Department of Physiology and Pharmacology, State University of New York Health Science Center at Brooklyn, Brooklyn, New York 11203
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ABSTRACT |
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Bianchi, Riccardo,
Steven R. Young, and
Robert K. S. Wong.
Group I mGluR activation causes voltage-dependent and -independent
Ca2+ rises in hippocampal pyramidal cells.
Application of the metabotropic glutamate receptor (mGluR) agonist
(1S,3R)-1-aminocyclopentane-1,3-dicarboxylic acid
(ACPD) or the selective group I mGluR agonist
(S)-3,5-dihydroxyphenylglycine (DHPG) depolarized both CA3
and CA1 pyramidal cells in guinea pig hippocampal slices. Simultaneous
recordings of voltage and intracellular Ca2+ levels
revealed that the depolarization was accompanied by a biphasic
elevation of intracellular Ca2+ concentration
([Ca2+]i): a transient calcium rise followed
by a delayed, sustained elevation. The transient
[Ca2+]i rise was independent of the membrane
potential and was blocked when caffeine was added to the perfusing
solution. The sustained [Ca2+]i rise appeared
when membrane depolarization reached threshold for voltage-gated
Ca2+ influx and was suppressed by membrane
hyperpolarization. The depolarization was associated with an increased
input resistance and persisted when either the transient or sustained
[Ca2+]i responses was blocked. mGluR-mediated
voltage and [Ca2+]i responses were blocked by
(+)--methyl-4-carboxyphenylglycine (MCPG) or
(S)-4-carboxy-3-hydroxyphenylglycine (4C3HPG). These data
suggest that in both CA3 and CA1 hippocampal cells, activation of group
I mGluRs produced a biphasic accumulation of
[Ca2+]i via two paths: a transient release
from intracellular stores, and subsequently, by influx through
voltage-gated Ca2+ channels. The concurrent mGluR-induced
membrane depolarization was not caused by the
[Ca2+]i rise.
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INTRODUCTION |
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Recent studies show that metabotropic
glutamate receptor (mGluR) activation elicits hyperpolarization
(Jaffe and Brown 1994; Shirasaki et al.
1994
) and/or depolarization in hippocampal pyramidal cells
(Charpak et al. 1990
; Congar et al. 1997
;
Crépel et al. 1994
; Guérineau et al.
1994
, 1995
; Pozzo Miller et al.
1995
, 1996
; Shirasaki et al.
1994
). There has been considerable interest in elucidating the
second-messenger mechanisms for the mGluR-induced membrane responses.
Initial studies indicate the involvement of G-proteins in these
responses (Congar et al. 1997
; Pozzo Miller et
al. 1995
; Shirasaki et al. 1994
; but see
Guérineau et al. 1995
). In particular, the role of
intracellular Ca2+ in mediating mGluR-induced membrane
responses has been evaluated, because this ion is a second messenger
for one class of mGluRs (see below). One study reports that
intracellular Ca2+ directly sustains an mGluR-mediated
depolarization by activating a nonspecific cation current in CA1
hippocampal neurons (Congar et al. 1997
). However, other
studies performed in CA3 pyramidal cells suggest that mGluR-induced
depolarizations do not require an intracellular Ca2+ rise
(Guérineau et al. 1995
; Pozzo Miller et al.
1995
). In CA1 neurons it has been suggested that transient
intracellular Ca2+ increases induced by mGluR agonists
elicit hyperpolarizing currents (Jaffe and Brown 1994
;
Shirasaki et al. 1994
). It is possible that these
different findings reflect the existence of multiple components of
mGluR-induced membrane responses that are differentially expressed in
CA1 and CA3 neurons. In an attempt to clarify the causal relationship
between intracellular Ca2+ and voltage responses to mGluR
stimulation, we simultaneously recorded intracellular Ca2+
levels ([Ca2+]i) and membrane potential in
hippocampal cells.
Elevation of [Ca2+]i following mGluR
activation may occur via release from intracellular stores
(Pozzo Miller et al. 1996) or through transmembrane
influx. Stimulation of group I mGluRs activates phosphoinositide
hydrolysis and secondarily elevates [Ca2+]i
by inducing release from inositol-1,4,5-trisphosphate
(IP3)-sensitive stores (Abe et al. 1992
;
Masu et al. 1991
). Transmembrane influxes of
Ca2+ may also contribute to the mGluR-induced
[Ca2+]i rise. This influx can occur via
nonspecific cation channels (Chen et al. 1997
;
Partridge et al. 1994
) or voltage-gated Ca2+ channels.
