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The Journal of Neuroscience, August 15, 2001, 21(16):6058-6068
Activation of Metabotropic Glutamate Receptor 1 Accelerates NMDA
Receptor Trafficking
Jian-yu
Lan,
Vytenis A.
Skeberdis,
Teresa
Jover,
Xin
Zheng,
Michael V. L.
Bennett, and
R. Suzanne
Zukin
Department of Neuroscience, Albert Einstein College of Medicine,
Bronx, New York 10461
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ABSTRACT |
Regulation of neuronal NMDA receptors (NMDARs) by group I
metabotropic glutamate receptors (mGluRs) is known to play a critical role in synaptic transmission. The molecular mechanisms underlying mGluR1-mediated potentiation of NMDARs are as yet unclear. The present
study shows that in Xenopus oocytes expressing
recombinant receptors, activation of mGluR1 potentiates NMDA channel
activity by recruitment of new channels to the plasma membrane via
regulated exocytosis. Activation of mGluR1 induced (1) an increase
in channel number times channel open probability, with no change in
mean open time, unitary conductance, or reversal potential; (2) an increase in charge transfer in the presence of NMDA and the open channel blocker MK-801, indicating an increased number of
functional NMDARs in the cell membrane; and (3) increased NR1 surface
expression, as indicated by cell surface Western blots and
immunofluorescence. Botulinum neurotoxin A or expression of a dominant
negative mutant of synaptosomal associated protein of 25 kDa
molelcular mass (SNAP-25) greatly reduced mGluR1-mediated
potentiation, indicating that receptor trafficking occurs via a
SNAP-25-mediated form of soluble N-ethylmaleimide sensitive
fusion protein attachment protein receptor-dependent exocytosis.
Because group I mGluRs are localized to the perisynaptic region in
juxtaposition to synaptic NMDARs at glutamatergic synapses in the
hippocampus, mGluR-mediated insertion of NMDARs may play a role in
synaptic transmission and plasticity, including long-term potentiation.
Key words:
metabotropic glutamate receptors; NMDA receptors; receptor trafficking; channel gating; protein kinase C; (1S,3R)-1-amino-cyclopentane-1,3-dicarboxylic
acid; ACPD
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INTRODUCTION |
Activation of metabotropic glutamate
receptors (mGluRs) modulates NMDA receptor (NMDAR) activity and is
implicated in synaptic transmission and activity-dependent synaptic
plasticity (Pin and Duvoisin, 1995 ; Conn and Pin, 1997; Nicoletti et
al., 1999). mGluRs are encoded by a gene family of eight members;
further diversity is generated by alternative RNA splicing of the
receptor C-terminal domains. By sequence homology, agonist selectivity,
intracellular signaling mechanisms, and differential targeting in
neurons, mGluRs can be divided into three classes. Group I mGluRs
(mGluR1 and mGluR5) couple to diverse intracellular signaling
transduction pathways. Group I mGluRs can couple positively to
phospholipase C, activation of which leads to stimulation of protein
kinase C (PKC) and release of intracellular
Ca2+ (Conn and Pin, 1997), or to adenylyl
cyclase, activation of which stimulates cAMP formation (Aramori and
Nakanishi, 1992; Joly et al., 1995). Group I mGluRs couple negatively
to NMDARs via a G-protein-dependent, membrane-delimited signaling
pathway in cortical neurons (Yu et al., 1997) and depolarize CA3
neurons via a G-protein-independent, Src-dependent signaling cascade
(Heuss et al., 1999).
Immunocytochemical studies reveal strikingly different distributions of
group I mGluRs within the hippocampus. Although mGluR1 is localized
primarily to somatostatin-positive GABAergic interneurons, mGluR5 is
highly expressed in pyramidal neurons (Shigemoto et al., 1997). At
excitatory glutamatergic synapses, mGluR1 and mGluR5 are concentrated
in the perisynaptic zone at the periphery of postsynaptic densities
(PSDs; Baude et al., 1993; Lujan et al., 1996). Localization of group I
mGluRs in close proximity to NMDARs and AMPA receptors (AMPARs), which
are concentrated at PSDs, and to critical signaling proteins is
consistent with a modulatory role in glutamatergic transmission.
Activation of group I mGluRs induces potentiation of NMDA EPSCs in
hippocampal (Aniksztejn et al., 1992; Challiss et al., 1994), striatal
(Pisani et al., 1997), and subthalamic nucleus (Awad et al., 2000)
slices and potentiation of NMDA currents in lamprey motoneurons
(Krieger et al., 2000) and frog spinal cord neurons (Holohean et al.,
1999). However, the molecular mechanisms underlying potentiation of
NMDARs by group I mGluRs are as yet unclear. Mice with null mutations
of either mGluR1 (Aiba et al., 1994a,b; Conquet et al., 1994; Bordi,
1996) or mGluR5 (Lu et al., 1997) display reductions of hippocampal
long-term potentiation (LTP) and abnormalities of motor coordination
and associative learning. Metabotropic glutamate receptor signaling is
also implicated in neurodegenerative diseases (Conn and Pin, 1997;
Nicoletti et al., 1999), cortical development (Kaczmarek et al., 1997),
and addiction (Wolf, 1998).
Activation of PKC induces rapid delivery of NMDARs to the cell surface
of Xenopus oocytes and hippocampal neurons (Lan et al.,
2001). The present study was undertaken to examine the hypothesis that
potentiation of NMDARs by mGluR1 occurs at least in part by receptor
trafficking. We used Xenopus oocytes to express a homogeneous population of receptors of known subtype in a geometrically simple system. Moreover, the molecular machinery for protein
trafficking is highly conserved from yeast to mammals (Bennett and
Scheller, 1993). Experiments involving patch-clamp recording, charge
transfer measurements, cell surface Western blots, and
immunofluorescence reveal that activation of mGluR1 promotes
delivery of new NMDA channels to the plasma membrane by regulated
exocytosis, a mechanism that accounts entirely for potentiation of
NMDAR activity. These studies provide molecular insight into synaptic
function of group I mGluRs and define a novel mechanism for
potentiation of NMDAR function.
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MATERIALS AND METHODS |
Expression constructs. Rat NMDAR subunit
NR1-4b (NR1100; Zukin and Bennett, 1995)
cDNA was cloned in this laboratory (Durand et al., 1992); rat NR1-1a
(NR1011) and mGluR1 cDNAs were gifts from Dr.
