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The Journal of Neuroscience, June 15, 1998, 18(12):4490-4499
Neuronal Expression of the Glutamate Transporter GLT-1 in
Hippocampal Microcultures
Steven
Mennerick1,
Rupali P.
Dhond1,
Ann
Benz1,
Wanyan
Xu1,
Jeffrey D.
Rothstein2,
Niels C.
Danbolt3,
Keith E.
Isenberg1, and
Charles F.
Zorumski1
1 Department of Psychiatry, Washington University
School of Medicine, St. Louis, Missouri 63110, 2 Department
of Neurology, Johns Hopkins School of Medicine, Baltimore, Maryland
21287, and 3 Anatomical Institute, University of Oslo,
Oslo, Norway
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ABSTRACT |
To address the question of the relative contributions of glial and
neuronal glutamate transport in the vertebrate CNS, we studied the
distribution of forebrain glutamate transporters in rat hippocampal
microcultures, a preparation in which physiological functions of
glutamate transporters have been well characterized. Two of the three
transporters, GLAST (EAAT1) and EAAC1 (EAAT3), are localized to
microculture glia and neurons, respectively, as expected. However, we
find strong immunoreactivity for the third glutamate transporter GLT-1
(EAAT2), a putatively glial transporter, in microculture neurons and in
a small subset of microculture glia. Indistinguishable
immunohistochemical staining patterns for GLT-1 were obtained with
antibodies directed against both the N terminal and C terminal of the
GLT-1 protein. Double-labeling experiments suggest that neuronal GLT-1
protein is primarily localized to the dendrites of excitatory neurons.
Neuronal electrogenic transport currents in response to
D-aspartate applications were occluded by the selective
GLT-1 inhibitor dihydrokainate. In contrast, glia exhibited a larger
transporter current density than did neurons, and the glial transport
current was less sensitive to dihydrokainate. Neuronal transport
currents were potentiated less than were glial currents when the
chaotropic anion thiocyanate was substituted for gluconate in the
whole-cell recording pipette, consistent with the previously reported
lower anion permeability of EAAC1 and GLT-1 compared with that of
GLAST. After microculture glia were rendered nonviable, excitatory
autaptic currents (EACs) were prolonged in the presence of
dihydrokainate, suggesting that neuronal GLT-1 is capable of
participating in the clearance of synaptically released glutamate. Our
results suggest that the initially proposed characterization of GLT-1
as a purely glial transporter is too simplistic and that under certain
conditions functional GLT-1 protein can be expressed in brain neurons.
The study suggests that changes in GLT-1 levels that occur with
pathology or experimental manipulations cannot be assumed to be
glial.
Key words:
glutamate transport; glutamate uptake; astrocyte; postsynaptic; dendrite; microculture; hippocampus
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INTRODUCTION |
Glutamate transporters participate
in limiting the effective lifetime of synaptically released glutamate
(Barbour et al., 1994 ; Mennerick and Zorumski, 1994; Tong and Jahr,
1994) and maintain ambient glutamate concentrations below toxic levels
(Rothstein, 1996a; Tanaka et al., 1997). The relative contributions of
glial versus neuronal glutamate transport remain unclear, although
recent electrophysiological studies suggest that electrogenic transport currents can be recorded during synaptic events in both glia and neurons in situ (Bergles and Jahr, 1997; Clark and Barbour,
1997; Otis et al., 1997). Understanding the differential contributions of various transporters is likely to lead to a better understanding of
the role of transporters in both normal and pathological states like
amyotrophic lateral sclerosis, in which downregulation of the
putatively glial glutamate transporter GLT-1 has been correlated with
neurodegeneration (Rothstein, 1996b).
The recent cloning of glutamate transporters has permitted localization
studies suggesting that, in the adult forebrain, two transporters,
GLAST (EAAT1) and GLT-1 (EAAT2), localize to glial cells, whereas EAAC1
localizes to neurons (e.g., Rothstein et al., 1994; Lehre et al.,
1995). These original distinctions have been slightly blurred recently
by the observation of GLT-1 mRNA (but not protein) expression in
hippocampal pyramidal cells (Schmitt et al., 1996; Torp et al., 1997),
observations of transient GLT-1 protein expression in white matter of
embryonic brain (Furuta et al., 1997), GLT-1 protein expression in
certain neurons of the retina (Grunert et al., 1994; Rauen and Kanner,
1994), and apparent neuronal expression of GLT-1 in a porcine ischemia
model (Martin et al., 1997).
Studying the localization of transporter subtypes in a preparation for
which a physiological role of glutamate transporters has been
characterized is one approach to understanding possible differential
contributions of the various glutamate transporters. We have been
studying the role of glutamate transporters in microcultures of
postnatal rat hippocampus. These cultures provide a highly controlled
environment that allows experimental manipulation of the cellular
growth environment. Additionally, because neurons and glia in culture
can be easily distinguished at the light microscopic level, it is
possible to perform independent experimental manipulations on one cell
type versus the other. We recently examined the effects on synaptic
currents of both broad-spectrum and glial-specific glutamate transport
inhibition (Mennerick and Zorumski, 1994). These results suggested a
predominant role for glia in clearance and/or buffering of glutamate
after presynaptic release. As part of an effort to determine the
molecular identity of the transporter(s) responsible for these effects,
we have undertaken an immunohistochemical localization of the three
forebrain transporters in hippocampal microcultures.
Our results suggest that although two of the transporters, GLAST and
EAAC1, show the expected distribution in glia and neurons, respectively, GLT-1 expression shows a surprising pattern of low glial
levels and robust expression in postsynaptic neuronal processes. The
results suggest that this putatively glial transporter can be
functionally expressed in neurons and suggest that changes in GLT-1
expression in pathological or experimental situations need to be
interpreted with caution.
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MATERIALS AND METHODS |
Cell culture. Hippocampal cells were prepared from
1-3 d postnatal Sprague Dawley rats. Slices of hippocampus 500-800
µm thick were treated with 1 mg/ml papain in oxygenated Leibovitz's
L-15 medium. Single-cell suspensions were obtained by trituration in modified Eagle's medium containing 5% horse serum, 5% fetal calf serum, 17 mM D-glucose, 400 µM
glutamine, 50 U/ml penicillin, and 50 µg/ml streptomycin. Cells
(75/mm2 for microcultures;
2100/mm2 for mass cultures) were plated onto plastic
culture dishes coated with a layer of collagen (mass cultures) or
0.15% agarose and collagen droplets (microcultures) as described
previously (Mennerick et al., 1995). Cultures were treated with
cytosine arabinoside (10 µM) after 3 d in
vitro and were used for experiments 1-16 d after plating. For
serum-free conditions, culture medium was replaced at 3 d in
vitro with the above medium, minus serum, and supplemented with 10 µg/ml insulin, 10 µg/ml transferrin, and 10 ng/ml sodium
selenite.
