Hypoxia modulates nitric oxide-induced regulation of NMDA
receptor currents and neuronal cell death
Muyiwa
Gbadegesin1,
Stefano
Vicini1,2,
Sandra J.
Hewett3,
David A.
Wink4,
Michael
Espey4,
Ryszard M.
Pluta5, and
Carol A.
Colton1,2
1 Interdisciplinary Program in
Neuroscience and 2 Department of
Physiology and Biophysics, Georgetown University Medical Center,
Washington, District of Columbia 20007;
3 Department of Pharmacology,
University of Connecticut Health Center, Farmington, Connecticut 06030;
and 4 Radiation Biology, National
Cancer Institute, and 5 Surgical
Neurology Branch, National Institute of Neurological Disorders and
Stroke, National Institutes of Health, Bethesda, Maryland 20892
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ABSTRACT |
Nitric oxide (NO)
released from a new chemical class of donors enhances
N-methyl-D-aspartate
(NMDA) channel activity. Using whole cell and single-channel
patch-clamp techniques, we have shown that
(Z)-1-[N-(3-ammoniopropyl)-N-(n-propyl)amino]-NO
(PAPA-NO) and diethylamine NO, commonly termed NONOates, potentiate the glutamate-mediated response of recombinant rat NMDA receptors (NR1/NR2A) expressed in HEK-293 cells. The overall effect is an increase in both peak and steady-state whole cell currents induced by
glutamate. Single-channel studies demonstrate a significant increase in
open probability but no change in the mean single-channel open time or
mean channel conductance. Reduction in oxygen levels increased and
prolonged the PAPA-NO-induced change in both peak and steady-state
glutamate currents in transfected HEK cells. PAPA-NO also enhanced cell
death in primary cultures of rodent cortical neurons deprived of oxygen
and glucose. This potentiation of neuronal injury was blocked by
MK-801, indicating a critical involvement of NMDA receptor activation.
The NO-induced increase in NMDA channel activity as well as NMDA
receptor-mediated cell death provide firm evidence that NO modulates
the NMDA channel in a manner consistent with both a physiological role
under normoxic conditions and a pathophysiological role under hypoxic conditions.
N-methyl-D-aspartate; oxygen-glucose deprivation; PAPA-NO
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INTRODUCTION |
THE N-methyl-d-aspartate
(NMDA) receptor is one of the subtypes of excitatory amino
acid receptors found on neurons in the central nervous system and plays
important physiological roles in both the fetal and adult central
nervous systems. Activation of the NMDA receptor affects processes
ranging from migration of neurons in the developing brain to learning
and memory (8, 15, 43, 53, 63). The NMDA receptor also mediates
pathophysiological states such as glutamate-mediated excitotoxicity,
which is commonly associated with ischemia-reperfusion injury,
persistent seizures, and chronic neurodegenerative disease (7, 11, 13,
27, 37, 67).
The precipitating events during excitotoxicity after
ischemia-reperfusion are linked to an influx of calcium, in
part via the NMDA receptor, and to the subsequent activation of a
variety of pathways inside the neuron (11, 13, 49). Increased
production of reactive oxygen species and reactive nitrogen species is
one of the early cellular responses (10, 12, 46, 50). Because the NMDA
receptor is known to contain regulatory sites that respond to changes
in oxidation and reduction (2, 14, 53), these sites provide a means by
which NMDA receptor function itself can be altered during the
ischemia-reperfusion events.
Oxidative stressors such as hydrogen peroxide and superoxide anion
decrease NMDA-mediated responses in a variety of preparations (2, 21,
53-56). However, exposure of the NMDA receptor to another major
class of reactive oxygen species, i.e., nitric oxide (NO), has produced
contradictory results. NO is produced in endothelial cells, neurons,
glia, and microglia/macrophages by a family of calcium/calmodulin-linked enzymes known as NO synthases (NOSs) (42).
The neuronal isoform of NOS (nNOS) is localized to postsynaptic regions
in many neurons in the central nervous system and is activated by an
influx of calcium (5, 8, 9, 16, 25). Mice deficient in nNOS have been
used to demonstrate the role of nNOS in ischemia-reperfusion injury and in excitotoxicity in general. Dawson et al. (17) have shown
that NMDA-mediated cell death is ameliorated in cortical cultures from
mice deficient in nNOS. Infarct size is reduced in
nNOS-deficient mice after cerebral ischemia (26, 48). Selective inhibition of nNOS by pharmacological agents such as 7-nitroindazole also reduces infarct size, reinforcing the idea that nNOS contributes to excitotoxic neuronal damage (24). However, recent in vitro studies
by Vidwans et al. (61) and Maynard et al. (39) demonstrate that NO can
protect against NMDA-mediated neuronal death most likely through
inhibition of NMDA receptor activity (20, 31, 33, 35, 61). The closure
of the NMDA channel in response to NO would necessarily preclude a
direct role for NO in NMDA-mediated excitotoxicity by this route.
