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1Department of Biological Sciences, Louisiana State University, Baton Rouge, Louisiana; and 2West Virginia University School of Medicine, Morgantown, West Virginia
Submitted 14 December 2005; accepted in final form 30 January 2006
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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; Taylor and Vaney 2003), so understanding the function and regulation of amacrine cells is critical to understanding retinal signal processing.
Nitric oxide (NO) has been implicated as a signaling molecule at all levels of the visual system (for review, see Cudeiro and Rivadulla 1999). Nitric oxide synthase (NOS) catalyzes the production of NO and citrulline from arginine. The neuronal form of this enzyme (nNOS) is constitutively expressed and stimulated by elevations of cytosolic Ca2+. In the canonical pathway, NO stimulates soluble guanylate cyclase (sGC) activity. NO is also known to affect target protein function through S-nitrosylation (Davis et al. 2001; Stamler et al. 1997) and by other direct interactions including formation of dinitrosyl iron complexes (Rogers et al. 2003).
Neuronal NOS has been found in subsets of each class of neuron in the vertebrate retina (Blute et al. 1997; Chun et al. 1999; Fischer and Stell 1999; Kim et al. 1999; Rios et al. 2000; Shin et al. 1999), and NO itself has been detected in a similar array of retinal cells (Blute et al. 2000, 2003; Neal et al. 1998). In the chicken retina, a subset of amacrine cells expresses high levels of nNOS, and distinct sublaminae of nNOS labeling occur throughout the inner plexiform layer of the retina (Crousillac et al. 2003; Fischer and Stell 1999; Rios et al. 2000). Furthermore, measurements at the inner surface of the retina indicate that this tissue can generate high (micromolar) levels of NO (Donati et al. 1995; Groppe et al. 2003; Heiduschka and Thanos 1998).
The physiological effects of NO production have been examined for some cell types that signal in the inner retina. In the rabbit retina, NO reduces gap-junction coupling between AII amacrine cells and cone bipolar cells (Mills and Massey 1995). Wexler and colleagues (1998) have shown that NO depresses GABAA receptor function in cultured rat amacrine cells. In the ferret retina, recordings of ganglion cell activity reveal a dramatic NO-dependent decrease in light responses (Wang et al. 2003). Although effects of NO have been identified in multiple retinal cell types, the full range of NO function in the inner retina remains unknown.
To further examine how NO influences amacrine cell signaling, whole cell recordings were made from cultured GABAergic amacrine cells and amacrine cell synaptic pairs derived from embryonic chick retinae. We show that NO alters amacrine cell activity by affecting the functional properties of GABAA receptors and, much more dramatically, by converting inhibitory synapses to excitatory synapses through a GABA receptor-independent mechanism. This mechanism involves an NO-induced redistribution of Cl–. Our results indicate that the redistribution of Cl– does not stem from Cl– moving across the plasma membrane but is instead due to release of Cl– from an internal compartment.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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Retinal cultures were prepared from 8-day-old chick embryos and maintained as previously described (Hoffpauir and Gleason 2002). Experiments were performed on isolated amacrine cells or isolated pairs of amacrine cells 6–14 days (EE 14 to PE 1) after plating. After 6 days in culture, these cells are considered to be mature, based on several physiological criteria including expression of the appropriate voltage- and ligand-gated ion channels (Huba and Hofmann 1991; Huba et al. 1992) and the formation of functional GABAergic synapses with other GABAergic amacrine cells. The properties of the amacrine-to-amacrine cell GABAergic synapses formed in culture have been previously characterized (Gleason et al. 1993). The uniformity of neurotransmitter phenotype indicates that the amacrine cells in culture represent a subset of amacrine cell phenotypes found in the intact retina.
Hippocampal cultures were made from 18-day-old rat (Fischer 344) embryos. Micro-dissected hippocampal tissue was obtained from Neuromics (Bloomington, MN) and prepared and maintained according to the supplier's protocols. Recordings from single hippocampal neurons were made after 4–7 days in culture. No physiological assessments of the relative maturity of these cells have been made. According to their developmental timeline alone, they would be considered embryonic (EE 22–EE 25).
