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The Journal of Neuroscience, May 15, 2001, 21(10):3303-3311
p38 Activation Is Required Upstream of Potassium Current
Enhancement and Caspase Cleavage in Thiol Oxidant-Induced Neuronal
Apoptosis
BethAnn
McLaughlin1,
Sumon
Pal1,
Minhnga P.
Tran1,
Andrew A.
Parsons2,
Frank C.
Barone2,
Joseph A.
Erhardt2, and
Elias
Aizenman1
1 Department of Neurobiology, University of Pittsburgh
School of Medicine, Pittsburgh, Pennsylvania 15261, and
2 Department of Cardiovascular Pharmacology, SmithKline
Beecham Pharmaceuticals, Philadelphia, Pennsylvania 19406
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ABSTRACT |
Oxidant-induced neuronal apoptosis has been shown to involve
potassium and zinc dysregulation, energetic dysfunction, activation of
stress-related kinases, and caspase cleavage. The temporal ordering and
interdependence of these events was investigated in primary neuronal
cultures exposed to the sulfhydryl oxidizing agent
2,2'-dithiodipyridine (DTDP), a compound that induces the intracellular
release of zinc. We previously observed that tetraethylammonium (TEA),
high extracellular potassium, or cysteine protease inhibitors block
apoptosis induced by DTDP. We now report that both p38 and extracellular signal-regulated kinase phosphorylation are
evident in neuronal cultures within 2 hr of a brief exposure to
100 µM DTDP. However, only p38 inhibition is capable of
blocking oxidant-induced toxicity. Cyclohexamide or
actinomycin D does not attenuate DTDP-induced cell death, suggesting
that posttranslational modification of existing targets, rather than
transcriptional activation, is responsible for the deleterious
effects of p38. Indeed, an early robust increase in
TEA-sensitive potassium channel currents induced by DTDP is attenuated
by p38 inhibition but not by caspase inhibition. Moreover, we found
that activation of p38 is required for caspase 3 and 9 cleavage,
suggesting that potassium currents enhancement is required for caspase
activation. Finally, we observed that DTDP toxicity could be blocked
with niacinamide or benzamide, inhibitors of poly (ADP-ribose)
synthetase. Based on these findings, we conclude that oxidation of
sulfhydryl groups on intracellular targets results in intracellular
zinc release, p38 phosphorylation, enhancement of potassium currents,
caspase cleavage, energetic dysfunction, and translationally
independent apoptotic cell death.
Key words:
oxidation; apoptosis; zinc; p38; potassium; caspase
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INTRODUCTION |
Substantive advances have been made
in elucidating the cellular and molecular signaling pathways
contributing to neuronal apoptosis. However, it remains largely
unknown how apparently disparate events implicated in apoptosis, such
as oxidative stress, ionic dysregulation, and activation of
mitogen-activated protein kinases (MAPK) and caspases, signal among one
another to initiate and propagate cell death. For instance, although it
has been known for some time that potassium ionophores promote
apoptosis, that appreciable loss of intracellular potassium occurs
during apoptosis (Perregaux and Gabel, 1994 ; Walev et al., 1995;
Bortner et al., 1997; Dallaporta et al., 1998; Bortner and
Cidlowski, 1999), and that increased extracellular potassium
can attenuate cell death, the precise role of the dysregulation of this
ion in promoting caspase-dependent cell death has not been well defined
(Ojcius et al., 1991; Beauvais et al., 1995; Hughes et al., 1997).
The loss of intracellular potassium that occurs after a variety of
apoptotic stimuli can be caused by increased activation of
voltage-gated potassium channels, and blocking these channels with
tetraethylammonium (TEA) is neuroprotective (Yu et al., 1997). This
suggests that potassium efflux may be a requisite step for propagating
apoptotic signaling. Given that increased potassium conductance can be
observed within 2 hr of staurosporine or tumor necrosis
factor- treatment (Maeno et al., 2000), it seems likely that
potassium channel opening may be both a conserved and proximal event
induced by multiple apoptotic stimuli. Although the factors that
contribute to enhanced potassium channel opening and the mechanism by
which the loss of intracellular potassium triggers apoptotic signal
cascades have not been determined, it has been proposed that the
osmotic and energetic dysfunction brought on by the loss of the cation
could result in p38 activation and cell death (Yu and Choi, 2000).
p38 is a member of the MAPK family that is sensitive to a number of
osmotic, oxidative, and environmental stressors. On activation, p38
catalyzes the phosphorylation and activation of specific transcription factors, including cyclic AMP-responsive element binding protein, DNA
damage-inducible genes, and ATF-2 (for review, see Cobb, 1999). Although the precise mechanism by which p38 induces cell death is
unclear, blockade of this kinase has been shown to be neuroprotective against a variety of apoptotic stimuli (Kawasaki et al., 1997; Clerk et
al., 1998).
The purpose of this work was to define the cellular mechanisms that
contribute to p38-induced cell death by determining the temporal
ordering of the signaling pathways that enhance potassium channel
opening, caspase activation, and cell death in neurons exposed to
oxidative stress. We have demonstrated previously that exposure to the
cell permeant thiol oxidant DTDP results in intracellular zinc release
and apoptotic cell death in neurons that can be attenuated with the
broad-spectrum cysteine protease inhibitor Boc-aspartate-fmk (BAF), the potassium channel blocker TEA, or high levels of
extracellular potassium (Aizenman et al., 2000). In this study, we have
used a novel and highly selective p38 inhibitor and found that
oxidative stress induces early increases in p38 phosphorylation that
initiate translationally independent apoptosis by increasing potassium channel activation upstream of caspase cleavage.
