* Cellular Neurobiology Laboratory, The Salk Institute for Biological Studies, La Jolla, California 92037; and Department of
Cell Biology, The Scripps Research Institute, La Jolla, California 92037
Glutathione depletion occurs in several forms of apoptosis and is associated with Parkinson's disease and HIV toxicity. The neurotransmitter glutamate kills immature cortical neurons and a hippocampal nerve cell line via an oxidative pathway associated with glutathione depletion. It is shown here that soluble guanylyl cyclase (sGC) activity is required for nerve cell death caused by glutathione depletion. Inhibitors of sGC block glutamate toxicity and a cGMP analogue potentiates cell death. Glutamate also induces an elevation of cGMP that occurs late in the cell death pathway. The resultant cGMP modulates the increase in intracellular calcium that precedes cell death because sGC inhibitors prevent calcium elevation and the cGMP analogue potentiates the increase in intracellular calcium. These results suggest that the final pathway of glutamate induced nerve cell death is through a cGMP-modulated calcium channel.
GLUTATHIONE is a cysteine-containing tripeptide ( GSH depletion is observed in models of apoptotic cell
death, including glucocorticoid-induced thymocyte apoptosis (Beaver and Waring, 1995 The steps linking GSH depletion to neuronal cell death
are largely unknown. We have been studying this pathway
using glutamate treatment of immature cortical neurons
and a neuronal cell line. Recently, we found that depletion
of GSH activates neuronal 12-lipoxygenase (12-LOX), which
leads to the production of reactive oxygen species, the increase in intracellular Ca2+, and ultimately to cell death
(Li et al., 1997 Cell Cultures
Mouse hippocampal HT22 cells (Davis and Maher, 1994 Cell Viability Assay
Cell viability was determined by either visual cell counting or MTT assays
in 96-well plates. The MTT assays measure the ability of cells to metabolize 3-(4,5-dimethyldiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT)
and is often used to measure cellular proliferation (Mosmann, 1983 Glutathione, cGMP, and Ca2+ Measurements
For the glutathione, cGMP, and Ca2+ measurements to be described below, HT22 cells were seeded at 5 × 105 cells/100 mm and the next day
were treated according to the experimental design. Cells were then collected at indicated times for the above assays. Glutathione was determined by the recycling assay based on the reduction of 5,5-dithiobis(2-
nitrobenzoic acid) with glutathione reductase and NADPH (Tietze, 1969 For cGMP determination, HT22 cells were treated with glutamate for
various times, collected, and resuspended in 0.1 N HCl. After 1 h on ice,
the samples were centrifuged in a microfuge. The supernatant was neutralized and assayed for cGMP content using an EIA cGMP assay kit from
Amersham International (Buckinghamshire, England). The pellet was
dissolved in 0.1 N NaOH, and protein content was determined using a
commercial kit from Pierce Chemical Co. (Rockford, IL). cGMP content
was calculated per milligram protein and presented relative to the controls.
The intracellular ionized calcium concentration was determined by
flow cytometry using ratiometric analysis. HT22 cells were loaded with 1 µM
Indo-1 at 37°C for 30 min in the presence of 0.005% Pluronic F-127 in
DME containing 10% FCS. After incubation, cells were collected,
washed, and resuspended in phenol red-free Hepes-buffered DME supplemented with 2% dialyzed FBS. Cells were allowed a 15-min recovery
period to hydrolyze the ester bond before being analyzed with a FACStarplus® flow cytometer (Becton Dickinson, Mountain View, CA). The fluorescence light from the two emission peaks of Indo-1, 410 nm (violet) and 485 nm (green), was collected, and the ratio of violet to green, which is
proportional to Ca2+ concentrations (Grynkiewicz et al., 1985 Reagents
Tissue culture reagents were obtained from GIBCO BRL (Gaithersburg,
MD). The fluorescent dye Indo-1 acetoxymethylester was obtained from
Molecular Probes (Eugene, OR). LY83583, NG-methyl-L-arginine, nifedipine, NG-nitro-L-arginine, 7-nitroindazole, and tin protophorphorin IX were from LC Laboratories (Woburn, MA). 8-(4-chlorophenylthio) guanosine-3 Inhibitors of sGC Prevent Glutamate-induced Neuronal
Cell Death
The addition of glutamate to the hippocampal cell line