Electrophysiology
Whole cell recordings were made using an Axopatch 1-D amplifier, Digidata 1322A data-acquisition board, and Clampex 9.2 software (Axon Instruments, Union City, CA). A reference Ag/AgCl pellet in 3 M KCl was connected to the culture dish via an agar bridge containing 3 M KCl. Patch electrodes were pulled from thick-walled borosilicate glass (1.5 mm OD, 0.86 mm ID; Sutter Instruments, Novato, CA) using a P-97 Flaming/Brown Puller (Sutter Instruments). Tip resistance values were 5–10 M
for ruptured-patch recordings and 3–5 M for perforated-patch recordings as measured in the bath. All recordings were made at room temperature (22–24°C) and were corrected for junction potential error. Voltage ramp data were leak-subtracted. Because we used both perforated- and ruptured-patch to study the NO-shift in EGABA (see RESULTS), we wanted to determine whether the recording configuration had an effect on the amplitude of the shift. To achieve this, we recorded NO-induced shifts in EGABA on 1 day with one batch of NO solution. For these experiments, we alternated between the two recording configurations and found no significant differences in the amplitude of the shifts (n = 3 in each configuration; P = 0.999).
Solutions
Unless otherwise indicated, all reagents were purchased from Sigma, St. Louis, MO. GABA application was achieved in 10–20 ms through computer-controlled perfusion barrel movements. This method was also used to switch between normal and zero external Cl– for the experiment depicted in Fig. 8A. All other solution changes were achieved in 
500 ms by opening and closing the valves upstream of a manifold feeding one common barrel. The compositions of external and internal solutions are shown in Tables 1 and 2, respectively. TTX (300 nM) and LaCl3 (25–50 µM) were added to external solutions (with the exception of the 0 Cl– external solution) for single-cell voltage clamp (but not current clamp) recordings to block voltage-gated Na+ and Ca2+ currents, respectively. For ruptured-patch recordings, the following reagents were added to internal solutions (Cs+-A, unless otherwise indicated): 50 U/ml creatine phosphokinase, 3 mM adenosine 5'-triphosphate (ATP) dipotassium salt, 1 mM ATP-disodium salt, 20 mM phosphocreatine (Calbiochem, La Jolla, CA), 2 mM guanosine 5'-triphosphate (GTP) sodium salt. For perforated-patch recordings, either amphotericin B (synaptic pairs, Fig. 4) or gramicidin (Figs. 5 and 6) were added to normal internal or Cs+ internal, respectively, to a final concentration of 10 µg/ml.
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NO-bubbled solutions
Pure NO solutions were prepared by bubbling solutions with argon for 15 min followed by 15 min of bubbling with pure soda lime-filtered NO. NO solutions were tightly sealed, protected from light, and stored at 4°C. Although the method for preparing NO was standardized, the efficacy of different batches was variable. For most experiments, injections between 10 and 50 µl were able to elicit shifts of 15–35 mV. When standard volumes of a batch of NO solution became less efficient, a new batch was obtained. The NO solutions were delivered into the perfusion system manually with a Hamilton syringe via in-line injections (10–50 µl depending on potency, see following text). In-line injection of dye-containing solutions indicated that NO exposure was limited to 1–3 s with most of the NO arriving in the first 500 ms. NO was bubbled into either H2O or TEA external. No differences in the effects of NO in the two solutions were observed. For synaptic recordings, NO was bubbled into a 40 mM HEPES solution to minimize low pH effects on Ca2+ channels and other synaptic proteins. The concentration of NO in these solutions was measured using an ISO-NO meter (NOMK2 system) with an ISO-NOP electrode (World Precision Instrument, Sarasota, FL) for a batch and volume that was demonstrated to be effective in eliciting cellular responses. The mean NO concentration measured at the perfusion outlet was 2.0 ± 0.3 µM. This value represents the upper limit of NO coming in contact with a cell. For SNAP solutions, 100 nM NO was detected in external solution containing 250 µM SNAP (2 h after preparation). With the exception of Fig. 1, the data shown were obtained using NO-bubbled solutions.