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MATERIALS AND METHODS |
Chemicals and reagents. All cysteine protease
inhibitors were purchased from Enzyme Systems Inc. (Livermore, CA).
Commercially available MAPK inhibitors were acquired from Calbiochem
(La Jolla, CA). All media and media supplements were from Life
Technologies (Grand Island, NY). Western blotting gels, transfer
apparatus, and standards were from Bio-Rad (Hercules, CA), and MAPK and
p38 antibodies were from Cell Signaling (Beverly, MA). Polyvinylidene difluoride (PVDF) membranes and ECL reagents for immunoblotting were from Amersham Pharmacia Biotech (Piscataway, NJ). All other chemicals were from Sigma (St. Louis, MO).
Cell culture and toxicity assays. Forebrain
neuronal-enriched cultures were prepared from embryonic day 17 rat
fetuses as previously described (McLaughlin et al., 1998). Dissociated
cells were plated on poly-L-ornithine-treated
tissue culture plates in a growth medium composed of 80% DMEM (high
glucose with L-glutamine and without sodium
pyruvate), 10% Ham's F12-Nutrients (Sigma), 10% bovine calf serum
(heat-inactivated) with antimycotic-antibiotic mixture (with
amphotericin B and streptomycin sulfate). Cultures were maintained in
an incubator at 37°C, 5% CO2. Glial cell
proliferation was inhibited after 48 hr in culture with 1-2
µM cytosine arabinoside. After 3 d
in vitro, the serum-containing medium was replaced with a
serum-free solution composed of neurobasal medium (without
L-glutamine), B27 supplement, and
antimycotic-antibiotic mixture. At 2 weeks in vitro, these
cultures were composed of >95% neurons as assessed by Hoescht and
glial fibrillary acidic protein staining.
Toxicity assays were performed on 2-week-old cultures (14-17 d
in vitro) unless otherwise noted. Immediately before drug
treatment, the cells were rinsed in MEM with Earle's salts
(200:1) containing 0.01% bovine serum albumin, 25 mM HEPES, and N2 media supplement. For some of
the experiments, cells were exposed to various compounds 1 hr before,
during, and in the 18-20 hr after DTDP exposure when assessing their
efficacy in altering DTDP toxicity. DTDP treatment was performed for 10 min at 37°C, 5% CO2, and terminated by serial dilution (200:1) in MEM as above. Cells were then returned to the
incubator, and neuronal viability was determined 18-20 hr after
exposure using a lactate dehydrogenase (LDH)-based in vitro toxicology assay kit. Media samples (40 µl) were analyzed
spectrophotometrically (490:630), according to the
manufacturer's protocol, to obtain a measure of cytoplasmic LDH
release from dead and dying neurons. Toxicity was assessed as the ratio
of LDHDTDP/LDHvehicle and, as are all data, expressed as the mean ± SEM.
Electrophysiological measurements. Recordings were conducted
on a total of 78 neurons using the whole-cell configuration of the
patch-clamp technique as described previously (Leszkiewicz et al.,
2000). The extracellular solution contained (in
mM): 115 NaCl, 2.5 KCl, 2.0 MgCl2, 10 HEPES, 0.1 1,2-bis(2-aminophenoxy)ethane-N,N,N,N,-tetraacetic acid (BAPTA), 10 D-glucose, pH was
adjusted to 7.2; 0.1 µM tetrodotoxin was added
to inhibit voltage-gated sodium channels. The intracellular (electrode)
solution contained (in mM): 120 KCl, 1.5 MgCl2, 1 CaCl2, 2.0 Na2ATP, 1 BAPTA, 10 HEPES, pH 7.2. Measurements
were obtained under voltage clamp with an Axopatch 200 amplifier (Axon Instruments, Foster City, CA) and pClamp software (Axon Instruments) using 2 M electrodes. Partial compensation ( 80%) for series resistance was performed in all cases. Currents were filtered at 2 kHz
and digitized at 10 kHz (Digidata; Axon Instruments). Drugs were
dissolved in the extracellular solution and applied to the cells using
a multibarrel fast perfusion system (Warner Instruments, Hamden, CT).
Potassium currents were evoked with a series of incremental 80 msec
voltage steps to 35 mV from a holding potential of  70 mV.
Steady-state current amplitudes were measured relative to baseline 70 msec after the initiation of each voltage step and normalized to cell capacitance.
Caspase activity assays. Caspase 3 and 9 activities were
measured using fluorometric caspase activity detection kits (R & D
Systems, Minneapolis, MN). Cells were treated as above for 10 min with
100 µM DTDP, rinsed, and returned to the
incubator. At various time points, cells were harvested after a wash in
ice-cold PBS, and pellets were collected by centrifugation at 250 × g for 10 min at 4°C. Pellets were resuspended in
standard cell lysis buffer provided by the manufacturer. Lysates were
then incubated on ice for 10 min. A small aliquot of the protein
suspension was removed and stored at 20°C for protein
quantification using the BCA protein assay (Pierce Chemical Co.,
Rockford, IL). After the 10 min incubation, 50 µl of cell lysate was
added to a 96 well flat-bottom plate along with an equal volume of 2×
reaction buffer with freshly prepared DTT to a final concentration of
10 µM. Finally, 5 µl of caspase 9 fluorogenic substrate (LEHD-AFC) or the caspase 3 fluorogenic substrate
(DEVD-AFC) was added, and the plate was incubated in the dark at
37°C for 1 hr. Fluorescence was then measured using a CytoFluor II
plate reader (PerSeptive Biosystems, Framingham, MA) equipped with a
400 nm excitation filter and a 505 nm emission filter. Data are
expressed as fluorescent units per microgram of protein.
