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The Journal of Neuroscience, June 15, 2000, 20(12):4582-4595
Potentiation of NMDA Receptor Function by the Serine Protease
Thrombin
Melissa B.
Gingrich,
Candice E.
Junge,
Polina
Lyuboslavsky, and
Stephen F.
Traynelis
Department of Pharmacology, Emory University School of Medicine,
Atlanta, Georgia 30322
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ABSTRACT |
Although serine proteases and their receptors are best known for
their role in blood coagulation and fibrinolysis, the CNS expresses
many components of an extracellular protease signaling system including
the protease-activated receptor-1 (PAR1), for which thrombin is the
most effective activator. In this report we show that activation of
PAR1 potentiates hippocampal NMDA receptor responses in CA1 pyramidal
cells by 2.07 ± 0.27-fold (mean ± SEM). Potentiation of neuronal NMDA receptor responses by thrombin can be
blocked by thrombin and a protein kinase inhibitor, and the effects of
thrombin can be mimicked by a peptide agonist (SFLLRN) that activates
PAR1. Potentiation of the NMDA receptor by thrombin in hippocampal
neurons is significantly attenuated in mice lacking PAR1. Although high
concentrations of thrombin can directly cleave both native and
recombinant NR1 subunits, the thrombin-induced potentiation we observe
is independent of NMDA receptor cleavage. Activation of recombinant
PAR1 also potentiates recombinant NR1/NR2A (1.7 ± 0.06-fold) and
NR1/NR2B (1.41 ± 0.11-fold) receptor function but not NR1/NR2C or
NR1/NR2D receptor responses. PAR1-mediated potentiation of recombinant
NR1/NR2A receptors occurred after activation with as little as 300 pM thrombin. These data raise the intriguing possibility
that potentiation of neuronal NMDA receptor function after entry of
thrombin or other serine proteases into brain parenchyma during
intracerebral hemorrhage or extravasation of plasma proteins during
blood-brain barrier breakdown may exacerbate glutamate-mediated cell
death and possibly participate in post-traumatic seizure. Furthermore,
the ability of neuronal protease signaling to control NMDA receptor
function may also have roles in normal brain development.
Key words:
serine protease; thrombin; NMDA receptor; protease
receptor; PAR1; hippocampal neurons
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INTRODUCTION |
It is becoming increasingly apparent
that the CNS expresses many serine proteases and zymogen precursors, as
well as unique members of the serpin class of selective serine protease
inhibitors (Nakajima et al., 1992 ; Sumi et al., 1992; Chen et
al., 1995; Gschwend et al., 1997; Hastings et al., 1997; Krueger et
al., 1997; Luthi et al., 1997; Pindon et al., 1997; Scarisbrick et al.,
1997; Shimizu et al., 1998; Yamamoto and Loskutoff, 1998). The
G-protein-coupled thrombin receptor protease-activated receptor-1 (PAR1) is also present in the developing and mature CNS, with expression in specific cells such as the CA1 pyramidal neurons of the
hippocampus (Weinstein et al., 1995; Niclou et al., 1998). Although
endogenous activators for this receptor in the brain have not been
defined, the mRNA for the thrombin precursor prothrombin and the
protein that converts prothrombin to thrombin (Factor Xa) are both
expressed in CNS tissue (Dihanich et al., 1991, Yamada and Nagai,
1996). In addition, blood-derived thrombin and other PAR1 activators
such as plasmin will directly enter brain tissue during penetrating
head wound, hemorrhagic stroke, rupture of cerebral aneurysms or
arteriovenous malformations, and possibly therapeutic treatment with
tissue plasminogen activator (tPA). Furthermore, if thrombin (36.7 kDa)
is generated in sufficient quantities during occlusive stroke or other
insults that increase blood-brain barrier permeability, it would be
expected to penetrate brain parenchyma given the permeability of larger
molecular weight markers such as albumin (66 kDa) (Laursen et al.,
1993) and dextran (71 kDa) (Du et al., 1996). Some blood-derived serine
proteases can directly increase blood-brain barrier permeability (Nagy
et al., 1998). The total quantity of active thrombin that can be produced from blood is 260-360 U/ml (Seegers, 1962; Arand and Sawaya,
1986), suggesting that prothrombin circulates at high concentrations
(>1 µM). Plasminogen circulates in blood at ~2 µM (Majerus et al., 1996).
The possibility that thrombin and other blood proteases might enter
brain tissue under pathological conditions raises the potential for
aberrant activation of protease receptors with potentially detrimental
consequences. This possibility is supported indirectly by several
observations. Thrombin infusion into rat caudate nucleus can produce a
glial scar similar to that typical of traumatic head injury (Nishino et
al., 1993; Motohashi et al., 1997). Furthermore, thrombin activation of
PAR1 triggers neurite retraction, glial proliferation, and apoptosis
(Gurwitz and Cunningham, 1988; Cavanaugh et al., 1990; Grabham and
Cunningham, 1995; Donovan et al., 1997). Thrombin directly injected
into the rat basal ganglia causes edema and precipitates seizures (Lee
et al., 1996, 1997), suggesting that thrombin, in addition to
heme-derived iron, may contribute to post-traumatic seizures
(Willmore et al., 1978). The best indicators of post-traumatic
epilepsy subdural hematoma and intracerebral hemorrhage (Willmore
1990; Lee and Lui, 1992; Annegers et al., 1998)could both potentially
lead to entry of serine proteases such as thrombin from the vasculature
into the brain.
In this study we have evaluated the possibility that thrombin actions
on PAR1 might modify NMDA receptor function, which is an important
contributor to both seizure initiation and neurodegeneration subsequent
to cerebrovascular insult or traumatic brain injury (Bradford 1995;
Whetsell, 1996; Obrenovitch and Urenjak, 1997; Dirnagl et al., 1999;
Lee et al., 1999).
Some of these results have been published previously in abstract form
(Butler and Traynelis, 1996; Gingrich et al., 1997, 1998; Gingrich and
Traynelis, 1998; Junge et al., 1999).
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MATERIALS AND METHODS |
Electrophysiological recording from rat and mouse
hippocampal slices. Mice or rats [postnatal day 12-21
(P12-P21)] were anesthetized using isoflurane and decapitated, and
the hippocampus was rapidly dissected. All procedures involving animals
have been approved by the Emory University Institutional Animal Care
and Use Committee. Transverse hippocampal slices (250-300 µm) were
cut in ice-cold artificial CSF (ACSF) using a vibratome and
secured in a submerged recording chamber perfused with 1 µM tetrodotoxin and 10 µM bicuculline in
ACSF. ACSF was composed of (in mM): 124 NaCl, 26 NaHCO3, 2.5 KCl, 1 CaCl2,
1.4 MgCl2, 1 NaH2PO4, and 10 glucose,
and was saturated with 95% O2-5%
CO2, pH 7.4. In some experiments the
extracellular recording solution was supplemented with 10 µM nifepidine (in 0.2% DMSO) to reduce
Ca2+ currents. Blind and visually guided
whole-cell patch recordings were obtained at 23°C from CA1 pyramidal
neurons using thin-walled 2.8-5.5 M glass pipettes filled with a
solution composed of (in mM): 110 Cs-gluconate, 40 HEPES, 5 MgCl2, 2 Na-ATP, 0.6 EGTA, and 0.3 Na-GTP, with
the pH adjusted to 7.3 using CsOH; osmolality was 275-290 mOsm. In
some experiments, EGTA was omitted, 40 mM HEPES was
replaced with 5 mM HEPES plus 30 mM CsCl, and
the solution was supplemented with 1 mM QX-314 (Sigma, St.
Louis MO); similar results were obtained with both internal solutions.
The presence of intracellular Cs+ should
block GABAB receptor-mediated currents. Brief
(<100 msec) pulses of NMDA (0.3-2 mM) plus glycine
(0.1-0.3 mM) were applied via pressurized pipette placed
either in or just above stratum radiatum; the pressurized pipette was
positioned to apply drug to the proximal third of the CA1 pyramidal
cell dendrite; dilution at the tip was minimized before recording, and
the tip was checked for blockage at the end of the experiment.