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10–30 min. For both fresh and air-exposed NO solutions, pH values typically ranged between 2 and 3. Injections of the appropriate amount of pH 2.5, NO-free solutions were routinely used as controls and none of the effects reported here were reproduced by low pH alone. Although extreme care has been taken to standardize the preparation, handling and injections of these volatile solutions, some variability remains. As such, only those cells receiving the same volumes, from the same batch of NO-bubbled solutions were used for quantitative comparisons. Data are reported as means ± SD and statistical significance was determined using the t-test. Data shown in Figs. 5, B and D, and 6F were analyzed using the paired t-test. |
RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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To explore the effects of NO, we recorded whole cell GABAA receptor currents (Hoffpauir and Gleason 2002) from individual amacrine cells. For these experiments, Cs+-A internal and TEA-A external solutions were used, and recordings were made in the ruptured-patch recording configuration. Under these conditions, we find that NO produces three effects in amacrine cells. First, moderate NO donor concentrations (250 µM SNAP, 100 nM NO, see METHODS) caused a small (15%) enhancement of the GABA-gated current amplitude that was not due to alteration in the reversal potential of the current (EGABA, Fig. 1). Similar effects were observed with NOC 12 (250 µM) and NOC 18 (300 µM, not shown). The SNAP-dependent enhancement was significantly inhibited in the presence of the NO scavenger carboxy-PTIO, indicating that NO rather than donor end products underlie the GABAA current enhancement (Fig. 1C, P = 0.002).
Second, higher concentrations of NO (hundreds of nanomolar to low micromolar, NO-bubbled solutions, see METHODS) produced a transient (1–3 s), GABA-independent inward current (Figs. 2, A and C, and 7Bi, asterisks). Based on reversal potential measurements, this NO-dependent, GABA-independent, inward current is a cation current (Erev = +23.4 ± 6.6mV, n = 5).
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). Note that the time course of recovery for the (higher concentration) NO-induced current enhancement (Fig. 2A) is consistent with the recovery time course for EGABA (Fig. 2E). Similar shifts in EGABA were obtained with high concentrations (2 mM) of the NO donor NOC 12 (not shown) indicating that the mechanism responsible for the shift in EGABA can be activated by either NO delivery method as long as the concentration of NO is in the correct concentration range. A small increase in the slope of the I-V relationship for the GABA-gated current (1st NO effect) was also detectable with higher NO concentrations (Fig. 2D, inset). The role of soluble guanylate cyclase (sGC) activation in the NO-induced shift in EGABA was investigated using the sGC inhibitor ODQ. Prolonged (>10 min) preincubation with the inhibitor did not block the NO-induced shift in EGABA indicating that sGC activity is not involved in this mechanism (Fig. 2, F and G). The nonsignificant trend toward larger responses in ODQ suggests that basal sGC activity might have a suppressive effect on the mechanism underlying the shift in EGABA. Finally, the NO-dependent current (2nd NO effect) may be related to the shift in EGABA but does not seem to be an absolute requirement because we have observed the shift in the absence of this current. NO-induced shift in EGABA is due an elevation in cytosolic Cl–
Three possible explanations for the NO-induced shift in EGABA were examined: a change in ion selectivity of the GABAA receptors, the introduction of another permeant anion into the cell, or an increase in intracellular Cl– concentration. To determine whether NO increases the selectivity of GABAA receptors to acetate (the only other anion in the internal solution), internal acetate was substituted with methanesulfonate, a bulky anion unlikely to permeate the receptors. This substitution, however, did not prevent the NO-induced shift in EGABA as indicated by the several-fold enhancement of the GABA-gated current (Fig. 3A). This suggests that the NO-induced shift in EGABA is not due to a change in the ion selectivity of the channels.
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; Bormann et al. 1987), it is possible that trapped cytosolic NO3– (from inwardly diffusing NO) contributes to the shift in EGABA. Furthermore, although the recording solutions are HCO3– -free, it is also possible that NO transiently stimulates the production of HCO3–, another anion that can permeate GABAA receptors (Bormann et al. 1987). In zero Cl– internal and zero Cl– external solutions, no GABA-gated currents were observed either before or after application of NO (Fig. 3B). This argues against the possibility that the shift is due to an increase in intracellular NO3–, or HCO3–, or any other permeant anion.