Western blot assessment of MAPK activation. At various times
after DTDP treatment, cells were harvested for detection of activated MAPK proteins. Cultures were placed on ice; after media aspiration, cells were washed twice in ice-cold PBS, then lysed in 50 mM Tris buffer, pH 8.0, with 1% Triton X-100, 2 mM EDTA, 150 mM NaCl, 8 mM  -glycerophosphate, and 100 µM sodium orthovanadate. Residual cells were
harvested, and an aliquot of this suspension was removed for later
protein determination. Then, an equal volume of laemmli buffer was
added to cell lysates, and samples were sonicated for 10 sec to remove
viscosity and shear DNA. Samples were heated then to 95°C for 5 min
and stored at 20°C until blots were run.
Equal protein concentrations were separated using 10 or 12% SDS-PAGE
minigels with prestained kaleidoscope molecular weight markers.
Proteins then were transferred to PVDF membranes and blocked for 1 hr
at room temperature (RT) in CHEMIblocker (Chemicon, Temecula, CA)
diluted 1:1 with PBS with 0.1% Tween (PBST). Membranes were washed
four times in PBST for a total of 30 min, then incubated overnight in primary antibody diluted 1:1000 in blocking
solution. The following day, primary antibody was removed; blots were
washed as above and then incubated for 1 hr at RT with an
HRP-conjugated anti-rabbit secondary antibody (Santa Cruz
Biotechnology, Santa Cruz, CA) diluted 1:2000 in blocking buffer.
Membranes were washed again as above and placed in 2 ml of ECL
chemiluminescent substrate for 1 min at RT. Membranes were exposed then
to Kodak X-OMAT x-ray film (VWR Scientific, Bridgeport, NJ).
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RESULTS |
DTDP-induced cell death in primary neuronal cultures can be
attenuated with antioxidants
Brief exposure to the cell permeant oxidant DTDP induces neuronal
apoptotic cell death (Aizenman et al., 2000). We observed that cell
death induced by DTDP can be attenuated by the antioxidant mixture B27
(Table 1). B27 contains a number of free
radical scavengers and reducing equivalents, including glutathione, tocopherol, selenium, and ascorbate. The presence of B27 both during
the 10 min exposure to DTDP and in the subsequent 18-20 hr increased
survival by almost 60% in the cells exposed to 75 µM
DTDP and by ~40% in cells treated with 100 µM DTDP.
This effect is most likely attributable to the antioxidant
properties of B27 because the neuroprotective action of the free
radical spin trap N-tert-butyl--phenylnitrone (PBN) was
very similar to that of B27. These findings are consistent with our
previous observation that DTDP results in accumulation of intracellular
zinc (Aizenman et al., 2000), because alteration in zinc homeostasis
and sequestration have been implicated as a causative factor in
neuronal oxidative stress via production of free radicals (for review,
see Weiss et al., 2000).
In additional experiments, we observed that 3 mM
niacinamide provided significant protection against DTDP-induced
apoptosis (Table 1). Niacinamide can exert neuroprotective action as an oxyradical scavenger, a precursor for NAD+
synthesis, an iNOS inhibitor, as well as a competitive inhibitor of
NAD+-catabolizing enzymes, including poly
(ADP-ribose) synthetase (PARS) (for review, see Szabo and
Dawson, 1998). Furthermore, niacinamide has recently been shown to
abrogate the toxicity induced by exposing cells to high levels of
extracellular zinc (Sheline et al., 2000). An additional PARS
inhibitor, benzamide (3 mM), was also observed to afford a
significant level of neuroprotection against 100 µM DTDP
(Table 1).
Oxidative stress induces rapid MAPK activation
Exposure to oxidative stress activates members of the
stress-activated protein kinase family (Clerk et al., 1998). We thus measured the activation of two MAPK pathways after 100 µM
DTDP treatment. Cells were harvested at various time points after DTDP exposure, and immunoblots were performed using phospho-specific antibodies to both p38 and extracellular signal-regulated kinase (ERK)
p42/44. Within 30 min of exposure to DTDP, a substantial increase in
p38 phosphorylation was evident (Fig. 1).
After 2 hr, p38 phosphorylation decreased appreciably, and levels were indistinguishable from baseline. Similarly, phospho-ERK immunoblotting demonstrated that there was a pronounced early rise in p42/44 activation within 30 min of exposure to oxidant that decreased dramatically by 1 hr. Use of the nonphosphorylated p38 and ERK antibodies was performed as a control to ensure that equal amounts of
each of the inactive MAPK proteins were present at all time points.
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Figure 1.
Activation of p38 and ERK after exposure to DTDP.
Whole-cell extracts of neuronal cultures were harvested at various time
points after 10 min exposure to 100 µM DTDP. Proteins
were separated on 12% SDS-PAGE gels and probed with antibodies
specific to the phosphorylated and nonphosphorylated forms of both p38
and ERK p42/44. Note that there was early increased p38 activation in
the first 2 hr after exposure to DTDP and a similar increase in ERK
activation within the first 45 min of oxidant exposure. Similar results
were obtained in two additional independent experiments.
