Iron alters glutamate secretion by regulating cytosolic aconitase activity

M. Christine McGahan, Jill Harned, Marilyn Mukunnemkeril, Malgorzata Goralska, Lloyd Fleisher, and Jenny B. Ferrell

Department of Molecular Biomedical Sciences, North Carolina State University, Raleigh, North Carolina

Submitted 9 September 2004 ; accepted in final form 17 December 2004


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Glutamate has many important physiological functions, including its role as a neurotransmitter in the retina and the central nervous system. We have made the novel observations that retinal pigment epithelial cells underlying and intimately interacting with the retina secrete glutamate and that this secretion is significantly affected by iron. In addition, iron increased secretion of glutamate in cultured lens and neuronal cells, indicating that this may be a common mechanism for the regulation of glutamate production in many cell types. The activity of the iron-dependent enzyme cytosolic aconitase (c-aconitase) is increased by iron. The conversion of citrate to isocitrate by c-aconitase is the first step in a three-step process leading to glutamate formation. In the present study, iron increased c-aconitase activity, and this increase was associated with an increase in glutamate secretion. Inhibition of c-aconitase by oxalomalate decreased glutamate secretion and completely inhibited the iron-induced increase in glutamate secretion. Derangements in both glutamate secretion and iron metabolism have been noted in neurological diseases and retinal degeneration. Our results are the first to provide a functional link between these two physiologically important substances by demonstrating a significant role for iron in the regulation of glutamate production and secretion in mammalian cells resulting from iron regulation of aconitase activity. Glutamatergic systems are found in many nonneuronal tissues. We provide the first evidence that, in addition to secreting glutamate, retinal pigment epithelial cells express the vesicular glutamate transporter VGLUT1 and that regulated vesicular release of glutamate from these cells can be inhibited by riluzole.

retinal pigment epithelial cells; lens epithelial cells


AMONG ITS NUMEROUS PHYSIOLOGICAL FUNCTIONS, glutamate is a major excitatory neurotransmitter in the central nervous system and the retina. Indeed, proper regulation of glutamate levels is essential to normal neuronal function, with astrocytes playing an important role (48). At high concentrations, glutamate can be a dangerous excitotoxin, and elevated glutamate concentration can occur as a result of defects in cellular reuptake and/or excess production and secretion. High concentrations of glutamate have deleterious effects on retinal function (9, 52, 54). Retinal pigment epithelial (RPE) cells contain glutamate transporters (41, 42, 47), and it has been hypothesized that uptake of glutamate by RPE cells is important in regulating extracellular retinal glutamate concentration to protect the neural retina from excitotoxic damage caused by high levels of glutamate (52, 54). However, to our knowledge, glutamate secretion by RPE cells has not been evaluated. In addition, mechanisms by which glutamate production and secretion might be altered are poorly defined.

The iron-regulatory protein (IRP) is a dual-function cytoplasmic protein. When iron is scarce, IRP binds to the mRNA of a number of proteins involved in iron metabolism; when iron is abundant, IRP acquires aconitase activity (21, 26, 34). In the latter situation, iron is used to complete an iron sulfur cluster in IRP, which confers aconitase activity on this protein. Therefore, iron availability regulates cytosolic aconitase (c-aconitase) activity. Mitochondrial aconitase does not have this dual function and is regulated differently from its cytosolic counterpart (29, 32, 46). Aconitase is an important mitochondrial enzyme, but the function of c-aconitase is not as clear (51). Cytosolic isocitrate dehydrogenase (CICD), present in most cell types examined, recently has been identified in neurons, astrocytes, oligodendrocytes, and microglia (45). CICD converts isocitrate, formed by the action of aconitase, to {alpha}-ketoglutarate with concomitant reduction of NADP (Fig. 1). It has been suggested that an increase in iron load, with its potential for causing oxidative damage, could trigger a protective increase in NADPH production by increasing aconitase activity and thus could provide the substrate for CICD (51). Indeed, cells that overexpress CICD are protected against oxidative damage and UV phototoxicity (30, 36). However, this cytosolic pathway may also play a role in the regulation of cytoplasmic glutamate production, because {alpha}-ketoglutarate is converted to glutamate by glutamate dehydrogenase (GDH). Indeed, in yeast, aconitase deficiency causes glutamate auxotrophy because of the lack of {alpha}-ketoglutarate generation (53). GDH, an enzyme with numerous isoforms in mammalian tissues (56), is found in mitochondrial and other cellular compartments, including cytosolic extracts of retina (2, 10, 37, 57, 59). GDH is present in both the retina and the retinal pigment epithelium (42). Therefore, by regulating aconitase activity, iron could also regulate the downstream production of glutamate. It should be noted that some cells do not make citrate (neurons) and therefore depend on neighboring cells such as astrocytes for glutamate (27). The importance of a link between iron metabolism and glutamate secretion is underscored by the separate findings of dysregulation of glutamate and iron metabolism in neural and retinal diseases (1, 3, 11, 20).



