June 2002
Volume 43, Issue 6
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Glaucoma  |   June 2002
Protective Effect of Arachidonic Acid on Glutamate Neurotoxicity in Rat Retinal Ganglion Cells
Author Affiliations
  • Atsushi Kawasaki
    From the Departments of Ophthalmology and Visual Science and
  • Ming-Hu Han
    From the Departments of Ophthalmology and Visual Science and
  • Ji-Ye Wei
    From the Departments of Ophthalmology and Visual Science and
  • Keiji Hirata
    Cell Biology, Yale University School of Medicine, New Haven, Connecticut.
  • Yasumasa Otori
    From the Departments of Ophthalmology and Visual Science and
  • Colin J. Barnstable
    From the Departments of Ophthalmology and Visual Science and
Investigative Ophthalmology & Visual Science June 2002, Vol.43, 1835-1842. doi:
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      Atsushi Kawasaki, Ming-Hu Han, Ji-Ye Wei, Keiji Hirata, Yasumasa Otori, Colin J. Barnstable; Protective Effect of Arachidonic Acid on Glutamate Neurotoxicity in Rat Retinal Ganglion Cells. Invest. Ophthalmol. Vis. Sci. 2002;43(6):1835-1842.

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      © ARVO (1962-2015); The Authors (2016-present)

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Abstract

purpose. Low concentrations of excitotoxic agents such as glutamate decrease survival of retinal ganglion cells (RGCs) and may be an important cause of RGC death in a variety of retinal diseases. Arachidonic acid (AA), an intercellular messenger in the central nervous system, has been reported to have multiple effects on glutamate receptors, including an inhibitory effect on non-N-methyl-d-aspartate (NMDA) receptors. The purpose of this study was to test the hypothesis that AA could protect RGCs from glutamate neurotoxicity.

methods. RGCs were purified from the rat retina on postnatal days 7 and 8 by a modified two-step panning method. Survival of RGCs after exposure to glutamate, with or without AA treatment, was measured after 3 days in culture. To visualize calcium signals, RGCs were loaded with a calcium indicator dye, fluo-3 acetoxymethyl ester, and the fluorescence was measured by laser scanning confocal microscopy. Electrophysiological effects of AA on non-NMDA ionotropic receptors were examined by using whole-cell patch clamp configurations.

results. Incubation of RGCs with 25 μM glutamate caused 60% loss of RGCs. This glutamate neurotoxicity was significantly ameliorated by low concentrations of AA. Concentrations of AA above 10 μM were toxic to RGCs. Calcium imaging showed that glutamate-, α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid- (AMPA) and kainate-induced intracellular calcium accumulation in these cells was reduced by AA. Electrophysiological recordings revealed that currents mediated by non-NMDA ionotropic receptors were inhibited by AA in a dose-dependent manner.

conclusions. Low concentrations of AA can reduce glutamate neurotoxicity to RGCs by the inhibition of non-NMDA ionotropic receptors. These results suggest that endogenous or exogenous AA may be used to protect RGCs from glutamate neurotoxicity and that AA may be one potential treatment for RGC loss in a variety of eye diseases, including glaucoma.