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To address the contribution of MAPK activation to the observed cell
death after DTDP exposure, the effects of specific inhibitors of
MEK 1/2 and p38 on DTDP toxicity were assessed. Cells were preincubated for 1 hr with either 10 µM U0126 (Satoh et
al., 2000) or a second-generation, highly selective, and
neuroprotective inhibitor of p38, SB 239063 (20 µM)
(Barone et al., 2001), and then exposed to 75 or 100 µM
DTDP for 10 min in the presence of the MAPK inhibitors. Kinase
inhibitors were also present during the 18-20 hr period after oxidant
exposure. Neuroprotective effects of these and other compounds were
assessed against both concentrations of DTDP to ensure that any
protective effects of these agents were not masked by the potentially
overwhelming neurotoxic action of 100 µM DTDP. Although
we observed relatively large increases in p42/44 phosphorylation after
DTDP exposure, this pathway did not appear to contribute to
oxidant-induced cell death because U0126 did not abrogate DTDP toxicity
(Fig. 2). However, treatment with the p38
inhibitor provided significant protection against both doses of DTDP
tested (Fig. 2). The majority of neurons treated with DTDP in the
presence of p38 inhibitor maintained a healthy phase-bright soma, yet
we did observe some disruption of neuronal processes. Cells treated
with SB 239063 alone did not display this phenotype (data not
shown).
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Figure 2.
p38 activation contributes to DTDP-induced
neurotoxicity but is not dependent on new protein synthesis. Neuronal
cultures exposed to 75 or 100 µM DTDP for 10 min in the
presence of inhibitors of ERK and p38 activation and protein synthesis.
Both U0126 (10 µM) and SB 239063 (20 µM)
were present for 60 min before, during, and 24 hr after exposure to
DTDP, whereas CHX (1 µg/ml) was present only during and after the
DTDP incubation. Cell viability was assessed 18-20 hr after DTDP
exposure by measuring the amount of LDH released into the culture
media. Data are expressed as mean percentage neuroprotection compared
with the same dose of DTDP without inhibitor ± SEM. Hence,
neuroprotection reflects the degree of viability relative to the extent
of death induced by each concentration of DTDP. Data represent the
mean ± SEM of three to six experiments performed in duplicate.
*p < 0.05; paired t test.
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Like other MAPKs, p38 is thought to exert its effect on cell survival
through modification of transcription factors. We were surprised
therefore to find that treatment with the protein synthesis inhibitor
cyclohexamide (1 µg/ml) did not significantly alter the neurotoxic
properties of DTDP (Fig. 2). Additionally, the transcription inhibitor
actinomycin D (5 µg/ml) was completely ineffective in protecting
neurons against 100 µM DTDP-induced toxicity
(n = 3; data not shown). This suggests that direct or indirect modification of existing intracellular targets, rather than
new protein synthesis, is the mechanism of the deleterious action of
p38 after oxidant exposure.
Oxidant-induced apoptosis is accompanied by an enhancement of
voltage-gated potassium currents downstream from p38 activity, but
upstream from caspase activation
In a previous study (Aizenman et al., 2000), we observed that
DTDP-mediated neuronal apoptosis could be abrogated by either high
extracellular potassium or TEA, suggesting that activation of
voltage-gated potassium channels and a concomitant putative loss of
intracellular potassium were involved in the cell death process
(Bortner et al., 1997; Yu et al., 1997, 2000). In the present study, we
investigated whether DTDP-induced cell death was indeed accompanied by
an enhancement of voltage-gated potassium currents and whether
inhibitors of cell death, including SB 239063, could attenuate channel
activity. Using whole-cell recordings, we detected a robust increase in
the amplitudes of outward voltage-gated potassium currents ~3-4 hr
after a 10 min exposure to 100 µM DTDP, compared with
vehicle-treated cells (Fig.
3A). Potassium currents in
both DTDP-treated cells and controls had relatively slow kinetics and
were sensitive to block by 10 mM TEA, consistent
with the hypothesis that apoptotic stimuli enhance delayed rectifier
potassium currents in dying neurons (Yu et al., 2000).
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Figure 3.
Enhanced voltage-gated potassium currents during
DTDP-induced apoptosis. A, Whole-cell potassium currents
obtained in two separate cortical neurons ~3 hr after a 10 min
exposure to either vehicle (0.1% DMSO) or 100 µM DTDP.
Cells were maintained in MEM at 37°C for the interval between drug
exposure and the recordings. Potassium currents were evoked by a series
of voltage steps to +35 mV from a holding potential of 70 mV. Note
that currents are substantially larger in the DTDP-treated cell, when
compared with the vehicle-treated neuron. TEA (10 mM)
blocked ~50% of the currents in both cases. B, Acute
application of 100 µM DTDP or 4 mM
dithiothreitol (DTT; a disulfide reducing agent) did not
produce appreciable changes in the amplitude of potassium
currents.
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In a separate set of experiments, we confirmed that the effects of DTDP
on the potassium currents were associated with the apoptotic process
and were not caused by a direct interaction of the redox reagent with
the channels (Aizenman et al., 1989; Gulbis et al., 2000). Acute
exposure of cells to 100 µM DTDP during the recording
process produced negligible changes in the amplitude of the
potassium currents (Fig. 3B). No effect was seen after incubation with the reducing agent dithiothreitol (3 mM), suggesting that the aforementioned increase
in potassium currents observed in dying cells is not caused by a direct
redox effect of DTDP on potassium channels.
A detailed time course analysis demonstrated that the upregulation of
potassium currents could be detected first ~3 hr after DTDP exposure
and continued to increase in a time-dependent fashion (Fig.
4A). Recordings became
particularly difficult to perform 4 hr after DTDP treatment because
cell membranes became somewhat fragile and unstable. Furthermore, in
most instances, the largest voltage steps in DTDP-treated cells
produced currents for which amplitudes saturated our patch-clamp
amplifier (>20 nA). As a result, we could not determine the
absolute extent of the enhancement of these currents. Hence, all
quantitative measurements were limited to a 3-4 hr postexposure time
window and performed on currents evoked by voltage steps no larger than
60 mV (70 to 10 mV), which we could reliably maintain under voltage
clamp.