NMDA-evoked currents were recorded at  70 mV (corrected for the +10 mV
measured junction potential) before, during, and after thrombin
application. In some experiments, membrane potential was changed to
40 mV during alternate agonist applications, or briefly jumped to
70 and 40 mV from a holding potential of 0 mV before and during
agonist application. Series resistance (mean 23.4 ± 2.3 M) was
monitored from the instantaneous current response to a 5 mV jump
applied before agonist application, and the membrane resistance (mean 1.4 ± 0.2 G) was estimated from the leak current at 70 mV
assuming a reversal potential of 0 mV. Series resistance compensation
was not used because the mean response amplitude (49 pA) will cause only a 1 mV error in the holding potential, and the slow response time
course eliminates the capacitative component of series resistance filtering. Experiments with substantial changes in membrane or series
resistance, regenerative currents, or development of leak currents
exceeding 200 pA at 70 mV were excluded from analysis. After 3-10
stable baseline measurements were taken, 3 U/ml  -thrombin (Calbiochem, La Jolla CA; Sigma, St. Louis MO; Hematological
Technologies, Essex Junction, VT) was applied through the bath solution
for 10-18 min. In control experiments, ACSF was applied through the same perfusion line as thrombin. The perfusion line and recording chamber were washed extensively after experiments involving thrombin treatment because low picomole levels of -thrombin are capable of
inactivating PAR receptors before recording (Vu et al., 1991). The
specific activity of the -thrombin from various vendors ranged between 1720 and 3200 NIH U/mg by comparison to Lot J of the NIH standard. To estimate the concentration of active -thrombin that corresponds to 1 U/ml activity, we calculated a conversion factor using
our most pure -thrombin (3200 U/mg). Because the protein in this lot
was reported by the manufacturer to be >95% -thrombin as
determined by gel electrophoresis, a solution with 1 U/ml -thrombin should be 9 nM using a molecular weight for thrombin of
36.7 kDa. For simplicity, we used a conversion factor of 1 U/ml = 10 nM -thrombin throughout the text to estimate the
concentration of active -thrombin (hereafter referred to as
thrombin) from various vendors. The peak potentiation of NMDA responses
by thrombin was calculated over 20 min after thrombin application as
the ratio of the highest running average of three consecutive response
amplitudes (excluding the first 3 min after thrombin application) to
the average of the three responses surrounding the time of thrombin application. Peak potentiation was used as a measure of the actions of
thrombin to remove the variability associated with the time required
for thrombin to reach cells at different depths in the slice.
Imaging of Fluo-3 fluorescence from cultured hippocampal
neurons. Embryonic day (E)17-E19 rat pups were taken from
CO2-asphyxiated pregnant rats, and the
hippocampus was dissected. Neurons were dissociated by trituration
through a sterile fire-polished Pasteur pipette, centrifuged, and
plated in Neurobasal (Life Technologies, Gaithersburg, MD) defined
media at a density of 60,000/ml on polylysine-coated (10 µg/ml) 12 mm
glass coverslips. Media was supplemented with B27 nutrients (Life
Technologies) plus penicillin/streptomycin, and cells were maintained
for 3-5 d at 37°C in humidified 5% CO2. Neurons were incubated for 30-45 min in a solution composed of (in
mM): 150 NaCl, 3 KCl, 10 HEPES, 2 CaCl2, 20 mannitol, 10 glucose supplemented with
0.1% pluronic acid, 0.5% DMSO, and 3 µM Fluo-3 acetoxymethyl ester (Molecular Probes, Eugene OR); dye-loaded cells
were placed in an identical solution lacking DMSO, pluronic acid, and
Fluo-3. Images were acquired every 15 sec after 1 sec exposure to
450-490 nm light, and fluorescence was recorded through a bandpass
filter (500-550 nm) using a Photometrics CC200 CCD camera; 3 nM thrombin (0.3 U/ml) ± 50 nM
D-Phe-Pro-Arg-chloromethylketone (PPACK; an irreversible
thrombin inhibitor) (Tapparelli et al., 1993) as well as 10 µM PAR agonist peptide SFLLRN were applied in the
presence of 0.5 µM tetrodotoxin and 50 µM
APV. NMDA (50 µM) and glycine (10 µM) were
subsequently applied in the presence of 0.5 µM
tetrodotoxin. Fluorescence intensity was measured in cell bodies using
image analysis software (Scion Corporation, Frederick, MD) and
expressed as F/Fo where
Fo is the fluorescence intensity before drug treatment. Increases in fluorescence greater than
1.2-fold were considered to be real changes because untreated cells
possessed a peak F/Fo ratio
of 1.1 ± 0.04 over a typical experiment.
Transmitted light measurements in hippocampal slices. Rat
hippocampal slices were prepared as described above and were bathed in
a submerged chamber with 0.5 µM TTX and 3-7 U/ml
thrombin in 0.5 µM TTX applied for 10-20 min. The
intensity of transmitted light (450-490 nm) through hippocampal slices
was monitored as an indication of extracellular volume fraction (Andrew
and MacVicar, 1994). Images were recorded using a Princeton Micromax
CCD camera (Trenton, NJ) (0.2 sec exposure every 20-40 sec) and
analyzed using Axon Imaging Workbench 2.1 software (Foster City, CA).
Intensity was expressed as
I/Io, where
Io is the intensity of transmitted light before treatment. Solutions were made hyperosmotic by addition of
30 mM mannitol, which can produce a 10%
expansion of the extracellular volume fraction (McBain et al.,
1990). Hypo-osmotic solutions were obtained by addition of 10%
v/v water.
Genotyping of PAR1 / mice.Male PAR1 +/ mice were
obtained from University of California San Francisco (gift from Dr.
Shaun Coughlin) and have been described elsewhere (Connolly et al., 1996). PAR1 +/ (C57Bl/6 background) mice were bred with female C57Bl/6 wild-type mice from Jackson Laboratories (Bar Harbor, ME), and
the presence of PAR1 or neomycin gene was determined using PCR of
genomic tail DNA obtained by digesting tails in 0.7 ml of a
solution containing 0.1 M EDTA, 50 mM
Tris, 0.5% SDS, 1 mg/ml proteinase K, pH 8.7, at 50-55°C for 4-5
hr; the digest was subsequently extracted with phenol/chloroform. The
DNA was precipitated and resuspended at 1 µg/µl in Tris/EDTA (10 mM/1 mM). PCR reactions were run with 1 µg
genomic DNA using the following protocols repeated through 40-45
cycles: 95°C (30 sec), 85°C (5 sec), 60°C (2 min), 72°C (2 min)
for PAR1 +/+ mice, and 95°C (30 sec), 85°C (5 sec), 58.5°C (45 sec), 72°C (1 min) for the neomycin resistance gene. The primers for
PAR1 have been described (Connolly et al., 1996); for neomycin the
primer pair GAAGGGACTTGCTATTGG, GCTCTTCAGCAATATCACGGG was used, which
generates a 431 bp fragment. PCR reactions testing the genotype of
unknown animals were performed in parallel with control DNA from PAR1
+/+, PAR1 +/, and PAR1 / mice. Mice used in this study were
derived from PAR1 / breeding pairs (11), PAR1 +/+ breeding pairs
(4), or Jackson Laboratories wild-type mice (11). PCR results were
repeated three times for all mice included in the study.
Expression and recording of NMDA and PAR1 receptor function in
Xenopus laevis oocytes. cRNA was synthesized from
linearized template cDNA according to manufacturer specifications
(Ambion, Austin, TX). Xenopus oocytes were removed from
ovaries of female frogs anesthetized with 0.3% 3-aminobenzoic acid
ethylester. Groups of 20-30 stage V-VI oocytes were incubated in 292 U/ml Worthington (Freehold, NJ) Type IV collagenase for 2 hr with slow
shaking. The oocytes were rinsed in Barth's solution and stored at
17°C in Barth's solution supplemented with 100 µg/ml gentamycin
and 40 µg/ml streptomycin. Control oocytes were injected with 3 ng NR1 subunit and 7 ng NR2B subunit. Other oocytes were injected with the
NMDA receptor subunits plus 3 ng PAR1 cRNA. Two-electrode voltage-clamp
recordings were made 3-7 d after injection. Oocytes were continually
perfused with recording solution (90 mM NaCl, 1 mM KCl, 10 mM HEPES, pH
7.4) and held under voltage clamp at 30 to 50 mV with an OC-725B
amplifier (Warner Instruments, Hamden, CT). Recording solution was
supplemented with 1.0 mM
CaCl2 during initial impalement of the oocytes
and subsequently 0.5 mM
BaCl2 during recording to reduce the
calcium-activated chloride current endogenous to oocytes. Voltage and
current electrodes had a resistance of 3-6 M when filled with
0.3-3 M KCl. NMDA receptor currents were evoked
by 20 µM glutamate/10
µM glycine perfused for 0.5-1 min. PAR1
receptor activation was elicited by either 2.5 U/ml thrombin
(Calbiochem) or 10-30 µM agonist peptide
SFLLRN (Bachem, Torrance, CA) for 2-3 min; the recording chamber and
perfusion lines were washed extensively between thrombin applications
because low picomole levels of thrombin are capable of inactivating PAR receptors before recording (Vu et al., 1991). Expression of PAR1 protein was verified by a thrombin- or SFLLRN-stimulated rapidly desensitizing inward current at 30 mV that reflects the
Ca2+-activated
Cl current endogenous to the oocyte
(Miledi, 1982). To test the ability of thrombin to modify NMDA receptor
function in oocytes, recordings of receptor function were made both
before and after a 15-60 min incubation in recording solution
supplemented with thrombin in Eppendorf tubes at room temperature. Only
oocytes that showed <10% change in current between a pair of
responses within each 10 min recording period were included in the
analysis, to eliminate possible rundown of response. Oocytes with
responses <50 nA were not included in the analysis.
