Candidate Review:

The Influence of GABA Metabolism on GABA Neurotransmission: The Role of Metabolic Regulatory Points and a Neuronal Glutamate Transporter

Ernesto Solis, Jr.

Neuroscience Graduate Program, Vanderbilt University School of Medicine, U1205 Medical Center North, Nashville, TN 37232, USA.
Correspondence e-mail: ernesto.solis@vanderbilt.edu

Abstract | Full Text | PDF

Excessive excitatory drive in the brain is thought to underlie diseases such as epilepsy.  One approach in the development of novel treatments for conditions characterized by hyper-excitability is the enhancement of GABA-mediated inhibition.  While most current medical interventions target GABAergic neurotransmission postsynaptically (e.g. benzodiazepines, barbiturates), much less is known about potential presynaptic therapeutic targets at the GABAergic synapse. This review describes recent findings that have exemplified presynaptic mechanisms that may provide the basis for the development of novel treatments to alter inhibitory neurotransmission.  GABA metabolism is summarized with an emphasis on the role of presynaptic regulatory points in GABA synthesis.  In addition, the excitatory amino acid transporter 3 (EAAT3), which is thought to provide the substrate for GABA synthesis, will be described in detail.  Finally, EAAT3 is presented as a potential therapeutic target to modulate GABA-mediated inhibition presynaptically, and the most recent findings on EAAT3’s functional regulation by several key players are reviewed.

GLUTAMATE AND GABA METABOLISM

Glutamate and GABA are the major excitatory and inhibitory neurotransmitters in the brain, respectively.  Unlike other neurotransmitter systems, such as monoamines, reuptake and recycling of glutamate and GABA does not appear to be as important as new synthesis to replenish the pool of neurotransmitter for synaptic vesicle filling.  Despite the critical importance of neurotransmitter supply, either to prevent depletion and maintain stable transmission or perhaps to dynamically adjust in response to demand, the metabolic pathways by which these transmitters are continuously supplied to synaptic terminals have not been resolved.

GABA is synthesized by the decarboxylation of glutamate, which is catalyzed by the enzyme glutamic acid decarboxylase (GAD).  Neurons are not capable of synthesizing glutamate1 on their own; therefore inhibitory neurons, like excitatory neurons, need a supply of glutamate.  At least two possible pathways through which glutamate may be acquired exists:  (1) the direct uptake of extracellular glutamate or (2) the uptake of glutamine, which can be converted to glutamate by neurons.  Transporters serving both of these roles are expressed by GABAergic neurons, and both have been demonstrated to play roles in the synthesis of GABA2-5.

GABA synthesis and synaptic vesicle filling are tightly coupled processes as revealed by biochemical assays.  GAD65, the synaptically localized isoform of GAD, is associated with a complex of proteins on synaptic vesicles that includes the vesicular GABA transporter6.  GABA synthesized from glutamate is taken up into synaptic vesicles preferentially over pre-existing GABA7.  Electrophysiological studies demonstrated that inhibiting GAD results in a reduction in the size of miniature synaptic events, which represent the amount of GABA released from a single synaptic vesicle8.  In contrast, knock-out of the predominant membrane transporter for GABA reuptake does not influence the size of these miniature events9.  Taken together, these findings suggest that new synthesis is more important than recycling of existing GABA.  Moreover, they demonstrate that any factors influencing GABA synthesis are likely to play an important role in maintaining, and possibly regulating, inhibitory synaptic transmission.  Finally, GABA is catabolized by the action of GABA transaminase (GABA-T), which deaminates GABA to make succinic semialdehyde (SSA), and then SSA dehydrogenase (SSADH) converts SSA to succinate, which enters the TCA cycle.  SSA can also be converted to γ–hydroxybutyrate (GBH) by the action of SSA reductase10 (Figure 1).

HOW IS SUBSTRATE OBTAINED FOR PRODUCTION OF GLUTAMATE AND GABA?

Supply of substrate to inhibitory neurons for GABA synthesis is mediated by highly regulated, sodium-dependent solute transporters, which carry metabolites against their concentration gradients.  EAAT3, a member of the high affinity glutamate transporter (excitatory amino acid transporter, EAAT) family, is expressed on somatodendritic compartments of excitatory neurons, and at axon terminals of GABAergic neurons11, 12.  SNAT1 and 2 are members of the sodium coupled neutral amino acid transporter (SNAT) family and are expressed on excitatory and inhibitory neurons13.  Because of their substrate affinity and the high ambient concentration of glutamine in the brain, SNAT1 and 2 are likely the major glutamine uptake pathway for neurons14.

