Glutamate transporter type 3 attenuates the activation of N-methy-D-aspartate receptors co-expressed in Xenopus oocytes
Department of Anesthesiology, University of Virginia Health System, Charlottesville, Virginia 22908-0710, USA
* Author for correspondence (e-mail: zz3c{at}virginia.edu)
Accepted 14 March 2005
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Summary |
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Key words: glutamate receptor, glutamate transporter, neurotransmission, oocytes
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Introduction |
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The regulation of extracellular glutamate homeostasis by EAATs has been
investigated by two approaches: molecular biology manipulations, in which the
expression of a selective EAAT is disrupted, and pharmacological blockade, in
which transporter function is inhibited with appropriate inhibitors. By using
molecular biology technique, it has been shown that mice that lack EAAT1 or
EAAT2 expression had increased extracellular concentrations of glutamate and
that the animals were susceptible to seizures and excitotoxic cell death
(Rothstein et al., 1996).
Although such an approach provides important functional information on EAATs
to regulate extracellular concentrations of glutamate in the central nervous
system, the chronic inhibition of the gene expression may induce compensatory
mechanisms and will not provide information on the dynamics of extracellular
glutamate homeostasis in response to acute disruption of the uptake
system.
Pharmacological blockade of EAATs has been used to study the role of EAATs
in the dynamics of glutamate homeostasis. The inhibition of EAAT activity has
been shown to prolong the glutamate-induced current, leading to a slowed
excitatory post-synaptic current (EPSC) decay at some synapses
(Barbour et al., 1994;
Diamond and Jahr, 1997
;
Kinney et al., 1997
;
Mennerick and Zorumski, 1994
;
Otis et al., 1996
;
Takahashi et al., 1995
;
Tong and Jahr, 1994
). A recent
study further suggests that glutamate translocation by EAATs is important in
control of EPSC decay (Mennerick et al.,
1999
).
We used a different approach, co-expression of N-methy-D-aspartate receptor (NMDAR) and EAATs, to investigate the role of EAATs in the control of NMDAR activation. We artificially expressed NMDAR with or without a neuronal EAAT, EAAT3, in Xenopus oocytes. This approach allowed us to compare the activation of NMDAR in the presence or absence of EAAT3 without the need to use multiple inhibitors for EAATs or glutamate receptors. By using this model, we tested the hypothesis that EAATs regulate NMDAR activation/current induced by glutamate.
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Materials and methods |
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Materials
All reagents, unless specified below, were obtained from Sigma Chemical (St
Louis, MO, USA).
cRNA preparation
Rat EAAT3 cDNA in a commercial plasmid vector (BluescriptSKM) was provided
by Mattias A. Hediger. Rat NMDAR1-1a in pBS SK(-) and rat NMDAR2A in pBS SK(+)
were from Steve F. Heinemann. The cDNAs were linearized with restriction
enzymes that were suggested by the people who made the constructs. The capped
cRNAs were transcribed using a commercial T7 polymerase (Ambion, Austin, TX,
USA).
Oocyte preparation and injection
These procedures were performed as we described before (Do et al.,
2001,
2002b
;
Fang et al., 2002
). Briefly,
one day before cRNA injection, stage V-VI oocytes were isolated from adult
female Xenopus laevis (Daudin) frogs (Ann Arbor, MI, USA anesthetized
with 0.2% 3-aminobenzoic acid ethyl ester. After being surgically removed from
the frog, oocytes were defolliculated with 20 mg collagenase (Type 1a) in 20
ml of Ca2+ free OR2 solution (containing in mmol l-1:
NaCl 82.5, KCl 2.0, MgCl2 1.0, and HEPES 5.0, pH adjusted to 7.4)
for 2 h at room temperature (22°C). Oocytes were injected (Drummond
`Nanoject', Drummond Scientific Co., Broomall, PA, USA) with 40 ng cRNA of
EAAT3 or 10 or 40 ng cRNA of NMDAR1-1a and NMDAR2A (1:4 weight ratio). Oocytes
were then incubated at 16°C in modified Barth's solution (containing in
mmol l-1: NaCl 88, KCl 1, NaHCO3 2.4, CaCl2
0.41, MgSO4 0.82, Ca(NO3)2 0.3, gentamicin
0.1, and HEPES 15, pH adjusted to 7.4) before voltage clamping was
performed.
