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The Journal of Neuroscience, October 15, 2002, 22(20):8838-8849
The Group I Metabotropic Glutamate Receptor Agonist
(S)-3,5-Dihydroxyphenylglycine Induces a Novel Form of
Depotentiation in the CA1 Region of the Hippocampus
Wei-Ming
Zho*,
Jia-Lin
You*,
Chiung-Chun
Huang, and
Kuei-Sen
Hsu
Department of Pharmacology, College of Medicine, National
Cheng-Kung University, Tainan 701, Taiwan
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ABSTRACT |
The ability of activation of group I metabotropic glutamate
receptor (mGluR) to induce depotentiation was investigated at Schaffer
collateral-CA1 synapses of rat hippocampal slices. Brief bath
application (5 min) of group I mGluR agonist
(S)-3,5-dihydroxyphenylglycine (DHPG) (10 µM) induced a long-term depression of synaptic
transmission or depotentiation (DEP) of previously established
long-term potentiation (LTP), which was independent of NMDA or
A1 adenosine receptor activation. This DHPG-DEP was
observed when DHPG was delivered 3 min after LTP induction. However,
when DHPG was applied at 10 or 30 min after LTP induction,
significantly less depotentiation was found. DHPG-DEP (1) is reversible
and has the ability to unsaturate LTP, (2) is synapse specific, (3)
does not require concurrent synaptic stimulation, (4) is
mechanistically distinct from NMDA receptor-dependent depotentiation,
(5) requires mGluR5 activation, (6) requires rapamycin-sensitive mRNA
translation signaling, (7) does not require phospholipase C or protein
phosphatase activation, and (8) is not associated with a change in
paired-pulse (PP) facilitation. In addition, the ability of DHPG to
reverse LTP was mimicked by a long train of low-frequency (1 Hz/15 min)
PP stimulation. Moreover, the expression of DHPG-DEP is associated with
a reduction in the increase of the surface expression of AMPA receptors
seen with LTP. These results suggest that the activation of mGluR5 and
in turn the triggering of a protein synthesis-dependent internalization of synaptic AMPA receptors may contribute to the DHPG-DEP in the CA1
region of the hippocampus.
Key words:
long-term potentiation (LTP); depotentiation; DHPG; metabotropic glutamate receptor (mGluR); AMPA receptor; hippocampus
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INTRODUCTION |
Activity-dependent modulation of
synaptic strength is an important component in the current
understanding of the cellular mechanisms underlying learning and memory
of the brain (Abraham and Bear, 1996
). Much of our understanding of
activity-dependent synaptic modification and its functional relevance
comes from studies on the mammalian hippocampus. In the hippocampus,
brief trains of activity can trigger long-lasting increases or
decreases in synaptic strength (Bliss and Lømo, 1973; Mulkey and
Malenka, 1992; Bliss and Collingridge, 1993; Dudek and Bear, 1993).
Although both long-term potentiation (LTP) and long-term depression
(LTD) are remarkable for their stability, recent work has shown that they are initially labile and sensitive to disruption by various interfering events or agents. For example, the hippocampal CA1 LTP can
be reversed by afferent low-frequency stimulation (LFS) (Barrionuevo et
al., 1980; Fujii et al., 1991; O'Dell and Kandel, 1994; Stäubli
et al., 1996, 1998; Huang et al., 1999, 2001), episodes of hypoxia
(Arai et al., 1990), or pharmacological treatments that interrupt
cell-cell or cell-matrix interactions (Bahr et al., 1997), when given
shortly after LTP induction. This reversal of synaptic strength from
potentiated state to pre-LTP level is referred to as depotentiation
(DEP) (Barrionuevo et al., 1980; Fujii et al., 1991; O'Dell and
Kandel, 1994; Wagner and Alger, 1996; Huang and Hsu, 2001).
Because LTP, in the mammalian brain, is generally assumed to be a
synaptic mechanism underlying learning and memory formation (Bliss and
Collingridge, 1993), the processes involved in the depression of
synaptic potentiation may contribute to the mechanisms of memory loss
or forgetting. Although over the past two decades a number of
electrical stimulation protocols have been reported to effectively
induce depotentiation, none of these protocols have yet been tested for
their ability to cause forgetting in behaving animals (Martin et al.,
2000). The major problem with doing so is that it is difficult to
induce depotentiation on all relevant pathways of the hippocampal
formation after an individual learning experience. Moreover, the
characterization of the intrinsic pathways involved in behaving
learning has been technically difficult. In comparison with electrical
stimulation approaches, a pharmacological induction protocol would have
the advantage that it is easier to target all of the relevant synapses
with a drug and may allow the evaluation of the functional relevance of
depotentiation in behaving animals. Thus, the primary goal of this
study is to establish a pharmacological depotentiation protocol and to
elucidate its induction mechanisms. Here we characterize a group I
metabotropic glutamate receptor (mGluR) agonist
(S)-3,5-dihydroxyphenylglycine (DHPG) induction
protocol that reliably induces a time-dependent depotentiation at
Schaffer collateral-CA1 synapses of rat hippocampal slices. We found
that mGluR5 and a rapamycin-sensitive protein synthesis-dependent
internalization of AMPA receptors are required for the development of
DHPG-induced depotentiation.
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MATERIALS AND METHODS |
Hippocampal slice preparation. Animal care was
consistent with the guidelines set by the Laboratory Animal Center of
National Cheng-Kung University (NCKU). All experimental procedures were approved by the NCKU Institutional Animal Care and Use Committee. Horizontal hippocampal slices (400 µm thick) were prepared from 4- to
5-week-old male Sprague Dawley rats after decapitation under halothane
anesthesia (Pike et al., 2000). After their preparation, slices were
placed in a holding chamber of artificial CSF (ACSF) oxygenated
with 95% O2-5% CO2 and
kept at room temperature for at least 1 hr before recording. The
composition of the ACSF solution was (in mM):
NaCl 117, KCl 4.7, CaCl2 2.5, MgCl2 1.2, NaHCO3 25, NaH2PO4 1.2, and glucose 11 at pH 7.3-7.4, equilibrated with 95% O2-5%
CO2.
Electrophysiological recordings. For the extracellular field
potential recordings, a single slice was then transferred to a
submerged-type recording chamber and held between two nylon nets. The
chamber consisted of a circular well of a low volume (1-2 ml) and was
perfused continuously with oxygenated ACSF at a flow rate of 2-3
ml/min at 32.0 ± 0.5°C. Standard extracellular field recording
techniques were used (Huang et al., 1999, 2001). Extracellular
recordings of field EPSPs (fEPSPs) were obtained from the
stratum radiatum using microelectrodes filled with 1 M NaCl (resistance 2-3 M
). A bipolar
stainless steel stimulating electrode was placed in stratum radiatum to
activate Schaffer collateral/commissural afferents at 0.033 Hz. The
stimulation strength was set to elicit responses equivalent to 30-40%
of the maximal fEPSP. In all experiments, baseline synaptic
transmission was monitored for 30 min before drug administration or
delivering high-frequency tetanic stimulation (TS). The strength of
synaptic transmission was quantified by measuring the slope of fEPSP.
