1Howard Hughes Medical Institute, 2Department of Biochemistry, and 3Department of Physiology and Neuroscience, New York University Medical Center, New York, New York 10016
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ABSTRACT |
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Boxer, Adam L., Herman Moreno, Bernardo Rudy, and Edward B. Ziff. FGF-2 Potentiates Ca2+-Dependent Inactivation of NMDA Receptor Currents in Hippocampal Neurons. J. Neurophysiol. 82: 3367-3377, 1999. Peptide growth factors such as the neurotrophins and fibroblast growth factors have potent effects on synaptic transmission, development, and cell survival. We report that chronic (hours) treatment with basic fibroblast growth factor (FGF-2) potentiates Ca2+-dependent N-methyl-D-aspartate (NMDA) receptor inactivation in cultured hippocampal neurons. This effect is specific for the NMDA-subtype of ionotropic glutamate receptor and FGF-2. The potentiated inactivation requires ongoing protein synthesis during growth factor treatment and the activity of protein phosphatase 2B (PP2B or calcineurin) during agonist application. These results suggest a mechanism by which FGF-2 receptor signaling may regulate neuronal survival and synaptic plasticity.
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INTRODUCTION |
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At excitatory, glutamatergic synapses, release of
the neurotransmitter glutamate stimulates a family of ionotropic
glutamate receptors. These receptors are transmembrane proteins that
are assembled from four to five structurally related integral membrane subunits. One member of this family is the
N-methyl-D-aspartate receptor (NMDAR), a cation
channel that is activated by simultaneous binding of the ligand,
glutamate, and depolarization of the postsynaptic cell (reviewed by
Hollmann and Heinemann 1994). The receptor is composed
of a ubiquitous NR1 subunit and one of four NR2 subunits termed NR2A-D
(reviewed by Mori and Mishina 1995
). Under
physiologically normal conditions, entry of Ca2+
ions through the NMDAR triggers biochemical and electrophysiologic changes that underlie synaptic plasticity. However, overactivation of
NMDARs, which may follow anoxia or glucose deprivation, has pathological consequences and is an important component of a large number of CNS disorders including excitotoxic cell death (Lipton and Rosenberg 1994
). Excitotoxic death is thought to result
from excessive Ca2+ influx through the receptor
following pathological receptor activation (reviewed by Mody and
MacDonald 1995
).
Peptide growth factors are regulators of CNS development, synaptic
plasticity, and neuronal survival after ischemia (reviewed in
Baird 1994; Eckenstein 1994
; Lewin
and Barde 1996
) and they exert both short and long term effects
on synaptic physiology (reviewed in Finkbeiner 1996
;
Katz and Shatz 1996
; Thoenen 1995
). The
expression of the growth factor, basic fibroblast growth factor (FGF-2), in the CNS increases during early postnatal development (Caday et al. 1990
; Kuzis et al. 1995
).
In the adult CNS, the concentration of FGF-2 exceeds those of certain
neurotrophins, such as nerve growth factor (NGF) (Eckenstein
1994
). These findings suggest that FGF exerts important
long-term effects in the adult as well.
Studies of excitotoxic cell death show that long term FGF-2 treatment
can prevent NMDAR-mediated cell death both in vitro (Férnandez-Sánchez and Novelli 1993;
Mattson et al. 1989
) and in vivo (Fisher et al.
1995
; Kirschner et al. 1995
; Koketsu et al. 1994
; MacMillan et al. 1993
; Nozaki
et al. 1993a
,b
). FGF can also regulate the internal cellular
Ca2+ concentration after NMDAR activation
(reviewed in Lindholm 1994
), and its neuroprotective
effects correlate with an attenuation of NMDAR-induced increases in
[Ca2+]i (Cheng and
Mattson 1991
; Mattson et al. 1989
) and with the expression of a putative NMDAR protein (Mattson et al.
1993
) and the Ca2+-binding protein,
calbindin D28 (Mattson et al. 1991
; Peterson et
al. 1996
). However, it is not known if FGF-2 affects NMDAR electrophysiologic function.
To determine whether FGF-2 can regulate the function of the NMDAR ion channel, we have carried out electrophysiological and molecular biological studies in cultured rat hippocampal neurons. Treatment with FGF-2 for hours to days significantly enhanced the ability of extracellular Ca2+ to inactivate NMDAR currents. Enhancement of receptor inactivation required protein synthesis during FGF-2 treatment, and the activity of calcineurin during agonist application, but did not involve changes in NMDAR protein subunit expression.
