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The Journal of Neuroscience, December 1, 2002, 22(23):10153-10162
Comparison of Coupled and Uncoupled Currents during Glutamate
Uptake by GLT-1 Transporters
Dwight E.
Bergles,
Anastassios V.
Tzingounis, and
Craig E.
Jahr
Vollum Institute, Oregon Health and Science University, Portland,
Oregon 97201-3098
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ABSTRACT |
The transport of glutamate across the plasma membrane is coupled to
the movement of cations (Na+, K+,
and H+) that are necessary for glutamate uptake and
transporter cycling as well as anions that are uncoupled from the flux
of glutamate. Although the relationship between these coupled
(stoichiometric) and uncoupled (anion) transporter currents is poorly
understood, transporter-associated anion currents often are used
to monitor transporter activity. To define the kinetic relationship
between these two components, we have recorded transporter currents
associated with stoichiometric and anion charge movements occurring in
response to the rapid application of L-glutamate to
outside-out patches from human embryonic kidney cells expressing GLT-1
transporters. Transporter-associated anion currents were approximately
twice as slow to rise and decay as stoichiometric transport currents, but the presence of permeant anions did not slow transporter cycling. A
kinetic model for GLT-1 was developed to simulate the behavior of both
components of the transporter current and to estimate the capture
efficiency of GLT-1. In this model the K+
counter-transport step was defined as rate-limiting, consistent with
the slowing of transporter cycling after the substitution of internal
K+ with Cs+ or
Na+. The model predicts that in physiological
conditions ~35% of GLT-1 transporters function as buffers, releasing
glutamate back into the extracellular space after binding.
Key words:
glutamate transporter; GLT-1; EAAT2; uptake; astrocyte; patch clamp
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INTRODUCTION |
Glutamate transporters support the
movement of glutamate across cell membranes. In the CNS this transport
is necessary to remove the glutamate that is released during excitatory
synaptic transmission, because glutamate is not subject to
extracellular enzymatic degradation. The movement of glutamate into
cells against a concentration gradient is achieved by harnessing energy
stored in the electrochemical gradients for
Na+, K+, and
H+; the translocation of each molecule of
glutamate is accompanied by the inward movement of 3 Na+ and 1 H+
and the outward movement of 1 K+ (Zerangue
and Kavanaugh, 1996
; Levy et al., 1998), an unbalanced movement of
charge that results in an inward current. In addition to this movement
of coupled charges, glutamate transporters allow certain anions to flow
across the membrane uncoupled from the movement of glutamate (Fairman
et al., 1995; Wadiche et al., 1995a; Eliasof and Jahr, 1996). Because
many more anions than coupled charges traverse the membrane during each
cycle of transport, this feature has been exploited to resolve
transporter currents in outside-out patches in which the density of
transporters is too low to resolve reliably the currents mediated by
the movement of coupled charges (Bergles and Jahr, 1997; Otis et al.,
1997; Otis and Jahr, 1998; Wadiche and Kavanaugh, 1998; Mennerick et al., 1999). By combining outside-out patch recording with rapid solution exchange techniques, we and others have found that it is
possible to record charge movements associated with discrete steps in
the transport cycle, revealing the speed at which glutamate and ion
binding occur and conformational changes in the protein are induced.
However, relating anion-potentiated transporter currents to the
movement of glutamate relies on detailed knowledge about the
relationship between the movement of coupled and uncoupled charges
during transport.
The flux of both anions and substrate appear to be catalyzed by the
transporter itself, because both components are present when
transporters are expressed heterologously. Nevertheless, although the
binding of glutamate activates the anion conductance, accumulation of
substrate is neither dependent on anions nor affected by the direction
of anion flux (Wadiche et al., 1995a). Furthermore, like many other
neurotransmitter transporters (Sonders and Amara, 1996), glutamate
transporters allow anions to traverse the membrane in the absence of
substrate (Bergles and Jahr, 1997; Wadiche and Kavanaugh, 1998). Once
bound by glutamate, the two pathways for charge transfer appear closely
coupled as both currents rise to a peak rapidly and decay to a
steady-state level in the continued presence of substrate (Bergles and
Jahr, 1997; Grewer et al., 2000; Otis and Kavanaugh, 2000). However,
the relative kinetics of these two currents vary between transporters
(Bergles and Jahr, 1997; Auger and Attwell, 2000; Grewer et al., 2000),
consistent with the differences in anion permeability (Wadiche et al.,
1995a) and unitary conductance (Larsson et al., 1996; Wadiche and
Kavanaugh, 1998) between transporters.
GLT-1 transporters are responsible for the majority of glutamate uptake
in the mammalian brain (Rothstein et al., 1996; Tanaka et al., 1997).
To determine the relationship between the movement of anions and
coupled charges through GLT-1 during glutamate transport, we measured
the kinetics of glutamate-induced anion and coupled transporter
currents in outside-out patches removed from human embryonic kidney
(HEK) cells that expressed a high density of GLT-1 transporters.
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MATERIALS AND METHODS |
Cell culture. GLT-1 transporters were expressed
heterologously in HEK 293 cells. We used a stably transfected inducible
cell line in which expression of GLT-1 was controlled by the ecdysone promoter (Dunlop et al., 1999). Cells were maintained in DMEM with 10%
fetal calf serum, 0.4 mg/ml G418 (Invitrogen, San Diego, CA), 0.4 mg/ml
Zeocin (Invitrogen), and 1% penicillin/streptomycin (Invitrogen) in
tissue culture flasks at 37°C with 5% CO2. To prepare cells for experiments, we plated them at a low density on glass coverslips coated with poly-L-lysine and collagen
(Invitrogen), allowed them to grow for 1 d, and then exposed them
to 10 µM ponasterone A (Invitrogen) for 18-24 hr before recording.