The fEPSP slopes were measured from ~20-70% of the rising phase
using a least-squares regression. LTP was induced by high-frequency TS,
at the test pulse intensity, consisting of two 1 sec trains of stimuli
at 100 Hz, delivered with an interval of 20 sec. In some cases, as
indicated in Results, depotentiation was induced by application
of 15 min paired-pulse low frequency trains of stimuli (PP-LFS) (with a
50 msec interstimulus interval) at 1 Hz, and the stimulation intensity
was the same as the test pulse intensity. The responses during the
trains were not recorded, and for convenience these periods are not
shown on the graph. To stimulate independent inputs to the same cell
population, two bipolar stimulating electrodes were positioned on both
sides of the recording microelectrode, alternating every 15 sec. Their positions were arranged so that the same amount of current evoked two
responses that did not differ from each other by >10%. The absence of
cross-pathway paired-pulse facilitation (PPF) was used to ensure that
the two inputs were independent of each other. The stimulation strength
was set to elicit response for which the amplitude was 30-40% of the
maximum spike-free response. All values of residual potentiation
reported here were calculated as the changes in fEPSP slope measured 40 min after washout of DHPG. Intracellular recordings were made from CA1
pyramidal cells using glass microelectrodes filled with potassium
acetate (4 M) having resistances ranging from 60 to 80 M (Huang et al., 1999). The membrane input resistance of the
recording neurons was calculated from the voltage deflection produced
by a transient hyperpolarizing current pulse (0.1 nA, 80 msec) passed
through the recording microelectrodes. The EPSP slopes were
measured from ~20-70% of the rising phase using a least-squares
regression. Microelectrodes were pulled from microfiber 1.0 mm
capillary tubing on a Brown-Flaming electrode puller (Sutter
Instruments, San Rafael, CA). Electrical signals were collected with an
Axoclamp-2B (Axon Instruments, Foster, CA) filtered at 1 kHz and
sampled at 10 kHz, and an Intel Pentium-based computer with pCLAMP
software (Version 7.0; Axon Instruments, Foster City, CA) was used to
acquire the data on-line and analyze it off-line.
Biochemical measurements of surface-expressed AMPA
receptors. Biotinylation experiments were performed as described
previously (Chung et al., 2000). Hippocampal slices were prepared and
treated with high-frequency TS with or without DHPG treatment exactly as described in the electrophysiological experiments. At the end of the
experiments, the CA1 subregion of the hippocampal slices between the
positions of the stimulating and recording electrodes was dissected out
and immediately frozen on dry ice. Two to three microdissected CA1
subregions (from control, 50 min after LTP induction, 42 min after
washout of DHPG with or without previous LTP induction) were pooled
together. In each experiment, an entire set of control, LTP, or
depotentiation pooled slices was taken from the same animal. The
microdissected subregions were then incubated with ACSF containing 1 mg/ml sulfo-succinimidyl-6-(biotinamido) hexanoate (Pierce
Chemical Company, Rockford, IL) for 30 min on ice. Unreacted
biotinylation reagent was washed once with ice-cold ACSF and quenched
by two successive 20 min washes in ACSF containing 100 mM glycine, followed by two washes in ice-cold
TBS (50 mM Tris, pH 7.5, 150 mM NaCl). The microdissected subregions were lysed in ice-cold homogenate buffer (50 mM
Tris-HCl, 100 mM NaCl, 15 mM sodium pyrophosphate, 50 mM sodium fluoride, 5 mM
EGTA, 5 mM EDTA, 1 mM
phenylmethylsulfonyl fluoride, 0.5% Triton X-100, 2 mM benzamidine, 60 µg/ml aprotinin, and 60 µg/ml leupeptin) and ground with a pellet pestle (Kontes Glassware,
Vineland, NJ). Samples were sonicated and spun down at 14,000 × g at 4°C for 15 min. Thirty microgram lysates of the
resulting supernatant were removed to measure total GluR1, and 90 µg
lysates of the supernatant was incubated with 100 µl of 50%
Neutravidin agarose (Pierce Chemical Company) for 3 hr at 4°C to
measure the isolated biotinylated proteins. After the Neutravidin
agarose was washed five times with homogenate buffer, bound proteins
were eluted with SDS sample buffer by boiling for 15 min. Total protein
and isolated biotinylated proteins were analyzed by quantitative
immunoblotting with polyclonal anti-GluR1 C-terminal antibody (1:1000,
Upstate Biotechnology, Lake Placid, NY). It was then probed with
HRP-conjugated secondary antibody for 1 hr and developed using the ECL
immunoblotting detection system. Immunoblots were quantified by
densitometric measurement using BioLight software.
Drug application. All drugs were applied by dissolving them
to the desired final concentrations in the ACSF and by switching the
perfusion from control ACSF to drug-containing ACSF. Appropriate stock
solutions of drugs were made and diluted with ACSF just before
application. 8-Cyclopentyl-1,3-dipropylxanthine (DPCPX), cycloheximide,
anisomycin, actinomycin D, rapamycin, FK506, and U73122 were dissolved
in dimethylsulfoxide (DMSO) stock solutions and stored at 
20°C
until the day of experiment. The concentration of DMSO in the perfusion
medium was 0.05%, which alone had no effect on the induction of either
LTP or DHPG-induced depotentiation in the CA1 region of rat
hippocampus. LY341495 and LY367385 were prepared by first dissolving
them in an equimolar amount of NaOH as a concentrated stock solution
and then diluting to their final concentration in ACSF. DHPG,
2-methyl-6-(phenylethynyl)pyridine (MPEP),
D(-)-2-amino-5-phosphonopentanoic acid
(D-APV), cycloheximide, anisomycin, actinomycin
D, rapamycin, U73122, and okadaic acid were purchased from Tocris
Cookson Ltd. (Bristol, UK); FK506 was purchased from Calbiochem (La
Jolla, CA).
Statistical analysis. The data for each experiment were
normalized relative to baseline. Data are presented as mean ± SEM. The significance of the difference between the mean was calculated by a paired or unpaired Student's t test as appropriate.
Numbers of experiments are indicated by n. Probability
values of p < 0.05 were considered to represent
significant differences.
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RESULTS |
Brief bath application of group I mGluR agonist DHPG causes
synaptic depression
Consistent with previous studies (Palmer et al., 1997; Huber et
al., 2000, 2001), application of group I mGluR agonist DHPG (1-50
µM) for 5 min caused an acute, dose-dependent reduction of field potentials (Fig.
1A). At concentrations
10 µM, the fEPSP did not fully recover after
washout of drug. Instead, the synaptic response remained at a depressed
level (10 µM: 87.6 ± 5.7% of pre-DHPG
baseline; n = 7; 20 µM:
76.8 ± 4.7%; n = 8; 50 µM: 62.5 ± 5.6%; n = 5;
measured at 30 min after DHPG washout). This DHPG-induced long-term
synaptic depression was also confirmed by intracellular recordings. As
shown in Figure 1B, DHPG (10 µM) application resulted in an initial large
reduction of EPSP followed by a slow recovery, but the EPSPs remained
at a depressed level after washout of the drug (86.8 ± 4.7% of
pre-DHPG baseline; n = 5). Although DHPG application
also produced a transient membrane depolarization from 71.5 ± 2.2 to 63.5 ± 2.1 mV (n = 5; p < 0.05; paired Student's t test) that was accompanied by a
increase in membrane input resistance from 48.6 ± 2.6 to
56.8 ± 2.3 M (n = 5; p < 0.05; paired Student's t test), these changes were not long
lasting and did not correlate with the change in EPSPs. At 30 min after
DHPG washout, the resting membrane potential and membrane input
resistance were 71.2 ± 2.7 mV and 47.8 ± 2.6 M,
respectively, which were not significantly different from those found
during the control baseline period (p > 0.05;
paired Student's t test). Thus, DHPG-induced long-term synaptic depression seems not to be a mere reflection of the change of
membrane excitability of CA1 pyramidal neurons after DHPG treatment. Because a 5 min application of 10 µM DHPG
consistently produced a long-lasting synaptic depression without
significant long-term changes in neuronal excitability, we chose this
protocol to examine the role of group I mGluRs in the induction of
depotentiation at Schaffer collateral-CA1 synapses.
