B in Off-bipolar cell terminals may control glutamate uptake in the synapses of bipolar cells and third-order neurons. Since EAAT2 has been found as the sole glutamate transporter expressed in cone-linked Off-bipolar cells in mouse retinas (Hanna & Calkins, 2007), these and future results obtained from salamander retina should have general importance for higher vertebrates. Furthermore, since EAAT2 is a major transporter responsible for glutamate uptake in the CNS, using amphibian retina allows us to pursue the possible function of EAAT2 in pre- and postsynaptic locales. Thus, our findings on the roles of pre- and postsynaptic EAATs in the distal retina may provide a better understanding of the function of EAATs throughout the CNS where they may be of general importance in the control of fast synaptic signalling, and possibly involved in synaptic plasticity.

Acknowledgments

We thank Dr Susan Amara for the generous gift of the sEAAT2B antibody and Dr Baiqin Li for providing sEAAT2B antibody labelling data in Fig. 2. This work was supported by the National Eye Institute (grant EY 14161, W.S.).

Glossary

Abbreviations

DHKA

dihydrokainic acid

EAAT

excitatory amino acid transporter

sEAAT

excitatory amino acid transporter cloned from salamander

INL

inner nuclear layer

IPL

inner plexiform layer

ONL

outer nuclear layer

OPL

outer plexiform layer

TBOA

dl-threo-b-benzyloxyaspartic acid

Author contributions

M.J.R. and W.S. were involved in the conception and design of the experiments and collection and analysis of data; M.J.R. conducted the experiments; H.R. was involved in revising the article critically for intellectual contents and data analysis. The manuscript was read and revised by all authors. The experiments were conducted in the Department of Basic Science, Charles E. Schmidt College of Medicine at Florida Atlantic University.