Western blots. Rat hippocampus was homogenized in ice-cold
buffer containing 20 mM Tris-HCl, pH 7.4, with 10%
sucrose, 10 µg/ml antipain, 5 µg/ml aprotinin, 5 µg/ml leupeptin,
0.1 mM PMSF, 1 mM EDTA, and 1 mM
EGTA. Cultures were washed twice with PBS before solubilization.
Hippocampal homogenates and cultures were solubilized in 2% SDS, 5%
2-mercaptoethanol, 65 mM Tris-HCl, pH 6.8, 10% glycerol,
and protease inhibitors. Aliquots of samples were subjected to 4-20%
SDS-PAGE electrophoresis and electroblotted onto nitroceullulose
membrane using a minigel and mini transblot apparatus (Bio-Rad,
Hercules, CA). Membranes were blocked (0.1% Tween and 5% nonfat
powdered milk in Tris-buffered saline) for 1 hr followed by
room-temperature incubation in primary antibodies diluted in
Tris-buffered saline [anti-B12 antibody (Lehre et al., 1995) (from
rabbit 68518), 0.25 µg/ml; C-terminal antibody (Rothstein et al.,
1994), 0.011 µg/ml]. Antibody binding was detected by a horseradish
peroxidase-conjugated anti-rabbit secondary and an ECL Western blotting
detection kit (Amersham, Arlington Heights, IL).
Immunohistochemistry. Cultures were rinsed in PBS and fixed
with 4% paraformaldehyde and 0.2% glutaraldehyde. After treatment with 1.5% hydrogen peroxide, cells were permeabilized and blocked in
10% normal goat serum and 0.1% Triton X-100. All antibodies were
characterized previously or obtained from commercial suppliers as
indicated, except for the anti-C510 EAAC1 antibody. The
affinity-purified anti-C510 (rabbit 69738) EAAC1 antibody was raised
against a C-terminal peptide of EAAC1 as described previously (Lehre et
al., 1995).
Cultures were incubated in primary antibodies diluted in blocking
solution at the following concentrations: 6.7 ng/ml [C-terminal affinity-purified anti-GLT-1 (Rothstein et al., 1994)], 20 ng/ml [N-terminal affinity-purified anti-B12 antibody from rabbit 68518 (Lehre et al., 1995)], and 1:20 working dilution of EAAT2 monoclonal antibody (Novo Castra, Newcastle upon Tyne, UK). For peptide block experiments, GLT-1 anti-B12 antibody was incubated overnight in 10-100-fold molar excess of free GLT-1 oligopeptide. Monoclonal antibody against GFAP (Chemicon, Temecula, CA) was used at a 1:100,000 working dilution. Monoclonal antibody against GABA (Chemicon) was used
at 2 µg/ml. Affinity-purified antibodies against EAAC1 (anti-C510;
rabbit 69738) and GLAST (anti A522; rabbit 68488) (Lehre et al., 1995)
were used at 20-200 ng/ml, diluted in blocking solution.
Visualization of antibody binding was routinely achieved using a Vector
ABC Elite Kit (Vector Laboratories, Burlingame, CA) using a horseradish
peroxidase/diaminobenzidine tetrahydrochloride chromogenic reaction
with nickel intensification. For some experiments, bound primary
antibody was visualized with a rhodamine-conjugated secondary antibody
(Chemicon) used at 2.5 µg/ml.
Electrophysiology. The extracellular bath solution for
synaptic physiology contained (in mM): NaCl, 140; KCl, 4.0;
CaCl2, 3.0; MgCl2, 1.0; HEPES,
10; D-APV, 0.05; and cyclothiazide, 0.01. The whole-cell
recording pipette solution for synaptic studies contained (in
mM): potassium gluconate, 130; NaCl, 4.0;
CaCl2, 0.5; EGTA, 5.0; HEPES, 10;
MgATP2, 2.0; and GTP, 0.5. The pH of solutions was
adjusted to 7.25. Whole-cell, voltage-clamp recordings of excitatory
autaptic currents (EACs) were performed from solitary neurons using
pipettes with an open-tip resistance of 2-5 M and with series
resistance compensated 90-100% using the compensation circuitry of an
Axopatch 1-D amplifier (Axon Instruments, Foster City, CA). Neurons
were stimulated with a brief (1.5 msec) voltage command pulse to 0 mV
from a holding potential of  70 mV. Solutions were exchanged with a
gravity-driven local perfusion system. Averages of two to five sweeps
in each condition were used for display and analysis. Although EACs
under the conditions of these experiments often decay with complex
kinetics (Mennerick and Zorumski, 1995), for purposes of comparison
among the various experimental conditions, EAC decays were fit with a
single-exponential function, generated from a Chebyshev-transform
algorithm (PClamp 6.0; Axon Instruments).
For exploring responses to applications of the transporter substrate
D-aspartate, the composition of extracellular solution was
altered as follows: cyclothiazide was omitted, and MgCl2
was increased to 2 mM. Additionally, 250 nM
tetrodotoxin, 10 µM MK-801, 50 µM
D-APV, 25 µM bicuculline methobromide,
10 µM 6-nitro-7-sulfamoylbenzo(f)quinoxaline-2,3-dione, and 50 µM GYKI 52466 were added. Large, presumably
excitatory neurons (Mennerick et al., 1995) and small, usually singly
nucleated islands of glia were examined for D-aspartate
responses. For some experiments, the intracellular pipette solution
contained thiocyanate in place of gluconate to examine
transporter-mediated anionic currents (Wadiche et al., 1995). For
estimation of D-aspartate current densities, current
amplitude was divided by total cell membrane capacitance, estimated
from biexponential fits to the decays of neuronal and glial membrane
responses to 15 mV hyperpolarizing voltage pulses (Mennerick et al.,
1995).
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RESULTS |
Localization of GLAST, EAAC1, and GLT-1 immunoreactivity
Staining of microcultures for the glial GLAST transporter and
neuronal EAAC1 transporter revealed the expected complementary staining
patterns in the two cell types. The vast majority of glia (>80%) in
microcultures exhibited detectable immunoreactivity when stained with
anti-A522 GLAST antibody. However, there was often considerable
variability in staining intensity, even among glia comprising an
individual microculture (Fig.
1A). Detectable neuronal staining was not present with this antibody (Fig.
1A). In contrast, staining with the anti-C510 (EAAC1)
antibody revealed staining of neuronal cell bodies and processes but
not of glia. Again, there was considerable variability in the levels of
EAAC1 staining among different neurons, often with the largest neurons staining most intensely (Fig. 1B).
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Figure 1.
