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Volume 17, Number 19,
Issue of October 1, 1997
pp. 7351-7358
Copyright ©1997 Society for Neuroscience
Redistribution and Stabilization of Cell Surface Glutamate
Receptors during Synapse Formation
Andrew L. Mammen1, 2,
Richard L. Huganir1, 2, and
Richard J. O'Brien1, 2, 3
1 Howard Hughes Medical Institute, and Departments of
2 Neuroscience and 3 Neurology, Johns Hopkins
University School of Medicine, Baltimore, Maryland 21205
ABSTRACT
- INTRODUCTION
- MATERIALS AND METHODS
- RESULTS
- DISCUSSION
- FOOTNOTES
- REFERENCES
ABSTRACT 
Although the regulation of neurotransmitter receptors during
synaptogenesis has been studied extensively at the neuromuscular junction, little is known about the control of excitatory
neurotransmitter receptors during synapse formation in central neurons.
Using antibodies against extracellular N-terminal (N-GluR1) and
intracellular C-terminal (C-GluR1) domains of the AMPA receptor subunit
GluR1, combined with surface biotinylation and metabolic labeling
studies, we have characterized the redistribution and metabolic
stabilization of the AMPA receptor subunit GluR1 during synapse
formation in culture. Before synapse formation, GluR1 is distributed
widely, both on the surface and within the dendritic cytoplasm of these neurons. The diffuse cell surface pool of receptor appears to be mobile
within the membrane and can be induced to cluster by the addition of
N-GluR1 to live neurons. As cultures mature and synapses form, there is
a redistribution of surface GluR1 into clusters at excitatory synapses
where it appears to be immobilized. The change in the distribution of
GluR1 is accompanied by an increase in both the half-life of the
receptor and the percentage of the total pool of GluR1 that is present
on the cell surface. Blockade of postsynaptic AMPA and NMDA receptors
had no effect on the redistribution of GluR1. These results begin to
characterize the events regulating the distribution of AMPA receptors
and demonstrate similarities between synapse formation at the
neuromuscular junction and at excitatory synapses in cultured
neurons.
Key words:
GluR1;
synaptogenesis;
spinal cord;
metabolism;
glutamate
receptors;
rat;
tissue culture
INTRODUCTION
The processes involved in the
generation and maintenance of synapses are critical for neuronal
development and synaptic plasticity. To date, information regarding the
regulation of neurotransmitter receptors during synapse formation has
largely been derived from studies of the developing vertebrate
neuromuscular junction (Hall and Sanes, 1993
). On the surface of
embryonic myotubes before synapse formation, acetylcholine receptors
(AChRs) are distributed diffusely at a density of
102/µm2. Under the influence of
molecules secreted by the motor nerve terminal, AChRs become highly
enriched at sites of nerve-muscle contact (~104
AChRs/µm2) and depleted at extrajunctional areas
(Anderson and Cohen, 1977; Frank and Fischbach, 1979; Salpeter and
Harris, 1983; Salpeter and Marchaterre, 1988). The synaptic enrichment
of AChRs is attributable to the clustering of preexisting extrasynaptic
receptors and the directed insertion of newly synthesized receptors
(Role et al., 1985). In addition to the dramatic changes in receptor
localization that accompany synaptogenesis at the neuromuscular
junction, the metabolic half-life of receptors increases from 1 to 10 d
(Berg and Hall, 1975b; Reiness and Weinberg, 1981; Steinbach,
1981).
Ionotropic glutamate receptors (GluRs) mediate excitatory synaptic
transmission between neurons in the CNS. These receptors have been
cloned and divided into AMPA, kainate, and NMDA subclasses on the basis
of their electrophysiological and pharmacological properties (Seeburg,
1993; Hollman and Heinemann, 1994). Each subclass of ionotropic GluRs
includes several distinct subunits that exist as heteromeric complexes.
Iontophoretic mapping, immunocytochemistry, and immunoelectron
microscopy have demonstrated that these receptors are localized at
excitatory synapses (for review, see Ehlers et al., 1996). Little is
known, however, about the mechanisms regulating clustering and
maintenance of GluRs at central synapses.
We have previously characterized the development and subunit
composition of GluR clusters in cultured spinal neurons (O'Brien et
al., 1997). Over a period of 10 d, the AMPA receptor subunit GluR1, visualized with an antibody that recognizes the intracellular C
terminus, changes its distribution from diffuse to highly clustered at
sites of synaptic contact, similar to the pattern seen with GABAA (Killisch et al., 1991) and glycine receptors
(Bechade et al., 1996). Here, we use an extracellular antibody to GluR1
to study the distribution of surface receptors during synaptogenesis in
the same system. When applied to live neurons, the bivalent antibody
induces the aggregation of extrasynaptic receptors, rendering them
easily visualizable, while simultaneously staining preexisting synaptic
clusters. We demonstrate that a pool of mobile, extrasynaptic receptors
is expressed on the surface of immature dendrites, which diminishes as
GluR1 redistributes to synapses. Coincident with the redistribution of
GluR1 to synapses, there is a metabolic stabilization of AMPA
receptors, evidenced by an increase in their half-life, determined both
by surface biotinylation and metabolic labeling. In addition to the
surface receptors, a second pool of GluR1 exists within the cytoplasm
of dendrites before and after the period of maximal synaptogenesis. The
relative distribution of GluR1 between the surface and cytoplasmic
pools of receptors also changes with time, because of an increase in
the number of surface receptors.
