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The Journal of Neuroscience, November 1, 2001, 21(21):8328-8338
Neuronal Glutamate Transporters Limit Activation of NMDA
Receptors by Neurotransmitter Spillover on CA1 Pyramidal Cells
Jeffrey S.
Diamond
Synaptic Physiology Unit, National Institute of Neurological
Disorders and Stroke, National Institutes of Health, Bethesda, Maryland
20892-4066
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ABSTRACT |
Glutamate released at synapses in the CA1 region of the hippocampus
escapes the synaptic cleft and activates extrasynaptic targets; it also
may "spill over" into neighboring synapses and activate receptors
there. Glutamate transporters in glial membranes restrict extrasynaptic
diffusion, but it is unclear whether neuronal glutamate transporters
also limit transmitter diffusion and receptor activation by spillover.
I examined the effects of a low-affinity competitive NMDA receptor
antagonist on EPSCs in acute hippocampal slices to distinguish
receptors activated within active synapses from those activated by
spillover. Glutamate spillover is observed between Schaffer collateral
fiber synapses onto CA1 pyramidal cells only when transporters in the
postsynaptic neuron are inhibited. Because glutamate transporters
operate most effectively at negative membrane potentials, these results
suggest that activation of NMDA receptors by spillover may depend on
postsynaptic activity.
Key words:
spillover; glutamate transporters; diffusion; Monte Carlo
simulation; hippocampus; NMDA receptor
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INTRODUCTION |
Communication at excitatory central
synapses is mediated by the neurotransmitter glutamate, which, after
the fusion of a synaptic vesicle with the presynaptic membrane,
diffuses across a narrow synaptic cleft and activates postsynaptic
receptors. At many central synapses, synaptically released glutamate
acts at sites beyond the immediately apposed postsynaptic density. This
phenomenon takes various forms but is commonly referred to as
"spillover." At mossy fiber and calyceal synapses, glutamate can
spill over between individual release sites and activate postsynaptic
receptors in different active zones within the same synaptic structure
(Trussell et al., 1993 ; Rossi et al., 1995; Silver et al., 1996;
Scanziani et al., 1997; Overstreet et al., 1999). In other cases,
glutamate spillover has been shown to activate extrasynaptic
metabotropic glutamate receptors (Scanziani et al., 1997; Schrader and
Tasker, 1997; Min et al., 1998; Vogt and Nicoll, 1999; Mitchell and
Silver, 2000; Semyanov and Kullmann, 2000) and glial transporters
(Bergles and Jahr, 1997; Bergles et al., 1997; Clark and Barbour,
1997).
In the present study, "spillover" refers to glutamate diffusion
between single-site synapses, like that reported to occur between
parallel fiber-stellate cell synapses in the cerebellum (Carter and
Regehr, 2000) and between Schaffer collateral fiber-pyramidal cell
synapses in the CA1 region of the hippocampus (Kullmann et al., 1996;
Asztely et al., 1997; Lozovaya et al., 1999). Spillover in CA1 was
examined here with D- -amino adipate (D-AA),
a low-affinity competitive NMDA receptor (NMDAR) antagonist
whose efficacy depends on the synaptic concentration of glutamate
(Clements et al., 1992). At room temperature and 34°C,
D-AA preferentially blocked slower components of the NMDAR
EPSC in pyramidal cells, suggesting that the NMDARs underlying those
components were activated by lower [Glu] than those contributing to
the peak of the EPSC. Reducing extrasynaptic glutamate transport
enhanced activation of those receptors encountering the lowest [Glu],
suggesting that they were activated by transmitter that had traversed
extrasynaptic territory. The effects of D-AA depended only
weakly on the strength of electrical stimulation, suggesting that
spillover may not require the coincident activation of multiple synapses.
In CA1, glutamate spillover is restricted by glial glutamate
transporters (Asztely et al., 1997). Glial transporters mediate the
large majority of glutamate uptake in this region (Rothstein et al.,
1996; Tanaka et al., 1997; Bergles and Jahr, 1998; Lehre and Danbolt,
1998; Kojima et al., 1999; Diamond and Jahr, 2000). CA1 pyramidal cells
also express glutamate transporters (Rothstein et al., 1994), but the
physiological purpose of neuronal transporters in the hippocampus,
including any role in limiting transmitter diffusion, remains unclear.
In the present study, spillover was observed only when neuronal
transporters were inhibited, either at room temperature or at 34°C.
These results indicate that neuronal glutamate transporters on CA1
hippocampal neurons help limit NMDAR activation by glutamate spillover.
Because glutamate transport is most efficient at negative membrane
potentials (Brew and Attwell, 1987; Barbour et al., 1991; Wadiche et
al., 1995), the degree of spillover between synapses may depend on the
activity of the postsynaptic cell.
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MATERIALS AND METHODS |
Slice preparation and solutions. Hippocampal slices
(400 µm) were prepared from 11- to 14-d-old Sprague Dawley rats, as
described previously (Sakmann and Stuart, 1995) and in accordance with
National Institute of Neurological Disorders and Stroke Animal Care and Use Committee guidelines. Slices were prepared and stored in artificial CSF (ACSF) containing (in mM):119 NaCl,
2.5 KCl, 1.3 MgCl2, 2.5 CaCl2, 1 NaH2PO4, 26.2 NaHCO3, and 11 glucose (bubbled with a 95%
O2-5% CO2 mixture). After
being cut in ice-cold ACSF on a vibratome (Leica, Nussloch, Germany),
slices were stored at 34°C for 30 min and at room temperature for up
to 7 hr thereafter. In experiments requiring
Mg2+-free conditions, slices were cut in
normal ACSF but were stored in solution in which
MgCl2 was replaced with
CaCl2; kynurenic acid (1 mM) was added to the
Mg2+-free storage solution to diminish
excitotoxicity. ACSF, equilibrated with the
O2-CO2 mixture, superfused
the recording chamber at a rate of 2 ml/min. Except where noted,
experiments were performed at room temperature (21-23°C).