Outside-out patch recordings. Coverslips were transferred to
a Plexiglas chamber mounted on a Zeiss FS1 fixed stage upright microscope equipped with differential interference contrast optics. Cells were superfused with artificial CSF (ACSF) consisting of (in
mM): 160 NaCl, 2.5 KCl, 2.5 CaCl2,
1.3 MgCl, 10 HEPES, and 10 glucose, pH 7.2 (NaOH), 330 mOsm. Whole-cell
recordings were made from HEK cells with the use of established
techniques, using internal solutions of the following composition (in
mM): 160 KA
(where
A corresponds to the anions gluconate,
nitrate, or thiocyanate), 10 EGTA, 10 HEPES, and 1 MgCl2, pH 7.3. TEA (20 mM) was
substituted for K-gluconate for the paired pulse recovery experiments
shown in Figure 4 to reduce the noise associated with
K+ channel gating. Holding potentials have
been corrected for the different junction potentials of the internal
solutions. Patch electrodes had resistances of 1.5-3 M
when filled
with the internal solution. After the establishment of whole-cell
configuration the pipette was pulled away slowly from the cell,
resulting in the removal of a patch of membrane in the outside-out
configuration. Patches were brought in front of a multibarreled glass
flow pipe that had control and test solutions flowing out. The standard patch ACSF solution contained (in mM): 170 NaCl, 2.5 KCl,
2.5 CaCl2, 1.3 MgCl2, and
10 HEPES, pH 7.2 (NaOH), 330 mOsm. All solutions were made by using
HPLC grade water and ultrapure salts. Reagents (agonists, antagonists,
etc.) were added directly to this solution. Rapid exchange of the
solution at the patch was achieved via the use of a piezoelectric
bimorph (Tong and Jahr, 1994). Exchange times of <200 µsec
(10-90%) were measured at the end of each recording by recording the
junction current produced at the tip of the electrode by switching
between solutions of different ionic strength. These "open tip"
responses are displayed above each trace to indicate the duration of
agonist/antagonist application. All responses were recorded at room
temperature (22-24°C) at a membrane potential of 
90 mV, unless
otherwise noted. A range of glutamate concentrations was applied to
patches by connecting a miniature manifold (Warner Instruments, Grand
Haven, MI) to one barrel of the flow pipe.
Patch currents were recorded with an Axopatch 200A amplifier (Axon
Instruments, Foster City, CA), filtered at 5 kHz, and digitized at 50 kHz by using programs written in Axobasic (Axon Instruments). Analysis
was performed by using programs written in Axobasic or Origin
(Microcal, Northampton, MA) software. Data are represented as ± SEM, unless otherwise noted. Rise time was calculated from 20 to 80%
of the peak, and half-decay was measured from the peak to the
steady-state level. Significance was measured with a Student's t test, and significance reflects a p value of
<0.05. Sweeps are averages of 5-30 responses.
Modeling. A chemical-kinetic model of glutamate transport
was constructed in the Simulation Control Program environment
(Simulation Resources, Redlands, CA) to describe the behavior of GLT-1.
This cyclical model incorporates the discrete binding of 3 Na+, H+,
glutamate, and K+, as required by the
known stoichiometry of transport (Zerangue and Kavanaugh, 1996), anion
conducting states, and voltage-dependent transitions. The binding order
of Na+ and glutamate incorporates the
finding that at least one Na+ binds before
glutamate and at least one Na+ binds after
glutamate (Wadiche et al., 1995a; Watzke et al., 2001). We also place
H+ translocation with glutamate and
Na+ (Zerangue and Kavanaugh, 1996) as
opposed to with the K+ translocation step
(Auger and Attwell, 2000), because cysteine is transported as a neutral
zwitterion (A. Tzingounis, personal communication). We have chosen to
make the K+ translocation step
rate-limiting, based on the work of Kanner (Kanner and Sharon, 1978;
Kanner and Bendahan, 1982) and because of the dramatic slowing of the
cycle when Cs+ or
Na+ is substituted for internal
K+ (see Results). The rates of the
K+ translocation step were set by the
paired pulse recovery times. Model parameters were adjusted to provide
the best fit to a set of averaged transporter currents recorded under
each of the conditions described. All simulations include all of the
anion conducting states and, in addition, use the same rate constants
except where rate changes are described explicitly in the text.
Simulated transporter responses in the presence of permeant anions are
in units of open probabilities; in the absence of anions, transporter
responses are in units of amperes per single transporter.
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RESULTS |
GLT-1 expression in HEK cells
The charge movement initiated by the binding of glutamate to
transporters is highly dependent on the species of ions present. Because of the presence of an associated anion conductance, transporter currents elicited by glutamate are much larger in the presence of
chaotropic anions such as thiocyanate
(SCN) and nitrate
(NO3) (Fairman et al.,
1995; Wadiche et al., 1995a; Eliasof and Jahr, 1996; Bergles and Jahr,
1997). Our initial experiments were performed in the presence of
intracellular SCN to maximize the size
of the glutamate transporter current. L-Glutamate (10 mM) elicited large inward currents (peak amplitude,
206.2 ± 21.3 pA; n = 20) in patches from HEK
293 cells that were exposed to ponasterone A for >24 hr to induce the
expression of GLT-1 (Fig.
1A). In contrast,
L-glutamate (10 mM) did not
evoke currents in outside-out patches from cells that had not been
exposed to ponasterone A (n = 5), indicating that the
expression of endogenous glutamate transporters in this cell line was
below the limit of detection of this technique (Dunlop et al., 1999).
Glutamate transporter currents rose to a peak rapidly (20-80% rise
time, 248 ± 10 µsec) and then decayed to a quasi-steady-state
level in 1.7 ± 0.1 msec (half-decay time) in the continued
presence of L-glutamate. With the removal of
glutamate this current decayed to baseline with a biexponential decay
(
1 = 1.4 ± 0.11 msec, 74%; 2 = 21.9 ± 2.1 msec). The kinetics of these transporter currents were dependent on the
concentration of L-glutamate, with the rise and
decay kinetics slower with lower concentrations of
L-glutamate (Fig. 1A). In addition, the EC50 of
L-glutamate was much lower when measured at the
peak of the response (137.8 ± 11.8 µM;
n = 9) than when measured at steady state (12.4 ± 2.5 µM) (Fig. 1B). These
glutamate-evoked transporter currents closely resembled glutamate
transporter currents recorded in outside-out patches from astrocytes in
hippocampal slices (Bergles and Jahr, 1997, 1998) and from HEK cells
expressing EAAT2 transporters (Otis and Kavanaugh, 2000).
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Figure 1.
Characteristics of GLT-1 transporter currents in
outside-out patches. A, Response of an outside-out patch
removed from a HEK cell expressing GLT-1 to a range of
L-glutamate concentrations. Inset, The peak
responses to 0.01 and 0.1 mM L-glutamate have
been scaled to the peak of the response to 10 mM
L-glutamate to illustrate the concentration dependence of
the rise time of the transporter currents. B,
L-Glutamate dose-response relationship of the peak
(open circles) and the steady-state
(filled circles) amplitudes of GLT-1-mediated
transporter currents. Data were fit with the logistic equation;
KSCN-based internal solution.
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A model for transport by GLT-1
A chemical-kinetic model developed to account for the properties
of the EAAT3 transporter (A. Tzingounis, personal communication) was
adapted to simulate the kinetics of GLT-1-mediated transporter currents
(Fig. 2A). This model
is based on an alternating access scheme (Läuger, 1991;
Kavanaugh, 1998) and incorporates the dependence of glutamate transport
on Na+, K+,
and H+ as well as transmembrane voltage.
