Paraquat-induced phosphatidylserine oxidation and apoptosis
are independent of activation of
PLA2
James P.
Fabisiak1,
Valerian E.
Kagan2,
Yulia Y.
Tyurina2,
Vladimir A.
Tyurin2, and
John S.
Lazo1
Departments of 1 Pharmacology
and 2 Environmental and
Occupational Health, Schools of Medicine and Public Health,
University of Pittsburgh, Pittsburgh, Pennsylvania 15261
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ABSTRACT |
Paraquat is a pneumotoxin that causes lung
injury by enhancing oxidative stress; however, the cellular responses
to these redox events are undefined. We previously showed that paraquat produced selective peroxidation of phosphatidylserine that preceded apoptosis in 32D cells. We now report that the phospholipase
A2 (PLA2) inhibitor quinacrine can
attenuate phosphatidylserine oxidation and also block paraquat-induced
apoptosis. Therefore, we investigated the potential for
PLA2 to mediate apoptosis after
paraquat. We found that, in contrast to quinacrine, the
PLA2 inhibitors manoalide, aristolochic acid, and arachidonyl trifluoromethylketone failed to
prevent paraquat-induced apoptosis. Moreover, no evidence of PLA2 activation was observed
within 7 h after paraquat exposure. Finally, quinacrine failed to
inhibit basal and 4-bromo-A-23187-induced release of
[3H]arachidonic acid
at concentrations that protected paraquat-induced apoptosis. We
conclude that paraquat-induced phosphatidylserine oxidation and
apoptosis occurred in the absence of
PLA2 activation and that
quinacrine protected phosphatidylserine and cell viability after
paraquat in a PLA2-independent
manner.
phospholipase A2; oxidative
stress; lipid peroxidation; quinacrine; arachidonic acid
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INTRODUCTION |
PARAQUAT (PQ), a bipyridyl herbicide, serves as a
prototypical environmental pneumotoxin, the toxic effects of which have been well described in both humans and animals (38). The mechanisms of
cellular damage after PQ involve the
P-450 reductase-dependent formation of
reactive oxygen species and subsequent interactions with lipids,
proteins, and nucleic acids (10). Because reactive oxygen species have
been implicated in the final common pathway that triggers apoptosis
(4), peroxidation of membrane lipids during oxidative stress represents
one potential target for signaling apoptosis (14). It has previously
been demonstrated that PQ induces apoptosis in 32D cells (8). In
addition, it was found that the selective oxidation of
phosphatidylserine (PS) precedes PQ-induced apoptosis. The
externalization of PS to the outer leaflet of the plasma membrane, as
measured by annexin V binding (9), now appears to be one hallmark of
apoptosis, although the mechanism responsible for its translocation is
not known. Inhibition of PS oxidation by overexpression of the
antiapoptotic gene bcl-2 (8) suggests that Bcl-2 can protect cells from one of the
consequences of oxidative stress, namely, lipid peroxidation, and
further supports the hypothesis that PS oxidation is an important
signal during apoptosis.
Membrane lipid peroxidation can affect membrane fluidity, alter the
function of membrane-bound proteins, perturb ion fluxes, and generate
toxic oxidized lipid products (18). Of particular interest is the
potential for oxidative stress to activate phospholipase A2
(PLA2). Lipid peroxidation
accelerates phospholipid hydrolysis in vascular endothelial cells (30),
and oxidized phospholipids provide better substrates for
PLA2 than native phospholipids
(35). In addition,
H2O2
appears to directly activate a signal-responsive PLA2 independent of lipid
peroxidation and changes in intracellular Ca2+, perhaps through stimulation
of enzyme phosphorylation (3). Activation of phospholipid hydrolysis
can further amplify membrane damage (25) and generate potent eicosanoid
and other signaling molecules (7).
Tumor necrosis factor (TNF) is a cytokine with potent cytolytic
properties capable of inducing apoptosis in sensitive cells (21, 22).
The ability of TNF to kill cells has been linked to its ability to
activate intracellular oxidative stress (37) and
PLA2 (13, 40, 42). The role of
PLA2 in apoptosis after oxidants
and other stimuli, however, remains unexplored. It is thus possible
that PLA2 activation coincident
with and resulting from lipid peroxidation can participate in the early
signaling of apoptosis after PQ-induced oxidative stress.
We used our previously characterized model of PQ-induced apoptosis in
32D cells to examine the potential role of
PLA2 (8). We tested the ability of
multiple pharmacological PLA2
inhibitors to protect cells from PQ-induced apoptosis. We also assessed
PLA2 activation after PQ exposure
to determine any association between phospholipid hydrolysis and PS
oxidation. Our results support the hypothesis that
PLA2 is not required to mediate
specific oxidation of PS and subsequent apoptosis after exposure to PQ.
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METHODS |
Materials.