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METHODS |
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Cell culture
Hippocampal cultures were prepared similarly to those used by
Mattson et al. (1991). E18 Sprague-Dawley embryos
(pregnant rats obtained from Taconic) were dissected from rats
anesthetized with pentobarbital sodium and placed in ice-cold,
Ca2+/Mg2+-free
phosphate-buffered saline (PBS), containing 0.6% (wt/vol) D-glucose. Hippocampi were dissected and incubated in
1 × trypsin/EDTA (Gibco) plus 10 µg/ml pancreatic DNAse I
(Boehringer) at room temperature for 15 min. Cells were dissociated by
trituration through a series of fire-polished pasteur pipets and seeded
on polyornithine (50-100 µg/ml, [MW 30,000-70,000],
Sigma)/laminin (10 µg/ml; natural mouse, Gibco) at ~1 × 106 cells per 60 mm dish. Culture medium
consisted of minimum essential medium containing Earle's salts and
L-glutamine (Gibco No. 11095-080), supplemented with 10%
heat-inactivated (30[min], 56°C) fetal bovine serum, 20 mM KCl, 1 mM Na Pyruvate, 1% (vol/vol) penicillin-streptomycin, and 0.6%
(wt/vol) D-glucose and was changed 4 h and 3 days
after plating. Mitotic inhibitors were not added because these became neurotoxic in the presence of FGF-2. Cells were grown in a 5% CO2-98% humidity incubator in 3 ml of media. In
some initial experiments, medium was changed to neurobasal medium
(Gibco) plus 1 × B27 supplement (Gibco) and 0.5 mM
L-glutamine after 4 h. Growth in this medium suppressed glial cell growth in control cultures but not in cultures grown in the presence of FGF-2. Results obtained in neurobasal/B27 were
similar to those in serum containing medium. Growth factors were added
directly to the medium from stock solutions prepared and stored as
suggested by the manufacturer [R + D Systems for BDNF, FGF-2 (157aa.
isoform) and NGF; Promega or R + D systems for NT-3]. Cultures were
used at 8-10 days in vitro, at which time ~40% of cells were glia
(as judged by GFAP immunofluorescence), and little neuronal cell death
was evident.
Electrophysiology
Recording electrodes were formed on a Narishige PP-83 puller
from 1.5 mm diam hard glass (A-M Systems, No. 6030) and polished on a
Narishige MF-83 microforge. Pipet solution contained (in mM) 155 Cs
MeSO3, 10 Cs HEPES, 5 BAPTA, 0.4 CaCl2, 2 MgCl2, and 5 Na
ATP (pH 7.3 with CsOH). Standard ECS contained (in mM) 167 NaCl, 2.4 KCl, 10 Na HEPES, 1 CaCl2, 0.01-0.02 glycine,
and 10 D-glucose (pH 7.3 with NaOH). In 5 mM
CaCl2 ECS, NaCl concentration was reduced to 157 mM. In some experiments 1 µM TTX (Research Biochemicals, RBI) was
included, and did not produce any observable effect on glutamate
receptor currents. When filled with pipet solution, electrodes had
resistances of 3-7 M. Typical seal resistances were 2-4 G
.
Whole cell recordings (Hamill et al. 1981
) were carried out at 22-24°C in tissue culture dishes on the stage of an inverted microscope with the use of an Axopatch 200A amplifier (Axon
Instruments). Neurons were identified on the basis of morphology and
voltage-activated sodium conductances at the beginning of each
recording (see below). Series resistance was 70-90%
compensated and most of the capacitance was cancelled. Access
resistance was monitored with a 20-ms, 10-mV hyperpolarizing pulse at
the start of each episode. Currents were filtered at 2 kHz, digitized,
stored on hard disk, and analyzed with pClamp 6 software (Axon
Instruments). Curve fitting was done by Chebeshev method using pClamp 6 software or least squares method using DeltaGraph Pro, with equations
taken from pClamp 6 manual. Statistical analysis was performed with
StatView 4.5 (Abacus Concepts) on a Macintosh computer. Data are
expressed as the mean ± SE.
Agonist and antagonist application
Recordings were carried out in 6 cm tissue culture dishes and
were perfused continuously by peristaltic pump at a rate of 2-3 ml/min
(total bath volume was ~2 ml). Agonists and antagonists were
dissolved in extracellular solution and applied by pressure ejection
(1-300 mmHg) from a flow pipe ~50 µm from the cell, using a
computer-controlled solenoid valve, 12 reservoir drug application device (DAD-12; Adams and List Associates, Westbury, NY). The solution
exchange time was estimated to be ~200-300 ms based on the rise time
of kainate-evoked currents and perfusion of 10% normal ECS. After
gaining whole cell access, cells were held at 90 mV and stepped in 10 mV increments to +30 mV to verify neuronal phenotype (large Na-channel