MATERIALS AND METHODS
Generation of N-terminal GluR1 (N-GluR1). N-GluR1 was
made by injecting rabbits with the synthetic peptide
KQWRTSDSRDHTRVDWKRPK (Molnar et al., 1994) coupled to albumin. The
antisera was affinity-purified against the same peptide coupled to
thyroglobulin, using MgCl2 for elution. Fab fragments of
N-GluR1 were generated using papain according to the Pierce (Rockford,
IL) protocol (no. 44885). Each batch of Fab fragments was tested for
its ability to detect receptor clusters, using C-terminal GluR1
(C-GluR1) as a control.
Patching of surface GluRs. Cultures of embryonic day 19 to
postnatal day 3 rat spinal cords were prepared as described in O'Brien
et al. (1997). At the appropriate times in culture, the antibody
N-GluR1 was added to growth medium at a concentration of 2 µg/ml for
90 min. Cultures were then rinsed in four changes of HBSS and fixed and
processed for immunohistochemistry as described (O'Brien et al.,
1997). Fab fragments of N-GluR1 were used at a concentration of 10 µg/ml. In cases in which C-GluR1 and C-GluR2/3 antibodies were used
jointly with N-GluR1, the C-terminal antibodies were primarily labeled
with Cy3 (CyDye Kit, Amersham, Arlington Heights, IL).
Immunohistochemical control experiments consisted of incubating a 20×
concentrate of either the N- or C-GluR1 antibodies with appropriate
peptide (0.2 mg/ml) for 1 hr at room temperature. This solution was
then diluted 1:20 to its final concentration and added to the
coverslips as above.
Surface half-life experiments. Spinal cord cultures were
washed once with PBS/Ca2+/Mg2+
(10 mM phosphate buffer, 2.7 mM KCl, 137 mM NaCl, 1 mM CaCl2, 0.5 mM MgCl2, pH 7.4) at 37°C and then
cooled gradually to 4°C before they were washed twice with cold
PBS/Ca2+/Mg2+. Cultures were
incubated with 2 ml of biotinylation reagent (1 mg/ml NHS-SS-biotin in
PBS/Ca2+/Mg2+) while being shaken
gently at 4°C for 12 min, and then washed three times with cold
PBS/Ca2+/Mg2+ including 0.1%
BSA. Growth media was then added back, and the plates were returned to
the 5% CO2 incubator at 37°C. Cell extracts were
collected at regular intervals beginning 30 min after return of the
cultures to the incubator by washing the plates with PBS and scraping
the cells into 166 µl of warm precipitation buffer (PB; 10 mM NaPO4, pH 7.4, 5 mM EDTA,
5 mM EGTA, 100 mM NaCl, 1 mM
Na3VO4, 10 mM sodium
pyrophosphate, 50 mM NaF, and 10 µ/ml aprotinin) with 1%
SDS (PB/SDS) and 10 mM lysine. The resulting solution was
then diluted with 833 µl of cold PB with 1% Triton X-100 (PB/Triton)
and 10 mM lysine, sonicated, and microfuged for 20 min at
14,000 rpm. The supernatants, which include >95% of the total GluR1,
were frozen at 
80°C. To precipitate biotinylated proteins, samples
were thawed, mixed with 200 µl UltraLink immobilized streptavidin
beads (Pierce), and rotated for 2 hr at 4°C. The beads were washed
twice with PB/Triton, twice with PB/Triton including 600 mM
NaCl, and twice with PB alone. After the addition of SDS-PAGE sample
buffer to the beads, the resulting slurry was boiled for 3 min. The
biotinylated proteins were resolved by SDS-PAGE, transferred to
Immobilon-P (Millipore, Bedford MA), and probed with either the C-GluR1
or neuronal glutamate transporter antibodies. Proteins were visualized
with enhanced chemiluminescence (ECL) (Amersham). The volume of sample
loaded for each half-life experiment was adjusted such that all time
points would be within the linear range of the film. Serial dilutions
of the t = 0 sample were run on each gel to provide a
standard curve for each experiment.
Surface expression experiments. Cell extracts were prepared
from spinal cord cultures immediately after the biotinylation reaction
described above by washing the plates with PB/1% BSA, adding 166 µl
of PB/SDS, and diluting with 833 µl of PB/Triton. These 1 ml samples
were mixed with 200 µl of streptavidin beads and rotated at 4°C for
2 hr. From this slurry, 240 µl (20% of total) was removed and added
to 100 µl of 3× sample buffer. The beads (80% of total) were
precipitated from the remaining slurry, and the supernatant was added
to 400 µl of 3× sample buffer. The beads were washed in 800 µl
each of PB/Triton, PB/Triton with 600 mM NaCl, and PB
alone. These washes were saved and added to 400 µl of 3× sample
buffer. The biotinylated proteins were eluted from the streptavidin
beads into 1200 µl of 1× sample buffer. All samples were boiled for
3 min and spun in a microcentrifuge before an equal volume of each was
loaded per lane. Serial dilutions of the total extract were run on each
gel to generate a standard curve for each experiment. After transfer of
the proteins, the immunoblots were probed with C-GluR1, stripped, and
reprobed with tubulin antibody.