Extracellular solutions contained picrotoxin (100 µM) and 6,7-dinitroquinoxaline-2,3-dione
(DNQX) (10 µM, except for the
experiments described in Fig. 3A) to block GABAA receptors
(GABAARs) and AMPA receptors
(AMPARs), respectively. In recordings of synaptic transporter currents
(see Figs. 3B, 4), 5 µM
(±)-3-(2-carboxypiperazin-4-yl)-propyl-1-phosphonic acid [(±)-CPP]
was also included to block NMDARs. The extracellular solution in
excised patch experiments contained 130 mM NaCl,
3.8 mM CaCl2, 10 mM HEPES, and 10 µM
glycine and was adjusted to pH 7.4 with NaOH. Internal pipette
solutions contained 120 mM
X+methanesulfonate, 10 mM EGTA, 0.2 mM NaGTP, 2 mM MgATP and 20 mM HEPES,
in which the cation X+ was
NMDG+ for the experiments described in
Figure 8D, Cs+ for all
other neuronal and excised patch recordings, and
K+ for astrocyte recordings. In the
experiments performed at 34°C in the absence of external
Mg2+, QX-314 (5 mM)
was included in the patch pipette to reduce excitability of the
postsynaptic cell. All internal and external solutions were adjusted
with sucrose, if necessary, to 290-300 mOsm. Reagents were obtained
from Sigma (St. Louis, MO), except for kynurenic acid and glutamate
(Tocris Cookson, Ballwin, MO) and
D,L-threo- -benzyloxyaspartate (TBOA), which
was a generous gift from Keiko Shimamoto (Suntory Institute for
Bioorganic Research, Osaka, Japan).
Electrophysiology. Slices were visualized on an upright
fixed-stage microscope (Zeiss, Thornwood, NY) equipped with
infrared-differential interference contrast optics. Recordings were
made with an Axopatch 1D amplifier (Axon Instruments, Foster City, CA),
and data were acquired (sample frequency, 5-10 kHz; filter frequency,
2-5 kHz) and analyzed with software written in Igor Pro (WaveMetrics
Inc., Lake Oswego, OR). Patch electrodes had tip resistances of 2-5 M [higher with
N-methyl-D-glucamine (NMDG)-based
internal solution]; access resistance was monitored throughout each
experiment, was typically <20 M (measured by the peak of the
charging current induced by a 1 mV step), and was not compensated.
EPSCs and synaptically activated transporter currents (STCs) were
elicited with a blunted bipolar stimulating electrode (115 µm
spacing) placed in stratum radiatum 100-200 µm from the soma of the
recorded cell. In excised patch experiments, rapid application of
glutamate was achieved with a multibarreled flow pipe attached to a
piezoelectric bimorph (Piezo Systems, Cambridge, MA). Twenty to 80%
solution exchange was typically 100 µsec (see Fig. 3C).
Voltages have not been corrected for a ~10 mV junction potential.
Moving the membrane potential from negative to positive potentials
resulted in a temporary reduction in input resistance and EPSC
amplitude, likely because of activation of voltage-dependent
conductances. The input resistance and EPSC amplitude increased
together and stabilized within several minutes (see Fig. 8). Unless
otherwise indicated, all data are reported as mean ± SD, and
p values are from paired t tests.
Simulations. NMDAR-D-AA kinetic
simulations (Fig. 1) were performed using
the Simulation Control Program (Simulation Resources Inc., Redlands,
CA); diffusion simulations (see Fig. 9) were performed using MCell
(Stiles et al., 1996), Monte Carlo simulation software designed
specifically to model chemical synaptic transmission. In both cases,
NMDARs were represented by a Markov model (Clements et al., 1992;
Lester and Jahr, 1992); neuronal transporters were configured by
modifying existing transporter models (Wadiche and Kavanaugh, 1998; Otis and Kavanaugh, 2000), such that
glutamate was removed from the simulation during transport and reverse
transport did not occur. The transporter Markov model comprised four
states, connected by the following rates (forward, backward; units are sec 1 or
M1
sec1):
Tout <>
GTout (2 × 107 · [Glu], 300);
GTout <>
GTin (500, 0);
GTin <>
Tin (2000, 0); and
Tin <>
Tout (40, 0). In an effort to
detect transporter-mediated currents in pyramidal cells, Bergles and
Jahr (1998) recorded synaptic responses with
SCN as the major internal anion to
maximize the conductance through the transporter-associated anion
conductance (Wadiche et al., 1995). Those experiments were simulated
here by convolving the time course over which transport events occurred
during the simulation with an exponential waveform (peak, 0.427 fA;  of 3 msec, i.e., 8 qe/cycle) to
reflect the deactivation of transporter currents in patches (Bergles
and Jahr, 1998) and the enhanced current with SCN (Watzke et al., 2001). To simulate
currents with an impermanent anion, the exponential waveform was scaled
down to reflect only the net inward flux resulting from the
electrogenic transport cycle (2 qe/cycle) (Zerangue and Kavanaugh,
1996).
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Figure 1.
Simulating the effect of D-AA on NMDAR
EPSCs. A, A kinetic model of the NMDAR (Clements et al.,
1992; Lester and Jahr, 1992), incorporating the binding and unbinding
of D-AA. Rates (M1
sec1 or sec1):
ka of 5 × 106;
k-a of 5; kb of
7 × 106; k-b of
210; of 91.6; of 46.5; kd of 8.4;
and k-d of 1.8. B, Model
prediction of receptor response to an exponentially decaying glutamate
concentration transient ([Glu]peak of 1 mM;
decay of 1 msec; top panel) in
"control" conditions (thin black line) and in 70 µM D-AA (thick black line).
The D-AA trace is also scaled (thick gray
line) to the same amplitude as control. C, As in
B, except the glutamate transient is 2 µM,
300 msec. D, Contour plot of the blockade of
D-AA of the simulated EPSC amplitude.
[Glu]peak values were 0.001, 0.003, 0.01, 0.03,0.1, 0.3, 1, 3, and 10 mM; decay values were 0.1, 0.3, 1, 3, 10, 30, 100, 300, and 1000 msec. All combinations were simulated,
resulting in a 9 × 9 array that was used to create the contour
map in Igor Pro. E, As in B, except for
[Glu]peak of 1 µM and decay
of 100 msec. F, As in B, but with the
glutamate transients from B and E
combined to drive the simulation. G, As in
B, except that the responses from B and
E were added in a 1:6 ratio to simulate a multisynapse
EPSC.