HT22 causes a rapid depletion of GSH, which in turn activates 12-LOX, leading to a form of programmed cell death
that is similar to but distinct from apoptosis (Tan, S., M. Wood, and P. Maher, manuscript submitted for publication). It has recently been shown that the products of
12-LOX enzymatic activity are required for oxidative glutamate toxicity (Li et al., 1997
To determine whether the above observations with
HT22 cells reflect the mechanism of oxidative glutamate
toxicity on central nervous system cells, embryonic rat cortical neurons were prepared, and 1 d later the cells were
exposed to glutamate as with HT22 cells. These young cultures are devoid of functional glutamate receptors and are
killed by nonreceptor-mediated glutamate-induced oxidative stress (Murphy and Baraban, 1990 Inhibitors of sGC Also Prevent Neuronal Cell Death
Induced by Buthionine Sulfoximine
Since the activation of 12-LOX has been experimentally
linked to the reduced level of intracellular GSH (Li et al.,
1997
sGC Inhibitors Do Not Affect the Ability of Glutamate
or BSO to Deplete GSH
To determine whether sGC inhibitors block the toxicity of
glutamate and BSO by affecting their ability to deplete
GSH, we compared intracellular glutathione levels in HT22
cells treated with these agents as a function of time. Fig. 2
shows that sGC inhibitor LY83583 did not block the depletion of GSH by either glutamate or BSO. None of the
other sGC inhibitors blocked glutathione depletion either
(data not shown). In the case of BSO treatment, the addition of LY83583 accelerated the depletion of GSH (Fig. 2). This is probably due to the fact that LY83583 is also capable of inhibiting glutathione reductase (Luond et al.,
1993 The Activation of sGC Occurs Near the Time of
Cell Death
Since sGC inhibitors block toxicity, it is likely that the enzyme is activated to produce cGMP. To determine if there
is an increase in the level of cGMP, intracellular cGMP
levels of HT22 cells treated with 5 mM glutamate for various
times were assayed. Fig. 3 A shows that the level of cGMP
starts to increase 8 h after glutamate treatment. Under the
conditions used, cells also begin to die at ~8 h after the addition of glutamate. The increase in cGMP was prevented
by treatment of cells with 1 µM LY83583 (data not shown).
Therefore, sGC activation occurs near the time of cell death.
To independently determine when in the cytotoxicity
pathway sGC is activated, HT22 cells were treated with
glutamate, and at various times afterwards LY83583 was
added, followed by a cell viability assay at 20 h. Alternatively, glutamate was added and then removed at various
times after its initial addition to the cultures. LY83583 was
able to inhibit glutamate toxicity even when applied up to
8 h into the cell death process (Fig. 3 B). The protection
conferred by LY83583 started declining at 8 h after addition of glutamate, which is the same time that glutamate
was no longer required for cell death (Fig. 3 B). After ~8
h of exposure to glutamate, HT22 cells gradually initiate
cell lysis, which is complete after ~15 h. However, surviving cells can still be rescued by LY83583 even during the
late period of cell death. For example, if 1 µM LY83583 is
added to cultures when 73% of cells are dead, the remaining 27% of the cells survive when examined 24 h later.
These observations suggest that the activation of sGC occurs at a point that is very close to cell lysis.
A Cell-permeable cGMP Analogue Potentiates
Glutamate Toxicity
If cGMP is involved in the cell death pathway, then exogenous cGMP should potentiate glutamate toxicity. The ability
of CPT-cGMP, a cell permeable and phosphodiesterase-resistant cGMP analogue, to potentiate glutamate-induced
cell death was examined. Overnight exposure of HT22
cells to 2 mM glutamate led to a 35% decrease in cell viability (Fig. 4 A). Coapplication of CPT-cGMP resulted in
further decreases in cell viability in a dose-dependent manner. For example, 2 mM glutamate caused a 35% cell death,
while 500 µM CPT-cGMP together with 2 mM glutamate
caused nearly a 90% decrease in cell viability (Fig. 4 A).
At 500 µM, CPT-cGMP alone was not significantly toxic
to HT22 cells. However, marked cell death was observed
when these cells were treated with CPT-cGMP at concentrations over 1 mM (Fig. 4 A, inset). Application of the less
stable cGMP analogues, dibutyl-cGMP and 8-bromo-cGMP,
did not result in cell death at concentrations up to 5 mM.