By monitoring both the voltage and intracellular Ca2+ responses in single CA3 and CA1 pyramidal cells, we were able to address two questions: 1) What are the origin and temporal pattern of [Ca2+]i rise elicited by mGluR agonists? 2) What is the causal relationship between the [Ca2+]i rise and mGluR-induced membrane potential changes?
Parts of this study have been presented in abstract form
(Bianchi et al. 1997; Young et al. 1996
).
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METHODS |
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Slice preparation
Transverse hippocampal slices (~300 µm thick) were prepared
from adult guinea pigs as described previously (Bianchi and Wong 1995). Slices were placed submerged in a coverslip-bottomed
recording chamber and superfused at 3-5 ml/min at 30-32°C. The
composition of the perfusing control solution was (in mM) 124 NaCl, 26 NaHCO3, 5 KCl, 1.6 MgCl2, 2.0 CaCl2, and 10 D-glucose. The solution was continuously gassed with a 95% O2-5% CO2
mixture (pH 7.4). One slice at a time was placed in the chamber and was
held down by nylon threads glued to a platinum ring. The arrangement
provided sufficient mechanical stability to allow extended
intracellular recordings but did not prevent solution exchange at the
bottom surface of the slice. The chamber was mounted on the stage of a
Nikon Diaphot inverted microscope. Optical access to the slice was
through the glass coverslip at the bottom of the chamber.
Electrophysiological recordings
Intracellular recordings were performed in the pyramidal cell
layer of the CA3 and CA1 regions with sharp glass electrodes containing
potassium acetate (0.4 M) and calcium green-1 (0.5 mM). In some
experiments the quaternary lidocaine derivative
N-(2,6-dimethylphenylcarbamoylmethyl)triethylammonium bromide (QX-314; 50 mM) was added to the electrode-filling solution. The electrode resistance was typically 50-80 M. Recordings in current-clamp mode were amplified (Axoclamp-2A; Axon Instruments, Foster City, CA), displayed on an oscilloscope (DSO 400; Gould, Valley
View, OH) and chart recorder (Gould TA240), and stored on FM tape
(Racal Store 4DS; Southampton, England). When recorded concurrently
with synchronized optical recordings (see Optical recordings), membrane voltage signals were filtered at
30-120 Hz for slow events [e.g., responses to
(1S,3R)-1-aminocyclopentane-1,3-dicarboxylic acid
(ACPD)] or at 1,500 Hz for faster events (e.g., responses to
intracellular current pulse injections), and stored directly in a
computer. Intracellular square-wave current pulses were timed by a
digital stimulator (PG 4000; Neuro Data Instruments, New York, NY).
Hyperpolarizing pulses (
0.1 to
1.0 nA, 150-300 ms) were applied
throughout an experiment to monitor the condition of the cell membrane
and to adjust the bridge balance (e.g., Fig. 12A).
Optical recordings
Changes in [Ca2+]i were detected by
filling cells with calcium green-1 (0.5 mM; C3010, Molecular Probes,
Eugene, OR) from the intracellular recording electrodes. The dye was
injected by passing 350-ms, 0.5-nA current pulses at 50% duty cycle
for 20-25 min. Epi-illumination was provided by a 150 W xenon short
gap bulb (XBO 150W CR OFR; Osram, Germany) linked to the microscope by a fused silica fiberoptic bundle (77578; Oriel, Stratford, CT). Filled
cells were viewed through a Nikon ×10 Fluor objective (n.a. = 0.5) and
standard fluorescein filter set (Nikon B-2E). The fluorescent images
shown in Figs. 1A and
2B were divided into three
and four areas, respectively, and gray-level scales were independently adjusted for the different regions. This was necessary to display both
dendrites and soma, which had widely different fluorescence intensities. The computer program used for simultaneous acquisition of
calcium green-1 fluorescence and transmembrane potential has been
described previously (Lasser-Ross et al. 1991
).