Astrocytes express glutamate transporters EAAT1 and 2 (also known as GLAST and GLT-1, respectively), of which EAAT2 is the major transporter for the clearance of synaptically released glutamate15, as well as SNAT3 and 513.  A glutamine-glutamate-GABA cycle has been proposed1 in which glutamate released from neurons is taken up by astrocytes, converted to glutamine with the enzyme glutamine synthetase and subsequently transported back to neurons (Figure 1).  In neurons, glutaminase type I (GLS1) has been proposed to be the enzyme that converts glutamine to glutamate 1.  Studies have suggested that the majority of neurotransmitter glutamate is recycled by GLS1 with minimal contribution of de novo synthesis of glutamate from α-ketoglutarate in excitatory neurons16, 17.  Interestingly, when GLS1 was knocked out, GABA levels were not reduced18, suggesting alternative metabolic pathways for GABA synthesis must be present in inhibitory neurons.

Figure 1 | Metabolic pathways of GABA. Acquisition of substrate for GABA synthesis, GABA packaging into synaptic vesicles, and breakdown of GABA by respective proteins

In acute hippocampal slices, pharmacological inhibition of either EAATs or SNATs results in a rapid reduction of GABA vesicle content, as measured by changes in miniature inhibitory postsynaptic current (mIPSC) amplitudes3, 19.  These results suggest that a dynamic equilibrium exists between GABA synthesis and vesicular filling, and consequently that inhibitory synaptic strength is directly regulated by two different substrate supply pathways.  These results also suggest that substrate supply is a key regulatory point in the determination of inhibitory synaptic strength.

THERAPEUTIC IMPLICATIONS OF REGULATING INHIBITORY NEUROTRANSMISSION: WHAT KNOCKOUT STUDIES TELL US ABOUT THE ROLE OF KEY PROTEINS INVOLVED IN THE REGULATION OF GABA SYNTHESIS (summarized in Table 1)

The importance of the regulatory points of GABA synthesis is revealed by knock-out mouse studies.  After glutamate is cleared by glutamate transporters on glia, glutamine synthetase (GS) converts glutamate to glutamine to recycle neurotransmitter for both excitatory and inhibitory neurotransmission.  In the context of an entire organism, the conversion of glutamate to glutamine results in detoxification of ammonia.  As expected, knocking out GS has a severe phenotype, with death occurring at embryonic day 3.520.  Since GS has a global role beyond the central nervous system, it is not likely to be a good therapeutic target to alter GABA metabolism. 

As mentioned above, GLS1 is the enzyme thought to convert glutamine to glutamate in neurons.  The GLS1 knock-out mouse dies within a day of birth, is slightly smaller than wild type, has impaired respiratory function, and is deficient in goal-directed behavior18.  It is thought that respiratory acidosis causes respiratory impairment and subsequent death. 

There are 2 isoforms of GAD, GAD65 and GAD67, named after their molecular weights of 65 and 67 kDa, respectively.  These 2 isoforms are encoded by independent genes and have different subcellular localizations in inhibitory neurons.  As mentioned earlier, GAD65 is found at synapses.  One study showed that GAD65 knock-out mice are more likely to develop seizures than wild type mice21.  A different GAD65 knock-out mouse had an epileptic phenotype characterized by spontaneous seizures that led to death22.  This mouse also showed increased anxiety-like behaviors and diminished response to anxiolytics23, pre-pulse inhibition deficits24, upregulation of the vesicular GABA transporter, and increased cytosolic GABA transport into synaptic vesicles25.

GAD67 localizes to the cell soma of inhibitory neurons.  The GAD67 knock-out mouse shows a reduction in GABA levels throughout the brain, a reduction in GAD activity, and severe cleft palate, which leads to death within 24 hours of birth26.  It is thought that the reduction of GABA levels in the GAD67 knock-out mouse brainstem to 30% of wild type leads to a malfunction in the respiratory control system and subsequent death27.  The GAD65/GAD67 double knock-out mouse dies after birth due to cleft palate.  GABA levels are low in this mouse28 and another study with a different GAD65/GAD67 double knock-out mouse determined that GABA synthesis is absent29.

The vesicular GABA transporter (vGAT) fills synaptic vesicles with both inhibitory neurotransmitters GABA and glycine.  Study of a vGAT knock-out mouse showed that this mouse is incapable of executing vesicular release of GABA and glycine29.

GAT1 is the predominant transporter responsible for GABA reuptake into inhibitory terminals, which allows for termination of GABA transmission.  The GAT1 knock-out mouse showed reduced anxiety-like and depression-like behaviors30, as well as decreased aggression31, tremor, ataxia, nervousness, and increased extracellular GABA levels, which led to enhanced tonic inhibition and diminished phasic inhibition9.  GAT1 has been targeted therapeutically by drugs, such as tiagabine, to treat epilepsy and anxiety32.

GABA-T and SSADH perform a 2-step enzymatic breakdown of GABA.  While there is no knock-out mouse for GABA-T, drugs that inhibit GABA-T, such as vigabatrin, have been used to increase GABA levels in the brain.  Vigabatrin is not a drug of choice for epilepsy treatment because it often causes visual field defects33.  The SSADH knock-out mouse exhibits ataxia, absence-like seizures with ictal behavior characterized by facial myoclonus, vibrissal twitching, and frozen immobility at 2 weeks.  At this time, the absence seizures become more severe evolving into generalized convulsive seizures that progress into lethal status epilepticus34, 35.  This mouse provides a good model for SSADH deficiency seen in humans36, which is characterized by absence seizures and mental retardation.