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Figure 4.
Potassium current enhancement during DTDP-induced
apoptosis is downstream from p38 activity but precedes caspase
activation. A, Temporal profile of DTDP-induced
enhancement of potassium currents during apoptosis. Whole-cell
potassium currents were evoked by a voltage step to 10 mV from a
holding voltage of 70 mV in 31 cells exposed for 10 min to 100 µM DTDP at time 0. Cells were maintained in MEM at 37°C
between DTDP exposure and the establishment of the recording at various
time points. Current amplitudes were normalized to cell capacitance.
The line represents the average potassium current
density for eight cells recorded at several random time points after
exposure to vehicle alone (0.1% DMSO). Note that potassium currents
begin to become enhanced, relative to baseline, ~3 hr after DTDP
exposure. B, In a separate set of recordings, neurons
were exposed for 10 min to vehicle (n = 9), 100 µM DTDP alone (n = 9), or DTDP in the
presence of 10 µM TPEN (n = 6), 20 µM SB 239063 (n = 14), or 10 µM BAF (n = 6). Potassium currents
were evoked in these cells 3-4 hr after DTDP exposure by voltage-steps
from 70 to 10 mV. Steady-state amplitudes were normalized to cell
capacitance. TPEN and SB 239063, but not BAF, inhibited the enhancement
of potassium currents observed in DTDP-treated cells. Values represent
the mean ± SEM; asterisks denote a significant
difference versus vehicle controls (p < 0.01; ANOVA followed by a Dunnett multiple comparisons test).
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With this paradigm, we examined whether substances known to inhibit
DTDP toxicity, namely the zinc chelator
N,N,N',N'-tetrakis(2-pyridylmethyl)ethylenediamine (TPEN), the p38 inhibitor SB 239063, and the caspase inhibitor BAF,
could block the enhancement of the potassium currents. Coexposure of
100 µM DTDP with either 10 µM TPEN or 20 µM SB
239063 completely prevented the enhancement of the potassium currents,
whereas 10 µM BAF did not (Fig.
4B). These data indicate that the intracellular release of zinc (Aizenman et al., 2000) and activation of the p38
cascade precede the enhancement of channel activity. In contrast, because BAF inhibits toxicity but not potassium channel activity, this
suggests that caspase activation follows potassium efflux. Importantly,
these results also indicate that the enhancement of potassium currents
during apoptosis is, in and of itself, not sufficient to cause neuronal
cell death.
Caspases 3, 8, and 9 contribute to oxidant-induced apoptosis
The results of the potassium channel recordings strongly suggested
that DTDP-induced caspase activation likely occurs subsequent to
enhanced potassium efflux. Zinc neurotoxicity has been associated with
mitochondrial dysfunction and, in some instances, apoptotic cell death
(Skulachev et al., 1967; Treves et al., 1994; Y. H. Kim et al.,
1999; Virag and Szabo, 1999; Sheline et al., 2000; Weiss et al., 2000).
Thus, we sought to determine whether caspases associated with
mitochondrial dysfunction were activated by DTDP, as well as the time
course of their activation. We assessed the efficacy of inhibiting
caspase 3, given that this protease lies downstream of caspase 9, which
is activated by mitochondrial cytochrome c release, as well as a
caspase 8, which has recently been localized to the mitochondria and
shown to be released during cellular dysfunction (Qin et al., 2000).
Cells were treated with cysteine protease inhibitors for 1 hr before,
during, and 24 hr after a 10 min exposure to DTDP. As expected from
previous work with relatively younger cultures (Aizenman et al., 2000),
the broad-spectrum cysteine protease inhibitor BAF (10 µM) provided appreciable neuroprotection against the two
doses of DTDP tested (Fig. 5). Moreover,
the novel nonpeptide inhibitor of caspases 3, 7, and 9, isatin-sulfonamide-4 (IS-4; 3 µM) (Erhardt et al., 2000;
Lee et al., 2000), provided 40 ± 6 and 39 ± 10% protection
against 75 and 100 µM DTDP, respectively. We next treated
cells with the caspase 8 inhibitor
z-Leu-Glu-(OMe)Thr-Asp(OMe)-CH2F (LETD; 10 µM), which also provided appreciable neuroprotection against both doses of DTDP (p < 0.05;
n = 4-5). Addition of
Biotin-Phe-Ala-CH2F (zFA; 20 µM), which contains an fmk group without
a caspase inhibitor peptide sequence, provided no protection on its own
(data not shown).
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Figure 5.
Inhibition of caspases 3, 8, and 9 blocks
oxidant-induced cell death. Neuronal cultures exposed to 75 or 100 µM DTDP for 10 min in the presence of peptide and
nonpeptide caspase inhibitors. The broad spectrum cysteine protease
inhibitor BAF (10 µM), caspase 3, 7, and 9 selective
nonpeptide inhibitor IS-4 (3 µM), and the caspase 8 selective inhibitor zLETD (10 µM) were present for 60 min
before, during, and 24 hr after exposure to DTDP. Cell viability was
assessed as above, and data are expressed as mean
percentage neuroprotection compared with the same dose of DTDP without
inhibitor ± SEM. Data represent the average of five experiments
performed in duplicate. All compounds provided significant protection
at both doses of DTDP tested. *p < 0.05; paired
t test.