current, not shown). All cells were also subject to an 100 µM kainate
I-V analysis to verify cell type dependence (Ozawa et
al. 1991
). Growth factors were not present in the ECS except as
indicated. Extracellularly applied drugs were present only during the
inactivation protocol (see below), except cycloheximide which was
present in the bath, wash and inactivation solutions, but not in the
agonist solutions. Intracellularly dialyzed drugs were present
throughout the experiment. All agonists and antagonists were obtained
from RBI except
-agatoxin IVA (Alamone Labs, Jerusalem),
cycloheximide (Sigma), calcineurin autoinhibitory peptide (Bachem),
control calcineurin inhibitory peptide (generous gift of B. Perrino and
T. Soderling, Vollum Institute), and phalloidin (Boehringer).
Molecular biology
RT-PCR analysis of NMDAR subunit mRNA was carried out by the
method of Sheng et al. (1994). Total RNA was isolated
from rat tissue or cultured cells by the guanidinium isothiocyanate/SDS method and digested with RNase-free DNAse I (Boehringer, 1 U/2.5 µg)
for 1 h at 37°C and phenol/chloroform extracted, ethanol
precipitated, and resuspended in water. cDNA was synthesized with the
Invitrogen cDNA cycle kit. cDNA reaction (1-2 µl) was included in 50 µl PCR reactions containing 60 mM Tris · Cl, 15 mM
NH4Cl, 1-7 mM MgCl2 (empirically optimized), and 2 mM each dNTP, plus 250 nM 5' and 3'
oligo and 1.5 U Amplitaq (Perkin-Elmer) polymerase. In some reactions,
8.3 µCi alpha 32P -dCTP was included. The
number of amplification cycles to remain in the linear range was
empirically determined for each oligonucleotide pair. Reactions were
visualized on 1.8% agarose gels, stained with ethidium bromide, or for
phosphorimager (Molecular Dynamics) quantitation, on 4% nondenaturing
polyacrylamide gels. Identity of bands was verified by size and by in
situ hybridization of adult rat brain sections (C. Kentros, A. Boxer,
B. Rudy, and E. B. Ziff, unpublished observations).
Oligonucleotide primers used to amplify NR1, N1, and C1 exons were
identical to those described in Sheng et al. (1994)
.
Neurofilament-L primers amplified nucleotides 1174-1719 of the rat
sequence: 5' oligo, TGGACATCGAGATTGCAGC; 3' oligo,
GGTTGGTGATGAGGTTGACC. NMDAR2A primers amplified nucleotides 4076-4453
of the rat sequence: 5' oligo, GGATTAACCGACAGCACTCC; 3' oligo,
ATGATGCTTGACCTCAAGG. NMDAR2B primers amplified nucleotides 3960-4347
of the rat sequence: 5' oligo, GCATTCCTACGACACCTTCG; 3' oligo, GACCACCACTGGCTTATTGG.
For western analysis, cells were washed twice with PBS and harvested in
ice-cold RIPA buffer containing protease and phosphatase inhibitors
(Sheng et al. 1994) for 5-15 min. Protein concentration was determined by Bradford Assay (BioRad) and equal amounts were loaded
onto SDS-polyacrylamide gels, run, and transferred to nitrocellulose by
standard methods. Blots were probed with antibody and visualized using
an ECL Kit (Amersham) according to manufacturer's directions. Antibodies were used according to manufacturer's directions as follows: mouse monoclonal anti-calbindin D28 and anti-neurofilament L
(Sigma); rabbit polyclonal antisera to NMDAR1, NMDAR2A; and rabbit
polyclonal NMDAR1, GluR1, GluR2/3, and GluR4 antisera from (Chemicon).
Immunocytochemistry
Cells were grown in eight-well, permanox chamber slides (Nunc), coated with polyornithine and laminin as above. Cells were washed twice with PBS and fixed in Vilim's fixative: 4% paraformaldehyde, 0.2% picric acid, 0.1 M Na2PO4, and 3% sucrose for 15-30 min. Cells were then washed three times in PBS for 5 min, permeabilized in 0.1% Triton X-100 + 0.1% BSA for 10-30 min, and blocked with 5% normal goat serum (Sigma) in PBS for 30 min or overnight. Immunocytochemistry was performed as directed with a Vectastain Elite immunoperoxidase kit. Antibodies were purchased from Chemicon.
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RESULTS |
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Ca2+ concentration and activity-dependent inactivation of NMDA-evoked currents in FGF-2 treated cells
The NMDAR undergoes several forms of activity-dependent current
inactivation (reviewed by Jones and Westbrook 1996). Two
of these forms, glycine-dependent and glycine-independent
desensitization, are induced by glutamate and NMDA in the presence of
low or high concentrations of the co-agonist glycine, respectively. A
third form of inactivation, Ca2+-dependent
inactivation (CDI), is induced by the increase in
[Ca2+]i (Legendre
et al. 1993
; Medina et al. 1996
) that follows
the entry of Ca2+ through the NMDAR receptor or
other channels. CDI involves the Ca2+-dependent
binding of calmodulin to the C0 domain of the NR1
C terminus and the displacement of
-actinin, which reduce NMDAR currents (Ehlers et al. 1996
; Krupp et al.
1999
; Wyszynski et al. 1997
; Zhang et al.
1998
). CDI is also sensitive to
[Ca2+]o (Legendre
et al. 1993
).
To determine whether FGF-2 changes activity-dependent physiological
properties of NMDARs, whole cell patch-clamp recordings (Hamill
et al. 1981) were elicited from pyramidal shaped neurons that
had been placed in culture for 8-11 days and subjected to different
growth factor treatments (Fig. 1). Cells
were analyzed that displayed 100 µM kainate-evoked I-V
profiles characteristic of type I, glutamatergic cells (Fig.
1G) (Ozawa et al. 1991
). NMDA-evoked currents
were detected in all cells tested. Cultures were treated with 10 ng/ml
FGF-2 for 24-48 h, or for controls were left untreated with growth
factor, or were treated with 10 ng/ml BDNF for 72-120 h before
recording.
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In the first experiments, we exposed FGF-2-treated cells, BDNF-treated
cells, and control cells to a 10-min conditioning pulse train
consisting of 3-s pulses of 50 µM NMDA in 1 mM
Ca2+ ECS applied for 2 min1. Each agonist application was preceded by
a 20-ms, 10-mV hyperpolarizing pulse from the holding potential of
60
mV to monitor access resistance. In addition to CDI, decreases in NMDAR
currents can take place in an irreversible,
Ca2+-independent, ATP-sensitive manner (washout)
(MacDonald et al. 1989
; Wang et al.