Efficiency of biotinylation. In experiments designed to
determine the efficiency of biotinylation, cells were prepared for biotinylation as described, scraped into the biotinylation reagent, sonicated, and spun for 15 min at 100,000 × g; the pellets
were washed three times with
PBS/Ca2+/Mg2+ including 0.1%
BSA. Detergent-soluble extracts of the pellets were prepared as
described above, by adding PB/SDS and diluting with PB/Triton. These
extracts were sonicated and spun in a refrigerated microcentrifuge at
14,000 × g for 15 min, and the supernatants were
processed as described above for the surface expression
experiments.
Quantitation of Western blots. All proteins were visualized
by ECL and analyzed using a Molecular Dynamics Personal Densitometer SI. Protein band intensity was quantitated using ImageQuant. A standard
curve was generated for each film, such that band intensity could be
expressed as a percentage of the biotinylated material at
t = O (for the half-life experiments) or a percentage
of the total extract (for the surface expression experiments). To
calculate half-lives, we plotted the natural logarithm of the
percentage of protein remaining as a function of time and found the
slope of the resulting regression line. Assuming first-order decay
kinetics, the half-life is equal to ln 2/slope.
Metabolic labeling and half-life determination of total
GluR1. The growth media from spinal cord cultures was removed and replaced with 1 ml of methionine and cysteine-free media containing 0.8 mCi Tran35S-label. Cells were incubated in this
media for 30 min, washed with PBS, and then returned to the incubator
in their original growth media. At various time points, cell extracts
were prepared by adding 1 ml of ice-cold lysis buffer (50 mM sodium phosphate, pH 7.5, with 1.0% Triton X-100, 0.5%
deoxycholate, 0.2% SDS, 50 mM NaF, 10 mM
sodium pyrophosphate, 5 mM EDTA, 5 mM EGTA, and 10 U/µl aprotinin) to the plates. The cells were scraped off the plates, transferred to a microfuge tube, mixed by inversion, and centrifuged at 4°C for 15 min at 15,000 × g. The
supernatants were stored at 80°C before use. Once all samples had
been collected, 100 µl of a 1:1 slurry of protein A-Sepharose CL-4B
(Pharmacia, Piscataway, NJ) and lysis buffer with 1% bovine serum
albumin was added to each tube. After rotating for 1 hr at 4°C, the
mixtures were spun at 2000 × g for 2 min, and the
supernatants were transferred to microfuge tubes containing 150 µl of
protein A-Sepharose that had been prebound with 4 µl of crude C-GluR1
antibody. After rotating for 2 hr at 4°C, the beads were washed
sequentially with the following solutions: lysis buffer alone (twice);
lysis buffer with 750 mM NaCl (three times); 10 mM sodium phosphate, pH 7.5, with 0.1% Triton X-100, 50 mM NaF, and 5 mM EDTA (twice). After removal of
the final wash, SDS-PAGE sample buffer was added directly to the beads
to elute antibody-bound GluR1. Eluted proteins were boiled for 3 min
before being loaded on 7.5% polyacrylamide gels. After completion of
electrophoresis, gels were dried and visualized on a Molecular Dynamics
Phosphoimager. Half-lives were calculated after quantitation of labeled
GluR1 in each sample using ImageQuant software.
RESULTS
An N-terminal antibody induces patches of surface GluR1
To study the distribution of surface GluRs during synaptogenesis,
we generated an antibody against an N-terminal domain of GluR1 (amino
acids 251-269) (Molnar et al., 1994), a region thought to be on the
extracellular face of the plasma membrane (Hollmann et al., 1994; Roche
et al., 1994). This antibody, N-GluR1, recognized a number of proteins
in total extracts of cultured spinal cord neurons (Fig.
1A); however, when used
to probe a sample including only cell surface proteins that had been
biotinylated and precipitated with streptavidin beads, N-GluR1
recognized a single predominant protein. This protein comigrated with
GluR1, as recognized by our previously described intracellular
antibody, C-GluR1 (Blackstone et al., 1992). Presumably, the additional
species recognized by N-GluR1 in total extracts of spinal cord neurons
are intracellular proteins. When N-GluR1 was added to mature, live
spinal cord cultures for 1 hr and then processed for double labeling
with C-GluR1 as described in Materials and Methods, N- and C-GluR1
staining completely colocalized (Fig. 1B).
Preincubation of N-GluR1 with its cognate peptide completely abolished
its immunoreactivity (Fig. 1C). Studies with HEK-293 cells
transiently transfected with recombinant GluR1 gave similar results
(data not shown). These results confirm the specificity of the N-GluR1
antibody when used for staining surface GluR1 receptors in live
neurons.
Fig. 1.