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Fluorescence images were acquired from a thermoelectrically cooled
charge-coupled device camera (CH230; Photometrics, Tucson, AZ) run in
frame-transfer mode. A software-driven timer/pulser (Master-8; AMPI,
Israel) provided for precise synchronization of optical and electrical
recordings. Fluorescence values were measured and averaged over an area
of interest selected within the fluorescence image. Relative
intracellular Ca2+ levels in the areas of interest were
expressed as change in fluorescence divided by the resting fluorescence
(F/F). The applied equation was
F/F (%) = 100 × (Fi
Fb)/(Fb
Fa), where Fi is the
average fluorescence of the area of interest in each image,
Fb is the baseline fluorescence of the area of
interest averaged over at least 10 images before stimulation, and
Fa is the autofluorescence measured at an
equivalent location in the slice at which no stimulus-associated signal
was detected. Normalized in this way, the dye fluorescence is a
monotonic function of the concentration of intracellular Ca2+ (Callaway et al. 1993
). To reduce
bleaching and phototoxic damage to the cell, neutral density filters
were introduced in the excitation light pathway. In a few cases
(n = 6 of 76 cells), traces were corrected for
bleaching (2-7% of baseline fluorescence over 5-6 min) by
subtracting a recording made without stimulation (e.g., Fig.
6B). Cells were accepted for optical recordings only when their fluorescence responses (
F/F) to brief
trains of action potentials were >10%.
Consistent with previous reports, action potentials elicited
Ca2+ increases that were substantially larger in the
proximal apical and basal dendrites than in somata (Fig. 2) (cf.
Jaffe et al. 1994; Regehr and Tank 1992
).
Due to light scattering from intervening tissue, most of the cells we
recorded were too deep within a slice to clearly distinguish dendritic
signals. Thus, in this paper we report Ca2+ signals
measured in somata (except for Fig. 2). In dye-filled cells near the
slice surface, however, dendrites were clearly visualized, and
dendritic responses to ACPD were also apparent (e.g., Fig. 2).
Pharmacological agents
Agonists of mGluRs were injected directly into the bath via a narrow tube from a syringe (pulse application). A pulse application lasted on average 7.1 ± 0.6 s (mean ± SE; range, 2.5-12.0 s; n = 20). Antagonists of mGluRs were added to the perfusate (bath application). For pulse applications, 100 µl of 2 mM ACPD (or 1 mM DHPG) was typically used. We attempted to characterize the time course of the agonist concentration after a pulse injection by applying 3 M KCl via the syringe tubing and monitoring the junction potential change in the recording pipette. The average time-to-peak of the potential change was 19.1 ± 1.2 s (range, 7-29 s) from the start of the injection and the half-amplitude duration (see Data analysis) was 208.5 ± 25.4 s (range, 23-487 s; n = 17). During the time required to reach peak agonist concentration at the cell (7-29 s), the bolus of 100 µl of 2 mM ACPD was diluted by perfusing solution that continued to flow into the chamber. The dilution can be estimated by considering the volume of the bath (0.9-1 ml) and a volume of solution of 0.3-2.4 ml perfusing in 7-29 s (at a flow rate of 3-5 ml/min), and yielded peak ACPD concentrations of 50-150 µM (25-75 µM for DHPG). Caffeine was either bath-applied (2-5 mM; e.g., Fig. 8, A and B) or pulse-applied (estimated concentration, 2-5 mM; e.g., Fig. 8C). Agonist applications to the same cell were separated by intervals of 30-40 min to allow complete recovery of the responses. In most experiments tetrodotoxin (0.3-0.6 µM) was present in the perfusate during the recordings.
ACPD (50-150 µM), (S)-3,5-dihydroxyphenylglycine (DHPG;
25-75 µM), (+)--methyl-4-carboxyphenylglycine (MCPG; 0.5-1 mM),
and (S)-4-carboxy-3-hydroxyphenylglycine (4C3HPG; 0.5 mM)
were purchased from Tocris Cookson (Ballwin, MO); QX-314 was obtained
from Research Biochemical International (Natick, MA); all the other
chemicals were from Sigma (St. Louis, MO).