The potentiation by CPT-cGMP was dependent on glutamate concentration (Fig. 4 B) but was largely unaffected
by the time when CPT-cGMP was added up to 12 h after glutamate addition (Fig. 4 C). This result is consistent with the above data, suggesting that cGMP functions near the
time when cells die.
cGMP-dependent Protein Kinase Is Not Involved in
Glutamate Toxicity
Since the molecular targets of cGMP action include cGMP-gated ion channels and cGMP-dependent protein kinases
(PKG) and phosphodiesterases (Goy, 1991 sGC/cGMP Regulates the Elevation of Ca2+
Another potential target for cGMP is the activation of
Ca2+ channels, which play an important role in oxidative
glutamate toxicity. The removal of extracellular Ca2+ or
the addition of the Ca2+ channel blocker CoCl2 inhibits
glutamate toxicity (Murphy et al., 1989
It was then asked if there is a temporal relationship between sGC activation and Ca2+ elevation. Cells were exposed to LY83583 and CoCl2 at various times after the addition of glutamate, and cell viability was assayed 20 h
after the initial addition of glutamate. Fig. 3 B shows that
LY83583 and CoCl2 function in parallel in terms of maximal protection. Both agents maintain maximal protection
up to 8 h past the addition of glutamate, after which cell lysis occurs gradually over the next few hours. Therefore,
both sGC activation and Ca2+ elevation occur near the
time of cell death.
If sGC activation is required for Ca2+ elevation, then its
inhibition should block the glutamate-induced accumulation of intracellular Ca2+. We therefore investigated the
effect of sGC inhibitors on the intracellular Ca2+ profile
during the glutamate treatment. The intracellular Ca2+ response to glutamate was first determined by flow cytometry using ratiometric analysis. Intracellular Ca2+ content is
proportional to the ratio of fluorescence intensities from
the two emission peaks of the Ca2+-bound and unbound
Indo-1 dye (Grynkiewicz et al., 1985 sGC-mediated Glutamate Toxicity Is through a
Mechanism Independent of NO and Carbon Monoxide
Since several of the inhibitors used above do not inhibit
the basal activity of sGC (see Discussion), the activation of
sGC is probably not due to its transcriptional upregulation, but rather due to the induction of sGC activators. NO
and carbon monoxide are activators of sGC. If they are responsible for the sGC activation observed in the cell
death, these activators should also be a target for blocking
glutamate toxicity. Therefore, we determined if the inhibition of their synthesis is able to block glutamate toxicity.
Various inhibitors of NO synthase (NOS) were tested at
several concentrations. NG-monomethyl-L-arginine methyl
ester, a specific inhibitor of NOS, did not attenuate glutamate or BSO toxicity in HT22 cells or in primary cortical
neurons (Fig. 1). Other NOS inhibitors, including 7-nitroindazole (a specific inhibitor for neuronal NOS-1) (up to
0.25 mM), NG-methyl-L-arginine (up to 1.0 mM), and NG-nitro-L-arginine (up to 1.0 mM), also did not show any
protective effect on glutamate toxicity (data not shown).
Similarly, tin protoporphyrin, an inhibitor of heme oxygenase and carbon monoxide production, only had minimal
effects on glutamate toxicity at near toxic concentrations
(data not shown).
The data presented above indicate a requirement for sGC/
cGMP in nerve cell death caused by GSH depletion. First,
three structurally unrelated inhibitors of sGC all block
glutamate or BSO toxicity (Fig. 1) at concentrations that
inhibit sGC activity or cause cGMP reduction in vitro. Second, glutamate induces an increased production of cGMP
(Fig. 3). Third, a membrane-permeable cGMP analogue,
CPT-cGMP, potentiates glutamate toxicity (Fig. 4). Lastly, sGC inhibitors block the glutamate-induced increase in intracellular Ca2+ required for toxicity (Fig. 5). Therefore,
the generation of cGMP and the subsequent activation of
a Ca2+ channel are events occurring in GSH depletion-
induced nerve cell death.
Of the sGC inhibitors used here, hydroxylamine and its
analogue, N-methyl-hydroxylamine, competitively bind to
the heme group associated with sGC, inhibiting sGC activation (Deguchi et al., 1978 Both cytoprotective and cytotoxic roles for cGMP have
been described in the nervous system. For example, cGMP
prevents motor neuron degeneration (Weill and Greene,
1984 It is likely that cGMP is responsible for the increase in
intracellular Ca2+ required for glutamate toxicity. The elevation of cGMP occurs late in the death pathway (Fig. 3),
and the increase in intracellular Ca2+ also occurs late.