NMDA receptor subunit immunoblots. Human embryonic kidney
(HEK) cells were maintained in DMEM supplemented with 0.5 mM glutamine, 1 mM pyruvate, penicillin,
streptomycin, and 10% fetal bovine serum in a humidified environment
with 5% CO2, and transfected with 1 µg/ml NR1
or 1:2 µg/ml NR1/NR2 cDNA (Chen and Okayama, 1987). Cells were washed
in ice-cold HEPES-buffered saline (HBS) and scraped on ice, and
membranes were isolated by centrifugation at 12,000 × g. N-terminal myc-tagged NR1-1a was a gift from
Dr. R. Huganir (Johns Hopkins University, Baltimore, MD) and was
constructed by inserting a myc tag (MEQKLISEEDLN) after
Asn50, 29 residues after the signal peptide. Adult rat brain regions
were dissected, frozen in liquid nitrogen, homogenized in ice-cold HBS,
centrifuged, and stored at 20°C. Membranes were resuspended in HBS
and treated with thrombin or buffer for 30 min at 37°C. Samples
were centrifuged, and the membrane pellet was resuspended in 2% SDS,
62.5 mM Tris, 10% glycerol, 5%
 -mercaptoethanol, 0.05% bromophenol blue, pH 6.8; some samples were
treated with 10 U DNase for 10 min at 37°C. SDS-PAGE and protein
blotting to Immobilon-P membranes were performed as described (Lau and
Huganir, 1995). Membranes were blocked for 30 min in 0.2 M Tris base, 1.37 M NaCl,
pH 7.4, containing 5% nonfat dry milk, and incubated overnight at
4°C in primary antibodies: NR1 mAb54.1 (gift from Dr. S. Heinemann,
Salk Institute, La Jolla, CA), NR1 alt-C-terminal (Upstate
Biotechnology, Lake Placid, NY), NR2A C-terminal (Chemicon, Temecula,
CA), or NR2B C-terminal (Upstate Biotechnology). Membranes were washed
three times and incubated with HRP-conjugated goat anti-mouse or goat
anti-rabbit antibodies (1:10,000), washed three times, and developed
using enhanced chemiluminescence. The percentage of receptor cleaved by
thrombin was determined using densitometry.
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RESULTS |
Thrombin potentiation of NMDA receptor currents in rat
hippocampal neurons
Messenger RNA encoding the thrombin receptor PAR1, which is known
to activate GQ- and
GI-linked intracellular signaling systems (Grand
et al., 1996; Dery et al., 1998), is expressed in hippocampal neurons
including CA1 pyramidal cells (Weinstein et al., 1995; Niclou et al.,
1998). We have studied the effects of thrombin on whole-cell
voltage-clamp recordings of rat CA1 hippocampal pyramidal neuron
responses to pressure application of NMDA plus glycine into the
dendritic field to test whether serine proteases can alter NMDA
receptor function through PAR1 activation. Figure 1A illustrates our
experimental recording arrangement. NMDA receptor current responses
were evoked in the presence of 1 µM
tetrodotoxin and 10 µM bicuculline. Typical
NMDA-evoked current responses recorded at 70 mV are shown as a time
course before and during the application of 30 nM
thrombin (3 U/ml) for a representative cell (Fig.
1B). In this cell, thrombin treatment potentiated the
current response with a time course consistent with thrombin diffusion
into the tissue. To confirm that the enhanced inward current response
arises from activation of NMDA receptors rather than
strychnine-sensitive glycine receptors or sensitization of
stretch-activated channels that might respond to pressure ejection of
agonist, responses to NMDA-glycine were blocked by 100 µM of the competitive NMDA receptor antagonist
APV (n = 7 cells). Thrombin potentiation of NMDA
responses in neurons with the pressurized pipette positioned <100 µm
above the slice ruled out the possibility that thrombin sensitized the
tissue to pressure. Thrombin produced on average a 2.07 ± 0.27-fold
peak potentiation (mean ± SEM; n = 21) of NMDA responses at 70 mV within 20 min compared with cells in which thrombin was not applied (Fig. 1C). Although potentiation
was often rapid (within a few minutes) (Fig.
1B,C), in some cells thrombin
potentiation continued to increase with time, reaching peak levels
between five- and eightfold. The thrombin-induced peak potentiation of
NMDA responses appeared to possess a bimodal distribution in which a
subset of the cells showed little response to thrombin (Fig.
1D). This differential effect of thrombin is consistent with the expression of PAR1 mRNA observed in some but not
all CA1 pyramidal cells (Weinstein et al., 1995; Niclou et al.,
1998).
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Figure 1.
A, The diagram illustrates
our experimental arrangement. The photomicrographs show typical
placement of the pressurized pipette (N)
in relation to recording pipette (R) in
stratum pyramidale. The typical placement of the NMDA containing
pipette (black arrow) is shown in relation to a single
CA1 pyramidal cell (white arrow). The black
arrowhead shows the CA1/CA3 boundary. B, Time
course of current-response amplitude to NMDA during 3 U/ml thrombin
application. Inset, Representative current-responses
from this same cell to pressure-ejected 1 mM NMDA and 100 µM glycine were recorded at 70 mV before and during
perfusion with 3 U/ml thrombin (30 nM). APV (100 µM) blocked the response to NMDA/glycine.
C, Mean time course (±SEM) of current responses to NMDA
plus glycine in cells treated with 3 U/ml thrombin or 3 U/ml thrombin + 100 ATU hirudin. The number of cells is indicated in
parentheses. The mean response time course is also
shown for control cells that were superfused with ACSF.
D, Mean potentiation (±SEM) of NMDA receptor responses
by thrombin (open bar; 70 mV) was blocked by an
irreversible serine protease inhibitor with high selectivity for
thrombin over plasmin. PPACK (500 nM) (~15× thrombin
concentration) was preincubated for 5-15 min with 30 nM
thrombin before application to slices. Some slices were pretreated with
10 µM bisindolylmaleimide (BIS) for 10 min. *p < 0.05 compared with thrombin
potentiation; Kruskal-Wallis ANOVA, Dunn post hoc test.
Open circles show results from individual neurons.
E, Scatter plot of the NMDA response amplitude
(top panel) versus fold-potentiation by thrombin
and membrane resistance versus fold-potentiation by thrombin
(bottom panel). There was no significant
correlation between these parameters or the level of thrombin-induced
potentiation (p < 0.5). Data from rat and
mouse were pooled.
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On average there were no significant changes in series resistance
(111 ± 7%; p < 0.2; paired t test)
during thrombin application. Average membrane resistance (1.2 ± 0.1 M) decreased modestly throughout the course of the experiment;
however, this decrease was not significantly different between
thrombin-treated (12 ± 6%) and control slices (16 ± 7%;
p < 0.2; t test). In addition, there was no
significant correlation (p < 0.5 in all cases)
between the levels of thrombin potentiation and the following
parameters: the series resistance (r = 0.37), the
change in series resistance (r = 0.00), the membrane
resistance (r = 0.24) (Fig. 1E), the change in membrane resistance (r = 0.18), or the
response amplitude (r = 0.04) (Fig.
1E). Current responses recorded under voltage clamp
to pressure application of NMDA reversed sign at +0.1 mV (n = 6). Together these results suggest that the
thrombin-induced potentiation of NMDA receptor function that we
observed does not reflect a thrombin-induced compromise of the voltage
clamp. Indeed, an identical degree of thrombin potentiation of the NMDA
response (1.8 ± 0.2-fold) was observed in cells not included in Figure 1D because either their input resistance was too low
or there was an increase in series resistance throughout the experiment (n = 12). Furthermore, thrombin-induced potentiation
was observed when cells were held at 0 mV to inactivate
Ca2+ channels, and the holding potential
briefly stepped to 70 mV before and during agonist application
(2.7 ± 0.7-fold potentiation; n = 7). There was
no significant difference between the degree of potentiation seen under
normal conditions or in cells held at 0 mV (p > 0.05; Mann-Whitney test). These results argue against the possibility
that asynchronous activation of PAR1/PKC-potentiated (e.g., Hall et
al., 1995) Ca2+ channels during
NMDA-induced loss of voltage clamp in distal dendrites could account
for the potentiation that we observed.