Many of the studies examining the modulation of NMDA receptor function
by NO have relied on various chemical agents that release NO, i.e., NO
donors. The most common donors that have been used include sodium
nitroprusside (SNP), S-nitrosocysteine
(SNOC), S-nitroso-N-acetylpenicillamine
(SNAP), and 3-morpholinosydnonimine (SIN-1). Exposure of neurons to
these agents results in a reduction in NMDA receptor responses (20, 31,
33, 35, 61). However, NO may not be the primary species produced by
these donors, since other reactive nitrogen species that act as
nitrosonium ion (NO+) donors
have been implicated in their actions (33). In support of this, the
actual level of NO generated by SIN-1, SNOC, or SNAP is very low (62).
Recently, a new generation of pure NO donors, the NONOates, has been
developed, which produce up to 15 µM NO with variable
half-lives ranging from seconds to hours (36, 62). NONOates, therefore,
can serve as a reliable, long-lasting source of NO. The aim of the
present study was to characterize the effects of the NO generated by
this new class of agents on NMDA receptor activity and neuronal cell
death under normoxic and hypoxic conditions.
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METHODS |
Cell cultures. Murine mixed cortical
cell cultures were prepared as described in Refs. 22 and 61. First,
cerebral cortices of 1- to 3-day-old mouse pups (CD-1; Charles River)
were removed under sterile conditions, minced, and incubated for
20-30 min (37°C) in 0.025% trypsin diluted in Hanks'
balanced salt solution supplemented with glucose (15 mM) and sucrose
(20 mM). Cells were pelleted by light centrifugation (1,850 rpm, 3 min), the supernatant was discarded, and the cell pellet was
resuspended in plating medium of modified Eagle's medium (Earle's
salts; Mediatech) supplemented with 2 mM glutamine, 20 mM glucose
[media stock (MS)], 10% fetal bovine serum (FBS), 10%
horse serum (HS), and epidermal growth factor (10 ng/ml). The cells
were plated in 24-well dishes (Primaria; Falcon) at a density of 1 hemisphere · plate
1 · 10 ml
1. Once confluent
(9-11 days), the cells, primarily astrocytes, were shifted into a
maintenance medium (MS + 10% HS).
Cortical neurons were obtained in a similar manner from embryonic
day 15 (E15) fetuses and then plated
at a density of 3 hemispheres per plate per 10 ml on an established bed
of astrocytes (12-24 days in vitro) in MS supplemented with 5% HS
and 5% FBS. After 5-7 days in vitro, cultures were exposed to 10 µM cytosine arabinoside for 2 days to eliminate any mitotic cells
(microglia, oligodendrocytes). Cells were then shifted into maintenance
medium, and the medium was changed twice weekly. Experiments were
performed on mixed cultures between 14 and 16 days in vitro. All
cultures were kept at 37°C in a humidified 5%
CO2-containing atmosphere.
Human embryonic kidney-293 (HEK-293) cells (ATCC No. CRL1573; American
Type Culture Collection, Manassas, VA), were grown at 37°C in a 6%
CO2-94% air atmosphere in MEM
(GIBCO BRL, Gaithersburg, MD), supplemented with 10% FBS and 100 U/ml
penicillin-streptomycin (GIBCO BRL).
Transfection of NMDA receptor
subunits. Exponentially growing HEK-293 cells were
dispersed with trypsin 15-20 h in advance of transfection,
resuspended at 2 × 105 cells
in 1.5 ml of culture medium, and plated on
poly-L-lysine (Sigma, St. Louis,
MO)-coated coverslips. HEK-293 cells were transfected with rat NMDA
receptor cDNAs using the calcium phosphate precipitation method as
described in Ref. 60. Mixed plasmids (3 µg total) coding for NR1a,
NR1b, and NR2A subunits of the NMDA receptor were added to the dish
containing the 1.5-ml culture medium. cDNA for pGreenLantern-1 protein
(GIBCO BRL) was cotransfected with the NMDA receptor cDNAs to allow for
visualization of the transfected cells. Greater than 90% of the cells
that expressed GreenLantern protein also expressed NMDA receptors. The
transfection mixture was removed after 8-10 h, the cells were
washed twice with PBS, and then the PBS was replaced with fresh culture
media containing 500 µM ketamine to reduce glutamate-mediated cell
toxicity. Transfected cells were allowed to equilibrate overnight
before use in the experimental protocols. Experiments were performed on
a minimum of three different transfected cell cultures
(n = the number of cells studied).
Statistical significance was determined using an unpaired or paired
Student's t-test according to the
experimental conditions.
Electrophysiology. With the use of the
patch-clamp technique, transfected HEK-293 cells were voltage clamped
at
60 mV in the whole cell or outside-out patch configuration.
Electrodes were pulled in two stages on a vertical pipette puller from
borosilicate glass capillaries (Wiretrol II; Drummond, Broomall, PA).
Typical pipette resistance was 5-7 M
. Intracellular (patch
pipette) solutions contained (in mM) 145 potassium gluconate, 5 EGTA, 2 MgCl2, 10 HEPES, 2 NaATP, and 0.2 NaGTP, pH to 7.2 with KOH. The cells were bathed in extracellular
solution containing (in mM) 145 NaCl, 5 KCl, 1 CaCl2, 5 HEPES, and 5 glucose. The
extracellular solution contained saturating concentrations of glycine
(10 µM).