The permeability to anions, in physiological conditions, is accounted
for primarily by anion conducting states directly connected to three
states in the cycle: ToNa2H,
ToNa3GH, and
TiK (although very minor components of the conductance were from open states connected to the other
Na+-bound external states) (Fig.
2B). In the absence of substrate, glutamate
transporters exhibit a "leak" conductance to anions that is shut
off by antagonists such as kainate, dihydrokainate (Bergles and Jahr,
1997; Otis and Jahr, 1998), and
D,L-threo-
-benzyloxyaspartate (TBOA) (see below; Watzke et al., 2001). This standing leak conductance is accounted for by the open state connected to
ToNa2H (in addition to a
very small component added via
ToNa2). With the rapid
application of substrate (10 mM
L-glutamate) to the external face of the
transporter, a fast transient anion current is evoked, primarily from
the occupancy of the open state attached to
ToNa3GH. After several
milliseconds a steady-state current is reached that is a combination of
occupation of the open states attached to TiK and
ToNa3GH; the open state attached to ToNa2H, the
main leak conductance, is practically unoccupied during the presence of
substrate. At the end of the pulse of substrate the slow return to the
leak conductance level is the result of the relatively slow
deoccupation of TiK because of the rate-limiting,
potassium-dependent translocation step. Were it not for the
TiK open state, an apparent outward current, relative to the resting leak, is produced at the end of the substrate application. This outward current is actually the result of a decreased
occupancy of all anion conductance states, below that observed at rest.
Such a current is seen in patch recordings from hippocampal astrocytes
(Bergles and Jahr, 1997) and Purkinje cells (Otis and Jahr, 1998);
however, it was not observed in EAAT2 (Otis and Kavanaugh, 2000) or in
the current study of GLT-1. The lack of this "overshooting"
decrease in anion conductance requires a compensating open state that
precedes the rate-limiting step, which in the present model is served
by the TiK open state. As shown in Figure
2C, this model mimics the peak and steady-state dose-response curves of the anion currents through GLT-1 (Fig. 1).
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Figure 2.
Kinetic model of the GLT-1 transporter.
A, Illustration of the discrete states and transition
rates present in the model. Four transitions are voltage-dependent:
ToNa2GH to ToNa3GH
(z = 0.55), TiNa3GH to
TiNa2GH (z = 0.4, asymmetry of 0.1), TiK to ToK
(z = 0.59, asymmetry of 0.9), and To
to ToNa1 (z = 0.46).
Anion conducting states (data not shown) are attached to the following
states: ToNa1,
ToNa2, ToNa2G,
ToNa2H, ToNa2GH (all
with opening rates of 50/sec and closing rates of 4700/sec),
ToNa3GH (opening rate, 1500/sec; closing rate,
10,000/sec), TiNa2 (opening rate, 80/sec;
closing rate, 4700/sec), and TiK (opening rate, 55/sec;
closing rate, 4700/sec). Simulated anion conductances are the sum of
all of these states. The numbers in the columns
correspond to the rates for the transitions noted in the model.
B, Occupancies of the three states that are the main
contributors to the anion conductance in control conditions in response
to a 30 msec application of 10 mM L-glutamate.
A downward deflection indicates increased occupancy of the state.
C, Simulated dose-response of the anion conductance for
0.01-10 mM L-glutamate (sum of all anion
conducting states). Inset, Simulated peak responses to
0.01 and 0.1 mM glutamate have been scaled to the peak
response of 10 mM glutamate.
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Comparison of coupled versus uncoupled transporter currents
To determine the relationship between the anion current associated
with glutamate transport (Fairman et al., 1995; Wadiche et al., 1995a)
and the current produced by the movement of charges stoichiometrically
linked to transport, we recorded glutamate transporter currents in the
absence of permeant anions in the intracellular solution. Transporter
currents recorded with internal gluconate
(K+ salt), which does not permeate the
transporter-associated anion channel (Wadiche et al., 1995a; Eliasof
and Jahr, 1996; Wadiche and Kavanaugh, 1998), were smaller (peak
amplitude, 28.5 ± 2.0 pA; n = 16) and exhibited
faster kinetics (rise time, 138 ± 8 µsec; half-decay time,
860 ± 41 µsec) than currents recorded with the permeant anion
thiocyanate (Fig. 3A,B).
However, the shape of the responses was similar under the two
conditions, with the stoichiometric current rising to a peak, decaying
to a steady-state level in the continued presence of glutamate, and
returning to baseline with a biexponential decay with the removal of
glutamate. The ratio of the amplitude measured at steady state to that
measured at the peak was similar for both anion and stoichiometric
currents (SS/peak ratio: KSCN, 0.10 ± 0.01, n = 20; K-gluconate, 0.15 ± 0.01, n = 16).
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Figure 3.
Comparison of coupled and uncoupled responses
elicited by L-glutamate. A,
L-Glutamate-evoked (10 mM) transporter current
from an outside-out patch recorded in the absence of permeant anions
(K-gluconate-based internal solution). B,
L-Glutamate-evoked (10 mM) transporter current
from an outside-out patch recorded in the presence of permeant anions
(KSCN-based internal solution). C, Simulation of the
GLT-1 model in response to 10 mM L-glutamate in
the absence of permeant anions (plot of charge transfer vs time for a
single transporter). D, Simulation result of the GLT-1
model in response to 10 mM L-glutamate in the
presence of permeant anions.
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The stoichiometric current is accounted for in the model (Fig.
3C) by the net flux of transporters through the
voltage-dependent steps of Na+ binding to
two external states (To and
ToNa2GH),
Na+ unbinding from the cytoplasmic face
(TiNa3GH), and from the
return potassium-dependent translocation step
(TiK to ToK). The forward translocation step (ToNa3GH
to TiNa3GH) is modeled as a
voltage-independent transition and thus does not participate in the
stoichiometric current. The voltage dependence and the rationale for
these assumptions are addressed below.
The slower kinetics of the transporter currents recorded with internal
SCN may indicate that conformational
changes associated with charge transfer and anion channel gating were
slowed in the presence of this anion. To address this question, we
compared the time necessary for the peak amplitude of the response to
recover from the steady-state level (paired pulse recovery) in
recordings with gluconate as the primary internal anion (Fig.
4B) with recordings in
which SCN was the primary internal anion
(Fig. 4A). The time constant of recovery was not
significantly different for the two conditions (KSCN = 33.1 ± 0.55 msec,
n = 6; gluconate = 32.5 ± 0.7 msec, n = 10) (Fig. 4C), indicating
that internal thiocyanate does not slow down the cycling rate of GLT-1
significantly. The model faithfully reproduces the paired pulse
recovery times in both conditions (Fig.