Characterization of the N-GluR1 antibody. An
antibody was raised against an N-terminal region of GluR1 as described
in Materials and Methods. A, Immunoblots of either total
(T) or biotinylated- and
streptavidin-immunoprecipitated (B) spinal cord
cultures were probed with antibodies to either N-GluR1 or C-GluR1.
Although N-GluR1 recognized several proteins, in addition to GluR1, in total extracts of spinal cord, there was only a single, appropriately sized protein in the pool of surface molecules. B,
C, Live cultures of spinal cord neurons were labeled
with N-GluR1 (B) or N-GluR1 plus peptide (0.1 mg/ml) (C) for 1 hr and then fixed and labeled with C-GluR1 as described in Materials and Methods. Surface labeling with N-GluR1 completely overlapped with C-GluR1 and was completely blocked by peptide.
[View Larger Version of this Image (55K GIF file)]
In our previous study, we examined the distribution of GluR1 in spinal
cord neurons using the C-terminal antibody C-GluR1, which in
fixed and permeabilized cells detects receptors both on the surface and
within the cytoplasm of dendrites. Before synaptogenesis, GluR1
staining was distributed diffusely throughout the cell. As synapses
formed over the first 10 d in vitro, GluR1 clusters appeared only at sites of synaptic vesicle accumulation, presumably representing synaptic contacts. In contrast, when N-GluR1 was added to
live spinal cord cultures before synaptogenesis (days 3 and 4 in
vitro), a significant and widely distributed population of
nonsynaptic, surface receptors was observed in the form of microaggregates (Fig.
2A). These aggregates
likely represent antibody-induced receptor patching, because only a
diffuse staining pattern was seen when neurons were incubated with a
Fab fragment of N-GluR1 (Fig. 2E-H) or when
cells were fixed but not permeabilized before the addition of N-GluR1
(data not shown). The specificity of the N-GluR1 surface staining at
these sites is revealed by corresponding patches detected by the
C-terminal antibody C-GluR1 (Fig. 2B). These
antibody-induced clusters of GluR1 are not found at sites of cell-cell
contact (Fig. 2D,L), nor do they colocalize with the
synaptic vesicle protein synaptophysin (Fig. 2C,K),
indicating that they are not associated with synapses. To confirm that
the patches induced by N-GluR1 were indeed present on the cell surface, we stained N-GluR1-induced patches with an FITC-labeled secondary antibody without previous permeabilization (Fig. 2I).
Under these conditions, staining for intracellular epitopes such as
C-GluR1 was absent (Fig. 2J). The surface population
of receptors detected with N-GluR1 represents only a fraction of the
total dendritic pool of GluR1. A 1 hr incubation with N-GluR1 appeared
to patch all the surface GluR1 (Fig. 2A) but left a
significant portion of the total pool, defined by C-GluR1, unclustered
(Fig. 2B). This latter population likely reflects a
pool of GluR1 within the dendritic cytoplasm but not on the surface
(see below).
Fig. 2.
Induction of extrasynaptic receptor patches by the
N-GluR1 antibody. Live, 3-d-old cultures of spinal cord neurons were
incubated with whole N-GluR1 (A-D, I-L) or a Fab
fragment of N-GluR1 (E-H) and then fixed
and processed as described. Antibody-induced surface patches of N-GluR1
(A), which are not seen with a Fab fragment of
N-GluR1 (E), have corresponding clusters of
C-terminal GluR1 staining (compare B with
F). In nonpermeabilized neurons
(I) surface N-GluR1 staining is observed,
whereas C-terminal staining is not detectable. The nonsynaptic location
of this staining is confirmed by the lack of associated synaptophysin
stain (D, H, L), and the absence of cell-cell contact
at these sites (C, G, K).
[View Larger Version of this Image (71K GIF file)]
Redistribution of GluR1 to synapses during neuronal maturation
The pool of mobile, extrasynaptic, surface GluR1 decreased
dramatically as neurons matured during the first 2 weeks in culture. As
shown in Figure 3, antibody-induced
extrasynaptic patches of surface GluR1 (Fig. 3A-C) are not
seen in mature cultures (Fig. 3D-F). In contrast, at
these latter time points, receptors were seen to localize primarily at
sites of presynaptic vesicle accumulation, defined by the presence of
the synaptic vesicle protein synaptophysin. Because fixation of the
mature cultures before staining with N-GluR1 resulted in an identical
distribution, with clustered receptor observed only at synapses (data
not shown), we conclude that these synaptic clusters are immobile and
not antibody-induced. In Figure 4, the
replacement of the diffuse, mobile receptor pool by a largely synaptic
and immobile pool is evaluated semiquantitatively by examining a large
number of cells from cultures of various ages. GluR1-positive neurons,
which make up 70% of the total population, were categorized as
"extrasynaptic" if they had mostly N-GluR1-induced, nonsynaptic
receptor patches, "synaptic" if they had mostly
synaptophysin-associated receptor clusters, and "transitional" if
they had both synaptic and nonsynaptic GluR1 clusters (see Figure 4
legend for definitions). On day 3 in vitro, 72% of neurons
had predominantly extrasynaptic receptors, 8% had predominantly
synaptic receptors, and 19% were transitional. By day 11 the
distribution had changed to 0% extrasynaptic, 80% synaptic, and 20%
transitional. Thus, during the maturation of spinal cord neurons in
culture, there is a significant redistribution of surface GluR1 to
synapses, which is nearly complete by day 11. Growing these cultures in
CNQX (10 µM) and APV (1 mM), concentrations sufficient to completely block synaptic transmission (O'Brien et al.,
1997), had no effect on GluR1 redistribution.