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Fig. 1. Schematic of the cytosolic pathway for the production of glutamate. VGLUT1, vesicular glutamate transporter 1; c-aconitase, cytosolic aconitase.

 
Glutamatergic systems have recently been found in numerous nonneuronal tissues, including the gastrointestinal tract and the testes (24), osteoblasts (49), and pinealocytes (49). Glutamate has been implicated as having autocrine, paracrine, and intracrine functions in neuronal and nonneuronal cells. Thus there is a precedent for studying this system in other nonneuronal cells such as RPE and lens epithelial cells (LEC). The purpose of this study was to determine whether RPE and LEC secrete glutamate, whether iron availability affects this secretion, and whether glutamatergic systems are present in the RPE.


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Tissue culture. Dogs were obtained from the Johnston County, NC, animal shelter after they had been euthanized. The eyes were removed from the orbits within 3 h of death and cut 7–8 mm posterior to the limbus, and the posterior section of the globe was placed in a cup-shaped dissection dish to maintain the normal shape of the globe. After removal of the vitreous and the retina, 0.05% trypsin and 0.53 mM EDTA in Hanks' balanced salt solution without calcium or magnesium (catalog no. 25300-54; GIBCO) were added to the exposed RPE (44). After 37°C incubation for 20–40 min, patches of RPE were lifted and were transferred to a centrifuge tube containing growth medium, Ham's F-12-DMEM (1:1; GIBCO) with 20% fetal bovine serum [FBS; American Type Culture Collection (ATCC), Manassas, VA], and 1% antibiotic antimycotic solution (GIBCO). The cells were then collected by centrifugation at 125 g for 5 min. The pellets were transferred to 35-mm wells containing the growth medium and then incubated at 37°C in 95% air-5% CO2 and 95% humidity. Generally, cells from each eye filled three to five wells. The cultures become confluent after 8–12 days. At that point, the cells were trypsinized and plated in 6- or 12-well clusters, depending on the planned experiments. Confluent cultures were used for all experiments unless otherwise noted.

Cytokeratin immunoreactivity is a consistent marker for canine RPE. Therefore, to verify the identity of cultured cells as epithelial cells and the absence of fibroblast contamination, cells were stained for cytokeratins using mouse monoclonal anti-cytokeratin-8,13 (Sigma) (44). All of the cultured cells stained positively and there were no unstained cells, indicating that pure epithelial cell cultures were obtained.

Fresh RPE cells were obtained from enucleated dog eyes. The posterior segment had the vitreous and retina removed, and growth medium was added. Using microdissection forceps, large strips of the RPE layer were gently pulled away from Bruch's membrane. These strips were rinsed and transferred to ice-cold growth medium and collected by centrifugation at 125 g for 5 min. Protein was extracted from these freshly obtained RPE strips for immunoblot detection of vesicular glutamate transporter 1 (VGLUT1) protein as described below.

To culture canine LEC, the lens was removed from the anterior portion of the globe and the anterior capsule, with adherent epithelial cells dissected free and placed in a tissue culture dish with DMEM (GIBCO), 10% FBS (Hyclone, Logan, UT), and 1% antibiotic antimycotic solution. After significant outgrowth of epithelial cells from the capsule, the cells were dispersed and grown to confluence. They were then plated in six-well plates for the experiments.