5). Because the recovery times were
slower than those recorded in patches from astrocytes (Bergles and
Jahr, 1997) that express both GLT-1 and GLAST, we reassessed the
recovery times in the present study by using a
SCN-based internal solution more similar
to that used in the previous study (i.e., without TEA). In the absence
of TEA, recovery was speeded by ~35%
(KSCNTEA = 21.8 ± 0.8 msec;
n = 10) (Fig. 4C, open squares),
suggesting that TEA slowed the rate-limiting steps of the cycle. Such a
slowing was mimicked in the model by slowing the binding rate of
internal K+ from
106 to 104
per M/sec and increasing the opening rate from
TiK from 55 to 100/sec
(anion+TEA = 29.9 ± 0.4 msec;
stoichiometric = 29.0 ± 0.1 msec;
anionTEA = 17.6 ± 0.2 msec) (Fig. 5). These data suggest that TEA interferes with the binding of
K+ to the internal face of the
transporter, as has been reported for the
Na+/K+ pump
(Eckstein-Ludwig et al., 1998). With the exception of these experiments, TEA was not included in internal solutions used in this
study. The model predicts that the anion current
(SCN) has slower kinetics than the
stoichiometric current (gluconate) because the open state linked to
ToNa3GH is reached after
Na+ binds within the membrane field (to
ToNa2GH), and this open
state is revisited several times, on average, before glutamate unbinds. In the absence of internal TEA, Na+, or
glutamate, the model predicts that the cycling rate of GLT-1 is 44/sec,
whereas the paired pulse recovery requires 17.6 msec (a rate of
57/sec). The paired pulse recovery rate is faster because some of the
bound glutamate is not transported and released into the cytoplasm;
instead, glutamate unbinds to the outside, providing a second pathway
for recovery. In conditions that may mimic the astrocytic cytoplasm
environment more accurately (10 mM
Na+, 50 µM
glutamate inside; Chatton et al., 2000), the model predicts that the
cycling rate will slow to 37/sec (and the paired pulse recovery to
51/sec). Thus at physiological temperature, assuming a
Q10 of ~2-3 (Bergles and Jahr, 1998; Wadiche
and Kavanaugh, 1998), the cycling rate of GLT-1 transporters is likely
to be ~10-20 msec.
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Figure 4.
The paired pulse recovery rate of GLT-1
transporters is not affected by the presence of permeant anions.
A, Response of a patch to pairs of applications of 10 mM L-glutamate (KSCN-based internal solution).
The interval between control (30 msec duration) and test (20 msec
duration) applications was 1-120 msec. B, Response of a
patch to paired applications of 10 mM
L-glutamate recorded without permeant anions in the
internal solution (K-gluconate-based internal solution).
C, Plot of the ratio of the peak amplitude of the second
(P2) to the first (P1) response to paired
applications of 10 mM L-glutamate recorded with
(filled squares) and without anions (open
circles) in the internal solution (solid lines
are single exponential fits to the data). Removal of TEA-Cl (10 mM) from the internal solution (open
squares) speeded the decay by ~35% (dashed
line).
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Figure 5.
Simulations of the paired pulse recovery rate of
GLT-1 transporters. A, Simulated uncoupled (anion
conductance) responses of a patch to paired applications of 10 mM L-glutamate. The protocol is the same as in
Figure 4. B, Simulated coupled responses of a patch to
paired applications of 10 mM L-glutamate.
C, Plot of the ratio of the peak amplitude of the second
(P2) to the first (P1) response to paired
applications of 10 mM L-glutamate recorded with
(filled squares) and without anions (open
circles) in the internal solution (solid lines
are single exponential fits to the data). The dashed
line is a single exponential fit to the model adjusted to
reflect the removal of TEA-Cl (open squares; see
Results).
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Internal Cs+ slows counter-transport
Previous studies of transporter currents in isolated retinal
Müller cells demonstrated that Cs+
could substitute for K+ at the
counter-transport site to support glutamate transporter cycling;
however, transporter currents elicited with
Cs+ in the pipette were only two-thirds as
large as those elicited with K+ in the
pipette (Barbour and Attwell, 1991). To determine whether Cs+ also supports the cycling of GLT-1, we
made a complete substitution of K+ with
Cs+ in the internal solution and recorded
transporter currents in response to 10 mM
L-glutamate. Nitrate-based
(NO3) internal
solutions were used for these experiments because the Cs+ salt of thiocyanate is not soluble at
physiological pH. The peak amplitude of the GLT-1 transporter current
recorded with Cs+ was significantly
smaller than that recorded with K+
(CsNO3: 16.8 ± 2.6 pA, n = 8; KNO3: 49.2 ± 7.3 pA,
n = 6; p < 0.0001) (Fig.
6A,B). In addition, the
steady-state amplitude was reduced nearly to zero with internal
Cs+, suggesting that cycling occurs much
more slowly with this cation and that anion conducting states are
visited infrequently. In support of this conclusion the peak of the
transporter current recovered by only 47 ± 5% (n = 6) in 120 msec in Cs+ compared with
105 ± 3% (n = 8) with
K+. This is consistent with the effects of
Cs+ on the
Na+/K+ pump
in which Cs+ has a 20-fold lower affinity
than K+ (Omay and Schwarz, 1992).
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Figure 6.
Internal Cs+ slows the cycling
rate of GLT-1 transporters. A, Response of a patch to
paired applications of L-glutamate (10 mM)
(interval, 120 msec); KNO3-based internal solution.
B, Response of a patch to the paired application of
L-glutamate (10 mM) as in A but
recorded with a CsNO3-based internal solution.
C, Simulation of uncoupled conductance (anion current)
as in A. D, Simulation of uncoupled
conductance to reflect slower binding of Cs+ as in
B. E, Response of a patch to paired
applications of L-glutamate (10 mM) separated
by 120 msec recorded with a K-gluconate-based internal solution.
F, Response of a patch to paired applications of
L-glutamate (10 mM) separated by 120 msec
recorded with a Cs-gluconate-based internal solution. G,
Simulations of the coupled current in E.
H, Simulations of the coupled current adjusted to
reflect the slower binding of Cs+ as in
F.
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Similar experiments were performed with K-gluconate and Cs-gluconate to
determine whether the stoichiometric current exhibited a similar
behavior after the replacement of internal
K+ with Cs+.
L-Glutamate-evoked transporter currents were smaller with
Cs-gluconate (9.72 ± 2.15 pA; n = 6) than with
K-gluconate (28.5 ± 2.0 pA; n = 16), they
exhibited a smaller steady-state current, and they cycled more slowly
(paired pulse recovery at 120 msec: Cs-gluconate, 52 ± 4%,
n = 4; K-gluconate, 104 ± 2%, n = 4) (Fig. 6E,F), similar to the effects
observed on the anion-potentiated transporter currents.