Retinal ganglion cell (RGC) death is the unifying feature of all diseases of the optic nerve, including glaucomatous optic neuropathy. Glaucoma is one of the most common causes of blindness and is associated with certain risk factors, such as high intraocular pressure or blood-flow dysregulation. Nevertheless, many cases of glaucoma, especially normotensive glaucoma, do not correlate with these risk factors. The mechanisms mediating RGC death in these cases of glaucoma are not well known, although apoptosis is thought to be an important final step. 1 There may be more than one pathway leading to RGC death, but much attention has been focused on the possible involvement of glutamate. It has been proposed that some cases of glaucoma may involve glutamate-mediated excitotoxicity. 2  
In many neuronal tissues, the predominant form of glutamate neurotoxicity is mediated by overstimulation of the N-methyl-d-aspartate (NMDA) subtype of glutamate receptors, which in turn causes excessive concentrations of intracellular Ca2+. In agreement with this, it has been reported that RGCs in mixed retinal cell cultures are susceptible to NMDA-induced cell death in certain culture conditions. 3 4 5 6 It has also been shown that activation of α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) and kainate subtypes of glutamate receptors, may play a role in glutamate neurotoxicity in many types of central nervous system neurons, 7 8 9 10 including RGCs. 11 In situ hybridization studies have shown that both AMPA-kainate receptors 12 and NMDA receptors 13 are expressed in the inner retina. In patch-clamp experiments, RGCs in mixed retinal cell culture display relatively small NMDA-evoked currents but fairly robust kainate currents. 14 15 In general, AMPA-kainate receptors of mature neurons are not permeable to Ca2+. However, it has been reported that isolated RGCs from rats aged between 3 and 8 days express Ca2+-permeable AMPA-kainate receptors in relatively large amounts. 16 AMPA-kainate receptor channels with high Ca2+ permeability have been detected in dissociated RGCs, and this has been correlated with the edited form of the GluR2 subunit expressed. 17 18 Recently, we showed that glutamate could activate Ca2+-permeable AMPA-kainate receptors in RGCs, which caused increases in intracellular calcium ([Ca2+]i) and decrease in cell survival of RGCs. 19 Whether such receptor channels are expressed in adult RGCs or can be expressed under pathologic conditions, is unknown. 
Arachidonic acid (AA), a cell diffusible fatty acid, is thought to serve as an intercellular messenger in many parts of the central nervous system. 20 The liberation of AA from membrane phospholipids by neural activity occurs through either activation of phospholipase A2 alone or the combined activation of phospholipase C and diacylglycerol lipase. 20 21 22 23 A number of neurotransmitters, including glutamate, serotonin, acetylcholine, and catecholamines, can initiate release of AA. At the receptor level, the activation of NMDA receptors or both AMPA and metabotropic glutamate receptors leads to release of AA from cultured neurons. 20 22 Release of AA has also been observed in preparations such as brain slices and cultured astrocytes. 20 AA released during neuronal activity exerts its effects directly, through activation of protein kinase C and formation of free radicals, or through its derivatives formed by the action of cyclooxygenase, lipoxygenase, and epoxygenase. 20 23 In physiological or pathologic conditions, AA can modulate ion channels, transporters, and receptors, 23 24 for example, inhibiting AMPA and kainate receptors in freshly dissociated cerebellar granule cells 25 and dorsal root ganglion neurons. 26 27 However, the functional significance of the inhibition of these non-NMDA ionotropic receptors by AA is unknown. An increasing body of evidence has shown the beneficial effects of fatty acids on various brain functions, such as epileptic seizures, 28 29 depression, 30 and other behavioral diseases. 31 It has been suggested that fatty acids may exert their beneficial effects by decreasing neuronal excitability through inhibition of the neurotransmitter receptors, such as non-NMDA ionotropic receptors that underlie this excitability, 29 32 indicating the clinical potential of AA as a neuroprotector. 
In the present study, we investigated whether AA could protect RGCs from glutamate neurotoxicity. We studied the effects of AA on purified postnatal RGC survival and monitored [Ca2+]i signals and AMPA-kainate receptor-mediated membrane currents. The results showed that low concentrations of AA could protect RGCs from glutamate-induced RGC death by decreasing the calcium influx through non-NMDA ionotropic receptors. 
Materials and Methods
Experimental animals were treated in accordance with the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research. Cell culture reagents were obtained from Gibco (Grand Island, NY), papain from Worthington Biochemical (Freehold, NJ), and recombinant neurotrophic factors from R&D Systems (Minneapolis, MN; human brain-derived neurotrophic factor, BDNF) and Peprotech (Rocky Hill, NJ; rat ciliary neurotrophic factor, CNTF). Unless noted, all other reagents were obtained from Sigma (St. Louis, MO). 
Preparation of Retinal Suspensions
Retinal ganglion cells were purified, as previously described. 19 33 Briefly, retinas from 7- to 8-day-old Long-Evans rats were dissected and incubated at 37°C for 30 minutes in 10 U/mL papain and 70 U/mL collagenase in Hanks’ balanced salt solution containing 0.2 mg/mL bovine serum albumin (BSA) and 0.2 mg/mL dl-cysteine. To yield a suspension of single cells, the tissue was then triturated sequentially through a narrow-bore Pasteur pipette in a solution containing 2 mg/mL ovomucoid, 0.004% DNase, and 1 mg/mL BSA. After centrifugation at 600 rpm for 5 minutes, the cells were rewashed in another ovomucoid-BSA solution (10 mg/mL of each). After centrifugation, the cells were resuspended in 0.1% BSA in phosphate-buffered saline (PBS). 
Panning Procedure
MAC1 or 2G12 (anti-Thy1) were purified from hybridoma supernatants by salt precipitation and affinity chromatography on protein A columns (Bio-Rad, Hercules, CA), according to the manufacturer’s protocols. The specificity of these antibodies and preparation of antibody-coated tubes, as well as the panning procedure, have been described previously. 19 Adherent cells on 2G12-coated tubes were washed with serum-free culture medium (described later). After centrifugation at 600 rpm for 5 minutes, the cells were seeded on 12-mm glass coverslips that had been coated, first with 50 μg/mL poly-l-lysine and then with 10 μg/mL laminin. 
Culture of Purified Retinal Ganglion Cells
Purified RGCs were plated at a low density of approximately 200 cells/cm2 of growth substrate. This plating density provided cultures in which most RGCs grew in physical isolation from other cells. The purified RGCs were cultured in serum-free medium (Neurobasal; Gibco), containing 1 mM glutamine, 10 μg/mL gentamicin, B27 supplement (1:50), 40 ng/mL each of BDNF and CNTF, and 5 μM forskolin (RGC culture medium). Cultures were maintained at 37°C in a humidified atmosphere containing 5% CO2 and 95% air. For long-term survival studies, culture medium was changed every 2 weeks. 
For survival assays, cultures were changed into culture medium containing 10% dialyzed fetal bovine serum. Glutamate at concentrations of 0 to 100 μM was added at the beginning of the experiment. 
Assay of Retinal Ganglion Cell Survival
The viability of RGCs, 2, 7, 14, 28, or 56 days after purification was determined using 1 μM calcein-AM, as previously described. 19 Because all ganglion cells were postmitotic when isolated and we have never seen increases in cell number in control cultures, the measurements represent cell survival rather than proliferation. In this study, a surviving RGC was defined as a cell with calcein-stained cell body and a process extending at least two cell diameters from the cell body. The percentage of surviving RGCs was determined for each time point. 