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Figure 1.
Effects of DHPG on EPSPs. A, A
brief application of DHPG for 5 min produced an acute, dose-dependent
depression of extracellular fEPSP. At concentrations 10
µM, the fEPSPs did not fully recover after washout of
drug. Example traces in the top portion are taken from a
slice treated with 10 µM DHPG. B, Summary
of experiments (n = 5) showing that DHPG (10 µM; 5 min) application caused a persistent depression of
intracellular EPSP. Although DHPG application also produced a transient
membrane depolarization and an increase in membrane input resistance,
these changes are not long lasting and did not correlate with the
change of EPSPs. Example traces in the top portion are
taken from a slice treated with 10 µM DHPG.
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DHPG induces depotentiation
Having confirmed the existence of a DHPG-induced LTD (DHPG-LTD) at
Schaffer collateral-CA1 synapses, we next asked whether the activation
of group I mGluRs by DHPG can trigger depotentiation. We initially
confirmed that our 100 Hz tetanization protocol reliably induced robust
LTP, similar to that reported previously (Fig. 2A) (Huang et al.,
1999, 2001). The slope of the fEPSP measured 50 min after
high-frequency TS was 153 ± 12% (n = 8) of
baseline (Fig. 2B). We next examined the effect of a
brief (5 min) application of DHPG (10 µM) on
the previously established LTP. As expected, DHPG applied 3 min after
LTP induction exerted a significant suppression of fEPSPs. After
washout of DHPG, the fEPSP recovered to near the pretetanus baseline
(Fig. 2C). On average, the fEPSP slope measured 40 min after
washout of DHPG was 105 ± 10% (n = 6) of baseline (Fig. 2D). In contrast, when DHPG was
applied 10 or 30 min after LTP induction, the magnitude of
depotentiation was reduced markedly (Fig.
2E,G). The mean residual
potentiation measured 40 min after DHPG washout was 136.8 ± 8.8%
(n = 6) and 138 ± 11% (n = 6) of
baseline, respectively (Fig.
2F,H). Comparison of the effect of DHPG applied to naive synapses (DHPG-LTD) or applied 3, 10, or 30 min after LTP induction was summarized in Figure 2I, in which the magnitude of depression measured 40 min after washout of DHPG was calculated relative to the baseline just
before DHPG application. The magnitude of depotentiation in which DHPG was applied 3 min after LTP induction was calculated by comparing the
synaptic responses at 40 min from the experiments illustrated in Figure
2D with the magnitude of LTP at 48 min from the
experiments illustrated in Figure 2B. As shown in
Figure 2I, the magnitude of depotentiation in which
DHPG was given 3 min (41.5%) after LTP was significantly larger than
the LTD elicited at naive synapses (13.2 ± 4.7%;
p < 0.05; unpaired Student's t test).
However, when DHPG was given 10 or 30 min after LTP induction, the
magnitude of depotentiation was the same as for LTD at naive synapses
(14.3 ± 9.4 and 16.2 ± 8.9%, respectively, versus
13.2 ± 4.7%; p > 0.05; unpaired Student's
t test). These results suggest that hippocampal CA1 LTP is
vulnerable to disruption by DHPG application within a brief period
after its induction.
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Figure 2.
DHPG induces a time-dependent reversal of LTP.
A, An example of the time course and magnitude of CA1
LTP. High-frequency TS to the Schaffer collateral afferent fibers
resulted in a large post-tetanic potentiation lasting for several
minutes after TS, followed by stable LTP expression. B,
Summary of data from eight experiments performed as in A. C, E,
G, Examples in which DHPG (10 µM; 5 min) was
applied at various times after LTP induction: C, 3 min
after; E, 10 min after; G, 30 min after.
D, F, H, Summary of experiments similar to those shown
in C, E, and G.
I, Histogram comparing the effect of DHPG applied to
naive synapses (DHPG-LTD) or applied 3, 10, or 30 min after LTP
induction. The magnitude of DHPG-LTD was calculated at 40 min after
washout of DHPG at naive synapses. The magnitude of 3 min delay
depotentiation was calculated by comparing the synaptic responses at 40 min after washout of DHPG from the experiments illustrated in
D with the magnitude of LTP at 48 min from the
experiments illustrated in B. Because both sets of data
have variance, it is not possible to calculate an SE of this
depotentiation value. Statistical analysis using ANOVA indicates that
this value is significantly different from LTP at naive synapses
(*p < 0.05). The magnitude of 10 and 30 min delay
depotentiation was calculated by comparing the synaptic responses at 40 min after washout of DHPG from the experiments illustrated in
F or H with the individual baseline
magnitude just before each DHPG application. Note that DHPG erased
potentiation when delivered 3 min after TS but was without effect when
applied 10 or 30 min after. The superimposed fEPSP in the
inset illustrates respective recordings from example
experiments taken at the time indicated by number.
Horizontal bars denote the period of the delivery of
DHPG. Calibration: 0.5 mV, 10 msec.
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Induction of DHPG-DEP unsaturates LTP
Having confirmed the existence of a time-dependent reversal of LTP
by brief application of DHPG (DHPG-DEP) at Schaffer collateral-CA1 synapses, we next examined whether the synapses that had undergone depotentiation could subsequently exhibit LTP. We adopted the approach
used to address the same question after the induction of homosynaptic
LTD at these synapses (Dudek and Bear, 1993; Mulkey et al., 1993). As
shown in Figure 3A,
high-frequency TS led to LTP that stabilized at a value of 159 ± 11% (30 min after TS; n = 8; p < 0.05; paired Student's t test) of baseline. In control experiments, application of a second TS delivered 30 min later resulted
in only 13.6 ± 8.1% additional LTP (n = 8;
p < 0.05; paired Student's t test). Figure
3B illustrates the experiments that were identical to those
in Figure 3A, with the only difference that DHPG (10 µM) was delivered 3 min after the first TS. The second TS that followed DHPG washout caused significantly more LTP than
in control experiments (P < 0.05; unpaired Student's t test) (Fig. 3B,C). A
second TS 30 min later caused the fEPSP amplitude to increase by
58.5 ± 9.4% (n = 8) of baseline. These results
suggest that DHPG-DEP is reversible and has the ability to unsaturate
LTP.
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Figure 3.
The depressed synapses can be potentiated.
A, Summary of eight experiments in which TS was applied
twice with a 30 min interval. B, Summary of eight
experiments identical to those shown in A with the
exception that DHPG (10 µM; 5 min) was delivered 3 min
after the first TS. In this case, subsequent TS 30 min later was able
to reverse the synaptic depression caused by DHPG. C,
Summary data in which the magnitude of the potentiation measured 30 min
after each TS was calculated relative to the baseline period before
each TS was applied. In control experiments, as illustrated in
A, we found that the first TS produced a potentiation of
65 ± 12% above baseline, whereas the second TS produced an
additional increase of only 13.6 ± 7.8%. In the experiments as
shown in B, in which the second TS followed a brief DHPG
application, it caused a potentiation of 58.5 ± 9.4% above
baseline (p < 0.05). Calibration: 0.2 mV,
10 msec.