Expression of GLAST and EAAC1 in hippocampal
microcultures. A, Bright-field photomicrograph of GLAST
expression in microcultures. GLAST was confined to glia, often
exhibiting a patchwork of staining intensities on a single
microculture. Note a cluster of unstained neurons
(arrow). Anti-A522 antibody, 100 ng/ml, was used for the
stain. B, Bright-field photomicrograph of EAAC1
staining. Staining was confined to neuronal somata and processes.
Anti-C510 antibody was used at 54 ng/ml. Scale bars: A,
80 µm; B, 40 µm.
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Immunostaining for the third forebrain transporter GLT-1 exhibited a
surprising pattern of labeling. Figure
2C shows immunoreactivity found on a microculture stained with an antibody raised against an
N-terminal region of the GLT-1 glutamate transporter (anti-B12 antibody) (Lehre et al., 1995). Only a minority of microculture glia
was immunoreactive for GLT-1. Immunoreactive clusters of a few glial
cells were often present on individual microcultures (Fig.
2B,C). These low levels of GLT-1
expression in microculture glia are consistent with descriptions of low
GLT-1 mRNA and protein levels in immature hippocampus (Shibata et al.,
1996; Ullensvang et al., 1997) and in other culture systems (Kondo et
al., 1995; Swanson et al., 1997).
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Figure 2.
Immunohistochemical evidence of GLT-1 expression
in microculture neurons. A, Phase-contrast
photomicrograph of a microculture of postnatal hippocampal neurons
8 d in vitro. The photomicrograph was obtained
after fixation and immunohistochemical processing. Note the numerous
neuronal somata, two of the largest of which are indicated by
arrows, and processes (e.g., arrowheads)
present. B, Immunofluorescence photomicrograph of the
microculture in A. The microculture was stained using an
antibody against GFAP and was visualized with a rhodamine-conjugated
secondary antibody to demonstrate the population of astrocytes
underlying the microculture. C, Bright-field
photomicrograph of GLT-1-like immunoreactivity. Note the
immunoreactivity of a small, clustered population of astrocytes on the
left of the microculture and the strong immunoreactivity
of many neuronal processes (e.g., arrowheads) but not
somata (e.g., arrows). Anti-B12 antibody, 20 ng/ml, and
anti-GFAP, 1:100,000, were used for the staining. Scale bar, 76 µm.
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Surprisingly, the most prominent GLT-1-immunoreactive
com- ponents of the microcultures were processes of microculture
neurons (Fig. 2C). In contrast to EAAC1 immunoreactivity,
neuronal somata were primarily devoid of immunoreactivity. GLT-1
immunoreactivity was most prominent in the proximal portions of
neuronal processes, and only a subset of processes appeared to be
labeled (Fig. 2A,C; see below).
Immunoreactivity was not detectable in neurons 1 d after plating,
when cells are primarily devoid of processes (data not shown).
Immunoreactivity gradually increased in subsequent days in
vitro, during the period of axon and dendrite elaboration.
Specificity of GLT-1 immunolocalization
Although mRNA for GLT-1 has been detected in neurons from several
brain regions by in situ hybridization (Schmitt et al., 1996; Torp et al., 1997), GLT-1 protein has not been detected in brain
neurons. Therefore, we performed several experiments to examine the
robustness and validity of the GLT-1-like immunoreactivity in
microculture neurons. To determine whether properties unique to the
microculture environment are responsible for GLT-1 immunoreactivity, we
examined GLT-1 immunoreactivity in conventional mass cultures of
postnatal hippocampal neurons. Figure 3
shows that mass-culture neurons, like microculture neurons, exhibit
prominent GLT-1 immunoreactivity in neuronal processes, with little
somatic immunoreactivity (Fig. 3B). As observed in
microcultures, only a minority of mass-culture glia was immunoreactive
for GLT-1. We also found that growing cultures in serum-free medium
beginning at day 3-6 in vitro did not alter neuronal GLT-1
immunoreactivity when examined between days 7-11 in vitro
(data not shown). These results indicate that neuronal GLT-1
immunoreactivity is a robust phenomenon, present in neurons grown in a
variety of culture conditions.
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Figure 3.
Mass-culture processes also display GLT-1
immunoreactivity. A, Phase-contrast photomicrograph of a
hippocampal mass culture 9 d in vitro.
B, Bright-field photomicrograph of the same field of
view shown in A. Staining with the anti-B12 antibody
reveals prominent staining of neuronal processes with little glial or
neuronal soma staining. Anti-B12 antibody was used at 20 ng/ml. Scale
bar, 76 µm.
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We compared Western blots using antibodies raised against N- and
C-terminal regions of the GLT-1 protein. Because of the larger amounts
of available tissue per culture dish, mass cultures rather than
microcultures were used for these experiments. The N-terminal anti-B12
antibody was compared with an antibody raised against a C-terminal
GLT-1 peptide (Rothstein et al., 1994). Both GLT-1 antibodies stained a
band of ~66 kDa in homogenates prepared from adult hippocampus and
from cultures (Fig. 4). GLT-1 in cultures possessed a much lower relative abundance than in adult hippocampus (Fig. 4), consistent with its restricted localization in
immunohistochemistry experiments.
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Figure 4.
Immunoblots of GLT-1. Immunoblots were prepared as
described in Materials and Methods either using the N-terminal anti-B12
antibody (left; 250 ng/ml) (Lehre et al., 1995) or using
a C-terminal antibody (right; 11 ng/ml) (Rothstein et
al., 1994). Lanes represent spleen (S; 20 µg of protein), adult hippocampus (H; 0.1 µg of
protein), and hippocampal cultures (HC; 50 µg of
protein). Molecular mass migration is indicated between the
panels in kDa.
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Several GLT-1 antibodies were also used in immunohistochemical
experiments and yielded indistinguishable patterns of staining. Figure
5 shows a comparison of the patterns of
immunostaining observed with the N-terminal (anti-B12) and C-terminal
GLT-1 antibodies in single-neuron microcultures. A monoclonal antibody
raised against a C-terminal portion of the human GLT-1 homolog EAAT2
also yielded staining indistinguishable from the other two antibodies
(data not shown).
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Figure 5.
Different GLT-1 antibodies yield similar cellular
staining patterns. Phase-contrast (A, C)
and bright-field (B, D) photomicrographs
of single-neuron microcultures immunostained for GLT-1 are shown.
A, B, A microculture immunostained using
an affinity-purified antibody directed against an N-terminal peptide
(anti-B12 antibody; 20 ng/ml). C, D, An
immunostain obtained using an affinity-purified antibody directed
against a C-terminal GLT-1 peptide (7 ng/ml). Scale bar, 76 µm. Note
that, with both antibodies, staining is localized to a nest of neuronal
processes and is primarily excluded from neuronal somata and glial
cells.