Data analysis
Data were analyzed using pClamp (Axon Instruments), Transform (Fortner Research LLC, Sterling, VA), and SigmaPlot (SPSS, Chicago, IL) software. Figures 1A and 2A are montages of different display scales applied to single images of CA3 pyramidal neurons filled with calcium green-1 (Photoshop, Adobe Systems, San Jose, CA). This is required for the display of unnormalized images because of the vastly greater dye-containing volume of the cell body compared with the dendrites.
Input resistance (Rin) was calculated from the amplitudes of the voltage responses divided by the intensity values of the injected current steps. In five cells (Fig. 9B), Rin was calculated as the slope of voltage-current (V-I) plots obtained from responses to depolarizing ramp current injections. Half-amplitude durations were measured from the time the rising phase of a response crossed one half the peak amplitude until the decay phase crossed the same line. Measured parameters are reported as means ± SE. Student's t-tests were applied for statistical comparisons, and differences were considered significant when P < 0.05.
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RESULTS |
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Intracellular recordings were performed on 73 CA3 and 20 CA1
pyramidal neurons, all with overshooting action potentials. The average
resting membrane potential (Vrest) and input
resistance (Rin) of CA3 cells were 62.7 ± 0.7 (SE) mV and 49.8 ± 2.5 M
, respectively. In CA1 cells
Vrest was
63.3 ± 1.1 mV and
Rin was 36.2 ± 2.4 M
. Of all neurons,
60 CA3 and 16 CA1 cells were successfully injected with the fluorescent
dye calcium green-1 for simultaneous electrical and optical recordings
(see Fig. 1). In general, CA1 cells responded to mGluR agonists in the
same way as CA3 cells, and the main conclusions of this paper apply to
both regions. Where regional differences were observed, they will be
illustrated in the section on mGluR responses in CA1.
ACPD-induced intracellular voltage and calcium responses in CA3 neurons
Membrane potential and intracellular Ca2+ responses were simultaneously recorded in CA3 cells from slices superfused with control solution (n = 6). Application of ACPD (50-150 µM; n = 12) elicited depolarization and action potentials (Fig. 1). Following an initial phase of intense firing, action potentials decreased in frequency and the membrane potential gradually recovered to the resting level.
Concurrent fluorescence measurements from the soma showed that ACPD applications produced an abrupt increase in the fluorescence of calcium green-1, indicating an increase in intracellular Ca2+ concentration ([Ca2+]i). Brief increases in the Ca2+ signal occurred in phase with action potential firing (Fig. 1). In addition, there was a baseline increase in [Ca2+]i with a time course parallel to that of the depolarization.
Under control conditions, both the optical and voltage records of the ACPD-induced response were clearly dominated by action potentials. To examine the voltage and [Ca2+]i responses directly elicited by mGluR activation, tetrodotoxin (TTX, 0.3-0.6 µM) was added to the perfusing solution in all remaining experiments. TTX suppressed fast Na+-mediated action potentials and their associated voltage-dependent Ca2+ increases. Under these conditions, ACPD induced a more gradual depolarization and increase in [Ca2+]i (Fig. 2B). The increase in [Ca2+]i often occurred in two phases: an initial transient phase and a later sustained phase. The transient phase consisted of a rapid rise that peaked before the voltage response (Fig. 2B, arrow). On average, the transient Ca2+ component peaked at 31.3 ± 3.6 s after ACPD application (n = 25), whereas the depolarization peaked at 118.6 ± 9.5 s (n = 25). The transient Ca2+ component was followed by a sustained phase of Ca2+ rise (Fig. 2B). The depolarization and the sustained Ca2+ response returned to baseline levels in 5-20 min (n = 4).
ACPD-induced [Ca2+]i increases were not restricted to the soma. In cells where Ca2+ responses could be visualized in the dendrites (Fig. 2, C and D), the biphasic [Ca2+]i response induced by ACPD was observed both in the soma and in the proximal apical and basal dendrites (Fig. 2, A and B).
Characteristics of the biphasic [Ca2+]i rise
To identify the sources of [Ca2+]i rise,
we tested the voltage sensitivity of the ACPD-induced Ca2+
responses (Fig. 3). The threshold for
voltage-gated Ca2+ influx, as shown by increase in calcium
green-1 fluorescence, was determined by applying a depolarizing current
to the cell. The mean threshold for Ca2+ influx was
60.3 ± 0.7 mV (n = 29 cells; Fig.