PQ dichloride, quinacrine (QN) hydrochloride, and manoalide were
obtained from Sigma (St. Louis, MO). Arachidonyl trifluoromethylketone (AACOCF3) was from ICN (Costa
Mesa, CA), and aristolochic acid was from Biomol (Plymouth Meeting,
PA). Proteinase K, RNase A, and RNase T1 were from Boehringer Mannheim
(Indianapolis, IN). [3H]arachidonic acid
(AA)
([5,6,8,9,11,12,14,15-3H]AA;
180-240 Ci/mmol) and BetaMax scintillation fluid were obtained from DuPont NEN (Boston, MA).
cis-Parinaric acid
(cis-PNA),
4-bromo-A-23187 (4-Br-A-23187), and Hoechst 33342 were obtained from
Molecular Probes (Eugene, OR). All tissue culture media and reagents
were from GIBCO BRL (Gaithersburg, MD)
except for fetal bovine serum (FBS), which was purchased from Hyclone
(Logan, UT). HPLC-grade solvents and ScintiVerse scintillation cocktail
were from Fisher Scientific (Pittsburgh, PA). All other chemicals and
reagents were of molecular biology or ultrapure grade.
Cell culture.
The murine myeloid cell lines 32D (clone 23) and WEHI-3B were kindly
provided by Dr. Robert Redner (Univ. of Pittsburgh) and were cultured
essentially as previously described (8). We used 32D cells transfected
with an empty mammalian expression vector containing the neomycin
resistance gene (32D/neo cells) that had served as negative
control cells in previous studies examining Bcl-2 overexpression (8).
These cells are phenotypically identical to wild-type 32D cells with
the exception that they are capable of growth in G-418-containing
media. Before drug exposure, 32D cells were seeded at 1.5 × 105 cells/ml in complete media
(RPMI with 10% FBS) and cultured for 48 h. The complete RPMI growth
media contained 10% media conditioned by WEHI-3B as a source of
interleukin (IL)-3, a required survival factor for 32D cells. PQ and QN
were freshly prepared as concentrated stocks in serum-free media and
were then added to cells at indicated concentrations.
PLA2 inhibitors
AACOCF3, manoalide, and
aristolochic acid were administered from stock solutions prepared in
DMSO and stored at
70°C until use. DMSO concentrations never
exceeded 0.4% (vol/vol), and control treatments contained DMSO vehicle alone. For IL-3 withdrawal experiments, 32D cells were cultured in
complete growth media for 48 h. Cells were then collected, washed three
times with serum-free media in the absence of WEHI-conditioned medium,
resuspended, and cultured in the original volume of RPMI, with 10% FBS
but lacking WEHI-conditioned medium.
Nuclear morphology.
Samples of 32D cells (0.5-1 ml) were obtained at the indicated
times after treatment and were collected by centrifugation (1,000 g, 2 min). Cells were then washed in
PBS, fixed with 2% paraformaldehyde in PBS, and stained with Hoechst
33342 (1 µg/ml) as previously described (8). At least 300 cells were
viewed under fluorescent microscopy, with excitation at 340-380
nm, and nuclear morphology was scored as either normal or apoptotic.
Apoptotic nuclei were characterized by small, bright-staining nuclei,
often very rounded and sometimes fragmented into distinct sections.
DNA fragmentation.
Internucleosomal DNA fragmentation was determined by conventional gel
electrophoresis in 2% agarose as previously described (8). Briefly,
DNA was extracted from 1 × 106 cells by lysis in 10 mM EDTA,
0.5% sarkosyl, and 50 mM Tris (pH 8.0) and sequential digestion in
proteinase K (1 mg/ml, 55°C) followed by RNase A (0.5 mg/ml) and
RNase T1 (1,000 units/ml) at 37°C. Electrophoresis was performed at
60 V for ~4 h. Gels were then stained with ethidium bromide (1 µg/ml) and then photographed under ultraviolet illumination.
PLA2 activation and AA release.
Labeling of cellular phospholipids with
[3H]AA and release of
radioactivity into tissue culture media were performed essentially as
described by Jäättellä (15). 32D/neo cells were
obtained from exponential growth-phase cultures, centrifuged (400 g, 10 min), and resuspended (0.8 × 106/ml) in fresh RPMI
containing 10% FBS and 10% WEHI-conditioned medium.
[3H]AA was added (0.2 µCi/ml), and cells were incubated overnight at 37°C in a 5%
CO2 incubator. The following
morning, the radiolabeled cells were harvested and washed three times
with 10 ml of serum-free medium. Radioactivity in aliquots of the total
cell suspension obtained before and after the first centrifugation
indicated that ~70% of the radiolabeled AA was incorporated into
32D/neo cells. Cell number and viability (routinely >95%) were
determined in the last wash suspension by trypan blue exclusion and
cell counting, and the final cell pellet was resuspended in complete
medium at 4 × 105 cells/ml.
Cells were then seeded into 24-well plates (0.5 ml/well) that contained
100 µl of medium containing test substances or vehicle controls.
Individual plates were removed from the incubator at the indicated
times and centrifuged (400 g, 10 min),
and aliquots (300 µl) of the cell-free medium were placed in
scintillation vials. Radioactivity was then determined by scintillation
counting after addition of 3 ml of ScintiVerse counting fluid.