Three days after exposure to various concentrations of AA, glutamate, or both, cell viability was determined as described. Approximately 200 cells were counted in the no-treatment experiment. The percentage of surviving RGCs was determined for each condition and was normalized to control specimens examined in parallel under the same conditions. The average relative percentage of cell survival in 10 experiments conducted under each condition is expressed in the text and figure as the mean ± SD. Statistical comparisons were made with Student’s t-test. 
[Ca2+]i Measurements
The membrane-permeant fluorescent calcium indicator dye, fluo-3 acetoxymethyl ester, was dissolved in dimethyl sulfoxide to produce a 500-μM stock solution. For dye loading, cells were incubated in Hanks’ balanced salt solution (HBSS) containing 5 μM fluo-3 for 10 minutes at 37°C and then were washed three times with HBSS at 37°C. All Ca2+ imaging was performed at room temperature. A laser scanning confocal system (MRC-1024; Bio-Rad, Hercules, CA) attached to an inverted microscope (Axiovert S100; Carl Zeiss, Thornwood, NY) and equipped with a krypton-argon ion laser, was used to visualize Ca2+-mediated fluorescence in the RGCs. The excitation illumination was 488 nm, and emitted fluorescence was collected through a 515-nm long-pass filter. Images were collected in standard confocal mode and in phase contrast using a transmitted light detector (Bio-Rad). Time-series images were made by collecting fluorescence images at a rate of 0.6 second. Confocal-image files were analyzed by computer (Laser Sharp software; Bio-Rad). The relative increase in fluorescence was calculated by dividing the pixel intensities of the image during stimulation by the pixel intensity of the control image before stimulation. Cells were treated sequentially with AA, then glutamate, AMPA, or kainate, in which case cells were exposed to AA for 2 minutes before and then during stimulation with glutamate, AMPA, or kainate. We waited for more than 5 minutes for the cells to return to the resting level between experiments on the same cells. These experiments were performed in the presence of 1.8 mM Ca2+ and 0.8 mM Mg2+ (the same concentration of Ca2+ and Mg as in the culture medium). All values are expressed as mean ± SD. 
Electrophysiological Recordings
Purified RGCs were maintained in low-density culture for 7 to 15 days. The cells on coverslips were held in a recording chamber and bathed in a Ringer’s solution of the following composition (in millimolar): 135 NaCl, 4.3 KCl, 1.7 CaCl2, 1.2 MgSO4, 0.5 KH2PO4, 2 NaHCO3, 10 HEPES, 15 glucose, 0.1 ascorbate, and 0.5 glutamine; pH was adjusted to 7.4 with NaOH. Thick-wall borosilicate glass was used for recording electrodes and were drawn on an electrode puller (Brown Flaming P-87; Sutter Instruments, San Rafael, CA). The typical resistance of the recording electrodes ranged from 4 to 9 MΩ when filled with an intracellular solution containing (in millimolar) 1 NaCl, 145 KCl, 2 MgCl2, 1 CaCl2, 10 HEPES, 2 EGTA, 2 adenosine triphosphate (ATP)-Mg, and 0.5 GTP-Na (pH 7.4). Compensations of electrode capacitance and series resistance were used for all recordings. Recordings were performed at a holding potential of −70 mV, with a patch clamp amplifier (3900A; Dagan, Minneapolis, MN) connected to a computer (Compaq, Houston, TX) by an -interface (TL-1 DMA). Recordings were controlled by computer (pClamp, ver. 6; Axon Instruments, Foster City, CA), filtered at 2 kHz, and stored on computer hard disk for off-line data analysis. 
Several software tools were used for off-line data analysis. Data of agonist-induced currents were exported by computer (Clampfit, ver. 6.0; Axon Instruments, Inc.). The exported data were processed with graphics software (CorelDRAW, ver. 8.232; Corel Corporation, Ottawa, Ontario, Canada). Statistical analysis and related figures were completed on computer (Origin, ver. 4.10; Microcal Software, Inc., Northampton, MA), and results presented as the mean ± SD. 
Results
RGC Viability with Time
To check the viability of RGCs, cells were labeled with calcein 2, 7, 14, 28, or 56 days after purification, and the percentage of surviving RGCs determined for each time. Even after 2 months of culture, the survival of RGCs was 44% ± 11.6%. 
Effect of AA on Retinal Ganglion Cell Survival
To investigate the effects of AA on purified RGCs, we incubated cultures with 1 to 50 μM AA. Increasing concentrations of AA caused a dose-dependent increase in cell death after 3 days of culture with an ED50 of 22.0 μM (Fig. 1) . Among the concentrations, 20 μM and 50 μM AA significantly reduced the survival of RGCs (P < 0.001). 
Effects of AA on Glutamate-Induced RGC Death
We previously reported that low concentrations of glutamate had neurotoxic effects on purified RGCs. 19 33 In the present study, the effect of AA on RGCs exposed to glutamate was examined. Results showed that the neurotoxic effect of 25 μM glutamate was significantly ameliorated by 3 μM (P < 0.001), but not 1 μM, AA (Fig. 2)
[Ca2+]i Measurements
To explore the mechanism underlying the neuroprotective effect of AA, we determined whether low doses of AA altered glutamate-induced increase in ([Ca2+]i) in purified RGCs, by using the fluorescent indicator dye fluo-3. 
Glutamate at 25 μM caused a rapid increase in [Ca2+]i in RGCs that was reversibly inhibited by AA. Results of a typical experiment examining the effects of AA on glutamate-induced [Ca2+]i change are shown in Figure 3 . Treatment of RGCs with 25 μM glutamate increased the [Ca2+]i level (Fig. 3b 3B 3C) . The glutamate-induced increase in [Ca2+]i was inhibited by treatment with 3 μM AA treatment (Fig. 3b 3D 3E) . After AA was washed out, glutamate reversibly increased [Ca2+]i again (Fig. 3b 3G 3F) . Furthermore, we calculated how much [Ca2+]i was increased by glutamate from the resting state. Glutamate induced a 3.41 ± 0.82-fold [Ca2+]i increase over the control level (Fig. 3c ; n = 8). Treatment with 3 μM AA significantly reduced the increment in fluo-3 fluorescence induced by glutamate, which was significant by paired t-test (2.31 ± 0.37 times less than that of control in the same RGCs; n = 8; P < 0.01; Fig. 3c ). 
We also investigated whether the increase in [Ca2+]i was dependent on Ca2+ influx from the extracellular space or release from intracellular Ca2+ stores. The cells were perfused with Ca2+-free HBSS with 1 mM EDTA (J. T. Baker, Phillipsburg, NJ) for 30 seconds before and then during stimulation with glutamate. Ca2+-free perfusion blocked the glutamate-induced increase in [Ca2+]i, indicating a necessary role for Ca2+ influx. 
Results of changes in fluo-3 fluorescence by glutamate receptor agonists, with or without AA, are shown in Figure 4 . AMPA (10 μM) and kainate (KA; 50 μM) increased [Ca2+]i (Figs. 4a 4c) , although NMDA did not (data not shown). Pretreatment with 3 μM AA significantly reduced the [Ca2+]i increase induced by 10 μM AMPA (AMPA, 3.88 ± 0.44 times; AMPA+AA, 3.19 ± 0.47 times; n = 8, P < 0.01, Fig. 4b ) or 50 μM kainate (kainate, 3.84 ± 0.58 times; kainate+AA, 2.50 ± 0.59 times; n = 8, P < 0.01, Fig. 4d ). These observations confirm our previous suggestion that glutamate neurotoxicity may be mediated by AMPA-kainate receptors on purified RGCs, and also suggest that AA may exert its protective effects by inhibiting these receptors. 
Electrophysiological Analysis