Fig. 3.
The distribution of surface GluR1 changes over
time in culture. Antibody-induced, nonsynaptic patches of GluR1 are
abundant on the dendrites of neurons after 4 d in culture
(A-C). On day 11, however
(D-F), all live cell staining with
N-GluR1 is confined to synapses, defined by the presence of presynaptic
synaptophysin stain (F). Note that
N-GluR1-induced antibody patching appears to cluster all the surface
immunostaining at both day 4 (A) and day 11 (D) but leaves a significant portion of the total
GluR1 signal, seen with the C-terminal antibody, unclustered (B,
E).
[View Larger Version of this Image (54K GIF file)]
Fig. 4.
Time course of surface GluR1 redistribution
in vitro. Synaptic clusters and extrasynaptic patches of
GluR1 were visualized as described in Figures 2 and 3.
GluR1-immunopositive neurons were categorized as either synaptic,
extrasynaptic, or transitional on the basis of the following scheme.
"Extrasynaptic" neurons averaged more than eight extrasynaptic
patches and one or fewer synaptic cluster per major dendrite.
"Synaptic" neurons averaged two or more synaptic clusters and two
or fewer extrasynaptic patches per major dendrite. "Transitional"
neurons fell between these two categories. Neurons were taken from a
series of three platings. The total number of neurons at each point is
listed above that point. For every neuron, the length of each major
dendrite examined was usually 60 µm.
[View Larger Version of this Image (13K GIF file)]
As noted above, little surface staining of GluR1 appeared outside of
antibody-induced patches on day 4, or outside synaptic contacts on day
11. In contrast, C-GluR1, which stains both intracellular and plasma
membrane GluR1, recognized a significant pool of diffuse GluR1
throughout the dendrite and cell body at both time points (Fig.
3B,E), even after the surface receptors were patched with N-GluR1. Thus, we conclude that although there is a decrease in extrasynaptic surface GluR1 as synapses form, there remains a substantial pool of diffuse GluR1 located throughout the dendritic cytoplasm. The specificity of this diffuse staining for GluR1 was
demonstrated in two ways (Fig. 5). First,
in the 30% of neurons that were GluR1 immunonegative both at the cell
body and at synaptic contacts, there was little diffuse dendritic
staining (Fig. 5B, open arrows). Second, nearly
all the diffuse dendritic stain could be abolished by preincubation
with the appropriate peptide (Fig. 5C,D). Therefore, before
synaptogenesis two pools of GluR1 exist in cultured dendrites: one
diffuse and freely mobile on the cell surface, and another, larger pool
(see below) within the cytoplasm.
Fig. 5.
The diffuse dendritic staining for GluR1 is
specific. For A and B, an 11 d
spinal cord culture was stained with Cy3-labeled C-GluR1, revealing two
neurons, one of which is GluR1 immunopositive and one of which is GluR1
immunonegative. Note the differences in diffuse dendritic stain between
the immunopositive (closed arrows) and immunonegative
(open arrows) neurons. The categorization of these
cultured neurons as immunopositive and immunonegative is based on cell
body and synaptic stain, and correlates well with in
situ hybridization signal for GluR1 mRNA. In C
and D, a GluR1-immunopositive neuron from a 5 d
culture, identified by live N-GluR1 stain (C),
shows minimal dendritic stain with Cy3-labeled C-GluR1 when the
antibody has been preincubated with peptide (for comparison, see Fig.
3B,E). In all control experiments, neuronal cell bodies
showed low-level, nonspecific staining, which did not extend into the
dendrites.
[View Larger Version of this Image (103K GIF file)]
Distribution of GluR1 on the surface of isolated neurons
To study the distribution of surface GluR1 in neurons devoid of
their normal synaptic input, we grew single, isolated spinal cord
neurons on islands of glia for up to 2 weeks in vitro (Segal and Furshpan, 1990). In our accompanying paper in this issue (O'Brien et al., 1997), we showed that 70% of these isolated neurons express GluR1 but that only 30% of these have GluR1 clusters at sites of
autaptic connections. These GluR1-expressing isolated neurons continued
to have N-GluR1-clusterable, extrasynaptic GluR1 on their surfaces for
up to 2 weeks in vitro, a time at which almost all neurons
in mass cultures have suppressed extrasynaptic GluR1 expression. This
was true even in neurons that had clusters of synaptic GluR1 (Fig.
6). Possible explanations for these
results could lie in the density of either excitatory or inhibitory
synapses on neurons grown in isolation [compare Fig. 1 with Fig. 6
here, or compare Fig. 2 with Fig. 5 in O'Brien et al. (1997)].