1996
), or in a reversible,
Ca2+-dependent, ATP-sensitive manner (rundown)
(Rosenmund and Westbrook 1993a
,b
). In this experiment a
concentration of ATP (5 mM) sufficient to prevent washout and retard
rundown was included in the pipet solution.
During the administration of the pulse train, all cells displayed an inactivation of NMDA-evoked current (average amplitude during 3 s pulses) whose magnitude increased with successive stimulations. However the inactivation was greater in FGF-2 treated cells than in control or BDNF-treated cells (Fig. 1, A and B). After 10 min of stimulation, control and BDNF-treated cell currents decreased to 75.2 ± 6.4% and 76.6 ± 4.9% of initial values, respectively [not significantly different, analysis of variance (ANOVA), Sheffé post hoc test], whereas currents in FGF-2-treated cells decreased to a much greater extent, to 31.5 ± 6.4% (P < 0.001) of initial values. This showed that FGF-2 treated cells displayed enhanced activity-dependent current inactivations relative to untreated or BDNF treated controls during a NMDA pulse train that were highly statistically significant at physiological concentrations of Ca2+ (1 mM Ca2+).
The decay of the NMDA-evoked currents in FGF-2-treated cells was best
fit by a single exponential with a time constant of 323 ± 79 s (n = 8). To determine whether extracellular
Ca2+ was required to maintain this current
inactivation, the ECS bathing the cell was switched to a nominally
Ca2+-free (1.5 mM EGTA) ECS after the 10 min of
stimulation in 1 mM Ca2+ ECS and cell stimulation
with 50 µM NMDA was continued. Figure 1C shows that a
representative FGF-2-treated cell displayed almost complete recovery of
NMDAR currents after 10 min in Ca2+-free ECS. On
average (Fig. 1D), NMDA-evoked currents recovered to ~90%
of initial, 1 mM Ca2+ ECS, values in 3 min for
control cells (n = 4), and in 6 min for FGF-2 treated
cells (n = 4). This demonstrates that the
inactivation of NMDAR currents during the administration of the pulse
train requires extracellular Ca2+, is greater in FGF-2
treated cells than in BDNF-treated or untreated control cells, and is
reversible in both control and FGF-2 treated cells when extracellular
Ca2+ is removed. Furthermore, the decline of the current in
the presence of extracellular Ca2+, as well as the recovery
of the current in the absence of extracellular Ca2+,
involves relatively slow processes (1/2 = ~300 s)
.
Mechanism of the FGF-2 effect
To determine the mechanism of action of FGF-2, we analyzed several
parameters of FGF-2 effects, including the specificity of the effects
for FGF-2 relative to other growth factors and the specificity for the
NMDAR versus the -amino-3-hydroxy-5-methyl-4-isoxazole propionic
acid receptor (AMPAR). We also determined the length of exposure to
FGF-2 necessary to alter NMDAR currents, the roles of new protein
synthesis and of the Ca2+-regulated phosphatase,
calcineurin (PP2B), and of voltage regulated Ca2+
channels. We asked if FGF-2 acts through modification of NMDAR subunit
composition. Finally we examined the effects of FGF-2 on currents
elicited by single agonist pulses.
Specificity for NMDAR and chronic FGF-2 treatment
To determine whether AMPARs are also regulated by FGF-2, 3-s
pulses (2 min1) of 100 µM kainate were
applied to cells in 1-2 mM Ca2+ ECS. After 25 min of stimulation, there was no difference in the amount of
inactivation in kainate-evoked currents in control (49.6 ± 9.0%
relative to initial value) versus FGF-2 treated cells (49.6 ± 9.0%) (Fig. 1, E and F). Thus the effects of
FGF-2 treatment are specific to the NMDAR versus the AMPAR.
To determine the length of FGF-2 treatment required to potentiate the
inactivation of NMDAR currents, 0.5-s test pulses (1 min1) of NMDA were applied to cells, before and
after 50 conditioning pulses 3 s each (2 min
1) of 10 µM L-glutamate in 1 mM Ca2+ ECS (see Fig.
2). The responses of cells treated with
FGF-2 for different times were compared. Figure 2A shows
NMDA-evoked currents recorded from a representative control and FGF-2
(24 h) treated cell. Peak NMDA-evoked currents were reduced to 66 and
68% of initial values (50 and 100 µM; compare preconditioning pulse
records with postconditioning) by the conditioning pulses in the FGF-2 treated cell, but not in the control cell. NMDA-evoked currents decreased to 71.3 ± 3.6% of initial values in cells that had
been treated with FGF-2 for 24 h (Fig. 2B). Control
cells displayed NMDA-evoked currents that were 116 ± 11.4% of
initial values and thus showed no current change. The effect in the
FGF-2 treated cells was significant (P
0.006, ANOVA,
Fisher's PLSD post hoc test), and also occurred in cells treated with
FGF-2 for 4 h and 120 h before recording, but not in cells
treated with FGF-2 only during the 25 min of conditioning pulses (0 h).
A similar result was obtained when the conditioning pulses used NMDA
(10 µM) instead of glutamate (data not shown). These data suggest
that the greater Ca2+-dependent NMDAR current
inactivations seen in FGF-2 treated cells require >0.5 h of growth
factor treatment, and are not an acute effect of FGF-2.