To investigate how AA reduced glutamate-induced ganglion cell death and exerted its neuroprotection, the effect of AA on glutamate receptor-mediated whole-cell currents was examined. The results showed that AA inhibited 25 μM glutamate and 50 μM kainate induced currents in a dose-dependent manner (Fig. 5) . Currents induced by 25 μM glutamate were reduced by AA to 88.9% ± 11.1% (0.3 μM AA), 56.2% ± 22.3% (1 μM AA), 52.5% ± 23.3% (3 μM AA), and 28.1% ± 18.7% (10 μM AA) of control, respectively (Figs. 5a 5c) . From these data a median inhibitory concentration (IC50) of 2.30 μM was calculated. The inhibition by 1 to 10 μM AA was significant (P < 0.01; n = 10), but that of 0.3 μM AA was not (P > 0.01, n = 10). 2-Amino-5-phosphonovaleric acid (APV; 100 μM), an NMDA receptor antagonist, had no effect on 25 μM glutamate-induced currents, but 20 μM 6,7-dinitroquinoxeline-2,3-dione (DNQX), an non-NMDA receptor antagonist, totally blocked these currents, as we have reported, 19 confirming that, in these cultures, 25 μM glutamate-induced currents were mediated by non-NMDA receptors. We have not attempted to identify the individual contributions of AMPA and kainate type receptors to these responses. Kainate-induced currents at 50 μM were inhibited to 84.7% ± 18.9%, 75.5% ± 14.6%, 57.2% ± 7.2%, and 17.2% ± 13.9% by 0.3 μM, 1 μM, 3 μM, and 10 μM AA, respectively (0.3 μM AA, n = 5; 1–10 μM AA, n = 14). From these data, an IC50 of 4.55 μM was calculated. Kainate currents were significantly inhibited by 1 to 10 μM AA (P < 0.01; n = 10), but not by 0.3 μM AA (P > 0.01, n = 10). These data show that AA had similar inhibitory effects on kainate- and glutamate-induced responses. 
The time dependence of the AA effect was also examined. Glutamate (25 μM) was applied to the test cells every 0.5 minute when the cells were incubated with 3 μM AA (Fig. 6a) . Glutamate-induced currents were not significantly changed by coapplication of 3 μM AA to the test cells (104.8% ± 17.9% of control; P > 0.01, n = 8) or by incubation of 3 μM AA for approximately 0.5 minute (75.5% ± 17.4% of control; P > 0.01, n = 8). Inhibition of glutamate-induced currents by AA varied with the duration of treatment, first being detectable after 1 minute (P < 0.01, n = 8) and reaching a maximum of approximately 50% after 4 minutes (Fig 6c) . Currents induced by 50 μM kainate were also significantly inhibited by complication of 3 μM AA (Fig. 6b) , but by less, being reduced to around 70% of control after 4 to 5 minutes (Fig. 6c)
Discussion
In the present study, we used purified RGCs, and an accurate and sensitive measurement of ganglion cell survival, to explore the effects of AA on glutamate toxicity. 19 33 A number of studies have been published on the relationship between RGC death and glutamate application in vitro, but the different methods used and the rapid and extensive RGC death noted in many control cultures make interpretation and comparison difficult. In mixed retinal cell cultures, the treatment with drugs may stimulate release of other factors that may act on RGCs. For example, blocking nitric oxide production or action decreases glutamate-induced or NMDA-mediated cell death in cultured retinal neurons. 34 35 In the presence of retinal Müller cells, glutamate is rapidly removed, 4 33 36 37 and nitric oxide toxicity is reduced. 32 Growth factors released from other cells in mixed cultures may also have complicated effects on the test cells. This makes mechanistic interpretation of changes in cell survival difficult. 
Several neurotransmitters cause the release of AA by activation of phospholipase A2 and phospholipase C in neuronal and glial cells. The physiological concentration of AA released in the intact tissue is unknown, but is probably below the critical micellar concentration of 30 μM. 20 Because it is very lipophilic, it may well be concentrated in membranous compartments and thus the effective concentration may be very different from the overall tissue concentration. Accumulating evidence shows that AA has biological effects at concentrations between 1 μM and 1000 μM. Knowing the real concentration of AA in intact tissue is important, because different concentrations of AA have complicated actions, sometimes even opposite effects at low and high concentrations, respectively. 38 We observed that low concentrations of AA had little effect on RGCs, but concentrations higher than 10 μM significantly decreased RGC survival. In the present study, we found that 3 μM AA alone had no detectable effect on RGCs, but significantly prevented glutamate-induced RGC death. We also selected this concentration because AA in the low micromolar range does not act through protein kinase C or through cyclooxygenase, lipoxygenase, or epoxygenase metabolites of AA. 25 26  
AA has positive or negative effects on both NMDA and non-NMDA subtypes of glutamate ionotropic receptors in variety of preparations. 26 39 Homologies in sequences of NMDA receptor subunits and fatty acid-binding proteins, suggest that there may be a fatty acid-binding domain on NMDA receptors. 39 However, the sequences of most non-NMDA ionotropic receptor subunits have no similar homology with fatty acid binding proteins. 26 The GluR6 subunit shows a weak homology, and homomeric GluR6 receptors expressed in HEK cells can be inhibited by AA and other fatty acids. 26 27 There is no evidence that such homomeric receptors are expressed in RGCs, and thus it is unlikely that AA has a direct effect on non-NMDA ionotropic receptors in these cells. In support of this, we found that AA had no significant effects on glutamate-induced inward currents when applied simultaneously with glutamate, but that preincubation with AA significantly inhibited Glu-induced currents (Figs. 6a 6c)
The increase in [Ca2+]i seen after glutamate or agonist application required external calcium but may also have a component from internal stores through calcium-induced calcium release. Because AA inhibited the glutamate receptor currents, we think that this is likely to explain the AA-mediated inhibition of the increase in [Ca2+]i. We cannot, at present, rule out an additional effect of AA on release from internal stores. The AMPA and kainate-induced [Ca2+]i responses were different between control (before treatment with AA) and recovery (after AA washing out; Figs. 4a 4c ). The reduced peak in the washout response may be due, in part, to depletion of calcium stores. Because the AA treated response was smaller than the washout response, and because this difference between control and washout responses was not seen with the natural ligand glutamate, the conclusion that AA reduces calcium responses remains valid. 
During brain anoxia or ischemia, there is a large release of glutamate into extracellular space, due to the redistribution of ions across cell membranes. As a result, [Ca2+]i increases to a level that can lethally activate calcium-dependent enzymes and leads to neuronal apoptosis. Simultaneously, ischemia promotes release of AA from neurons and glial cells. 20 The released AA inhibits glutamate uptake and also desensitizes postsynaptic AMPA and NMDA receptors, the latter effect perhaps accounting for the neuroprotective function of AA in ischemia. Ischemia, especially of the distal optic nerve and RGC, is one of prominent stress factors identified in the eyes of patients with glaucoma. 40 On the basis of our findings that a low concentration of AA protected RGCs from glutamate neurotoxicity, we expect that this agent would have similar neuroprotective effects in vivo. Steroidal anti-inflammatory drugs have been used after glaucoma filtering surgery. 41 These drugs prevent the formation of AA, a precursor of potent inflammatory mediators, such as prostaglandins, thromboxane, and leukotrienes, by inhibiting the action of phospholipase A2. 41 42 Therefore, steroids may actually have harmful effects by reducing AA neuroprotective effects in ischemia of glaucoma. 
Our results suggest several approaches that may be of benefit to patients with glaucoma who have progressive visual field loss, despite satisfactory control of intraocular pressure. First, drugs selectively blocking downstream pathways of AA metabolism should be used in glaucoma treatment. Nonsteroidal anti-inflammatory drugs such as ferulic acid, piroxicam, and phenidone, especially inhibit the lipoxygenase and/or the cyclooxygenase pathways and consequently prevent the formation of inflammatory mediators. 41 Second, a suitable concentration of AA should be maintained to promote survival of RGCs. 
 