The above-mentioned technique was modified to include an analysis of
[3H]AA hydrolysis from
individual phospholipids after PQ and
Ca2+-ionophore stimulation. For
these studies, cellular phospholipids were labeled as described above.
After overnight incubation, cells were washed as described above with
the exception that the first wash in serum-free media contained 0.5 mg/ml of fatty acid-free human serum albumin to assist in the removal
of free AA. Washed cells were resuspended to 4 × 105 cells/ml, and 15 ml were
seeded into a T-25 flask for each condition. Test substances were then
applied, and cells were incubated at 37°C for 3 h, at which time
two 7-ml aliquots were removed from each flask and centrifuged (400 g, 10 min). Supernatants were decanted, and cell pellets were resuspended in 1.5 ml of PBS and quick-
frozen in a methanol-dry ice bath followed by storage at
70°C until assay. Lipids were extracted from thawed samples
by addition of 10 ml of chloroform-methanol followed by vigorous mixing
(4 min), centrifugation, and collection of the lower organic phase. The
remaining aqueous phase was reextracted with 4 ml of chloroform and
combined with the first organic phase. Organic phase was then
evaporated to dryness under nitrogen, and the residue was reconstituted
in 250 µl of chloroform-methanol (2:1).
Pi was determined on 30 µl of
this sample as described (1), and the remainder was applied to a 5 × 5-cm silica G TLC plate (Whatman, Clifton, NJ). Lipid classes
were separated by two-dimensional high-performance TLC using a solvent
system of chloroform-methanol-28% ammonium hydroxide (65:35:5) in the
first direction and chloroform-acetone-methanol-glacial acetic
acid-water (50:20:10:10:5) in the second. Lipids were visualized by
exposure to iodine vapor and identified by comigration relative to
purified standards. Identified spots were then scraped from the plate
into scintillation vials, lipids were extracted from gel by addition of
250 µl of chloroform-methanol (1:1), and radioactivity was determined
after the addition of 3 ml of BetaMax scintillation fluid.
Lipid peroxidation.
The use of cis-PNA to measure
oxidation of specific phospholipid classes has previously been
described (8). Briefly, 32D/neo cells in log-phase growth were allowed
to incorporate cis-PNA (4 µg/ml,
final concentration) in serum-free RPMI containing 5% WEHI-conditioned
medium for 2 h (1 × 106
cells/ml). cis-PNA was given in
complex with human serum albumin previously prepared by addition of 500 µg of cis-PNA to 50 mg of human
serum albumin in 1 ml of PBS. After repeated washing of cells to remove
unincorporated cis-PNA, cells were
placed in complete media and incubated with the various test substances for two additional hours. Aliquots of cells were then removed, centrifuged at 1,000 g for 2 min, and
washed once with PBS. Cell suspensions were immediately transferred to
cold methanol containing butylated hydroxytoluene (0.1 mg/ml). Total
lipids were extracted with a modified Folch procedure and subjected to
HPLC for separation of phospholipids as previously described (8). The
amount of cis-PNA fluorescence in
individual phospholipid classes was normalized to the amount of
Pi contained within the total
lipid extract. Pi was determined
spectrophotometrically using the method of Chalvardjian and
Rubnicki (5).
Statistical analyses.
The effects of various PLA2 inhibitors on cell viability,
apoptotic morphology, and lipid peroxidation treatments were first assessed by a one-way ANOVA followed by Dunnett's multiple comparisons with control group, which in most cases represented the PQ
alone-treated group. Levels for significant difference were set at
P < 0.05.
 |
RESULTS |
Effect of PLA2 inhibitors on PQ-induced
apoptosis and PS oxidation.
Apoptosis in response to the prototypical oxidant PQ, which is
characterized by chromatin condensation and fragmentation, internucleosomal DNA cleavage, and loss of cell viability within 24 h
of toxin exposure, was previously described (8). We first determined
whether PQ-induced apoptosis was affected by
PLA2 inhibitors.
Figure 1 shows the concentration-dependent
ability of the PLA2 inhibitor QN
to inhibit the changes in nuclear morphology and cell viability after
exposure to PQ. Figure 1A shows that
400 µM PQ alone killed ~30% of the 32D/neo cells within 24 h. In
contrast, when QN was simultaneously included during the PQ exposure,
there was a concentration-dependent preservation of cell viability that achieved >50% protection at 5 µM QN. Similarly, Fig.
1B demonstrates that QN also inhibited
the PQ-induced nuclear changes. Viability of cells receiving 1 and 5 µM QN alone were unchanged compared with untreated control cells
(89.3% for 5 µM QN, 89.6% for 1 µM), and changes in nuclear
morphology were not observed. Preliminary studies revealed that higher
concentrations of QN (10-50 µM) were toxic by themselves (data
not shown) and not included for further analyses. Inhibition of
PQ-induced apoptosis by QN was confirmed by analysis of
internucleosomal DNA fragmentation. Figure
2 shows that 5 µM QN reduced the
formation of low-molecular-weight DNA fragments after PQ exposure.