3B, left vertical dotted line). Intracellular
Ca2+ levels were transiently depressed by hyperpolarizing
pulses applied at membrane potentials above the threshold for
Ca2+ entry.
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After determining the threshold for voltage-gated Ca2+ entry, ACPD was applied to the slice. ACPD elicited a depolarization (Fig. 3, A and B, Vm) and a biphasic [Ca2+]i increase (Fig. 3B, Ca2+). Hyperpolarizing pulses applied via the recording electrode affected the two phases of the Ca2+ response differently: the [Ca2+]i during the early transient phase was not affected, whereas the [Ca2+]i during the second phase was suppressed.
Comparison of the responses induced by current injection with those induced by ACPD showed that the initial transient Ca2+ rise occurred subthreshold to voltage-gated Ca2+ entry (below the horizontal dotted line in Fig. 3B, Vm). In contrast, the sustained phase of Ca2+ rise occurred only at membrane potentials above threshold (horizontal dotted line in Fig. 3B, Vm).
Time course of the initial transient [Ca2+]i rise
Because the transient [Ca2+]i rise was not suppressed by membrane hyperpolarization, we isolated this component by applying ACPD at increasing levels of hyperpolarization. At resting membrane potentials, the ACPD-induced Ca2+ response in some cells consisted of a continuous increase instead of a biphasic response (Fig. 4A). The transient Ca2+ component was then revealed when the cell was hyperpolarized by current injection (Fig. 4B, arrow). Upon increasing the level of hyperpolarization, complete separation of the initial transient component from the sustained component of [Ca2+]i increase was observed (Fig. 4, C and D).
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The time course of the isolated transient [Ca2+]i rise was analyzed in experiments in which ACPD was applied at hyperpolarized levels (Fig. 5; n = 5). The [Ca2+]i rise started within a few seconds after the beginning of the ACPD application, reached a peak in 10.8 ± 1.5 s, and then recovered to baseline in 59.4 ± 4.0 s (Fig. 5A, top trace; see also Fig. 5B, top graph). The half-amplitude duration of the transient [Ca2+]i rise was 26.8 ± 2.6 s.
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Pharmacology of the ACPD-induced responses
MCPG, an antagonist effective against both group I and group II mGluR-mediated responses, prevented the development of the ACPD-induced depolarization and the intracellular Ca2+ responses when added to the perfusing solution (0.5-1 mM; n = 4; Fig. 6). 4C3HPG (0.5 mM), an agonist of group II mGluRs and an antagonist of the group I mGluRs, also prevented the development of the ACPD-induced intracellular Ca2+ and voltage responses (n = 3). These results indicate that the responses activated by ACPD are mediated by activation of group I mGluRs. We used the selective group I mGluR agonist DHPG (50 µM; n = 6) to confirm the involvement of these receptors. DHPG induced depolarizations and biphasic [Ca2+]i rises. When the responses were recorded at hyperpolarized levels, DHPG elicited a transient [Ca2+]i rise similar to that elicited by ACPD (Fig. 7).
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Effect of caffeine on the voltage and [Ca2+]i responses induced by ACPD
One possible source for the voltage-independent transient
[Ca2+]i increase is release from
intracellular stores. Caffeine has been reported to affect this process
in a reversible way (Friel and Tsien 1992; Henzi
and MacDermott 1992
). We tested the effect of caffeine on the
ACPD-induced voltage and Ca2+ responses in CA3 pyramidal
cells. ACPD was applied in the absence, in the presence, and after wash
out of caffeine (bath-applied 2-5 mM; Fig.
8). In the presence of caffeine, ACPD
induced the expected depolarization and sustained
[Ca2+]i rise, but the initial transient
[Ca2+]i response was blocked [Fig.