If the effect of NO is to raise internal Cl–, then a similar shift in reversal potential should also be observed when glycine receptors are activated. Agonist-gated currents were elicited with pulses of 20 µM glycine or 20 µM GABA delivered during voltage ramps. Addition of NO caused a shift in the reversal potential to more positive values for both agonists (Fig. 3C; GABA shift: 33.6 ± 11.5 mV; glycine shift: 30.0 ± 14.0 mV; P = 0.67; n = 5), indicating that NO is stimulating a redistribution of Cl– in these cells. An increase in the slope of the glycine-gated current was also observed, indicating an enhancing effect of NO on glycine receptor function. The ability to change the internal Cl– concentration in the face of diffusion from the pipette may be unexpected, but it is not unprecedented. Cl– transport has been shown to be effective in opposing diffusion from the recording pipette (Staley et al. 1996) and changes in cytosolic Cl– have been measured by optical methods in the ruptured patch recording configuration (Isomura et al. 2003). It is also possible that the redistribution of Cl– is not a global event, but one that occurs locally, possibly in neuronal processes.
Current-clamp recordings were made to determine how the NO-induced shift in ECl– affects GABA- and glycine-dependent changes in membrane potential (Fig. 3D). Resting membrane potentials were –60.1 ± 13.2 mV (n = 5). Under control conditions, application of GABA or glycine hyperpolarized these cells to –70.9 ± 2.7 and –68.7 ± 5.3 mV, respectively. For the first agonist pulse after NO application, GABA depolarized the cells to –31.8 ± 25.2 mV and glycine depolarized the cells to –34.4 ± 18.4 mV. One cell responded to both GABA (not shown) and glycine (Fig. 3D) with a single action potential followed by a steady depolarization. This is the typical voltage response for these cells given sufficient depolarization (Gleason et al. 1993). It is important to note, however, that action potentials are not required for Ca2+-dependent neurotransmitter release from these cells (Gleason et al. 1993). These current-clamp recordings demonstrate that NO can change the sign of whole–cell voltage responses to GABA and glycine.
NO-induced shift in ECl– occurs at synapses
To examine the influence of NO specifically at retinal amacrine cell synapses, dual perforated-patch recordings were made from pairs of cultured GABAergic amacrine cells. It is well established that GABAergic amacrine cell synapses are normally inhibitory, both in the retina (Lagnado 1998) and in these cultures (Gleason et al. 1993). Recordings from isolated pairs of amacrine cells demonstrate that brief exposure to NO can transiently change the sign of GABAergic synapses (Fig. 4). With both pre- and postsynaptic cells in the voltage-clamp configuration, a depolarizing voltage step in the presynaptic cell produced a small outward current in the postsynaptic cell. After NO, postsynaptic currents become inward indicating an excitatory effect on the postsynaptic cell (Fig. 4A, gray trace). The relatively small, noisy postsynaptic currents are consistent with the low release rates described for these synapses in culture (Gleason et al. 1993) and for GABAergic amacrine cell synapses in the intact retina (Zheng et al. 2004). When the postsynaptic cell is switched to current clamp, the postsynaptic response is barely detectable because under control conditions, the resting potential of the cell (–67 mV) is very near the calculated ECl– (–70 mV). After NO, the postsynaptic voltage response is depolarizing (Fig. 4B, gray trace). NO-dependent, excitatory synaptic responses were detected in all pairs examined (n = 5). Postsynaptic responses to subsequent presynaptic depolarizations were omitted from Fig. 4 for clarity, but they show that recovery from the NO-dependent shift in ECl– occurs over a similar time frame as observed in recordings of whole cell GABA-gated currents in single amacrine cells (see Fig. 2, A and E). It is possible that NO has other, as yet uncharacterized effects on amacrine cell synapses (Ahern et al. 2002; Hölscher 1997), but the key observation here is that the shift in ECl– found in whole cell recordings also occurs at synapses.