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DHPG-DEP is synapse specific
The following experiment was designed to investigate whether
DHPG-DEP is synapse specific. To address this question, two stimulating electrodes were placed on both sides of the recording microelectrode in
the CA1 region to activate two groups of synapses to the same dendritic
field. As a typical example illustrated in Figure
4A, independence of
inputs activated by the two electrodes was assessed by verifying the
absence of heterosynaptic facilitation between the two inputs using
paired stimuli applied at 40 msec intervals. By having confirmed the
independence of afferents activated, we then induced LTP at the
synapses of one input (test pathway) by giving a high-frequency tetanic
stimulation; this was followed by a brief application of DHPG (10 µM). Application of the high-frequency tetanic
stimulation to the test pathway generated homosynaptic LTP without
noticeable modifications of the control, naive pathway. When DHPG was
applied 3 min after LTP induction, LTP was reversed almost completely.
The mean residual potentiation measured 40 min after washout of DHPG
was 115.3 ± 7.9% (n = 6) of baseline. At the
control pathway, the mean residual synaptic response measured 40 min
after washout of DHPG was 86.8 ± 5.6% (n = 6) of
pre-DHPG baseline, which was not significantly different from that
found in slices without receiving tetanic stimulation (87.7 ± 5.7%; n = 7) (Fig. 1A). These
results indicated that tetanizing of one pathway does not affect the
DHPG-mediated synaptic depression in the untetanized pathway.
Therefore, DHPG-DEP at Schaffer collateral-CA1 synapses is a
synapse-specific phenomenon.
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Figure 4.
DHPG-induced depotentiation does not require
concurrent synaptic stimulation: summary graph of six experiments. Two
stimulatory electrodes were used to activate two independent groups of
afferents (confirmed by no heterosynaptic facilitation). After a stable
baseline was established, LTP was induced on two afferent pathways by
coactivation, followed by application of DHPG (10 µM; 5 min). In the test pathway, the stimulation was stopped during the
application of DHPG and did not restart stimulation until 10 min after
the washout of DHPG. Depotentiation was successfully induced in both
control and test stimulation pathways. The superimposed fEPSP in the
inset illustrates respective recordings from example
experiments taken at the time indicated by number.
Horizontal bars denote the period of the delivery of
DHPG. Calibration: 0.5 mV, 10 msec.
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Concurrent synaptic stimulation is not required for DHPG-DEP
In all of above experiments, synaptic stimulation was administered
throughout the application of DHPG, and it cannot be concluded that
group I mGluR activation alone is sufficient to induce depotentiation. To determine whether concurrent synaptic stimulation is necessary for
the triggering DHPG-DEP, two independent pathways converging into the
same postsynaptic population of CA1 neurons were stimulated. After a
stable baseline was established for the two pathways, LTP was induced
by high-frequency TS to both pathways simultaneously. This was followed
by a brief bath application of DHPG to reverse LTP. In the control
pathway, synaptic transmission was monitored throughout the application
of DHPG, whereas the afferent stimulation in the test pathway was
terminated as soon as DHPG was applied, and the stimulation was
restarted 10 min after washout of DHPG. In the test pathway, absence of
stimulation during DHPG application was still able to reverse LTP. The
average fEPSP amplitude measured 40 min after washout of DHPG was
103.4 ± 8.5% (n = 6) of baseline, which was not
significantly different from that measured in the control pathway
(105.7 ± 8.6% of baseline; p > 0.05; paired
Student's t test) (Fig. 5).
Thus, activation of group I mGluRs alone is sufficient to induce
depotentiation without concurrent synaptic stimulation.
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Figure 5.
DHPG-DEP is synapse specific. A, An
example showing that two stimulating electrodes were used to activate
two independent groups of afferents. Field EPSPs were evoked by paired
stimulations applied at 40 msec intervals to the first or second
afferent, or both. Paired-pulse facilitation was present when
stimuli were applied twice to the same afferent (homosynaptic
facilitation) but not when the stimuli were applied to different
afferents (no heterosynaptic facilitation). B, Example
of an experiment showing that two pathways were recorded simultaneously
and LTP was induced only in the test pathway. Application of DHPG after
the induction of LTP reverses the enhancement. The superimposed fEPSP
in the inset illustrates respective recordings from
example experiments taken at the time indicated by
number. Horizontal bars denote the period
of the delivery of DHPG. Calibration: 0.5 mV, 10 msec.
C, Summary of data from six experiments performed as in
B.
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mGluR5 mediates DHPG-DEP
Because group I mGluRs include mGluR1 and mGluR5 subtypes (Anwyl,
1999), both of which are activated by DHPG, we next examined which
group I mGluR subtype mediates the DHPG-DEP. To address this issue, we
performed a series of experiments in which we analyzed the effect of
mGluR antagonists on the development of DHPG-DEP. Figure
6 summarizes our pharmacological
examination of DHPG-DEP. Initially, we examined the effect of the
broad-spectrum mGluR antagonist LY341495 (10 µM) on
DHPG-DEP. As shown in Figure 6A, when LY341495 was
applied during DHPG application, both the acute response and the
resulting depotentiation were inhibited significantly (p < 0.05; unpaired Student's t
test). The mean residual potentiation measured 40 min after DHPG
washout was 154 ± 11% (n = 8) of baseline. We
next tested the effect of the highly selective mGluR1 antagonist LY367385 (100 µM) on the DHPG-DEP. Although
LY367385 exerted a partial but significant reduction of the acute
response during DHPG application (p < 0.05;
unpaired Student's t test), it did not affect the magnitude
of DHPG-DEP (p > 0.05; unpaired Student's t test). The mean residual potentiation measured 40 min
after DHPG washout was 108.9 ± 7.8% (n = 6) of
baseline (Fig. 6B). Finally, we examined the effect
of the selective mGluR5 antagonist MPEP (10 µM)
on the DHPG-DEP. As shown in Figure 6C, MPEP also produced a
partial but significant reduction of the acute response during DHPG
application (p < 0.05; unpaired Student's
t test) and completely prevented the induction of DHPG-DEP
(p < 0.05; unpaired Student's t
test). The mean residual potentiation measured 40 min after DHPG
washout was 149 ± 11% (n = 8) of baseline. These
results suggest that the activation of the mGluR5 subtype is an
absolute requirement for the induction of DHPG-DEP at Schaffer
collateral-CA1 synapses.
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Figure 6.
Application of mGluR5 antagonist selectively
prevents the induction of DHPG-DEP. A, Summary of eight
experiments in which broad-spectrum mGluR antagonist LY341495 (10 µM) was applied 10 min before and left until the end of
DHPG application (10 µM; 5 min). Both the acute response
and the resulting depotentiation induced by DHPG were effectively
blocked by LY341495. B, Summary of six experiments
showing that the selective mGluR1 antagonist LY367385 (100 µM) did not affect DHPG-DEP, although it can partially
inhibit the acute synaptic depression by DHPG. C,
Summary of eight experiments showing that the induction of DHPG-DEP was
inhibited by a selective mGluR5 antagonist MPEP (10 µM).