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The time course of caspase 9 and 3 activation after 10 min treatment
with DTDP was then assessed using enzymatic caspase activity assays
with 7-amino-4-trifluoromethyl coumarin (AFC)-labeled substrate peptides. As shown in Figure 6, there is
a time-dependent increase in caspase 9 and caspase 3 activities after
DTDP exposure. Notably, only very minor changes in activity were
evident before the 7 hr time point, indicating that DTDP-induced
caspase activation likely occurs after p38 phosphorylation and
potassium efflux. This possibility was tested by examining whether
antioxidants, niacinamide, and p38 inhibitors block caspase cleavage.
The efficacy of each of these agents at blocking caspase activity was
assessed 7 hr after treatment. The elevation in caspase 9 activity that was observed at this time point was significantly diminished by 10 µM BAF, 20 µM SB 239063, or 3 mM niacinamide (Fig.
7A). Similarly, the increase
in caspase 3 activity was also blocked when cells were treated with
these three compounds (Fig. 7B). Additionally, the
antioxidant mixture B27 and free radical spin trap PBN significantly attenuated both caspase 9 and 3 activity by 41 ± 13 and 61 ± 4%, respectively (n = 4-5;
p < 0.05; one sample t test). These
findings indicate that reactive oxygen species (ROS), p38, and
possibly energetic stress signals are upstream effectors that elicit
caspases 3 and 9 cleavage after DTDP exposure.
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Figure 6.
Caspase 9 and 3 are activated relatively late
after exposure to DTDP. Neuronal cultures exposed to 100 µM DTDP for 10 min and then washed and harvested at
various time points to assess the extent of caspase cleavage using
fluorometric peptide substrates. Lysed cells were incubated in the
presence of either the fluorogenic caspase 9 substrate (LEHD-AFC) or
the caspase-3 substrate (DEVD-AFC). Activity was then calculated as
fluorescent units per microgram of protein and expressed as
activity in relative units compared with control. Data represent the
average ± SEM of at least six experiments performed in duplicate.
p < 0.05; one-way ANOVA.
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Figure 7.
NAD+ depletion and p38
activation occur before caspase cleavage. Neuronal cultures were
exposed to 100 µM DTDP for 10 min in the presence of
niacinamide (Nia; 3 mM), BAF (10 µM), or SB 239063 (20 µM) as above and
harvested 7 hr later. Data were analyzed and expressed as in Figure 6
and represent the mean ± SEM of four to six independent
experiments performed in duplicate. There was a statistically
significant decrease in caspase cleavage by blocking either
NAD+-catabolization or p38 on caspase cleavage,
suggesting that activation of these pathways occurs before caspase
involvement in DTDP toxicity. Values represent the mean ± SEM;
asterisks denote a significant difference versus matched
DTDP treatment (*p < 0.05; paired t
test). All DTDP treatment groups were pooled for presentation
purposes.
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Oxidant and zinc dysregulation are the earliest apoptotic signals
induced by DTDP, and their dysfunction leads to p38
activation
The sum of the biochemical and electrophysiological data suggested
that p38 activation is both a proximal and necessary event for both
potassium channel opening and caspase cleavage. However, we had not yet
determined the temporal ordering and interdependence of oxidative
stress and zinc dysregulation in relation to MAPK phosphorylation.
Therefore, we performed immunoblots for p38 activation on cells that
had been treated with neuroprotective agents. Because 2 hr after DTDP
treatment was the latest time point at which we observed maximal p38
phosphorylation, we chose this time to assess the efficacy of the
neuroprotective agents at blocking p38 activation. Cells exposed for 10 min to 25 µM TPEN in the presence of DTDP were provided
the greatest inhibition (>90%) of p38 induction (Fig.
8). The free radical spin trap PBN (500 µM) also substantially decreased p38 phosphorylation
(60%). This finding indicates that oxidative stress and zinc release
from intracellular stores are required for DTDP induction of p38.
However, niacinamide (3 mM) decreased p38 phosphorylation
to a much lesser extent (25%), as did the potassium channel blocker
TEA (25 mM; <20%), suggesting that although these drugs
can block cell death, their effects mostly lay downstream of p38
phosphorylation.
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Figure 8.
p38 activation occurs upstream of
K+ channel opening and is induced by zinc release
and oxidative stress. Neuronal cultures were exposed to 100 µM DTDP for 10 min in the presence of niacinamide (3 mM), TEA (25 mM), TPEN (25 µM),
or PBN (500 µM) as above, and whole- cell extracts were
harvested 2 hr later. Proteins were separated by SDS-PAGE and
transferred to PVDF membranes that were then probed with an antibody
selective for the phosphorylated form of p38. Extracts from C-6 glioma
cells after anisomycin (Anis) treatment were
used as a positive control. Note that p38 activation induced by DTDP
was substantially attenuated by both ion chelation
(TPEN) and free radical scavenging
(PBN), but only marginally by TEA blockade of
K+ efflux. Similarly, inhibition of
NAD+ catabolism by niacinamide provided only
marginal attenuation of p38 activation. Quantification of levels with
NIH Image software revealed the following mean band densities (± SD):
control, 109.9 ± 36.9; DTDP, 226.1 ± 13.9; DTDP + niacinamide, 196.6 ± 21.9; DTDP + TEA, 202.3 ± 18.2; DTDP + TPEN, 114.4 ± 18.3; DTDP + PBN, 175.5 ± 26.3. Comparable
results were obtained in two other independent experiments.