NKCC does not mediate the increase in cytosolic Cl-
What mechanism underlies the increase in cytosolic Cl–? The plasma membrane Na+-K+-Cl– co-transporter (NKCC) and the K+-Cl– co-transporter (KCC2) are Cl– co-transport mechanisms known to regulate the distribution of Cl– across neuronal plasma membranes (Kakazu et al. 2000; Russell 2000). It is now well established that changes in Cl– co-transporter expression determine the effects of GABA during development (for review, see Payne et al. 2003). In the adult retina, ECl– varies among cell types and subcellular locations, and this correlates with the expression pattern of NKCC and KCC2 (Vardi et al. 2000). Furthermore, the response properties of retinal starburst amacrine cells are dependent on the activity of NKCC and KCC2 (Gavrikov et al. 2003). Given the established role of these transporters in determining ECl–, we investigated the role of NKCC and KCC2 in the NO-induced shift in ECl–.
NKCC typically transports 2 Cl–, 1 Na+, and 1 K+ into the cell and the neuron specific K+-Cl– co-transporter, KCC2, transports 1 Cl– and 1 K+ out of the cell. Thus the NO-induced shift in EGABA could be due to an increase in NKCC activity or a decrease in KCC2 activity. To determine if these mechanisms were involved, perforated-patch recordings were made with Cl–-impermeant gramicidin in the pipette and the co-transport inhibitors, bumetanide (300 µM) and furosemide (300 µM) were used to block NKCC and KCC2 activity. Both inhibitors shifted ECl– to more negative potentials, indicating a dominant inhibitory effect on NKCC (Fig. 5, A and C). Neither blocker, however, inhibited the NO-induced shift in ECl– (Fig. 5, B and D). To asses the variability of responses under control conditions, the effects of two separate NO applications were also examined and we found no significant difference between the two trials (P = 0.69, n = 5, not shown). At lower concentrations (10 µM), bumetanide is selective for NKCC (Russell 2000). In similar experiments, bumetanide at 10 µM was also ineffective in blocking the shift in ECl– (not shown).
Removal of the externally required co-transported ions shifted ECl– to more negative potentials. Furthermore, for those cells that were exposed to both zero K+ and zero Na+ solutions, ECl– was considerably more negative in zero K+ solutions than in zero Na+ solutions (Fig. 6A). In some cases, switching to zero K+ shifted ECl– by tens of millivolts in as few as 15 s (Fig. 6B). These results are consistent with the prediction that zero external K+ would prevent inward Cl– transport through NKCC and promote outward Cl– transport through KCC2. Despite these effects, none of these manipulations blocked the NO-induced shift in ECl– (Fig. 6, C–E). Figure 6F shows that removal of co-transported ions did not cause a statistically significant (P = 0.26; 0.17) change in the magnitude of the NO-induced shift in ECl–. Interestingly, a nonsignificant trend toward larger shifts was observed in the absence of co-transported ions. This may relate to an enhanced Cl– gradient between the cytosol and an internal compartment (see following text). The roles of two other known plasma membrane Cl– transport mechanisms have also been investigated. Neither 4,4'-diisothiocyanatostilbene-2,2'-disulfonic acid (DIDS, 40 µM, n = 5) a HCO3/Cl– transport inhibitor, nor ethacrynic acid (10 µM, n = 4), an inhibitor of ATP-dependent, Na+-independent Cl– transport, blocked the shift in ECl– (not shown).