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To simulate direct release (see Fig. 9B), 5000 transmitter
molecules were released from a point source at the presynaptic membrane
into a 500 × 500 × 20 nm synaptic cleft. The diffusion coefficient (D) was set to 1 × 106
cm2/sec. The middle 225 × 225 nm
(0.05 µm2) square of the apposed 0.25 µm2 postsynaptic membrane was occupied
by NMDARs (500 µm2, i.e., ~25
receptors). The cleft glutamate concentration was measured in the cleft
space directly above this 0.05 µm2
square. When included, transporters occupied the remainder of the
postsynaptic membrane (5000 µm2,
~1000 transporters). To simulate indirect release (see Fig. 9A), glutamate was released from a distant point to create a
spatially homogeneous transmitter concentration; clearance at the
periphery of the simulation was adjusted such that the cleft glutamate
concentration peaked at ~1 µM and decayed
with an exponential time course of 100 msec (see Fig. 9A).
To simulate extrasynaptic activation of NMDARs and transporters (see
Fig. 9C), the synapse was replaced with a cube (200 nm on a
side) on which NMDARs (500 µm2) and
transporters (5000 µm2) were evenly
distributed; extrasynaptic glutamate concentration was calculated by
counting the number of glutamate molecules in the 10 nm layer of
space surrounding the cube. Glutamate release was the same as in the
indirect release simulation (see Fig. 9A). Control
simulations confirmed that the number of NMDARs (0-500 µm2) did not affect significantly the
glutamate concentration at the membrane (data not shown). The
whole-cell extrasynaptic transporter current (see Fig. 9C,
right panel) was derived by scaling the simulated
transport current measured across the 0.24 µm2 surface of the cube to reflect
transport over 2100 µm2, equivalent to
10% of the total membrane surface area within CA1 stratum radiatum of
a single pyramidal cell (Bannister and Larkman, 1995a,b).
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RESULTS |
If NMDARs at some inactive synapses were activated by glutamate
spillover, one would predict that these receptors would encounter a
lower [Glu] during a synaptic event than those located in an active
synapse. Directly activated receptors could then be distinguished from
spillover-activated receptors by examining the actions of a
low-affinity, competitive antagonist of the NMDAR. Some weak antagonists, such as D-AA, dissociate so rapidly that they
unbind, while synaptically released transmitter remains in the cleft, allowing glutamate to replace the antagonist at some fraction of
receptors (Clements et al., 1992). The extent to which this replacement
occurs (and therefore the extent of antagonist block) depends on the
amplitude and time course of [Glu]. This general approach has been
used to estimate the [Glu] time course in the synaptic cleft under
different experimental conditions (Clements et al., 1992; Tong and
Jahr, 1994; Diamond and Jahr, 1997; Choi et al., 2000).
One limitation of this method is that the antagonist effect, as
measured by the decrease in EPSC amplitude, does not indicate a unique
transmitter concentration time course. For example, a kinetic model of
the NMDAR (Clements et al., 1992; Lester and Jahr, 1992) (Fig.
1A) predicts that 70 µM
D-AA would block a relatively large, fast
transient ([Glu]peak, 1 mM; decay, 1 msec) (Fig. 1B) and a smaller, slower transient (2 µM, 300 msec) (Fig. 1C) equally
well. Systematically varying both the peak [Glu] and the time
constant of decay indicates that a particular degree of block by 70 µM D-AA could reflect any
in a range of [Glu] transients (Fig. 1D).
D-AA generally does not change the time course of
simulated NMDAR responses, except for a slight slowing of responses to
very slow [Glu] transients (Fig. 1C).
The model predicts that, if receptors in some synapses encounter much
less glutamate than receptors in other synapses, D-AA could
actually speed the NMDAR EPSC decay. For example, NMDARs encountering a
relatively large, fast transient (Clements et al., 1992) (Fig.
1B) would be blocked to a lesser extent by
D-AA than receptors activated by a smaller,
slower transient (1 µM, 100 msec) (Fig.
1E). The model also predicts that small transients, if permitted to decay gradually, would activate receptors quite slowly
and could give rise to a significantly slowed conductance (Fig.
1E). Therefore, D-AA, by
blocking slower components of the EPSC to a greater extent than faster
components, could speed the decay of an EPSC comprising directly and
indirectly activated synapses (Fig. 1G). Note that, when
fast and slow components are combined at a single synapse,
D-AA does not affect the simulated EPSC decay
(Fig. 1F). The two components must occur at separate synapses for D-AA to speed the EPSC decay (Fig.
1G). The effects of D-AA on the
simulated EPSC decay are abolished by eliminating spillover altogether
or by enlarging and/or prolonging the slow component so that the degree
of block is similar in the indirect and direct cases (Fig. 1, compare
B, C).
To test whether NMDARs at different synapses encounter a range of
[Glu] during a synaptic response, NMDAR EPSCs were recorded from CA1
pyramidal neurons in acute slices of rat hippocampus. Responses were
elicited by stimulating Schaffer collateral fibers in stratum radiatum
in the presence of DNQX (10 µM) and picrotoxin (100 µM), to block AMPARs and GABAARs,
respectively. EPSCs exhibited a J-shaped current-voltage relationship
that depended on the presence of external
Mg2+ and were blocked by the NMDAR
antagonist D-CPP (5 µM; data not shown). At
room temperature (22-24°C), when pyramidal cells were voltage
clamped at +50 mV to relieve the voltage-dependent
Mg2+ block of the channel, the NMDAR EPSC
decayed in a multiexponential manner, with a half-decay time
(t1/2) of 185 ± 34 msec
(mean ± SD; n = 28) (Fig.
2A).
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Figure 2.
D-AA speeds the decay of the NMDAR
EPSC. A, NMDAR EPSCs (Vh of
+50 mV) recorded under control conditions (i,
iii, v) and in the presence of 1 µM D-CPP (ii) or 70 µM D-AA (iv).
Traces represent averages of 20 consecutive responses.
Response vi recorded at Vh of
+10 mV. B, NMDAR EPSC amplitudes over the course of the
experiment in A. C, EPSCs
ii, iii, and iv from
A, scaled to the same amplitude. D,
Half-decay times of NMDAR EPSCs recorded in control solution, in the
presence of 70 µM D-AA, or in the presence of
1 µM D-CPP. n = 8.