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Omission of primary antibody from the incubation solution caused a
complete loss of staining in both microculture glia and neurons (data
not shown). Similarly, Figure 6 shows
that when primary antibody was preabsorbed with an excess of the free
N-terminal peptide used to raise the antibody, both neuronal and glial
immunostaining was abolished. Together the above results support the
idea that the staining pattern of neuronal processes observed is
attributable to neuronal expression of GLT-1 protein.
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Figure 6.
Peptide block of immunoreactivity.
A, B, Phase-contrast
(A) and bright-field (B)
photomicrographs of a microculture fixed after 10 d in
vitro and stained with the anti-B12 GLT-1 antibody (20 ng/ml).
C, D, Phase-contrast
(C) and bright-field (D)
photomicrographs of a microculture from the same plating of the
culture shown in A and B. The
microculture was fixed and processed for immunohistochemistry at the
same time and using the same reagents as the microculture in
A and B except that the primary antibody
was preabsorbed with an excess of the free-peptide immunogen. Scale
bar, 76 µm.
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Cellular localization of GLT-1 immunoreactivity
Postnatal hippocampal cultures comprise both excitatory
(glutamatergic) and inhibitory (GABAergic) neurons (Mennerick et al., 1995). To determine whether GLT-1 protein is differentially expressed in one neuronal phenotype versus another, we performed double-labeling studies with a GLT-1 antibody and with an antibody against GABA, to
identify inhibitory neurons. Figure 7
shows that neuronal processes labeled for GLT-1 were never double
labeled with the antibody against GABA. This result indicates that
neuronal GLT-1 expression is limited primarily, if not exclusively, to
excitatory neurons in hippocampal microcultures.
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Figure 7.
Segregation of GLT-1 immunoreactivity and GABA
immunoreactivity. A, Phase-contrast photomicrograph of a
microculture 11 d in vitro. B,
Fluorescence photomicrograph showing GABA immunoreactivity of the
microculture depicted in A. The anti-GABA antibody was
used at 2 µg/ml. C, Bright-field photomicrograph
showing GLT-1 immunoreactivity (C-terminal antibody, 7 ng/ml) of the
same microculture. A GABA-positive process (presumed dendrite) is
indicated by the arrow in A-C. Scale
bar, 38 µm.
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Although axon terminals of excitatory neurons can be shown
biochemically to possess strong glutamate transport activity (Gundersen et al., 1993), immunocytochemical localization studies have failed to
detect evidence of a presynaptic transporter (Rothstein et al., 1994;
Chaudhry et al., 1995; Yamada et al., 1996). To determine whether axons
or dendrites might preferentially exhibit GLT-1 immunoreactivity, we
combined staining for the dendritic marker MAP2 (Caceres et al., 1986)
and GLT-1. Results showed extensive overlap of the staining patterns
yielded by the MAP2 and GLT-1 antibodies (Fig.
8). Additionally, many processes that
showed no MAP2 immunoreactivity were also GLT-1 negative (Fig. 8).
However, there were also instances of MAP2-positive processes that were GLT-1 negative. These processes likely arose from GABAergic cells (see
Fig. 8). Finally, we also found some examples of GLT-1-positive processes that showed no detectable MAP2 immunoreactivity. These could
represent axons or could represent dendrites expressing MAP2 at levels
below the threshold for detection. These experiments suggest that
neuronal GLT-1 expression localizes to specific cellular and
subcellular compartments in hippocampal microcultures.
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Figure 8.
GLT-1 immunoreactivity is primarily colocalized
with a dendritic marker. A, Phase-contrast
photomicrograph of microculture neurons. B, The same
field depicting MAP2 immunofluorescence. C, Bright-field
view showing GLT-1 immunoreactivity (C-terminal antibody, 7 ng/ml) of
the same microculture. The arrow indicates a
double-labeled process. The arrowhead indicates a
neuronal process labeled for MAP2 but devoid of GLT-1 immunoreactivity.
Note that an occasional neuronal process positive for GLT-1
immunoreactivity but negative for MAP2 immunoreactivity can also be
found. Scale bar, 76 µm.
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Evidence that neuronal GLT-1 is functional
Immunostaining does not give a clear picture of the density of
functional transporters in neuronal or glial membranes. Because glutamate transport is electrogenic, it is possible to monitor the
function of glutamate transporters using standard electrophysiological techniques (Brew and Attwell, 1987). To obtain a comparative estimate of the density of functional transporters in neurons and glia and to
determine the potential contribution of GLT-1 to the glutamate transport, we recorded from neurons and glia voltage clamped at 70
mV. In the presence of competitive and noncompetitive glutamate receptor blockade, 100 µM D-aspartate
elicited small inward currents in both neurons and glia. The currents
were sensitive to substitution of lithium for extracellular sodium
(n = 10 neurons and 6 glia; data not shown), indicating
the currents were attributable to sodium-dependent glutamate transport.
We used the GLT-1-selective inhibitor dihydrokainate (DHK) to assess
the potential contribution of GLT-1 to transport currents. The
amplitude of neuronal D-aspartate responses was occluded by
56 ± 6% (n = 7) in the presence of 500 µM DHK (Fig. 9). In
contrast to neurons, glial D-aspartate currents were less
sensitive to DHK, consistent with the low levels of GLT-1 expression
and high levels of GLAST expression observed in microculture glia (Fig.
9B). In neurons but not in glial cells, DHK applied alone
resulted in a small inward current (14.4 ± 3.8 pA;
n = 7; data not shown) despite the presence of high
concentrations of postsynaptic receptor antagonists. The nature of this
current was not explored further.
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Figure 9.
DHK-sensitive D-aspartate currents in
neurons and glia. A, A response of a neuron to 100 µM D-aspartate
(d-Asp) applied for the
duration indicated by the bars above the
traces. D-Aspartate was applied either alone
(left) or with 500 µM DHK
(right). Traces are averages of five
interleaved responses in each condition and were filtered at 50 Hz. In
the DHK condition, DHK was preapplied for 2 sec before and after
D-aspartate application. B, DHK (500 uM) more strongly inhibited neuronal
D-aspartate (100 uM) responses (filled
bar) than glial D-aspartate responses (open bar;
asterisk denotes p < 0.01, independent t
test, n = 7 neurons and 6 glia).
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Glutamate transporters can also be functionally distinguished on the
basis of a recently described anion permeability (Wadiche et al., 1995;
Eliasof and Jahr, 1996). The rank order of anion permeability for the
forebrain transporters is EAAT1 (GLAST) > EAAT3 (EAAC1) > EAAT2
(GLT-1) (Wadiche et al., 1995). As in the previous experiment, we first
recorded from neurons and glia with gluconate as the main pipette anion
and at a holding potential of 70 mV, near ECl.