Shift in ECl– is independent of extracellular Cl-
The lack of involvement of known plasma membrane Cl– transporters indicates that NO is either stimulating an unexamined Cl– transport mechanism or releasing Cl– from inside the cell. Endosomal compartments are known to maintain relatively high Cl– concentrations to offset the membrane potentials generated by inward proton pumping and endosomal acidification (Faundez and Hartzell 2004; Sonowane and Verkman 2003). To determine whether NO was releasing Cl– from an internal compartment, external Cl– was reduced to match the internal Cl– concentration (14.2 mM). NO was applied while cells were held at the reversal potential for Cl–. Under these conditions, subsequent records collected during pulses of GABA demonstrated that the NO-induced redistribution of Cl– persisted in the absence of an electrochemical gradient for chloride across the plasma membrane (Fig. 7A, see also Fig. 2C). To determine whether external Cl– was required at all, NO was also applied in the absence of external Cl–. For these experiments (Fig. 7B), control ramps (Bi) and the NO applications (Bii) were conducted in zero Cl– external solution. The arrival of NO in zero Cl– was marked by the presence of the NO-dependent inward current (asterisk). After switching to normal Cl– external solution (TEA-A), so that the reversal potential of the current could be measured, another set of ramps was delivered (Biii). Values for the reversal potentials of the GABA-gated currents after NO application (mean shift: 26.6 ± 9.9 mV, n = 9) indicate that an increase in internal Cl– occurred in the absence of external Cl–. Control experiments were performed in the same manner but without NO application (Fig. 7, D and E). Only slight fluctuations (1–2 mV) in reversal potential resulted from just switching between 0 Cl– external and normal external solutions.
The switch back to normal external Cl– concentration before measuring the GABA-gated current reversal potentials leaves open the possibility that rapid Cl– influx could occur just prior to measuring the reversal potential. To avoid this possibility, we also examined the large NO-induced change in the amplitude of the GABA-gated currents at –70 mV that results from the positive shift in reversal potential. In this way, the effects of NO can be evaluated without re-introducing external Cl–. In this experiment, both NO and GABA are applied in zero external Cl–. Substantial increases (2.9 ± 0.7-fold) in the GABA-gated current amplitude were observed in all cells tested (n = 5; Fig. 8A ). Because external Cl– is absent for both the NO injection and the enhancement of the GABA-gated current, the Cl– must be coming from the inside of the cell. Why, in the absence of internal and external Cl– (Fig. 3B), did the NO-induced release of Cl– from internal stores not produce an inward GABA-gated current? It is plausible that under Cl–-free conditions, the Cl– store becomes depleted. Clearly, much remains to be understood about the dynamics of intracellular Cl– fluxes.
Aside from the removal of both internal and external Cl–, the only manipulation that blocked the NO-induced shift in Cl– distribution was to raise the internal Cl– concentration to 114 mM. Under these conditions, application of NO produced virtually no shift in the reversal potential of the GABA-gated currents (mean shift = 0.5 ± 0.2 mV, n = 5, Fig. 8B). This result is consistent with internal Cl– release if the 114 mM internal Cl– reduces the gradient for Cl– between an internal Cl– compartment and the cytoplasm.
NO-induced shift in ECl– is not confined to the avian retina
Does the NO-induced shift in Cl– distribution rely on an amacrine cell-specific mechanism or is the underlying mechanism more widely expressed? To explore this, we made similar recordings from cultured rat hippocampal neurons. We find that NO also induces a shift in ECl– in hippocampal neurons (Fig. 9A; mean shift 38 ± 19.6 mV, n = 6). An inward GABA-independent/NO-dependent current was also observed in hippocampal neurons (Fig. 9C, asterisk). To determine whether the shift in ECl– was also due to internal release of Cl–, we repeated the experiment shown in Fig. 7B. Removal of extracellular Cl– during NO application did not inhibit the shift in ECl–, suggesting that, as for amacrine cells, NO stimulates the release of Cl– from an internal store in hippocampal neurons (Fig. 9D).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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100 nM) release of NO from donors enhances GABA-gated currents by modifying receptor activity. Brief pulses of higher concentrations of NO (hundreds of nanomolar to low micromolar, NO-bubbled solutions) produce sGC-independent increases in intracellular Cl– and shift EGABA to more positive potentials. Furthermore, we show that the shift in the Cl– reversal potential occurs at synaptic sites and is sufficient to promote excitation at GABAergic synapses. Finally, our results indicate that the redistribution of Cl– underlying this change results from a release of Cl– from an internal compartment. Role of NO in the retina
Measurements indicate that the concentration of NO at the cell in NO-bubbled solutions is in the hundreds of nanomolar to low micromolar range. Is this range of concentrations relevant to the levels of NO generated in the retina? NO electrode measurements at the inner retinal surface yield values ranging from 6 to 15 µM (Donati et al. 1995; Groppe et al. 2003). Consistent with the electrode measurements, a biochemical assay (the Griess method) also shows retinal NO production well into the micromolar range (Heiduschka and Thanos 1998). These intraocular measurements indicate that NO concentrations can be substantial and that the doses of NO used in our experiments are within the physiological range.