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Fig. 1.
Quinacrine (QN) protected 32D cells transfected with an empty mammalian
expression vector containing the neomycin resistance gene (32D/neo
cells) from paraquat (PQ)-induced cytotoxicity and apoptosis. 32D/neo
cells were seeded at 1.5 × 105/ml in T-25 flasks and cultured
for 48 h. Cells were then treated with or without PQ (400 µM) in
presence or absence of QN (1 and 5 µM). Twenty-four hours later,
cells were harvested, viability was determined by trypan blue exclusion
(A), and percentage of apoptotic
nuclei was determined using Hoechst 33342 fluorescence
(B). Data are means ± SE of 4 individual observations. Each group was compared relative to PQ
treatment alone using 1-way ANOVA and Dunnett's multiple comparison
with control cells (* P < 0.05, ** P < 0.01).
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Fig. 2.
QN reduced internucleosomal DNA fragmentation in 32D/neo cells after PQ
exposure. 32D/neo cells were cultured and treated as described in Fig.
1. After 24 h of drug treatment, 1 × 106 cells were obtained, and DNA
was prepared by sequential proteinase K and RNase digestion. DNA was
subjected to electrophoresis in a 2% agarose gel. Gel was stained with
ethidium bromide and photographed under ultraviolet illumination. Note
formation of extensive internucleosomal DNA fragmentation,
corresponding to 180- to 200-bp ladders, that occurs with PQ (400 µM)
and was substantially reduced with simultaneous inclusion of 5 µM QN.
+, Presence; , absence.
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To mechanistically attribute the protective actions of QN to
PLA2 inhibition, it was important
to assess the protective potential of multiple and structurally diverse
PLA2 inhibitors. Because the
specificity of QN for various PLA2
types [secretory and cytosolic PLA2
(sPLA2 and
cPLA2,
respectively)] is not firmly established, we chose
pharmacological agents with varying specificity for
sPLA2 and
cPLA2. Manoalide produces
selective inhibition of sPLA2,
whereas AACOCF3 specifically
antagonizes cPLA2. Aristolochic
acid possesses mixed activity toward both enzyme types. Figure
3 shows the effects of these three specific
PLA2 inhibitors on PQ-induced
apoptosis. 32D/neo cells were treated with 400 µM PQ, which produced
~40% apoptotic cells within 24 h. None of these
PLA2 inhibitors attenuated PQ-induced apoptosis. Surprisingly, two agents,
AACOCF3 (50 µM) and aristolochic
acid (50 and 250 µM), enhanced apoptosis to over 80% when combined
with PQ treatment, which is similar to the apoptosis seen with
AACOCF3 and aristolochic acid
alone. None of the compounds reduced DNA fragmentation compared with PQ
alone. Thus, among inhibitors of
PLA2, QN was unique in its ability
to ameliorate the apoptotic process after exposure to the prototypical
oxidant PQ.

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Fig. 3.
Specific inhibitors of phospholipase
A2
(PLA2) failed to protect 32D
cells from PQ-induced apoptosis. 32D/neo cells were seeded as described
in METHODS and cultured for 48 h.
Cells were treated with PQ (400 µM) in absence or presence of
indicated concentrations of PLA2
inhibitors (brackets): arachidonyl trifluoromethyl ketone
(AACOCF3), aristolochic acid,
and manoalide. Twenty-four hours later, nuclear morphology was
evaluated using Hoechst 33342 fluorescence (similar to Fig.
1B), and genomic DNA was analyzed by
gel electrophoresis (similar to Fig. 2). Note that none of these
PLA2 inhibitors could inhibit
apoptosis similar to QN and, in several instances, enhanced PQ-induced
apoptosis. Nos. on top, concentrations of respective
inhibitors in µM.
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PLA2 activity is not modulated by PQ or
QN.
Because oxidative stress can potentially regulate
PLA2 activity, we next determined
the activity of this important regulatory enzyme after PQ exposure. We
also examined whether the concentration of QN used to protect 32D cells
from PQ apoptosis was sufficient to inhibit
PLA2 activity in these cells. For
these experiments, PLA2 activity
was assessed by the release of metabolically incorporated [3H]AA in the presence
and absence of PQ-induced oxidative stress. The
Ca2+ ionophore 4-Br-A-23187 was
used as a positive control. Figure 4 shows
the time-dependent release of
[3H]AA into the medium
over 7 h of treatment in the absence (Fig. 4A) and presence (Fig.
4B) of 400 µM PQ. In addition,
Fig. 4C depicts the activation of
PLA2 after 1 µM 4-Br-A-23187.
Note the rapid eightfold increase in AA release within 1 h of
4-Br-A-23187 treatment compared with control treatment, which indicated
the expression of functional
Ca2+-dependent
PLA2 within these cells. In
contrast, oxidative stress induced by 400 µM PQ failed to lead to
significant enzyme activation. The inclusion of 5 µM QN in all cases
failed to attenuate the basal or
Ca2+-activated release of
[3H]AA and indicated
that antiapoptotic concentrations of QN were below the level required
to inhibit PLA2 activity.