Under these conditions, the response recorded to
D-aspartate applications should be nearly completely
mediated by transport current rather than by anion current. The current density in glial cells filled with gluconate was nearly threefold larger than that in neurons (Fig. 10),
suggesting a higher transport rate per transporter or a higher density
of functional transporters in glia. We next substituted thiocyanate for
gluconate in the pipette solution. Thiocyanate is the most permeant of
the chaotropic anions through the transporter anion conductance
(Eliasof and Jahr, 1996). With thiocyanate-filled cells, the difference
between current density in glia and neurons became much more dramatic (Fig. 10), indicating that glial transporters have a much higher anion
permeability than do neurons. These results are consistent with GLAST
(EAAT1) being the predominant transporter in microculture glia and with
EAAC1 (EAAT3) and GLT-1 (EAAT2) functioning predominantly in neurons.
The results also make it unlikely that either EAAT4 or EAAT5, two
recently cloned transporters from cerebellum and retina (Fairman et
al., 1995; Arriza et al., 1997), contribute significantly to neuronal
transport currents in hippocampal microcultures. These two transporters
have a much larger anion conductance than do the forebrain
transporters.
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Figure 10.
Anion permeability of glial and neuronal
transporters. A, Membrane currents in a glial cell in
response to 100 µM D-aspartate
(d-Asp) applied for the
duration indicated by the bars above the
traces. The cell was first filled with a gluconate-based
whole-cell pipette solution. The pipette was gently removed and
replaced with a whole-cell pipette containing thiocyanate substituted
for gluconate. Membrane capacitance of the cell was 47 pF.
B, The same protocol described in A
performed on a microculture neuron. Membrane capacitance was 80 pF.
C, D, A comparison of neuronal and glial
D-aspartate current density measured in cells filled either
with gluconate (C) or with thiocyanate
(D). A two-way (cell type by pipette solution)
ANOVA revealed a significant interaction between cell type and pipette
solution (p < 0.001; n = 10 gluconate-filled glia and 5 thiocyanate-filled glia;
n = 13 gluconate-filled neurons and 9 thiocyanate-filled neurons). Glial cells exhibited a larger current
density than did neurons in both conditions
(p < 0.01, Bonferroni-corrected
t tests), but the difference was much larger with a
thiocyanate-based pipette solution. Statistically significant
differences between the current density obtained with different
internal solutions were also detected in both cell types
(p < 0.01, Bonferroni-corrected
t tests). Not obvious from the bar graphs
is the difference between neuronal current density with a gluconate
pipette solution (0.13 ± 0.03 pA/pF) versus that with a
thiocyanate pipette solution (0.30 ± 0.06 pA/pF).
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At some glutamate synapses, including microculture synapses, transport
inhibition can prolong the decay of EPSCs (Barbour et al., 1994;
Mennerick and Zorumski, 1995; Otis et al., 1996). However, the
molecular identity of the transporters responsible for these effects
remains to be elucidated. To test for a role of GLT-1 in modulating the
time course of synaptic glutamate, we examined decays of EACs mediated
by AMPA receptors in single-neuron microcultures. We compared the
effects of DHK, a selective blocker of GLT-1 (Pines et al., 1992;
Arriza et al., 1994), with the effects of the broad-spectrum
transporter substrate L-trans-pyrollidine dicarboxylic acid (PDC) (Arriza et al., 1994). The AMPA receptor potentiator cyclothiazide (CYZ; 10 µM) was included in
bath solutions to enhance our ability to detect effects of glutamate
uptake inhibition (Mennerick and Zorumski, 1995). At 500 µM, DHK had a consistently weaker effect on the decay of
the EAC than did a low (50 µM) concentration of PDC (Fig.
11A). In six cells,
DHK increased EAC decay time constants by 21 ± 3%, whereas PDC
increased the decay time constants by 47 ± 8%. Essentially
identical results were obtained when the effects of as much as 1 mM DHK were compared with that of another broad-spectrum
substrate inhibitor, threo-3-hydroxy aspartate (THA; 30 µM; Fig. 11B). Based on the experiments
of Figure 11, we conclude that although neuronal and glial GLT-1
contribute measurably to glutamate transport during a synaptic event,
GLT-1 is not the primary transporter involved.
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Figure 11.
DHK prolongs EACs, but less than broad-spectrum
substrate inhibitors. A, Superimposed
traces showing the averages of responses from a solitary
excitatory neuron in the absence of transport inhibition
(Control) interleaved with responses obtained in
the presence of 500 µM DHK and of 50 µM
PDC. B, Experiment similar to that shown in
A except that 1 mM DHK and 30 µM THA, another broad-spectrum transporter substrate,
were used.
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The effects of DHK on EACs in Figure 11 suggest that GLT-1 participates
in limiting the lifetime of synaptically released glutamate, but these
results do not address the question of whether neuronal GLT-1
specifically is capable of participating in the clearance of synaptic
glutamate. Toward this end, we designed an experiment to examine
neuronal GLT-1 contributions to the decay of the synaptic glutamate
transient in the absence of glial transport. Previously, we have shown
that microculture glia make a strong contribution to synaptic glutamate
transport (Mennerick and Zorumski, 1994). This conclusion was based on
the ability to record from and depolarize microculture glia during a
synaptic event, thereby exploiting the voltage dependence of glutamate
transport (Brew and Attwell, 1987) to slow the rate of glial transport.
However, toward the goal of abolishing glial transport, this technique
suffers from its dependence on a high degree of spatial voltage control
over the glial cells, which may not be achieved in recordings from multiglial microcultures.
Therefore, to abolish glial glutamate transport in the present
experiments, we killed microculture glia using a sharp glass pipette to
stab glial nuclear and cytoplasmic regions repeatedly before recording
from an overlying excitatory neuron. This treatment resulted in the
immediate swelling of glial nuclei, blebbing of the glial plasma
membrane, and retraction of the glial membrane from the edges of the
microculture. Glia stabbed in the presence of trypan blue showed
immediate accumulation of the dye into the nucleus, indicating that the
cell membrane was effectively rendered nonviable (e.g., Fig.
12B,C).
Similarly, stabs performed during a whole-cell, current-clamp recording
from the stabbed cell resulted in the immediate abolishment of the
membrane potential (61.4 ± 5.2 to 0.7 ± 5.9 mV;
n = 3). For reasons that are unclear, the baseline
glial resting potential was more positive than that observed in glia in
more intact preparations (Bergles and Jahr, 1997; Clark and Barbour,
1997). However, the experiment indicates that the stabbing procedure
effectively abolished the ion gradients necessary to drive glutamate
transport.
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[in a new window]
|
Figure 12.
DHK prolongs EPSCs in the absence of viable glial
cells. A, Left, Traces
showing averages of interleaved EACs in the absence
(Control) or presence of 500 µM DHK.
Mechanical stab wounds with a sharp glass pipette had rendered
underlying glia nonviable (see B, C).
Right, The best-fit time constants of 13 interleaved
responses in each condition; filled bar, control; open
bar, DHK (*p < 0.01, unpaired two-tailed
t test; data from the same cell shown on the
left). B, C, Phase
contrast (B) and bright-field
(C) photomicrographs of the microculture from
which the traces in A were recorded.