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D-AA speeds the decay of the NMDAR EPSC
D-AA (70 µM) reversibly reduced the
NMDAR EPSC amplitude to 32 ± 11% of control (n = 8) (Fig. 2A,B) and also decreased
t1/2 to 56 ± 12% of control
(n = 8; p = 6 × 105). To control for possible changes in
space clamp attributable to the decrease in EPSC amplitude,
parallel experiments were performed with D-CPP, a
high-affinity antagonist that dissociates slowly (koff ~1
sec1) (Benveniste and Mayer, 1991) such
that its efficacy is independent of the glutamate concentration during
a synaptic event. Applied at a concentration of 1 µM, D-CPP blocked the
NMDAR EPSC to a similar extent (37 ± 12%; n = 8)
as 70 µM D-AA
(p = 0.1; n = 8). Each drug was
applied and then washed out (Fig. 2B), and the order of application was shuffled such that each drug was applied first in
half of the experiments. Although D-CPP did
decrease t1/2 slightly (to 85 ± 14%
of control; n = 8; p = 0.016) (Fig.
2D), the effect of D-CPP on
t1/2 was only one-third that of
D-AA (p = 0.0014; n = 8). The speeding of the NMDAR EPSC by
D-AA, interpreted in the context of the kinetic
model (Fig. 1), suggests that a wide range of peak [Glu] occurs
across the synapses contributing to an NMDAR EPSC.
D-AA does not affect release, glutamate transport, or
NMDAR kinetics
Possible actions of D-AA aside from NMDAR antagonism
could complicate the interpretation of the data in Figure 2. To test for presynaptic actions of D-AA, its effects on AMPAR EPSCs
were examined in the absence of AMPAR and NMDAR antagonists at
Vh of  60 mV, a potential at which
NMDARs are blocked by external Mg2+.
D-AA did not change the amplitude (102 ± 3% of control; n = 3) or paired-pulse facilitation
(94 ± 6% of control; n = 3) of AMPAR EPSCs (Fig.
3A), indicating that the drug
exerts no detectable presynaptic effects. Moreover,
D-AA did not affect STCs (Otis et al., 1997)
recorded in CA1 astrocytes (amplitude, 101 ± 5% of control;
n = 3) (Fig. 3B), responses that are acutely
sensitive to changes in release probability (Bergles and Jahr, 1997).
In addition to arguing against any presynaptic effects of
D-AA, this result demonstrates that the drug does
not interfere with glial glutamate transport.
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Figure 3.
D-AA does not affect release
probability, glutamate transport, or NMDAR kinetics. A,
AMPAR EPSCs (Vh of 70 mV; no AMPAR or
NMDAR antagonists present). Control trace (top
panel) reflects average of 25 EPSCs before application
of D-AA. Traces in control and D-AA are
overlaid in the right panel for comparison.
B, STCs recorded from astrocytes
(Vh of 95 mV; AMPAR and NMDAR antagonists
present). Responses in control solution (left
panel) and in D-AA (middle
panel) are overlaid for comparison (right
panel). C, NMDAR currents elicited in an
outside-out excised patch by a brief application of 1 mM
L-glutamate. Top panel shows "open-tip"
current, obtained at the end of the experiment after patch rupture to
check the speed of solution exchange across the patch pipette.
Vh of +60 mV.
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D-AA might also shorten the decay of the NMDAR EPSC by
speeding deactivation of the receptor, although previous work has shown this not to be the case (Clements et al., 1992). These results were
confirmed by eliciting NMDAR-mediated currents with glutamate in
outside-out patches excised from pyramidal cells (Fig. 3C). D-AA caused no significant change in the decay of
the response to a brief application of 1 mM
L-glutamate (114 ± 16% of control; n = 3) in patches voltage clamped at +50 mV. Similar
results were observed at 60 mV (94 ± 6% of control;
n = 3; data not shown).
Distinguishing between spillover and "whispering" synapses
The speeding of the NMDAR EPSC decay by D-AA suggests
that the NMDARs activated during an EPSC are exposed to a range of
transmitter concentration profiles. Some receptors appear to encounter
high glutamate concentrations and, as a result, are blocked relatively weakly by D-AA (Fig. 1B). Other receptors
encounter lower glutamate concentrations, and, as a result, their
activation is slower and more strongly blocked by
D-AA. One interpretation is that NMDARs at some
synapses encounter a lower concentration of glutamate because the
transmitter spills over from another, distant synapse. However, it is
also possible that different synapses (or even individual events at the
same synapse) could present widely different glutamate concentrations
to synaptic receptors as a result of variations in vesicular
transmitter content or the rate at which transmitter escapes the
vesicle during exocytosis (Choi et al., 2000; Fisher et al., 2001). One
way to distinguish between spillover and low-glutamate, so-called
"whispering" synapses would be to examine the extent to which
glutamate transport limits NMDAR activation. High-affinity,
sodium-dependent glutamate transporters restrict extrasynaptic
glutamate diffusion (Asztely et al., 1997; Bergles and Jahr, 1998).
Transport has a much smaller effect on [Glu] within an active synapse
(Diamond and Jahr, 1997), perhaps because most transporters are located
extrasynaptically in glial membranes (Lehre and Danbolt, 1998) and a
steep concentration gradient drives the rapid clearance of glutamate
from the cleft. Therefore, although transport ought to limit activation
of NMDARs by glutamate spillover (Asztely et al., 1997), it would be
unlikely to affect significantly the [Glu] time course at a
whispering synapse.
Glutamate transport was diminished with TBOA (Shimamoto et al., 1998),
a competitive transporter antagonist that does not interact with NMDARs
(Jabaudon et al., 1999). To gauge the degree of transporter blockade by
TBOA (and the consequent slowing of glutamate clearance), STCs were
recorded from astrocytes in CA1 stratum radiatum. TBOA reduced the
amplitude of the STC (and prolonged its decay) in a dose-dependent
manner (Fig. 4). TBOA at 30 µM caused a fourfold decrease in the
STC amplitude (to 27 ± 4% of control; n = 4) and
a fourfold increase in the exponential decay time constant (440 ± 60% of control; n = 4). TBOA at 30 µM exerted a greater blockade of the STC than
300 µM dihydrokainic acid (Bergles and
Jahr, 1997; Diamond and Jahr, 2000), which completely blocks GLT-1
(Arriza et al., 1994), the transporter subtype that constitutes 80% of
the glial transporter population (Lehre and Danbolt, 1998). Nonetheless, the remaining transporters cleared synaptically released glutamate within a few hundred milliseconds of its release (exponential of 78 ± 10 msec; n = 4). The notion that a
small fraction of the available transporters could effectively remove
synaptically released glutamate from the extracellular space was
supported by the observation that, in recordings from pyramidal cells,
the addition of 30 µM TBOA did not increase
background activation of NMDARs (no change in holding current at
Vh of +50 mV; n = 4; data not shown). Thus, this degree of transporter blockade caused little increase in basal glutamate levels.