THE ROLE OF EAAT3 ON INHIBITORY NEUROTRANSMISSION, SEIZURES, AND EPILEPSY

A role for EAAT3 as a critical regulator of neuronal excitability in vivo was demonstrated using antisense knock-down of the transporter37-42.  In this study, antisense oligonucleotides to EAAT3 were infused into the lateral ventricles of rats, adjacent to the hippocampus.  The animals developed spontaneous seizures corresponding to the time course of EAAT3 protein level reduction.  Biochemical analysis confirmed reduced GABA content and impaired GABA synthesis in hippocampal tissue from treated animals.  These results suggest that EAAT3 mediates an endogenous negative feedback mechanism whereby increased extracellular glutamate enhances GABA synthesis and inhibitory synaptic strength. The EAAT3 knock-out mouse did not have an epileptic phenotype, but it did develop dicarboxylic aminoaciduria and behavioral abnormalities43.  In addition, this mouse develops glutathione deficiency and shows age-dependent neurodegeneration44.  The discrepancy in the phenotypes between the knock-out mouse and the knock-down rats, in particular the epileptic phenotype, has been attributed to a compensatory upregulation of a glutamate transporter homologous to EAAT3 in the knock-out mouse4

Table 1 | Summary of knock-out studies of key regulatory points in GABA metabolism.



Several studies have investigated changes in EAAT3 expression in a variety of chronic epilepsy models and in human epileptic brain. Most studies of seizure models have looked at EAAT3 expression changes at least 24 hours after seizure induction45, and these studies have examined changes at the tissue level which would predominantly reflect expression changes in the more numerous excitatory neurons. Moreover, the results of these studies were inconsistent, possibly due to differences in the epilepsy models, measurements (mRNA vs. protein) and regions examined.  The only study of acute changes showed that EAAT3 protein in hippocampal pyramidal neurons appears to internalize 6 hours after kainic acid seizure induction46. No study has examined the expression of EAAT3 by inhibitory neurons in seizures and epilepsy.  Because EAAT3 is expressed at postsynaptic sites on excitatory neurons and presynaptically on inhibitory neurons, it seems reasonable to hypothesize that seizure activity will have distinct effects on these two pools of transporters.

Recent evidence suggests that one function of glutamate transporters on inhibitory neurons, potentially EAAT3, may be the dynamic regulation of inhibition by extracellular glutamate levels47.  Glutamate has been reported to increase extracellularly prior to seizure onset in human brain48.  Therefore, EAAT3 may function to prevent the onset of seizures or to curtail seizure activity once started through enhancement of inhibition.

REGULATION OF EAAT3 SURFACE EXPRESSION AND GLUTAMATE UPTAKE BY SIGNALING CASCADE MOLECULES

Signaling cascade-mediated regulation of EAAT3 in inhibitory neurons could allow for modulation of inhibitory neurotransmission.  A number of studies reported that EAAT3 activity is regulated by signaling cascade molecules.  Most of these studies used heterologous settings including the C6 glioma cell line, which expresses EAAT3 endogenously.  Regulation of neuronal EAAT3 in an endogenous setting and its effects on neuronal glutamate uptake are less well described.  One study demonstrated functional upregulation of EAAT3 activity following induction of long term potentiation in the hippocampus and subsequent translocation of the protein from a cytosolic to a membrane compartment49.

Glutamate uptake via EAAT3 is increased by non-specific activation of protein kinase C (PKC) with phorbol-12,13-myristate (PMA) in C6 glioma cells50 and in primary neuronal cultures46, 50.  This increase in glutamate uptake is associated with a rapid increase in EAAT3 surface expression in both C6 glioma cells and neurons46.  Bisindolylmaleimide II (Bis II), a PKC inhibitor, completely blocks the PMA-induced glutamate uptake, but has no effect on basal glutamate uptake levels51.  PKC activation mimicked the LTP induced upregulation of EAAT349.

PKC activity is mediated by a family of three subgroups (classic, novel, and atypical PKCs) each having unique properties.  Classic PKC (cPKC) subtypes, which include three members (α, β, and γ), require calcium as a co-factor and are activated by diacylglycerol (DAG) and phorbol esters.  In both C6 glioma cells and cortical neurons, the PMA-induced increase in EAAT3 activity was blocked with Gö6976 (10 μM), a selective inhibitor of cPKC subtypes52.  Of the three cPKC subtypes, C6 glioma cells only express PKCα, suggesting that PKCα is the cPKC subtype that plays a role in the regulation of EAAT3 activity.  In addition, C6 glioma cells treated with PMA showed a direct interaction between PKCα and EAAT3 on the cell surface52.  Rat brain synaptosomes show basal EAAT3-PKCα association in the absence of PMA, while PMA treatment induced additional EAAT3-PKCα association.  Both effects in synaptosomes are blocked by PKC antagonists suggesting the association may be triggered by endogenous stimulation of PKC activity under physiological conditions53.