Figure 1.
 
Dose-dependent effects of AA on RGC survival. Purified RGCs were cultured for 2 days in serum-free medium and then exposed for 3 days to various concentrations of AA. Survival of treated RGCs is expressed as a percentage of surviving cells in parallel untreated cultures. Each data point appears as the mean ± SD (n = 10 experiments). Application of increasing concentrations (1–50 μM) of AA caused a dose-dependent decrease in survival. The calculated ED50 for AA was 22.0 μM. *Significant difference compared with control (P < 0.001).
Figure 1.
 
Dose-dependent effects of AA on RGC survival. Purified RGCs were cultured for 2 days in serum-free medium and then exposed for 3 days to various concentrations of AA. Survival of treated RGCs is expressed as a percentage of surviving cells in parallel untreated cultures. Each data point appears as the mean ± SD (n = 10 experiments). Application of increasing concentrations (1–50 μM) of AA caused a dose-dependent decrease in survival. The calculated ED50 for AA was 22.0 μM. *Significant difference compared with control (P < 0.001).
Figure 2.
 
The effects of AA on RGC death induced by 25 μM glutamate. Three days after exposure to AA, glutamate, or both, cell viability was determined. The number of surviving RGCs was determined for each condition and was normalized to control specimens. Glutamate (25 μM) significantly reduced RGC survival (*P < 0.001), and the neurotoxic effect of 25 μM glutamate was significantly ameliorated by 3 μM AA (#P < 0.001). Significant neuroprotection by 1 μM AA was not detected.
Figure 2.
 
The effects of AA on RGC death induced by 25 μM glutamate. Three days after exposure to AA, glutamate, or both, cell viability was determined. The number of surviving RGCs was determined for each condition and was normalized to control specimens. Glutamate (25 μM) significantly reduced RGC survival (*P < 0.001), and the neurotoxic effect of 25 μM glutamate was significantly ameliorated by 3 μM AA (#P < 0.001). Significant neuroprotection by 1 μM AA was not detected.
Figure 3.
 
(a) Measurement of [Ca2+]i by fluo-3 and confocal microscopy. Glutamate (25 μM ) induced a rapid increase in [Ca2+]i from the baseline (B) to the peak (C) level. AA (3 μM) alone did not evoke a change in [Ca2+]i (D) and 3 μM AA pretreatment for 2 minutes abolished the 25 μM glutamate-induced [Ca2+]i increase (E). The inhibitory effect of AA was reversible. Repeated treatment with glutamate again elicited a peak in [Ca2+]i (G) from the baseline level (F). (a, arrows) Times at which the images in (b, B–G) were captured. Micrographs show the change in fluorescence of fluo-3-AM-loaded retinal ganglion cells in response to 25 μM glutamate, without or with treatment with 3 μM AA. A phase-contrast image of two typical RGCs is shown in (b, A). Fluorescence images of the two RGCs were shown before (B, D, F) and during application of 25 μM glutamate without (C, G) or with (E) 3 μM AA. Pretreatment with AA significantly reduced the increase in fluo-3 fluorescence produced by application of 25 μM glutamate. Scale bar: (A) 20 μm. (c) Summary data showing the increase in fluo-3-AM fluorescence induced by 25 μM glutamate in the presence or absence of AA. AA reduced the glutamate-induced increase in [Ca2+]i. Bars, ±SD. *Significant difference compared with control (P < 0.01, n = 8 cells).
Figure 3.
 