S. Nakanishi (Kyoto University, Kyoto, Japan); mouse  1, 2, and
3 (corresponding to rat NR2A, NR2B, and NR2C, respectively) cDNAs
were gifts from Dr. M. Mishina (Niigata University, Tokyo, Japan).
cDNAs were subcloned into the pBluescript SK( ) vector (Stratagene, La
Jolla, CA) for oocyte expression. Synaptosomal associated protein of 25 kDa molecular mass (SNAP-25) and SNAP-25( 20) cDNAs were gifts
from Dr. R. Y. Tsien (Howard Hughes Medical Institute, University
of California, San Diego, CA). Capped mRNAs were synthesized as runoff
transcripts from linearized plasmid cDNAs with T3 or T7 polymerase
(mMessage mMachine transcription kit; Ambion, Austin, TX; 2 hr at
37°C). The concentration and integrity of mRNAs were assessed after
staining with ethidium bromide by direct comparison of sample mRNAs
with an RNA standard ladder (Life Technologies, Gaithersburg, MD).
Electrophysiology of recombinant NMDA receptors expressed in
Xenopus oocytes. Selected stage V and VI oocytes from
adult female Xenopus laevis (Xenopus I, Ann
Arbor, MI) were injected with a mixture of in vitro
transcribed mRNAs (20 ng of mRNA/cell; NR1:NR2:mGluR1 ratio, 1:2:1)
using a Nanoject injector (Drummond Scientific, Broomall, PA) as
described previously (Zheng et al., 1997). Oocytes were maintained at
18°C in culture buffer (in mM: 103 NaCl, 2.5 KCl, 2 MgCl2, 2 CaCl2, 5 HEPES, pH 7.5).
Whole-cell recording. Whole-cell currents were recorded from
oocytes (2-6 d after injection) at ambient temperature using a
GeneClamp 500 amplifier (Axon Instruments, Foster City, CA) by
two-microelectrode voltage clamp, filtered at 20 Hz, and digitized on-line at 100 Hz (Zheng et al., 1997). Currents were elicited by bath
application of NMDA (300 µM with 10 µM
glycine) at a holding potential of 60 mV. Oocytes were perfused in
Mg2+-free, normal frog Ringer's solution
consisting of (in mM): 116 NaCl, 2.0 KCl, 1.0 CaCl2, and 10 HEPES, pH 7.2 (Zheng et al., 1997).
Because Ca2+ influx through the NMDAR can
cause Ca2+ amplification (Zheng et al.,
1997), Ca2+ inactivation (Legendre et al.,
1993), and activation of chloride channels endogenous to the oocyte
(Leonard and Kelso, 1990) and thereby affect measurements of PKC
potentiation, all experiments except those illustrated in Figures 1-3
were performed in Ringer's solution in which
CaCl2 was replaced by
BaCl2. The pipette resistance ranged from 0.5 to
2 M with an internal solution consisting of (in mM):
3000 KCl and 10 HEPES, pH 7.2. For mGluR1 activation, oocytes were
incubated in
(1S,3R)-1-amino-cyclopentane-1,3-dicarboxylic acid (ACPD; 100 µM, 2 min; Sigma, St. Louis,
MO) and washed. ACPD potentiation is defined as the ratio of the
NMDA-elicited current measured after ACPD application to that measured
before ACPD application. For direct PKC activation, oocytes were
incubated in 100 nM
12-O-tetradecanoylphorbol 13-acetate (TPA; Sigma) for 10 min. TPA potentiation is defined as the ratio of the NMDA-elicited
current measured after TPA application to that measured before TPA application.
Single-channel recording. Single-channel currents were
recorded from outside-out patches excised from devitellinized oocytes (2-7 d after injection) as described previously (Araneda et al., 1999). Pipette resistance ranged from 7 to 12 M with an internal solution consisting of (in mM): 115 CsCl, 5.0 EGTA, and 10 HEPES, pH 7.2. External solutions were as described for whole-cell
recording to permit direct comparisons between NMDA-elicited whole-cell and single-channel currents. NMDA (100 µM with 10 µM glycine) solution was delivered to the patch from a
multibarrel array fed by gravity.
Single-channel current amplitudes were determined from means of
Gaussian fits to all-point amplitude histograms. The number of active
channels times channel open probability
(npo) was calculated as the total
channel open time divided by recording time. Open time durations were
calculated from single-channel openings above baseline but may include
events from more than one channel in the same patch. All data are
presented as mean ± SEM for 4-12 experiments performed with
different oocytes. Statistical significance was assessed by the
Student's t test (SigmaPlot 3.0) and ANOVA (CLR ANOVA
1.3).
Surface Western blot analysis. Control and ACPD-treated
Xenopus oocytes expressing mGluR1 and NR1-4b/NR2A
receptors were screened for NMDA currents in the range of 100-300 nA
and ACPD potentiation of ~2-3. Oocytes were incubated in external
recording solution in the absence or presence of ACPD (100 µM, 2 min) and washed twice, and surface
proteins were biotinylated with the membrane-impermeant reagent
sulfosuccinimidyl 2-(biotinamido) ethyl-1,3'-dithiopropionate
(sulfo-NHS-SS-biotin, 1.5 mg/ml, 30 min at 4°C; Pierce, Rockford, IL)
according to the method of Chen et al. (1999). Cell extracts were
prepared as described by Hollmann et al. (1994). To isolate
biotinylated surface proteins from nonsurface proteins, cell extracts
were incubated with Neutravidin-linked beads (Pierce) for 2 hr at
4°C, centrifuged, and washed. Bound proteins were eluted from beads
by incubation with SDS-PAGE gel loading buffer containing DTT (which
releases the proteins from the biotin moiety) and subjected to gel
electrophoresis together with aliquots of total protein.
Cell surface immunolabeling of oocytes. Xenopus
oocytes expressing NR1-4b/NR2A receptors with mGluR1 were labeled
with monoclonal antibody 54.1 directed to the extracellular loop of the
NR1 subunit (Gazzaley et al., 1996). Oocytes were screened for NMDA
currents in the range of 100-300 nA and ACPD potentiation of
approximately twofold to threefold. Oocytes were incubated in 20 ml of
external recording solution in the absence or presence of ACPD (100 µM, 2 min), devitellinized, and fixed in 4%
paraformaldehyde and 2% sucrose (1 hr at room temperature). Oocytes
were then incubated with NR1 antibody (10 µg/ml, 4°C overnight),
followed by biotinylated horse anti-mouse IgG (Vector Laboratories,
Burlingame, CA; 10 µg/ml, 1.5 hr at room temperature) and
FITC-conjugated avidin (Vector Laboratories; 10 µg/ml, 1.5 hr at room
temperature). Oocytes were rinsed, placed in a chamber 1.5 mm deep with
a coverslip forming the bottom (MatTek, Ashford, MA), and covered with
ProLong mounting medium (Molecular Probes, Eugene, OR) to reduce
fluorescence quenching. Cross-sectional and tangential images of
oocytes were viewed by a Bio-Rad MRC 600 Kr/Ar laser scanning confocal
microscope and acquired using COMOS software (Bio-Rad, Hercules, CA).