HT22 cells start to die ~8 h after the addition of glutamate, while sGC inhibitors are able to rescue cells even when the majority of cells are dead, suggesting that sGC
activation occurs very shortly before actual cell death. The
elevation of Ca2+, which was blocked by sGC inhibitors,
occurs only in 10-20% of cells at any time during the period of disintegration (Fig. 5 B), suggesting that Ca2+ accumulation immediately precedes cell death in individual
cells. Additional evidence supporting the relevance of
Ca2+ elevation with sGC/cGMP comes from the observations that: (a) CPT-cGMP potentiates the elevation of
Ca2+ induced by glutamate (Fig. 5 A); (b) other potential
regulators of cGMP such as PKG (Fig. 4) and cGMP-regulated phosphodiesterases are not involved in cell death;
and (c) inhibitors of the other classes of identified Ca2+
channels are unable to prevent cell death. Therefore,
HT22 cells probably possess an unidentified cGMP-modulated Ca2+ channel (Finn et al., 1996 Ca2+ is known to be involved in programmed cell death
(McConkey and Orrenius, 1994 In summary, it is shown in a clonal cell line and primary
cortical neurons that the elevation of cGMP, which likely
results from sGC activation, is a critical step in the nerve
cell death caused by GSH depletion. The activation of sGC
is via an NO-independent mechanism. cGMP then induces
Ca2+ influx, which immediately precedes cell death. If the
observation can be extended to in vivo situations, then intervention of the sGC/cGMP pathway could be beneficial
to individuals suffering from PD or other pathologies associated with GSH depletion.
-glutamylcysteinylglycine) that exists in both the
reduced (GSH)1 and the oxidized states (GSSG).
GSH is the predominant form within the cell, usually accounting for greater than 99% of the total glutathione
(Meister and Anderson, 1983
). GSH plays an important role in protecting cells from oxidative damage and regulates several aspects of cellular metabolism. The synthesis
of GSH is regulated by the enzyme
-glutamyl-cysteine
synthetase and its precursor molecule, cysteine, which is
present at low concentration within the cell. Low levels of
intracellular GSH are linked to a variety of pathological
conditions, such as HIV (Herzenberg et al., 1997
) and Parkinson's disease (Perry et al., 1982
; Sofic et al., 1992
; Sian
et al., 1994
). This association appears to be significant because HIV-infected lymphocytes that contain decreased
intracellular GSH are more likely to undergo apoptosis
(Staal et al., 1992
; Ameisen et al., 1995
). Artificially elevating GSH by N-acetylcysteine prolongs the survival of the
HIV-infected cells (Herzenberg et al., 1997
). In the nervous system, there is an early and highly specific decrease
in GSH in the substantia nigra of Parkinson's disease patients (Perry et al., 1982
; Sofic et al., 1992
; Sian et al.,
1994
), and the artificial depletion of GSH both induces dopaminergic neuronal cell death in vitro (Jenner and Olanow, 1996
) and potentiates the toxicity of 6-hydroxydopamine and 1-methyl-4-phenylpyridinium (MPP+) in
vivo (Pileblad et al., 1989
; Wullner et al., 1996
). Together, these data suggest that GSH depletion directly contributes
to cell death in a variety of cell types.
; Slater et al., 1995
) and
anti-Fas/APO-1-induced T lymphocyte apoptosis (van den
Dobbelsteen et al., 1996
). Several mechanisms account for
the depletion of GSH. (a) The production of reactive oxygen species and the increased requirement for glutathione peroxidase may lead to the consumption of GSH. (b) The
activation of a transmembrane export channel appears to
be responsible for the decline in intracellular GSH during
apoptosis in lymphocytes (van den Dobbelsteen et al.,
1996
) and U937 human monocytic cells (Ghibelli et al., 1995
).
(c) When persistently exposed to the neurotransmitter
glutamate, immature cortical neurons, which do not have
ionotropic glutamate receptors, are depleted of intracellular GSH through the competition by glutamate for cystine
uptake (Murphy and Baraban, 1990
; Murphy et al., 1990
).
Elevated levels of extracellular glutamate interfere with
the exchange of glutamate with cystine through a cystine/
glutamate antiporter, which normally carries cystine into
the cell. This leads to a decrease in the level of intracellular cystine and its reduction product cysteine, thereby causing a decrease in the level of GSH. This mechanism,
called oxidative glutamate toxicity, has also been described in a hippocampus nerve cell line HT22 (Davis and
Maher, 1994
), in a nerve-glial hybrid cell line N18-RE-105
(Murphy et al., 1989
), and in primary oligodendrocytes
(Oka et al., 1993
). When artificially depleted of GSH, immature cortical neurons also undergo programmed cell death (Ratan et al., 1994
).