Electrophysiological recordings
As we described before (Do et al.,
2002b; Fang et al.,
2002
), 4-5 days after injection of the cRNA, oocytes were
superfused by gravity flow with Mg2+- and Ca2+-free
Ringer's solution (containing in mmol l-1: NaCl 96, KCl 2,
BaCl2 1.8, and HEPES 10, pH adjusted to 7.5) containing glycine 10
µmol l-1. The flow is about 3 ml min-1 and the oocyte
chamber volume is about 1 ml. Clamping microelectrodes were pulled from
capillary glass (10 µl Drummond Microdispenser, Drummond Scientific
Company, Broomall, PA, USA) on a micropipette puller (model 700C; David Kopf
Instruments, Tujunga, CA, USA). Electrodes were broken at the tip whose
diameter was approximately 10 µm and filled with 3 mol l-1 KCl
obtaining resistance of 3 M
. Oocytes were voltage-clamped using a
two-electrode voltage clamp amplifier (OC725A; Warner Corporation, New Heaven,
CT, USA), which was connected to a DAS-8A/D conversion board
(Keithley-Metrabyte, Taunton, MA, USA) on an IBM-compatible PC. Data
acquisition and analysis were performed using the OoClamp program
(Durieux, 1993
). Current was
examined for 70 s (25 s of application of glutamate or NMDA, 45 s of recovery
with a glutamate- and NMDA-free superfusate) at a holding potential of -70 mV.
Flow-stopped experiments, as modified from a previous study
(Supplisson and Bergman,
1997
), were performed by applying glutamate or NMDA for 25 s and
then flow stopping for 15 s before the flow was resumed for 5 s with a
glutamate- or NMDA-containing solution followed by a 35 s of recovery with a
glutamate- and NMDA-free superfusate. At least 2 min interval time was allowed
after each measurement. Response was quantified by measuring the peak current
using OoClamp program. All experiments were performed at room temperature.
Administration of experimental chemicals
Since EAATs are Na+-co-transporters, in some experiments
Na+ in the bath solution was replaced by Li+ to inhibit
the EAAT3 function. To prevent NMDAR activation in the co-expression oocytes,
glycine-free Ringer's solution containing 5 mmol l-1
MgCl2 was used.
Statistical analysis
Due to the variation in the expression level of EAAT3 and NMDAR proteins in
oocytes, glutamate- or NMDA-induced response was normalized to the maximal
response of the oocytes to the agents (300 µmol l-1 glutamate or
1 mmol l-1 NMDA). Results are mean
±S.D. from 6-16 oocytes from at least three
frogs. Statistical analysis was performed by unpaired t-test. A
P<0.05 was accepted as significant. EC50 was derived by
analyzing data with Graphpad Prism 3.0 (Graphpad Software, Inc, San Diego, CA,
USA).
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Results |
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Reduction of NMDAR currents in EAAT3+ oocytes under stopped-flow condition
In oocytes expressing NMDAR only (EAAT3-), the glutamate-induced
current remained stable when the flow of the L-glutamate-containing
superfusate was stopped. By contrast, co-expressing oocytes developed a marked
decrease of the inward current when the superfusion was stopped
(Fig. 2). Reestablishing the
flow restored the full amplitude of the current (Figs
2 and
3). This phenomenon is called
stopped-flow reduction of NMDAR current in this paper.
|
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The degree of stopped-flow reduction of NMDAR current depended on the concentrations of glutamate in the superfusates (Fig. 4). When glutamate concentrations were more than 100 µmol l-1, stopping the flow produced a small reduction or no reduction at all (Fig. 4). This phenomenon is expected for a saturable uptake process operating in the presence of substrates at supra-saturable concentrations.