8A, Caffeine (bath); n = 5]. The transient
[Ca2+]i response reappeared after wash out of
caffeine (Fig. 8A, Wash). These observations were confirmed
in four cells (Fig. 8B). Thus the transient
[Ca2+]i rise was not required for the
ACPD-induced depolarization. Bath application of caffeine alone
sometimes caused a small depolarization (2-6 mV; n = 3 of 5). However, when the holding current was adjusted to keep the
membrane potential constant, caffeine did not cause steady-state
changes in [Ca2+]i. We next confirmed that,
as previously reported (Garaschuk et al. 1997
;
Glaum et al. 1990
), pulse application of caffeine caused
transient Ca2+ release. Figure 8C shows that
pulse application of caffeine elicited a transient
[Ca2+]i rise similar to the transient
[Ca2+]i rise elicited by ACPD
(n = 10).
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Input resistance change underlying the ACPD-induced depolarization
Input resistances (Rin) of CA3
pyramidal neurons were assessed by current injections. Ramp current
injections were applied over ranges of 80 to
45 mV before and after
ACPD application (Fig. 9). Figure
9B shows the average voltage-current relationship (V-I plot) from five experiments. The V-I plot
suggests that in control conditions a decrease in membrane resistance
occurred at membrane potentials more depolarized than
60 mV (inward
rectification). This rectification was suppressed by ACPD. ACPD also
increased the input resistance of the cells from 67.0 ± 5.9 M
to 96.3 ± 8.4 M
(P < 0.001; n = 5; Student's t-test for paired data). The extrapolated
intersection of the regression lines for the control and the ACPD
responses was at
102.9 ± 4.0 mV, near the potassium equilibrium
potential. Simultaneous monitoring of the Ca2+ response
suggests that the threshold for voltage-gated Ca2+ entry
was not altered by ACPD (n = 4; P = 0.36; Student's t-test for paired data; Fig. 9). The
V-I plot obtained during the ACPD response also showed no
significant change in slope at the potential when
[Ca2+]i began to increase (Fig.
9B, ACPD plot).
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Intracellular voltage and calcium responses to mGluR agonists in CA1 cells
Our data indicate that the mGluR-mediated depolarization in CA3
neurons was not caused by an increase of
[Ca2+]i because neither sustained (Fig.
4D) nor transient (Fig. 8) [Ca2+]i
rise was required for the depolarization. To the contrary, studies in
CA1 cells have shown that a depolarizing response to mGluR stimulation
is sustained by a calcium-activated nonspecific cation current
(ICAN) (Congar et al. 1997;
Crépel et al. 1994
). We carried out additional
experiments in CA1 cells to evaluate whether the mGluR responses we
recorded in CA3 could also be found in CA1 and whether the sustained
depolarization in CA1 required a [Ca2+]i rise.
In all the CA1 cells examined at resting potential (n = 13), ACPD elicited [Ca2+]i increases and membrane potential changes. As in CA3 cells, the [Ca2+]i response consisted of a transient rise followed by a sustained elevation. The sustained phase of the [Ca2+]i response was suppressed by hyperpolarization (Fig. 10). The voltage response consisted of an initial hyperpolarization followed by a long-lasting depolarization (Fig. 10). DHPG elicited intracellular Ca2+ and voltage responses similar to the ACPD-induced ones (n = 4; not shown).
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The initial hyperpolarization elicited by mGluR agonists was observed in all CA1 cells (n = 16; Figs. 10 and 11), whereas it was only occasionally seen in CA3 cells (10 of 67; e.g., Fig. 8A). A comparison of the averaged transient intracellular Ca2+ response recorded in CA1 cells with that of the averaged hyperpolarizing voltage response shows that the two events peaked at approximately the same time (Fig. 11).
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The effect of membrane potential on the mGluR-mediated responses in CA1
cells was examined. ACPD was applied at different initial membrane
potentials (Fig. 12B).
However, CA1 cells were more difficult to hyperpolarize than CA3 cells,
presumably because hyperpolarization activated the "Q" current
(Fig. 12A, left), which acted to shunt the injected current.
For this reason, some experiments were performed using QX-314 in the
recording pipette to block the Q current (Perkins and Wong
1995) (Fig. 12A, right). In Fig. 12B,
increasing levels of hyperpolarization gradually suppressed the
sustained Ca2+ response. With sufficient hyperpolarization,
the sustained Ca2+ component was completely blocked,
leaving behind an isolated transient [Ca2+]i
rise, whereas the depolarization remained (Fig. 12Bc). As in CA3 cells, this depolarization far outlasted (almost 20 min; not shown)
the isolated transient [Ca2+]i rise (~10 s;
Fig. 12Bc). An additional effect of QX-314 illustrated in
Fig. 12B was the suppression of the ACPD-induced transient
hyperpolarization (n = 7).