). The experiments outlined below extend
these studies to show that the soluble guanylyl cyclase (sGC)/
cGMP pathway is involved in nerve cell death caused by
GSH depletion. This investigation was predicated upon
the following observations. First, Ca2+ elevation is required for oxidative glutamate-induced neuronal cell death
since the removal of extracellular Ca2+ prevents cell death
(Murphy et al., 1989
; Davis and Maher, 1994
) and the intracellular Ca2+ level increases during cell death (Li et al.,
1997
). One possible regulator of Ca2+ elevation is cGMP
since cGMP can directly or indirectly modulate some ion
channels (Yau, 1994
; Kaupp, 1995
; Finn et al., 1996
). Second, LOX activation is required for glutamate toxicity (Li et al., 1997
), and LOX metabolites activate sGC (Snider et
al., 1984
; Brune and Ulrich, 1991). This paper reports the
first evidence for a role of sGC in mediating programmed
cell death caused by glutathione depletion.
Materials and Methods
) are derived from
the HT4 cell line (Morimoto and Koshland, 1989
) and are propagated in
DME (Vogt and Dulbecco, 1963
) supplemented with 10% FBS. Primary
cortical neurons were prepared from embryonic day 17 Sprague-Dawley
rats as described (Abe et al., 1990
). After dissociation from brain tissues
with trypsin and DNase I, cells were maintained in MEM supplemented
with 30 mM glucose, 2 mM glutamine, 1 mM pyruvate, and 10% FBS.
). In
this system, it correlates with cell viability as determined by trypan blue
exclusion and a colony-forming assay (Davis and Maher, 1994
). The assay
medium contained 5% dialyzed FBS for the glutamate toxicity assay or
horse serum for the assay of buthionine sulfoximine (BSO) toxicity (Li et
al., 1997
). Fresh preparations of BSO were used. For the MTT assay, cells
were dissociated and seeded into 96-well microtiter plates at a density of
2.5 × 103 or 5 × 104 cells per well in 100 µl medium for HT22 cells and cortical neurons, respectively. The next day cells were treated with various
reagents according to the experimental design. 20 h after the treatment, 10 µl of MTT solution (5 mg/ml) was added to each well and incubated for
3 h. 100 µl of solubilization solution (50% dimethylformamide, 20% SDS,
pH 4.8) was then added to the wells, and the next day the absorption value
at 570 nm was measured. Results are expressed relative to the indicated
controls and were analyzed using a Mann-Whitney test.
).
Sample preparation and assay procedures were described elsewhere (Li et
al., 1997
).
), was obtained. 10,000 viable cells were analyzed in each assay.
-5
-cyclic monophosphate (CPT-cGMP) was obtained from Biolog
(La Jolla, CA). Other reagents, including L, D-buthionine sulfoximine, hydroxylamine, methylene blue, N-methyl-hydroxylamine, and NG-monomethyl-L-arginine methyl ester, were purchased from Sigma Chemical Co.
(St. Louis, MO).
Results
). One target for LOX
metabolites is sGC (Snider et al., 1984
; Brune and Ulrich,
1991). To determine if sGC is involved in glutamate toxicity, we first tested the effect of various inhibitors of sGC
on the survival of HT22 cells after exposure to glutamate.
HT22 cells were incubated with 5 mM glutamate in the
presence of several concentrations of the inhibitors for 20 h.
Cell viability was then determined by MTT reduction, a viability assay that correlates in this system with trypan blue
exclusion and colony formation assays (Davis and Maher, 1994
). Under these conditions, glutamate alone caused the
complete lysis of cells as assayed by both MTT reduction
(Fig. 1 A) and microscopic examination. Treatment of the
cells with 6-anillino-5,8-quinolinedione (LY83583), a specific inhibitor of sGC (Mulsch et al., 1988
), inhibited the
toxicity of glutamate in a concentration-dependent manner (Fig. 1 A and data not shown). The IC50 for the inhibition of glutamate toxicity was 0.5 µM, which is in good
agreement with the concentration of LY83583 required
for sGC inhibition in isolated tissues (Mulsch et al., 1988
).
Inhibitors with very different structures from LY83583, including methylene blue (Gruetter et al., 1981
), hydroxylamine, and N-methyl-hydroxylamine (Deguchi et al., 1978
),
all inhibited glutamate-induced cell death in a dose-dependent manner (Fig. 1 A and data not shown). The concentrations giving the maximal protection were all in the ranges
that inhibit the activation of the purified sGC and/or block
cGMP elevation in isolated tissues (Deguchi et al., 1978
;
Gruetter et al., 1981
). Although these structurally unrelated inhibitors may have other effects on cells, their only
shared target is sGC.
Fig. 1.
sGC inhibitors prevent nerve cell death caused by
glutamate and BSO. Experiments were performed as described
in the Materials and Methods. Results are expressed as relative to
controls treated with agents alone. The results shown are the
mean ± SD of a typical experiment with five determinations.