All drugs were applied to the bath using a gravity-fed Y-tube delivery
system placed within 500 µm of the cell under study. The tip of the
Y-tube had a diameter of ~100 µM and allowed solution exchange in
<20 ms. In addition, the chamber (0.5 ml total volume) was
continually perfused at a rate of ~4-5 ml/min. All drugs were dissolved in the same extracellular solution used to bathe the cells.
For experiments in low oxygen (hypoxia), control solution or solutions
containing glutamate or glutamate plus the test drugs were vigorously
aerated with 100% N2 at ambient
pressure in separate enclosed and vented chambers. These solutions were
then used to perfuse the transfected HEK cells. Exact oxygen tensions
were not determined under these conditions.
Data acquisition and analysis. Whole
cell currents were monitored with an EPC-7 patch-clamp amplifier (List
Electronics, Darmstadt/Eberstadt, Germany), filtered at 1.5 kHz with an
eight-pole low-pass Bessel filter (Frequency Devices, Haverhill, MA),
and digitized with Axotape 2 software (Axon Instruments, Foster City,
CA). Clampfit (Axon Instruments) was used for off-line analysis, and
Origin (MicroCal Software, Northampton, MA) was used for figure
preparation and statistical analysis. Single-channel recordings were
filtered at 1.5-2.5 kHz and digitized at 10-20 kHz. Analysis
and curve fitting for the single-channel recordings were performed
using Fetchan and pStat analysis programs (Axon Instruments). The
threshold for measuring open and shut intervals was set at one-half the maximal amplitude of the main conductance level of the channel. Log-binned interval histograms for shut and open duration as well as
burst duration were plotted with a square root vertical axis and fitted
using maximum likelihood fitting (52). For display purposes, traces
were filtered at 50 Hz.
NO donors.
(Z)-1-[N-(3-ammoniopropyl)-N-(n-propyl)amino]-NO
(PAPA-NO) and diethylamine-NO (DEA-NO) were gifts from Dr. David Wink
(National Institutes of Health) or were obtained from Alexis Biochemical (San Diego, CA).
2-(4-Carboxyphenyl)-4,4,5,5-tetramethylimidazoline-1-oxyl-3-oxide (C-PTIO; potassium salt) and SIN-1 were obtained from Alexis
Biochemicals. All NONOates were prepared as stock solutions of 10 or
100 mM in alkaline solutions and were diluted into the desired media immediately before use. Expended solutions were prepared by allowing the NO donor solutions to sit for 24 h at room temperature.
Combined oxygen-glucose deprivation.
Cultures were placed in an anaerobic chamber (Forma Scientific,
Marietta, OH) that contained a gas mixture of 5%
CO2-10%
H2-85%
N2 (<0.2%
O2). Culture media were replaced
by thorough exchange with a deoxygenated, glucose-free balanced salt
solution alone or a balanced salt solution containing PAPA-NO
(0.3-10 µM) and then were placed in a 37°C humidified incubator within the chamber for 40-45 min. Cultures were
subsequently removed from the incubator, and 100 µl of cell culture
supernatant were collected to assess early cell death using the lactate
dehydrogenase (LDH) assay (see below). Exposure medium was then
exchanged with oxygenated MEM, and the cells were returned to a
normoxic (21% O2) incubator at
37°C containing 5%
CO2. Delayed neuronal
death was assessed 20-24 h later. The balanced salt solution
contained (in mM) 116 NaCl, 5.4 KCl, 0.8 MgSO4, 1 NaH2PO4,
26.2 NaHCO3, 1.8 CaCl2, and 0.01 glycine.
For exposure under normoxic conditions, cultures were exposed for 45 min to PAPA-NO (0.3-10 µM) diluted into MS alone (or in the
presence of 10 µM NMDA) at 37°C in a 5%
CO2-containing incubator. Cultures
were subsequently removed from the incubator, and 100 µl of cell
supernatants were collected for assessment of early neuronal injury.
The exposure medium was then exchanged with fresh MS and the cells
returned for an additional 20-24 h, whereupon delayed neuronal
death was determined.
Assessment of neuronal cell death.
Neuronal cell death was estimated by examination of cultures under
phase-contrast microscopy and quantitatively assessed by the
measurement of LDH released by damaged or destroyed cells into the
bathing medium following treatment. LDH activity was quantified by the
rate of oxidation of NADH, which was followed spectrophotometrically at
340 nm (29). The small amount of LDH present in the medium of sister
cultures subjected to sham wash (generally <15% of total) was
subtracted from the levels in experimental conditions, to yield the LDH
signal specific to experimental injury. Data are expressed as the
percentage of total neuronal LDH (=100%) that was determined for each
experiment by assaying the supernatant of sister cultures after 24 h of
exposure to 300 µM NMDA.