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DISCUSSION |
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In hippocampal pyramidal cells, stimulation of group I mGluRs by ACPD or by DHPG produces a depolarization that is accompanied by an elevation of intracellular Ca2+. The intracellular Ca2+ rise consists of a transient, voltage-independent component followed by a voltage-dependent, sustained component.
Source and possible roles of the transient [Ca2+]i rise
The inability of membrane hyperpolarization to suppress the
transient [Ca2+]i rise suggests that the
source of Ca2+ is unlikely to be influx through
voltage-gated Ca2+ channels. A more likely source is
release from intracellular stores. The reversible blockade of the
transient [Ca2+]i rise by caffeine also
suggests an involvement of intracellular Ca2+ stores.
Activation of group I mGluRs has been shown to generate IP3
and to release Ca2+ from IP3-sensitive stores
(Abe et al. 1992; Masu et al. 1991
; Pozzo Miller et al. 1996
). Although caffeine and
IP3 may act on different intracellular
Ca2+-permeable channels, co-localization of these channels
in hippocampal cells has been demonstrated (Seymour-Laurent and
Barish 1995
; Sharp et al. 1993
), suggesting the
possibility that caffeine and IP3 act on the same
intracellular Ca2+ store. We showed that pulse application
of caffeine caused transient [Ca2+]i increase
(Fig. 8C). Bath application of caffeine may deplete a common
pool of releasable Ca2+ (Seymour-Laurent and Barish
1995
) and thereby prevent the occurrence of the mGluR-mediated
[Ca2+]i transient. Alternatively, a direct
antagonistic action of caffeine on IP3 receptors
(Ehrlich et al. 1994
; Parker and Ivorra
1991
; Seymour-Laurent and Barish 1995
) may be involved.
The amplitude of the ACPD-induced transient
[Ca2+]i rise was small compared with that
elicited by a burst of action potentials. In CA1 cells, however, it was
probably sufficient to activate a calcium-dependent K+
current (see below). Another possible role for the transient [Ca2+]i rise would involve initiation of
[Ca2+]i waves (Berridge 1997).
Agonist-induced release of Ca2+ from intracellular stores
can set up propagating [Ca2+]i waves
(Jaffe and Brown 1994
). Propagating
[Ca2+]i increases may provide the link
between receptor responses and protein synthesis (Berridge
1998
; Phenna et al. 1995
). Other studies from
this laboratory have shown that group I mGluR activation elicits
protein synthesis-dependent long-term changes in the activity of
hippocampal CA3 cells (Merlin et al. 1998
). It would be
interesting to test whether the transient
[Ca2+]i rise we have observed plays a role in
this phenomenon.
Sustained [Ca2+]i rise is mediated by influx through voltage-gated Ca2+ channels
The contribution of voltage-dependent Ca2+ channels to
the sustained [Ca2+]i rise is supported by
the observation that the accumulation of
[Ca2+]i during the sustained component
occurred only when the neuron was depolarized above the threshold for
voltage-dependent Ca2+ influx (see Fig. 3B).
Hyperpolarization during the sustained calcium rise to levels below the
threshold also reversibly and totally suppressed the Ca2+
accumulation (see Fig. 9A). The average threshold for
voltage-dependent [Ca2+]i increase was about
60 mV. Although low-threshold T-type Ca2+ channels are
present in hippocampal pyramidal cells (Magee et al.
1995
; Soong et al. 1993
), one might expect these
channels to be inactivated at the potentials at which our recordings
were carried out (Kavalali et al. 1997
). Sustained
low-voltage-activated Ca2+ currents have also been
described in both CA1 (Magee et al. 1996
) and CA3
(Avery and Johnston 1996
) pyramidal cells; such currents are a more likely source of the Ca2+ in the group I
mGluR-mediated sustained calcium response. Pharmacological characterization of these currents was not attempted in this study, because blockers of voltage-gated Ca2+ channels were
reported to affect the function of intracellular Ca2+
stores, presumably by preventing refilling of depleted stores (Garaschuk et al. 1997
).