FITC fluorescence was measured (excitation wavelength, 488 nm;
emission, 515 nm) with a 40× objective lens (aperture, 1.3); laser
intensity, photomultiplier gain, and pinhole aperture were kept constant.
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RESULTS |
Activation of mGluR1 potentiates NMDA responses
To examine the interaction between group I mGluRs and NMDARs, we
coexpressed mGluR1 with NR1-4b/NR2A receptors in Xenopus oocytes and examined the effect of mGluR activation on NMDA responses by whole-cell recording under voltage clamp. NR1-4b/NR2A and -B have
the highest cell surface expression (Okabe et al., 1999) and highest
degree of PKC potentiation (Zheng et al., 1999). Application of the
mGluR agonist ACPD (100 µM) in
Ca2+-free Ringer's solution to the oocyte
elicited a large inward current (~750 nA) that decayed to near
baseline in <30 sec (Fig. 1A). The ACPD-elicited
current is ascribable to Cl efflux
through Ca2+-activated
Cl channels endogenous to the oocyte
(Saugstad et al., 1996). It is well documented that activation of
mGluR1 can lead to an increase of
[Ca2+]i by
activation of phospholipase C and release of
Ca2+ from inositol triphosphate
(IP3)-sensitive intracellular stores (Pin and
Duvoisin, 1995). Hence, the ACPD-evoked current indicates effective
expression of mGluR1 and targeting of functional receptors to the
cell surface of the oocyte.
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Figure 1.
Activation of mGluR1 potentiates NMDA
whole-cell currents; alternative splicing of the NR1 subunit has little
effect on ACPD potentiation. A, Typical sequence showing
NMDA-activated whole-cell currents recorded in Ca2+
Ringer's solution at Vh = 60 mV from
oocytes expressing mGluR1 and NR1-4b/NR2A receptors before and after
application of ACPD (100 µM, 2 min), followed by
application of TPA (100 nM, 10 min). NMDA, 300 µM; glycine, 10 µM. ACPD elicited a large
inward current that decayed rapidly to near baseline within 30 sec,
ascribable to Cl efflux through
Ca2+-activated Cl channels
endogenous to the oocyte. ACPD significantly potentiated NMDA
responses. Application of the phorbol ester TPA to oocytes after ACPD
treatment further potentiated NMDA responses. B,
Schematic representation of the NR1 splice variants. N1,
C1, C2, and C2' are alternatively
spliced cassettes; C0 is the region between the fourth
transmembrane domain and the first splice site in the C terminal (Zheng
et al., 1999). C, Summary of several experiments
illustrating that ACPD potentiation did not differ significantly for
NR1-4b/NR2A versus NR1-1a/NR2A receptors. In contrast, potentiation by
TPA (applied to oocytes after ACPD) was significantly greater for
NR1-4b/NR2A receptors compared with NR1-1a/NR2A receptors
(p < 0.01, one-way ANOVA followed by
Bonferroni's t test). Each data point is
from a single experiment; horizontal bars represent the
means.
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To examine the effect of mGluR1 activation on NMDAR responses, we
recorded whole-cell currents elicited by NMDA (300 µM
with 10 µM glycine) before and after bath application of
ACPD (100 µM, 2 min) to the oocyte. Treatment with ACPD
significantly increased NMDA-elicited responses (Fig.
1A; compare first and second NMDA responses). In
Ca2+ Ringer's solution, ACPD potentiation
(defined as the ratio of NMDA current amplitude measured after ACPD to
that measured before ACPD) was 2.97 ± 0.30 (n = 7) (Fig. 1C).
To investigate the effect of alternative splicing of the NR1 subunit on
mGluR1-mediated potentiation of NMDAR responses, we coexpressed
mGluR1 with either NR1-1a/NR2A (NR1011/NR2A)
or NR1-4b/NR2A (NR1100/NR2A) receptors in
Xenopus oocytes (Fig. 1B). NR1-1a/NR2A and
-B have the lowest cell surface expression (Okabe et al., 1999) and
lowest degree of PKC potentiation (Zheng et al., 1999). Application of
ACPD potentiated NMDA responses of NR1-4b/NR2A receptors to 2.97 ± 0.30 times the control response in Ca2+
Ringer's solution (n = 7) (Fig.
1A,C). ACPD potentiated responses of NR1-1a/NR2A
receptors to 2.03 ± 0.18 times the control response in
Ca2+ Ringer's solution (n = 7) (Fig. 1C). Differences in ACPD-induced potentiation
between splice variants did not reach statistical significance in
either Ca2+- containing or
Ca2+-free
(Ba2+) Ringer's solution (one-way ANOVA
followed by Bonferroni test, p > 0.05 for comparisons
between each receptor pair). The smaller magnitude of the mGluR1
potentiation may have obscured an effect of NR1 splicing similar to
that observed for PKC potentiation (Zheng et al., 1999).
To investigate the effects of the NR2 subunit on ACPD-induced
potentiation, we coexpressed mGluR1 with NR1-4b/NR2A, NR1-4b/NR2B, or NR1-4b/NR2C receptors in Xenopus oocytes. The mGluR1
agonist ACPD (100 µM) significantly potentiated NMDA-induced currents of NR1-4b/NR2A and NR1-4b/NR2B receptors (n = 3) (Fig.
2A,B). Potentiation was
to 3.33 ± 0.88 (n = 3) (Fig.
1A) and 2.57 ± 0.14 (n = 3)
times the control responses in Ca2+
Ringer's solution, respectively (Fig. 2D). The
degree of ACPD potentiation did not differ significantly for
NR1-4b/NR2A versus NR1-4b/NR2B receptors. As observed for TPA
potentiation (Mori et al., 1993), ACPD did not potentiate responses of
NR1-4b/NR2C receptors (n = 3) (Fig. 2C);
potentiation was to 1.02 ± 0.04 times the control response in
Ca2+ Ringer's solution (n = 3) (Fig. 2D). Similar results were observed for the
three receptor subtypes in Ba2+ Ringer's
solution. Thus, the NR2 subunit composition of NMDAR affected the
degree of ACPD potentiation.
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Figure 2.
The NR2 subunit alters ACPD potentiation.