|
![]() | (1) |
where [Glu]s is the sensed glutamate concentrations, I is the glutamate-evoked current, Imax is the current response to a saturating glutamate concentration (300 µmol l-1), and the EC50 and the Hill coefficient (n) refer to the mean values measured in oocytes expressing NMDAR only. As shown in Fig. 5, the [Glu]s was about one half to one third of the glutamate concentrations (ranging from 0.3 to 30 µmol l-1) in the superfusates. The [Glu]s values were 0.10±0.07, 0.35±0.18, 1.41±0.71, 4.53±2.52, 13.85±6.13 µmol l-1 (means ±S.D., N=12) for glutamate concentrations in the superfusates = 0.3, 1, 3, 10 and 30 µmol l-1, respectively.
|
Reduction of NMDAR current in EAAT3+ oocytes under continuous flow condition
Under continuous flow condition, L-glutamate at concentrations
lower than 30 µmol l-1 (no saturating concentrations for NMDAR)
induced smaller currents in co-expressing oocytes than those in oocytes
expressing NMDAR only (Fig. 6).
This reduction can be quantitatively expressed as lower local glutamate
concentrations being sensed by NMDAR in co-expressing oocytes than those in
oocytes expressing NMDAR only (Fig.
5).
|
This reduction can also be quantitatively expressed as EC50 being right-shifted (Fig. 7). As a control study, no EC50 shift of the EAAT3 responses was noticed between the oocytes expressing EAAT3 only (31.8±6.5 µmol l-1, N=12) and co-expressing oocytes (30.9±6.0 µmol l-1, N=9, P>0.05) (Fig. 7A). In addition, no EC50 shift was observed between the oocytes expressing NMDAR only (32.0±9.8 µmol l-1, N=6) and co-expressing oocytes (30.4±5.2 µmol l-1, N=6, P>0.05) when NMDA was used as the agonist for NMDAR (Fig. 7B). However, significant EC50 shift was found between the oocytes expressing NMDAR only (oocytes injected with 40 ng mRNA of NMDAR, 3.6±0.9 µmol l-1, N=16) and co-expressing oocytes (oocytes injected with 40 ng mRNA of NMDAR and 40 ng mRNA of EAAT3, 10.5±4.3 µmol l-1, N=15, P<0.05) when glutamate was used as the agonist for NMDAR (Fig. 7C). The EC50 was shifted even more in oocytes that had higher quantitative ratio of EAAT3 proteins/NMDAR proteins (oocytes injected with 10 ng mRNA of NMDAR and 40 ng mRNA of EAAT3, 17.1±6.2 µmol l-1, N=10, P<0.05 compared with oocytes injected with 40 ng mRNA of NMDAR and 40 ng mRNA of EAAT3) (Fig. 7C). A linear correlation between the EC50 of glutamate-induced current responses and the ratio of EAAT3 current/total glutamate-induced current in the co-expression oocytes was apparent (Fig. 7D). However, the EC50 shift was abolished when Na+ in the superfusates was replaced by Li+ in the co-expressing oocytes injected with 40 ng mRNA of NMDAR and 40 ng mRNA of EAAT3 (3.8±1.2 µmol l-1, N=6, P>0.05 compared with oocytes expressing NMDAR only) (Fig. 7C).
|
These results suggest that EAAT3 can decrease glutamate concentrations sensed by NMDAR in the co-expressing oocytes even under the continuous flow conditions, generating a steady-state glutamate concentration gradient between the bath solution and the cell membrane. The apparent affinities for glutamate in EAAT3+ oocytes appeared different from those in EAAT3- oocytes even at early response time (Fig. 6), suggesting that the glutamate gradient was established as fast as the change of glutamate concentrations in solution.