Human RPE cells were obtained from ATCC (catalog no. CRL-2302). This spontaneously arising human cell line is designated ARPE-19. The culture and subculture conditions recommended by the ATCC were followed. The cells were cultured in DMEM-Ham's F-12 (catalog no. 30-2006; ATCC) with 10% FBS (catalog no. 30-2020; ATCC). Confluent cultures in six-well plates were used in the experiments. A human cortical neuronal cell line (HCN-2, catalog no. CRL-10742; ATCC) was cultured in DMEM (catalog no. 30-2002; ATCC) and 15% FBS (catalog no. 2020; ATCC). They were subcultured once and plated in 12-well clusters for the experiments. Cultures that were 80% confluent were used for the experiments.

Experimental conditions. When glutamate was measured in cell conditioned medium (CCM), all treatments were performed in serum-free, glutamine-free MEM (GIBCO). This medium does not have added iron. The cells are first rinsed with this medium before the experiments are started. It is important to perform these experiments in glutamine-free medium because glutamine can be taken up by cells and converted into glutamate. Riluzole, L-trans-pyrrolidine-2,4-dicarboxylic acid (PDC), and cis-1-aminocyclobutane-1,3-dicarboxylic acid were obtained from Tocris (Ellisville, MO). All other chemicals were obtained from Sigma.

The amount of iron added as ferric ammonium citrate (FAC) and ferric chloride (FeCl3) was 1.8 mg/l. This concentration is in the range of the pathological levels of iron in the intraocular fluids that we measured during intraocular inflammation (43). Furthermore, the amount of iron added as FAC was similar to that used by other investigators to create iron overload (38).

Samples of CCM were obtained at various times and frozen for future analysis of glutamate concentration. At the end of the experiments, cells were lysed with 10 mM Tris buffer, pH 7.4, containing 10 µl/ml phenylmethylsulfonyl fluoride (57 mM) and 6 µl/ml protease inhibitor cocktail (Sigma). After the cells were scraped from the wells and subjected to 20-min incubation on ice, the cell lysates were centrifuged at 14,700 g for 5 min. The supernatants were frozen for later measurement of protein using the bicinchoninic acid method (Pierce Biotechnology, Rockford, IL).

Glutamate analysis. Amplex Red assay kits (catalog no. A-12221; Invitrogen, Carlsbad, CA) were used to measure L-glutamate in the CCM (7). Standards and diluted CCM (minimum 12.5x dilution) were added to black-walled 96-well plates. The reaction mixture was added to the samples, followed by 30-min incubation at 37°C. Fluorescence was measured in a Fluoroskan Ascent FL fluorometer (Thermo Electron, Milford, MA) at 530-nm excitation and 590-nm emission. The fluorescence is proportional to the amount of glutamate in the sample. Glutamate concentration in the serum-free, glutamine-free MEM (not exposed to RPE) was <4 µM (the detection limit), and neither FAC, FeCl3, nor hemin affected the results of the glutamate assay (data not shown). In control experiments for FAC treatment, ammonium citrate was added and was found to have no effect on glutamate accumulation in CCM (data not shown). In experiments in which serial samples of the CCM were obtained from the same well over time, glutamate concentration was determined and expressed in micromoles. When cell lysates were obtained at the end of the experiment, the protein concentration of the lysates was measured and the data were normalized to micromoles per milligram of protein. This normalization allowed for more direct comparison to other cell types and systems.

Cytosolic aconitase. Mitochondria-free cytosolic extracts were obtained after the various treatments, and aconitase activity was measured. Absence of mitochondria was assessed by determination of the absence of cytochrome c. Plates containing treated RPE were placed on ice and washed twice with PBS. They were lysed in 50 mM HEPES (pH 7.4) containing 0.25 M sucrose and 0.007% digitonin (Sigma) for 5 min. Cells were scraped off the plates and ultracentrifuged at 230,000 g for 20 min at 5°C.