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To rule out the possibility that the effects of FGF-2 are general
effects of RTK stimulation, cells were treated with 10 ng/ml of BDNF or
NT-3 for 120 h and subject to the above assay. Only FGF-2-treated
cells displayed an inactivation of NMDA-evoked currents after
conditioning pulses with glutamate, although both BDNF and FGF-2 were
able to induce the expression of the Ca2+ binding
protein, Calbindin D28 (Ghosh and Greenberg 1995;
Vicario-Abejon et al. 1995
) (Fig. 2, C and
D). These data confirm the results of Fig. 1 that BDNF does
not induce the effect and indicate that the inactivation of NMDA-evoked
currents is a specific effect of activation of the FGF RTK (FGF
receptor 1 or FGFR1) (Eckenstein 1994
) and not a general
effect of neurotrophic growth factor RTK activation.
Dependence on Ca2+ and calcineurin and ongoing protein synthesis
To address biochemical aspects of the mechanism, conditioning
pulses of glutamate were applied under different extracellular ionic
conditions or in the presence of different pharmacological agents.
Cells were then tested with 0.5-s pulses of 50 and 100 µM NMDA
(response ratios averaged, Fig.
3C). As above, the
inactivation of the current was dependent on repetitive activation of
the NMDAR and was not a nonspecific effect of whole cell dialysis
(MacDonald et al. 1989) because conditioning pulses
lacking agonist did not decrease the peak current ratio in FGF-2
treated cells. Also consistent with previous results, the inactivation
of the current was dependent on Ca2+ entry
because application of the conditioning pulses in
Ca2+-free (1.5 mM EGTA) ECS abolished the effect
(P = 0.05). The inactivation did not result from
rundown, a Ca2+-dependent decrease in NMDAR
currents involving local depolymerization of the actin cytoskeleton,
with a time course similar to the FGF-2-potentiated inactivation of
NMDAR currents (Rosenmund and Westbrook 1993b
). Rundown
is blocked by dialysis of the cell with 1 µM phalloidin, a
filamentous actin stabilizing drug. However, inclusion of phalloidin in
the pipet solution had no effect on the NMDA-evoked inactivation.
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Ca2+-dependent inactivations of NMDAR function
may be mediated by the Ca2+-dependent protein
phosphatase 2B, calcineurin (Lieberman and Mody 1994;
Tong and Jahr 1994
; Tong et al. 1995
). To
address whether calcineurin was required for the FGF-2 effect, a
specific calcineurin inhibitor peptide that mimics the autoinhibitory
domain (10 µM) (Hashimoto et al. 1990
) was included in
the pipet. The peptide significantly inhibited the FGF-2-treated cell
Ca2+-dependent inactivation of peak 50 µM
NMDA-evoked current (P < 0.01), whereas a control
inhibitor peptide with two mutant amino acid residues had no effect.
This shows that calcineurin is required for the
Ca2+-dependent NMDAR current inactivation
potentiated by FGF-2-treatment.
In Fig. 2B, >25 min of cell treatment with FGF-2 (the time of FGF-2 treatment for t = 0 h) was necessary for FGF-2's effects on NMDAR currents. The requirement for extended treatment raised the possibility that FGF-2 was acting indirectly, perhaps through the induction of the synthesis of a new protein. To determine the requirement for new protein synthesis, neurons were treated with FGF-2 (10 ng/ml) for 24 h in the absence and presence of 1 µM cycloheximide and assayed for changes in NMDAR inactivation. Figure 3A shows that the effect of FGF-2 on peak 50 µM NMDA-evoked current ratios is abolished in the presence of cycloheximide (also observed at all NMDA concentrations tested, data not shown). This concentration of cycloheximide is sufficient to block the FGF-2-induced increase in calbindin D28 protein level, as assayed by western blot (Fig. 3B). Thus the FGF-2 effect requires ongoing protein synthesis during FGF-2 exposure.
Independence from voltage gated calcium channels and from NMDAR subunit change
It has been shown that FGF-2 increases the expression of L-type
voltage-gated Ca2+ channels (Shitaka et
al. 1996). Increased Ca2+ fluxes through
voltage-gated channels in neurons in the distal neurites could
contribute to the altered NMDAR responses observed in FGF-2 treated
cells. However, specific inhibitors of L-type (50 µM nifedipine),
N-type (10 µM
-conotoxin GVIA), and P/Q-type Ca2+ channels (200 nM
-agatoxin IVA), either
alone or in combination (data not shown) had no effect on the
inactivation of NMDAR currents in FGF-2-treated cells (Fig.
3C). This indicates that these voltage-gated Ca2+ channels do not have a role in the FGF-2 effect.
The requirement for ongoing protein synthesis during FGF-2
treatment suggests that a change in gene expression may be required for
the FGF-2 effect. To determine whether the FGF-2 effect results from
changes in the expression of NMDAR protein subunits (Hollmann and Heinemann 1994), the levels of these subunits were assayed in control cells and cells treated with FGF-2 for 120 h. No
significant FGF-2-induced changes were detected in NR1 (constitutively
spliced portion of transcript), NR1 (N1 exon), NR1 (C1 exon), or NR2A mRNA levels, as analyzed by RT-PCR. There was a ~30% decrease in
NR2B mRNA levels, relative to an NF-L internal control (Fig. 4Bii), however western
analysis (Fig. 4C, ii and iii) failed to demonstrate a change in NR2B protein levels. There were also no changes
in protein levels or distribution induced by FGF-2 treatment, as
assayed by immunocytochemistry (NR1-full protein, NR2A/B, GluR1, GluR2/3, andGluR4; data not shown). FGF-2 treatment induced an increase
in calbindin D28 protein expression (Figs. 2-4Cvi), but there were no changes in calcineurin
subunit levels (Fig.