The superimposed fEPSP in the inset illustrates
respective recordings from example experiments taken at the time
indicated by number. Horizontal bars
denote the period of delivery of DHPG or mGluR antagonists as
indicated. Calibration: 0.5 mV, 10 msec.
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NMDA and A1 adenosine receptor activation are not
required for DHPG-DEP
We and other groups (Larson et al., 1993; O'Dell and Kandel,
1994; Stäubli and Chun, 1996; Fujii et al., 1997; Huang et al., 1999, 2001) have shown previously that the induction of depotentiation by LFS at Schaffer collateral-CA1 synapses is dependent on the activation of NMDA and A1 adenosine receptors. It
was therefore of interest to examine the role of NMDA and
A1 adenosine receptor activation in the
development of DHPG-DEP. If activation of these receptors is critical
for DHPG-DEP, their blockade should inhibit the induction of DHPG-DEP.
To test this idea, we examined the effect of the NMDA receptor
antagonist D-APV (25 µM) and
A1 adenosine receptor antagonist DPCPX (1 µM) on the development of DHPG-DEP. We found that the
induction of DHPG-DEP was not significantly affected by either
D-APV or DPCPX pretreatment (Fig.
7). On average, the residual potentiation
measured 40 min after DHPG washout was 115.5 ± 8.2%
(n = 6; p > 0.05 when compared with
control DHPG-DEP slices; unpaired Student's t test) and
119 ± 12% (n = 6; p > 0.05 when
compared with control DHPG-DEP slices; unpaired Student's t
test) of baseline, respectively. Moreover, the induction of LTP was not
significantly affected by either D-APV or DPCPX
treatment. The slope of the fEPSP measured 50 min after LTP induction
was 153 ± 11% (n = 5; p > 0.05 when compared with control LTP slices; unpaired Student's t
test) and 158 ± 12% (n = 5; p > 0.05 when compared with control LTP slices; unpaired Student's
t test) of baseline, respectively (Fig. 7). These results
suggest that the induction of DHPG-DEP is not attributable to the
activation of either NMDA or A1 adenosine
receptors.
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Figure 7.
NMDA and A1 adenosine receptor
activation are not required for DHPG-DEP. A, Summary of
experiments showing that neither DHPG-DEP (n = 6)
nor LTP (n = 5) was affected by the NMDA receptor
antagonist D-APV (25 µM) that had been
applied immediately after tetanic stimulation (TS).
B, Summary of experiments showing that A1
adenosine receptor antagonist DPCPX (1 µM) did not affect
the induction of LTP (n = 5) or DHPG-DEP
(n = 6). The superimposed fEPSP in the
inset illustrates respective recordings from example
experiments taken at the time indicated by number.
Horizontal bars denote the period of the delivery of
DHPG, D-APV, or DPCPX as indicated. Calibration: 0.5 mV, 10 msec.
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PLC signaling is not required for DHPG-DEP
Because mGluR5 is coupled to Gq-protein,
which activates PLC and therefore can activate phosphoinositide
hydrolysis (Anwyl, 1999), we explored the possibility that an
mGluR5-mediated activation of PLC is essential for triggering DHPG-DEP.
To test this idea, we directly examined the effect of U73122, a
broad-spectrum PLC blocker, on the induction of DHPG-DEP. A recent
study has reported that U73122, when applied at a dose of 10 µM, specifically blocked PLC
1-dependent cationic
conductance elicited by DHPG in the hippocampal CA3 neurons (Chuang et
al., 2001). We therefore examined this dose of U73122 on the induction
of DHPG-DEP. As shown in Figure 8,
application of U73122 (10 µM) alone failed to affect the
LTP induction and did not affect the degree of DHPG-DEP. The slope of
fEPSP measured 50 min after LTP induction was 152.6 ± 8.2%
(n = 6; p > 0.05 when compared with
control slices; unpaired Student's t test) of baseline.
Moreover, the residual potentiation measured 40 min after DHPG washout
was 107.3 ± 7.2% (n = 7; p > 0.05 when compared with control DHPG-DEP slices; unpaired Student's t test). These results indicate that mGluR5-mediated
DHPG-DEP is not attributable to the activation of PLC signaling.
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Figure 8.
DHPG-DEP is independent of the PLC signaling
pathway. Shown is a summary of experiments (n = 6)
in which a broad-spectrum PLC blocker U73122 (10 µM) was
applied previous to the application of TS and DHPG (10 µM; 5 min). Note that U73122 had no consistent effect on
the induction of either LTP or DHPG-DEP. The superimposed fEPSP in the
inset illustrates respective recordings from example
experiments taken at the time indicated by number.
Horizontal bars denote the period of the delivery of
DHPG or U73122. Calibration: 0.5 mV, 10 msec.
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Protein phosphatase activation is not required for DHPG-DEP
Because hippocampal CA1 LTP is generally thought to be caused, at
least in part, by the activation of several postsynaptic protein
kinases, including
Ca2+/calmodulin-dependent protein kinase
II, protein kinase C (PKC), mitogen-activated protein kinase, protein
kinase A, and Src family protein tyrosine kinases (Bliss and
Collingridge, 1993; Malenka and Nicoll, 1999; Sanes and Lichtman,
1999), it is therefore reasonable to suggest that DHPG may elicit
depotentiation by interfering with the protein phosphorylation
processes required for LTP, perhaps by activating PP-coupled cascades.
This proposal was supported by our previous observation showing that
pretreatment of the hippocampal slices with serine/threonine-PP
inhibitors, okadaic acid or calyculin A, prevents the LFS-induced
depotentiation at Schaffer collater-CA1 synapses (Huang et al., 1999,
2001). If PP is critical for DHPG-DEP, blockade of PP activity should
inhibit the induction of depotentiation by DHPG. To test this idea, we
examined the effect of potent PP1/2A inhibitors, okadaic acid and
calyculin A, on the development of DHPG-DEP. We found that the degree
of DHPG-DEP was not significantly affected by a 2-4 hr preincubation
of hippocampal slices with either okadaic acid (1 µM) or
calyculin A (1 µM). On average, the residual potentiation
measured 40 min after washout of DHPG was 112.7 ± 8.6%
(n = 6; p > 0.05 when compared with
control DHPG-DEP slices; unpaired Student's t test) and
109.5 ± 7.9% (n = 6; p > 0.05 when compared with control DHPG-DEP slices; unpaired Student's t test) of baseline, respectively (Table
1). Moreover, these treatments alone
failed to affect the LTP induction. The fEPSP slope measured 50 min
after LTP induction was 165 ± 11% (n = 5; p > 0.05 when compared with control LTP slices;
unpaired Student's t test) and 162.6 ± 9.1%
(n = 5; p > 0.05 when compared with
control LTP slices; unpaired Student's t test) of baseline,
respectively (Table 1).
To examine the possible contribution of PP2B (calcineurin) to
depotentiation, slices were preincubated for 2-4 hr in FK506 (10 µM). We found that FK506 possessed no significant effect
on DHPG-DEP; the degree of depotentiation did not differ from the control slices. The mean residual potentiation measured 40 min after
washout of DHPG was 112.6 ± 9.3% (n = 5;
p > 0.05 when compared with control DHPG-DEP slices;
unpaired Student's t test) of baseline (Table 1, Fig.