Aconitase activity in the lysates was measured by determining the disappearance of cis-aconitate from the cell lysates at 240 nm. Briefly, after the cells were treated and lysed, the lysates were concentrated using Centricon-30 filters centrifuged at 5,000 g for 1 h at 5°C, which reduced the volume by 80–90% and removed most of the sucrose. One-third of each lysate was added to 50 mM Tris·HCl buffer, pH 7.4, for a total volume of 490 µl. The temperature was equilibrated to 37°C. To start the reaction, 10 µl of 20 mM cis-aconitate (Aldrich) were added and absorbance was measured at 240 nm for 10 min at 37°C.

RT-PCR. Total RNA was isolated from canine LEC, canine RPE, and ARPE-19 using the Qiagen RNeasy Mini Kit (catalog no. 74101). Mouse brain total RNA was purchased from BD Biosciences Clontech (catalog no. 636601). Five micrograms of total RNA from canine LEC, canine RPE, and ARPE-19, as well as 1 microgram of total RNA from mouse brain, were used for RT-PCR with Ready-to-Go RT-PCR beads (catalog no. 27-9266-01; Amersham Biosciences). Primers for VGLUT1 amplification were 5'-GAGAAACAGCCGTGGGCAGAG-3' and 5'-TCAGTAGTCCCGGACAGGGGGTGG-3'. The expected product size was 207 bp. Primers for VGLUT2 amplification were 5'-AGCAAGGTTGGCATGTTGTCTG-3' and 5'-CGGTCCTTATAGGAGTACGCGT-3'. The expected product size was 698 bp.

Immunoblotting. Membrane fractions were prepared from canine RPE fresh tissue, canine RPE cultured cells, and ARPE-19 cells using SDS solubilization buffer consisting of 2% SDS, 1% DTT, and 1% 2-(N-hexylamino)ethanesulfonic acid. Cells were washed in PBS and homogenized in SDS solubilization buffer containing protease inhibitors. The homogenate was spun for 1 h at 200,000 g at 20°C. The resulting supernatant containing the membrane fraction was used for Western blot analysis. Mouse brain tissue extract was used as a positive control (Stressgen Biotechnologies, Victoria, BC, Canada). Total protein (20 µg) from mouse brain tissue extract, canine RPE cultured cells, and ARPE-19 cells as well as 90 µg of total protein from canine RPE fresh tissue, were subjected to 10% SDS-PAGE. After electrophoresis, the proteins were transferred to nitrocellulose membrane. The membrane was blocked with 2% nonfat milk powder in Tris-buffered saline containing Tween 20 (TBST) and then probed with antibodies against VGLUT1 (1 µg/ml; Alpha Diagnostic, San Antonio, TX). After being washed with TBST, the membrane was incubated with horseradish peroxidase-conjugated anti-rabbit IgG antibodies (1:5,000 dilution; Alpha Diagnostic). Proteins were visualized using the enhanced chemiluminescence detection system (Amersham, Little Chalfont, UK) according to manufacturer's protocol.


    RESULTS AND DISCUSSION
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Although RPE cells have glutamate receptors (39, 42) and transporters (5, 47), secretion of glutamate by these cells has not been demonstrated previously. Cultured RPE were placed in glutamine-free/serum-free MEM (an iron-free medium) for these experiments to eliminate the possibly confounding contribution of glutamine uptake and metabolism to glutamate in these cells. In Fig. 2A, the 2-, 6-, and 24-h time points represent data obtained in one set of experiments. This figure clearly shows that glutamate accumulates in the CCM over time. The 24- and 48-h time points are normalized to protein content (Fig. 2B) to allow for comparison with other studies. Note that the 48-h time point represents results from a separate set of experiments. Therefore, because of variation in control secretion between experimental sets, the 48-h time point cannot be compared directly with the 24-h time point.



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Fig. 2. Iron and glutamate secretion by retinal pigment epithelial (RPE) cells. The effect of ferric ammonium citrate (FAC; 33 µM) and ferric chloride (FeCl3; 33 µM) or hemin (10 µM) on the accumulation of glutamate in the medium overlying cultured RPE cells is shown. A: samples of the medium were taken at the indicated times after the beginning of treatment. Glutamate concentration (µM) in the medium samples was determined. B: results of two independent sets of experiments are shown, with the amount of glutamate in the cell conditioned medium (CCM) normalized to the amount of protein present at the end point of the experiments (either 24 or 48 h). Histogram bars represent means ± SE of six samples. Error bars <1.0 (A) or <0.1 (B) are not shown. *P < 0.05, significantly different from control at each time point (ANOVA with Tukey's test).