Alternately, autaptic synapses may not be as proficient as heterotopic
synapses in forming excitatory synapses. We suspect that some component of appropriate synaptogenesis may be limiting under these
conditions.
Fig. 6.
Lack of suppression of extrasynaptic GluR1 in
island cultures. Island cultures containing isolated spinal cord
neurons were grown for 10 d. Under these conditions all synaptic
contacts are autaptic. In A, the staining pattern of
N-GluR1 continues to show extrasynaptic patches of GluR1
(arrowheads) despite autaptic clusters of GluR1
(arrows).
[View Larger Version of this Image (67K GIF file)]
The half-life of surface GluR1 increases as neurons mature
To correlate changes in receptor distribution with changes in
receptor turnover, we used membrane-impermeant biotinylation, combined
with streptavidin precipitation, to distinguish surface from subsurface
pools of GluR1. Figure 7A
demonstrates that the biotinylation reaction proceeds quickly, peaking
in 10 min, and that all biotinylated GluR1 may be recovered by a single
streptavidin precipitation. Moreover, virtually no tubulin is detected
in the streptavidin-precipitated material, indicating that only cell surface proteins were recovered. To examine the efficiency of the
biotinylation reaction, neurons were lysed in the presence of the
biotinylating reagent, and membranes were prepared immediately from the
lysed cells. After streptavidin precipitation of the detergent-soluble
membrane extracts, no GluR1 could be detected in the supernatant (Fig.
7B), indicating that the biotinylation reaction was complete
under these conditions. It should be noted, however, that fewer regions
of GluR1 are accessible to biotin in the surface biotinylation
experiments described below. Consequently, it is possible that the
efficiency of surface biotinylation is not 100%.
Fig. 7.
Time course and efficiency of biotinylation of
GluR1 in cultured spinal cord neurons. Cultures of spinal cord neurons
were biotinylated at 4°C with 1 mg/ml NHS-SS-biotin. In
A, cultures were exposed to the biotinylating reagent
for different times, and the detergent soluble fraction of each culture
was added to streptavidin-conjugated beads and incubated for 2 hr. The
supernatant recovered from the precipitation of the 15 min-treated
neurons was reprecipitated with streptavidin beads, and the additional streptavidin-precipitated material was loaded as well. All samples were
loaded such that each lane represents 1% of the total material per
plate. The gel was transferred to immobilon and probed with both
C-GluR1 and anti-tubulin antibodies. In B, one plate of
neurons was scraped into the biotinylating reagent, sonicated, spun at 14,000 × g for 15 min, and resuspended in
precipitation buffer including 0.2% SDS and 1% Triton X-100. A
fraction of this was saved and loaded as total extract. The remainder
was incubated for 2 hr with streptavidin-conjugated beads. The
supernatant, streptavidin-precipitated, and total extracts were loaded
such that each lane represents 1% of the material from the plate. The gel was transferred and probed with the C-GluR1 antibody.
[View Larger Version of this Image (20K GIF file)]
To determine the half-life of surface GluR1 before and after excitatory
synapse formation, plates of spinal cord neurons from day 4 and day 11 in vitro were biotinylated, returned to the incubator, and
solubilized at varying times after the initial biotinylation. Surface
GluR1 was recovered with streptavidin and visualized by immunoblot
(Fig. 8A). Quantitative
analysis of the immunoblots was performed to determine the fractional
decrease of biotinylated (surface) GluR1 over time (see Materials and
Methods), from which the receptor half-life could be calculated (Fig.
8B). Table 1 shows that
between days 4 and 11 in vitro, a time at which GluR1 goes
from mostly extrasynaptic to almost entirely synaptic, the half-life of
surface GluR1 increased from 10.4 ± 2.2 to 30.5 ± 12.7 hr
(paired t test; p < 0.05).
Fig. 8.
Half-life of cell surface GluR1 in spinal cord
neuronal cultures at day 4 and day 11 in vitro. Plates
of spinal cord neurons were biotinylated at day 4 and day 11 and
recultured for 0-24 hr, at which time cell extracts were harvested,
sonicated, and frozen. Subsequently, these samples were thawed and
incubated with streptavidin-linked beads, and the
streptavidin-precipitated material was loaded onto gels
(A). A standard curve including serial dilutions
of the t = 0 streptavidin-precipitated material was
included on each gel for purposes of quantitation. After transfer, gels
were probed with the C-GluR1 antibody. In B, the natural log of the percent of remaining surface GluR1 was plotted against time,
and half-lives were calculated from the regression slopes of the
resulting lines. The results of the experiment in A are shown.
[View Larger Version of this Image (36K GIF file)]
Table 1.