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Fig. 4.
PQ and QN did not affect PLA2
activation. 32D/neo cells were labeled overnight with
[3H]arachidonic acid
(AA) as described in METHODS.
Radiolabeled cells were then harvested, washed, and seeded into 24-well
plates (4 × 105 cells/0.5
ml) containing 100 µl of vehicle control
(A), PQ (final concentration, 1 mM;
B), or 4-bromo-A-23187
(4-Br-A-23187; 10 µM; C) with or
without QN (5 µM). At indicated times after drug treatment, plates
were removed and centrifuged. Aliquots of cell-free media were then
removed for quantification of radioactivity. Data are means ± SE
(SE within size of symbol) of triplicate observations from 1 experiment
that was replicated a second time. Note 8-fold increase in release of
[3H]AA from cells into
medium after 4-Br-A-23187 stimulation. PQ produced no release of
[3H]AA above that seen
in untreated cells. QN had no effect on basal or
Ca2+ ionophore-induced
PLA2 activity.
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We also directly evaluated phospholipid hydrolysis after PQ and QN
treatment of 32D cells by preparing total lipid extracts from
[3H]AA metabolically
labeled cells. The amount of
[3H]AA-derived
radioactivity was then determined in individual phospholipid and free
fatty acid pools after separation by TLC to assess
PLA2 activation. A typical
chromatogram of lipids extracted from 32D/neo cells is illustrated in
Fig. 5. Phosphatidylcholine (PC)
represented about one-half (51.1 ± 2.1%) of the total
phospholipid, with phosphatidylethanolamine (PEA) being the next most
prominent phospholipid (20.1 ± 1.6%). Additional phospholipids
detectable on the plates of abundance were sphingomyelin (8.9 ± 0.8%), PS (8.0 ± 1.0%), phosphatidylinositol (PI; 8.0 ± 0.9%), diphosphatidylglycerol (2.8% ± 0.4%),
lysophosphatidylcholine (0.6% ± 0.2%), and phosphatidic acid (0.3 ± 0.1%). No significant differences in the pattern of distribution
of phospholipid classes were apparent between control, PQ-treated, and
4-Br-A-23187-treated cells. As shown in Table
1, essentially no release of
[3H]AA was observed
from any of the phospholipid classes studied after a 3-h PQ exposure.
These data substantiate the conclusion that PQ failed to activate
PLA2 before apoptosis. In
contrast, the Ca2+ ionophore
4-Br-A-23187 induced release from several major phospholipid classes.
After a 3-h treatment with 1 µM 4-Br-A-23187, ~50% of the
[3H]AA was lost from
PEA and 40% was lost from PC and PI. Thus the Ca2+-regulated
PLA2 activity in 32D/neo cells
appeared to recognize multiple phospholipid substrates. Notably, PS was
resistant to hydrolysis by PLA2
despite preferential oxidation of this phospholipid during PQ-induced
oxidative stress. Essentially identical results were observed when QN
was included during the 4-Br-A-23187 treatment, with 43, 47, and 33%
of the [3H]AA released
from PEA, PI, and PC, respectively. In addition, the neutral lipid
fraction containing triglycerides lost ~30% of its content of
[3H]AA after
4-Br-A-23187 treatment. Despite the significant release of
[3H]AA from multiple
lipid sources within 32D/neo cells after 4-Br-A-23187 treatment, no
significant increase in radioactivity was observed within the free
fatty acid pool within these cells (data not shown), indicating that
hydrolyzed AA was rapidly transported from the cell into the
extracellular medium.

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Fig. 5.
A typical high-performance TLC 2-dimensional chromatogram of a total
lipid extract from 32D cells. Lipids were isolated and subjected to
2-dimensional TLC as described in
METHODS. FFA, free fatty acids; NL,
neutral lipids; DPG, diphosphatidylglycerol; PI, phosphatidylinositol;
PEA, phosphatidylethanolamine; PS, phosphatidylserine; PC,
phosphatidylcholine; SPH, sphingomyelin; PA, phosphatidic acid; LPC,
lysophosphatidylcholine.
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Table 1.
Release of [3H]AA from individual phospholipid classes
after paraquat and Ca2+-ionophore treatment of
32D/neo cells
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Table 2 shows the specific activity of
[3H]AA incorporation
normalized to Pi content of each
lipid for four major classes of phospholipid and demonstrates that
differences in specific incorporation of AA did not determine substrate
specificity. Note that PEA is approximately eightfold enriched in AA
compared with PC, and each appears to lose a similar percentage of
[3H]AA after
4-Br-A-23187 stimulation. PS was found to be relatively resistant to
hydrolysis; however, PS and PC showed similar specific incorporation of
AA. Thus, although PS and PC are similarly enriched in AA, PS does not
represent as efficient a substrate for
PLA2 as does PC in 32D cells.
QN inhibits early PS oxidation after PQ.