NO measurement. NO release from the NO
donors was measured continuously for 12 h with an electrochemical probe
(Diamond General) placed in a 3-ml sample of the perfusion media. After
stabilization, the probe was calibrated against known concentrations of
NO with the use of in vitro methods previously described (38). NO stock solutions used for calibration of the probe were prepared by bubbling NO into a solution of 1 M NaOH to eliminate other nitrogen oxide species and then through a sealed container with deoxygenated normal
saline. The concentration of NO in the stock solution was confirmed
using colorimetric methods.
 |
RESULTS |
Whole cell response to PAPA-NO. The
effects of the NO donors PAPA-NO and DEA-NO on NMDA receptor activity
were studied in HEK-293 cells transiently transfected with NMDA NR1a,
NR1b, and NR2A subunit cDNAs. Glutamate (30 µM) was applied in 10-s
pulses to the cells and whole cell currents were recorded. As shown in Fig. 1 for HEK cells
transfected with NR1a and NR2A subunits, the evoked currents
demonstrated slow onset and offset kinetics similar to those recorded
using native NMDA receptors (40). On average, the peak responses of
NR1a/NR2A transfected cells to 30 µM glutamate were 300 ± 74 pA
(SE; n = 8 cells), whereas average
steady-state currents, determined after 10 s of application of agonist,
were 217 ± 48 pA (n = 8). The
response to glutamate was reexamined in the presence of the NO donors
by adding the donor directly to the glutamate-containing perfusion
media and applying this mixture in 10-s pulses onto the cell using the
Y-tubing system. Because NO production by PAPA-NO rises quickly then
falls over 20 min (Fig. 1B), PAPA-NO
was added to the glutamate solution <1 min before application to the
cell. This insured maximal NO generation. The presence of glutamate and
PAPA-NO (200 µM) significantly increased both the peak current and
the steady-state current compared with glutamate alone. The effect of
PAPA-NO was reversible by removing the NO donor from the perfusion
media or by the simultaneous addition of C-PTIO, an NO scavenger that
rapidly and irreversibly inactivates NO (Fig.
1A) (4). DEA-NO, a similar NO
donor but with a shorter half-life than PAPA-NO (~7 min) also
produced an increase in glutamate-mediated whole cell currents (Table
1).

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Fig. 1.
Modulation of whole cell
N-methyl-D-aspartate
(NMDA) receptor responses by
(Z)-1-[N-(3-ammoniopropyl)-N-(n-propyl)amino]-nitric
oxide (PAPA-NO). NMDA whole cell responses were elicited in HEK-293
cells transiently transfected with the NMDA receptor subunits NR1a and
NR2A. Traces indicate responses from a typical transfected HEK cell to
30 µM glutamate applied via a Y-tube.
A: responses to sequential application
of 30 µM glutamate (GLUT) and 30 µM glutamate + 200 µM PAPA-NO.
Responses to glutamate + PAPA-NO were elicited 40 s after addition of
PAPA-NO to glutamate solution. A 2-min wash period was interposed
between each application of drugs. Right
trace: removal of effect of PAPA-NO by addition of
2-(4-carboxyphenyl)-4,4,5,5-tetramethylimidazoline-1-oxyl-3-oxide
(C-PTIO), an NO scavenger. Bars, time of drug application.
B: electrochemical detection of NO
production by PAPA-NO over time. NO production by PAPA-NO was measured
in separate control experiments using an NO-sensitive electrode in same
media as that used for electrophysiological recordings.
C: whole cell current recordings
obtained from a representative HEK cell transiently transfected with
NMDA receptor subunits NR1a and NR2A in response to 30 µM glutamate
in presence and absence of 1 mM 3-morpholinosydnonimine (SIN-1). Cells
were washed for 2 min between each drug application and were voltage
clamped at 60 mV. D:
electrochemical detection of NO production from SIN-1 using an
NO-sensitive electrode. Measurement of NO production from SIN-1 was
carried out in same media used for electrophysiological recordings.
Final concentration of SIN-1 was 1 mM.
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Table 1.
Effects of PAPA-NO and DEA-NO on whole cell currents of cells
transfected with different NMDA receptor subtypes
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The putative NO donor SIN-1 did not generate NO under the same
conditions as PAPA-NO and did not increase the glutamate-mediated response in transfected HEK cells (Fig. 1,
C and
D). In contrast, the presence of 1 mM SIN-1 decreased the whole cell currents produced by application of
30 µM glutamate to the bath.
To rule out the possibility that the amine backbone of PAPA-NO mimicked
the glycine-independent potentiation of the NMDA receptor by spermine,
we tested the effect of PAPA-NO on whole cell currents in HEK-293 cells
transfected with the NR1b and NR2A subunit cDNAs. The NR1b splice
variant contains a 21-amino acid
NH2-terminal insert generated by
exon 5 and shows no potentiation by spermine at saturating
concentrations of glycine (66). If PAPA-NO or the other NONOates mimics
the effect of spermine, potentiation should not be observed in
recombinant receptors containing the NR1b subunit splice variant. As
shown in Table 1, recombinant NMDA receptors containing the splice
variant demonstrated increased whole cell currents in response to
PAPA-NO. The average control peak current value for the NR1b/NR2A
variant was 296 ± 141 pA (n = 5), and the average control
steady-state current value was 207 ± 84 pA
(n = 5). In the presence of 200 µM
PAPA-NO, these values were increased by 33 ± 15% and 33 ± 14%, respectively, compared with glutamate alone (Table 1). The NR1b
splice variant was chosen for all subsequent experiments, thereby
eliminating both the spermine and pH sites involved in NMDA channel
regulation (57) and allowing a more direct analysis of the effect of NO.