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|
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DISCUSSION |
We show here that oxidative stress induced by DTDP induces
apoptotic cell death via activation of the stress-related protein kinase p38. We present evidence that p38 phosphorylation is a proximal
event leading to the enhancement of voltage-gated potassium currents, a
phenomenon that has previously been associated with apoptosis (Yu et
al., 1997). This process occurs in the absence of new protein
synthesis, suggesting that p38 causes direct or indirect
phosphorylation of the potassium channels themselves, or an associated
structure. We also confirm a previous report that the enhancement of
the potassium currents precedes the activation of caspases (Maeno et
al., 2000). Furthermore, the observation that caspase inhibitors block
cell death but not potassium current enhancement suggests that the
putative loss of intracellular potassium after DTDP treatment is not,
in and of itself, a lethal event.
We reported previously that DTDP induces intracellular zinc release
(Aizenman et al., 2000). This process contributes to the observed p38
activation because it is inhibited by TPEN, a high-affinity zinc
chelator. Zinc release may also be critically important in other forms
of oxidation-induced apoptosis in which p38 has been implicated, such
as that occurring after NO exposure (Ghatan et al., 2000). Indeed, NO
has been shown previously to trigger zinc release from metallothionein
(Pearce et al., 2000), a metalloprotein that also releases zinc after
DTDP oxidation (Jiang et al., 1998; Maret and Vallee, 1998). We also
find evidence that the intracellular signaling cascades activated by
DTDP appear to be similar to those observed by other researchers after
exposure to extracellular zinc. Samet et al. (1998) reported p38 and
ERK activation in human bronchial cells after zinc exposure. Although
we observed increases in both ERK and p38 after DTDP treatment, only
p38 activation contributes significantly to the observed neurotoxicity.
Exposure to zinc may induce energetic stress via inhibition of
glycolytic enzymes (Ikeda et al., 1980; Krotkiewska and Banas, 1992;
Kukimoto et al., 1996), because the resultant cell death can be
attenuated by addition of 3 mM niacinamide (Sheline et al.,
2000). Similarly, we found that niacinamide provides significant protection against DTDP toxicity. The most widely described
neuroprotective properties of niacinamide have been attributed to the
fact that it is a precursor for NAD+
synthesis and an inhibitor of the
NAD+-catabolizing enzyme PARS. PARS is
activated by DNA damage after oxidative stress (for review, see Pieper
et al., 1999), and depletion of NAD+ by
PARS places an energetic stress on cells (Satoh and Lindahl, 1992).
Niacinamide inhibits PARS with an IC50 of 100 µM in vitro (Rankin et al., 1989). However, at
the doses used in our work and the work by Sheline et al. (2000),
niacinamide can also act as an oxyradical scavenger, an iNOS inhibitor,
and an inhibitor of other
NAD+-catabolizing enzymes (Ziegler et al.,
1996; for review, see Szabo and Dawson, 1998). We believe that this and
our previous work suggest that the neuroprotective action of
niacinamide can be attributed most likely to its ability to increase
cellular energetic status. That is, niacinamide is far less effective
than the free radical spin trap PBN in blocking p38 activation,
suggesting that the oxyradical scavenging activity of niacinamide is
not overly dramatic. Furthermore, we have previously observed
appreciable DNA damage within 3 hr of exposure to DTDP (Aizenman et
al., 2000), suggesting that PARS activation and subsequent energetic
dysfunction may also be triggered by the production of free radicals
after DTDP exposure. Finally, another PARS inhibitor, benzamide, was also effective in inhibiting DTDP-induced cell death.
Our findings that the earliest manifestations of DTDP-induced cell
stress are associated with oxidative injury are supported by the
neuroprotective action of both PBN and B27, as well as by the finding
that PBN partially blocks p38 activation. This result supports previous
studies in which PBN attenuates oxidative stress-induced p38 activation
in other in vitro systems (Floyd, 1999). There is evidence
suggesting that rapid rise in intracellular zinc, such as that which
occurs after DTDP exposure (Aizenman et al., 2000), may itself lead to
ROS generation and lipid peroxidation (Sensi et al., 1999). The ability
of zinc to directly elicit oxidative stress remains somewhat
controversial however, given that the ability of antioxidants to
attenuate zinc toxicity has been inconsistent (E. Y. Kim et al.,
1999; Y. H. Kim et al., 1999; but also see Sensi et al., 1999;
Sheline et al., 2000). It is also possible that critical cellular redox
agents, including thioredoxin, glutathione, cysteine, and
N-acetyl cysteine, are directly altered by DTDP. These cellular antioxidants are important not only as free radical scavengers but also as metal chelators and substrates for redox reactions (for review, see Deneke, 2000).
The observation that DTDP induces apoptotic cell death, which can be
attenuated with caspase inhibitors, high levels of extracellular potassium, or TEA, is consistent with the observation that loss of
intracellular potassium and cell shrinkage are early requisite features
of apoptosis (Barbiero et al., 1995; Benson et al., 1996; Bortner et
al., 1997; Hughes et al., 1997). However, the signaling pathways that
contribute to enhanced potassium efflux are ill defined. Kinase
signaling pathways have been implicated in apoptotic ionic
dysregulation as Gomez-Angelats et al. (2000) recently demonstrated that suppression of PKC occurs upstream of potassium channel opening in
Fas-induced apoptosis. Furthermore, Fas-induced cell shrinkage could be
blocked by PKC stimulation or enhanced by PKC inhibition. Preliminary
findings in our laboratory suggest moderate protection against 100 µM DTDP can be induced by the PKC activator PMA (40 nM) (B. A. McLaughlin and E. Aizenman, unpublished
observation). The role of PKC and other kinases on p38 signaling
remains to be elucidated because the temporal ordering and reliance of
these phosphorylation events is unclear in other systems (Jun et al., 1999a,b; Shimizu et al., 1999; Yu et al., 2000). For instance, PKC
activation has been linked to increased reactive oxygen species generation and has been suggested to be deleterious to neuronal cultures exposed to zinc (Noh et al., 1999). This discrepancy may be an
important point of divergence between intracellular and extracellular
zinc-induced toxicity (Cuajungco and Lees, 1997).