Immediately after the experiment in A, 0.1% trypan blue
was added to the culture dish for 10 min. The arrow in
C indicates a trypan-positive glial nucleus. Two other
trypan-positive glial nuclei are also visible. After photography was
completed, the microculture was stabbed again repeatedly, and trypan
blue was added a second time to the dish. No other trypan-positive
nuclei were observed after the second insult, indicating that no other
viable glia were present in this microculture (data not shown). The
arrowhead indicates the remnants of the soma of the
recorded neuron, primarily destroyed after removing the whole-cell
recording pipette. Scale bar: B, C, 32 µm. D, E, Phase-contrast
(D) and bright-field (E)
photomicrographs of another microculture not used for experiments but
in the same dish as that represented in A-C. Note the
absence of trypan-positive nuclei. Scale bar: D,
E, 32 µm.
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Recordings from solitary excitatory neurons were performed 5-10 min
after stabbing underlying glia, again in the presence of 10 µM CYZ to enhance our ability to detect residual
glutamate. Under these conditions, DHK application resulted in a
statistically significant prolongation of EAC decays (Fig. 12).
Mean time constants in eight neurons changed from 17.4 ± 3.4 to 20.3 ± 4.8 msec (p < 0.05, paired
one-tailed t test). Immediately after the experiment, trypan
blue was added to the culture dish to assess cell viability. In all
eight cases, all underlying glia were compromised by the stabbing
procedure (e.g., Fig. 12B,C).
Unstabbed glia from the same dish exhibited no trypan-positive nuclei
(Fig. 12D,E), indicating that
trypan staining was specific to nonviable cells.
In five of the above eight cases, THA was also applied to cells in
which DHK was tested. Application of a moderate concentration of THA
had a smaller effect than did DHK (15 ± 5% prolongation with 500 µM DHK; 6 ± 3% prolongation with 50 µM THA), in contrast to the results obtained in Figure
11. Importantly, these results suggest that DHK prolongs synaptic
currents via a strong effect on GLT-1 rather than via a weak effect on
another transporter. The diminished THA effects after killing glia are
also consistent with a strong contribution of glial uptake during
evoked synaptic events in intact microcultures (Mennerick and Zorumski,
1994). In conclusion, these experiments demonstrate that neuronal GLT-1 is functional and capable of participating in the clearance of synaptic
glutamate.
| |
DISCUSSION |
The present results represent evidence that brain neurons can
express the glutamate transporter protein GLT-1 (EAAT2) (Pines et al.,
1992; Arriza et al., 1994). GLT-1 was localized primarily to the
dendrites of presumed excitatory neurons in microcultures and
conventional mass cultures of postnatal hippocampal cells. Although
there is evidence of expression of GLT-1 protein in certain classes of
retinal photoreceptors and bipolar cells (Grunert et al., 1994; Rauen
and Kanner, 1994), GLT-1 mRNA, but not protein, has been detected in
synaptically mature hippocampal neurons (Schmitt et al., 1996; Torp et
al., 1997). Interestingly, a recent genetic knock-out of GLT-1 in mice
revealed a drastic diminution of synaptosomal transport, suggesting the
possibility of neuronal expression of GLT-1 (Tanaka et al., 1997).
Although glial contamination of synaptosomes cannot be excluded,
another very recent paper described detectable levels of GLT-1
expression in glial-poor cultures of cortical neurons and a large
contribution of DHK-sensitive transport to neuronal glutamate uptake
(Wang et al., 1998).
There are several possible reasons for the detection of neuronal GLT-1
protein in the present study compared with previous studies (Danbolt et
al., 1992; Levy et al., 1993; Rothstein et al., 1994; Chaudhry et al.,
1995; Lehre et al., 1995; Schmitt et al., 1996). First, the failure to
detect neuronal GLT-1 in light and electron microscopic studies on
intact brain tissue does not exclude neuronal expression of GLT-1. The
detection failure only implies that the highest GLT-1 concentrations
are found in brain astrocytes in situ. The GLT-1
concentration in brain tissue is extremely high, with GLT-1
representing as much as 1% of brain membrane protein (Danbolt et al.,
1990). Therefore, it is possible that in situ comparatively
low levels of neuronal GLT-1 protein, although perhaps functionally
significant, have gone unappreciated. Extremely high levels of
astrocyte expression of GLT-1 may have prompted use of antibody
concentrations below those that would reveal significant neuronal
labeling. Therefore, the low levels of GLT-1 expression in cultured
astrocytes may have helped facilitate our detection of neuronal
GLT-1.
Second, because previous electron microscopy has not revealed neuronal
labeling in situ, it is also possible that neuronal GLT-1
expression is a reflection of the ex vivo environment of the
cultured cells. Thus, GLT-1 expression in brain neurons may be
suppressed by novel regulatory mechanisms. This possibility was
originally suggested after detection of GLT-1 mRNA in certain neuronal
populations (Schmitt et al., 1996; Torp et al., 1997). This idea has
not previously gained support because of the failure to detect neuronal
GLT-1 protein. The findings presented here are therefore important as
the first demonstration that under appropriate conditions, brain
neurons express functional GLT-1 protein. Consequently, when
biochemical studies of human pathological tissue or of mammalian cell
cultures are interpreted, it should not be assumed that GLT-1 protein
is always glial. The findings presented here suggest the cellular
localization may change under certain conditions.
Third, the possibility exists that the neuronal expression of GLT-1 in
our cultures could be a reflection of the immature state of the neurons
used. This seems unlikely because the levels of GLT-1 are undetectable
at birth and gradually increase in the first 2 weeks after birth
(Ullensvang et al., 1997).
The present results suggest that neuronal GLT-1 protein is expressed
primarily, if not exclusively, in non-GABAergic microculture cells.
Because glutamatergic cells comprise the other major transmitter phenotype in postnatal hippocampal microcultures (Segal and Furshpan, 1990; Bekkers and Stevens, 1991; Mennerick et al., 1995), we conclude that neuronal GLT-1 expression is limited to excitatory cells. These
results are consistent with in situ hybridization studies of
brain, in which neuronal GLT-1 mRNA is found primarily in the principal
neurons of the hippocampus, especially CA3 pyramidal cells (Schmitt et
al., 1996; Torp et al., 1997). Although the reasons for the selective
expression in excitatory neurons are unclear, principal cells of the
hippocampus are highly susceptible to cell death in many models of
glutamate excitotoxicity (Choi, 1994). It is possible that selective
expression of GLT-1 and other glutamate transporters in these
glutamatergic cells helps maintain low ambient glutamate concentrations
in the microenvironment of the cells, thereby protecting against
glutamate excitotoxicity.