To confirm that the potentiating effects of thrombin that we observed
reflect the proteolytic actions of thrombin rather than nonspecific
effects or the actions of other contaminant proteases, we evaluated
whether the potentiation of NMDA receptor responses by thrombin could
be blocked by two selective thrombin antagonists. We first examined
whether hirudin (Calbiochem), an inhibitor that binds with high
affinity to the anion binding exosite
(KD 20 fM) of
thrombin, could alter the time course of thrombin potentiation of NMDA
receptor function. Hirudin significantly reduced the potentiation by
thrombin of NMDA responses (0.96 ± 0.23-fold; n = 5) (Fig. 1C) (p < 0.05;
Mann-Whitney test for average potentiation 12-19 min after
treatment). In a separate experiment, we premixed thrombin with an
irreversible inhibitor of thrombin that covalently modifies the
fibrinopeptide catalytic site for cleavage of substrate (500 nM PPACK) (Tapparelli et al., 1993) before
application to the slice. The average peak potentiation of the NMDA
response after thrombin-PPACK application was also significantly
reduced (1.28 ± 0.17; n = 8) compared with
thrombin treatment (2.07 ± 0.27-fold; n = 21)
(Fig. 1D) (Kruskal-Wallis ANOVA, Dunn post
hoc test; p < 0.05). Nonparametric statistical
tests were used because of the non-normal distribution of peak fold
potentiation by thrombin (Fig. 1D). These experiments
confirm that the potentiation of NMDA receptor currents that we
observed are caused by the proteolytic actions of thrombin on either a
protease receptor such as PAR1 or some other substrate.
Because PAR1 is expressed in the CA1 region and is known to couple to
the GQ family of G-proteins, which can
stimulate phosphoinositide hydrolysis, increase intracellular
Ca2+, and activate intracellular protein
kinases (Grand et al., 1996), we tested whether the serine/threonine
protein kinase inhibitor bisindolylmaleimide (BIS) could occlude
thrombin potentiation of NMDA receptor function. When BIS (10 µM) was included in ACSF, potentiation of NMDA receptor
currents by application of 30 nM (3 U/ml) thrombin was
reduced to 1.20 ± 0.13-fold (n = 10, p < 0.05; ANOVA) (Fig. 1D),
indicating that the potentiating effect of thrombin requires at least
one functional protein kinase. NMDA responses were unchanged during
preapplication of BIS, suggesting that BIS does not induce an
inhibition of the NMDA response that is additive to potentiation.
Thrombin does not alter extracellular volume fraction in
hippocampal slices
In vivo injection of thrombin into the brain can cause
edema and tissue swelling (Lee et al., 1996), raising the possibility that the potentiation we observed of NMDA receptor responses to agonist
applied from a point source might reflect a change in the spatial
diffusion profile of NMDA and glycine secondary to thrombin-induced
changes in extracellular volume fraction. For example, any changes in
the extracellular volume fraction that amplified the concentration of
pressure-ejected NMDA in the narrow clefts or expanded the tissue
volume reached by NMDA would potentiate the NMDA response. To ensure
that the diffusional characteristics of pressure-applied NMDA are not
influenced by thrombin treatment, we measured the effects of thrombin
application on transmitted light, which has previously been shown to be
a sensitive measure of extracellular volume fraction (Andrew and
MacVicar, 1994). Hippocampal slices (300 µm) were prepared and
incubated in 0.5 µM TTX to abolish synaptic
activity. Perfusion of these slices with solutions made hyperosmotic by
addition of 30 mM mannitol reduced the
transmitted light by 10% in both CA1 stratum radiatum and stratum
pyramidale, consistent with expansion of the extracellular space
(McBain et al., 1990; Andrew and MacVicar, 1994). Conversely, hypo-osmotic solutions produced a 20% increase in transmitted light
(Fig. 2A), consistent
with induction of cell swelling. Treatment of slices with 3-7 U/ml of
thrombin for 10-20 min produced no significant change in the intensity
of transmitted light (I) in either stratum pyramidale
(I/Io = 1.00 ± 0.01)
or stratum radiatum (I/Io = 1.01 ± 0.01) compared with untreated slices (Fig.
2B). These data suggest that neither thrombin
activation of PAR1 nor cleavage of other substrates significantly
alters extracellular volume fraction in acute slices. We interpret
these data to suggest that the potentiation of NMDA receptor responses
that we observe after activation of PAR1 does not reflect a
thrombin-induced change in the temporal-spatial diffusion profile of
NMDA-glycine released into or above the tissue from our pressurized
pipette.
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Figure 2.
A, Left
panel, Photograph of a 300 µm rat hippocampal slice with the
region indicated in blue expanded to the
right. Right panels, Pseudocolor
representation of the relative intensity of transmitted light (500-550
nm) in a slice bathed in ACSF after treatment with 0.5 µM
TTX (Control), 0.5 µM TTX plus 7 U/ml thrombin, or 0.5 µM TTX with ACSF made hypo-osmotic
by addition of 10% v/v water. Boxes show regions from
which average intensity measurements were made in this slice.
SP, Stratum pyramidale; SR, stratum
radiatum. The color code on the right
indicates a 3.5-fold relative range of intensity. B,
Left panel, Mean time course (±SEM) of the effects of
thrombin or hypo-osmotic treatment on transmitted light in seven cells.
Increased transmittance is correlated with reduced extracellular volume
fraction (see Results) (Andrew and MacVicar, 1994). Right
panel, Summary of experiments showing no effect of thrombin on
mean relative transmitted light compared with hypo-osmotic ACSF or ACSF
made hyperosmotic by addition of 30 mM mannitol.
Bars show the mean ratio of transmittance after
treatment (I) to control
transmittance (I0).
* indicates significantly different from control
measurements (p < 0.05; t
test). The number of slices is indicated in
parentheses.
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Thrombin proteolysis of the NR1 subunit
Because one defining feature of thrombin is its proteolytic
action, we examined whether NMDA receptor protein subunits were substrates for thrombin cleavage. Incubation of brain membranes with
thrombin for 1 hr at 37°C resulted in the appearance of a new band
that was 12 kDa (n = 10) lower in molecular weight than the parent band in immunoblots probed with an antibody to NR1 that
recognizes the M3 M4 loop (mAb54.1; epitope between residues 660-811)
(Siegel et al., 1994) (Fig.
3A). However, similar
treatment of membranes did not induce any observable molecular weight
shift of either the NR2A (n = 8) (Fig. 3B)
or NR2B subunits (n = 10; 3-300 U/ml; data not shown).
NR1 subunits in all brain regions tested were cleaved by high
concentrations of thrombin (1-3 µM; 100-300
U/ml; n = 5-7 for each region). The degree of cleavage by both high and low concentrations of thrombin varied across regions,
as shown in Figure 3A. Although cerebellar
(n = 7), cortical (n = 5), and
brainstem (n = 5) NR1 subunits were insensitive to cleavage by 300 nM thrombin (30 U/ml), 30 nM thrombin (3 U/ml) cleaved 20% of the NR1
subunit in hippocampal membranes (n = 7) and 50% of
NR1 in striatal membranes (n = 10), suggesting that receptors in these membranes are more sensitive to the effects of
thrombin. In addition, 1-3 µM thrombin cleaved
100% or subunits in all brain regions (Fig. 3C) except
striatum, where the maximal observed cleavage of 52% suggests that a
thrombin-resistant population of NR1 subunits exists. Thrombin-induced
NR1 cleavage does not involve PAR1 receptor-activated second messenger
systems because 0.1-1 mM PAR1 peptide agonist
SFLLRN did not stimulate cleavage (n = 3) (Fig.
3C). In addition, 100 ATU hirudin, a thrombin
inhibitor, blocked cleavage of NR1 in striatal membranes by 30 nM thrombin (n = 3), suggesting
that the shift in the molecular weight of the NR1 subunit does not
reflect the actions of contaminant proteases.
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Figure 3.
A, Left panel, Immunoblot
analysis showing thrombin cleavage of striatal NR1 subunit in membrane
homogenates detected with mAb54.1. Right panel, The
concentration dependence of thrombin cleavage of the NR1 subunit is
shown for various brain regions. The degree of cleavage has been
normalized to 1.0 to facilitate comparison of the thrombin sensitivity.
One hundred percent of NR1 was cleaved in all brain regions except
striatum, which showed maximum 52% cleavage. Measurements are the mean
of five experiments. B, Immunoblot analysis showing the
thrombin insensitivity of hippocampal NR2A detected with a C-terminal
antibody. No new bands were observed after the treatment of NR2A with
low or high concentrations of thrombin. C, Thrombin
treatment of hippocampal NMDA receptors reduces the molecular weight of
NR1, detected with mAb54.1. Treatment of membranes with a supramaximal
concentration of the PAR1 peptide agonist SFLLRN did not alter the
molecular weight of NR1. D, Thrombin treatment of
recombinant NR1-1a subunits engineered to include a sequence encoding
a myc tag at the N terminal and expressed in HEK
cells (see Materials and Methods) detected with a myc
antibody caused a similar molecular weight shift as for
thrombin-treated neuronal receptors. E, Thrombin
treatment did not alter the molecular weight of recombinant NR1-1b
subunits with the C terminal deleted after residue 785 (NR1 785).