 
The accumulation of glutamate in the CCM is significantly increased by addition of iron in three different forms, with each form having a unique time course. FAC had no effect at 2 h but increased glutamate accumulation in CCM at the 6-, 24-, and 48-h time points. The increase at 6 h was only 10% greater than that of the control (Fig. 2A); however, at 24 h, the increase was 158% of control, and by 48 h, FAC had increased accumulation of glutamate to five times that of control (Fig. 2B). FeCl3 also significantly increased glutamate accumulation as early as 6 h and continued to increase it at 24 h; however, glutamate accumulation was no different from control at 48 h. Yet another pattern emerged when cells were treated with hemin as the iron source. At the early time points, glutamate accumulation was decreased by almost 50% (Fig. 2A), while at the later time points, glutamate accumulation was significantly increased. The reasons for the depression of glutamate accumulation caused by hemin at the early time points remain unclear. However, the gradual increase in glutamate accumulation at the later time points may be due to a lag period between the addition of hemin and its catabolism by heme oxygenase, which would make iron available by releasing it from the porphyrin ring. In contrast to the effects of addition of iron to RPE cells, treatment of cells with the iron chelator {alpha}-{alpha}-dipyridyl caused a significant decrease in glutamate accumulation in the CCM (from 26 ± 3.0 to 22 ± 2.6 µM at 4 h; P < 0.003, paired t-test).

All effects on glutamate accumulation in the CCM caused by changes in iron load were induced relatively slowly compared with the effects of receptor agonists on synaptic release of glutamate. The movement of iron between sites of storage and utilization and the kinetics of this movement are very complex and poorly understood. Our conclusion on the basis of the present study is that the consequences of altering intracellular iron pools is gradually translated into effects on the numerous systems affected by iron availability, including aconitase activity.

It is unlikely that glutamate secreted by RPE cells came from intracellular stores. The amount of glutamate within the cells (from 9.5-cm2 wells; n = 6) was 18 nmol, while the total amount of glutamate that accumulated in the CCM was 86 nmol at 24 h (n = 6) and was 159 nmol after FAC treatment (n = 6). Therefore, most of the glutamate that accumulated in the CCM was newly formed.

The results of our previous studies of iron uptake by LEC (18) indicated that there was more than one intracellular pool of iron. The size of these pools was dependent on the source of iron, which was differentially available for incorporation into ferritin. For example, FAC stimulated iron uptake and iron incorporation into ferritin when transferrin was the source of radioactively labeled iron. In contrast, FAC decreased iron uptake and incorporation into ferritin when FeCl3 was the source. It appears that FAC competed with FeCl3 for uptake and incorporation into ferritin, but not with iron entering the cell from transferrin. Therefore, iron availability for incorporation into the iron sulfur cluster as well as the activation of c-aconitase and subsequent increase in glutamate production may also be source dependent and may be influenced differentially by different pools of iron within the cells.

Three additional cell types were tested to determine whether the FAC effect was limited to primary canine RPE cultures (Table 1). Glutamate accumulated in CCM from a cultured human RPE cell line (ARPE-19), primary cultures of canine LEC, and a human cortical neuronal cell line (HCN-2). In all cases, this accumulation was augmented by FAC treatment.


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Table 1. FAC increases glutamate secretion in a human RPE cell line, primary cultures of canine lens epithelial cells, and cultures of a human cortical neuronal cell line

 
To determine the contribution of c-aconitase to the accumulation of glutamate in the CCM, RPE were exposed to oxalomalate (OMA), an inhibitor of aconitase and CICD (16, 60), in the presence and absence of FAC. OMA decreased basal glutamate secretion by ~60% and completely inhibited the FAC-induced increase in glutamate accumulation in the CCM. Because iron is also known to regulate mitochondrial aconitase, a possible contribution of this enzyme to glutamate production cannot be ruled out. These findings indicate that much of glutamate accumulation in CCM results from production via the aconitase pathway and that FAC's effects on glutamate accumulation in the CCM were mediated entirely through an increase in aconitase activity (Fig. 3A).