4Cvii). Significant levels of NR1, NR2B, and GluR1-3, but
not N2RA protein (Fig. 4Cii) were detected in our cultures,
consistent with reports of NMDAR subunit mRNA expression in cortical
neuron and glial cultures grown under similar conditions (Zhong
et al. 1994
). NMDAR NR1 subunit mRNA contained approximately a
20:1 ratio of N1-exon (absent):N1-exon (present) and a 20:1 ratio of
C1-exon (present):C1-exon (absent) splice variants (Sheng et al.
1994
) in both control and FGF-2 treated cells (Fig. 4A,
ii and iii). These results strongly suggest that FGF-2
does not act by altering the levels of known neuronal glutamate
receptor proteins.
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FGF-2 alters currents elicited by single agonist pulses
To determine if the effects of FGF-2 could be observed
during single NMDA pulses, cells were held at 60 mV and a pulse of 50 µM NMDA was applied via pressure ejection for 9 or 3 s in
Ca2+-free (1.5 mM EGTA) or in 1 mM
Ca2+-containing, Mg2+-free
extracellular solution (ECS), containing 20 µM glycine. In 0 mM or 1 mM extracellular Ca2+, there was no significant
difference (P > 0.1, Student's t-test, two
tailed) in the peak amplitude of the NMDA-evoked currents in
FGF-2-treated versus control cells (Fig.
5A, i-iv). In ECS containing
1 mM Ca2+, the currents inactivated after the
initial peak. In the presence of 50 µM NMDA for 3 s, there was a
greater inactivation of the current in FGF-2-treated (34.4 ± 5.6%) relative to control cells (21.0 ± 2.2%) in 1 mM
Ca2+ extracellular solution (P = 0.06; Fig. 5B). This suggested that the effect of FGF-2
treatment is observed during single pulses and is enhanced with
increasing concentrations of Ca2+. To determine
whether a higher extracellular Ca2+ concentration
would accentuate the difference in NMDAR currents during single pulses
in FGF-2 treated versus control cells, 9-s pulses of 50 µM NMDA were
applied in ECS containing 5 mM Ca2+ to control
and FGF-2 treated cells, as above. Elevation of the extracellular
Ca2+ concentration (Fig. 5A, v and
vi) led to a larger inactivation of the current in both
FGF-2-treated and control cells. Although the peak, NMDA-evoked
currents were not significantly different under these conditions
[630.6 ± 56.6 (FGF) vs. 734.5 ± 66.7 pA (control),
P > 0.1, Student's t-test, two tailed],
the mean I3s and mean current at
9 s (I9s) were significantly
(P = 0.05 at 3 s and P = 0.007 at
9 s) reduced in FGF-2 treated cells to a greater extent than in
control cells. The mean I9s in
FGF-2-treated cells was ~50% of controls: 205.9 ± 52.8 versus
438.1 ± 61.2 pA. This difference was not related to a difference
in cell size because the mean whole cell capacitance of the cells was
similar under these conditions (24.2 ± 3.3 vs. 27.8 ± 1.2 pF). The difference in I9s was caused
by a greater degree of time-dependent current inactivation in the
FGF-2-treated than in control cells (Fig. 5B): 69.6 ± 5.7% versus 42.3 ± 3.2%, after 9 s (P = 0.0004; I3s's also significantly
different, P = 0.0007). There was no difference in
relative mean current amplitude responses to 1-s applications of 1-500
µM NMDA (P > 0.1, ANOVA; Fig. 5C),
suggesting that FGF-2 treatment does not alter the receptor's agonist
affinity. These data show that NMDARs in FGF-2 treated hippocampal
neurons display a greater Ca2+-dependent
inactivation of NMDAR currents during single 3- or 9-s applications of
NMDA in comparison with these receptors in control cells.
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DISCUSSION |
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A novel action of basic FGF-2 on embryonic hippocampal neurons is described here. We show that FGF-2 treatment of 8 d.i.v. hippocampal neurons enhances the capacity of the NMDAR to undergo a form of Ca2+-dependent inactivation. The change is specific for both FGF-2 and the NMDA receptor. The change requires new protein synthesis during growth factor treatment, which is consistent with a need for >25 min of FGF-2 stimulation. Changes in NMDAR subunit composition, however, do not appear to be involved. The inactivation is dependent on extracellular Ca2+, and is reversible in the absence of extracellular Ca2+. The inactivation was highly statistically significant following a train of pulses administered under physiological concentrations of extracellular Ca2+ ([Ca2+] = 1 mM). The magnitude of the current inactivation relative to the initial current during a pulse train became greater with successive pulses. This suggests that, the inactivation may be a consequence of increased accumulation of intracellular Ca2+ during the train. In agreement, during single pulses, the extent of the FGF-2 effect was elevated by increases in Ca2+ in the bath. The inactivation, however, does not depend on voltage regulated Ca2+ channels. A possible target for Ca2+ is the Ca2+-dependent protein phosphatase, calcineurin, whose activity was required for receptor inactivation.