*Significantly different from glutamate treatment (P < 0.01, Mann-Whitney test). Similar results were obtained in three independent experiments. (A) Dose effect of sGC inhibitors and NOS
inhibitor NG-monomethyl-L-arginine methyl ester (NAME) on
glutamate toxicity in HT22 cells. LY, LY83583; MB, methylene
blue; NMH, N-methyl-hydroxylamine; HA, hydroxylamine. The
concentrations given above each compound are in micromolar.
The glutamate concentration used was 5 mM. Note that more
than five doses were tested but only the optimum and a lower
dose were presented. (B) Dose effect of sGC inhibitors and
NAME on glutamate toxicity in rat cortical neurons. (C) Dose effect of sGC inhibitors and NAME on BSO toxicity in HT22 cells. BSO was added to the growth medium at 50 µM. (D) Dose effect
of sGC inhibitors and NAME on BSO toxicity in rat cortical neurons.
[View Larger Version of this Image (38K GIF file)]
; Murphy et al.,
1990
). Fig. 1 B shows that the sGC inhibitors LY83583,
methylene blue, and N-methyl-hydroxylamine also block
glutamate toxicity in primary neuronal cell cultures. These
results are identical to those obtained with HT22 cells, except that the optimal concentrations of the inhibitors are
lower. Hydroxylamine itself was toxic to primary cortical
neurons, which is probably because hydroxylamine gives rise to nitric oxide (NO). Several other NO-releasing compounds induce apoptosis in cortical neurons (Palluy and
Rigaud, 1996
). The above results suggest that sGC is a critical enzyme in oxidative glutamate toxicity.
), it was asked if GSH depletion is also sufficient to
activate sGC-mediated cell death. To determine if sGC is
also involved in neuronal cell death associated with GSH
depletion, the effect of sGC inhibitors on BSO toxicity was
examined. BSO, a specific inhibitor of
-glutamylcysteine synthetase, the rate-limiting enzyme in GSH synthesis,
depletes intracellular GSH and causes cell death in a time-
and dose-dependent manner in both HT22 cells and primary cortical neurons (Li et al., 1997
). An overnight exposure of HT22 cells or primary cortical neurons to 50 µM
BSO causes glutathione depletion (Fig. 2) and a dramatic loss
of cell viability (Fig. 1, C and D). sGC inhibitors LY83583,
methylene blue, and N-methyl-hydroxylamine all block BSO
toxicity in a manner similar to that of glutamate, suggesting that sGC is directly linked to neuronal cell death caused
by GSH depletion. There are, however, differences in the
pharmacology of sGC inhibitors that prevent glutamate-
and BSO-induced cell death. These differences may be
due to the fact that at least the early events are different
for these two models of cell death, because glutamate interferes with the cystine uptake while BSO directly inhibits GSH synthesis.
Fig. 2.
sGC inhibitor
LY83583 does not prevent
the depletion of glutathione
by glutamate or BSO. HT22
cells were treated with either 5 mM glutamate or 50 µM
BSO in the presence or absence of 2 µM LY83583 for
various times. The cells were
collected and assayed for glutathione levels and protein
content. The data are presented relative to controls (0 h, no treatment). The level of glutathione was 12.4 ± 0.62 nmol/mg
protein and 8.08 ± 0.08 nmol/mg protein for cells grown in FCS-containing medium and horse serum-containing medium, respectively. Horse serum was used when BSO toxicity was studied
because it gave more sensitive and consistent results (Li et al., 1997).
[View Larger Version of this Image (27K GIF file)]
) and therefore the regeneration of GSH from GSSG.
These data, showing that sGC inhibitors prevent cell death
but not GSH depletion, support the previous argument
that GSH depletion caused by glutamate is not in itself
sufficient to cause cell lysis (Murphy et al., 1989
).
Fig. 3.
Glutamate induces
the increased production of
cGMP. (A) Time course
analysis of cGMP production. HT22 cells were treated
with 5 mM glutamate for various times, collected, and assayed for cGMP and protein
levels. Data are presented
relative to controls. The cGMP level for control cells
is 190 ± 23 fmol/mg protein.
Similar results were obtained
in two independent experiments, each with multiple
samples. (B) The ability of
various reagents to block glutamate toxicity as a function of time when they are
added to medium. HT22 cells
were treated with 5 mM
glutamate, and at various times after the addition, reagents were added to the medium and cell viability was
assayed next day. Cell viability is presented relative to controls without any treatment.