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Fig. 3. The effect of inhibition of c-aconitase (oxalomalate, OMA; 5 mM) at 24 h (A), inhibition of regulated release of glutamate by riluzole (R) at the times and concentrations indicated (B), and inhibition of glutamate transporter by L-trans-pyrrolidine-2,4-dicarboxylic acid (PDC; 5 mM) at 24 h (C) on the accumulation of glutamate in the medium overlying cultured RPE cells. Samples of the medium were obtained at the indicated times after treatment was initiated. Glutamate concentration in the medium samples was determined, and the amount of protein per well was determined as indicated in A and C. Histogram bars represent means ± SE of at least five samples. A: *P < 0.05, significantly different from control and FAC treated cells; **significantly different from all other groups. B: *P < 0.05, significantly different from control; **P < 0.05, significantly different from control and 0.063 mM riluzole. C: *P < 0.05, significantly different from control and FAC + PDC-treated cells; **P < 0.05, significantly different from all other groups. Statistical tests for all data included ANOVA and Tukey's test.

 
While the effects of iron on aconitase activity are well known (21, 31), it was important to demonstrate that changes in the activity of this enzyme in response to alterations in iron availability paralleled changes in glutamate secretion in RPE. Therefore, the activity of c-aconitase was determined in mitochondria-free cell lysates by measuring the disappearance of added cis-aconitate. Aconitase activity in the cytosolic extracts of RPE was significantly increased by FAC (Table 2). Furthermore, incubation of the cells with the iron chelator dipyridyl significantly reduced aconitase activity, and OMA treatment completely abolished the activity of this enzyme (Table 2). These changes in aconitase activity paralleled changes in glutamate secretion resulting from the same treatments. On the basis of these results, we conclude that iron-induced changes in aconitase activity drive the production of glutamate and its subsequent secretion.


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Table 2. Measurement of aconitase activity in RPE

 
In neurons, glutamate is released at the synapse from preformed storage vesicles. A recent report indicated that RPE cells express significant amounts of syntaxins 1A and 1B and synaptobrevin-2, all of which are typically restricted to neurons and neuroendocrine cells (40). These proteins are involved in calcium-regulated exocytosis of synaptic vesicles. To determine whether regulated release of glutamate from RPE contributed to glutamate accumulation in CCM, riluzole was used to inhibit voltage-gated channel-regulated glutamate release (14). At the 2-h time point, both doses of riluzole significantly reduced the accumulation of glutamate in the CCM (Fig. 3B). This reduction was still apparent at 4 and 6 h, at which times the effects were concentration dependent.

There are three types of vesicular glutamate transporters found in neuronal cells: VGLUT1, VGLUT2, and VGLUT3 (17, 28). These transporters are responsible for the uptake of glutamate into vesicles in preparation for transport to the membrane for release. Further evidence supporting the presence of a glutamatergic system in nonneuronal cells comes from our data showing the presence of VGLUT1 protein expression (Fig. 4A) in both RPE and LEC; VGLUT2 was not found in these cells. An immunoblot of VGLUT1 was positive for cultured canine RPE and human RPE (ARPE-19) as well as freshly obtained canine RPE (Fig. 4B). The fact that freshly obtained canine RPE and cultured canine RPE both contained VGLUT1 protein indicates that the presence of this vesicular protein in cultured canine RPE is not an artifact of culture conditions. These results support the hypothesis that at least some of the glutamate released by RPE and LEC is regulated by mechanisms similar to those occurring in neuronal cells.



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Fig. 4. A: RT-PCR analysis of VGLUT1 and VGLUT2 expression in canine lens epithelial cells (LEC) and RPE cells as well as in human RPE cells. Mouse brain total RNA was used as a positive control for both VGLUT1 and VGLUT2. B: immunoblot analysis of VGLUT1 with proteins obtained from the membrane fraction of mouse brain (control), canine RPE (freshly obtained from enucleated eyes), cultured canine RPE cells, and cultured human RPE cells (ARPE-19).