Forms of use-dependent decrease of NMDAR function
The NMDAR undergoes several forms of activity-related decrease in
function which are distinguished from one another by their dependence
on agonist and Ca2+ (reviewed by Jones and
Westbrook 1996). Two forms of desensitization may take place
following glutamate (or NMDA) binding (Benveniste et al.
1990
; Mayer et al. 1989
; Vyklicky et al.
1990
). In the first, glycine-independent desensitization, a
high (micromolar) concentration of the co-agonist glycine, limits
receptor entry into the most highly desensitized states. At suboptimal
glycine concentrations (in the nanomolar range), the receptor undergoes a greater, glycine-dependent desensitization. Because the FGF-2 induced
effects studied here take place in the presence of 10 µM glycine,
they are unlikely to involve the first of these processes, glycine-dependent desensitization. Moreover, because both control and
FGF-2 treated cells showed identical current profiles when stimulated
with 50 µM NMDA and 10 µM glycine in the absence of extracellular
Ca2+, the extent of the second desensitization
process, glycine-independent desensitization which takes place under
these conditions, is not altered by FGF-2. Finally, the FGF-2 effect is
unlikely to involve either rundown, a decrease in conductance resulting
from depolymerization of the actin cytoskeleton (Rosenmund and
Westbrook 1993a
,b
) because it was not affected by phalloidin, a
stabilizer of actin filaments, or washout because of washout's
irreversibility (MacDonald et al. 1989
; Wang et
al. 1996
).
Relationship of Ca2+-dependent inactivation to the FGF-2 effect
The NMDAR undergoes yet another process, CDI (Krupp et al.
1999; Legendre et al. 1993
; Mayer and
Westbrook 1985
; Medina et al. 1994
;
Zilberter et al. 1991
). CDI is a glycine-insensitive inactivation whose rate (
inact = 4.7 s at
[Ca2+]0 = 1.3 mM)
increases with increasing
[Ca2+]o
and which plateaus at 45-50% (Legendre et al. 1993
).
CDI is diminished by buffering intracellular Ca2+
with BAPTA (Legendre et al. 1993
). Following CDI, if
[Ca2+]o is lowered,
currents recover biphasically, with kinetic components of
1/2 = 0.5-5 s and of
1/2 = 10-50 s (Legendre et al.
1993
; Medina et al. 1994
, 1996
). This recovery
is closely correlated with the decay of
[Ca2+]i (Medina et
al. 1996
). Because inactivation develops on the inner part of
the plasma membrane (Medina et al. 1996
) and because tonic intracellular perfusion of the cell with elevated
Ca2+ occludes inactivation (Legendre et
al. 1993
), CDI is thought to be induced by elevation of
[Ca2+]I rather than by
ligand binding alone.
The activity-dependent inactivation of NMDA receptor currents
described here in FGF-2 treated cells resembles CDI in several respects. These include its Ca2+-dependence, its
reversibility, its time course and its kinetic parameters. At 0 µM
[Ca2+]o, there was no
difference between FGF-2 treated and control cell current profiles. As
[Ca2+]o was increased,
current inactivations were seen in both cell groups, but were greater
for FGF-2 treated cells. Thus the FGF-2 effect was manifested only in
the presence of extracellular Ca2+ and is larger
in 5 mM than in 1 mM Ca2+, both characteristics
of CDI. Furthermore, the current inactivation reversed fully in the
absence of extracellular Ca2+, as does CDI. Also,
the time constant of the current inactivation during an individual
agonist pulse, ~1-5 s, is similar to that of CDI (Legendre et
al. 1993; Medina et al. 1994
), when the two are
measured under comparable conditions, but is greater than that of
glycine-independent desensitization (~100 ms) (Benveniste et
al. 1990
).
During administration of a train of agonist pulses to FGF-2 treated
cells, we observed inactivation of mean receptor currents with an
apparent 1/2 = 323 s. The currents
recovered when the cells were exposed to 0 mM
Ca2+ ECS, with an apparent
1/2 of ~360 s. These values for
1/2 are apparent time constants that depend on
the parameters of the protocol. However, they indicate that the FGF-2
potentiated effect has a long time constant component, as does CDI. The
Ca2+-dependence, reversibility and kinetic
parameters of the FGF-2 potentiated effect all suggest that either CDI
itself, or another process with similar properties, is enhanced in
cells which have been treated with FGF-2.
Role of calcineurin, -actinin, and Ca2+ homeostasis
FGF-2 could alter the biochemical response of the NMDAR to
Ca2+ fluxes. Peptide inhibitor experiments
indicate that potentiation by FGF-2 requires calcineurin. Calcineurin
has been implicated previously in receptor control (Lieberman
and Mody 1994; Tong and Jahr 1994
; Tong
et al. 1995
), although the dependence of inactivation on a
phosphatase has been questioned (Legendre and Westbrook
1990
; Medina et al. 1996
; Rosenmund and
Westbrook 1993a
). The effects of FGF-2 also require new protein
synthesis prior to agonist stimulation of the receptor. The new protein
could function as a receptor regulatory factor that enhances the
ability at Ca2+ to control receptor currents.