The effects of internal Cs+ could be
mimicked in the model by using modifications similar to those used to
fit the data recorded with internal TEA. For internal
Cs+ the K+
binding rate to Ti was slowed to 8 × 103 per M/sec, the
translocation rate from TiK to
ToK was slowed from 40 to 8/sec, and the opening
rate from TiK was increased from 55 to 85/sec.
These modifications were sufficient to mimic the kinetic changes of
both anion and stoichiometric currents (Figs. 6C,D,
7C,D). The model predicts that
internal Cs+ will slow the cycling rate to
~6/sec, over sevenfold slower than with internal
K+. Because many physiological studies of
synaptic transmission use internal Cs+ to
block K+ channels and increase membrane
resistance, this action of internal Cs+ on
transport could decrease neuronal uptake and alter the strength of
synaptic excitation. One deficiency of the model is that it does not
predict the more than twofold decrease in the peak of the current seen
with internal Cs+. This discrepancy could
be accounted for if Cs+, in addition to
its effect on the rate of counter-transport, acted as a noncompetitive
antagonist, effectively decreasing the pool of available transporters
and further diminishing the capacity of uptake.
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Figure 7.
Replacement of internal K+ with
Na+ inhibits transporter cycling. A,
Response of a patch to the paired application of
L-glutamate (10 mM) separated by 120 msec,
recorded with a NaSCN-based internal solution. B,
Response of a different patch to the paired application of
L-glutamate (10 mM) recorded with a NaSCN-based
internal solution containing 10 mM L-glutamate.
C, Simulation of the response to the paired application
of L-glutamate (10 mM) separated by 120 msec
with a NaSCN-based internal solution. D, Simulation
response to the paired application of L-glutamate (10 mM) separated by 120 msec with a NaSCN-based internal
solution containing 10 mM L-glutamate.
E, Response of a patch to the paired application of
L-glutamate (10 mM) recorded without permeant
anions in the internal solution (Na-gluconate-based internal solution).
F, Response of a patch to the paired application of
L-glutamate (10 mM) recorded with a
Na-gluconate-based internal containing 10 mM
L-glutamate. G, Same as C but
without internal permeant anions (Na-gluconate-based internal
solution). H, Same as D but without
permeant anions (Na-gluconate-based internal solution and 10 mM L-glutamate).
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Internal Na+ blocks counter-transport
The dependence of counter-transport on intracellular
K+ (Kanner and Sharon, 1978; Kanner and
Bendahan, 1982) suggested that removal of internal
K+ would prevent transporter cycling.
Substitution of Na+ for internal
K+ therefore could isolate the charge
movements associated only with the
Na+-bound transitions. To investigate this
possibility, we replaced K+ with
Na+ in the pipette solution (NaSCN).
Transporter currents persisted under these conditions but had
significantly smaller amplitudes (110.2 ± 12.6 pA;
n = 5) than those recorded with internal
K+ (206.2 ± 21.3 pA;
n = 20). These currents rose to a peak rapidly (340 ± 16 µsec) but decayed more slowly to a steady-state level (Fig. 7A). Surprisingly, the steady-state current persisted
despite the removal of L-glutamate, suggesting
that in the absence of internal K+ the
transporter continued to visit a conducting state. Recovery of the
response was very slow, because the peak response recovered by only
16 ± 3% (n = 5) after 120 msec. Indeed, for
these experiments the repetition rate had to be increased from every 5 sec to every 15 sec to allow the transporter current to recovery fully
between pulses of glutamate. That the response does recover, albeit
slowly, indicates that there is a mechanism by which the transporters can become competent to bind glutamate again, even in the absence of
internal K+.
If intracellular concentrations of Na+ and
glutamate rise, the transporter is forced to operate in an "exchange
mode" (Kanner and Bendahan, 1982; Otis and Jahr, 1998), decreasing
the net influx of glutamate and Na+. When
10 mM L-glutamate was added to the NaSCN
intracellular solution, transporter currents elicited by external
glutamate had an amplitude (142.3 ± 17.0 pA; n = 4) and rise time (320 ± 38 µsec) similar to those recorded
with NaSCN alone (Fig. 7B). However, the amplitude of
the steady-state response was increased dramatically (SS/peak
ratio = 0.73 ± 0.09), as expected if transporters are forced
to enter a conducting state repeatedly. With the removal of
extracellular L-glutamate this current decayed
with a of 35.6 ± 4.3 msec (single exponential fit) compared
with the biexponential fit (1 = 1.36 ± 0.11 msec, 74%;
2 = 21.94 ± 2.1 msec) observed with
K+ inside. The slow decay of this current
may, in part, reflect the relatively slow unbinding of glutamate to the
outside (see below). However, this decay was strongly voltage-dependent
and became faster at more depolarized potentials (data not shown), indicating a voltage-dependent transition in the path to unbinding. The
ability of intracellular glutamate to force unbinding to the outside
allowed the peak of the transporter current to recover by 96 ± 4% in 120 msec (n = 4).
The behavior of the stoichiometric current was analyzed under similar
conditions. Application of glutamate to patches with Na-gluconate
inside elicited transient inward currents (Fig. 7E). As with
Cs-gluconate, these responses were significantly smaller than those
recorded with K+ inside (peak amplitude,
6.8 ± 2.5 pA; n = 5). In contrast to the
responses recorded with SCN, transporter
currents recorded under these conditions decayed to almost zero,
suggesting that internal Na+ does not
support cycling readily. Consistent with this hypothesis, the peak
amplitude recovered by only 20 ± 4% in 120 msec, similar to that
observed with NaSCN. The addition of 10 mM
glutamate to the Na-gluconate solution did not change the size of the
peak response to 10 mM external glutamate
significantly (peak amplitude, 21.2 ± 12.0 pA;
n = 3) (Fig. 7F); however, as seen
for NaSCN, this manipulation accelerated the recovery from steady state
such that the peak amplitude now recovered by 91 ± 7% at 120 msec.
The model reproduces the effects of substituting internal
K+ with Na+
(Fig. 7C,D). In the absence of internal glutamate the model
predicts that transporters will accumulate into internal
Na+-bound states. As transporters collect
inside, the anion current disappears unless a conducting state can be
visited from the inside. The sustained anion current evoked in these
conditions is reproduced well by the conducting state reached from
TiNa2, the first internal state reached in the cycle that is not bound by glutamate. The accumulation of transporters in internal states is reversible, because
~20% of the current does recover after 120 msec (Fig. 8). The model can mimic this slow
recovery in two ways, either by having a slow transition between
TiNa2 and
ToNa2 or by allowing internal Na+ to substitute for the
normally K+-driven translocation. Although
the present data do not distinguish between these two possibilities,
the latter possibility has been depicted in the model (Fig.