 
To study potential pathways for reuptake of secreted glutamate, a specific glutamate transporter inhibitor, PDC (6), was used in the presence or absence of FAC. RPE were incubated for 24 h with the indicated treatments. As shown earlier in Fig. 2, A and B, FAC significantly increased glutamate accumulation in the CCM. Treatment with PDC also increased the accumulation of glutamate in the CCM and significantly enhanced FAC-induced glutamate accumulation (Fig. 3C). Another inhibitor of glutamate transporters, cis-1-aminocyclobutane-1,3-dicarboxylic acid (35), also increased glutamate accumulation in the CCM (from 17 ± 0.7 to 42 ± 2.5 µM at 24 h; P < 0.004, paired t-test). The increased accumulation of glutamate in the CCM in the presence of either inhibitor indicates that RPE cells can recycle the glutamate that they secrete. However, the data suggest that the recycling is not completely efficient, because glutamate continues to accumulate in the CCM over time, and this accumulation is further enhanced by iron.

Glutamate secretion by RPE and LEC could have both autocrine and paracrine functions. When added to preparations of RPE and retina, glutamate stimulated reattachment of photoreceptors to RPE and increased shedding and phagocytosis of photoreceptor discs (12, 19). Glutamate can also increase RPE proliferation (58). Glutamatergic systems have recently been found in other nonneuronal tissues, including pinealocytes (50), islets of Langerhans (25), stomach, intestine, and testis (25). The present study provides evidence for the presence of this system in two ocular epithelial cell types: RPE cells and LEC. Therefore, regulation of glutamate metabolism by iron may represent a universal phenomenon of primary importance to cells of diverse origin. The roles played by glutamate in the physiology and pathology of these tissues need further study.

The results of the present study clearly demonstrate that iron-induced changes in aconitase activity parallel changes in glutamate production and accumulation in the CCM. Interestingly, iron-deficient anemic rats have significantly lower serum glutamate and impaired behavioral function (33). There are other potential links between iron metabolism and glutamate secretion in pathological situations. For example, iron derived from heme deposited during hemorrhage in the brain and in the retina could also affect the activity of this pathway and increase glutamate secretion. The present results obtained with hemin indicate that there could be long-term effects on glutamate production, which would be dependent on the relatively slow release of iron from heme. In other studies, ischemia-reperfusion injury altered both iron availability (4) and glutamate secretion (55). In the retina, an imbalance in iron homeostasis has been associated with degeneration of this tissue (20, 61).

There are numerous neurodegenerative conditions in which iron (62)and glutamate metabolism, storage, and secretion are dysregulated. For example, the concentration of iron in the brain increases with age (3, 22) and in Alzheimer's disease (11, 23), Huntington's disease (8), and Parkinson's disease (13). All of these neurological diseases are also characterized by alterations in glutamate secretion (1). In this regard, patients with Parkinson's disease were found to have increased iron content in the nigrostriatum, but not increased ferritin levels (15). It was concluded that iron accumulation was compartmentalized in such a way that the elevated iron content was not available to influence ferritin synthesis. There are a number of important questions that need to be answered before investigators can get closer to making a firm connection between iron, glutamate, and neurodegeneration. For example, does the concentration of iron available to alter aconitase activity vary sufficiently to effect a physiologically significant change in glutamate concentration in the brain?

We suggest that a derangement in the normal movement of iron between intracellular compartments not only would increase the availability of iron for participation in damaging free radical reactions but also could alter the activity of c-aconitase and glutamate synthesis. Therefore, studies of the regulation of iron movement between compartments are needed to provide information important to understanding the interface between iron availability, aconitase activity, and glutamate formation. Such knowledge will enhance the ability to target more specifically the development of therapies.


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This work was supported by National Eye Institute Grant EY-04900-22 (to M. C. McGahan) and by funds from the State of North Carolina.


    FOOTNOTES
 

Address for reprint requests and other correspondence: M. C. McGahan, Dept. of Molecular Biomedical Sciences, North Carolina State Univ., 4700 Hillsborough St., Raleigh, NC 27606 (E-mail: chris_mcgahan{at}ncsu.edu)

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.


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