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Discussion |
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Our results showed that EAAT3+ oocytes had a fast recovery of
NMDAR current in the stopped-flow experiments and a smaller NMDAR current in
response to a non-saturating concentration of glutamate in the continuous flow
experiments than EAAT3- oocytes. These results suggest that EAAT3
regulates NMDAR currents. Previous studies have demonstrated that the
application of EAAT inhibitors in brain slices or cell cultures increased the
peak glutamate concentration in the synaptic cleft and prolonged EPSC decay at
some synapses (Barbour et al.,
1994; Diamond and Jahr,
1997
; Kinney et al.,
1997
; Mennerick and Zorumski,
1994
; Otis et al.,
1996
; Takahashi et al.,
1995
; Tong and Jahr,
1994
). The action of EAATs at controlling the amplitude of EPSC
has also been reported (Diamond,
2001
; Diamond and Jahr,
1997
; Tong and Jahr,
1994
). Thus, EAATs, by regulating glutamate concentration in
synapses, modulate glutamate neurotransmission such as that through NMDAR.
However, the modulation of glutamate neurotransmission by EAATs is not
effective in all synapses. For example, studies of small simple synapses like
hippocampal Schaffer collateral to pyramidal cell synapses have showed that
inhibition of EAATs did not change the decay of EPSC
(Hestrin et al., 1990;
Isaacson and Nicoll, 1993
;
Sarantis et al., 1993
). This
failure to modulate may be due to very few EAATs that are expressed nearby to
the glutamate receptors or a small quantitative ratio of EAATs/glutamate
receptors. Under these conditions, small amount of glutamate from the total
glutamate pool will bind to EAATs and the inhibition of EAATs may not
significantly change the amount of glutamate available to glutamate receptors.
To experimentally model this situation, we compared the effects of EAATs on
NMDAR current in oocytes with different expression ratios of EAAT3/NMDAR. Our
results showed that oocytes with smaller ratio of EAAT3/NMDAR had smaller
changes in amplitude of NMDAR currents compared to oocytes with NMDAR alone.
Consistent with our results, the synapses in which the inhibition of EAATs did
not affect the EPSC usually have limited glial covering or large distance
between the synaptic cleft and glial membrane
(Lehre and Danbolt, 1998
).
How do EAATs alter the availability of glutamate to glutamate receptors in
synapses and, thus, regulate glutamate neurotransmission? Each transport cycle
of glutamate by EAATs consists of at least three stages: glutamate binding,
glutamate translocation and an anion-conducting state
(Billups et al., 1998;
Grewer et al., 2000
). It was
calculated that the time constant for a complete cycle of transport at -80 mV
and 22°C is approximately 70 ms, which is significantly slower than the
estimated glutamate-decay time constant in hippocampal synapses (
1-2 ms)
(Clements et al., 1992
). This
difference was predicted to be true also at the physiological temperature
(Wadiche et al., 1995
). Thus,
whether EAATs really constitute a major mechanism for removing released
glutamate was questioned (Wadiche et al.,
1995
). However, binding and translocation of glutamate may have
rapid kinetics (Mennerick et al.,
1999
; Tong and Jahr,
1994
). It was estimated by applying the laser-pulse photolysis
technique of caged glutamate with a time resolution of 100 µs that
glutamate translocation occurs within a few milliseconds after being bound to
EAAT3 (Grewer et al., 2000
).
Johr and colleagues have proposed that the action of EAATs at controlling EPSC
is a consequence of rapid buffering of glutamate by a high density of binding
sites provided by EAATs near to glutamate receptors
(Diamond and Jahr, 1997
;
Tong and Jahr, 1994
).
Implications
As discussed in the above section, studies in the literature have suggested
the regulation of the amplitude of EPSC by EAATs. However, those studies were
performed with use of various combinations of inhibitors to isolate glutamate
receptor-mediated EPSC from that caused by other neurotransmitters. In
addition, due to the lack of non-transportable inhibitors, most of those early
studies were performed with transportable EAAT inhibitors. These EAAT
inhibitors can induce glutamate release from intracellular space via
heteroexchange (Volterra et al.,
1996). Thus, the use of these inhibitors may have resulted in
overestimation of the EAAT roles in maintaining extracellular glutamate
homeostasis. To overcome this problem, Jabaudon et al.