NMDA-activated whole-cell currents were recorded in
Ca2+ or Ca2+-free Ringer's
solution from oocytes expressing mGluR1 and NMDARs.
A-C, ACPD (100 µM, 2 min) potentiated
NMDA currents in oocytes expressing NR1-4b/NR2A or NR1-4b/NR2B
receptors; no potentiation was observed for NR1-4b/NR2C receptors.
N1, C0, and C2' are defined in the
legend to Figure 1. D, Summary of three experiments
illustrating ~3.33-fold potentiation by ACPD of NR1-4b/NR2A and
2.57-fold for NR1-4b/NR2B receptors in Ca2+
Ringer's solution but little or no potentiation of NR1-4b/NR2C
receptors.
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To rule out the possibility that ACPD directly activates NMDA receptors
(the mGluR agonist 2-(2',3')-dicarboxycyclopropylglycine at
high concentrations may act on the NMDA receptor as an agonist) (Opitz
et al., 1994; Yu et al., 1997), we applied a high concentration of ACPD
(100 µM) to oocytes expressing NR1-4b/NR2A receptors in the absence of mGluR1. Under these conditions, ACPD elicited no
response and did not potentiate NMDA currents (data not illustrated), indicating that the Cl response was
mGluR-dependent and that ACPD did not activate or potentiate NMDA
channels directly.
ACPD increases npo but not conductance or
open time
To examine effects of mGluR1 activation on NMDA channel
conductance and gating, we recorded channel activity in outside-out patches excised from oocytes expressing mGluR1 and NR1-4b/NR2A receptors before and after ACPD treatment. We compared patches excised
before and after ACPD treatment, because patch formation inhibits
exocytosis, presumably by deformation of the membrane and disruption of
association with cytoplasmic elements (Lan et al., 2001)
In control patches, NMDA (10 µM) activated single
channels with a conductance of  = 44 ± 1 pS at 60 mV,
which did not vary with voltage (n = 5) (Fig.
3A,D). Occasional transitions
from the main conductance state to sublevels (total dwell time of <1% of that of the main state; data not shown) were excluded from the
analysis. NMDA channel activity in patches excised after application of
ACPD (100 µM, 2 min) was markedly potentiated
(Fig. 3A,B). ACPD increased
npo by approximately fourfold from
0.050 ± 0.006 before ACPD (n = 5) to 0.209 ± 0.027 after ACPD (n = 5; p < 0.001) (Fig. 3C).
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Figure 3.
ACPD potentiates npo.
A, B, NMDA-activated single channels recorded from
outside-out patches excised from oocytes expressing recombinant
NR1-4b/NR2A and mGluR1 receptors; patches were excised before
(A) and after (B)
application of ACPD (100 µM, 2 min). NMDA, 100 µM; glycine, 10 µM. The main unitary
conductance was 44 ± 1 pS at Vh = 60 mV. C, ACPD treatment potentiated the
npo from 0.050 ± 0.006 recorded from
patches excised before ACPD (n = 5) to 0.204 ± 0.034 recorded from different patches excised from the same oocytes
after ACPD treatment (n = 5; p < 0.001). D, Single-channel current-voltage
relationships. ACPD did not alter the main unitary conductance of
NMDA-activated channels (44 ± 1 pS before ACPD;
n = 5; vs 44 ± 1 pS after ACPD;
n = 5). ACPD treatment did not affect the reversal
potential (Erev = 0 mV before ACPD;
n = 5; and 0 mV after ACPD; n = 5). E, F, Histograms of mean open time durations for
NMDA-activated channels before and after application of ACPD to the
oocyte. Mean open times were determined by a single exponential fit to
the event lists generated by single opening and closing events. ACPD
did not alter the mean open time.
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mGluR1 activation did not alter NMDA single-channel conductance
(n = 5) (Fig. 3A,B) or reversal potential
(Erev ~ 0), as evidenced by
comparison of current-voltage relationships (n = 5)
(Fig. 3D). Moreover, ACPD did not significantly change the
mean duration of openings (Fig. 3E,F). The
distribution of open times was fit by a single exponential, consistent
with the presence of a single open state (Fig. 3E,F)
( = 6.35 ± 0.26 msec before ACPD; n = 5;
vs = 6.08 ± 0.29 msec after ACPD; n = 5). Potentiation of npo without change
in was also induced by the mGluR agonist (S)-3,5-dihydroxyphenylglycine (100 µM; data not shown).
mGluR1 activation induces an increase in the number of active
NMDA channels
The results reported thus far indicate that ACPD increases the
number of active channels in the cell membrane times the open probability. To distinguish between effects of mGluR1 activation on the
number of functional channels in the membrane, n, and
channel open probability, po, we
recorded NMDA whole-cell currents in the presence of MK-801 (5 µM) from control (Fig.
4B, left
trace) and ACPD-treated (Fig. 4B, right
trace) oocytes by a modification of the method of Jahr (1992) as
adapted by Rosenmund et al. (1995). This method takes advantage of the
essentially irreversible block of NMDA-elicited currents by the
open-channel blocker MK-801. To determine the number of channels in the
whole oocyte, N, we calculated the cumulative charge
transfer, Q, which is the integral of NMDA-elicited current
during the time required for complete block by MK-801. The number of
channels, N, can be calculated from Q as follows:
where tbl is the time constant
for MK-801 block (tbl = 1/kbl [MK-801], and
kbl = 2.5 × 107 M/sec; Jahr,
1992). From the single-channel recordings (Fig. 3), ACPD does not
change single-channel conductance, and we would not expected it to
change tbl. The channel number,
N, for control and ACPD-treated oocytes was normalized to
the NMDA-elicited whole-cell current, which corrects for any
differences in levels of expression within and between the two groups.
For control oocytes, which exhibited control currents in the range of
100-300 nA, the mean number of channels per 100 nA was 6.8 ± 0.8 × 105, and the mean channel
density for 100 nA was 0.02 µm2,
assuming a surface area for the oocyte of 3 × 107 µm2
(Zampighi et al., 1999). ACPD increased N ~1.6-fold
(NACPD/Ncontrol = 1.6; p < 0.01). This analysis indicates that mGluR1
activation increases the number of functional NMDA channels per
cell.
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Figure 4.
mGluR1 activation increases NMDA channel
number and modestly (but not significantly) increases open probability.
A, ACPD (100 µM, 2 min), bath-applied,
potentiated whole-cell currents elicited by application of NMDA (1 mM with 50 µM glycine). To avoid
contributions by Ca2+ inactivation,
Ca2+ amplification to NMDA responses, or both,
recording was in Ba2+ Ringer's solution as the
extracellular solution. Vh = 60 mV.