Control studies were done to examine the effect of expended PAPA-NO on
the response to pulses of 30 µM glutamate (Fig.
2). PAPA-NO solutions equilibrated at
25°C for 24 h induced a depression of 27 ± 5%
(n = 10) in glutamate-mediated whole
cell currents in cells transfected with the NR1a/NR2A and NR1b/NR2A
subtypes, indicating that the increased response of the NMDA receptor
induced by PAPA-NO was not due to the parent molecule.

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Fig. 2.
Expended PAPA-NO inhibits NMDA receptor activity.
A: whole cell current recordings
obtained from a representative HEK cell transiently transfected with
NR1a/NR2A NMDA receptor subunit in response to sequential application
of 30 µM glutamate and 30 µM glutamate + expended PAPA-NO. PAPA-NO
solution was expended for 24 h at room temperature.
B: average reduction in whole cell
currents compared with wash in response to expended PAPA-NO.
* P < 0.05 (n = 10 cells).
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Modulation of NMDA single-channel activity by
NO. To examine the NO-mediated effect on NMDA receptor
function in more detail, we determined the effect of PAPA-NO on NMDA
channel activity in outside-out patches excised from transfected
HEK-293 cells. A representative recording is shown in Fig.
3 and demonstrates that, under control
conditions (5 µM glutamate, 10 µM glycine), the open probability
was 0.023 and, in the presence of 100 µM PAPA-NO plus glutamate, the
open probability increased to 0.041. The increased open probability
returned to the untreated value when the NO scavenger, C-PTIO, was
added to the perfusion media. The average open probability in the
presence of glutamate alone was 0.029 ± 0.008 (n = 11 patches) and was 0.072 ± 0.02 (n = 11) for PAPA-NO plus
glutamate.

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Fig. 3.
Modulation of NMDA single-channel activity by NO. Representative
current traces obtained in an outside-out patch from a transfected
HEK-293 cell in presence of 5 µM glutamate alone
(A); 5 µM glutamate + 100 µM
PAPA-NO (B); and 5 µM glutamate + 100 µM PAPA-NO + 100 µM C-PTIO
(C). Holding potential was 60
mV and the extracellular solution contained 10 µM glycine. PAPA-NO
was added to glutamate solution ~40 s before application to the patch
via the Y-tube. Note increase in burst duration in presence of PAPA-NO.
D: open probability
(Po) in the
patch during above treatments.
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Distributions of channel open time and burst duration were also
determined and are shown for a representative patch in Fig. 4, A-D.
PAPA-NO did not alter the distribution of channel open times (Fig. 4,
A and
B). Average values of the mean open
times for 11 patches were 3.7 ± 0.4 and 4.1 ± 0.4 ms in the
presence of glutamate alone and glutamate plus PAPA-NO, respectively.
Determination of the distribution of burst durations revealed an
increase in burst duration (Fig. 4, C
and D). The average burst duration
changed from 8 ± 1 ms (n = 8 patches) in glutamate alone to 15 ± 3 ms (n = 8 patches) in the presence of
PAPA-NO. Current-voltage relationships were also obtained for the main
conductance state over a range of membrane potentials from
80 to
+80 mV (Fig. 4F). Conductance and
reversal potential values for the main conductance state of the
channels under control conditions were 51.6 pS and 3.8 mV, respectively, whereas, for PAPA-NO-treated channels, conductance and
reversal potential of the main conductance state were 50.8 pS and 3.4 mV, respectively. Thus no change in single-channel conductance was
observed.

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Fig. 4.
Effect of NO on NMDA receptor single-channel parameters.
A and
B: single-channel open time was
determined from a representative transfected HEK cell in presence of 1 µM glutamate alone. Distribution of 473 open intervals was fit with 2 exponentials with time constant and relative areas of 0.43 ms (27.2%)
and 2.95 ms (72.7%), respectively
(A). Distribution of 1,700 open
intervals obtained from same patch as above in presence of 100 µM
PAPA-NO + 1 µM glutamate. Distribution was fit with 2 exponentials
with time constants and relative areas of 0.44 ms (21%) and 3.02 ms
(79%) (B). Intervals plotted in
these histograms were measured from currents analog filtered at 2.5 kHz
and sampled at 20 kHz. Glycine concentration = 10 µM.
C and
D: effect of NO on burst duration
recorded from a different cell in response to 1 µM glutamate in
presence and absence of 100 µM PAPA-NO. Distribution had duration and
relative areas of 1.46 ms (52.3%) and 6.90 ms (47.6%) for glutamate
alone (C) and 1.38 ms (44.9) and
9.67 ms (55.1) for glutamate + PAPA-NO
(D). Glycine concentration = 10 µM. E: effect of PAPA-NO on average
(±SE) percent change compared with control of channel open
probability, mean open time, and average burst duration. Burst analysis
was performed only on patches with no double openings. All patches were
from NR1a/NR2A transfected cells.