Thrombin treatment of the same deletion construct engineered to express
a thrombin cleavage site (LVPRGS) starting at residue 198 in exon 5 verified that thrombin was active in the experiment because a band
corresponding to the predicted molecular weight for cleavage at exon 5 was observed. F, Linear map of the NR1 gene product
showing the relative positions of the engineered myc and
mAb54.1 epitopes, the engineered thrombin cleavage site, the 785
deletion, and the region predicted from these results to harbor the
thrombin cleavage site. Black boxes are
membrane-associated domains.
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Recombinant NR1 subunits transiently expressed in HEK 293 cells were
also sensitive to thrombin cleavage. Thrombin (10-1000 U/ml) cleaved
both homomeric NR1-1b receptors (n = 9/10 experiments) and heteromeric NR1-1b/NR2B receptors (n = 6/6
experiments; data not shown). The appearance of a new NR1 band of
reduced molecular weight in thrombin-treated brain membranes was
detected using mAb54.1, which recognizes an epitope located near the M3
transmembrane region (Fig. 3F) (Siegel et al., 1994).
To evaluate the location of the thrombin cleavage site, we assessed the
effects of thrombin on a recombinant NR1-1a subunit that contained a
myc epitope inserted into the N terminal after Asn50.
Thrombin induced a similar shift in molecular weight for N-terminal
myc-tagged NR1 as observed for the recombinant and native
receptors detected with mAb54.1 (Fig. 3D) (n = 3), suggesting that cleavage likely occurs near the C terminal of the
receptor. Consistent with this idea, deletion of the C-terminal portion
of NR1-1b by replacement of the coding sequence starting at amino acid
position 785 with a cDNA encoding HLEGPIL-stop abolished any detectable
cleavage of the NR1 subunit (Fig. 3E) (n = 3). These data suggest that thrombin cleaves NR1 subunit somewhere near
the C terminal (Fig. 3F) and were consistent with
immunoblots performed using a C-terminal antibody (Fig.
4B). However, use of a
C-terminal antibody recognized a larger cleavage product than predicted
from data in Figure 3. The reasons for the discrepancy in the molecular
weight of the C-terminal NR1 cleavage product from brain membranes as
detected with different antibodies are under investigation.
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Figure 4.
A, The time course of the response
amplitude for a CA1 pyramidal cell that exhibits robust potentiation
during thrombin treatment is shown. Inset,
Current-responses before and after thrombin (3 U/ml) treatment of the
slice are superimposed. Application of 100 µM APV
completely inhibited the response. B, Immunoblot
analysis using a C-terminal NR1 antibody of microdissected CA1 region
after thrombin treatment from the slice shown in A
confirms that our thrombin treatment did not cause proteolytic cleavage
of the NR1 subunit. The control lane shows the position of the expected
cleavage band in brain membranes treated with 300 U/ml thrombin.
C, Treatment of NR1/NR2A and NR1/NR2B recombinant
receptors expressed in Xenopus oocytes with the same
concentration of thrombin (3 U/ml) and the same duration (15 min) used
in slice experiments did not significantly alter the mean response
amplitude (±SEM) compared with buffer-treated oocytes
(t test). The mean fold potentiation by thrombin is
shown and was calculated from each oocyte as the ratio of the
post-treatment current (I) to the
pretreatment current (I0).
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To test whether the potentiation of NMDA responses that we observed in
hippocampal neurons treated with low concentrations of thrombin for
short periods of time (12-20 min at 23°C) was related to proteolysis
of the NR1 subunit, the CA1 region was microdissected after
electrophysiological experiments and analyzed by SDS-PAGE followed by
immunoblot. The results of one such experiment are shown in Figure 4.
Despite robust potentiation of the NMDA receptor response during the 15 min of 3 U/ml thrombin application (Fig. 4A), no NR1
cleavage product was observed in immunoblots of membranes obtained from
the CA1 region microdissected immediately after the experiment (Fig.
4B). Similar results were obtained from six slices,
which are consistent with data in Figure 3A that show only
modest (~20%) cleavage of hippocampal NR1 subunit by four times
longer thrombin treatments (1 hr incubation at 37°C with 3 U/ml
thrombin). If cleavage of NR1 by 3 U/ml thrombin is linearly related to
time and temperature sensitive with a
Q10 of 2.5, we predict from the 20%
cleavage of hippocampal NR1 after 1 hr at 37°C that thrombin should
cleave 1% of the NR1 protein after 15 min treatment of our slices at
23°C, which is likely below our limit of detection. In addition,
there was no significant difference in the response amplitude of
recombinant NR1/NR2A and NR1/NR2B NMDA receptors expressed in
Xenopus oocytes, which lack thrombin receptors (Vu et al.,
1991) that were treated either with buffer or identical thrombin
concentrations and incubation conditions used in slice experiments (3 U/ml; 15 min; 23°C) (Fig. 4C). Incubation of oocytes with
higher concentrations of thrombin (300-1000 nM)
for 60 min reduced current-response amplitude to 42 ± 9%
(n = 7), compared with 78 ± 5%
(n = 6; p < 0.05, t test) for buffer-treated controls, suggesting that thrombin can inhibit NMDA
receptor function, presumably through cleavage of NR1. These and other
results (see below) suggest that the thrombin-mediated potentiation of
the NMDA current responses in CA1 hippocampal pyramidal neurons does
not reflect NR1 proteolysis or direct interaction of thrombin with NMDA receptors.
Thrombin potentiation of NMDA receptor function is
PAR1 dependent
To evaluate the working hypothesis that thrombin activation of the
protease receptor PAR1 leads to potentiation of NMDA receptor secondary
to activation of intracellular signaling pathways, we first verified
that hippocampal neurons contain functional protease receptors, which
are known to couple to the GQ family of
G-proteins (Yang, 1997). We measured increases in fluorescence of
the calcium-sensitive dye Fluo-3 in response to 3 nM
thrombin or 10 µM PAR1 peptide agonist SFLLRN. This
peptide matches the new N terminal of PAR1 that is revealed by thrombin
cleavage at Arg41 and is thought to act as a tethered activator of the
receptor (Fig.
5A,B).
Experiments were performed on cultured neurons in the presence of 0.5 µM TTX to reduce the synaptic activity and 50 µM APV to eliminate the possibility that
thrombin potentiation of tonically active NMDA receptors (Sah et al.,
1989) might increase intracellular Ca2+.
Images were acquired while the following sequence of solutions was
perfused into the chamber: TTX/APV, TTX/APV/thrombin or SFLLRN, TTX/APV, TTX/NMDA/glycine. Both thrombin and the PAR1 peptide agonist
SFLLRN elicited a robust increase in the Fluo-3 fluorescence in both
the soma and dendrites of a subset of neurons, suggesting that these
treatments increased intracellular Ca2+.
The thrombin inhibitor PPACK (50 nM) reduced the
percentage of neurons responding to thrombin and the response magnitude
to control levels (1.1 ± 0.04-fold; n = 46) (Fig.
5B). Because we could detect no proteolysis of NR1 in the
slices that we studied, the most likely explanation for the
potentiation that we observed after thrombin treatment was that
thrombin activated PAR receptors on CA1 pyramidal cells. To directly
evaluate this possibility, we tested whether the specific PAR1 agonist
peptide SFLLRN, which mimics the new N terminal on PAR1 revealed after
thrombin cleavage after Arg41, could also potentiate NMDA receptor
responses in CA1 pyramidal cells in acute hippocampal slices. Figure
5C summarizes the results of this experiment and shows that
application of 30 µM PAR agonist SFLLRN (10×
EC50) to the slice produced a 1.76 ± 0.19-fold (n = 9) potentiation of the response to
pressure-applied NMDA plus glycine, which was significantly different
from that observed in buffer-treated slices (1.05 ± 0.03-fold
potentiation; n = 8; p < 0.01 Mann-Whitney test). These data are consistent with the idea that
thrombin potentiates NMDA receptors through activation of a protease
receptor, most likely PAR1.
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Figure 5.
A, Thrombin and PAR1 peptide
agonist SFLLRN increase somatic Fluo-3 fluorescence in cultured
hippocampal neurons. The peak fluorescence increase in the experiments
shown was 3.45- and 2.23-fold for SFLLRN and thrombin, respectively.