At the subcellular level, antibody localization studies have usually
found neuronal transporters at the postsynaptic side of the synaptic
cleft but not at the presynaptic side (Rothstein et al., 1994; Yamada
et al., 1996). Notable exceptions to these studies are several studies
that have found GLT-1 immunoreactivity in certain photoreceptors and
bipolar cells of the mammalian retina. In these studies,
immunoreactivity is found throughout the cells, apparently including
the presynaptic terminals (Grunert et al., 1994; Rauen and Kanner,
1994). Despite this precedent for a presynaptic localization of GLT-1
in retina, our MAP2 colocalization experiments suggest that in
excitatory hippocampal cells, GLT-1 is primarily a postsynaptic
transporter.
Interestingly, neuronal GLT-1 can apparently participate in the
clearance of synaptically released glutamate. However, the effects are
weak compared with the pharmacological blockade of neuronal and glial
transporters with broad-spectrum substrates such as PDC. Our previous
results suggest that glial cells are strong contributors to synaptic
glutamate uptake in microcultures (Mennerick and Zorumski, 1994). Based
on the pharmacological and immunohistochemical evidence in the present
study, it is likely that these effects are mediated predominantly by
GLAST.
In summary, we have presented evidence that functional GLT-1 glutamate
transporter protein can be expressed in brain neurons. The expression
of GLT-1 in cultured neurons is specific to excitatory cells and is
specifically localized to the processes of these cells. The results
make it more likely that GLT-1 may be expressed by hippocampal neurons
in situ and that certain environmental conditions may alter
the expression of neuronal GLT-1 protein.
| |
FOOTNOTES |
Received Feb. 16, 1998; revised March 31, 1998; accepted April 1, 1998.
This work was supported by National Institutes of Health Grants
MH-00964 and MH-45493 and a grant from the Bantly Foundation (C.F.Z.).
S.M. was supported by a fellowship from the McDonnell Center for
Cellular and Molecular Neurobiology and a Lucille P. Markey
postdoctoral fellowship. We thank Drs. John Olney and Nuri Farber
(Washington University) for help with image processing.
Correspondence should be addressed to Dr. Steven Mennerick, Department
of Psychiatry, Washington University School of Medicine, 4940 Children's Place, St. Louis, MO 63110
| |
REFERENCES |
-
Arriza JL,
Fairman WA,
Wadiche JI,
Murdoch GH,
Kavanaugh MP,
Amara SG
(1994)
Functional comparisons of three glutamate transporter subtypes cloned from human motor cortex.
J Neurosci
14:5559-5569[Abstract].
-
Arriza JL,
Eliasof S,
Kavanaugh MP,
Amara SG
(1997)
Excitatory amino acid transporter 5, a retinal glutamate transporter coupled to a chloride conductance.
Proc Natl Acad Sci USA
94:4155-4160[Abstract/Free Full Text].
-
Barbour B,
Keller BU,
Llano I,
Marty A
(1994)
Prolonged presence of glutamate during excitatory synaptic transmission to cerebellar Purkinje cells.
Neuron
12:1331-1343[Web of Science][Medline].
-
Bekkers JM,
Stevens CF
(1991)
Excitatory and inhibitory autaptic currents in isolated hippocampal neurons maintained in cell culture.
Proc Natl Acad Sci USA
88:7834-7838[Abstract/Free Full Text].
-
Bergles DE,
Jahr CE
(1997)
Synaptic activation of glutamate transporters in hippocampal astrocytes.
Neuron
19:1297-1308[Web of Science][Medline].
-
Brew H,
Attwell D
(1987)
Electrogenic glutamate uptake is a major current carrier in the membrane of axolotl retinal glial cells.
Nature
327:707-709[Medline].
-
Caceres A,
Banker GA,
Binder L
(1986)
Immunocytochemical localization of tubulin and microtubule-associated protein 2 during the development of hippocampal neurons in culture.
J Neurosci
6:714-722[Abstract].
-
Chaudhry FA,
Lehre KP,
van Lookeren Campagne M,
Ottersen OP,
Danbolt NC,
Storm-Mathisen J
(1995)
Glutamate transporters in glial plasma membranes: highly differentiated localizations revealed by quantitative ultrastructural immunocytochemistry.
Neuron
15:711-720[Web of Science][Medline].
-
Choi DW
(1994)
Glutamate receptors and the induction of excitotoxic neuronal death.
Prog Brain Res
100:47-51[Web of Science][Medline].
-
Clark BA,
Barbour B
(1997)
Currents evoked in Bergmann glial cells by parallel fibre stimulation in rat cerebellar slices.
J Physiol (Lond)
502:335-350[Abstract/Free Full Text].
-
Danbolt NC,
Pines G,
Kanner BI
(1990)
Purification and reconstitution of the sodium- and potassium-coupled glutamate transport glycoprotein from rat brain.
Biochemistry
29:6734-6740[Medline].
-
Danbolt NC,
Storm-Mathisen J,
Kanner BI
(1992)
An [Na+ + K+] coupled L-glutamate transporter purified from rat brain is located in glial cell processes.
Neuroscience
51:295-310[Web of Science][Medline].
-
Eliasof S,
Jahr CE
(1996)
Retinal glial cell glutamate transporter is coupled to an anionic conductance.
Proc Natl Acad Sci USA
93:4153-4158[Abstract/Free Full Text].
-
Fairman WA,
Vandenberg RJ,
Arriza JL,
Kavanaugh MP,
Amara SG
(1995)
An excitatory amino-acid transporter with properties of a ligand-gated chloride channel.
Nature
375:599-603[Medline].
-
Furuta A,
Rothstein JD,
Martin LJ
(1997)
Glutamate transporter protein subtypes are expressed differentially during rat CNS development.
J Neurosci
17:8363-8375[Abstract/Free Full Text].
-
Grunert U,
Martin PR,
Wassle H
(1994)
Immunocytochemical analysis of bipolar cells in the macaque monkey retina.
J Comp Neurol
348:607-627[Web of Science][Medline].
-
Gundersen V,
Danbolt NC,
Ottersen OP,
Storm-Mathisen J
(1993)
Demonstration of glutamate/aspartate uptake activity in nerve endings by use of antibodies recognizing exogenous D-aspartate.
Neuroscience
57:97-111[Web of Science][Medline].
-
Kondo K,
Hashimoto H,
Kitanaka J-I,
Sawada M,
Suzumura A,
Marunouchi T,
Baba A
(1995)
Expression of glutamate transporters in cultured glial cells.
Neurosci Lett
188:140-142[Web of Science][Medline].
-
Lehre KP,
Levy LM,
Ottersen OP,
Storm-Mathisen J,
Danbolt NC
(1995)
Differential expression of two glial glutamate transporters in the rat brain: quantitative and immunocytochemical observations.
J Neurosci
15:1835-1853[Abstract].