We next examined whether QN could act during the initiation of
apoptosis after PQ. On the basis of the previous observation of
selective early oxidation of PS early during PQ-induced apoptosis (8),
we measured PQ-dependent oxidation of
cis-PNA incorporated into individual
phospholipid classes with or without QN. Figure 6 shows that QN blocked the selective
oxidation of PS after PQ. Within 2 h, PQ oxidized ~25% of the
cis-PNA covalently incorporated into
PS, whereas no significant loss was observed in other phospholipid classes, including PC, PEA, PI, and sphingomyelin. The simultaneous inclusion of 5 µM QN, however, completely prevented the PQ-dependent oxidation of PS. Exposure of 32D/neo cells to QN alone had no effect on
the peroxidation of any of the phospholipid classes studied here.

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Fig. 6.
QN blocked selective peroxidation of PS after PQ exposure. 32D/neo
cells with incorporated cis-parinaric
acid (cis-PNA) were treated with or
without PQ (1 mM for 2 h) in presence or absence of QN (5 µM),
followed by extraction of phospholipids and resolution by HPLC.
Unoxidized cis-PNA content of each
phospholipid class was quantified using arbitrary fluorescence units in
each peak and then normalized to
Pi content of each peak. Data are
means ± SE of cis-PNA content of
each phospholipid class [PC
(A), PEA
(B), PS
(C), PI
(D), and SPH
(E)] based on 7 observations
obtained from 2 separate experiments. Statistical analysis was
performed by 1-way ANOVA and Dunnett's multiple comparisons with
control group. * Significant difference from PQ treatment alone
(P < 0.01).
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QN does not inhibit apoptosis after IL-3
withdrawal.
We next studied whether the effects of QN were specific for PQ-induced
apoptosis or whether the cytoprotective effects of this compound
extended to other apoptotic stimuli. 32D cells depend on the cytokine
IL-3 for continued survival in culture and undergo extensive apoptosis
after the removal of this growth factor. Figure 7 shows the time course of apoptosis (Fig.
7A) and the formation of
internucleosomal DNA cleavage (Fig.
7B) after IL-3 withdrawal in the
presence and absence of 5 µM QN. Note that the time-dependent increase in cells that exhibited nuclear condensation and fragmentation 48 h after IL-3 withdrawal was similar in the absence and presence of
QN. Thus, in contrast to its ability to inhibit PQ-induced apoptosis,
QN failed to protect 32D/neo cells from apoptosis after IL-3
withdrawal.

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Fig. 7.
QN failed to inhibit apoptosis after interleukin (IL)-3 withdrawal.
32D/neo cells were seeded at 1.5 × 105 cells/ml for 48 h. Cells were
washed and resuspended in complete culture medium with or without
IL-3-containing WEHI-conditioned medium. QN (5 µM) was also supplied
to indicated cells at this time. After 24 or 48 h, aliquots of cells
were processed to assess nuclear morphology
(A) and internucleosomal DNA
fragmentation (B). Data are means ± SE of 3-4 individual observations. Note that QN had no
effect on induction of apoptotic nuclear morphology or internucleosomal
fragmentation after IL-3 withdrawal. Stds, standards.
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DISCUSSION |
PQ-induced PS oxidation and apoptosis are independent of early
PLA2 activation.
Considerable evidence has emerged supporting a role for
PLA2 in the potentiation of cell
injury and death. TNF is a cytokine capable of mediating apoptotic cell
death on sensitive cell types and serves as an example of
PLA2 involvement with cell death. First, Voelkel-Johnson et al. (42) reported a positive correlation between cPLA2 activity in various
cell lines and their ability to be killed by TNF. Second, inhibition of
cPLA2 activity with AACOCF3 and anti-sense constructs
ameliorates the cytotoxic effect of TNF on adenovirus-infected cells
(40). Third, TNF-resistant L929 cells isolated by Hayakawa et al. (13)
lack cPLA2, and sensitivity to TNF
can be partially restored by transfection of cPLA2 cDNA into these cells.
PLA2 has also been implicated in other examples of cell death, including glutamate neuronal toxicity (12) and ischemic injury (2, 32).
Therefore, we initially hypothesized that early
PLA2 activation could play a role
in mediating apoptosis after PQ exposure. Several observations,
however, refute this notion. First, several specific inhibitors of
PLA2, such as
AACOCF3, manoalide, and
aristolochic acid, could not inhibit PQ-induced apoptosis. Second,
using two distinct assays, we observed absolutely no evidence for
activation of PLA2 after PQ
exposure. Third, the concentrations of QN required to inhibit apoptosis
in our studies were significantly less than those frequently used by
others to inhibit PLA2 in vitro
(3, 23) and failed to exert an inhibitory effect on
PLA2 measured in these studies.
Thus PLA2 activation does not
appear to immediately follow PS oxidation and participate in apoptosis
after PQ. Therefore, PQ-induced apoptosis does not resemble
TNF-mediated cell death in terms of its dependence on
PLA2 activation.
It is important to note, however, that
PLA2 activation may play a role in
other forms of cell death, such as necrosis, after oxidant exposure.