2A). The addition of internal glutamate forces the
transporter into an exchange mode, continually revisiting the
Na+ and glutamate-bound internal and
external states. Because the open state attached to
ToNa3GH is occupied
repeatedly in this condition, a large steady-state current is achieved.
When external glutamate is withdrawn, the relatively slow glutamate
unbinding rate to the outside forces the transporter to exchange a few
times before glutamate unbinds at the external face. This slow
relaxation is responsible for the relatively slow decay of the anion
current at the end of the glutamate pulse. The voltage dependence of
this decay is the result of the voltage dependence of the binding and unbinding of the third Na+, which precedes
external glutamate unbinding.
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Figure 8.
Voltage dependence of coupled and uncoupled
GLT-1-mediated transporter currents. A, Response of a
patch to 10 mM L-glutamate recorded at membrane
potentials between 100 and 60 mV (KSCN-based internal solution).
B, Response of a patch to 10 mM
L-glutamate recorded at membrane potentials between 110
and 50 mV (K-gluconate-based internal solution). C,
Plot of the current to voltage (I-V)
relationship for the peak amplitude recorded in response to 10 mM L-glutamate in the presence (KSCN,
open circles) or absence (K-gluconate,
filled squares) of permeant anions.
D, Simulation of uncoupled conductance to 10 mM L-glutamate recorded at membrane potentials
between 100 and 60 mV. E, Simulation of coupled
current to 10 mM L-glutamate recorded at
membrane potentials between 110 and 50 mV. F,
Simulated current to voltage (I-V) relationships
for the peak amplitude simulated responses in D
(uncoupled conductance, filled circles) and
E (coupled current, filled
squares).
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The stoichiometric currents recorded with Na-gluconate-based internal
solutions can be replicated by this model without further modifications
(Fig. 7G,H). In the absence of internal glutamate the
transporters accumulate in internal states, and no steady-state current
is seen, reflecting a lack of cycling through the
K+ counter-transport transition. In the
presence of internal glutamate, after an initial transient net influx,
again the steady state is nonexistent as cycling is reduced to zero
through the K+ counter-transport transition.
Voltage dependence of transport
The uptake of glutamate by GLT-1 is strongly voltage-dependent
(Wadiche et al., 1995b; Levy et al., 1998). We examined the voltage
dependence of GLT-1 transporter currents by applying 10 mM
L-glutamate to patches held at range of potentials between 100 and 50 mV. The current to voltage (I-V)
relationship of the peak of the response exhibited prominent inward
rectification, and transporter currents did not reverse at potentials
as positive as 50 mV (Fig. 8A,C). A similar
rectification of peak currents was observed with recordings in the
absence of permeant anions (K-gluconate internal solution) (Fig.
8B,C). The model accounts for the voltage dependence
of transport by assigning voltage dependence to the first and third
Na+ binding step to the external face of
the transporter, the unbinding step of the first
Na+ after translocation
(TiNa3GH to
TiNa2GH), and the
K+-dependent translocation (A. Tzingounis,
personal communication). In addition, the voltage dependencies of the
internal Na+ binding step and the
K+-dependent translocation are
asymmetrical. These asymmetries are required to mimic the kinetics of
the currents across the range of holding potentials that were tested.
In the previous figures anion currents are simulated as open
probabilities (Po). However, when
different holding potentials are compared, the
Po should be scaled according to the
electrochemical gradient and the voltage dependence of the anion
conductances. Unfortunately, we have not been able to measure the
voltage dependence of the anion conductances independently of
Po. We have measured the
I-V relationship of the substrate-independent anion leak
current, as described below. If we assume that the voltage dependence
of the conductance states activated by glutamate is the same as that of
the leak current, we can scale the simulated
Po traces by using the I-V
relationship obtained for the leak current. When this scaling is
performed, however, the simulated voltage dependence of
glutamate-evoked anion currents is far too great, suggesting that the
voltage dependence of the leak current is not the same as that for
glutamate-evoked conductances. The simulations (Fig. 8D,F), therefore, are presented as the
unaltered Po.
Pharmacological inhibition of transporter currents
EAAT2 transporters are inhibited by the nontransported antagonist
dihydrokainate (DHK) with a Ki of
between 24 and 79 µM (Arriza et al., 1994;
Shimamoto et al., 1998), an affinity 100-fold greater than that for
other glutamate transporters. In the presence of permeant anions
(SCN internal) DHK alone produced an
outward shift in the holding current (Fig.
9A), consistent with
observations made in patches from astrocytes (Bergles and Jahr, 1997)
and from HEK cells expressing EAAT2 (Otis and Kavanaugh, 2000). This
outward shift suggests that there is a leak of anions through the
transporters that is maintained in the absence of glutamate and blocked
by DHK. The voltage dependence of the DHK-induced responses was the
inverse of that observed for L-glutamate (Fig.
9B), as expected if these responses were mediated by an
inhibition of a resting anion current. With the removal of DHK the
current returned to baseline with a of 23.2 ± 2.8 msec
(n = 4; single exponential fit). This decay provides an
estimate for the unbinding rate of DHK, if it is assumed that the
conductance opens without delay after the unbinding of DHK. The
application of TBOA (300 µM), a nontransported
antagonist that has an ~10-fold higher affinity for EAAT2
(Ki = 5.7 µM;
Shimamoto et al., 1998), produced a similar maximal shift in the
holding current but decayed with a of 247.2 ± 20.4 msec
(n = 4; single exponential fit) with removal,
indicating that TBOA unbinds much more slowly than DHK from GLT-1
transporters (Fig. 9A). It was possible to model these
decays by setting the unbinding rate of DHK as 165/sec and the
unbinding rate of TBOA as 6/sec (Fig. 9C).
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Figure 9.
Block of anion leak current
by nontransported GLT-1 antagonists. A, The application
of dihydrokainate (DHK; 300 µM) or
D,L-threo--benzyloxyaspartate
(TBOA; 300 µM) elicited outward shifts in
the holding current in patches recorded with permeant anions in the
internal solution (KSCN-based internal solution). The response to the
same patch to 10 mM L-glutamate is shown by the
bottom trace. The values for the vertical scale bar are
20 pA (top traces) and 60 pA (bottom
trace). B, Voltage dependence of the peak
amplitudes of responses to either 300 µM DHK
(filled circles; error bars within
symbols) or 10 mM
L-glutamate (filled squares).
Inset, responses to DHK at potentials between 100 and
60 mV. Calibration: 10 msec, 20 pA. C, Simulations of
responses to DHK (binding rate, 3 × 106 per
M/sec; unbinding rate, 165/sec) and to TBOA (binding rate,
3 × 106 per M/sec; unbinding rate,
6/sec).