* P < 0.001 (n = 11).
F: current
(I)-voltage
(V) relationship for mean
single-channel currents in presence and absence of 100 µM PAPA-NO.
Amplitude measurements were taken from a total of 4 patches in control
glutamate (50 nM to 1 µM) solutions ( ) and solutions containing
glutamate + 100 µM PAPA-NO ( ).
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Effect of low oxygen tensions on the changes in
glutamate-mediated whole cell currents by PAPA-NO. To
determine whether hypoxic conditions altered the response of the NMDA
receptor to NO, ambient oxygen tension in the solutions was decreased.
For these experiments, the glutamate or glutamate plus PAPA-NO
solutions perfusing the cells was vigorously and continuously agitated
with 100% N2 immediately before
application to the cells. In contrast to the previous experiments where
30 µM glutamate was used, PAPA-NO (200 µM) was added in the
presence of saturating concentrations of glutamate (200 µM). For that
reason, the potentiation of the peak current induced by PAPA-NO was not
pronounced. However, use of saturating concentrations of glutamate
allowed discernment of the effect of PAPA-NO in normoxic and hypoxic
conditions. In addition, use of saturating concentrations of glutamate allowed the study of the effect of PAPA-NO on
desensitization, which appeared to be a possible mechanism for the
action of PAPA-NO. This mechanism was suggested by the effect of
PAPA-NO on burst duration in the single-channel studies (see
DISCUSSION). The relative change in
the response to PAPA-NO was followed over time, and average values were
obtained by determining the ratio of the sequential responses to
glutamate plus PAPA-NO to the average response to glutamate alone. Only
cells with stable control responses to repetitive glutamate pulses were
used in these experiments. As shown in Fig. 5, the effect of PAPA-NO on both the peak
and steady-state current in response to 5-s pulses of 200 µM
glutamate is enhanced in a low-oxygen environment. A comparison of the
normalized values for peak and steady-state currents in normoxic and
hypoxic conditions is shown in Fig. 5,
C and
D. A significant
(P < 0.01) increase in both the peak
and steady-state current was seen initially but decayed with time.

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Fig. 5.
Changes in effect of NO in a low-oxygen environment.
A: effect of PAPA-NO on
glutamate-induced whole cell currents in an ambient oxygen (normoxic)
environment. Traces represent typical recordings from NR1b/NR2A
transfected cells. Glutamate (200 µM) applications were for 5 s
followed by a 5-s wash period in presence and absence of 200 µM
PAPA-NO. PAPA-NO was added to glutamate-containing solution and was
thus also applied in 5-s intervals with a wash between each
application. B: typical glutamate
responses (5 s, 200 µM) from a different cell in presence and absence
of 200 µM PAPA-NO in a low-oxygen (hypoxic) environment.
C and
D: relative change in response to
glutamate induced by exposure to PAPA-NO was followed over time. Values
were obtained by determining an average control (untreated) response to
glutamate and then determining the ratio of individual responses to the
average control response for a sequential series of responses to
glutamate in presence of PAPA-NO. Data points represent average ± SE relative change for 5 cells under normoxic condition and for 4 cells
under hypoxic conditions. C: average
relative peak currents
(Ipeak).
D: average relative steady-state
currents (Isteady
state). * P < 0.05.
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NMDA-mediated neuronal injury in normoxic and hypoxic
conditions with PAPA-NO. The relationship between the
increase in NMDA channel response induced by exposure to PAPA-NO and a
downstream functional end point, i.e., neuronal injury, was also
investigated. First, cortical neurons in primary culture were exposed
to PAPA-NO (0.3-10 µM) for 45 min under normoxic conditions to
determine whether it was directly cytotoxic. Under these control
conditions, exposure to PAPA-NO alone produced no early (data not
shown) or late cortical neuronal cell death over the concentration
range studied (Fig.
6A).
Furthermore, PAPA-NO did not enhance cell death induced by low doses of
NMDA (10 µM; Fig. 6B). However,
when the cortical cultures were deprived of oxygen and glucose in the
presence of PAPA-NO, cell killing was significantly increased over that produced by oxygen-glucose deprivation (OGD) alone (Fig.
7A). The
expended NONOate had no effect on OGD-mediated killing (data not
shown), indicating that the PAPA-NO-mediated enhancement of cell death
was not due to a nonspecific effect of a metabolic product produced by
PAPA-NO. To determine whether cell death in the low-oxygen, low-glucose
condition was dependent on a functional NMDA receptor response, the
irreversible NMDA antagonist MK-801 was utilized. As shown in Fig.
7B, the PAPA-NO-mediated enhancement of OGD-induced cell death was prevented by the addition of MK-801 (10 µM) to the exposure medium.

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|
Fig. 6.