Phospatidylinositol 3-kinase (PI3K) has also been shown to regulate EAAT3 activity.  Wortmannin, an irreversible PI3K inhibitor, decreases glutamate uptake and EAAT3 cell surface expression in C6 glioma cells within minutes.  Platelet-derived growth factor (PDGF), which stimulates PI3K activity, increases both the activity and cell surface expression of EAAT3.  The PDGF-mediated increase in EAAT3 activity is not blocked by the PKC antagonist BisII, and the PMA-mediated increase in glutamate uptake is not blocked by wortmannin suggesting that at least two independent signaling pathways regulate EAAT3 activity50.

Protein kinase A (PKA) has been shown to regulate EAAT3 activity.  In primary neuronal cultures glutamate uptake and EAAT3 surface expression decrease after treatment with H89, a PKA inhibitor.  The H89-mediated decrease in glutamate uptake was counteracted by pre-treating cells with forskolin, a PKA activator54.

As mentioned earlier, EAAT3 is expressed at GABAergic terminals and glutamatergic postsynaptic sites, but there are few studies that examined the functional regulation of EAAT3, and these studies have primarily looked at the postsynaptically localized EAAT3.  To our knowledge, no functional studies of EAAT3 at GABAergic terminals have been conducted.

REGULATION OF EAAT3 ACTIVITY BY PRESYNAPTIC RECEPTORS AT GABAERGIC TERMINALS: METABOTROPIC GLUTAMATE RECEPTORS AND OPIOID RECEPTORS

Metabotropic glutamate receptors (mGluRs) have multiple effects on interneurons through their actions on somata and axon terminals.  In general, group I mGluRs (mGlu1 and mGlu5) are located on somatodendritic compartments, and group II/III mGluRs are located on presynaptic terminals, although there are many exceptions55.  Presynaptic mGluRs on inhibitory terminals are activated by glutamate that is released from neighboring excitatory synapses56.  When extracellular glutamate levels are sufficient to reach transporters on inhibitory terminals, it is likely that presynaptic mGluRs would be activated as well.  mGlu1 agonists activate PKC in hippocampal pyramidal cells57, but the signaling pathways activated in interneurons are not known.  Interestingly, group I agonists are generally pro-convulsant in vitro and in animal models58.  Investigation of possible regulation of EAAT3 activity on GABAergic neurons by mGluRs may provide important insights into the endogenous signaling mechanisms underlying a crosstalk between excitatory and inhibitory neurotransmission.

Opioid receptors are members of the superfamily of G-protein-coupled receptors that utilize inhibitory G-proteins (Gi/o).  After Gi/o are stimulated by opioid receptors, multiple effectors are activated including adenylyl cyclase and mitogen-activated protein kinase54, 59.  Also, activation of Gi/o leads to inhibition of cAMP production and PKA activity60.  Inhibition of PKA has been associated with decreases in glutamate uptake and glutamate transporter surface expression levels by neurons50.  Of the three well-characterized opioid receptors (Mu-, Delta-, and Kappa-opioid receptors), Mu-opioid receptor (MOR) and Delta-opioid receptor (DOR) are highly expressed in cortex61, 62 and hippocampus63-65.  While it seems that DOR is expressed by both excitatory and inhibitory neurons66, MOR is preferentially expressed in axonal and somatodendritic compartments of GABAergic neurons in the hippocampus.  In addition, both MOR and DOR localize to GABAergic neurons in dissociated cortical and hippocampal cultures67.

Recently, it was shown in EAAT3-expressing Xenopus oocytes that co-expressing increasing amounts of DOR decreased glutamate uptake and EAAT3-mediated currents.  In addition, DOR and EAAT3 can be co-immunoprecipitated and co-localized in both Xenopus oocytes and in rat cultured hippocampal neurons, suggesting a direct interaction between EAAT3 and DOR.  Activation of DOR with pre-treatment of [D-Pen2,5]-enkephalin (DPDPE), a DOR agonist, counteracted the reduction in glutamate uptake and EAAT3-mediated current in Xenopus oocytes, and co-localization in both Xenopus oocytes and hippocampal neurons68.  It is possible that DOR inactivates EAAT3 by its direct interaction and when DOR is stimulated by DPDPE, EAAT3 is released and allowed to increase its activity.  This is further indication that EAAT3 could be regulated by G-protein coupled receptors.  In addition, strong evidence suggests that protein kinase B (Akt), a downstream target of PI3K, regulates EAAT3 activity69.  Akt is a downstream target of DOR in T cells making DOR a strong candidate for the regulation of EAAT370.  Whether MOR also regulates EAAT3 activity has not been investigated, however, its selective expression by GABAergic neurons suggests the possibility that, if it does, it may provide a signaling mechanism to selectively regulate EAAT3 on inhibitory neurons. Therefore, from a potential therapeutic perspective, MOR may be the most interesting candidate for the regulation of glutamate transporter activity in GABAergic neurons.