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Figure 4.
Effects of TBOA on astrocyte STCs.
A, TBOA at 10 µM reversibly reduced the
amplitude and slowed the decay of the STC. B, Effects of
30 µM TBOA on the STC (different astrocyte than in
A). C, Effects of 10 µM
(n = 4) and 30 µM
(n = 4) TBOA on STC amplitude
(PTBOA/PCON)
and exponential decay
(TBOA/CON). Amplitudes were
calculated by subtracting the peak from the amplitude of the
slowly-decaying potassium current (Bergles and Jahr, 1997) measured
400-450 msec after stimulation.
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TBOA reduces effects of D-AA on late component of
NMDAR EPSCs
Next, the effects of D-AA on NMDAR EPSCs in pyramidal
cells were compared in control conditions and in the presence of TBOA (Fig. 5). Neurons were voltage clamped at
+50 mV, a potential at which any glutamate transport into the
postsynaptic cell would be greatly diminished (Wadiche et al., 1995).
Application of 30 µM TBOA caused small increases in EPSC
amplitude (117 ± 7% of control; n = 4;
p = 0.018) and decay time
(t1/2 of 147 ± 34% of control;
n = 4; p = 0.06) (Fig. 5B).
Applied in the presence of TBOA, 70 µM
D-AA reduced the EPSC peak amplitude to the same
extent as control (D-AA alone: amplitude, 25 ± 2% of control; D-AA in TBOA: amplitude,
29 ± 5% of control; n = 4; p = 0.19) (Fig. 5, compare A, C), but it had no
effect on the decay time (t1/2 in TBOA
plus D-AA of 106 ± 15% of
t1/2 in TBOA alone; n = 4;
p = 0.45) (Fig.
5C,F,G). Application of
10 µM TBOA caused an intermediate reduction of
the D-AA effect on the EPSC decay
(t1/2 in TBOA plus D-AA of 70 ± 11% of
t1/2 in TBOA alone; n = 6;
p = 0.05) (Fig. 5G). As with 30 µM TBOA, 10 µM TBOA did
not affect the reduction by D-AA of EPSC
amplitude (n = 6; p = 0.36). These
results indicate that blocking transporters preferentially enhanced
activation of the receptors, contributing to slower, more
D-AA-sensitive components of the EPSC, suggesting
that these receptors were activated by glutamate spillover. Of course,
neighboring synapses are located at a range of distances from an active
synapse; as a result, reducing transport is likely to affect each
indirectly activated synapse differently. However, 30 µM TBOA appeared to enhance [Glu] at synapses
mediating the slow components of the EPSC such that, on average, the
[Glu] at those synapses competed with D-AA, as well as the [Glu] mediating the fast components.
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Figure 5.
TBOA preferentially enhances a slow component of
the NMDAR EPSC. A, NMDAR EPSCs
(Vh of +50 mV) recorded in control
conditions and in the presence of 70 µM D-AA.
B, EPSCs recorded in control conditions and in the
presence of 30 µM TBOA. Recovery response obtained at the
end of the experiment (see D). C, EPSCs
recorded in the presence of 30 µM TBOA alone and in the
additional presence of 70 µM D-AA.
Calibration in C also applies to A and
B. D, NMDAR EPSC amplitude throughout the
entire experiment in which all illustrated responses were obtained.
E, Control and D-AA responses from
A, scaled to the same amplitude for comparison of decay.
F, TBOA and TBOA plus D-AA responses from
C, scaled. G, Effect of D-AA
on NMDAR EPSC decay under different conditions. TBOA at 10 µM (n = 6) and 30 µM
(n = 4) was applied in different experiments.
Control values were not different in the two data sets
(p = 0.63), so the control bar reflects the
pooled results (n = 10).
|
|
D-AA speeds NMDAR EPSC at
near-physiological temperature
The kinetics of NMDA receptors and glutamate transporters are
strongly temperature dependent (e.g.,
Q10 of transport, 3) (Wadiche and
Kavanaugh, 1998; Auger and Attwell, 2000), although diffusion is not
(Q10 of 1.3) (Hille, 1984). To test
whether receptor activation by spillover is present at more
physiological temperatures (Asztely et al., 1997), the experiments
described in Figure 2 were repeated at 34°C (Fig.
6A). At 34°C, the
NMDAR EPSC decayed more rapidly (t1/2 of
62 ± 13 msec; n = 6) than at room temperature
(p < 109;
unpaired t test). Nonetheless, D-AA
(70 µM) sped the decay of the EPSC
significantly (t1/2 of 70 ± 17% of
control; n = 6; p = 0.009), whereas
D-CPP did not (t1/2
of 95 ± 20% of control; n = 4; p = 0.66) (Fig. 6B), suggesting that NMDARs are
activated by spillover even at near-physiological temperatures.
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Figure 6.
D-AA speeds the NMDAR EPSC decay at
34°C. A, NMDAR EPSCs recorded in control conditions
and in the presence of 70 µM D-AA or 1 µM D-CPP. Traces have been scaled to the same
amplitude for comparison of decay. Vh of +50
mV. B, Effects in a group of experiments of
D-AA (n = 6) and D-CPP
(n = 4).
|
|
Spillover does not require transmitter pooling between
active synapses
Is glutamate released at a single synapse sufficient to activate
receptors in neighboring synapses, or does activation of receptors by
spillover require pooling of glutamate released from several active
synapses? At cerebellar parallel fiber-Purkinje cell synapses, the
extent of spillover depends on the spatiotemporal density of synaptic
activity, because it is enhanced by high-frequency and high-intensity
stimulation (Carter and Regehr, 2000). If this were true in CA1,
increasing stimulus strength should enhance spillover and decrease the
effect of D-AA on the NMDAR EPSC decay, similar to the
effect of blocking transporters with TBOA (Fig. 5). This was tested by
varying the stimulus intensity over a threefold range, which in control
solution caused proportional changes in EPSC amplitude and
insignificant changes in half-decay time (p = 0.30; n = 6; one-way ANOVA) (Fig.