(1999
) applied
DL-threo-ß-benzyloxyaspartate (TBOA), a
non-transportable EAAT inhibitor developed recently, to rat hippocampal slice
culture and used the NMDAR of patched CA3 hippocampal neurons as `glutamate
sensors'. They found that under basal conditions, the activity of EAATs
compensates for the continuous, non-vesicular release of glutamate from the
intracellular compartment. The inhibition of EAAT activity by TBOA immediately
results in significant accumulation of extracellular glutamate
(Jabaudon et al., 1999
). The
inhibition of postsynaptic EAATs in CA1 pyramidal cells by TBOA has been shown
to enhance the activation of NMDAR by neurotransmitter spillover from
neighboring synapses onto the synapses of these pyramidal cells
(Diamond, 2001
). We
co-expressed NMDAR and EAAT3 in oocytes and did not need to use inhibitors to
isolate NMDAR responses for study.
Our results showed that the effects of EAAT3 on the NMDAR activation
decreased as the extracellular glutamate concentration increased. The effects
were minimal at 100 µmol l-1 or higher concentrations of
glutamate. Although glutamate concentration in the synaptic cleft during
excitation (which may vary in different synapses) is not known, it is believed
that the concentration is in micro molar level
(Danbolt, 2001). Thus, the
in vivo glutamate concentrations in the synaptic cleft may fall into
the concentration range that can be regulated by EAATs to effectively modulate
glutamate receptor activation as demonstrated in our study. However, the
degree of the effects of EAATs on glutamate receptor activation in
vivo is obviously dependent on the density of EAATs and the distance
between EAATs and glutamate receptors. The density of heterologous expression
of transporters in oocytes usually is at about 150-3000 transporters
µm2 (Mager et al.,
1993
; Supplisson and Bergman,
1997
; Zampighi et al.,
1995
). This density is lower than that estimated in the nervous
tissue because 15,000 and 21,000 glial EAAT molecules were calculated to be
present per µm2 in the stratum radiatum of hippocampus CA1 and
molecular layer of cerebellum, respectively
(Lehre and Danbolt, 1998
), and
EAAT4 is at an average density of
2000 molecules µm2 of the
molecular layer (Dehnes et al.,
1998
). Thus, the effects of EAATs on glutamate receptor activation
under in vivo conditions may be bigger than that in our study.
However, the distance between EAATs and glutamate receptors in vivo
may be larger than that in oocytes (we assume that NMDAR and EAAT3 proteins
are distributed evenly at the surface of the oocyte including the microvilli).
Only can EAATs located inside the synaptic cleft regulate glutamate
concentrations there. However, all five EAATs cloned so far appear to be
present outside synaptic cleft except for EAAT4 on the postsynaptic densities
of Purkinje cell spines (Danbolt,
2001
). The perisynaptical distribution of neuronal EAATs such as
EAAT3 may limit glutamate diffusing into synapses from outside
(Diamond, 2001
).
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Acknowledgments |
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References |
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---|
Arriza, J. L., Eliasof, S., Kavanaugh, M. P. and Amara, S.
G. (1997). Excitatory amino acid transporter 5, a retinal
glutamate transporter coupled to a chloride conductance. Proc.
Natl. Acad. Sci. USA 94,4155
-4160.