CONCLUSIONS

This review highlights the importance of the regulation of neurotransmitter GABA metabolism.  In order to make progress towards the development of novel therapeutic targets for inhibitory neurotransmission, candidate therapeutic targets at GABAergic terminals, such as EAAT3, must be studied.  Through EAAT3’s possible regulation by signaling cascade molecules and specific receptors, the activity of EAAT3 could be manipulated in order to alter glutamate uptake, GABA synthesis, and ultimately, inhibitory neurotransmission.  Additionally, regulatory points in GABA synthesis, such as GAD and vGAT, which can adjust GABA levels and alter inhibitory neurotransmission, need to be further explored.

REFERENCES

1.   Bak LK, Schousboe A and Waagepetersen HS (2006). The glutamate/GABA-glutamine cycle: aspects of transport, neurotransmitter homeostasis and ammonia transfer. J Neurochem. 98 (3): 641-653.

2.   Liang SL, Carlson GC and Coulter DA (2006). Dynamic regulation of synaptic GABA release by the glutamate-glutamine cycle in hippocampal area CA1. J Neurosci. 26 (33): 8537-8548.

A recent study that shows glutamine transporters play a role in the synthesis of neurotransmitter GABA under certain stimulation protocols.

3.   Mathews GC and Diamond JS (2003). Neuronal glutamate uptake Contributes to GABA synthesis and inhibitory synaptic strength. J Neurosci. 23 (6): 2040-2048.

Physiological study, which shows glutamate uptake by neurons leads to increased GABA synthesis.

4.   Sepkuty JP, Cohen AS, Eccles C, Rafiq A, Behar K, Ganel R, Coulter DA and Rothstein JD (2002). A neuronal glutamate transporter contributes to neurotransmitter GABA synthesis and epilepsy. J Neurosci. 22 (15): 6372-6379.

This study shows that knocking down EAAT3 yields an epileptic phenotype, which might be associated with decreased GABA levels in the hippocampus.

5.   Fricke MN, Jones-Davis DM and Mathews GC (2007). Glutamine uptake by System A transporters maintains neurotransmitter GABA synthesis and inhibitory synaptic transmission. J Neurochem. 102 (6): 1895-1904.

6.   Hsu CC, Davis KM, Jin H, Foos T, Floor E, Chen W, Tyburski JB, Yang CY, Schloss JV and Wu JY (2000). Association of L-glutamic acid decarboxylase to the 70-kDa heat shock protein as a potential anchoring mechanism to synaptic vesicles. J Biol Chem. 275 (27): 20822-20828.

7.   Jin H, Wu H, Osterhaus G, Wei J, Davis K, Sha D, Floor E, Hsu CC, Kopke RD and Wu JY (2003). Demonstration of functional coupling between gamma -aminobutyric acid (GABA) synthesis and vesicular GABA transport into synaptic vesicles. Proc Natl Acad Sci U S A. 100 (7): 4293-4298.

In addition to the finding that GABA synthesis is coupled to vGAT, this study shows that GABA that is newly synthesized from glutamate is taken up into synaptic vesicles preferentially over pre-existing GABA.

8.   Engel D, Pahner I, Schulze K, Frahm C, Jarry H, Ahnert-Hilger G and Draguhn A (2001). Plasticity of rat central inhibitory synapses through GABA metabolism. J Physiol. 535 (Pt 2): 473-482.

9.   Jensen K, Chiu CS, Sokolova I, Lester HA and Mody I (2003). GABA transporter-1 (GAT1)-deficient mice: differential tonic activation of GABAA versus GABAB receptors in the hippocampus. J Neurophysiol. 90 (4): 2690-2701.

10. Meldrum B (1982). Pharmacology of GABA. Clin Neuropharmacol. 5 (3): 293-316.

11. He Y, Janssen WG, Rothstein JD and Morrison JH (2000). Differential synaptic localization of the glutamate transporter EAAC1 and glutamate receptor subunit GluR2 in the rat hippocampus. J Comp Neurol. 418 (3): 255-269.

12. Rothstein JD, Martin L, Levey AI, Dykes-Hoberg M, Jin L, Wu D, Nash N and Kuncl RW (1994). Localization of neuronal and glial glutamate transporters. Neuron. 13 (3): 713-725.

13. Mackenzie B and Erickson JD (2004). Sodium-coupled neutral amino acid (System N/A) transporters of the SLC38 gene family. Pflugers Arch. 447 (5): 784-795.

14. Bröer A and Brookes N (2001). Transfer of glutamine between astrocytes and neurons. J Neurochem. 77 (3): 705-719.

15. Danbolt NC (2001). Glutamate uptake. Prog Neurobiol. 65 (1): 1-105.

16. Hertz L (2004). Intercellular metabolic compartmentation in the brain: past, present and future. Neurochem Int. 45 (2-3): 285-296.