References

  1. Arriza JL, Eliasof S, Kavanaugh MP, Amara SG. Excitatory amino acid transporter 5, a retinal glutamate transporter coupled to a chloride conductance. Proc Natl Acad Sci U S A. 1997;94:4155–4160. doi: 10.1073/pnas.94.8.4155. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Barbour B, Brew H, Attwell D. Electrogenic uptake of glutamate and aspartate into glial cells isolated from the salamander (Ambystoma) retina. J Physiol. 1991;436:169–193. doi: 10.1113/jphysiol.1991.sp018545. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Brew H, Attwell D. Electrogenic glutamate uptake is a major current carrier in the membrane of axolotl retinal glial cells. Nature. 1987;327:707–709. doi: 10.1038/327707a0. [DOI] [PubMed] [Google Scholar]
  4. Cadetti L, Bartoletti TM, Thoreson WB. Quantal mEPSCs and residual glutamate: how horizontal cell responses are shaped at the photoreceptor ribbon synapse. Eur J Neurosci. 2008;27:2575–2586. doi: 10.1111/j.1460-9568.2008.06226.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Cadetti L, Tranchina D, Thoreson WB. A comparison of release kinetics and glutamate receptor properties in shaping rod-cone differences in EPSC kinetics in the salamander retina. J Physiol. 2005;569:773–788. doi: 10.1113/jphysiol.2005.096545. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Danbolt NC. Glutamate uptake. Prog Neurobiol. 2001;65:1–105. doi: 10.1016/s0301-0082(00)00067-8. [DOI] [PubMed] [Google Scholar]
  7. Eliasof S, Werblin F. Characterization of the glutamate transporter in retinal cones of the tiger salamander. J Neurosci. 1993;13:402–441. doi: 10.1523/JNEUROSCI.13-01-00402.1993. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Eliasof S, Arriza JL, Leighton BH, Kavanaugh MP, Amara SG. Excitatory amino acid transporters of the salamander retina: identification, localization, and function. J Neurosci. 1998;18:698–712. doi: 10.1523/JNEUROSCI.18-02-00698.1998. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Furness DN, Dehnes Y, Akhtar AQ, Rossi DJ, Hamann M, Grutle NJ, Gundersen V, Holmseth S, Lehre KP, Ullensvang K, Wojewodzic M, Zhou Y, Attwell D, Danbolt NC. A quantitative assessment of glutamate uptake into hippocampal synaptic terminals and astrocytes: new insights into a neuronal role for excitatory amino acid transporter 2 (EAAT2) Neuroscience. 2008;157:80–94. doi: 10.1016/j.neuroscience.2008.08.043. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Gaal L, Roska B, Picaud SA, Wu SM, Marc R, Werblin FS. Postsynaptic response kinetics are controlled by a glutamate transporter at cone photoreceptors. J Neurophysiol. 1998;79:190–196. doi: 10.1152/jn.1998.79.1.190. [DOI] [PubMed] [Google Scholar]
  11. Grant GB, Werblin FS. A glutamate-elicited chloride current with transporter-like properties in rod photoreceptors of the tiger salamander. Vis Neurosci. 1996;13:135–144. doi: 10.1017/s0952523800007185. [DOI] [PubMed] [Google Scholar]
  12. Hanna MC, Calkins DJ. Expression of genes encoding glutamate receptors and transporters in rod and cone bipolar cells of the primate retina determined by single-cell polymerase chain reaction. Mol Vis. 2007;13:2194–2208. [PubMed] [Google Scholar]
  13. Hasegawa J, Obara T, Tanaka K, Tachibana M. High-density presynaptic transporters are required for glutamate removal from the first visual synapse. Neuron. 2006;50:63–74. doi: 10.1016/j.neuron.2006.02.022. [DOI] [PubMed] [Google Scholar]
  14. Innocenti B, Heidelberger R. Mechanisms contributing to tonic release at the cone photoreceptor ribbon synapse. J Neurophysiol. 2008;99:25–36. doi: 10.1152/jn.00737.2007. [DOI] [PubMed] [Google Scholar]
  15. Jackman SL, Choi SY, Thoreson WB, Rabl K, Bartoletti TM, Kramer RM. Role of the synaptic ribbon in transmitting the cone light response. Nat Neurosci. 2009;12:303–310. doi: 10.1038/nn.2267. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Jackson CR, Chaurasia SS, Zhou H, Haque R, Storm DR, Iuvone PM. Essential roles of dopamine D4 receptors and the type 1 adenylyl cyclase in photic control of cyclic AMP in photoreceptor cells. J Neurochem. 2009;109:148–157. doi: 10.1111/j.1471-4159.2009.05920.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Lasansky A. Organization of the outer synaptic layer in the retina of the larval tiger salamander. Philos Trans R Soc Lond B Biol Sci. 1973;265:472–789. doi: 10.1098/rstb.1973.0033. [DOI] [PubMed] [Google Scholar]