However, calcineurin itself is not likely to be the newly synthesized
protein since FGF-2 did not alter its level.
Recently, it has been shown (Halpain et al. 1998;
Wyszynski et al. 1997
) that brief (5 min) treatment of
cultured hippocampal neurons with NMDA induces an increase in the
levels of calcineurin in spines, and a calcineurin-dependent collapse
of spine structure that is associated with the depolymerization of
F-actin. These findings raise the possibility that the
calcineurin-dependent enhancement of receptor auto-regulation by FGF-2
described here involves a calcineurin-dependent reorganization of the
actin cytoskeleton. Indeed, the association of the actin cytoskeleton
with the NMDA receptor has been shown to affect channel currents and is
controlled by Ca2+ through
Ca2+-calmodulin, which displaces
-actinin from
NR1 (Ehlers et al. 1996
; Krupp et al.
1999
; Zhang et al. 1998
). Krupp et al.
(1999)
have shown that CDI is abolished when the
C0 region of C terminal domain of NR1, which
contains the binding site for
-actinin (Wyszynski et al.
1997
), is truncated. Krupp et al. (1999)
suggest that the receptor, when "latched" to the actin cytoskeleton
via
-actinin, is in the active state. Possibly the requirement that
we observe for protein synthesis during treatment with FGF-2 involves
expression of a protein that makes the receptor more susceptible to
dissociation from
-actinin by Ca2+- and
calcineurin-dependent process.
FGF-2 may also act by altering the regulation of intracellular
Ca2+. Indeed, FGF-2 induces L-type
Ca2+-channels in fetal hippocampal neurons (Shitaka
et al. 1996). However, this is an unlikely basis for the action
of FGF-2 since voltage gated Ca2+ channel (VGCC) blockers
did not attenuate the FGF-2 effect. The induction by NMDA, the
dependence on [Ca2+]o, and the lack of
sensitivity to VGCC blockers, all suggest that the NMDAR rather than
VGCC is the major point of Ca2+ entry. Neither the role of
endoplasmic Ca2+ stores, shown to be important in dorsal
horn neurons (Kyrozis et al. 1996
), nor the role of
retrieval mechanisms were studied.
Alteration of Ca2+ homeostasis by FGF-2 and a
consequent accumulation of intracellular Ca2+ could also
increase inactivation. Perhaps the Ca2+ elevation produced
by the conditioning train is larger or longer lasting in the FGF-2
treated cells, inactivating the receptor. Increased levels of
Ca2+ could enhance the Ca2+-calmodulin
dependent displacement of -actinin discussed above. Increased
receptor inactivation and calbindin induction, which was also observed
in FGF-2 treated cells, may be viewed as components of a more general
protective cellular response to increased intracellular Ca2+ transients. However, induction of calbindin is
unlikely to be the basis for the effects on the NMDA receptor because
BDNF did not alter receptor regulation although it induced calbindin.
Also, there is no evidence that the calbindin in these cells is in
sufficient proximity to the synaptic membrane of dendritic spines to
buffer acute rises in peri-synaptic calcium concentration.
Physiological significance of FGF-2-induced increases in NMDAR inactivation
Glutamate receptor inactivation is a process that maintains
physiological levels of signaling while preventing toxicity
(Brorson et al. 1995; David et al. 1996
;
Zorumski and Thio 1992
; Zorumski et al.
1989
). Treatment with FGF-2 protects neurons from toxic insults
involving NMDAR activation in vitro, and in vivo reduces focal ischemic
infarct size (Lin and Finklestein 1997
). Our results suggest that an enhancement of the capacity of the NMDAR to inactivate may contribute to the neuroprotective effect of FGF-2. The high levels
of FGF proteins and the ubiquitous neuronal expression of the FGF
receptor, FGFR1, in adult CNS neurons are consistent with such a role
(Caday et al. 1990
; Eckenstein 1994
;
Kuzis et al. 1995
; Yazakai et al. 1994
).
NMDAR desensitization and inactivation are important components
of basic synaptic physiology (reviewed by Jones and Westbrook 1996),
and their changes may influence the threshold for synaptic modification
(Abraham and Bear 1996
). Developmental increases in FGFR
signaling could regulate synaptic development or plasticity by shifting
the threshold for synaptic plasticity to favor LTD or related events.
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ACKNOWLEDGMENTS |
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We thank M. Chesler and P. Osten for critical reading of the manuscript; S. Burden, B. Perrino, and M. Sheng for helpful discussions; B. Perrino and T. Soderling for the control calcineurin inhibitory peptide; and M. Sheng for antibodies. A. L. Boxer is a trainee at the New York University Medical Center Medical Scientist Training Program and E. B. Ziff is an Investigator at the Howard Hughes Medical Institute.
This research was supported by National Institutes of Health Grants NS-30989 to B. Rudy and AG-13620 to E. B. Ziff.
Present address of H. Moreno: Dept. of Neurology, SUNY Brooklyn College of Medicine, Brooklyn, NY 11203-2098.
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FOOTNOTES |
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Address for reprint requests: E. B. Ziff, Howard Hughes Medical Institute, New York University Medical Center, Dept. of Biochemistry, 550 First Ave., New York, NY 10016.
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 13 April 1999; accepted in final form 17 August 1999.
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REFERENCES |
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