B, ACPD increased the number of functional NMDA channels
expressed at the cell surface (N). Currents were
elicited by application of NMDA (1 mM NMDA with 50 µM glycine) in the continuous presence of the open
channel blocker MK-801 (5 µM) from control
(left) and ACPD-treated (right) oocytes
at a holding potential of 60 mV. The NMDA inward current increased to
a peak value, after which it decayed exponentially as MK-801 entered
and blocked NMDA channels as they opened. The cumulative charge
transfer, Q, which is the total current flow during the
time interval for complete block by MK-801, was obtained by integration
of the current trace over time. The larger integrated current in
ACPD-treated oocytes indicated an increased number of functional
channels per cell. C, Agonist-evoked currents in
B were normalized to the same peak amplitude and used
for kinetic analysis. The more rapid decay of the NMDA current in
ACPD-treated oocytes versus control in this pair of oocytes indicates
an increased rate of channel opening, k ,
but the difference was not significant in the pooled data.
D-H, Quantization of data in A-C.
D, Potentiation of NMDA whole-cell current,
IACPD/Icontrol,
was 2.2 ± 0.2 times control (p < 0.001; n = 5). E, The increase in channel number, N, was significant
(p < 0.01), and
NACPD/Ncontrol = 1.6 (n = 5). F, The open
probabilities were not significantly different
(po, control = 0.13 ± 0.01; po, ACPD = 0.16 ± 0.01;
po, control/po,
ACPD = 1.2; n = 5). G,
Opening rates, k, for control (1.9 ± 0.2/sec) and ACPD-treated oocytes (2.1 ± 0.2/sec;
p < 0.01). H, Closing rates,
k , for control (12.7 ± 1.5/sec) and
ACPD-treated oocytes (11.1 ± 0.8/sec).
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The data in Figure 4B also enabled us to analyze the
effect of mGluR1 activation on NMDA channel gating. We first calculated channel open probability in control and ACPD-treated oocytes from the
NMDA-elicited whole cell current, I, the single-channel
current, i (which is not changed by mGluR1 activation) (Fig.
3D), and the number of channels per cell, N, as
follows:
from which po,
ACPD/po, control = 1.2. The ACPD-induced increase in channel open probability was not significant.
As an independent measure of the effect of mGluR1 activation on channel
gating, we analyzed the decay of NMDA-elicited current in the presence
of MK-801 from the data in Figure 4B normalized to
the same peak amplitude (Fig. 4C). Provided that
kbl [MK-801] 
k and
k (the closing and opening rates,
respectively), the decay can be described by a single exponential with
a rate constant, k, for channel
opening. Activation of mGluR1 did not significantly change
k (1.9 ± 0.2/sec for control
oocytes vs 2.1 ± 0.2/sec for ACPD-treated oocytes) (Fig.
4G). From measurements of
po and
k and the relation
po = k/(k + k), we calculated values of
k = 12.7 ± 1.5/sec in control
oocytes versus 11.1 ± 0.8/sec in ACPD-treated oocytes (Fig.
4H). These values of
k are substantially less than the
value of ~167/sec predicted from the ~6 msec mean open time of
single channels. The apparent discrepancy arises because the
po equation as applied to macroscopic
current assumes an equilibrium between unliganded and bursting states
and thus will yield a k value
corresponding to termination of bursting rather than of individual
openings. Moreover, k calculated in
this manner will be overestimated in that the probability that a
receptor is liganded is greater than the probability that it is open,
po, calculated from N
and I.
mGluR1 activation delivers NMDA channels to the cell membrane
via exocytosis
The results reported thus far indicate that mGluR1 activation
induces an increase in the number of active channels at the cell
surface but does not distinguish between insertion of new NMDA channels
and unmasking of silent channels. To distinguish between these
possibilities, we performed two additional experiments. First, we
examined the effects of loading oocytes with the light chains of type A
botulinum neurotoxin (BoNT A), which is known to inactivate SNAP-25 and
to prevent SNAP-25-dependent exocytosis (Montecucco and Schiavo, 1995).
Treatment of oocytes with BoNT A reduced the degree of mGluR1-mediated
potentiation of NMDA-elicited currents by ~50% (Fig.
5A-C). A mixture of type A,
B, and E BoNTs reduced ACPD potentiation by ~60% (Fig.
5C). No significant change in the resting potential, input
resistance, and basal NMDA response was caused by BoNTs (data not
shown). BoNT A also reduces TPA potentiation by ~50% (Lan et al.,
2001). No effects on mGluR1-mediated potentiation were observed when
only the vehicle (10 mM DTT) was injected.
Presumably, toxin treatment attenuated the number of new channels
inserted by mGluR1 activation.
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Figure 5.
mGluR1 promotes delivery of NMDA channels to
the cell membrane via exocytosis. A-C, Microinjection
of the light chain of botulinum toxin type A BoNT (50 ng), an enzyme
known to cleave SNAP-25, into oocytes 5 hr before recording reduced
ACPD potentiation of NMDA-elicited currents by ~50%. A,
B, Representative NMDA-elicited whole-cell currents in an
oocyte loaded with 50 nl of DTT (5 mM) in A
and 50 ng of BoNT A and DTT in B before and after ACPD
application. BoNT A reduced potentiation but not basal NMDA-elicited
currents or ACPD-elicited Cl currents.
C, Quantification of A and
B. A mixture of type A, B, and E BoNTs reduced ACPD
potentiation by 60%.
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Second, we examined the ability of a dominant negative mutant of
SNAP-25 to block mGluR1-mediated potentiation. SNAP-25(20), a
truncation mutant lacking the C-terminal 20 amino acids, corresponds to
yeast sec9-17, a dominant negative mutant of the yeast SNAP-25 homolog (Yao et al., 1999). Mutant and wild-type SNAP-25 were expressed
in oocytes by injection of their cRNAs 48 hr after injection of NMDAR
cRNAs, and ACPD potentiation was measured ~20 hr later. ACPD-induced
potentiation of NMDA currents in control oocytes (2.5 ± 0.3;
n = 3) (Fig.
6A,D) and in oocytes
expressing full-length SNAP-25 (2.9 ± 0.2; n = 4)
(Fig. 6B,D) was greater than in oocytes expressing
SNAP-25(20) (1.5 ± 0.2; n = 5;
p < 0.01) (Fig. 6C,D). Expression of
SNAP-25(20) did not affect the NMDA-elicited current or endogenous
Ca2+-activated
Cl currents. These findings indicate
that NMDA channels inserted at the cell surface by mGluR1 activation
are delivered, at least in part, by SNAP-25-mediated soluble
N-ethylmaleimide sensitive fusion protein attachment protein
receptor (SNARE)-dependent exocytosis.