7B). Although the effects of
D-AA on EPSC decay were not significantly
different at different stimulus intensities (p = 0.89; repeated-measures ANOVA; n = 6) (Fig.
7D), the trend in the data indicated a small reduction in the effects of D-AA on
t1/2 with stronger stimulation, suggesting that activation of more synapses may slightly exacerbate spillover. These effects of increasing stimulus intensity were, however, quite
subtle compared with those induced by blocking transporters (compare
Figs. 5G, 7D), consistent with glial cell
recordings that showed the decay of the STC to be slowed by competitive
transporter antagonists but unaffected by stimulus intensity (Diamond
and Jahr, 2000). These results do not demonstrate directly that
glutamate released from a single synapse is sufficient to activate
NMDARs in neighboring synapses, but the fact that
D-AA sped the decay of even the smallest NMDAR
EPSCs (Fig. 7D) supports the possibility that spillover does
not require activation of multiple synapses.
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Figure 7.
Effects of D-AA on NMDAR EPSCs
elicited by a range of stimulus intensities. A, NMDAR
EPSCs (Vh of +50 mV) were elicited by 75 (triangles), 150 (circles), and 225 (squares) µA stimulation. Stimuli were interleaved
throughout experiment. B, Effects of changing stimulus
intensity on EPSC amplitude (black bars) and half-decay
time (gray bars). Data pooled from six
experiments. C, Effects of D-AA on EPSC
amplitude at different stimulus intensities. D, Effects
of D-AA on EPSC half-decay time at different stimulus
intensities. Data in C and D were taken
from same six cells as B. Data from individual cells are
superimposed on bar graphs in C and
D.
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Postsynaptic transporters limit NMDAR activation by
glutamate spillover
All of the NMDAR EPSCs described thus far were recorded at +50 mV,
a potential at which glutamate transport into the postsynaptic neuron
is greatly diminished (Brew and Attwell, 1987; Barbour et al., 1991;
Wadiche et al., 1995). The physiological role of neuronal transporters
in the hippocampus is poorly understood. Hippocampal pyramidal cells
express glutamate transporters (Rothstein et al., 1994; He et al.,
2000), but the large majority of synaptically released glutamate in the
hippocampus is taken up by glial transporters (Rothstein et al., 1996;
Bergles and Jahr, 1998; Lehre and Danbolt, 1998; Kojima et al., 1999;
Diamond and Jahr, 2000) and STCs are not detected in pyramidal cells
(Bergles and Jahr, 1998). To test whether neuronal transporters limit
glutamate spillover, extracellular Mg2+
was replaced with Ca2+, allowing NMDAR
EPSCs to be recorded at both positive and negative potentials (Fig.
8A). NMDAR EPSCs
decayed significantly faster at negative potentials than at positive
potentials (t1/2,neg, 46 ± 15% of t1/2,pos;
n = 6; p = 0.0018).
D-AA reduced the EPSC amplitude at both
potentials, but it sped the EPSC decay only at positive potentials
(positive potentials: t1/2, 57 ± 20% of control; n = 6; p = 0.0049;
negative potentials: t1/2, 100 ± 10% of control; n = 6; p = 0.80)
(Fig. 8B,E).
D-CPP exerted little effect on the EPSC decay at
positive or negative potentials (n = 4) (Fig.
8C). Thus, no NMDAR activation by spillover was detected at
negative potentials. It appears unlikely that this result reflects any
voltage dependence of NMDAR affinity for either D-AA or glutamate. Competitive antagonists bind
and unbind the NMDAR in a voltage-independent manner (Benveniste and
Mayer, 1991), and the dose-response relationship of NMDARs to
glutamate in excised patches was not different at negative and positive
potentials (n = 4 patches; p = 0.2;
data not shown).
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Figure 8.
D-AA speeds NMDAR EPSC decay only when
postsynaptic transport is inhibited. A, NMDAR EPSCs were
recorded at 60 or +60 mV (as indicated in top
panel), and the effects of D-AA and
D-CPP were compared at both potentials. B,
EPSCs from the experiment shown in A, recorded in
control conditions and in the presence of 70 µM
D-AA. Responses in antagonist have been duplicated and
scaled (dashed lines) to control response for comparison
of decay time course. Inset, Comparison of control EPSCs
recorded at negative (i) and positive
(ii) potentials. C, as in
B, except recordings are in control and in the presence
of 1 µM D-CPP. D, Effects of
70 µM D-AA and 1 µM
D-CPP at Vh of 70 mV with NMDG
in the patch pipette. No synaptic current was observed when
Vh was +60 mV. Responses at 70 mV were
scaled to the control response for comparison of decay time course.
Control EPSC amplitude was 240 pA. E, Effects of
D-AA on NMDAR EPSC half-decay time at negative and positive
potentials with Cs+ in the recording pipette and at
negative potentials with NMDG in the pipette. Experiments were
performed at 22°C (black bars) or 34°C
(gray bars). Numbers in each
bar indicate number of cells tested.
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|
If the voltage dependence of NMDAR activation by spillover reflects a
role for postsynaptic glutamate transporters, then blocking transport
in the recorded cell should rescue the effect of D-AA at
negative potentials. Glutamate transporters bind intracellular potassium and transport it out of the cell to complete the transport cycle (Kanner and Bendahan, 1982; Barbour et al., 1991). Cesium can
replace potassium in the cycle (Barbour et al., 1991; Auger and
Attwell, 2000), but larger cations such as choline cannot (Barbour et
al., 1991). To block the complete cycling of neuronal transporters in
pyramidal cells, cesium was replaced in the intracellular patch
solution with NMDG. NMDAR EPSCs recorded at positive potentials were
abolished within 5-7 min after break-in with NMDG in the pipette (Fig.
8D), suggesting that NMDG diffused rapidly to
postsynaptic sites and failed to permeate NMDAR channels. At negative
potentials, the NMDAR EPSC decayed more slowly with NMDG in the pipette
(t1/2 of 147 ± 8 msec;
n = 5) than with cesium
(t1/2 of 95 ± 20 msec; n = 6; p < 0.001; unpaired
t test). Although within-cell comparisons were not made,
NMDG did not appear to affect EPSC amplitude at negative potentials.