Our observation that p38 activation is an early and requisite event for
potassium channel opening strongly suggests that SAPK phosphorylation
leads to a volume change in neurons. Maeno and coworkers (2000)
recently reported that apoptotic volume decrease occurs before DNA
fragmentation, cytochrome c release, and caspase activation, apoptotic
events that are reminiscent of those reported here and previously
(Aizenman et al., 2000). Nonetheless, because p38 is activated by
osmotic stress in other systems (for review, see Ono and Han, 2000),
one could easily envision a scenario in which the putative loss of
intracellular potassium activates p38 and other proapoptotic signaling
molecules, thus placing enhanced potassium channel activation upstream
of p38 phosphorylation. Indeed, Benson et al. (1996) have shown that
cell shrinkage correlates with activation of ERK and p38 in kidney
cells and that use of p38 blockers largely inhibits the resulting
protective cellular volume recovery, whereas ERK inhibitors do not.
However, our observation that p38 inhibitors completely block the
enhancement of TEA-sensitive potassium channels after DTDP exposure and
that TEA only marginally attenuates the observed increase in p38
activation supports a very different temporal ordering of MAPK
phosphorylation and potassium channel activation, at least in neurons.
It is likely that the combination of oxidative stress and zinc
dysregulation elicited by DTDP, and possibly other oxidants, is
sufficient to activate p38, which then leads to enhanced activation of
TEA-sensitive potassium channels and a subsequent, requisite volumetric
dysregulation observed in other forms of apoptosis.
We were intrigued by the observation that although p38 activation plays
a prominent role in DTDP toxicity, it does not require new protein
synthesis. The finding that p38 enhances voltage-gated potassium
currents associated with apoptosis suggests that these channel, or
closely associated proteins, may be direct or indirect targets of
phosphorylation by this kinase. It has been reported that p38 can alter
activity of intracellular proteins in a transcriptionally independent
manner. For example, Kusuhara and coworkers (1998) have shown that the
activity of the membrane-bound
Na+/H+
exchanger can be modified in a transcriptionally independent manner by
both ERK and p38. Given that the delayed rectifying potassium channels
have multiple phosphorylation sites that alter activity (Levitan,
1999), it is thus possible that they may also be modified by MAPKs.
Indeed, Adams et al. (2000) have shown that the Kv4.2 potassium channel
can be directly phosphorylated by ERK, suggesting that the activity of
this and the potassium channels associated with apoptotic cell
death may also be modified by p38.
In conclusion, our work suggests that oxidant-induced zinc
dysregulation and ROS generation are the most proximal events in an
apoptotic cascade in which p38 activation leads to potassium efflux,
and subsequently, energetic dysfunction and caspase activation. This
novel temporal ordering of apoptotic signal transduction is important
not only in furthering our understanding of the complex interplay
between posttranslational protein modification and ionic dysfunction in
apoptosis, but also as a model (Fig. 9)
in which to address the importance of these events in neurodegenerative conditions involving zinc release and oxidative stress, such as cerebral ischemia.
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Figure 9.
Model of early and late events in
oxidant-induced neuronal injury. Based on the findings in this and our
previous work, the cellular and molecular signaling pathways that
contribute to oxidant-induced neurotoxicity have been summarized. The
"early" events (left panel) in DTDP toxicity
include the rapid loss of zinc homeostasis within 10 min of exposure to
DTDP (Aizenman et al., 2000), a primary oxidative cell injury resulting
in a rapid increase in p38 phosphorylation within 30 min of oxidant
exposure, and then subsequent DNA damage within 3 hr of DTDP treatment
(Aizenman et al., 2000). p38 activation can be blocked by the zinc
chelator TPEN and, to some degree, by the free radical spin trap, PBN.
Also by 3 hr, enhanced activation of TEA-sensitive potassium channels
associated with apoptosis can be observed. p38 is directly responsible
for the loss of intracellular potassium because the p38 inhibitor SB
239063 can attenuate channel activity, whereas potassium channel
blockers have a very small effect on p38 activity. The "late"
events (right panel) in this cascade likely
include energetic dysfunction brought on by zinc dysregulation
(Skulachev et al., 1967; for review, see Weiss et al., 2000), NADH
depletion, and, ultimately, caspase activation within 7 hr. Blockade of
any of these events with compounds such as niacinamide, benzamide, and
peptide and nonpeptide caspase inhibitors provides substantial
neuroprotection.
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FOOTNOTES |
Received Dec. 7, 2000; revised Jan. 23, 2001; accepted Feb. 13, 2001.
This work was supported by a grant from the Alzheimer's Disease
Research Centre, University of Pittsburgh (B.A.M), National Institutes
of Health Grant NS29365 (E.A.), and an American Heart Association
Grant-in-Aid (E.A.). We thank Drs. Randall Pittman, Ian Reynolds,
Donald DeFranco, and Gregg Stanwood for helpful comments and
suggestions, and Daniel Leszkiewicz, Shen Du, and Annalhees Rodriguez
for assistance.
Correspondence should be addressed to Dr. Elias Aizenman, Department of
Neurobiology, University of Pittsburgh School of Medicine, E1456 BST,
Pittsburgh, PA 15261. E-mail: redox{at}pitt.edu.
| |
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ABSTRACT
INTRODUCTION