Summary of half-life and percent of receptor on surface
experiments
| Experiment |
Day 4 |
Day 11 |
n |
|
| GluR1 surface
half-life |
10.4 ± 2.2 hr |
30.5
± 12.7 hr |
3 |
| GluR1 total half-life |
12 ± 1.4 hr |
31
± 4.2 hr |
2 |
| NGT half-life |
10.3 ± 0.7 hr |
13.2
± 2.4 hr |
3 |
| % GluR1 on surface |
32.5 ± 4.5% |
68.6
± 12.9% |
5 |
|
|
The means ± SD for the half-lives of surface
GluR1 and total GluR1 on days 4 and 11 are shown. A paired t
test demonstrated a significant increase in the half-life of surface
GluR1 from day 4 to day 11 (p < 0.05). The
half-life of the neuronal glutamate transporter (NGT) did not change
significantly between day 4 and day 11. Also shown are values for the
percent of GluR1 on the surface of spinal cord neurons. There is a
significant increase in the percent of GluR1 on the surface from day 4 to day 11 (p < 0.05).
|
|
To determine the half-life of the total pool of GluRs, spinal cord
cultures were metabolically labeled with 35S-methionine.
Cell extracts were prepared at a number of time points after the
initial 35S-methionine pulse, and GluR1 was
immunoprecipitated and separated by SDS-PAGE (data not shown). The
metabolic half-lives of total GluR1 at days 4 and 11 were calculated to
be 12 ± 1.4 and 31 ± 4.2 hr, respectively (Table 1). Thus,
the incorporation of GluR1 into synapses correlates with an increase in
its half-life. In contrast, the half-life of the neuronal glutamate
transporter, which remains diffusely distributed on the neuronal
surface between days 4 and 11 in culture (O'Brien et al., 1997), does
not change (Table 1).
The fraction of GluR1 on the cell surface increases as
neurons mature
As mentioned previously, immunocytochemical studies had indicated
that there exists a sizable pool of intracellular GluR1 in cultured
spinal cord neurons (Figs. 2, 3, 4). We again used surface biotinylation
to determine the percentage of GluR1 that resides on the cell surface
at different times in vitro. In these experiments surface
GluR1 was biotinylated, precipitated with streptavidin, and compared
with total GluR1 using quantitative immunoblotting (Fig.
9). We found that only 32.5 ± 4.5%
of the total GluR1 was located on the cell surface at day 4, a time at which little synapse formation has taken place. By day 11, in contrast,
68.6 ± 12.9% of GluR1 could be found on the cell surface. This
represented a significant increase in the percentage of GluR1 on the
surface as neurons mature and GluR1 redistributes to synapses (Student's t test; p < 0.05). As mentioned
above, these estimates of the size of the surface pool of GluR1 may be
slightly high if the efficiency of surface biotinylation is actually
less than the 100% our controls suggested.
Fig. 9.
Determination of the fraction of GluR1 on the cell
surface at day 4 and day 11. Spinal cord neuronal cultures were
biotinylated, harvested, and incubated with streptavidin beads. Samples
of the total extract, streptavidin-supernatant (intracellular),
streptavidin-precipitate (surface), and washes were saved and loaded,
such that each lane represents protein from 1% of the plate. Serial
dilutions of the total extract were also loaded for purposes of
quantitation. The gels were transferred, probed with C-GluR1, stripped,
and reprobed with anti-tubulin antibody. In each of five experiments,
<1% of the total tubulin was precipitated as a "surface"
molecule.
[View Larger Version of this Image (34K GIF file)]
DISCUSSION
To investigate the regulation of excitatory transmitter receptors
during synapse formation in neurons from the CNS, we have studied
cultured spinal cord neurons using various antibodies against GluR
subunits. Previously, we found that synapse formation occurs over the
course of 10 d in these cultures, beginning on day 2 (O'Brien et
al., 1997). In the present study we have used an antibody that
recognizes an extracellular domain of GluR1, N-GluR1, to study the
distribution of surface receptors during this same period. Before
synapse formation, GluR1 is distributed widely over the surface of
dendrites in a form that is mobile and easily aggregated into patches
by the addition of N-GluR1. Biotinylation studies suggest that at this
time, the surface component represents approximately one-third of the
total receptor pool, with the remaining two-thirds in the cell body and
dendritic cytoplasm. Immunohistochemical studies also suggest the
presence of a significant cytoplasmic pool of dendritic GluR1 before
synapse formation. As synapses form over the subsequent 2 weeks,
surface GluR1 is gradually directed away from extrasynaptic sites and
toward sites of appropriate cell-cell contact. During this same time
period there is an increase in the percentage of total cellular GluR1 on the surface, although a substantial dendritic cytoplasmic pool remains. Whether the redistribution of GluR1 away from extrasynaptic sites is attributable to a directed insertion at synapses or to migration within the plane of the membrane is uncertain. Some conclusions can be drawn regarding the mechanisms leading to the redistribution of GluR1. First, synaptic activity had no effect on
receptor redistribution. Spontaneous synaptic activity begins on day 3 in vitro and increases rapidly over the subsequent 7 d
(O'Brien et al., 1997). Previous work showed that incubating cultures
with TTX (Craig et al., 1994) or CNQX and APV (O'Brien et al., 1997),
at concentrations sufficient to block all synaptic activity, had no
effect on synapse formation. Our present study suggests that synaptic
activity also plays no role in the redistribution of surface receptors.
In contrast, neurons grown in island cultures, with a paucity of
synaptic input, had no downregulation of extrasynaptic receptors.