9B). Pretreatment of the
slices with another inhibitor of PP2B, cyclosporin A (250 µM), was similarly ineffective on DHPG-DEP. On
average, the residual measured 40 min after washout of DHPG was
116 ± 11% (n = 4; p > 0.05 when
compared with control DHPG-DEP slices; unpaired Student's t
test) of baseline (Table 1). Additionally, the induction of LTP was not
significantly affected by either FK506 or cyclosporin A pretreatment.
The slope of the fEPSP measured 50 min after LTP induction was
152.3 ± 9.4% (n = 5; p > 0.05 when compared with control LTP slices; unpaired Student's t
test) and 158.8 ± 8.9% (n = 6; p > 0.05 when compared with control LTP slices; unpaired Student's
t test) of baseline, respectively (Table 1). These results
suggest that the induction of DHPG-DEP does not require the activation
of PP1/2A or PP2B.
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Figure 9.
DHPG-DEP is dependent on protein synthesis but not
mRNA synthesis. A, Example of experiments showing that
preincubation of slices with the protein synthesis inhibitor
cycloheximide (60 µM; 30-60 min) blocked the induction
of DHPG-DEP but not LTP. B, Summary of data from
experiments performed as in A (LTP,
n = 5; DHPG-DEP, n = 6).
C, Example of experiments showing that preincubation of slices
with another protein synthesis inhibitor, anisomycin (20 µM; 1 hr), also blocked the induction of DHPG-DEP
(n = 6) but not LTP (n = 5).
D, Summary of data from experiments performed as in
C (LTP, n = 5; DHPG-DEP,
n = 6). E, Example of experiments
showing that preincubation of slices with the transcription inhibitor
actinomycin D (25 µM; 30-60 min) had no effect on the
induction of either DHPG-DEP (n = 6) or LTP
(n = 5). F, Summary of data from
experiments performed as in E (LTP,
n = 5; DHPG-DEP, n = 6).
G, Histogram comparing the effect of transcription and
translation inhibitors on the DHPG-DEP. Asterisks represent
a significant difference from control group (p < 0.05). The magnitude of depotentiation was calculated by comparing the
synaptic response at 40 min after washout of DHPG from the experiments
illustrated in B, D, and F
with the magnitude of LTP at 48 min from the experiments illustrated in
B, D, and F. Because
both sets of data have variance, it is not possible to calculate an SE
of these depotentiation values. The superimposed fEPSP in the
inset illustrates respective recordings from example
experiments taken at the time indicated by number.
Horizontal bars denote the period of the delivery of
DHPG. Calibration: 0.5 mV, 10 msec.
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DHPG-DEP requires rapamycin-sensitive mRNA
translation signaling
Previous studies have shown that a rapid synthesis of dendritic
protein is primarily necessary for the induction of hippocampal CA1
DHPG-LTD because its induction can be blocked by translation inhibitors
(Huber et al., 2000, 2001). To explore the possible contribution of
mRNA translation to the DHPG-DEP, we performed a series of experiments
in which we analyzed the effect of two translation inhibitors,
cycloheximide and anisomycin, on the development of DHPG-DEP. As
expected, preincubation of the hippocampal slices in the cycloheximide
(60 µM) for 30-60 min blocked the induction of DHPG-DEP
(Fig. 9A). The mean residual measured 40 min after washout
of DHPG was 155 ± 12% (n = 6; p < 0.05 when compared with control DHPG-DEP slices; unpaired Student's
t test) of baseline (Fig. 9B). In addition,
pretreatment of anisomycin (20 µM) for 1 hr
also prevented the induction of DHPG-DEP (Fig. 9C). In
anisomycin pretreatment slices, the mean residual measured 40 min after
washout of DHPG was 145 ± 13% (n = 6;
p < 0.05 when compared with control DHPG-DEP slices;
unpaired Student's t test) of baseline (Fig. 9D). In contrast, the induction of LTP was not significantly
affected by either cycloheximide or anisomycin pretreatment. The slope of fEPSP measured 50 min after LTP induction was 155 ± 11%
(n = 5; p > 0.05 when compared with
control LTP slices; unpaired Student's t test) and 151 ± 10% (n = 5; p > 0.05 when compared with control LTP slices; unpaired Student's t test) of
baseline, respectively) (Fig.
9B,D). To examine the possible
contribution of mRNA synthesis to the DHPG-DEP, we examined the effect
of transcription inhibitor actinomycin D on DHPG-DEP. In contrast to
the translation inhibitors, actinomycin D (25 µM) pretreatment for 30-60 min had no effect
on the development of either LTP or DHPG-DEP (Fig. 9E). On
average, the residual potentiation measured 40 min after washout of
DHPG was 107 ± 10% (n = 6; p > 0.05 when compared with control DHPG-DEP slices; unpaired Student's
t test) of baseline. Moreover, the fEPSP slope measured 50 min after LTP induction was 149 ± 12% (n = 5;
p > 0.05 when compared with control LTP slices;
unpaired Student's t test) of baseline (Fig.
9F). Comparison of the effect of translation and
transcription inhibitors on DHPG-DEP was summarized in Figure
9G, in which the magnitude of depression measured 40 min
after washout of DHPG was calculated relative to the magnitude of LTP
measured at 48 min from each experiments. As shown in Figure 9G, the induction of DHPG-DEP was specifically prevented by
translation inhibitors. These results suggest that like DHPG-LTD, an
mRNA translation signaling is required in the establishment of DHPG-DEP at Schaffer collateral-CA1 synapses.
A recent study has characterized a rapamycin-sensitive translational
signaling pathway that regulates the translation of a specific subclass
of mRNAs and contributes to late-phase LTP and BDNF-induced synaptic
potentiation (Tang et al., 2002). We therefore conducted experiments to
assess the possible involvement of the rapamycin-sensitive pathway in
translation-dependent DHPG-DEP. As shown in Figure
10A, application of
rapamycin (200 nM) alone failed to affect the LTP
induction but effectively prevented the induction of DHPG-DEP. The
slope of fEPSP measured 50 min after LTP induction was 148 ± 13%
(n = 5; p > 0.05 when compared with control slices; unpaired Student's t test) of baseline. The
residual potentiation measured 40 min after DHPG washout was 143 ± 12% (n = 5; p < 0.05 when compared
with control DHPG-DEP slices; unpaired Student's t test).
In contrast, application of FK506, a drug that has a similar structure
and can bind to the same FKBP12 receptor as rapamycin but does not
inhibit the target of rapamycin (Raught et al., 2001), did not affect
the magnitude of either LTP or DHPG-DEP (Fig. 10B,
Table 1). These results demonstrate a role for rapamycin-sensitive mRNA
translation signaling in the development of DHPG-DEP.
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Figure 10.
A rapamycin-sensitive signaling pathway
contributes to DHPG-DEP. A, Summary of experiments
showing that preincubation of slices with rapamycin (200 nM; 2-4 hr) blocked the induction of DHPG-DEP
(n = 5) but not LTP (n = 5).
B, Summary of experiments showing that preincubation of
slices with FK506 (10 µM; 2-4 hr) did not affect either
LTP (n = 5) or DHPG-DEP (n = 5). The superimposed fEPSP in the inset illustrates
respective recordings from example experiments taken at the time
indicated by number. Horizontal bars
denote the period of the delivery of DHPG. Calibration: 0.5 mV, 10 msec.