The bar on the right shows pseudocolor
scale representation of fluorescence intensity
(black = lowest intensity,
yellow = highest intensity). B,
Bar graphs summarize the results from Fluo-3 imaging
experiments, which demonstrate the expression of thrombin- or peptide
agonist-responsive receptors in some cultured neurons. Left
panel shows mean (±SEM) peak Fluo-3 fluorescence as fold
increase (F/Fo)
induced by 0.3 U/ml (3 nM) thrombin, 10 µM
peptide agonist SFLLRN, or 3 nM thrombin preincubated with
50 nM of the thrombin inhibitor PPACK. *
indicates significantly greater than control
(p < 0.05; t test). Fluo-3
fluorescence increased in response to thrombin or SFLLRN within a few
seconds and relaxed toward baseline over the next 10 min. Right
panel summarizes number of neurons that responded to PAR1
activation; only cells that responded to NMDA with an increase in
Fluo-3 fluorescence were considered to be neurons and included in the
analysis. C, Left panel CA1 pyramidal
cell current-responses to pressure-applied NMDA/glycine before and
during SFLLRN application; circle shows time of
NMDA/glycine application. Right panel, Mean peak fold
potentiation (±SEM) of the NMDA receptor responses in rat CA1
pyramidal cells by 30 µM (~10×
EC50) (Gerszten et al., 1994) of the PAR peptide
agonist SFLLRN is compared with cells treated with ACSF. Holding
potential was 70 mV. Number of cells is indicated in
parentheses. *p < 0.05;
Mann-Whitney test.
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To further evaluate the role of PAR1 activation in thrombin-induced
potentiation of NMDA receptor responses, we studied the effects of
thrombin application on hippocampal CA1 pyramidal cells from wild-type
mice as well as mice engineered to lack the full-length PAR1 gene
(Connolly et al., 1996). Application of 3 U/ml of thrombin caused a
robust potentiation of responses to pressure-applied NMDA plus glycine
in wild-type C57Bl/6 mice that was significantly different at all times
from that observed in control experiments in which ACSF was applied
(Fig. 6). However, application of
thrombin to PAR1 / C57Bl/6 mice did not significantly increase NMDA
receptor responses. Series and membrane resistance were monitored
throughout the experiment and showed only modest changes that could not
account for the different effects of thrombin. As in rat hippocampal
slices, NMDA receptor response in wild-type and PAR1 / mice were
abolished by 100 µM APV (n = 9) after
thrombin treatment, suggesting that any potentiation observed reflected
NMDA receptor activation rather than sensitization of the cells to
glycine or pressure. Although these experiments provide direct evidence
for the involvement of PAR1 in thrombin-induced potentiation of NMDA
receptor function, modest levels of thrombin potentiation of NMDA
receptor responses still develop slowly in PAR1 / mice, becoming
significant only after thrombin application. This latent potentiation
in PAR1 / mice raises the possibility that thrombin cleavage
of as yet uncharacterized PARs or other signaling substrates in
hippocampal neurons might lead to modest levels of NMDA receptor
potentiation observed in PAR1 / mice.
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Figure 6.
A, B,
Left panels, Raw current traces from every fourth NMDA
application recorded at 70 mV (i.e., every 4 min) are shown for CA1
pyramidal cells from PAR1 +/+ C57Bl/6 mice (A)
and PAR1 / C57Bl/6 mice (B). Gray
boxes show periods of thrombin application. Calibration, 10 sec, 20 pA. Right panels, APV (100 µM) blocked NMDA responses recorded at 40
mV (calibration, 5 sec, 50 pA); NMDA responses reversed near 0 mV.
C, Mean time courses (±SEM) of thrombin (3 U/ml)-induced potentiation of NMDA responses in CA1 pyramidal cells
from PAR +/+ and PAR1 / C57Bl/6 mice were compared with that of
mice (PAR1 +/+, /) treated with ACSF. * indicates
significantly different from ACSF-treated slices
(p < 0.05; Kruskal-Wallis test; Dunn
post hoc test). There were minimal changes in the series
resistance or membrane resistance during the course of the experiment
(bottom panels).
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Potentiation of recombinant NMDA receptors by activation of the
thrombin receptor PAR1
To test whether PAR1 activation can directly alter NMDA receptor
function in recombinant systems, we injected Xenopus oocytes with cRNA encoding NMDA receptor subunits alone or together with cRNA
encoding the PAR1 receptor. The oocytes were placed under two-electrode
voltage clamp to record agonist-evoked currents at a membrane holding
potential of 30 mV in Mg2+-free
solutions (Fig. 7). Recordings were made
in a paired fashion such that two oocytes, one of which was coinjected
with PAR1, were recorded from simultaneously with the same solutions.
The presence of functional PAR1 receptor was confirmed by observation of the Ca2+-activated
Cl current during a 2-3 min application
of 0.03-3 U/ml (300 pM-30 nM) of thrombin. This inward current is observed
when Ca2+ is released from intracellular
stores after PAR1 activation of phosphoinositol-linked signaling
(Miledi, 1982; Miledi and Parker, 1984; Vu et al., 1991). NMDA receptor
responses in oocytes that were not coinjected with PAR1 cRNA
(n = 19) were not potentiated by thrombin application,
whereas oocytes that were injected with PAR1 cRNA typically displayed
thrombin-induced potentiation of the NMDA receptor-mediated current
(n = 23) (Fig.
7A,B). Potentiation of NR1-1a/NR2A
receptors coexpressed with PAR1 was observed after treatment with 300 pM (1.37 ± 0.08-fold; n = 7), 3 nM (1.68 ± 0.11-fold;
n = 6), or 30 nM thrombin (Fig.
7B). Low concentrations of thrombin (0.3-3
nM) that were capable of evoking PAR1-mediated potentiation of NMDA receptor function did not produce any cleavage of
NR1 subunit (Fig. 3), further suggesting that the potentiation that we
observed is independent of thrombin proteolysis of NR1.
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Figure 7.
A, Xenopus oocytes
were coinjected with NR1-1a/NR2B cRNAs or NR1-1a/NR2B/PAR1. Current
recordings were made under voltage clamp at 30 mV in the nominal
absence of extracellular Mg2+. The current-response
to 20 µM glutamate plus 10 µM glycine
(G in all panels) was potentiated after
activation of PAR1 by either 2.5 U/ml (25 nM) thrombin or
10 µM SFLLRN. Activation of PAR1 initiates an endogenous
Ca2+-activated Cl current; no
such responses were evoked by thrombin treatment of oocytes that were
not coinjected with PAR1 cRNA (n = 127; data not
shown). B, Although thrombin (2.5 U/ml)-stimulated or
SFLLRN (10-30 µM)-stimulated
Ca2+-activated Cl currents were
observed in oocytes coinjected with PAR1 and NR1 plus NR2A, NR2B, NR2C,
NR2D, or NR3A, only responses from NMDA receptors containing NR1 plus
NR2A, NR2B, NR2A/NR3A, or NR2B/NR3A were potentiated by activation of
PAR1 cDNA. C, Linear map showing alternative splice
variants of the NR1 subunit that were studied. NR1-4astop
was created by mutation of the codon for Q866 in the NR1 cDNA to a stop
codon. D, Alternative C- or N-terminal NR1 splicing or
creation of a new reading frame by use of an alternative splice site
within exon 22 does not markedly alter PAR1-mediated potentiation of
NMDA receptor responses. For all panels, * indicates
significantly different from oocytes without PAR1 coinjection
(p < 0.05 by t test); number
of oocytes indicated in parentheses. Data are mean ± SEM.
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To confirm that activation of PAR1 leads to NMDA receptor current
potentiation, we tested whether the PAR1-activating peptide (10-30
µM SFLLRN) could potentiate recombinant NMDA receptor
responses in oocytes coinjected with PAR1 cRNA. Oocytes expressing
NR1-1a/NR2B NMDA receptor subunits but not PAR1 had a ratio of the
NMDA receptor-mediated current after thrombin application to the
current response before thrombin application of 0.87 ± 0.04 (n = 9). By contrast, oocytes expressing the same NMDA
receptor subunits and PAR1 had a significantly larger ratio after
SFLLRN stimulation of PAR1 (Fig. 7A) (1.41 ± 0.11-fold
potentiation; p < 0.01; paired t test;
n = 9). Similar results were found for SFLLRN
potentiation of NR1-1a/NR2A receptors (1.85 ± 0.1-fold
potentiation; n = 6). These data showing SFLLRN potentiation of recombinant NMDA receptor responses rule out the possibility that thrombin cleavage of substrates other than PAR1 might
contribute to the observed NMDA receptor potentiation.
To evaluate the subunit dependence of PAR1 potentiation of NMDA
receptor responses, we coinjected oocytes with various subunit combinations. Figure 7C shows several NR1 splice variants
coexpressed with NR2B that were tested for their sensitivity to PAR1
potentiation of receptor function. The NR1-4astop mutation
eliminates 22 C-terminal amino acids included in the NR1-4a subunit as
a result of a frame shift associated with the use of an alternative
splice site in exon 22. Receptors containing all splice variants tested
were significantly potentiated by PAR1 activation (Fig. 7D).
The NR2 subunit had a more marked effect on PAR1 potentiation of NMDA receptor function. Oocytes coinjected with NR1-1a and either the NR2A
or NR2B subunit were potentiated by thrombin activation of PAR1 (Fig.
7B). Coexpression of NR3A with NR1-1a/NR2A or NR1-1a/NR2B subunits did not occlude PAR1-mediated potentiation. However, the
amplitude of NMDA receptor responses in oocytes coinjected with NR1-1a
and either NR2C or NR2D was unaffected by thrombin activation of PAR1.