  18. Levy LM, Warr O, Attwell D. Stoichiometry of the glial glutamate transporter GLT-1 expressed inducibly in a Chinese hamster ovary cell line selected for low endogenous Na+-dependent glutamate uptake. J Neurosci. 1998;18:9620–9628. doi: 10.1523/JNEUROSCI.18-23-09620.1998. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Maple BR, Zhang J, Pang JJ, Gao F, Wu SM. Characterization of displaced bipolar cells in the tiger salamander retina. Vision Res. 2005;45:697–705. doi: 10.1016/j.visres.2004.09.038. [DOI] [PubMed] [Google Scholar]
  20. Otis TS, Jahr CE. Anion currents and predicted glutamate flux through a neuronal glutamate transporter. J Neurosci. 1998;18:7099–7110. doi: 10.1523/JNEUROSCI.18-18-07099.1998. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Palmer MJ, Taschenberger H, Hull C, Tremere L, von Gersdoff H. Synaptic activation of presynaptic glutamate transporter currents in nerve terminals. J Neurosci. 2003;23:4831–4841. doi: 10.1523/JNEUROSCI.23-12-04831.2003. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Pang JJ, Gao F, Wu SM. Stratum-by-stratum projection of light response attributes by retinal bipolar cells of Ambystoma. J Physiol. 2004;558:249–262. doi: 10.1113/jphysiol.2004.063503. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Picaud SA, Larsson HP, Grant GB, Lecar H, Werblin FS. Glutamate-gated chloride channel with glutamate-transporter-like properties in cone photoreceptors of the tiger salamander. J Neurophysiol. 1995;74:1760–1771. doi: 10.1152/jn.1995.74.4.1760. [DOI] [PubMed] [Google Scholar]
  24. Rabl K, Cadetti L, Thoreson WB. Kinetics of exocytosis is faster in cones than in rods. J Neurosci. 2005;25:4633–4640. doi: 10.1523/JNEUROSCI.4298-04.2005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Rauen T, Wiessner M, Sullivan R, Lee A, Pow DV. A new GLT1 splice variant: cloning and immunolocalization of GLT1c in the mammalian retina and brain. Neurochem Int. 2004;45:1095–1106. doi: 10.1016/j.neuint.2004.04.006. [DOI] [PubMed] [Google Scholar]
  26. Roska B, Gaal L, Werblin FS. Voltage-dependent uptake is a major determinant of glutamate concentration at the cone synapse: an analytical study. J Neurophysiol. 1998;80:1951–1960. doi: 10.1152/jn.1998.80.4.1951. [DOI] [PubMed] [Google Scholar]
  27. Schnapf JL, Copenhagen DR. Differences in the kinetics of rod and cone synaptic transmission. Nature. 1982;296:862–864. doi: 10.1038/296862a0. [DOI] [PubMed] [Google Scholar]
  28. Thoreson WB, Rabl K, Townes-Anderson E, Heidelberger R. A highly Ca2+-sensitive pool of vesicles contributes to linearity at the rod photoreceptor ribbon synapse. Neuron. 2004;42:595–560. doi: 10.1016/s0896-6273(04)00254-5. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Thoreson WB, Stella SL, Jr, Bryson EI, Clements J, Witkovsky P. D2-like dopamine receptors promote interactions between calcium and chloride channels that diminish rod synaptic transfer in the salamander retina. Vis Neurosci. 2002;19:235–247. doi: 10.1017/s0952523802192017. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Vandenbranden CA, Yazulla S, Studholme KM, Kamphuis W, Kamermans M. Immunocytochemical localization of the glutamate transporter GLT-1 in goldfish (Carassius auratus) retina. J Comp Neurol. 2000;423:440–451. doi: 10.1002/1096-9861(20000731)423:3<440::aid-cne7>3.0.co;2-7. [DOI] [PubMed] [Google Scholar]
  31. Veruki ML, Mørkve SH, Hartveit E. Activation of a presynaptic glutamate transporter regulates synaptic transmission through electrical signaling. Nat Neurosci. 2006;9:1388–1396. doi: 10.1038/nn1793. [DOI] [PubMed] [Google Scholar]
  32. Wu SM. The off-overshoot responses of photoreceptors and horizontal cells in the light adapted retinas of the tiger salamander. Exp Eye Res. 1988;47:261–268. doi: 10.1016/0014-4835(88)90009-7. [DOI] [PubMed] [Google Scholar]
  33. Wersinger E, Schwab Y, Sahel JA, Rendon A, Pow DV, Picaud S, Roux MJ. The glutamate transporter EAAT5 works as a presynaptic receptor in mouse rod bipolar cells. J Physiol. 2006;577:221–234. doi: 10.1113/jphysiol.2006.118281. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Yang J, Wu SM. Characterization of glutamate transporter function in the tiger salamander retina. Vision Research. 1997;37:827–838. doi: 10.1016/s0042-6989(96)00231-3. [DOI] [PubMed] [Google Scholar]

Articles from The Journal of Physiology are provided here courtesy of The Physiological Society

RESOURCES