Glutamate, 5 mM; LY83583, 2 µM; CoCl2, 50 µM; Glu, replacement with normal DME. Similar results were obtained in
two independent experiments. Note that cell viability was compared with untreated controls 20 h after the initial seeding. HT22
cells have a short doubling time and grow to nearly four times the
original cell number in 20 h. The apparent initial low viabilities
conferred by the protectants reflect a lower rate of proliferation
rather than survival.
[View Larger Version of this Image (16K GIF file)]
Fig. 4.
The cell-permeable cGMP analogue CPT-cGMP potentiates glutamate-induced cell death. (A and B) Dose effect of
CPT-cGMP and glutamate on cell viability. HT22 cells were
treated overnight with glutamate and/or CPT-cGMP at various
concentrations, and viability was determined by the MTT assay.
Results, presented relative to controls without any treatment, are
the mean ± SD of a typical experiment with three determinations. Similar results were obtained in three independent experiments. (A, inset) Dose effect of CPT-cGMP on cell viability.
HT22 cells were treated for 20 h with CPT-cGMP at various concentrations, and viability was determined. Results are presented
relative to untreated controls. (C) A time course of the potentiation effect afforded by CPT-cGMP. Cells were treated with 2 mM
glutamate, and at various times after the addition, 500 µM CPT-cGMP was added. 24 h after the addition of glutamate, cell viability was assayed. Similar results were obtained in three independent experiments, each with three determinations. (D) A specific
inhibitor of protein kinase G, KT5823, does not prevent or attenuate cell death induced by glutamate. Experiments were performed as described in the legend to Fig. 1.
[View Larger Version of this Image (35K GIF file)]
), it is expected
that the inhibition of sGC will affect the activity of these
targets. If they are in the cell death pathway, then their inhibition will also block cell death. We examined whether
PKG is involved in cell death using KT5823, a highly selective inhibitor of PKG with an IC50 of 0.23 µM (Ito and
Karachot, 1990
). Fig. 4 D shows that KT5823 was unable
to block cell death even at concentration up to 100 µM,
suggesting that PKG is not involved in the cell death
caused by GSH depletion.
; Davis and Maher,
1994
), and glutamate induces the elevation of intracellular
Ca2+ (Li et al., 1997
). These data suggest that the increased
intracellular Ca2+ is derived from extracellular medium
rather than intracellular calcium pools. Initial experiments
were done to identify the specific channels involved in cell
death. The influx of Ca2+ may be via voltage-dependent
channels, cGMP-regulated channels, Na+/Ca2+ exchangers, or nonspecific membrane leakage. Membrane leakage is unlikely since CoCl2 prevents cell death (Murphy et al.,
1989
; Davis and Maher, 1994
). Replacement of Na+ by Li+
did not prevent cell death (data not shown), which excludes the involvement of a Na+/Ca2+ exchanger. Various
inhibitors of Ca2+ channels were then tested for their ability to prevent cell death induced by glutamate. These included the L-type channel inhibitors nifedipine, dilitizam,
verapamil, and nimodipine, and the N-type channel inhibitor
-conotoxin GVIA. None of these was protective at
concentrations up to toxic doses (data not shown). Therefore, it is unlikely that L- or N-type channels or Na+/Ca2+
exchangers are involved in the cell death. The involvement
of a cGMP-regulated Ca2+ channel can not be tested biochemically because no specific inhibitors exist (Finn et al.,
1996
). Although pimozide and D600 can block these channels in some cases (Finn et al., 1996
), they were not effective in preventing cell death caused by glutathione depletion (data not shown). If, however, it were possible to
demonstrate that CPT-cGMP potentiates the increase in
intracellular Ca2+ in a manner similar to toxicity, then it is
likely that cGMP-gated Ca2+ channels are opened. Fig. 5 A
shows that CPT-cGMP indeed potentiates Ca2+ elevation.
CPT-cGMP greatly enhances the level of intracellular Ca2+ in cells exposed to marginally toxic (2 mM) concentrations of glutamate.
Fig. 5.
sGC/cGMP regulates the influx of Ca2+. (A
and B) Representative scatter plots depicting the violet/
green ratio of intracellular Indo-1-emitted fluorescence
(proportional to Ca2+) versus
the side angel scatter (SSC-height, which is proportional to the gravity of cells). HT22
cells were treated as indicated for 12 h and Ca2+ content determined as described
in Materials and Methods.