Our goal was to specifically study apoptosis; therefore, we chose a
concentration of PQ that killed cells primarily by apoptosis (8). In
this previous study, ~50% of 32D/neo cells were nonviable after 24-h
exposure to 400 µM PQ. On analyzing apoptotic nuclei, we extrapolated
that over 80% of those dead cells had undergone apoptosis. Thus
nonapoptotic cell death contributes little to the overall cytotoxicity
of PQ under these conditions. It is possible that
PLA2 blockers may have protective
effects in blocking necrotic cell death; however, experimental testing of this hypotheses requires a model of cell death different from that
applied here.
It could be argued that the failure of
PLA2 inhibitors to attenuate
apoptosis resulted from using concentrations below those required for
enzyme inhibition. We therefore assessed the ability of these
PLA2 inhibitors to attenuate AA
release in our experimental system. Neither 2 h of basal or
Ca2+-induced AA release was
affected by 1 or 5 µM manoalide. This inhibitor, however, is
relatively specific for the 14-kDa
sPLA2 and has a reported
EC50 of 0.2-0.3 µM for
inhibiting Ca2+-induced eicosanoid
production in peritoneal macrophages (26). Thus the failure of
manoalide to inhibit PLA2 in our
studies suggests the predominance of an 85-kDa
cPLA2 in controlling
Ca2+-dependent AA release in 32D
cells. In contrast, 250 µM aristolochic acid reduced
4-Br-A-23187-induced AA release by ~40% but failed to protect
against PQ apoptosis. Although still somewhat selective for
sPLA2, aristolochic acid probably
possesses efficacy against cPLA2
at this concentration (43). The
cPLA2-specific drug
AACOCF3 paradoxically increased
basal AA release (2-fold at 10 µM and 9-fold at 50 µM). This
release of AA, however, could not be temporally separated from a
dramatic and rapid incidence in apoptosis in these cells (Fabisiak,
unpublished observations). The nature of this AA release
is currently under investigation but probably does not represent
cPLA2 activation, since the
AACOCF3 concentrations used here
were above those required to inhibit
cPLA2 in other cell types (33).
Instead, it may represent the shedding of membrane-bound apoptotic
bodies or sPLA2 activation that
could occur as a sequela to apoptosis. Regardless of how these data are
interpreted, they still support that notion that
PLA2 activation is not responsible for the initiation of PQ-induced apoptosis.
Surprisingly, we found no evidence for the activation of
PLA2 during PQ-induced oxidative
stress and lipid peroxidation in these cells. These data contrast those
of Salgo et al. (35), who found oxidized phospholipids to be preferred
substrates for PLA2. Thus PS
oxidation observed after PQ may not be a sufficient signal to enhance
PLA2 activity. It is possible,
however, that because PS incorporated relatively little AA, PS
oxidation occurs primarily within other polyenoic fatty acids. Thus
hydrolysis and repair of oxidized PS could have gone undetected using
AA release in these studies. It should be stressed, however, that even
small amounts of oxidized PS could participate in modulation of various
aspects of apoptosis. For example, PS oxidation could lead to PS
externalization by specific interaction and inhibition with
lipid-transporting activities such as aminophospholipid translocase. It
is also possible that PLA2
activities can be differentially regulated under various oxidative
stresses. Although cPLA2 appears activated after
H2O2
treatment (3), porcine pancreatic
sPLA2 appears to be directly
inhibited by PQ and other superoxide anion-generating systems (11).
Thus different lipid-based signaling pathways may be operative for the
induction of apoptosis: those that are PLA2 dependent, such as after TNF
exposure, and those dependent on PS oxidation, such as after PQ, that
are independent on PLA2 activation.
To our knowledge, this is the first report of
PLA2 substrate specificity in
these nontransformed 32D murine myeloid progenitor cells. Multiple
PLA2 types have been described
that differ in structure, substrate specificity, and
Ca2+ sensitivity (7). It is
possible that activation of a
Ca2+-independent
PLA2 could be involved in
signaling apoptosis, since most of the inhibitors chosen for this study
are restricted in their effects to the
Ca2+-requiring enzymes. This is
unlikely, however, in light of the fact that we detected no AA release
or phospholipid hydrolysis after PQ, which would reflect activity of
any activated PLA2. Although we
have not as yet characterized the specific
PLA2 types responsible for AA
liberation after 4-Br-A-23187 stimulation of 32D cells, it is possible
to state that Ca2+-dependent
PLA2 in 32D cells can utilize
AA-containing PEA, PI, and PC as substrates. In contrast, PS appears to
be a relatively poor substrate for hydrolysis by
PLA2 despite its preferential oxidation during PQ-induced apoptosis.
QN has antiapoptotic effects not shared by other
PLA2 inhibitors.
We have shown that PQ-induced apoptosis can be inhibited
by the agent QN. The cytoprotective effects of QN were clearly
demonstrated by its ability to maintain cell viability and inhibit
several characteristics of apoptosis, including internucleosomal DNA
fragmentation and chromatin condensation. Importantly, our data also
indicate that QN acted early after PQ exposure to disrupt a potential
proapoptotic signal, namely, PS oxidation. Selective oxidation of PS is
associated with apoptosis and is inhibited by bcl-2
overexpression (8). Thus QN and Bcl-2 may inhibit signal transduction
pathways that utilize oxidation of specific membrane phospholipids as
downstream mediators for apoptotic signaling after oxidative stress.