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We examined the kinetics of inhibition of GLT-1 by DHK by
measuring the effect of 300 µM DHK on the response to a
saturating dose of L-glutamate (10 mM). As
expected from the results above, DHK alone produced an outward shift in
the holding current. In addition, DHK inhibited the peak, but not the
steady-state current, in response to L-glutamate (Fig.
10A); presumably, 10 mM glutamate is sufficient to overwhelm this
competitive antagonist at steady state despite a high occupancy by DHK
in the absence of glutamate. In the absence of permeant anions
(K-gluconate internal) DHK (300 µM) also
inhibited the peak of the response to L-glutamate
(10 mM) (Fig. 10B). However, as
expected because of the absence of permeant anions and thus the leak
current, DHK produced no outward shift in the holding current.
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Figure 10.
Kinetics of inhibition of GLT-1 transporters by
DHK. A, Response of a patch to either 10 mM
L-glutamate alone (bottom trace, thin
line) or 10 mM L-glutamate plus 300 µM DHK (top trace, thick line); for the
latter response the patches were stepped from a solution with 300 µM DHK to one containing 10 mM
L-glutamate plus 300 µM DHK.
B, Responses recorded in the absence of permeant anions
in the internal solution (K-gluconate internal solution) as in
A. C, Simulations of uncoupled
conductance illustrating the effect of DHK (300 µM) on
the response of GLT-1 to 10 mM L-glutamate.
D, Simulations of coupled current illustrating the
effect of DHK.
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The major features of inhibition of both anion and stoichiometric
currents by DHK were reproduced by the model (Fig. 10C,D). The predicted microscopic affinity for DHK is 55 µM (on rate = 3 × 106 per M/sec; off
rate = 165/sec), a value in the range measured experimentally for
EAAT2 [Arriza et al. (1994), Ki = 23 µM; Shimamoto et al. (1998),
IC50 = 196 µM].
Agonist-dependent changes in kinetics
Different excitatory amino acids have been shown to elicit
transporter currents with dramatically different kinetics (Bergles and
Jahr, 1997; Wadiche and Kavanaugh, 1998), in part because of the
different affinities of the transporters for different agonists (Arriza
et al., 1994). In particular, transporter currents in outside-out
patches recorded from oocytes expressing EAAT1 transporters (Wadiche
and Kavanaugh, 1998), from astrocytes (Bergles and Jahr, 1997), and
from Bergmann glial cells (Bergles et al., 1997) exhibited much slower
kinetics in response to aspartate than to glutamate. Unexpectedly, the
kinetics of GLT-1 in response to L-glutamate and
D-aspartate were very similar (data not shown), and the
amplitudes of the steady-state currents were not significantly different (paired t test)
(L-glutamate, 19.4 ± 3.3 pA;
D-aspartate, 19.6 ± 3.7 pA;
n = 15). However, with permeant anions the peak amplitude of D-aspartate currents was 66 ± 1% (n = 15) of L-glutamate, resulting in a significantly larger SS/peak ratio
(D-aspartate, 0.22 ± 0.02;
L-glutamate, 0.12 ± 0.01; n = 15). In addition, D-aspartate-evoked currents
decayed more slowly at the end of the pulse. Coupled transporter
currents recorded with internal K-gluconate exhibited the same
features; the peak amplitude of responses to
D-aspartate was 79 ± 3% (n = 12) of the L-glutamate response (data not
shown), although the steady-state amplitudes were not significantly
different (paired t test)
(L-glutamate, 3.5 ± 0.5 pA;
D-aspartate, 3.5 ± 0.4 pA;
n = 12). These differences can be simulated, at least qualitatively, if the binding and unbinding rates of
D-aspartate are slowed relative to
L-glutamate, the forward translocation rate is
decreased by 35%, and the opening rate of the conductance from the
fully bound state (ToNa3GH)
is approximately halved. The slowed translocation rate is in line with
the observed lower flux rate of D-aspartate than
L-glutamate through EAAT2
(Imax D-aspartate/Imax
L-glutamate = 0.84; Arriza et al., 1994).
However, the lack of difference between steady-state stoichiometric
currents indicates that in these conditions the uptake of the two
substrates should be similar. Indeed, the model predicts that the
cycling rate achieved with an application of 10 mM D-aspartate is also 42/sec, indicating that for D-aspartate transport
the K+ counter-transport transition is
similarly rate-limiting.
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DISCUSSION |
The translocation of glutamate by the
Na+-dependent glutamate transporters is
associated with net movement of positive charge through the membrane
field. This unbalanced stoichiometry results from the movement of 3 Na+, 1 H+
into the cell with each negatively charged molecule of glutamate and
only 1 K+ out (Zerangue and Kavanaugh,
1996; Levy et al., 1998). In addition to this movement of cations
essential for glutamate translocation and transporter cycling,
glutamate transporters also allow anions to flow across the membrane
(Fairman et al., 1995; Wadiche et al., 1995a; Eliasof and Jahr, 1996).
In this study we used a heterologous expression system (Dunlop et al.,
1999) to create a high density of GLT-1 transporters in the plasma
membrane so that the kinetic properties of both the stoichiometric and
anion currents associated with glutamate uptake could be recorded in
outside-out patches. These studies revealed that the kinetics of the
anion currents are slower than stoichiometric currents but that anions
do not alter appreciably the recovery rate or the efficiency of GLT-1 transporters. We developed a chemical-kinetic model that mimics the
behavior of GLT-1, in which the anion conductance is associated with
multiple states in the cycle and the rate-limiting transition occurs
during the counter-transport of K+ (Kanner
and Sharon, 1978; Kanner and Bendahan, 1982).
Relationship between stoichiometric and anion currents
Although transporter currents recorded with and without permeant
anions had similar waveforms, the stoichiometric current rose faster
and decayed more rapidly from the peak than the anion current, as shown
previously for astrocyte transporters (Bergles and Jahr, 1997), for
EAAC1 (Grewer et al., 2000; Watzke et al., 2001), and for Purkinje cell
transporters (Auger and Attwell, 2000; Wadiche and Jahr, 2001).