Barbour, B., Keller, B. U., Llano, I. and Marty, A. (1994). Prolonged presence of glutamate during excitatory synaptic transmission to cerebellar Purkinje cells. Neuron 12,1331 -1343.[CrossRef][Medline]
Billups, B., Rossi, D., Oshima, T., Warr, O., Takahashi, M., Sarantis, M., Szatkowski, M. and Attwell, D. (1998). Physiological and pathological operation of glutamate transporters. Prog. Brain Res. 116,45 -57.[Medline]
Clements, J. D., Lester, R. A., Tong, G., Jahr, C. E. and Westbrook, G. L. (1992). The time course of glutamate in the synaptic cleft. Science 258,1498 -1501.[Medline]
Costa, A. C. S., Patrick, J. and Dani, J. (1994). Improved technique for studying ion channels expressed in Xenopus oocytes, including fast superfusion. Biophysical J. 67,395 -401.[Abstract]
Danbolt, N. C. (2001). Glutamate uptake. Progr. Neurobiol. 65,1 -105.[CrossRef][Medline]
Dehnes, Y., Chaudhry, F. A., Ullensvang, K., Lehre, K. P.,
Storm-Mathisen, J. and Danbolt, N. C. (1998). The glutamate
transporter EAAT4 in rat cerebellar Purkinje cells: a glutamate-gated chloride
channel concentrated near the synapse in parts of the dendritic membrane
facing astroglia. J. Neurosci.
18,3606
-3619.
Diamond, J. S. (2001). Neuronal glutamate
transporters limit activation of NMDA receptors by neurotransmitter spillover
on CA1 pyramidal cells. J. Neurosci.
21,8328
-8338.
Diamond, J. S. and Jahr, C. E. (1997).
Transporters buffer synaptically released glutamate on a submillisecond time
scale. J. Neurosci. 17,4672
-4687.
Do, S.-H., Fang, H. Y., Ham, B. M. and Zuo, Z.
(2002a). The effects of lidocaine on the activity of glutamate
transporter EAAT3: the role of protein kinase C and phosphatidylinositol
3-kinase. Anesthesia Analgesia
95,1263
-1268.
Do, S.-H., Kamatchi, G. L. and Durieux, M. E.
(2001). The effects of isoflurane on native and chimeric
muscarinic acetylcholine receptors: the role of protein kinase C.
Anesthesia Analgesia 93,375
-381.
Do, S.-H., Kamatchi, G. L., Washington, J. M. and Zuo, Z. (2002b). Effects of volatile anesthetics on glutamate transporter, excitatory amino acid transporter type 3. Anesthesiology 96,1492 -1497.[CrossRef][Medline]
Durieux, M. E. (1993). An IBM-compatible software system for electrophysiologic receptor studies in Xenopus oocytes. Comp. Meth. Programs Biomed. 41,101 -105.[CrossRef]
Fang, H., Huang, Y. and Zuo, Z. (2002). The different responses of rat glutamate transporter type 2 and its mutant (tyrosine 403 to histidine) activity to volatile anesthetics and activation of protein kinase C. Brain Res. 953,255 -264.[CrossRef][Medline]
Grewer, C., Watzke, N., Wiessner, M. and Rauen, T.
(2000). Glutamate translocation of the neuronal glutamate
transporter EAAC1 occurs within milliseconds. Proc. Natl. Acad.
Sci. USA 97,9706
-9711.
Hestrin, S., Sah, P. and Nicoll, R. A. (1990). Mechanisms generating the time course of dual componet excitatory synaptic currents recorded in hippocampal slices. Neuron 5, 247-253.[CrossRef][Medline]
Isaacson, J. S. and Nicoll, R. A. (1993). The
uptake inhibitor L-trans-PDC enhances responses to glutamate but
fails to alter the kinetics of excitatory synaptic currents in the
hippocampus. J. Neurophysiol.
70,2187
-2191.
Jabaudon, D., Shimamoto, K., Yasuda-Kamatani, Y., Scanziani, M.,
Gahwiler, B. H. and Gerber, U. (1999). Inhibition of uptake
unmasks rapid extracellular turnover of glutamate of nonvesicular origin.
Proc. Natl. Acad. Sci. USA
96,8733
-8738.
Kinney, G. A., Overstreet, L. S. and Slater, N. T.
(1997). Prolonged physiological entrapment of glutamate in the
synaptic cleft of cerebellare unipolar brush cells. J.
Neurophysiol. 78,1320
-1333.
Lehre, K. P. and Danbolt, N. C. (1998). The
number of glutamate transporter subtype molecules at glutamatergic synapses:
chemical and stereological quantification in young adult rat brain.
J. Neurosci. 18,8751
-8757.