These results suggest that only receptors that contain NR1/NR2A or
NR1/NR2B subunits can be potentiated by PAR1 activation. In addition,
these data are consistent with the idea that NR1 splicing alone does
not dominate the molecular determinants of PAR1 potentiation in our
Ba2+-containing recording solution (Zheng
et al., 1997; Logan et al., 1999).
The potentiation of recombinant NMDA receptor responses by thrombin
activation of PAR1 was independent of voltage over the range 80 to
+30 mV (n = 6; data not shown). The potentiation that
we observed occurred for responses to maximal concentrations of
glutamate and glycine, eliminating the possibility that potentiation reflects an increase in the glutamate or glycine
EC50. PAR1 potentiation could also be observed
for NR1-1a(C798A)/NR2B receptors (1.35 ± 0.07-fold;
n = 7; p < 0.05 by t test)
compared with control oocytes lacking PAR1 (0.98 ± 0.05;
n = 5), which rules out any contribution to the
potentiation that we observed of changes in reduction/oxidation state
of the disulfide linkage in NR1 (Sullivan et al., 1994). In addition,
PAR1 potentiation of recombinant NMDA receptor responses does not
significantly alter the ratio of current responses at pH 7.6 and pH 6.8 (Fig. 8C,D),
suggesting that PAR1 activation does not relieve tonic proton
inhibition (Traynelis and Cull-Candy, 1990). This result is consistent
with the lack of effect (Fig. 7) of alternate splicing of NR1 exon 5 on
PAR1 potentiation (Traynelis et al., 1995). Potentiation of
NR2A-containing receptors was observed both in the absence (Fig. 7) and
presence of 10 µM EDTA (2.01 ± 0.17-fold;
n = 12), eliminating any contribution of PAR1-mediated
relief of tonic inhibition by contaminant
Zn2+ in our recording solutions (Paoletti
et al., 1997; Zheng et al., 1998). Figure
8A,B shows that PAR1-mediated
potentiation did not reduce Mg2+ block of
recombinant NMDA receptors. The modest enhancement of Mg2+ blockade after thrombin activation of
PAR1 for NR1-1a/NR2B/NR3A receptors was not significantly different
from a modest enhancement of Mg2+
potentiation observed in thrombin-treated control oocytes that did not
express PAR1 (data not shown; n = 7), suggesting that it was not linked to PAR1 activation.
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Figure 8.
A, PAR1 activation by 3 U/ml of
thrombin (black bar) potentiated glutamate-evoked NMDA
receptor responses (gray bars) without altering
inhibition by 0.2 mM Mg2+ (open
bars). B, The effects of PAR1 activation on
inhibition by Mg2+ are summarized for different
combinations of NMDA receptor subunits. Number in
parentheses indicates the number of oocytes. The mean
inhibition by Mg2+ is shown and was calculated from
each oocyte as the ratio of the glutamate/glycine-evoked current during
Mg2+ application
(IMg) to the pretreatment current
(IO). Data are mean ± SEM. All oocytes showed robust potentiation of NMDA receptors responses
after PAR1 activation. C, PAR1 activation by the agonist
peptide 10 µM SFLLRN (black bar)
potentiated glutamate-evoked NMDA receptor responses
(gray bars) without significantly altering
inhibition produced by lowering pH from 7.6 (control) to 6.8 (open bars). D, The effects of PAR1
activation on proton inhibition are summarized for different
combinations of NMDA receptor subunits. Number in
parentheses indicates the number of oocytes. The mean
inhibition by pH 6.8 is shown and was calculated from each oocyte as
the ratio of the glutamate/glycine-evoked current at pH 6.8 (IpH 6.8) to the pretreatment current
(IpH 7.6). Data are mean ± SEM.
All oocytes showed robust potentiation of NMDA receptors responses
after PAR1 activation. For all panels, * indicates
p < 0.05 (t test).
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Thrombin potentiation of neuronal NMDA receptor function is
voltage dependent
Hippocampal NMDA receptors are under strong voltage-dependent
block by extracellular Mg2+, as
illustrated by the current-voltage curve from hippocampal CA1
pyramidal cells shown in Figure
9A. To investigate whether the
thrombin-induced potentiation of neuronal NMDA receptors is independent
of voltage-dependent Mg2+ blockade, we
compared the ratio of NMDA-evoked whole-cell currents recorded at 70
and 40 mV in control cells. We found that the ratio of current
recorded at 70 to 40 mV was significantly larger after thrombin
treatment when compared with pretreatment control (n = 22; Wilcoxon rank sum test; p < 0.01) (Fig.
9B). Moreover, in 16 of 21 CA1 pyramidal cells examined, the
potentiation of NMDA receptor responses by thrombin was larger at 70
mV (Fig. 9C, open symbols). Average peak
potentiation was 2.10 ± 0.29-fold at 70 mV and 1.48 ± 0.11-fold at 40 mV in 1.4 mM
Mg2+ (n = 19; Wilcoxon
rank sum test; p < 0.01) (Fig.
9D,E). Interestingly, the magnitude
of potentiation observed at 40 mV was virtually identical to that
observed in Xenopus oocytes in the absence of Mg2+ (Fig. 9E). We interpret
these data to suggest that there may be an additional voltage-dependent
component to the potentiation of neuronal NMDA receptors at
hyperpolarized potentials after thrombin activation of PAR1 that is not
present in Xenopus oocytes (Fig. 5) (Durand et al., 1993;
Wagner and Leonard, 1996; Zheng et al., 1997; Xiong et al., 1998). One
potential explanation of this result is that PAR1 activation modestly
reduces the voltage-dependent Mg2+ block
of neuronal NMDA receptors in addition to potentiating receptor
function in a voltage-independent fashion. Two previous reports of
PKC-mediated relief of Mg2+ blockade of
neuronal NMDA receptor responses (Chen and Huang, 1992; Zhang et al.,
1996) are consistent with this idea, because PAR1 is thought to couple
to signaling systems linked to the GQ family of
G-proteins, such as PKC (Grand et al., 1996). However, an
alternative explanation for the voltage dependence of PAR1 potentiation
of neuronal receptors is that PAR1-linked modification or thrombin
proteolysis of other channels or membrane proteins reduces our ability
to keep the dendrites under voltage control. The resulting reduction in
holding potential of distal dendrites during NMDA receptor activation
might modestly reduce NMDA receptor block by extracellular
Mg2+, causing an apparent supplemental
potentiation of the NMDA receptor response. Although we can detect no
thrombin-induced changes in membrane resistance and have blocked some
K+, Na+, and
Ca2+ channels with
Cs+, QX-314, and nifedipine, the complex
electrotonic structure of CA1 hippocampal pyramidal cells makes it
difficult to dismiss this alternative possibility. Ultimately,
single-channel studies in which individual
Mg2+-induced blockages of the NMDA
receptor are evaluated will be required to assess whether PAR1
activation also causes any reduction of
Mg2+ blockade.
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Figure 9.
A, The mean current-voltage
relationship from five outside-out patches excised from CA1 pyramidal
cells in hippocampal slices. In each patch, the voltage was ramped
between 100 and +30 mV (1-2 sec) between 10 and 50 times in the
absence and presence of 50 µM NMDA plus 10 µM glycine. The resulting traces from each patch were
averaged together and subtracted; data between patches were
subsequently averaged. The box highlights the region
analyzed in whole-cell recordings. B, The mean ratio
(±SEM) of the whole-cell current-response to pressure-applied
NMDA/glycine measured at 70 and at 40 mV is shown before
(Pre-treatment) and after 20-30 min of thrombin or
buffer treatment. * indicates significantly different
from pretreatment (p < 0.05; Wilcoxon rank
sum test). C, Plots show increased peak fold
potentiation at 70 mV compared with 40 mV for most cells studied
(open symbols); a few cells showed larger peak fold
potentiation at 40 mV (filled symbols).
Squares are data from rat, and circles
are data from mouse. D, The time course (±SEM) of the
fold potentiation recorded at both 70 and 40 mV in 19 cells in
response to the application of 3 U/ml (30 nM) thrombin.