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Figure 6.
A dominant negative mutant of SNAP-25 reduces ACPD
potentiation of NMDARs. A, ACPD potentiation of
NMDA-elicited currents in oocytes expressing mGluR1 and NR1-4b/NR2A
receptors as in Figure 2A. B,
Coexpression of wild-type SNAP-25 with NMDARs had no effect on ACPD
potentiation. C, Coexpression of SNAP-25(20), a
dominant negative mutant of SNAP-25, with NMDARs markedly reduced ACPD
potentiation. D, Quantification of the effects of
wild-type and mutant SNAP-25 on ACPD potentiation.
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Although the data indicate that ACPD increases the number of functional
NMDA channels at the cell surface via a SNARE-dependent mechanism, they
do not exclude a contribution from reduction in internalization (or
another mechanism of silencing). As an estimate of the rate of
constitutive exocytosis in the oocyte, we measured reappearance of
functional channels at the cell surface after quasi-irreversible block
by MK-801. MK-801 (1 µM) blocked completely the
NMDA-elicited current (Fig.
7A, first response). A test
application of NMDA at 3 min elicited a small response, indicative of a
small number of newly inserted channels, which were opened by the
agonist (Fig. 7A, second response). The small size of the
test response is indicative of a relatively slow rate of constitutive
exocytosis and is consistent with the half-time of ~40 min reported
for recovery of the acetylcholine response after irreversible block of
nicotinic acetylcholine receptors expressed in Xenopus
oocytes (Akabas et al., 1992). Because under steady-state conditions,
the rate of internalization equals the rate of exocytosis, the rate of
internalization is slow (Fig. 7A), and a reduction in the
rate of internalization (or of silencing) could account for, at most, a
very small fraction of the potentiation by ACPD.
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Figure 7.
mGluR1 potentiation increases the rate of
exocytosis of NMDA channels in the cell membrane. Whole-cell recordings
were obtained from Xenopus oocytes expressing
NR1-4b/NR2A receptors in Ca2+-free Ringer's
solution. The open channel blocker MK-801 (1 µM) was used
to estimate the rate of delivery of functional channels to the cell
surface in control and ACPD-treated oocytes. NMDA, 300 µM; glycine, 10 µM. A,
Application of MK-801 (1 µM) in the presence of agonist
completely blocked the NMDA response. Three minutes after block and
washout of NMDA, MK-801 was removed; then a test application of NMDA
elicited a very small response (~10% of control), attributable to
either recovery of a small number of channels from block or insertion
of new channels. B, After complete block of the NMDA
response by MK-801 (1 µM), ACPD (100 µM)
was applied for 2 min in the continuous presence of MK-801. After
washout of ACPD and MK-801, the peak of the NMDA-induced response was
slightly smaller than that of the control response. The greater decay
of this response is ascribable to residual MK-801. C,
Quantitation of data in A and B. Test
responses were normalized to initial currents. Bar 1,
IACPD/Icontrol,
where IACPD is the NMDA-elicited current
after ACPD. In this batch of oocytes. ACPD-induced potentiation was to
~2 times the control response. Bar 2,
IACPD,
MK-801/IMK-801, where
IACPD, MK-801 is the current after block of
the control response by MK-801 and potentiation by ACPD, and
IMK-801 is the current after block of the
control response by MK-801 followed by 3 min recovery. In the presence
of MK-801, ACPD potentiation was to 4.5 times the recovered response
observed after block by MK-801 in A, a value that could
not be accounted for by twofold potentiation of the recovered response.
Bar 3, IACPD,
MK-801/Icontrol. The potentiation
measured as the ratio of the potentiated response after MK-801 block to
the control response was somewhat smaller than the potentiated response
without MK-801 minus the control response.
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As a further test of ACPD-induced recruitment of new channels to the
cell surface, we blocked NMDA currents with MK-801 as in Figure
7A and then applied ACPD in the presence of MK-801 (Fig. 7B). In these oocytes, a test application of NMDA elicited a
much larger response, ~4.5 times that observed in control oocytes at 3 min after MK-801 block (Fig. 7A) (p < 0.01) and approximately the same size as the control response
(p < 0.001) (Fig. 7C). If ACPD
potentiation were attributable entirely to insertion of new channel
molecules (Fig. 4), and constitutive insertion were negligible, the
amplitude of the test NMDA current after ACPD application in the
presence of MK-801 (IACPD, MK-801)
would equal the amplitude of the test NMDA current after ACPD alone
(IACPD) minus the amplitude of the
initial control current (Icontrol).
IACPD, MK-801 was slightly smaller
than the predicted value, possibly an effect of residual MK-801 (Fig.
5D, bars 1, 3). These findings demonstrate that
ACPD causes a rapid increase in the number of functional NMDARs in the
cell surface.
ACPD increases surface expression of NMDARs
The electrophysiological experiments presented thus far argue for
mGluR1-mediated increase in the number of active channels via
exocytosis but do not provide a measure of the total number of channel
molecules expressed at the surface. To examine the effect of mGluR1
on the surface expression of NMDARs, we performed cell surface Western
blots by a modification of the method of Chen et al. (1999). Control
and ACPD-treated oocytes expressing mGluR1 and NR1/NR2A receptors
were surface-labeled with sulfo-NHS-SS-biotin, and surface proteins
were separated from intracellular proteins by reaction with Neutravidin
beads and centrifugation. Samples of surface and total cell protein
were treated with DTT (which releases the biotin moiety from surface
proteins) and subjected to electrophoresis. Membranes were probed with
antibody 54.1, a monoclonal antibody directed to the extracellular loop
of the NR1 subunit (Gazzaley et al., 1996) (Fig.
8A). Analysis of band densities indicated an increase in surface NR1 expression to 1.7 ± 0.2 times control (n = 4; p < 0.01)
(Fig. 8B,D), with no change in total cell NR1 protein
(Fig. 8C).
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Figure 8.
mGluR1 increases NMDAR surface expression. NR1
surface and total cell expression in control and ACPD-treated oocytes
is shown, as assessed by Western blot analysis of surface proteins
isolated by biotinylation. A, Representative Western
blot of surface protein from oocytes expressing NR1-4b/NR2A with
mGluR1 receptors probed with anti-NR1 antibody 54.1. 3, 6, 12, Micrograms of protein in samples of total cell extract
before Neutravidin bead extraction loaded on each lane;
surface, aliquot of Neutravidin bead-isolated receptors.