Mager, S., Naeve, J., Quick, M., Labarca, C., Davidson, H. and Davidson, L. (1993). Steady states, charge movements, and rates for a cloned GABA transporter expressed in Xenopus oocytes. Neuron 10,177 -188.[CrossRef][Medline]
Mennerick, S., Shen, W., Xu, W., Benz, A., Tanaka, K.,
Shimamoto, K., Isenberg, K. E., Krause, J. E. and Zorumski, C. F.
(1999). Substrate turnover by transporters curtails synaptic
glutamate transients. J. Neurosci.
19,9242
-9251.
Mennerick, S. and Zorumski, C. F. (1994). Glial contributions to excitatory neurotransmission in cultured hippocampal cells. Nature 368,59 -62.[CrossRef][Medline]
Otis, T. S., Wu, Y. C. and Trussell, L. O. (1996). Delayed clearance of transmitter and the role of glutamate transporters at synapses with multiple release sites. J. Neurosci. 16,1634 -1644.[Abstract]
Roskoski, R., Jr (1979). Net uptake of aspartate by high-affinity rat cortical synaptosomal transport system. Brain Res. 160,85 -93.[CrossRef][Medline]
Rothstein, J. D., Dykes-Hoberg, M., Pardo, C. A., Bristol, L. A., Jin, L., Kuncl, R. W., Kanai, Y., Hediger, M. A., Wang, Y., Schielke, J. P. et al. (1996). Knockout of glutamate transporters reveals a major role for astroglial transport in excitotoxicity and clearance of glutamate. Neuron 16,675 -686.[CrossRef][Medline]
Rothstein, J. D., Martin, L., Levey, A. I., Dykes-Hoberg, M., Jin, L., Wu, D., Nash, N. and Kuncl, R. W. (1994). Localization of neuronal and glial glutamate transporters. Neuron 13,713 -725.[CrossRef][Medline]
Sarantis, M., Ballerini, L., Miller, B., Silver, R. A., Edwards, M. and Atwell, D. (1993). Glutamate uptake from the synaptic cleft does not shape the decay of the non-NMDA component of the synaptic current. Neuron 11,541 -549.[CrossRef][Medline]
Supplisson, S. and Bergman, C. (1997). Control
of NMDA receptor activation by a glycine transporter co-expressed in
Xenopus oocytes. J. Neurosci.
17,4580
-4590.
Takahashi, M., Kovalchuk, Y. and Attwell, D. (1995). Presynaptic and postsynaptic determinates of EPSC waveform at cerebellular climbing fiber and parallel fiber to Purkinje cell synapses. J. Neurosci. 15,5693 -5702.[Abstract]
Tong, G. and Jahr, C. E. (1994). Block of glutamate transporters potentiates postsynaptic excitation. Neuron 13,1195 -1203.[CrossRef][Medline]
Volterra, A., Bezzi, P., Rizzini, B. L., Trotti, D., Ullensvang, K., Danbolt, N. C. and Racagni, G. (1996). The competitive transport inhibitor L-trans-pyrrolidine-2, 4-dicarboxylate triggers excitotoxicity in rat cortical neuron-astrocyte co-cultures via glutamate release rather than uptake inhibition. Eur. J. Neurosci. 8,2019 -2028.[Medline]
Wadiche, J. I., Arriza, J. L., Amara, S. G. and Kavanaugh, M. P. (1995). Kinetics of a human glutamate transporter. Neuron 14,1019 -1027.[CrossRef][Medline]
Zampighi, G. A., Kreman, M., Boorer, K. J., Loo, D. D., Bezanilla, F., Chandy, G., Hall, J. E. and Wright, E. M. (1995). A method for determining the unitary functional capacity of cloned channels and transporters expressed in Xenopus laevis oocytes. J. Membr. Biol. 148, 65-78.[Medline]
Zuo, Z. (2001). Isoflurane enhances glutamate uptake via glutamate transporters in rat glial cells. Neuroreport 12,1077 -1080.[CrossRef][Medline]