With NMDG in the pipette, 70 µM
D-AA sped the decay of the NMDAR EPSC at negative
potentials (t1/2 of 55 ± 11% of
control; n = 5; p = 0.0011) (Fig.
8D,E), whereas in the same cells, 1 µM D-CPP had no
significant effect (t1/2 of 93 ± 18%
of control; n = 5; p = 0.48) (Fig.
8D). Analogous results were observed at 34°C;
D-AA sped the decay of the NMDAR EPSC at negative
potentials with NMDG as the internal cation (80 ± 11% of
control; n = 5; p = 0.03) but not with
Cs (96 ± 8% of control; n = 5; p = 0.30) (Fig. 8E). Together, these results indicate
that postsynaptic, neuronal transporters limit glutamate diffusion into
quiescent synapses and that the postsynaptic membrane potential may
influence the degree to which NMDARs are activated by glutamate spillover.
| |
DISCUSSION |
The results presented here argue that glutamate diffusion between
excitatory synapses is sufficient to activate NMDARs in quiescent
synapses in the CA1 region of the hippocampus. Glutamate spillover was
observed after single stimuli and did not increase substantially at
higher stimulus intensities, consistent with previous reports in CA1
showing that spillover is unaffected by changes in release probability
(Asztely et al., 1997). Although a role for transmitter pooling between
synapses cannot be excluded, evidence for spillover even in relatively
small synaptic responses (Fig. 7) suggests that it may occur after
glutamate release from a single synapse. In contrast, spillover between
parallel fiber-stellate cell synapses in the cerebellum requires
high-frequency stimulation (HFS) and is enhanced by stronger
stimulation (Carter and Regehr, 2000).
For glutamate spillover to be detected with D-AA (as in
Fig. 2), indirectly activated synapses must give rise to a
significantly slowed NMDAR conductance. It is possible, therefore, that
this method detects only low levels of spillover at distant synapses and is insensitive to larger, more transient [Glu] waveforms at nearest-neighbor synapses that produce NMDAR conductances similar in
time course to those at directly activated synapses (Rusakov and
Kullmann, 1998). If NMDARs activated by spillover contributed to the
EPSC peak, however, one would predict that reducing glutamate uptake
with TBOA would increase the EPSC amplitude and diminish the effect of
D-AA on the peak. Both 10 and 30 µM TBOA made
the EPSC slightly larger, but neither concentration changed the effect of D-AA on the amplitude (Fig. 5) (see Results). Although
this result is inconsistent with a significant contribution of
spillover to the peak of the EPSC, it is also likely that spillover to
those synapses closest to the release site is the least sensitive to transporter blockade.
A physiological role for neuronal transporters
One common characteristic of glutamate spillover in CA1 and in
other regions is that it is enhanced by blocking glutamate uptake (Otis
et al., 1996; Asztely et al., 1997; Lozovaya et al., 1999; Overstreet
et al., 1999; Carter and Regehr, 2000). Neuronal transporters in
Purkinje cells take up a significant fraction of synaptically released
glutamate (Otis et al., 1997; Auger and Attwell, 2000). In CA1, a role
for neuronal transporters is unclear, because the large majority of
uptake is accomplished by glia (Rothstein et al., 1996; Bergles and
Jahr, 1998; Lehre and Danbolt, 1998; Kojima et al., 1999; Diamond and
Jahr, 2000). Accordingly, reducing glial transport significantly
enhanced glutamate spillover (Figs. 4, 5). The data presented in Figure
8 demonstrate, however, that neuronal transporters also limit the
extent to which NMDARs in inactive synapses are activated by glutamate
spillover. Spillover was observed only at positive potentials, when
transport is diminished, or when transporters in the postsynaptic cell
were disrupted by NMDG applied through the patch pipette (Fig.
8D,E). Because membrane potential
does not change the affinity of the transporter for glutamate
(Mennerick et al., 1999), the effect observed here may reflect actual
transport into the postsynaptic cell.
At first, the role for neuronal transporters described here seems at
odds with evidence that pyramidal cells do not exhibit measurable
transporter currents either in response to synaptic stimulation or in
excised patches (Bergles and Jahr, 1998). However, a simple diffusion
model of an excitatory synapse (Fig. 9)
(see Materials and Methods) suggests that strategic subcellular
localization may allow a relatively small number of transporters to
play an important role. If, for example, transporters were expressed
perisynaptically on CA1 pyramidal cells, as indicated by
ultrastructural studies (He et al., 2000), they could restrict NMDAR
activation by low levels of glutamate diffusing into the synapse from
outside (Fig. 9A). The 1 µM, 100 msec transient used here and Figure 1 was chosen merely as an example,
although diffusion simulations do predict that [Glu] decays very
slowly once it falls well below the affinity of glial glutamate uptake
(kd ~13 µM)
(Bergles and Jahr, 1997; Rusakov and Kullmann, 1998). The actual
[Glu] waveform is likely to vary widely across indirectly activated
synapses. Perisynaptic transporters have relatively little impact on
receptor activation when glutamate is released from within the synapse,
because diffusion plays the dominant role in clearing glutamate rapidly
from the cleft (Clements et al., 1992) (Fig. 9B). In either
case, the resulting neuronal transporter current would be small, even
under conditions maximizing the contribution of the
transporter-associated anion conductance, i.e., with a highly permeable
internal anion such as SCN (Bergles and
Jahr, 1998; Wadiche and Kavanaugh, 1998; Watzke et al., 2001). Thus,
neuronal transport could play an important role in limiting glutamate
spillover while remaining difficult to detect with standard
electrophysiological methods.
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Figure 9.
Monte Carlo simulation of indirect, direct, and
extrasynaptic NMDAR activation. A diffusion model (see Materials and
Methods) was designed to simulate the effects of neuronal transporters
in different scenarios. A, Indirect activation.
Glutamate diffused into the synaptic cleft from a distant release site.