Therefore it seems that some quantitative aspect of synapse formation,
independent of synaptic activity, causes both an increase in GluR1 at
synaptic sites and a decrease at extrasynaptic sites. Perhaps a growth
factor or cell adhesion molecule derived from the presynaptic neuron
directs the redistribution of AMPA receptors to synapses during
development. The absence of a significant effect of synaptic activity
on receptor redistribution in cultured spinal neurons is in contrast to
results observed at the neuromuscular junction where extrasynaptic AChR
levels are regulated by electrical activity. AChRs reappear
extrasynaptically after blockade of synaptic transmission or muscle
denervation, a process that can be antagonized by chronic electrical
stimulation (Lomo and Rosenthal, 1972; Berg and Hall, 1975a; Lomo and
Westgaard, 1975).
Another aspect of synapse formation, shared both by spinal cord neurons
and by the neuromuscular junction, is the stabilization of receptors at
synapses. Using biotinylation techniques, we found that GluR1 receptor
subunits on the cell surface have a half-life of ~10 hr at day 4. By
day 11, however, the half-life of surface GluR1 increased to ~30 hr,
coincident with the aggregation of surface receptors at synapses. Very
similar values were obtained for the half-lives of the total pool of
GluR1, determined by pulse-chase experiments using
35S-methionine as a label. The values obtained for GluR1
half-lives by these two methods are similar, suggesting that, as has
been found in other cell-surface biotinylation studies (Volz et al., 1995), the biotinylation of surface molecules has little effect on
GluR1 metabolism. The increase in surface GluR1 observed between days 4 and 11 in vitro is likely a consequence of both the
increased half-life of surface GluR1, attributable to its stabilization at synapses, and to the increasing number of synapses in these cultures. Further work will be required to determine what fraction of
synthesized GluR subunits ultimately reaches the cell surface and to
elucidate the degradative pathways involved in their metabolism. Studies examining the surface expression and metabolism of other GluR
subunits will also be of significant interest.
The current model for neurotransmitter receptor regulation during
synaptogenesis has emerged largely from work at the neuromuscular junction. In this system, preexisting and mobile surface AChRs are
trapped and stabilized at sites of nerve-muscle contact under the
influence of agrin, a basement membrane bound molecule (Bowe and
Fallon, 1995). In a second step, the presynaptic nerve terminal directs
the synthesis and insertion of locally formed receptors under the
influence of ARIA (Falls et al., 1993), a molecule that may also
initiate the synthesis of a novel receptor subunit. Third, nerve-induced electrical activity in the postsynaptic myotube causes a
downregulation of extrajunctional receptors (Avila et al., 1989). In
spinal cord cultures, the diffusely distributed population of surface
GluR1 found before synapse formation could conceivably aggregate at
synapses. Indeed, this possibility has been suggested by others who
have detected the presence of extrasynaptic surface AMPA receptor
subunits in both the hippocampus (Baude et al., 1995) and striatum
(Bernard et al., 1997); however, direct proof for or against this
theory is lacking. An alternative hypothesis, which we are currently
testing, involves the directed insertion of preformed receptors at
sites of synaptic contact. These receptors could be derived from the
large intradendritic pool of receptors that we observed before and
after synapse formation. These intradendritic receptors could either be
associated with the extensive sarcoplasmic reticulum seen in the
processes of many neurons (Spacek and Harris, 1997) or could be
directly linked to the cytoskeleton. Whether this pool of subsurface
receptors is eventually incorporated into synapses is unclear. Kharazia
et al. (1996) recently observed GluR1 containing subsurface vesicles
associated with the cytoplasmic face of many GluR1-containing cortical
synapses. In contrast, Baude et al. (1995) failed to observe similar
subsynaptic vesicles in the hippocampus. The disappearance of
extrasynaptic receptors seen in our present study could be a reflection
of either synaptic stabilization or localized insertion of GluR1, both
of which could be subunit dependent. The future isolation and
characterization of proteins involved in GluR clustering should help to
clarify the molecular mechanisms underlying excitatory synaptogenesis in these neurons.
FOOTNOTES
Received February 28, 1997; revised June 2, 1997; accepted June
10, 1997.
Correspondence should be addressed to Dr. Richard Huganir, Howard
Hughes Medical Institute, PCTB 900, Johns Hopkins Medical School, 725 N. Wolfe Street, Baltimore, MD 21205.
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E. Molnar, L. Pickard, and J. K. Duckworth
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N. Sans, C. Racca, R. S. Petralia, Y.-X. Wang, J. McCallum, and R. J. Wenthold
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L. Pickard, J. Noel, J. M. Henley, G. L. Collingridge, and E. Molnar
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L. Shen, F. Liang, L. D. Walensky, and R. L. Huganir
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C. D. Hohnke, S. Oray, and M. Sur
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J. R. Cottrell, G. R. Dube, C. Egles, and G. Liu
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D. L. Pettit and G. J. Augustine
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N. Filippova, A. Sedelnikova, Y. Zong, H. Fortinberry, and D. S. Weiss
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N. Filippova, R. Dudley, and D. S Weiss
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