-
Levy LM,
Lehre KP,
Rolstad B,
Danbolt NC
(1993)
A monoclonal antibody raised against an [Na+ + K+] coupled L-glutamate transporter purified from rat brain confirms glial cell localization.
FEBS Lett
317:79-84[Web of Science][Medline].
-
Martin LJ,
Brambrink AM,
Lehmann C,
Porteracailliau C,
Koehler R,
Rothstein JD,
Traystman RJ
(1997)
Hypoxia-ischemia causes abnormalities in glutamate transporters and death of astroglia and neurons in newborn striatum.
Ann Neurol
42:335-348[Web of Science][Medline].
-
Mennerick S,
Zorumski CF
(1994)
Glial contributions to excitatory neurotransmission in cultured hippocampal cells.
Nature
368:59-62[Medline].
-
Mennerick S,
Zorumski CF
(1995)
Presynaptic influence on the time course of fast excitatory synaptic currents in cultured hippocampal cells.
J Neurosci
15:3178-3192[Abstract].
-
Mennerick S,
Que J,
Benz A,
Zorumski CF
(1995)
Passive and synaptic properties of neurons grown in microcultures and in mass cultures.
J Neurophysiol
73:320-332[Abstract/Free Full Text].
-
Otis TS,
Wu Y-C,
Trussell LO
(1996)
Delayed clearance of transmitter and the role of glutamate transporters at synapses with multiple release sites.
J Neurosci
16:1634-1644[Abstract/Free Full Text].
-
Otis TS,
Kavanaugh MP,
Jahr CE
(1997)
Postsynaptic glutamate transport at the climbing fiber-Purkinje cell synapse.
Science
277:1515-1518[Abstract/Free Full Text].
-
Pines G,
Danbolt NC,
Bjørås M,
Zhang Y,
Bendahan A,
Eide L,
Koepsell H,
Storm-Mathisen J,
Seeberg E,
Kanner BI
(1992)
Cloning and expression of a rat brain L-glutamate transporter.
Nature
360:464-467[Medline].
-
Rauen T,
Kanner BI
(1994)
Localization of the glutamate transporter Glt-1 in rat and macaque monkey retinae.
Neurosci Lett
169:137-140[Web of Science][Medline].
-
Rothstein JD
(1996a)
Knockout of glutamate transporters reveals a major role for astroglial transport in excitotoxicity and clearance of glutamate.
Neuron
16:675-686[Web of Science][Medline].
-
Rothstein JD
(1996b)
Therapeutic horizons for amyotrophic lateral sclerosis.
Curr Opin Neurobiol
6:679-687[Web of Science][Medline].
-
Rothstein JD,
Martin L,
Levey AI,
Dykes-Hoberg M,
Jin L,
Wu D,
Nash N,
Kuncl RW
(1994)
Localization of neuronal and glial glutamate transporters.
Neuron
13:713-725[Web of Science][Medline].
-
Schmitt A,
Asan E,
Puschel B,
Jons T,
Kugler P
(1996)
Expression of the glutamate transporter GLT1 in neural cells of the rat central nervous system: non-radioactive in situ hybridization and comparative immunocytochemistry.
Neuroscience
71:989-1004[Web of Science][Medline].
-
Segal MM,
Furshpan EJ
(1990)
Epileptiform activity in microcultures containing small numbers of hippocampal neurons.
J Neurophysiol
64:1390-1399[Abstract/Free Full Text].
-
Shibata T,
Watanabe M,
Tanaka K,
Wada K,
Inoue Y
(1996)
Dynamic changes in expression of glutamate transporter mRNAs in developing brain.
NeuroReport
7:705-709[Web of Science][Medline].
-
Swanson RA,
Miller JW,
Rothstein JD,
Farrell K,
Stein BA,
Longuemare MC
(1997)
Neuronal regulation of glutamate transporter subtype expression in astrocytes.
J Neurosci
17:932-940[Abstract/Free Full Text].
-
Tanaka K,
Watase K,
Manabe T,
Yamada K,
Watanabe M,
Takahashi K,
Iwama H,
Nishikawa T,
Ichihara N,
Kikuchi T,
Okuyama S,
Kawashima N,
Hori S,
Takimoto M,
Wada K
(1997)
Epilepsy and exacerbation of brain injury in mice lacking the glutamate transporter GLT-1.
Science
276:1699-1702[Abstract/Free Full Text].
-
Tong G,
Jahr CE
(1994)
Block of glutamate transporters potentiates postsynaptic excitation.
Neuron
13:1195-1203[Web of Science][Medline].
-
Torp R,
Hoover F,
Danbolt NC,
StormMathisen J,
Ottersen OP
(1997)
Differential distribution of the glutamate transporters GLT1 and rEAAC1 in rat cerebral cortex and thalamus: an in situ hybridization analysis.
Anat Embryol (Berl)
195:317-326[Medline].
-
Ullensvang K,
Lehre KP,
StormMathisen J,
Danbolt NC
(1997)
Differential developmental expression of the two rat brain glutamate transporter proteins GLAST and GLT.
Eur J Neurosci
9:1646-1655[Web of Science][Medline].
-
Wadiche JI,
Amara SG,
Kavanaugh MP
(1995)
Ion fluxes associated with excitatory amino acid transport.
Neuron
15:721-728[Web of Science][Medline].
-
Wang GJ,
Chung HJ,
Schnuer J,
Pratt K,
Zable AC,
Kavanaugh MP,
Rosenberg PA
(1998)
High affinity glutamate transport in rat cortical neurons in culture.
Mol Pharmacol
53:88-96[Abstract/Free Full Text].
-
Yamada K,
Watanabe M,
Shibata T,
Tanaka K,
Wada K,
Inoue Y
(1996)
EAAT4 is a post-synaptic glutamate transporter at Purkinje cell synapses.
NeuroReport
7:2013-2017[Web of Science][Medline].
Copyright © 1998 Society for Neuroscience 0270-6474/98/18124490-10$05.00/0
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|
S. Kojima, T. Nakamura, T. Nidaira, K. Nakamura, N. Ooashi, E. Ito, K. Watase, K. Tanaka, K. Wada, Y. Kudo, et al.
Optical Detection of Synaptically Induced Glutamate Transport in Hippocampal Slices
J. Neurosci.,
April 1, 1999;
19(7):
2580 - 2588.
[Abstract]
[Full Text]
[PDF]
|
|
|
|
|
|
|
S. Li, G. A. R. Mealing, P. Morley, and P. K. Stys
Novel Injury Mechanism in Anoxia and Trauma of Spinal Cord White Matter: Glutamate Release via Reverse Na+-dependent Glutamate Transport
J. Neurosci.,
July 15, 1999;
19(14):
RC16 - RC16.
[Abstract]
[Full Text]
[PDF]
|
|
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Abstract
Introduction