Our laboratory has also observed that PS oxidation associated with apoptosis appears uniquely resistant to vitamin E analogs (unpublished observations).
It is important to note, however, that QN was ineffective at inhibiting
apoptosis after IL-3 withdrawal, whereas overexpression of
bcl-2 greatly attenuates apoptosis in this model (29) (data not shown). Thus significant differences must exist between these two
models with respect to the action of QN. Many diverse stimuli, including oxidative stress (4, 14, 8), growth factor withdrawal (29),
and TNF exposure (22) among others, presumably converge at some point
to initiate a common apoptotic signaling cascade. It is likely that
Bcl-2 acts early to disrupt the initiation of this final common pathway
of apoptosis that includes release of cytochrome
c from mitochondria (20), activation
of caspase proteases (31), and possibly PS oxidation. One potential
locus for the QN effect observed here is within an oxidant-specific
pathway not shared by other stimuli of apoptosis such as growth factor deprivation.
QN inhibits TNF cytotoxicity (21) and neuronal cell death after
glutamate exposure (12). In addition, QN protects myocytes and neurons
from ischemic cell death (2, 32) and the kidney from
cyclosporin-mediated toxicity (19). These studies usually attribute the
cytoprotective actions of QN to its ability to act as an inhibitor of
PLA2 (6, 24). QN,
however, is a prototypical cationic amphiphilic compound similar to
local anesthetics and chloroquine, with the potential to interact
directly with membrane phospholipids (16) and stabilize a variety of
organelle membranes including lysosomes (23). The presence of secondary
and tertiary amine groups within the QN molecule makes possible their
protonation at physiological pH and physical interaction specifically
with acidic phospholipids such as PS. In addition, QN can modulate a
variety of other membrane-associated functions such as
Ca2+ and
K+ channels (28, 34) and
acetylcholine receptors (16). QN itself, however, did not appear to
modulate Ca2+-dependent cellular
processes in these studies, since it failed to alter either basal or
Ca2+ ionophore-induced
[3H]AA release (Fig.
4). It is also possible that QN cytoprotection may be due to a direct
antioxidant action. QN can inhibit the release of oxygen radicals from
human alveolar macrophages (39) as well as suppress superoxide
production in a cell-free xanthine-xanthine oxidase system (23, 41).
The effects of QN on blocking PQ-induced PS oxidation support this
hypothesis. Studies are currently underway to further elucidate the
molecular mechanisms of QN cytoprotection during oxidative stress.
In summary, we showed that QN inhibited selective oxidation of PS and
subsequent apoptosis after PQ exposure. These effects appeared
independent of PLA2 inhibition. In
addition, no evidence of PLA2
activation was observed after PQ. Therefore, PS oxidation and other
components of apoptosis after PQ did not require
PLA2 activation and suggest that
multiple apoptotic signaling pathways exist. Some, as in the case of
TNF, appear to depend on PLA2
activation, whereas others such as PQ utilize selective oxidation of
membrane phospholipids and are
PLA2 independent. In addition,
these data point to a novel mechanism for QN cytoprotection not shared
by other PLA2 inhibitors. Clearly,
care must be taken in the interpretation of experiments utilizing QN as
a putative PLA2 inhibitor. Data suggest the potential usefulness of compounds structurally similar to
QN as tissue protective agents during exposure to oxidative agents such
as PQ.
 |
ACKNOWLEDGEMENTS |
We thank Dr. Daniel Johnson for the kind gift of the 32D/neo cells
and for helpful comments in the preparation of this manuscript.
 |
FOOTNOTES |
This work was funded, in part, by American Lung Association Research
Grant (to J. P. Fabisiak), National Cancer Institute Grant CA-61299 (to
J. S. Lazo), American Institute for Cancer Research Grants 9A50 (to J. S. Lazo) and 97B128 (to V. E. Kagan), Johns Hopkins Center for
Alternatives to Animal Testing Grant 96008 (to V. E. Kagan),
National Cancer Institute Oncology Research Faculty Development Program
(V. A. Tyurin), and International Neurological Science Fellowship
Program F05 NS10669, administered by National Institute of Neurological
Disorders and Stroke, National Institutes of Health in collaboration
with Unit of Neuroscience, Division of Mental Health and Substance
Abuse, World Health Organization (Y. Y. Tyurina).
V. A. Tyurin and Y. A. Tyurina are on leave from the Institute of
Evolutionary Physiology and Biochemistry, Russian Academy of Science,
St. Petersburg, Russia.
Address for reprint requests: J. P. Fabisiak, Dept. of Pharmacology,
E1313 Biomedical Science Tower, Univ. of Pittsburgh, Pittsburgh, PA
15261.
Received 15 September 1997; accepted in final form 6 February
1998.
 |
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