NMDARs (white ovals) were located in the center of the
postsynaptic density, and glutamate transporters (black
circles) were located perisynaptically. The number of glutamate
molecules released and rate of clearance from simulation were adjusted
to yield a homogeneous concentration that peaked at 1 µM
and decayed with an exponential time constant of 100 msec. Dense,
perisynaptic transporter expression (5000 µm2,
~1000 transporters total) reduced the amount of glutamate that
reached the NMDARs (second panel) and,
consequently, reduced NMDAR activation (third
panel). This reduction in [Glu]cleft
required the uptake of >600 glutamate molecules (fourth
panel). Traces reflect averages of 20 simulations.
B, Direct activation. Panels are the same as in
A, except that a quantum of glutamate (5000 molecules)
was released from within the synaptic cleft. Perisynaptic transporters
did not affect the glutamate concentration in the cleft or in NMDAR
activation. Traces reflect the average of 10 simulations.
C, Extrasynaptic activation, as in A and
B, except that NMDARs (500 µm2)
and transporters (5000 µm2) were distributed
randomly in the extrasynaptic membrane. Glutamate was released from a
distance, as in A. Whole-cell transporter currents
(right panel) were scaled to reflect transport
into the pyramidal cell during an evoked response if synaptically
released glutamate were to reach 10% of the pyramidal cell membrane in
stratum radiatum (see Materials and Methods). Traces reflect the
average of 10 simulations.
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Alternative explanations for the effects of D-AA
The effects of D-AA described above demonstrate that,
during an EPSC, NMDARs encounter a wide range of [Glu] waveforms. For the time course of the NMDAR conductance to be slower than that already
imposed by the slow kinetics of the channel (Lester et al., 1990), the
[Glu] waveform must peak at a much lower level and decay much more
slowly than thought to occur in an active synapse after exocytosis
(Clements et al., 1992) (Fig. 1). Peak [Glu] could vary significantly
within a single synapse, but large differences in decay time would be
unlikely to occur within even the largest synaptic contacts made
between Schaffer collateral fibers and CA1 pyramidal cells (~0.11
µm2) (Schikorski and Stevens, 1997). It
is probable that the [Glu] waveform in an active synapse comprises
fast and slow components, but such a combination does not predict an
effect of D-AA on the decay of the simulated EPSC (Fig.
1F). This result is not an artifact of nearly
complete NMDAR occupancy by the simulated fast component (Clements et
al., 1992) (Fig. 1B), because similar results are observed (data not shown) when the fast component achieves only 50%
NMDAR occupancy (Mainen et al., 1999).
NMDARs appear to be expressed primarily in postsynaptic densities
(Fritschy et al., 1998). Nonetheless, it is possible that the effects
observed here reflect activation of extrasynaptic NMDARs rather than
receptors located in neighboring synaptic contacts. Although it is
difficult to rule it out directly, this possibility seems unlikely.
Differences in NMDAR activation were observed by blocking the
transporters in a single pyramidal cell (Fig. 8), suggesting that
transporters on one neuron can reduce the glutamate concentration
encountered by NMDARs on the same cell. The diffusion model suggests,
however, that even a very large number of extrasynaptic transporters
would limit extrasynaptic NMDAR activation only slightly, and that this
uptake would result in a sizeable transporter current (Fig.
9C). This conflicts with synaptic recordings in pyramidal
cells, which show no detectable transporter-mediated component (Bergles
and Jahr, 1998). Moreover, blockade of GLT-1, the primary glial
glutamate transporter, does not decrease significantly the amount of
synaptically released glutamate transported by hippocampal astrocytes
(Diamond and Jahr, 2000), suggesting that the remaining glial
transporters (the GLAST subtype) are competent to take up transmitter
and that no other significant glutamate sink competes with glial
transporters. If neuronal transporters were expressed primarily
perisynaptically (He et al., 2000), even at very high density (e.g.,
5000 µm2) (Fig. 9), they would likely
be well outnumbered by GLAST on the surrounding glial membranes (Lehre
and Danbolt, 1998).
Implications for synaptic transmission
Glutamate spillover has been posed as an alternative explanation
(Asztely et al., 1997) for the experimental observation of NMDA-only,
"silent" synapses (Isaac et al., 1995; Liao et al., 1995). Indeed,
the dependence of the spillover observed here on the postsynaptic
membrane potential might explain, in part, why the quantal content of
NMDAR EPSCs is greater at positive potentials (Niu et al., 1998).
However, silent synapses are evident at negative holding potentials
(Liao et al., 1995) in conditions under which spillover was not
detected with the method used here, although spillover onto closely
neighboring synapses may elicit an NMDAR conductance that is not
significantly slower than at an active synapse, making it more
difficult to detect with D-AA.
The results presented here suggest that, when the postsynaptic cell
membrane remains relatively hyperpolarized during conditions of low or
moderate activity, neuronal transporters help limit NMDAR activation by
spillover. However, HFS, like that required to induce long-term
potentiation (LTP) (Bliss and Collingridge, 1993), could depolarize the
postsynaptic membrane sufficiently to decrease neuronal transport
(transport at 20 mV is only ~30% as efficient as at 70 mV)
(Wadiche et al., 1995) and encourage NMDAR activation by spillover. If
a number of synapses were activated even weakly by spillover during
HFS, collectively they could increase postsynaptic depolarization and,
consequently, calcium influx through NMDARs at both directly and
indirectly activated synapses. However, the low levels of NMDAR
activation at any one of the "indirect" synapses may not admit
sufficient calcium to induce LTP (Malenka, 1991); moderately activated
synapses may even undergo long-term depression (Cummings et al., 1996).
Thus, glutamate spillover could enhance the induction of LTP while
preserving, and possibly improving, its synapse specificity.
| |
FOOTNOTES |
Received June 26, 2001; revised Aug. 9, 2001; accepted Aug. 10, 2001.
This work was supported by the National Institute of Neurological
Disorders and Stroke Intramural Research Program. I thank Craig Jahr
for valuable discussions and Chris McBain, Tom Otis, and Josh Singer
for comments on this manuscript. I also thank Keiko Shimamoto (Suntory
Institute for Bioorganic Research) for the generous gift of TBOA.
Correspondence should be addressed to Dr. Jeffrey S. Diamond, National
Institutes of Health/National Institute of Neurological Disorders and
Stroke, Synaptic Physiology Unit, Building 36, Room 2C09, 36 Convent Drive, Bethesda, MD 20892-4066. E-mail:
diamondj{at}ninds.nih.gov.
| |
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TOP
ABSTRACT
INTRODUCTION