B-D, Quantitative analysis of the effects of ACPD on
surface expression (B), total cell protein
(C), and fractional surface expression
(D) of NMDARs. Surface expression was increased
to 1.7 ± 0.2 times control (n = 4;
p < 0.01). Total cell NR1 was not changed. The
proportion of NR1 expressed at the cell surface increased from 2.6 to
4.5%.
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In addition, we assessed the effect of mGluR1 activation on receptor
surface expression by immunofluorescence. Control and ACPD-treated
nonpermeabilized oocytes expressing NR1-4b/NR2A and mGluR1 receptors
were reacted with antibody 54.1, followed by a biotinylated secondary
antibody and fluorescein-conjugated avidin. Optical sections through
the oocyte were viewed by confocal microscopy and captured by COMOS
software. Control oocytes expressing NMDARs exhibited clear
immunofluorescence, which appeared concentrated at the external surface
(Fig. 9A,C). ACPD dramatically
increased NR1 surface immunolabeling (Fig. 9B,D). Little or
no immunofluorescence was observed for oocytes reacted with the
FITC-tagged secondary antibody in the absence of the primary antibody
(Fig. 9E) or for water-injected oocytes labeled with the
N-terminal antibody (Fig. 9F). These findings
indicate specificity of the immunofluorescence labeling. ACPD did not
increase the resting conductance of the oocytes after the
Ca2+-activated
Cl current subsided. Moreover, control
experiments indicate that TPA treatment does not increase the
permeability of the oocyte membrane to the antibody (Lan et al., 2001).
We therefore infer that ACPD is unlikely to affect the permeability of
the oocyte membrane.
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Figure 9.
mGluR1 increases NR1 surface
immunofluorescence. Oocytes expressing NR1100/NR2A
receptors were incubated in external recording solution in the presence
or absence of ACPD (100 µM, 10 min) and subjected to
immunocytochemistry. Immediately after incubation, intact oocytes were
devitellinized mechanically, transferred to glass coverslips, and fixed
with paraformaldehyde (4%) and sucrose (2%). To label surface NMDARs,
fixed oocytes were incubated with NR1 antibody 54.1, followed by
FITC-conjugated secondary antibody. NMDARs on the oocyte surface were
then visualized using confocal microscopy. Surface fluorescence was
expressed as the mean intensity of fluorescence per unit area. Shown
are representative oocytes expressing NR1-4b/NR2A receptors from
control (A, B) and ACPD (C, D) treatment
groups labeled by the extracellular epitope antibody. Control oocytes
labeled by a secondary antibody in the absence of primary antibody
(E) and water-injected oocytes labeled as in
A-D showed negligible fluorescence
(F).
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DISCUSSION |
Modulation of ligand-gated channels by G-protein-linked receptors
is likely to be an important mode of receptor regulation under
physiological conditions. The present study demonstrates that
mGluR1-mediated potentiation of NMDARs in Xenopus oocytes is mediated by recruitment of new channel molecules to the cell surface
via regulated exocytosis. We show that loading cells with BoNT A, which
interferes with exocytosis by cleaving SNAP-25, markedly reduces ACPD
potentiation of NMDAR responses. Moreover, expression of a dominant
negative mutant of SNAP-25, SNAP-25(20), nearly abolishes ACPD
potentiation. Specificity of mGluR1-stimulated insertion of NMDARs
at the cell surface is indicated by the observation that NR1/NR2A and
NR1/NR2B but not NR1/NR2C receptors exhibit ACPD-induced potentiation.
The present study is the first to examine biophysical mechanisms at the
single-channel level in a system in which activation of group I mGluRs
induces potentiation of NMDAR activity. We show that activation of
mGluR1 increases the number of functional NMDA channels at the cell
surface, with no change in single-channel conductance, mean open
duration, or open probability. Our study is also the first to
demonstrate mGluR-mediated rapid trafficking of NMDARs in oocytes via a
SNARE-dependent mechanism. A simple scenario is that activation of
mGluR1 regulates membrane fusion events of NMDAR-containing vesicles
by activation of endogenous protein kinases, which in turn
phosphorylate a protein involved in receptor trafficking. One potential
target is SNAP-25, which can be directly phosphorylated in an
activity-dependent manner (Shimazaki et al., 1996; Genoud et al.,
1999). The intracellular signaling pathway linking mGluR1 activation
with an increase in regulated exocytosis is, however, unknown, and
association of SNAP-25 or its binding partners with NMDARs remains to
be established.
Recent studies by our laboratory show that activation of PKC induces
rapid delivery of NMDARs to the cell surface of oocytes and hippocampal
neurons and an increase in the NMDA channel opening rate (Lan et al.,
2001). Several findings indicate that the molecular mechanisms
underlying mGluR1-mediated potentiation of NMDARs differ from those of
PKC potentiation. First, ACPD regulates receptor trafficking in the
absence of a significant effect on channel gating (although the present
data cannot exclude a proportionately smaller change in the channel
opening rate). Second, whereas the degree of TPA potentiation is
markedly affected by NR1 splicing (Zheng et al., 1999), the degree of
ACPD potentiation is affected only modestly by NR1 splicing, an effect
that was not statistically significant in the present study. Third,
whereas PKC-induced potentiation of NMDAR activity is observed in
oocytes and dissociated hippocampal neurons, ACPD potentiation is
observed in oocytes and in hippocampal slices but not in dissociated
neurons, suggesting that the intracellular signaling pathway linking
mGluR activation to NMDAR potentiation requires preservation of
neuronal circuitry (P. J. Conn, personal communication).
Finally, group I mGluRs couple to diverse intracellular
signaling pathways and potentiate NMDA EPSCs and NMDA-elicited currents by PKC-independent signal transduction in some neuronal cell types and
preparations and in expression systems. Electrophysiological studies of
CA3 pyramidal neurons from rat hippocampal slice cultures indicate that
synaptic activation of group I mGluRs (mGluR1) by mossy fiber
stimulation evokes an EPSC by a G-protein-independent signaling pathway
(Heuss et al., 1999). These studies show that an Src-family tyrosine
kinase is an integral component in a signaling pathway that
functionally links mGluR1 activation with a transient cationic
conductance increase. Because NR1/NR2A NMDARs are functionally modified
by tyrosine kinases such as Src (Wang and Salter, 1994), it is
reasonable to propose that mGluR1 potentiates NMDA currents via
activation of Src. Consistent with a role for tyrosine kinase-dependent modulation of NMDAR activity, we recently reported that insulin promotes rapid exocytosis of NMDARs in Xenopus oocytes
(Skeberdis et al., 2001a). Alternatively, group I mGluRs might signal
via a membrane-delimited signaling pathway, as has been observed for mGluR-mediated inhibition of NMDAR activity in embryonic mouse cortical
neurons (Yu et al., 1997).