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Postsynaptic expression of DHPG-DEP
We further characterized the location of DHPG-DEP expression
(presynaptic or postsynaptic) by testing its effect on PPF. When the
excitatory afferents to the central neurons are activated twice at
intervals of tens of milliseconds between each stimulus, the response
to the second stimulus is generally facilitated in relation to the
initial stimulus. This phenomenon is called PPF and is attributed to an
increase of the amount of transmitter release to the second stimulus
(Zucker, 1989). Thus, manipulations that affect transmitter release may
interact strongly with PPF. In agreement with previous studies (Manabe
et al., 1993), we found that LTP expression at Schaffer collateral-CA1
synapses was not associated with a significant change in PPF. Although
the reduction in PPF was significant during the initial minute after
tetanic stimulation, there was no change in PPF during the stable
expression of LTP. On average, the PPF ratio 50 min after LTP
induction was 1.48 ± 0.06, which was not significantly different
from that of the control baseline values (1.54 ± 0.05;
n = 6; p > 0.05; paired Student's
t test) (Fig.
11A). In DHPG-DEP
slices, the acute response of DHPG was associated with an increase in
PPF ratio from 1.52 ± 0.06 to 1.94 ± 0.06 (n = 6; p < 0.05; paired Student's
t test), but the resulting depotentiation had no change in
PPF (Fig. 11B). The average PPF ratio measured 40 min
after washout of DHPG was 1.55 ± 0.05, which was not
significantly different from the baseline PPF ratio (1.52 ± 0.06;
n = 6; p > 0.05; paired Student's
t test). These results suggest that the expression of
DHPG-DEP at Schaffer collateral-CA1 synapses is postsynaptic.
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Figure 11.
DHPG-DEP has no effect on PPF.
A, Summary of six experiments showing that the
expression of LTP was not associated with any change in PPF.
B, Summary of six experiments showing that the
expression of DHPG-DEP was also not associated with any change in PPF,
although the acute synaptic depression was accompanied by a significant
increase in PPF. The superimposed fEPSP in the top
portion illustrates respective recordings from example
experiments taken at the time indicated by number.
Horizontal bars denote the period of the delivery of
DHPG.
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DHPG-DEP is associated with a reduction in the surface expression
of AMPA receptors
Over recent years, evidence has accumulated that rapid changes in
the surface expression of postsynaptic ionotropic glutamate receptors
are responsible for the expression of long-term synaptic plasticity in the brain (Isaac et al., 1995; Liao et al., 1995; Hayashi et al., 2000; Carroll et al., 2001). In addition, a recent report indicates that expression of DHPG-LTD is associated with an
internalization of both AMPA and NMDA receptors from the surface of
hippocampal neurons (Snyder et al., 2001). We therefore examined the
effect of DHPG-DEP on the surface expression of AMPA receptors. To
examine membrane surface expression of AMPA receptors, we adapted a
biochemical surface biotinylation technique commonly used in cultured
cells for use in the hippocampal slices (Chung et al., 2000; Snyder et
al., 2001). Figure 12 shows that a
substantial increase in surface expression of AMPA receptors is evident
in LTP-established slices (128.5 ± 5.5% of control slices;
n = 6; p < 0.05 when compared with the
control slices; paired Student's t test). This result is
consistent with that reported by other investigators (Shi et al., 1999;
Hayashi et al., 2000). In DHPG-DEP slices, we found that there was a
significant reduction of surface expression of AMPA receptors
(105.8 ± 3.6% of control slices; n = 7;
p < 0.05 when compared with the LTP slices; paired
Student's t test). However, application of DHPG alone
without previous LTP induction also did not cause a significant
reduction of surface expression of AMPA receptors (96.9 ± 4.5%
of control slices; n = 6; p > 0.05 when compared with the control slices; paired Student's t
test). These results strongly support the notion that the expression of
DHPG-DEP is attributable to a reversal of LTP-associated increase in
the surface expression of AMPA receptors.
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Figure 12.
DHPG-DEP is accompanied by a loss of
surface GluR1. A, Representative blot showing the total
and biotinylated surface GluR1 from control (Con), LTP,
DHPG-DEP, and DHPG treatment slices. B, Densitometric
quantification revealed the change in surface GluR1 levels after LTP,
DHPG-DEP, and DHPG treatment (n = 6). Note that the
expression of DHPG-DEP is associated with a reduction in the increase
of the surface expression of GluR1 seen with LTP.
*p < 0.05 when compared with the control slices;
# p < 0.05 when compared with the LTP
slices.
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Synaptic release of glutamate activates mGluR5 to reverse LTP
The results thus far were obtained with exogenous application of
DHPG, and it is unclear whether the release of glutamate from Schaffer
collateral afferents can effectively activate mGluR5 to elicit
depotentiation. In the final set of experiments, we adapted a recently
reported protocol to test this hypothesis (Kemp and Bashir, 1999). As
shown in Figure 13A,
delivery of paired-pulse stimulation (50 msec interstimulus interval)
at 1 Hz for 15 min in the presence of D-APV (50 µM) produced a stable form of LTD. The slope of
fEPSP measured 30 min after PP-LFS was 77.9 ± 7.6% (n = 6; p < 0.05; unpaired Student's
t test) of baseline. In these slices, no LTD was evoked in
the presence of MPEP (10 µM) during PP-LFS
application. The slope of fEPSP measured 30 min after PP-LFS was
96.5 ± 7.3% (n = 6; p > 0.05;
unpaired Student's t test) of baseline (Fig.
13B). PP-LFS applied at 3 min after LTP induction, like DHPG
application, completely reversed LTP (Fig. 13C). The mean
residual potentiation measured 40 min after the end of PP-LFS was
107.2 ± 8.4% (n = 6) of baseline (Fig.
13D). Moreover, this PP-LFS-induced depotentiation was
completely prevented by MPEP (10 µM) during
PP-LFS application (Fig. 13E). The mean residual potentiation measured 40 min after the end of PP-LFS was 158 ± 12% (n = 6; p < 0.05 when compared
with PP-LFS-DEP slices; unpaired Student's t test) of
baseline (Fig. 13F). These results suggest that the
long trains of paired-pulse stimulation of Schaffer collateral afferents that cause the synaptic release of glutamate are adequate to
activate mGluR5 to trigger depotentiation.
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Figure 13.
Paired-pulse low-frequency stimulation
(PP-LFS) induces mGluR5-dependent LTD and
depotentiation. A, Example of an experiment showing that
delivery of PPF-LFS (50 msec interstimulus interval; 1 Hz/15 min) in
the presence of D-APV (50 µM) produced a
stable form of LTD at naive synapses. Pretreatment of the slices with
the mGluR5 antagonist MPEP (10 µM) blocked the induction
of LTD with PP-LFS. B, Summary of data from six
experiments performed as in A. C, Example
of an experiment showing that PP-LFS can induce depotentiation at
previous potentiated synapses in the presence of D-APV (25 µM). D, Summary of data from six
experiments performed as in C. E, Example
of an experiment showing that PP-LFS-induced depotentiation was
completely prevented by MPEP (10 µM) during PP-LFS
application. F, Summary of data from six experiments
performed as in E. The superimposed fEPSP in the
inset illustrates respective recordings from example
experiments taken at the time indicated by number.
Horizontal bars denote the period of the delivery of
PP-LFS, D-APV, or MPEP. Calibration: 0.5 mV, 10 msec.