17. Peng L, Hertz L, Huang R, Sonnewald U, Petersen SB, Westergaard N, Larsson O and Schousboe A (1993). Utilization of glutamine and of TCA cycle constituents as precursors for transmitter glutamate and GABA. Dev Neurosci. 15 (3-5): 367-377.

18. Masson J, Darmon M, Conjard A, Chuhma N, Ropert N, Thoby-Brisson M, Foutz AS, Parrot S, Miller GM, Jorisch R, Polan J, Hamon M, Hen R and Rayport S (2006). Mice lacking brain/kidney phosphate-activated glutaminase have impaired glutamatergic synaptic transmission, altered breathing, disorganized goal-directed behavior and die shortly after birth. J Neurosci. 26 (17): 4660-4671.

A surprising finding in this study was that the glutamine-glutamate pathway to recycle excitatory neurotransmitter does not seem to be as crucial as was expected for excitatory neurotransmission.

19. Fricke MN, Jones-Davis DM and Mathews GC (2007). Glutamine uptake by System A transporters maintains neurotransmitter GABA synthesis and inhibitory synaptic transmission. J Neurochem.

20. He Y, Hakvoort TB, Vermeulen JL, Lamers WH and Van Roon MA (2007). Glutamine synthetase is essential in early mouse embryogenesis. Dev Dyn. 236 (7): 1865-1875.

21. Asada H, Kawamura Y, Maruyama K, Kume H, Ding R, Ji FY, Kanbara N, Kuzume H, Sanbo M, Yagi T and Obata K (1996). Mice lacking the 65 kDa isoform of glutamic acid decarboxylase (GAD65) maintain normal levels of GAD67 and GABA in their brains but are susceptible to seizures. Biochem Biophys Res Commun. 229 (3): 891-895.

22. Kash SF, Johnson RS, Tecott LH, Noebels JL, Mayfield RD, Hanahan D and Baekkeskov S (1997). Epilepsy in mice deficient in the 65-kDa isoform of glutamic acid decarboxylase. Proc Natl Acad Sci U S A. 94 (25): 14060-14065.

23. Kash SF, Tecott LH, Hodge C and Baekkeskov S (1999). Increased anxiety and altered responses to anxiolytics in mice deficient in the 65-kDa isoform of glutamic acid decarboxylase. Proc Natl Acad Sci U S A. 96 (4): 1698-1703.

24. Heldt SA, Green A and Ressler KJ (2004). Prepulse inhibition deficits in GAD65 knockout mice and the effect of antipsychotic treatment. Neuropsychopharmacology. 29 (9): 1610-1619.

25. Wu H, Jin Y, Buddhala C, Osterhaus G, Cohen E, Jin H, Wei J, Davis K, Obata K and Wu JY (2007). Role of glutamate decarboxylase (GAD) isoform, GAD(65), in GABA synthesis and transport into synaptic vesicles-Evidence from GAD(65)-knockout mice studies. Brain Res. 1154: 80-83.

26. Asada H, Kawamura Y, Maruyama K, Kume H, Ding RG, Kanbara N, Kuzume H, Sanbo M, Yagi T and Obata K (1997). Cleft palate and decreased brain gamma-aminobutyric acid in mice lacking the 67-kDa isoform of glutamic acid decarboxylase. Proc Natl Acad Sci U S A. 94 (12): 6496-6499.

27. Kuwana S, Okada Y, Sugawara Y, Tsunekawa N and Obata K (2003). Disturbance of neural respiratory control in neonatal mice lacking GABA synthesizing enzyme 67-kDa isoform of glutamic acid decarboxylase. Neuroscience. 120 (3): 861-870.

28. Ji F, Kanbara N and Obata K (1999). GABA and histogenesis in fetal and neonatal mouse brain lacking both the isoforms of glutamic acid decarboxylase. Neurosci Res. 33 (3): 187-194.

29. Fujii M, Arata A, Kanbara-Kume N, Saito K, Yanagawa Y and Obata K (2007). Respiratory activity in brainstem of fetal mice lacking glutamate decarboxylase 65/67 and vesicular GABA transporter. Neuroscience. 146 (3): 1044-1052.

30. Liu GX, Cai GQ, Cai YQ, Sheng ZJ, Jiang J, Mei Z, Wang ZG, Guo L and Fei J (2007). Reduced Anxiety and Depression-Like Behaviors in Mice Lacking GABA Transporter Subtype 1. Neuropsychopharmacology. 32 (7): 1531-1539.

31. Liu GX, Liu S, Cai GQ, Sheng ZJ, Cai YQ, Jiang J, Sun X, Ma SK, Wang L, Wang ZG and Fei J (2007). Reduced aggression in mice lacking GABA transporter subtype 1. J Neurosci Res. 85 (3): 649-655.