(a) Measurement of [Ca2+]i by fluo-3 and confocal microscopy. Glutamate (25 μM ) induced a rapid increase in [Ca2+]i from the baseline (B) to the peak (C) level. AA (3 μM) alone did not evoke a change in [Ca2+]i (D) and 3 μM AA pretreatment for 2 minutes abolished the 25 μM glutamate-induced [Ca2+]i increase (E). The inhibitory effect of AA was reversible. Repeated treatment with glutamate again elicited a peak in [Ca2+]i (G) from the baseline level (F). (a, arrows) Times at which the images in (b, B–G) were captured. Micrographs show the change in fluorescence of fluo-3-AM-loaded retinal ganglion cells in response to 25 μM glutamate, without or with treatment with 3 μM AA. A phase-contrast image of two typical RGCs is shown in (b, A). Fluorescence images of the two RGCs were shown before (B, D, F) and during application of 25 μM glutamate without (C, G) or with (E) 3 μM AA. Pretreatment with AA significantly reduced the increase in fluo-3 fluorescence produced by application of 25 μM glutamate. Scale bar: (A) 20 μm. (c) Summary data showing the increase in fluo-3-AM fluorescence induced by 25 μM glutamate in the presence or absence of AA. AA reduced the glutamate-induced increase in [Ca2+]i. Bars, ±SD. *Significant difference compared with control (P < 0.01, n = 8 cells).
Figure 4.
 
(a) Measurements of relative increase in fluo-3 fluorescence induced by AMPA, with or without treatment with AA. Tested cells were treated sequentially with 10 μM AMPA, then 3 μM AA with 10 μM AMPA, and 10 μM AMPA only. Pretreatment with AA was performed for 2 minutes before the application of AMPA. AMPA (10 μM) increased [Ca2+]i rapidly. It was significantly reduced by pretreatment with 3 μM AA. (b) Changes induced by 10 μM AMPA in the presence and absence of AA. AA significantly suppressed the [Ca2+]i increase induced by AMPA (AMPA, 3.88 ± 0.44, and AMPA+AA, 3.19 ± 0.47 times control; n = 8, P < 0.01). Data are the mean ± SD. *Significant difference (P < 0.01). (c) Changes in fluo-3 fluorescence induced by kainate, with or without treatment with AA. Tested cells were treated sequentially with 50 μM kainate, 50 μM kainate with 3 μM AA, and 50 μM kainate alone. Pretreatment with 3 μM AA for 2 minutes significantly reduced the increase in fluo-3 fluorescence produced by application of kainate. (d) Summary of changes in [Ca2+]i induced by kainate in the presence and absence of AA. AA significantly suppressed the increase in [Ca2+]i induced by 50 μM kainate (kainate, 3.84 ± 0.58 times control; and kainate+AA, 2.50 ± 0.59 times control; mean ± SD, n = 8). *Significant difference (P < 0.01).
Figure 4.
 
(a) Measurements of relative increase in fluo-3 fluorescence induced by AMPA, with or without treatment with AA. Tested cells were treated sequentially with 10 μM AMPA, then 3 μM AA with 10 μM AMPA, and 10 μM AMPA only. Pretreatment with AA was performed for 2 minutes before the application of AMPA. AMPA (10 μM) increased [Ca2+]i rapidly. It was significantly reduced by pretreatment with 3 μM AA. (b) Changes induced by 10 μM AMPA in the presence and absence of AA. AA significantly suppressed the [Ca2+]i increase induced by AMPA (AMPA, 3.88 ± 0.44, and AMPA+AA, 3.19 ± 0.47 times control; n = 8, P < 0.01). Data are the mean ± SD. *Significant difference (P < 0.01). (c) Changes in fluo-3 fluorescence induced by kainate, with or without treatment with AA. Tested cells were treated sequentially with 50 μM kainate, 50 μM kainate with 3 μM AA, and 50 μM kainate alone. Pretreatment with 3 μM AA for 2 minutes significantly reduced the increase in fluo-3 fluorescence produced by application of kainate. (d) Summary of changes in [Ca2+]i induced by kainate in the presence and absence of AA. AA significantly suppressed the increase in [Ca2+]i induced by 50 μM kainate (kainate, 3.84 ± 0.58 times control; and kainate+AA, 2.50 ± 0.59 times control; mean ± SD, n = 8). *Significant difference (P < 0.01).
Figure 5.
 
Dose-dependent effects of AA on 25 μM glutamate- and 50 μM kainate-induced inward currents. After the rupture to make whole-cell recording, the agonists were applied to the recorded cells several times, and the cells that had stable responses to the repeated application of agonists were selected for further experiments. (a) Cells were pretreated with different concentrations of AA for approximately 2 minutes before application of 25 μM glutamate. The inhibition of AA on the currents induced by 25 μM glutamate increased with the AA concentration. (b) Similarly, the inhibition by AA of currents induced by 50 μM kainate also increased with the AA concentrations. (c) The statistical results show the dose-dependent inhibitory effects of AA on the currents induced by 25 μM glutamate (n = 10) and 50 μM kainate (0.3 μM AA, n = 5; 1–10 μM AA, n = 14). Significant difference between *glutamate or #kainate and control (P < 0.01).
Figure 5.
 
Dose-dependent effects of AA on 25 μM glutamate- and 50 μM kainate-induced inward currents. After the rupture to make whole-cell recording, the agonists were applied to the recorded cells several times, and the cells that had stable responses to the repeated application of agonists were selected for further experiments. (a) Cells were pretreated with different concentrations of AA for approximately 2 minutes before application of 25 μM glutamate. The inhibition of AA on the currents induced by 25 μM glutamate increased with the AA concentration. (b) Similarly, the inhibition by AA of currents induced by 50 μM kainate also increased with the AA concentrations. (c) The statistical results show the dose-dependent inhibitory effects of AA on the currents induced by 25 μM glutamate (n = 10) and 50 μM kainate (0.3 μM AA, n = 5; 1–10 μM AA, n = 14). Significant difference between *glutamate or #kainate and control (P < 0.01).
Figure 6.
 
Time-dependent inhibitory effect of AA on glutamate- and kainate-induced inward currents. To examine whether AA action is time-dependent, the agonists were applied every 0.5 minute, without or with the treatment of 3 μM AA. (a) Coapplication of 3 μM AA or preincubated only for 0.5 minute did not significantly change the current induced by 25 μM glutamate compared with control (P = 0.376 and P = 0.013; n = 8). Longer preincubation times led to substantial inhibition. (b) Inhibition by 3 μM AA of 50-μM kainate-induced currents was detected with coapplication of kainate and AA and increased with incubation time. (c) Data summary showing the time-dependence of the inhibition of glutamate and kainate currents. Significant difference between *glutamate or #kainate and control (P < 0.01).
Figure 6.
 