Mechanisms of the mGluR-mediated membrane potential changes
In CA1 cells, ACPD elicited an initial hyperpolarization that was
associated with the transient [Ca2+]i
increase. Hyperpolarizing responses elicited by ACPD have been observed
previously in CA1 cells and were attributed to the activation of
Ca2+-dependent K+ conductances (Jaffe
and Brown 1994; Shirasaki et al. 1994
). QX-314 has been shown to suppress a component of the
Ca2+-dependent K+ current (Oda et al.
1992
). Thus the block of the hyperpolarization by QX-314 (Fig.
12) is consistent with the notion that this response was caused by a
Ca2+-dependent K+ current triggered by the
transient [Ca2+]i rise. The mGluR-mediated
hyperpolarizing responses were more consistently expressed in CA1 cells
than in CA3 cells. Because CA3 pyramidal cells are known to exhibit a
Ca2+-dependent K+ current
(Schwartzkroin and Stafstrom 1980
; Storm
1990
), our observation suggests that the mGluR-mediated
[Ca2+]i rise in CA1 cells can more easily
access membrane K+ channels than can the
[Ca2+]i rise in CA3 cells.
Both CA1 and CA3 cells exhibited a sustained depolarization in response
to ACPD. It has been suggested that an mGluR-mediated depolarization in
CA1 pyramidal cells is due to a nonspecific cation current activated by
Ca2+ release from intracellular stores (Congar et
al. 1997; Crépel et al. 1994
). We found an
mGluR-mediated depolarization in both CA1 and CA3 cells that far
outlasted the transient Ca2+ elevation due to intracellular
release (Figs. 12B and 4). Moreover, the depolarization
persisted in the absence either of the transient (Fig. 8, A
and B) or of the sustained (Figs. 5 and 12) components of
increased [Ca2+]i, making it unlikely to be
the result of the Ca2+-activated current described previously.
The ACPD-induced depolarization led to a voltage-dependent Ca2+ influx. However, the ACPD V-I plot (Fig. 9) suggests that the voltage-dependent Ca2+ currents did not contribute significantly to the depolarization. If the depolarization caused by the Ca2+ current were large, one would expect a departure of the slope of the V-I curve from linearity.
mGluR-mediated depolarizations have been reported in both CA3 and CA1
that result from conductance increases (Congar et al. 1997; Crépel et al. 1994
;
Guérineau et al. 1995
; Pozzo Miller et al.
1995
). On the other hand, the long depolarization that we
recorded was associated with a conductance decrease (i.e., increase in
input resistance). mGluR-mediated depolarizations involving conductance
decreases have also been reported by others and attributed to blockade
of K+ channels (Charpak et al. 1990
;
Guérineau et al. 1994
; Lüthi et al.
1997
).
Apparently, the mGluR-mediated depolarization in hippocampal pyramidal
cells consists of multiple components. Differences in recording
conditions may affect the relative expression of the different
components. For instance, the depolarizing conductance increases were
recorded in the presence of potassium channel blockers (Congar
et al. 1997; Crépel et al. 1994
) or
elevated extracellular [K+] (Guérineau et
al. 1995
), whereas under normal ionic conditions, a
depolarization associated with a conductance decrease was more prominent (Guérineau et al. 1994
; and this study).
It is notable that mGluR-mediated depolarizations are sufficient to
activate voltage-dependent Ca2+ influx and thus to modulate
[Ca2+]i even near the resting membrane potential.
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ACKNOWLEDGMENTS |
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We thank L. R. Merlin and K. L. Perkins for reading the manuscript, W. Ross and N. Lasser-Ross for the kind gift of the data acquisition software, G. Frick and R. Law for help with data analysis software, and M. Avitable of the State University of New York Scientific/Academic Computing Center for help with statistics and image processing software.
This study was supported by National Institute of Neurological Disorders and Stroke Grant NS-35481.
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FOOTNOTES |
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Address for reprint requests: R. Bianchi, Dept. of Physiology and Pharmacology, State University of New York Health Science Center at Brooklyn, Box 29, 450 Clarkson Ave., Brooklyn, NY 11203.
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 6 November 1998; accepted in final form 10 February 1999.
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REFERENCES |
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