32. Schwartz TL and Nihalani N (2006). Tiagabine in anxiety disorders. Expert Opin Pharmacother. 7 (14): 1977-1987.

33. Kalviainen R, Nousiainen I, Mantyjarvi M, Nikoskelainen E, Partanen J, Partanen K and Riekkinen P, Sr. (1999). Vigabatrin, a gabaergic antiepileptic drug, causes concentric visual field defects. Neurology. 53 (5): 922-926.

34. Gibson KM, Schor DS, Gupta M, Guerand WS, Senephansiri H, Burlingame TG, Bartels H, Hogema BM, Bottiglieri T, Froestl W, Snead OC, Grompe M and Jakobs C (2002). Focal neurometabolic alterations in mice deficient for succinate semialdehyde dehydrogenase. J Neurochem. 81 (1): 71-79.

35. Hogema BM, Gupta M, Senephansiri H, Burlingame TG, Taylor M, Jakobs C, Schutgens RB, Froestl W, Snead OC, Diaz-Arrastia R, Bottiglieri T, Grompe M and Gibson KM (2001). Pharmacologic rescue of lethal seizures in mice deficient in succinate semialdehyde dehydrogenase. Nat Genet. 29 (2): 212-216.

36. Cortez MA, Wu Y, Gibson KM and Snead OC, 3rd (2004). Absence seizures in succinic semialdehyde dehydrogenase deficient mice: a model of juvenile absence epilepsy. Pharmacol Biochem Behav. 79 (3): 547-553.

37. Crino PB, Jin H, Shumate MD, Robinson MB, Coulter DA and Brooks-Kayal AR (2002). Increased expression of the neuronal glutamate transporter (EAAT3/EAAC1) in hippocampal and neocortical epilepsy. Epilepsia. 43 (3): 211-218.

38. Doi T, Ueda Y, Tokumaru J and Willmore LJ (2005). Molecular regulation of glutamate and GABA transporter proteins by clobazam during epileptogenesis in Fe(+++)-induced epileptic rats. Brain Res Mol Brain Res. 142 (2): 91-96.

39. Ghijsen WE, da Silva Aresta Belo AI, Zuiderwijk M and Lopez da Silva FH (1999). Compensatory change in EAAC1 glutamate transporter in rat hippocampus CA1 region during kindling epileptogenesis. Neurosci Lett. 276 (3): 157-160.

40. Gorter JA, Van Vliet EA, Proper EA, De Graan PN, Ghijsen WE, Lopes Da Silva FH and Aronica E (2002). Glutamate transporters alterations in the reorganizing dentate gyrus are associated with progressive seizure activity in chronic epileptic rats. J Comp Neurol. 442 (4): 365-377.

41. Miller HP, Levey AI, Rothstein JD, Tzingounis AV and Conn PJ (1997). Alterations in glutamate transporter protein levels in kindling-induced epilepsy. J Neurochem. 68 (4): 1564-1570.

42. Ueda Y, Doi T, Tokumaru J, Yokoyama H, Nakajima A, Mitsuyama Y, Ohya-Nishiguchi H, Kamada H and Willmore LJ (2001). Collapse of extracellular glutamate regulation during epileptogenesis: down-regulation and functional failure of glutamate transporter function in rats with chronic seizures induced by kainic acid. J Neurochem. 76 (3): 892-900.

43. Peghini P, Janzen J and Stoffel W (1997). Glutamate transporter EAAC-1-deficient mice develop dicarboxylic aminoaciduria and behavioral abnormalities but no neurodegeneration. Embo J. 16 (13): 3822-3832.

44. Aoyama K, Suh SW, Hamby AM, Liu J, Chan WY, Chen Y and Swanson RA (2006). Neuronal glutathione deficiency and age-dependent neurodegeneration in the EAAC1 deficient mouse. Nat Neurosci. 9 (1): 119-126.

This study shows the role of EAAT3 in glutathione deficiency and in neurodegeneration over time

45. Furuta A, Noda M, Suzuki SO, Goto Y, Kanahori Y, Rothstein JD and Iwaki T (2003). Translocation of glutamate transporter subtype excitatory amino acid carrier 1 protein in kainic acid-induced rat epilepsy. Am J Pathol. 163 (2): 779-787.

46. Davis KE, Straff DJ, Weinstein EA, Bannerman PG, Correale DM, Rothstein JD and Robinson MB (1998). Multiple signaling pathways regulate cell surface expression and activity of the excitatory amino acid carrier 1 subtype of Glu transporter in C6 glioma. J Neurosci. 18 (7): 2475-2485.

This study shows that EAAT3 can be regulated by at least 2 independent signaling pathways, one involves PKC and the other involves PI3K.

47. Hartmann K, Bruehl C, Golovko T and Draguhn A (2008). Fast homeostatic plasticity of inhibition via activity-dependent vesicular filling. PLoS One. 3 (8): e2979.

48. During MJ and Spencer DD (1993). Extracellular hippocampal glutamate and spontaneous seizure in the conscious human brain. Lancet. 341 (8861): 1607-1610.