Time-dependent inhibitory effect of AA on glutamate- and kainate-induced inward currents. To examine whether AA action is time-dependent, the agonists were applied every 0.5 minute, without or with the treatment of 3 μM AA. (a) Coapplication of 3 μM AA or preincubated only for 0.5 minute did not significantly change the current induced by 25 μM glutamate compared with control (P = 0.376 and P = 0.013; n = 8). Longer preincubation times led to substantial inhibition. (b) Inhibition by 3 μM AA of 50-μM kainate-induced currents was detected with coapplication of kainate and AA and increased with incubation time. (c) Data summary showing the time-dependence of the inhibition of glutamate and kainate currents. Significant difference between *glutamate or #kainate and control (P < 0.01).
The authors thank Keely Bumsted and Stephen Viviano for excellent technical assistance, and Michael Nathanson for helpful discussions. 
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Figure 1.
 
Dose-dependent effects of AA on RGC survival. Purified RGCs were cultured for 2 days in serum-free medium and then exposed for 3 days to various concentrations of AA. Survival of treated RGCs is expressed as a percentage of surviving cells in parallel untreated cultures. Each data point appears as the mean ± SD (n = 10 experiments). Application of increasing concentrations (1–50 μM) of AA caused a dose-dependent decrease in survival. The calculated ED50 for AA was 22.0 μM. *Significant difference compared with control (P < 0.001).
Figure 1.
 
Dose-dependent effects of AA on RGC survival. Purified RGCs were cultured for 2 days in serum-free medium and then exposed for 3 days to various concentrations of AA. Survival of treated RGCs is expressed as a percentage of surviving cells in parallel untreated cultures. Each data point appears as the mean ± SD (n = 10 experiments). Application of increasing concentrations (1–50 μM) of AA caused a dose-dependent decrease in survival. The calculated ED50 for AA was 22.0 μM. *Significant difference compared with control (P < 0.001).
Figure 2.
 
The effects of AA on RGC death induced by 25 μM glutamate. Three days after exposure to AA, glutamate, or both, cell viability was determined. The number of surviving RGCs was determined for each condition and was normalized to control specimens. Glutamate (25 μM) significantly reduced RGC survival (*P < 0.001), and the neurotoxic effect of 25 μM glutamate was significantly ameliorated by 3 μM AA (#P < 0.001). Significant neuroprotection by 1 μM AA was not detected.
Figure 2.
 
The effects of AA on RGC death induced by 25 μM glutamate. Three days after exposure to AA, glutamate, or both, cell viability was determined. The number of surviving RGCs was determined for each condition and was normalized to control specimens. Glutamate (25 μM) significantly reduced RGC survival (*P < 0.001), and the neurotoxic effect of 25 μM glutamate was significantly ameliorated by 3 μM AA (#P < 0.001). Significant neuroprotection by 1 μM AA was not detected.
Figure 3.
 
(a) Measurement of [Ca2+]i by fluo-3 and confocal microscopy. Glutamate (25 μM ) induced a rapid increase in [Ca2+]i from the baseline (B) to the peak (C) level. AA (3 μM) alone did not evoke a change in [Ca2+]i (D) and 3 μM AA pretreatment for 2 minutes abolished the 25 μM glutamate-induced [Ca2+]i increase (E). The inhibitory effect of AA was reversible. Repeated treatment with glutamate again elicited a peak in [Ca2+]i (G) from the baseline level (F). (a, arrows) Times at which the images in (b, B–G) were captured. Micrographs show the change in fluorescence of fluo-3-AM-loaded retinal ganglion cells in response to 25 μM glutamate, without or with treatment with 3 μM AA. A phase-contrast image of two typical RGCs is shown in (b, A). Fluorescence images of the two RGCs were shown before (B, D, F) and during application of 25 μM glutamate without (C, G) or with (E) 3 μM AA. Pretreatment with AA significantly reduced the increase in fluo-3 fluorescence produced by application of 25 μM glutamate. Scale bar: (A) 20 μm. (c) Summary data showing the increase in fluo-3-AM fluorescence induced by 25 μM glutamate in the presence or absence of AA. AA reduced the glutamate-induced increase in [Ca2+]i. Bars, ±SD. *Significant difference compared with control (P < 0.01, n = 8 cells).
Figure 3.
 
(a) Measurement of [Ca2+]i by fluo-3 and confocal microscopy. Glutamate (25 μM ) induced a rapid increase in [Ca2+]i from the baseline (B) to the peak (C) level. AA (3 μM) alone did not evoke a change in [Ca2+]i (D) and 3 μM AA pretreatment for 2 minutes abolished the 25 μM glutamate-induced [Ca2+]i increase (E). The inhibitory effect of AA was reversible. Repeated treatment with glutamate again elicited a peak in [Ca2+]i (G) from the baseline level (F). (a, arrows) Times at which the images in (b, B–G) were captured. Micrographs show the change in fluorescence of fluo-3-AM-loaded retinal ganglion cells in response to 25 μM glutamate, without or with treatment with 3 μM AA. A phase-contrast image of two typical RGCs is shown in (b, A). Fluorescence images of the two RGCs were shown before (B, D, F) and during application of 25 μM glutamate without (C, G) or with (E) 3 μM AA. Pretreatment with AA significantly reduced the increase in fluo-3 fluorescence produced by application of 25 μM glutamate. Scale bar: (A) 20 μm. (c) Summary data showing the increase in fluo-3-AM fluorescence induced by 25 μM glutamate in the presence or absence of AA. AA reduced the glutamate-induced increase in [Ca2+]i. Bars, ±SD. *Significant difference compared with control (P < 0.01, n = 8 cells).
Figure 4.
 