PAPA-NO is not toxic to primary cortical neurons under normoxic
conditions. A: primary cortical
cultures were exposed for 45 min to PAPA-NO (0.3-10 µM) under
normoxic conditions. PAPA-NO was removed by washing, and cell death was
assessed immediately using the lactate dehydrogenase (LDH) assay (none
detected; data not shown) as well as 20-24 h later. Data points
represent mean value of LDH release (±SE) for each condition scaled
to that induced by exposure of neurons to 300 µM NMDA for 24 h (taken
as = 100; n = 11-13 wells assayed
from 4 separate culture groups). B:
cortical cultures were exposed to a low dose of NMDA (10 µM) in
presence and absence of PAPA-NO (0.3-10 µM) and assayed for cell
death using the LDH assay immediately following washout (none detected;
data not shown) as well as 20-24 h later. Data points represent
mean value of LDH release (±SE) for each condition scaled to that
induced by exposure of neurons to 300 µM NMDA for 24 h (taken as = 100; n = 19-22 wells assayed from
6 separate culture groups). PAPA-NO did not significantly alter
NMDA-induced cell death as determined by ANOVA.
|
|

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|
Fig. 7.
PAPA-NO potentiates oxygen and glucose deprivation-induced neuronal
injury. A: primary cortical cultures
were deprived of oxygen and glucose in absence and presence of varying
concentrations of PAPA-NO (0.3-10 µM). At 20-24 h after 45 min of oxygen-glucose deprivation (OGD), LDH release into the bathing
medium was measured. Data are expressed relative to LDH signal
corresponding to neuronal death in sister cultures (taken as = 100;
measured after exposure to 300 µM NMDA for 24 h).
* P < 0.01 as assessed by
ANOVA followed by Dunnett's t-test
(n = 12-20 wells assayed for 5 separate culture groups). B: cell
cultures were deprived of oxygen and glucose alone or in presence of
NMDA receptor antagonist MK-801 (10 µM) alone, PAPA-NO (10 µM)
alone, or MK-801 (10 µM) + PAPA-NO (10 µM). At 20-24 h after
40-45 min of oxygen-glucose deprivation, LDH release into the
bathing medium was measured. Data are expressed relative to the LDH
signal corresponding to neuronal death in sister cultures (taken as = 100; measured after exposure to 300 µM NMDA for 24 h).
* P < 0.001 as assessed by
ANOVA followed by Student-Newman-Keuls test
(n = 7-8 wells assayed for 3 separate culture groups).
|
|
 |
DISCUSSION |
NO released from well-characterized, time-dependent, pure NO donors
such as the NONOates enhances NMDA channel activity. Using whole cell
and single-channel patch-clamp techniques, we have shown that the two
NONOates studied, i.e., PAPA-NO and DEA-NO, potentiate
glutamate-mediated responses of recombinant rat NMDA receptors
expressed in HEK-293 cells. The overall effect is an increase in both
the peak and steady-state currents induced by glutamate. Single-channel
kinetics demonstrate a significant increase in open probability and an
increase in burst duration but no change in channel open time or
single-channel conductance. The effect of NO could be readily reversed
by NO scavengers, and the addition of expended NONOate (both PAPA-NO
and DEA-NO) produced a slight depression in whole cell currents rather
than the enhancement seen in the presence of NO. The action of PAPA-NO
was highly time dependent and could be visualized most clearly within
10 min of addition to the cells. The falloff in activity most likely
represents the time-dependent release of NO from the NONOates. The
polyamine-like backbone that remains after release of NO could be
responsible for the inhibition of the NMDA response seen with expended
NONOate solutions, since polyamines induce a voltage-dependent
inhibition of the NMDA channel (40).
The enhancement of NMDA channel whole cell and single cell responses
are in contrast to the effect on the NMDA channel of other NO donors
reported in the literature. Previous studies with donors such as SNAP
(25), SIN-1 (35), SNOC (31), and SNP (25, 31, 47) have demonstrated
that the NMDA receptor is inhibited by NO. One potential explanation
for the discrepancy is that reactive species other than NO are produced
downstream of NO, which inhibit rather than potentiate receptor
activity (33). NO is well known to undergo a number of rapid
interactions with oxygen, forming reactive nitrogen-oxygen species
(RNOs) that may themselves have effects on a variety of molecules and
reactions within the cell (64, 65). RNOs can react with glutathione or
with sulfhydryl groups on proteins and lipids, producing independent effects on the NMDA receptor. In our experiments, the action of SIN-1
on NMDA channel activity is consistent with the action of these other
NO species. Using an NO electrode in separate control studies, we
showed that SIN-1 does not release measurable amounts of NO over 12 h.
SIN-1 has been reported, however, to produce superoxide anion. Thus any
NO generated by SIN-1 would rapidly combine with superoxide anion to
form the powerful oxidizing agent peroxynitrite (59). Colton et al.
(14), Tang and Aizenman (55, 56), and others (30, 54) have demonstrated
that oxidation depresses NMDA channel function and that both whole cell
currents and single-channel properties are reduced under oxidative
conditions. SIN-1 in our study inhibits whole cell glutamate-mediated
currents, a result that is consistent with oxidation of the NMDA channel.