However, the slower kinetics of the anion-potentiated transporter
currents were not translated into a slower overall paired pulse
recovery rate of GLT-1, suggesting that states in the transport cycle
are occupied for similar periods whether or not permeable anions are
present. The slower time course of the anion currents suggests that
conducting states are reached, and can be revisited, subsequent to the
movement of coupled charges through the membrane field. More dramatic
differences in the kinetics of anion and stoichiometric currents were
observed when internal K+ was substituted
with Na+. With permeant anions inside
(NaSCN-based internal solution), the current remained active long after
glutamate removal, whereas in the absence of permeant anions
(Na-gluconate-based internal solution) the stoichiometric current
decayed to zero. When Na+ is the only
internal cation, external glutamate causes transporters to accumulate
in Na+-bound internal states that conduct
anions. The lack of a steady-state component of the stoichiometric
current indicates that cycling occurs only very slowly in this
condition. Similarly, internal Cs+
substitution also dramatically slowed cycling, indicating that Cs+ is inefficient at supporting
counter-transport. It is possible that anions also slow transitions
between some states in the transport cycle, as suggested by Auger and
Attwell (2000) for Purkinje cell glutamate transporters. However, such
an effect would have to be small compared with the rate-limiting
K+ counter-transport step, because anions
affect neither paired pulse recovery (Fig. 4) nor glutamate flux in
steady-state conditions (Wadiche et al., 1995a).
Efficiency of transport by GLT-1
An important parameter of transport is the probability
that the binding of a molecule of substrate to the external face of the
transporter leads to release of a molecule of substrate into the
cytoplasm (i.e., transport) rather than back into the extracellular space. If substantial extracellular unbinding occurs in
physiological conditions, glutamate will have the opportunity to bind
to receptors long after release, although transporters still could
lower the concentration of glutamate in the extracellular space rapidly and slow diffusion through buffering (Rusakov and Kullmann, 1998; Diamond, 2001) (but see Barbour, 2001). We have estimated the efficiency of this transporter by simulating a very short (1 µsec) application of glutamate and measuring the net flux through the unbinding transitions to either the internal and external faces. The
ratio of unbinding to the inside to the sum of unbinding on both sides
provides a measure of the efficiency of the transporter. Under the
conditions used for the majority of the experiments in this study (0 glutamate, 0 Na+ internal), the predicted
efficiency of GLT-1 is 65%. This value is particularly sensitive to
the unbinding rate of glutamate to the outside, a rate that
unfortunately is not well defined. The dose-response relationship
(Fig. 1), for instance, can be simulated reasonably well with off rates
ranging from 250 to 1000/sec; however, the estimate of efficiency
changes from 79 to 48%, respectively, over this range. If we use an
unbinding rate of 500/sec, changing the internal composition to one
that may be more representative of astrocyte cytoplasm (10 mM Na+, 50 µM
glutamate) had little effect on efficiency (64%), whereas decreasing
the holding potential from 90 to 60 mV lowered the efficiency to
50%. Increasing internal glutamate to 10 mM further decreased the efficiency to 44%. Under these conditions efficiency becomes the probability that there will be a net flux of one glutamate molecule from the outside to the inside per cycle; when glutamate and
Na+ are present internally, the
translocation of one molecule of glutamate from outside to inside can
be followed by the reverse translocation of glutamate from the
cytoplasm to the extracellular fluid, thereby decreasing efficiency.
These results indicate that many transporters function as buffers only,
rather than as unidirectional transporters that force the net
accumulation of glutamate.
Comparison of GLT-1 to astroglial transporter currents
GLT-1 (EAAT2) is the predominant glutamate transporter in the
mammalian brain (Rothstein et al., 1994; Lehre et al., 1995) and is
essential for maintaining a low ambient level of glutamate (Rothstein
et al., 1996; Tanaka et al., 1997). This transporter is expressed at a
high density in astrocyte membranes (Lehre et al., 1995), a density
sufficient to allow electrogenic transporter currents to be resolved in
outside-out patches from these cells in acute brain slices (Bergles and
Jahr, 1997, 1998). GLT-1 transporter currents evoked by
L-glutamate exhibited many of the same characteristics as
those recorded in patches from astrocytes, including a rapid rise to a
peak, a decay to a steady-state level in the continued presence of
L-glutamate that was ~10% of the peak amplitude, and an
EC50 at steady state that was ~13
µM. This is in close agreement with the equilibrium
Km measurements made from GLT-1
expressed in Chinese hamster ovary cells (16.5 µM; Levy et al., 1998). In addition, both GLT-1
and astrocyte transporter currents were ~10-fold larger when recorded
in the presence of SCN in the internal
solution, consistent with the high permeability of these transporters
to chaotropic anions (Wadiche et al., 1995a). GLT-1 transporters also
exhibited a substrate-independent leak of anions that could be blocked
by the nontransported antagonists DHK and TBOA. However, GLT-1
transporter currents exhibited a slower rise time, a slower decay time
in the continued presence of L-glutamate, and a
prominent biexponential decay after the removal of
L-glutamate. This trend toward slower kinetics
differs from that observed for the EAAT2 transporter recorded in
patches from HEK cells, which had somewhat faster initial kinetics than those observed here (Otis and Kavanaugh, 2000). These differences are
somewhat surprising given the high sequence homology between EAAT2 and
GLT-1 (Kanner, 1993; Arriza et al., 1994). However, a similar
biexponential decay after the removal of glutamate was reported for
EAAT2 transporters (Otis and Kavanaugh, 2000).
A dramatic difference between GLT-1 and astrocyte
transporter currents was observed in the kinetics of the response to
D-aspartate (Fig. 7D) (Bergles and Jahr, 1997).
D-Aspartate-evoked transporter currents from
astrocytes exhibited a prominent steady-state component that was not
observed with GLT-1 (SS/peak ratio: GLT-1, 0.22 ± 0.02;
astrocyte, 0.52 ± 0.15). This difference may reflect the presence
of GLAST glutamate transporters that also are expressed in hippocampal
astrocytes (Rothstein et al., 1994; Lehre et al., 1995), because
transporter currents mediated by EAAT1, the human homolog of GLAST,
exhibit an enhanced steady-state current in response to
D-aspartate (Wadiche and Kavanaugh, 1998) and
transporter currents recorded from Bergmann glial cells, which express
a high density of GLAST transporters (Rothstein et al., 1994; Lehre et al., 1995), exhibit a large steady-state current in response to L-aspartate (Bergles et al., 1997). These
observations are consistent with the complete inhibition of the peak
amplitude of GLT-1 transporter currents by DHK (Fig. 10), an antagonist
with a >100-fold higher affinity for EAAT2 than for EAAT1 transporters
(Arriza et al., 1994), but only a partial inhibition of the peak
amplitude of astrocyte transporter currents [Bergles and Jahr (1997),
their Fig. 6E].
The ability to record both stoichiometric charge movement and anion
permeation mediated by GLT-1 has allowed for the creation of a kinetic
model of transport that reproduces both coupled and uncoupled currents
measured experimentally. The model has been used to estimate the
efficiency of glutamate transport and rates of cycling in a variety of
conditions those parameters of transport that have not been amenable to
direct experimental determination. These characteristics of transport
may be critical in determining the activation of glutamate receptors
both within and surrounding the synaptic cleft after exocytotic release
and during pathological conditions such as ischemia.
TOP
ABSTRACT
INTRODUCTION