49. Levenson J, Weeber E, Selcher JC, Kategaya LS, Sweatt JD and Eskin A (2002). Long-term potentiation and contextual fear conditioning increase neuronal glutamate uptake. Nat Neurosci. 5 (2): 155-161.

50. Guillet BA, Velly LJ, Canolle B, Masmejean FM, Nieoullon AL and Pisano P (2005). Differential regulation by protein kinases of activity and cell surface expression of glutamate transporters in neuron-enriched cultures. Neurochem Int. 46 (4): 337-346.

51. Gonzalez MI, Kazanietz MG and Robinson MB (2002). Regulation of the neuronal glutamate transporter excitatory amino acid carrier-1 (EAAC1) by different protein kinase C subtypes. Mol Pharmacol. 62 (4): 901-910.

Interesting article which shows a direct interaction between EAAT3 and PKCα not just in C6 glioma, but also in neurons.

52. Gonzalez MI, Bannerman PG and Robinson MB (2003). Phorbol myristate acetate-dependent interaction of protein kinase Calpha and the neuronal glutamate transporter EAAC1. J Neurosci. 23 (13): 5589-5593.

53. Sims KD, Straff DJ and Robinson MB (2000). Platelet-derived growth factor rapidly increases activity and cell surface expression of the EAAC1 subtype of glutamate transporter through activation of phosphatidylinositol 3-kinase. J Biol Chem. 275 (7): 5228-5237.

54. Connor M and Christie MD (1999). Opioid receptor signalling mechanisms. Clin Exp Pharmacol Physiol. 26 (7): 493-499.

55. Conn PJ (2003). Physiological roles and therapeutic potential of metabotropic glutamate receptors. Ann N Y Acad Sci. 1003: 12-21.

56. Mitchell SJ and Silver RA (2000). Glutamate spillover suppresses inhibition by activating presynaptic mGluRs. Nature. 404 (6777): 498-502.

57. Bashir ZI, Bortolotto ZA, Davies CH, Berretta N, Irving AJ, Seal AJ, Henley JM, Jane DE, Watkins JC and Collingridge GL (1993). Induction of LTP in the hippocampus needs synaptic activation of glutamate metabotropic receptors. Nature. 363 (6427): 347-350.

58. Wong RK, Bianchi R, Taylor GW and Merlin LR (1999). Role of metabotropic glutamate receptors in epilepsy. Adv Neurol. 79: 685-698.

59. Tso PH and Wong YH (2003). Molecular basis of opioid dependence: role of signal regulation by G-proteins. Clin Exp Pharmacol Physiol. 30 (5-6): 307-316.

60. Marinissen MJ and Gutkind JS (2001). G-protein-coupled receptors and signaling networks: emerging paradigms. Trends Pharmacol Sci. 22 (7): 368-376.

61. Drake CT and Milner TA (2002). Mu opioid receptors are in discrete hippocampal interneuron subpopulations. Hippocampus. 12 (2): 119-136.

62. Drake CT and Milner TA (1999). Mu opioid receptors are in somatodendritic and axonal compartments of GABAergic neurons in rat hippocampal formation. Brain Res. 849 (1-2): 203-215.

63. Bausch SB, Patterson TA, Appleyard SM and Chavkin C (1995). Immunocytochemical localization of delta opioid receptors in mouse brain. J Chem Neuroanat. 8 (3): 175-189.

64. Ding YQ, Kaneko T, Nomura S and Mizuno N (1996). Immunohistochemical localization of mu-opioid receptors in the central nervous system of the rat. J Comp Neurol. 367 (3): 375-402.

65. Mansour A, Fox CA, Burke S, Akil H and Watson SJ (1995). Immunohistochemical localization of the cloned mu opioid receptor in the rat CNS. J Chem Neuroanat. 8 (4): 283-305.

66. Commons KG and Milner TA (1997). Localization of delta opioid receptor immunoreactivity in interneurons and pyramidal cells in the rat hippocampus. J Comp Neurol. 381 (3): 373-387.

67. Eriksson PS, Hansson E and Ronnback L (1990). Opiate receptors in neuronal primary cultures. Neuropharmacology. 29 (9): 799-804.

68. Xia P, Pei G and Schwarz W (2006). Regulation of the glutamate transporter EAAC1 by expression and activation of delta-opioid receptor. Eur J Neurosci. 24 (1): 87-93.

Recent study that shows in C6 glioma cells there is a functional interaction between DOR and EAAT3.  It is suggested that this interaction might also occur in neurons.

69. Krizman-Genda E, Gonzalez MI, Zelenaia O and Robinson MB (2005). Evidence that Akt mediates platelet-derived growth factor-dependent increases in activity and surface expression of the neuronal glutamate transporter, EAAC1. Neuropharmacology. 49 (6): 872-882.

70. Shahabi NA, McAllen K and Sharp BM (2006). delta opioid receptors stimulate Akt-dependent phosphorylation of c-jun in T cells. J Pharmacol Exp Ther. 316 (2): 933-939.

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