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DISCUSSION |
In the present study, we have shown that activation of mGluR5,
either by a brief application of the selective group I mGluR agonist
DHPG or with synaptically released glutamate (PP-LFS), can trigger a
novel form of DEP by a mechanism that requires a de novo
protein synthesis from existing mRNA. This mGluR5-mediated depotentiation is (1) reversible, (2) able to unsaturate LTP, (3) time
dependent, (4) induced postsynaptically, (5) mechanistically distinct
from NMDAR-DEP, (6) independent of PLC signaling, and (7) not
associated with PP activation. In addition, the expression of DHPG-DEP
is caused, at least partly, by a rapid removal of AMPA receptors from
the postsynaptic membrane.
In addressing the locus of expression of synaptic plasticity, a change
in PPF is frequently used as an indication of presynaptic expression.
As in the case for CA1 LTP (Manabe et al., 1993), the expression of
DHPG-DEP caused no significant change in PPF (Fig. 11), which strongly
suggests that DHPG-DEP is attributed to postsynaptic mechanisms.
However, a recent report by Watabe et al. (2002) showed that a
combination of postsynaptic induction and presynaptic expression is
involved in DHPG-LTD in the hippocampal CA1 region. The reason that we
do not observe the involvement of a presynaptic mechanism in the
expression of DHPG-DEP is not clear but could be attributable partly to
the use of different doses (10 vs 100 µM) and time scales
of DHPG challenges (5 vs 10 min), resulting in activation of different
cellular processes that may vary in their mode of action. In contrast,
the DHPG-induced acute depression of synaptic transmission is expressed
presynaptically, because this depression was accompanied by a marked
increase in PPF. Although it is difficult, on the basis of the present
data, to estimate how DHPG acts presynaptically to reduce transmitter release and acts postsynaptically to induce DEP, a plausible
explanation of this finding may be that DHPG acts on a different
subtype of group I mGluRs to regulate CA1 pyramidal cell function. This
idea is supported by the finding that the induction of DHPG-DEP was specifically blocked by the mGluR5 antagonist MPEP but not by mGluR1
antagonist LY367385 (Fig. 6), although both MPEP and LY367385 can
partially inhibit the DHPG-induced acute synaptic depression. Taking
all of these results together, we conclude that DHPG acts on
postsynaptic mGluR5 to induce DEP and acts on presynaptic mGluR1 to
reduce transmitter release. These observations are in agreement with
recent anatomy data demonstrating that mGluR5 is the most abundant
group I mGluR in CA1 pyramidal cells, whereas mGluR1 is not detectable
(Baude et al., 1993; Romano et al., 1995).
How might mGluR5 activation lead to DEP? Although it is known that
mGluR5 couples to PLC and therefore can activate PKC and mobilize
Ca2+ from intracellular stores (Anwyl,
1999), it is unlikely that this conventional signal transduction
mechanism underlies DHPG-DEP. Indeed, we could not inhibit the
induction of DHPG-DEP by applying PLC blocker U73122 (Fig. 8). It
appears, however, that a novel protein synthesis signaling pathway is
involved in the induction of DHPG-DEP. Evidence supporting this is that
preincubation of the hippocampal slices with the protein synthesis
inhibitors, cycloheximide or anisomycin, but not the transcriptional
inhibitor, actinomycin D, effectively prevents the induction of
DHPG-DEP (Fig. 9). Such a result is in line with the observation that
new protein synthesis is stimulated by group I mGluR activation in hippocampal synaptoneurosomes (Weiler et al., 1997; Angenstein et al., 1998). Recent studies have also demonstrated that rapid dendritic protein synthesis is required for the induction of DHPG-LTD (Huber et al., 2000, 2001) or DHPG-mediated facilitation of the persistence of an intermediate phase of LTP (Raymond et al., 2000) in
the CA1 region of hippocampal slices. Although the signal transduction events that couple mGluR5 activation to protein synthesis control were
not examined here, a rapamycin-sensitive signaling pathway seems to
contribute to this process, because disruption of this signaling
pathway with rapamycin inhibits the expression of DHPG-DEP (Fig. 10).
Although a number of rapamycin-sensitive gene transcripts have been
characterized to date, including those encoding ribosomal proteins (S3,
S6, S14, and S24), translation initiation factors (eIF-4E, 4E-BP1, and
4E-BP2), and translation elongation factors (eEF1A and eEF2) (Brown and
Schreiber, 1996; Raught et al., 2001; Steward and Schuman, 2001; Tang
et al., 2002), the specific component of this signaling pathway
participating in the development of DHPG-DEP remains to be established.
Activity-dependent changes in AMPA receptor trafficking have been
proposed recently to play an important role in bidirectional synaptic
plasticity (Malinow et al., 2000; Carroll et al., 2001). For example,
it has been known that LTP induction in hippocampal slices is
associated with an increase in the delivery of AMPA receptors to
dendritic spines (Shi et al., 1999, 2001; Hayashi et al., 2000),
whereas the expression of LTD is associated with a decrease in the
surface expression of AMPA receptors (Kandler et al., 1998; Luthi et
al., 1999). In addition, with the use of biochemical analysis, we have
extended these findings by showing that the expression of DHPG-DEP is
associated with a reduction in the increase of the surface expression
of AMPA receptors seen with LTP (Fig. 12). This finding clearly
indicates that removal of synaptic AMPA receptors is a candidate
mechanism for the expression of DHPG-DEP in the hippocampus. Similarly,
it has been found that cerebellar (Wang and Linden, 2000) and
hippocampal CA1 (Snyder et al., 2001) LTD, which are also triggered by
activation of group I mGluRs, require postsynaptic internalization of
AMPA receptors. What mechanism might give rise to the removal of AMPA
receptors from synapses during DHPG-DEP? Calcineurin (PP2B) was found
to be involved in promoting AMPA receptor endocytosis in response to
application of insulin (Lin et al., 2000) or AMPA (Beattie et al.,
2000), suggesting that this signaling pathway might be widely involved
in the internalization of AMPA receptors. However, we found that DHPG
still induced depotentiation by applying the selective PP2B inhibitors,
FK506 and cyclosporin A (Fig. 10, Table 1), indicating that this
calcineurin-dependent signaling pathway is not involved in
DHPG-mediated regulation of AMPA receptor endocytosis in the
hippocampal slices. Furthermore, previous work has reported that PP1/2A
inhibitors, okadaic acid and calyculin A, can facilitate DHPG-LTD in
the hippocampal CA1 region (Schnabel et al., 2001), suggesting that
PP1/2A might have an important, active role in governing the processes
underlying DHPG-LTD. However, we found that neither okadaic acid nor
calyculin A treatment significantly influences DHPG-DEP. This result
argues that a PP1/2A-dependent signaling process is involved in the
generation of DHPG-DEP. The basis for the difference in the present
results and those of Schnabel et al. (2001) is uncertain. However, it
is unlikely that our treatment protocols were ineffective in blocking
PP1/2A activity because the same treatment protocols used in our
previous studies are found to be sufficient to prevent the LFS-induced
DEP at the same synapses (Huang et al., 1999, 2001).
One of the intriguing aspects of DEP is that the magnitude of DEP is
inversely proportional to the time lag of depotentiating stimulation
after LTP induction (Fujii et al., 1991; O'Dell and Kandel, 1994).
TOP
ABSTRACT
INTRODUCTION