(a) Measurements of relative increase in fluo-3 fluorescence induced by AMPA, with or without treatment with AA. Tested cells were treated sequentially with 10 μM AMPA, then 3 μM AA with 10 μM AMPA, and 10 μM AMPA only. Pretreatment with AA was performed for 2 minutes before the application of AMPA. AMPA (10 μM) increased [Ca2+]i rapidly. It was significantly reduced by pretreatment with 3 μM AA. (b) Changes induced by 10 μM AMPA in the presence and absence of AA. AA significantly suppressed the [Ca2+]i increase induced by AMPA (AMPA, 3.88 ± 0.44, and AMPA+AA, 3.19 ± 0.47 times control; n = 8, P < 0.01). Data are the mean ± SD. *Significant difference (P < 0.01). (c) Changes in fluo-3 fluorescence induced by kainate, with or without treatment with AA. Tested cells were treated sequentially with 50 μM kainate, 50 μM kainate with 3 μM AA, and 50 μM kainate alone. Pretreatment with 3 μM AA for 2 minutes significantly reduced the increase in fluo-3 fluorescence produced by application of kainate. (d) Summary of changes in [Ca2+]i induced by kainate in the presence and absence of AA. AA significantly suppressed the increase in [Ca2+]i induced by 50 μM kainate (kainate, 3.84 ± 0.58 times control; and kainate+AA, 2.50 ± 0.59 times control; mean ± SD, n = 8). *Significant difference (P < 0.01).
Figure 4.
 
(a) Measurements of relative increase in fluo-3 fluorescence induced by AMPA, with or without treatment with AA. Tested cells were treated sequentially with 10 μM AMPA, then 3 μM AA with 10 μM AMPA, and 10 μM AMPA only. Pretreatment with AA was performed for 2 minutes before the application of AMPA. AMPA (10 μM) increased [Ca2+]i rapidly. It was significantly reduced by pretreatment with 3 μM AA. (b) Changes induced by 10 μM AMPA in the presence and absence of AA. AA significantly suppressed the [Ca2+]i increase induced by AMPA (AMPA, 3.88 ± 0.44, and AMPA+AA, 3.19 ± 0.47 times control; n = 8, P < 0.01). Data are the mean ± SD. *Significant difference (P < 0.01). (c) Changes in fluo-3 fluorescence induced by kainate, with or without treatment with AA. Tested cells were treated sequentially with 50 μM kainate, 50 μM kainate with 3 μM AA, and 50 μM kainate alone. Pretreatment with 3 μM AA for 2 minutes significantly reduced the increase in fluo-3 fluorescence produced by application of kainate. (d) Summary of changes in [Ca2+]i induced by kainate in the presence and absence of AA. AA significantly suppressed the increase in [Ca2+]i induced by 50 μM kainate (kainate, 3.84 ± 0.58 times control; and kainate+AA, 2.50 ± 0.59 times control; mean ± SD, n = 8). *Significant difference (P < 0.01).
Figure 5.
 
Dose-dependent effects of AA on 25 μM glutamate- and 50 μM kainate-induced inward currents. After the rupture to make whole-cell recording, the agonists were applied to the recorded cells several times, and the cells that had stable responses to the repeated application of agonists were selected for further experiments. (a) Cells were pretreated with different concentrations of AA for approximately 2 minutes before application of 25 μM glutamate. The inhibition of AA on the currents induced by 25 μM glutamate increased with the AA concentration. (b) Similarly, the inhibition by AA of currents induced by 50 μM kainate also increased with the AA concentrations. (c) The statistical results show the dose-dependent inhibitory effects of AA on the currents induced by 25 μM glutamate (n = 10) and 50 μM kainate (0.3 μM AA, n = 5; 1–10 μM AA, n = 14). Significant difference between *glutamate or #kainate and control (P < 0.01).
Figure 5.
 
Dose-dependent effects of AA on 25 μM glutamate- and 50 μM kainate-induced inward currents. After the rupture to make whole-cell recording, the agonists were applied to the recorded cells several times, and the cells that had stable responses to the repeated application of agonists were selected for further experiments. (a) Cells were pretreated with different concentrations of AA for approximately 2 minutes before application of 25 μM glutamate. The inhibition of AA on the currents induced by 25 μM glutamate increased with the AA concentration. (b) Similarly, the inhibition by AA of currents induced by 50 μM kainate also increased with the AA concentrations. (c) The statistical results show the dose-dependent inhibitory effects of AA on the currents induced by 25 μM glutamate (n = 10) and 50 μM kainate (0.3 μM AA, n = 5; 1–10 μM AA, n = 14). Significant difference between *glutamate or #kainate and control (P < 0.01).
Figure 6.
 
Time-dependent inhibitory effect of AA on glutamate- and kainate-induced inward currents. To examine whether AA action is time-dependent, the agonists were applied every 0.5 minute, without or with the treatment of 3 μM AA. (a) Coapplication of 3 μM AA or preincubated only for 0.5 minute did not significantly change the current induced by 25 μM glutamate compared with control (P = 0.376 and P = 0.013; n = 8). Longer preincubation times led to substantial inhibition. (b) Inhibition by 3 μM AA of 50-μM kainate-induced currents was detected with coapplication of kainate and AA and increased with incubation time. (c) Data summary showing the time-dependence of the inhibition of glutamate and kainate currents. Significant difference between *glutamate or #kainate and control (P < 0.01).
Figure 6.
 
Time-dependent inhibitory effect of AA on glutamate- and kainate-induced inward currents. To examine whether AA action is time-dependent, the agonists were applied every 0.5 minute, without or with the treatment of 3 μM AA. (a) Coapplication of 3 μM AA or preincubated only for 0.5 minute did not significantly change the current induced by 25 μM glutamate compared with control (P = 0.376 and P = 0.013; n = 8). Longer preincubation times led to substantial inhibition. (b) Inhibition by 3 μM AA of 50-μM kainate-induced currents was detected with coapplication of kainate and AA and increased with incubation time. (c) Data summary showing the time-dependence of the inhibition of glutamate and kainate currents. Significant difference between *glutamate or #kainate and control (P < 0.01).
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