E, The mean peak fold potentiation is shown after PAR1
activation for NR1-1a/NR2B receptors in Xenopus oocytes
recorded in the absence of extracellular Mg2+. Data
are from Figure 7; # indicates p < 0.05 (t test). Solid bars show the mean peak
fold potentiation for thrombin-treated hippocampal CA1 pyramidal cells,
compared with ACSF-treated cells (Buffer). *
indicates significantly different from buffer-treated
cells (p < 0.05; Mann-Whitney test); **
indicates significantly different from neuronal results obtained at
40 mV (p < 0.05; Wilcoxon rank sum
test).
|
|
| |
DISCUSSION |
In this study we have demonstrated under multiple experimental
paradigms that the serine protease thrombin at nanomolar concentrations can potentiate recombinant and hippocampal NMDA receptor function. The
potentiation of neuronal NMDA receptor responses appears to reflect
activation of the G-protein-coupled protease receptor PAR1, which is
differentially expressed throughout the CNS. Although high
concentrations of thrombin can directly cleave the hippocampal NR1
subunit, multiple lines of evidence suggest that the PAR1-mediated potentiation that we observed is independent of NMDA receptor proteolysis. These results hold intriguing implications for both neuronal development and neuropathological situations in which compromise of the blood-brain barrier provides access for
blood-derived serine proteases that are capable of activating PAR1 to
brain parenchyma. Furthermore, these results raise the possibility that expression of thrombin by neural tissue (Dihanich et al., 1991) might
act on PAR1 to influence NMDA receptor function.
PAR1 potentiation of NMDA receptors in normal brain function
The control of NMDA receptor function by serine proteases may play
a role in normal brain function, given the important contribution to
synaptic plasticity and neuronal development suggested for NMDA
receptors. A role for the serine proteases tPA in late-phase long-term
potentiation of excitatory synaptic function has recently been
suggested (Baranes et al., 1998; Zhuo et al., 2000), and our results
raise the possibility that PAR1 activation might influence the
contribution of NMDA receptor activation to various forms of synaptic
plasticity. PAR1 has been suggested to play a role in neuronal
development on the basis of thrombin's stimulation of neurite
retraction (Gurwitz and Cunningham, 1988) and reversal of astrocyte
stellation (Grabham and Cunningham, 1995). The ability of thrombin and
other serine proteases such as plasmin (Junge et al., 1999) to
potentiate NMDA receptor function and thus enhance Ca2+ signaling could be important for
several developmental processes that are influenced by NMDA receptors,
including synapse stabilization (Scheetz and Constantine-Paton, 1994),
neuronal survival (Ikonomidou et al., 1999), neuronal migration (Komuro
and Rakic, 1993), as well as growth cone guidance (Baird et al., 1996;
Zheng et al., 1996). Several additional studies provide support for
serine protease and PAR1 involvement in synaptic (Liu et al., 1994;
Baranes et al., 1998) and neuronal development (Debeir et al., 1998).
Thus, our results are consistent with a number of potential roles of serine protease signaling in the CNS.
PAR1 potentiation of NMDA receptor function in
pathological situations
It is now well accepted that NMDA receptor overactivation plays a
role in expanding the region of neuronal injury after experimental ischemia (Whetsell, 1996; Dirnagl et al., 1999; Lee et al., 1999). Thus, it is possible that thrombin entry into the brain during hemorrhagic stroke and penetrating head wound may contribute to neuronal damage through potentiation of NMDA receptor function. If
thrombin is generated in sufficient quantities to enter brain tissue in
excess of endogenous serine protease inhibitors such as PN-1 (Luthi et
al., 1997) during other cardiovascular insults that compromise
blood-brain barrier integrity (e.g., occlusive stroke, closed head
injury, status epilepticus) (Laursen et al. 1993; Barzo et al., 1997;
Correale et al., 1998), thrombin might contribute to the neuropathology
of these conditions through potentiation of NMDA receptors. Similarly,
if other serine proteases capable of activating brain PARs such as
plasmin (Ishihara et al., 1997) also potentiate NMDA receptor function
(Junge et al., 1999), they too may have harmful effects should they
reach brain parenchyma. Extravasated plasminogen has been suggested to
be cleaved by tPA in brain tissue to produce abnormally high levels of
plasmin, which may be important for neuronal damage after experimental ischemia or injection of kainic acid (Tsirka et al., 1995, 1997a,b; Wang et al., 1998; Kim et al., 1999). The effects of serine proteases and their receptors on neuronal function is a timely topic given the
current use of the protease tPA in treatment of occlusive stroke (Hacke
et al., 1995; Wardlaw et al., 1997).
The recent demonstration that thrombin entry into the brain can evoke
seizures (Lee et al., 1997) together with the thrombin-mediated potentiation of NMDA receptor responses described here suggests that
thrombin together with heme-derived iron (Willmore et al., 1978)
could be a contributing factor to post-traumatic seizures. Unprovoked
seizures complicate 7-34% of civilian and combat trauma patients, and
2-25% of patients with cerebrovascular disease or intracerebral
bleeding suffer seizures, with a higher incidence in hemorrhagic than
ischemic stroke (Berger et al., 1988; Faught et al., 1989; Lancman et
al., 1993; Arboix et al., 1997). Interestingly, the best indicators of
post-traumatic epilepsy include subdural hematoma and intracerebral
hemorrhage, two conditions that should lead to thrombin entry into the
brain (Lee and Lui, 1992; Annegers et al., 1998). Our data showing
thrombin potentiation of NMDA receptor function plus plasmin-induced
reductions in GABAergic transmission (Mizutani et al., 1997) are
consistent with the idea that serine proteases can control neuronal
excitability (Luthi et al., 1997). Thus, although more work is needed
to elucidate the details of protease signaling in the brain, our
results strengthen the idea that serine proteases and their receptors
have important roles in both normal and pathological situations.
Comparison of results with previous work
The PAR1-induced potentiation of neuronal NMDA receptor function
that we observed at 40 mV is similar to potentiation of neuronal NMDA
receptors after activation of various G-protein-linked receptor
systems, including m1 muscarinic receptors (Markram and Segal,
1990; Marino et al., 1998), metabotropic glutamate receptors (mGluRs)
(Aniksztejn et al., 1991, 1992), and µ opioid receptors (Chen and
Huang, 1991). In addition, the PAR1-mediated potentiation of
recombinant NMDA receptor responses in Xenopus oocytes is
similar to 5-HT2 and mGluR receptor potentiation
of recombinant NMDA receptors (Kelso et al., 1992; Blank et al.,
1996 ). The potentiation of NMDA receptor function after activation of
mGluRs, 5-HT2 receptors, and µ opioid receptors
has been suggested to require PKC (Aniksztejn et al., 1991, 1992; Chen
and Huang, 1991; Kelso et al., 1992; Blank et al., 1996).
Furthermore, purified PKC can potentiate (Chen and Huang, 1991, 1992;
Xiong et al., 1998) as well as phosphorylate NMDA receptors (Tingley et
al., 1993, 1997; Leonard and Hell, 1997). Moreover, the NR2 subunit
dependence of PAR1 potentiation is consistent with the subunit
dependence of phorbol ester potentiation of heteromeric NMDA receptor
function in oocytes (Kutsuwada et al., 1992; Mori et al., 1993; Grant
et al., 1998). These findings, together with the coupling of PAR1
receptors to the GQ family of G-proteins,
which can lead to PKC activation, make PKC an attractive candidate
pathway for PAR1 potentiation of neuronal NMDA receptor responses. Our
data showing that the protein kinase inhibitor BIS can block PAR1
potentiation of neuronal NMDA responses are also suggestive of a role
for PKC, although we cannot rule out involvement of other
serine/threonine kinases because the concentration of BIS used to
ensure penetration into the slice was nonselective. Moreover, it is
possible that other signaling systems that couple to PAR1 (Dery et al.,
1998), GQ and GI family of
G-proteins (Bence et al., 1997; Umemori et al., 1997), or PKC (Le
Good et al., 1998; Marais et al., 1998) might mediate the effects of
PAR1 on NMDA receptor function. Furthermore, kinase regulation of the
binding of intracellular proteins to glutamate receptors (Matsuda et
al., 1999) raises another level of potential complexity to the
signaling that we observed. Thus, more work is required to elucidate
the pathway(s) underlying PAR1 potentiation of NMDA receptor function.
| |
FOOTNOTES |
Received Jan. 18, 2000; revised March 28, 2000; accepted April 7, 2000.
This work was supported by the National Institute of Mental Health
(M.B.G.), and the National Institute of Neurological Diseases and
Stroke (S.F.T.), The John Merck Fund (S.F.T.), and the Emory University
Research Committee (S.F.T.). We thank Dr. R. Dingledine for critical
comments on this manuscript, and D. Falls, D. Wigston, and A. Levey for
helpful comments during the course of this project. We thank Drs. S. Coughlin, A. Levey, S. Heinemann, R. Huganir, S. Nakanishi, and P. Seeburg for sharing molecular reagents with us. We also thank Drs. J. Conn, J. Doherty, A. Levey, M. Marino, S. Rouse, and E. Marchan for
help with some of these experiments, Dr. F. Jaramillo for help with
imaging experiments, Dr. M. Hollmann for advice on isolated oocyte
membranes, and N. Ciliax, N. Patel, and J. Therien for excellent
technical assistance.
Correspondence should be addressed to Dr. Stephen Traynelis, Department
of Pharmacology, 5025 Rollins Research Center, 1510 Clifton Road,
Atlanta, GA 30322-3090. E-mail:
strayne{at}emory.edu.
| |
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ABSTRACT
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