The role of PKC-mediated signaling cascades in mGluR
potentiation of NMDARs is, at best, controversial. mGluR-mediated
potentiation of NMDA EPSCs is reported to occur independently of PKC
signaling in hippocampal (Harvey and Collingridge, 1993) and cerebellar slice preparations (Kinney and Slater, 1993), as is group I
mGluR-mediated potentiation of NMDA currents in lamprey motoneurons
(Krieger et al., 2000) and frog spinal cord neurons (Holohean et al.,
1999). On the other hand, the intracellular pathway linking mGluRs to potentiation of NMDA responses appears to involve PKC in at least two
preparations, although neither the mGluR nor NMDAR subtype or receptor
class was identified in these preparations. PKC inhibitors block at
least in part mGluR-mediated potentiation of NMDA EPSCs in hippocampal
slice (Aniksztejn et al., 1992) and potentiation of NMDA currents in
oocytes expressing rat brain mRNA (Kelso et al., 1992). Thus, multiple
pathways might contribute to the group I mGluR-dependent regulation of
NMDARs observed in the present study. This concept is supported by
pharmacological studies; whereas mGluR-dependent potentiation of NMDARs
is only partly blocked by selective PKC inhibitors, it is completely
blocked by the broad-spectrum kinase inhibitor staurosporine (Skeberdis
et al., 2001b).
mGluR potentiation is via regulated exocytosis
In the present study, ACPD potentiation of NMDA-elicited currents
was inhibited by ~50% injection of BoNT A and nearly completely abolished by expression of SNAP-25(20) (Figs. 5, 6). These findings provide important new evidence for a mechanism involving regulated exocytosis of NMDARs to the cell surface of the oocyte. Inhibition of
exocytosis by BoNT A is incomplete in a number of other systems, including exocytosis of Isoc channels
(Yao et al., 1999) and NMDARs in Xenopus oocytes (Lan et
al., 2001). The action of BoNT A on regulated exocytosis of NMDARs was
relatively specific in that (1) ACPD-induced insertion of new channel
molecules was not inhibited by BoNT B or E; and (2) constitutive
expression of NMDARs and endogenous
Ca2+-activated
Cl channels (Yao et al., 1999; our
unpublished observations) and transfected epithelial
Na+ channels (Yao et al., 1999) in oocytes
is not reduced by BoNT A.
Structural basis of localization of group I mGluRs and NMDARs
Immunogold studies reveal a highly ordered arrangement of group I
mGluRs and NMDARs at the postsynaptic membrane of CA1 synapses and
indicate that the two classes of receptors are concentrated in
microdomains in close proximity to one another (Baude et al., 1993;
Lujan et al., 1996; Lujan et al., 1997; Shigemoto et al., 1997).
Studies involving the yeast two-hybrid system and immunoprecipitation suggest that mGluR1 and mGluR5 may be cross-linked to NMDARs at
synapses. The C-terminal tails of mGluR1 and mGluR5 contain a
recognition sequence for binding by members of the Homer family of
anchoring proteins (Tu et al., 1998). Homer proteins, in turn, link via
Shank and PSD to each other and to IP3 receptors,
NMDARs, and AMPARs (Tu et al., 1999). Thus, a scaffolding structure
consisting of Homer and Shank may directly connect group I mGluRs to
NMDARs, thereby providing strategic positioning for regulation of
NMDARs by mGluRs at hippocampal synapses.
Physiological significance of mGluR1 potentiation of NMDARs
The physiological relevance of mGluR1 actions is underscored by
observations that activation of group I mGluRs induces potentiation of
NMDA EPSCs in rat hippocampal (Aniksztejn et al., 1992; Challiss et
al., 1994), striatal (Pisani et al., 1997), and subthalamic nucleus
(Awad et al., 2000) slices and potentiation of NMDA currents in lamprey
motoneurons (Krieger et al., 2000) and frog spinal cord neurons
(Holohean et al., 1999). Turnover and biotinylation studies of
cerebellar granule neurons indicate a large pool of NR1 subunits
(~60% total NR1) in the cytoplasm of dendritic shafts and spines,
where they are poised for rapid assembly with NR2 subunits and
insertion at synaptic sites (Huh and Wenthold, 1999). Rapid trafficking
of AMPARs between the plasma membrane and an intracellular compartment
in response to synaptic activity (Nishimune et al., 1998; Osten et al.,
1998; Song et al., 1998) and synaptic plasticity (Carroll et al., 1999)
is thought to represent recycling.
Findings from the present study suggest a mechanism whereby group I
mGluRs can augment the number of synaptic NMDARs, thereby modulating
neuronal excitability and lowering the threshold for LTP and long-term
depression. Moreover, activation of NMDARs potentiates mGluR5 activity
(Challiss et al., 1994; Luthi et al., 1994; Alagarsamy et al., 1999);
mechanisms implicated in this potentiation include a rise in
intracellular Ca2+, activation of
calcineurin, and inhibition of PKC (De Blasi et al., 2001). Thus,
positive interactions between mGluRs and NMDARs may be reciprocal in
nature and important to NMDAR-dependent LTP. The present study extends
earlier studies by providing insight into the biophysical and molecular
mechanisms underlying mGluR1-mediated regulation of NMDARs in a pure
population of receptors. Insertion and retrieval of NMDARs to and from
the plasma membrane are likely to be common and powerful mechanisms for
regulating excitatory synaptic transmission.
| |
FOOTNOTES |
Received May 21, 2001; revised May 21, 2001; accepted May 31, 2001.
This work was supported by National Institutes of Health Grants NS
20752 and NS 31282 (R.S.Z.) and NS 07512 (M.V.L.B). M.V.L.B is the
Sylvia and Robert S. Olnick Professor of Neuroscience. We thank Alice
P. Wang for technical support. We acknowledge the Analytical Imaging
Facility of the Albert Einstein College of Medicine (Michael Cammer, director).
Correspondence should be addressed to Dr. R. Suzanne Zukin, Department
of Neuroscience, Albert Einstein College of Medicine, 1300 Morris Park
Avenue, Bronx, NY 10461. E-mail: zukin{at}aecom.yu.edu.
| |
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ABSTRACT
INTRODUCTION
![N=Q/[&ggr;(<UP>V</UP>−E<SUB><UP>rev</UP></SUB>)t<SUB><UP>bl</UP></SUB>],](/content/vol21/issue16/fulltext/6058/img001.gif)