Recently, NO electrode measurements have been made near the surface of an individual cell in the ganglion cell layer of the turtle retina (Eldred and Blute 2005). An NO concentration of 200 nM was detected. It is important to note, however, that imaging of NMDA-stimulated NO production in the same retina has demonstrated that the NO signals generated are often highly localized to individual cells and even discrete boutons in the inner synaptic layer (Blute et al. 2000). This observation suggests that some of the downstream effects of NO signaling occur primarily in nNOS-expressing cells and their immediate synaptic partners, where NO concentrations would be higher.
How might the NO-induced changes we show for cultured amacrine cells affect GABAergic signaling between amacrine cells in the inner retina? The diversity of synaptic partners and complex synaptic arrangements of amacrine cells in the IPL (Dowling and Boycott 1965; Hartveit 1999; Kolb 1997; Marc and Liu 2000) complicate this issue. The cartoon in Fig. 10 depicts a simplified subset of interactions in the inner retina. The cell on the left is a GABAergic or glycinergic, nNOS-expressing amacrine cell that receives a mixture of excitatory (glutamatergic, from bipolar cells) and inhibitory (GABA- and/or glycinergic, from other amacrine cells) inputs. The net effect of these inputs is to determine the strength of the inhibitory output from the cell onto its postsynaptic partners (ganglion cells, bipolar cells, and/or amacrine cells). If the cell is exposed to a relatively high concentration of NO (presumably via its own nNOS activity), the distribution of chloride shifts in that amacrine cell so the balance of its input becomes more excitatory and the inhibitory output of this cell is enhanced. Consistent with this proposal, Wang et al. (2003) demonstrate an NO-dependent suppression of ganglion cell light responses that is due to alterations in presynaptic (probably amacrine cell) input. This observation would be consistent with an NO-dependent shift in the amacrine cell ECl– that results in an enhancement of inhibitory output onto ganglion cells.
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Our results indicate that the NO-induced redistribution of Cl– is due to Cl– release from inside the cell. Endosomal compartments are a potential source of internal Cl–. Cl– is thought to enter these compartments as a counter ion for protons being pumped in by vacuolar proton pumps (Sonawane and Verkman 2003). Movement of Cl– across endosomal membranes is thought to be mediated by the CLC family of Cl– transport proteins. ClC3-7 are found on intracellular membranes (for review, see Faundez and Hartzell 2004; Jentsch et al. 2002). Of these, ClC3 may be the most relevant because it is highly expressed in the brain and has been shown to be expressed on endosomes, including synaptic vesicles. Interestingly, knockouts of the CIC3 gene resulted in marked tissue degeneration specifically in the hippocampus and the retina (Stobrawa et al. 2001). Although physiological evidence for a role for CLC transport proteins in the NO-induced shift in ECl– awaits further experimentation, we have found that a polyclonal antibody raised against a conserved peptide from ClC3 gives an intense and punctuate labeling pattern that appears to be located intracellularly (McMains and Gleason, unpublished observations). It may be that the efflux of Cl– from endosomes via ClC3 contributes to the NO-induced shifts in EGABA and Eglycine and the transitions between inhibition and excitation.
Interestingly, NO has been shown to reversibly inhibit the vacuolar proton pump through S-nitrosylation (Forgac 1999). Although the mechanism by which NO stimulates Cl– efflux is not yet known, given the established relationship between proton pumping and Cl– movement across endosomal membranes, it is plausible that proton flux is somehow involved. However the Cl– efflux is achieved, the demonstration that it does occur and that it may be a broadly expressed mechanism is sure to impact our view of GABA and glycine-dependent synaptic signaling in the CNS.
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ACKNOWLEDGMENTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Address for reprint requests and other correspondence: E. Gleason, Dept. of Biological Sciences, Rm. 202 Life Sciences Building, Louisiana State University, Baton Rouge, LA 70803 (E-mail: egleaso{at}lsu.edu)
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