Each plot represents 10,000 viable cells. Similar results
were obtained in two independent experiments with repeated samples. The level of
Ca2+ was arbitrarily defined
as low (the violet/green ratio
less than 50), medium (the
ratio between 50 and 100),
and high (the ratio above
100), and the following percentages of cells containing low, medium, and high Ca2+
are given in that order. (A)
97.8, 2.0, and 0.2 (control):
91.5, 6.7, and 1.8 (2 mM Glu);
66.0, 22.0, and 12.0 (2 mM Glu + 500 µM CPT-cGMP); and 96.9, 2.8, and 0.3 (500 µM CPT-cGMP). (B) 98.4, 1.5, and 0.1 (control); 90.2, 7.6, and 2.2 (5mM Glu); 98.9, 1.0, and 0.1 (5 mM Glu + 1 µM LY83583); and 99.5, 0.5, and 0 (1 µM LY83583). Note that the baseline for
the control in A was higher than that in B.
[View Larger Version of this Image (34K GIF file)]
) and was arbitrarily
defined as low (the ratio is less than 50), medium (the ratio
is between 50 and 100), and high (the ratio is between 100 and 250). In control cells, only about 2% of the cells had medium to high levels of Ca2+, with very few cells containing the high Ca2+ level (Fig. 5 B; percentiles of cells in
each class were derived from the scatter plots shown).
Treatment with 5 mM glutamate for 12 h resulted in a dramatic increase in the percentages of cells containing medium to high levels of Ca2+ (Fig. 5 B). 12 h after the exposure of cells to glutamate, ~20% of the cells were alive. Of
these, only 10-20% had medium to high levels of Ca2+
(Fig. 5 B and data not shown). The fact that the high Ca2+-
containing cells accounted for such a minority of cells at
the time when cells were dying rapidly again suggests that
Ca2+ elevation occurs very close to cell lysis. The rise of
Ca2+ level during the incubation of cells with glutamate
was prevented by sGC inhibitors, while these inhibitors
had no effect on control cells (Fig. 5 B and data not
shown). Therefore, sGC inhibitors block glutamate-induced
Ca2+ elevation in HT22 hippocampal cells. It was not,
however, possible to do these experiments in primary cultures because of unstable baselines due to spontaneous
cell death in the short term cultures.
Discussion
). However, the other sGC inhibitor, LY83583, appears to be relatively specific for sGC
in various studies (Schmidt et al., 1985
; Mulsch et al.,
1988
). The mechanism by which LY83583 inhibits sGC is
not completely known, but the inhibitory effect appears to
require the intracellular reduction of the compound
(Mulsch et al., 1988
). However, LY83583 inhibits glutathione reductase, thereby causing an increase in the level
of GSSG (Luond et al., 1993
). This effect may contribute
to the ability of LY83583 to inhibit sGC since high levels
of GSSG can cause an irreversible loss of sGC activity
(Graff et al., 1978
; Frey et al., 1980; Wu et al., 1992
; Mayer
et al., 1995
).
) and protects against excitatory amino acid-induced
damage in cerebellar slices (Garthwaite and Garthwaite,
1988
). cGMP may also be involved in the excitoprotective activities of the secreted forms of
-amyloid precursor in
hippocampal neurons (Barger et al., 1995
) and mediate
the survival effect of NO on trophic factor-deprived PC12
cells and sympathetic neurons (Farinelli et al., 1996
). In
contrast, the accumulation of cGMP is associated with retinal degeneration (Lolley et al., 1977
; Bowes et al., 1990
)
and possibly mediates excitatory amino acid-induced cytotoxicity in cortical neurons (Frandsen et al., 1992
; Lustig et al., 1992
). The experiments presented above demonstrate that sGC/cGMP is also a component in the cell
death pathway in nerve cells depleted of GSH.
).
). However, the regulator
and the effector molecule for these Ca2+ fluxes are largely
unknown. Recently, it has been shown that Ca2+ influx in
glucocorticoid-induced lymphocyte apoptosis is mediated through the inositol 1,4,5-triphosphate receptor pathway
(Khan et al., 1996
). It is therefore likely that various regulators of Ca2+ exist to control cell death in different environments.
Received for publication 28 July 1997 and in revised form 25 September 1997.
1. Abbreviations used in this paper: BSO, buthionine sulfoximine; CPT-cGMP, 8-(4-chlorophenylthio) guanosine-3We would like to thank Drs. H. Kimura, Y. Liu, Y. Sagara, and S. Tan for their helpful suggestions during the course of this study and critically reading the manuscript, and Dr. D. Chambers for expert assistance in the FACS® analysis.
This work was supported by National Institutes of Health grants R01 NS09658 (to D. Schubert) and 5PO1-NS28121 (to D. Schubert and P. Maher). Y. Li was supported in part by a postdoctoral fellowship from the de Hoffmann Foundation and a National Research Service Award 1 F32 AG 05769-01.
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