Both the whole cell and single-channel studies show a direct effect of
NO released from PAPA-NO on the NMDA channel. This is apparent from the
potentiation of the peak and steady-state current at the whole cell
level and the increase in open probability and burst duration at the
single-channel level. The exact site of NO action in our study is not
clear. Aizenman and Potthoff (3) have indicated that NO does not appear
to act on Cys-744 and Cys-798, the redox sites commonly associated with
the inhibition of NMDA channel function by oxidation. These authors
suggest that inhibition by NO utilizes an alternative site. Our data
also indicate alternative sites of action for NO. In this case,
NO-induced potentiation of the NMDA response may be due to a decrease
in desensitization, since burst duration is increased. Lester and Jahr
(32) have demonstrated that an increase in burst duration represents
entry of the NMDA receptor into a relatively nondesensitizing state.
NO-mediated enhancement of NMDA receptor activity under aerobic
conditions is likely to be a normal physiological function of NO. In
our experiments, the increase in whole cell currents in response to
PAPA-NO is generated using rapid and short-duration application pulses
of the NO donor. Although the exact concentration of NO is not known,
the cell is exposed in a transient fashion to low levels of NO. This
exposure pattern to NO is reminiscent of the action of nNOS in the
hippocampus and is consistent with a physiological role for NO. For
example, NO has been implicated in the regulation of synaptic
transmission and is released from postsynaptic neurons in a
calcium/calmodulin-dependent fashion (8, 12, 51). One potential role,
as proposed by Bredt et al. (8) and others (43, 51), is
that NO serves as a retrograde messenger, strengthening synaptic
connections during long-term potentiation. Localization of nNOS to the
synaptic region, as indicated by Brenman et al. (9) and Aoki et al.
(5), would provide easy access of NO to the NMDA receptor. Because the
overall level of NO is low, direct effects of NO would dominate (19, 65). Consistent with a physiological modulatory role, our data demonstrate that neither NO-mediated cell death nor an enhancement of
NMDA-induced neuronal injury occurs under these conditions. However,
during ischemia, the fall in oxygen level may significantly alter the response to NO, allowing a pathophysiological state to ensue.
Hypoxic conditions alter the effect of PAPA-NO on the NMDA receptor.
Reduction in oxygen levels increased and prolonged the PAPA-NO-induced
change in both peak and steady-state glutamate currents in the
transfected HEK cells. Although the mechanism of this effect is
unclear, separate studies using mixed cortical cultures indicate that
the action of PAPA-NO in a low-oxygen environment may have functional
consequences. Cell death mediated by oxygen and glucose deprivation was
increased by exposure to low concentrations of PAPA-NO. MK-801-induced
blockade of the enhanced toxicity further suggests that the effect of
PAPA-NO in OGD-treated cultures is due to activation of the NMDA
channel. An increased NMDA receptor activity would also provide an
explanation for the association between increased astrocytic inducible
NOS activity and increased neuronal death seen during OGD
(23). Alternatively, one might argue that NO-mediated enhancement of
OGD-induced neuronal injury could result from alterations in astrocyte
glutamate uptake. However, NO derived from DEA-NO at concentrations up
to 10 mM had no effect on glutamate uptake by purified or recombinant
high-affinity glutamate transporters (58). By contrast, peroxynitrite
and oxygen radicals effectively prevented glutamate uptake (58). Under
hypoxic conditions, the formation of peroxynitrite or other oxidizing
species is low. Because our data clearly demonstrate that neuronal cell
death is enhanced in the presence of low oxygen, i.e., under those
conditions where formation of RNOs is minimal, a direct effect on the
NMDA receptor is favored.
The source of NO in hypoxic episodes is not clear, but direct
measurement of NO using NO electrodes during rat carotid occlusion has
shown a burst of NO at the onset of the occlusion, followed by a slow
decline (34). A variety of studies using different techniques have also
indirectly implicated that NO is generated during the early phase of
ischemia (6, 18, 44, 45). Although continued lack of oxygen, as
would occur in the core of an ischemic infarct, would preclude the
generation of NO by NOS (1), the low oxygen tension that is maintained
in the penumbral region could be enough to sustain NO formation.
Further, the return of oxygen with reperfusion would promote a
different chemical profile of NO species, including the
generation of oxidizing RNO compounds that might be expected to
decrease NMDA channel function via its redox regulation site (31, 33).
Thus, although they may have other direct cytotoxic effects, the
formation of peroxynitrite or other oxidizing species could be
protective due to their ability to inhibit the NMDA receptor. Further
generation of NO by NOS during continued reperfusion and the return of
aerobic conditions could have beneficial actions due to its ability to
scavenge · OH and to reduce deleterious
reactions such as lipid peroxidation (12, 28, 41, 61, 63). However, the
initial anoxic events and the potential role of NO in
glutamate-mediated calcium entry during this time period may have
already initiated irreversible damage.
 |
ACKNOWLEDGEMENTS |
This work was supported in part by National Institutes of Health
Grants AG-16026-01 (to C. Colton) and NS-36812-01 (to S. J. Hewett).
 |
FOOTNOTES |
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. §1734 solely to indicate this fact.
Address for reprint requests and other correspondence: C. Colton, Dept.
of Physiology, Georgetown Univ. Medical School, 3900 Reservoir Rd. NW,
Washington, DC 20007 (E-mail: glia01{at}aol.com).
Received 4 March 1999; accepted in final form 3 June 1999.
 |
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