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,
Departments of
*
Pathology,
Medicine, and
Microbiology and Immunology, State University of New York Health Science Center, College of Medicine, Syracuse, NY 13210
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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m). We show that, in Jurkat human T cells and
peripheral blood lymphocytes, Fas-induced apoptosis is preceded by 1)
an increase in reactive oxygen intermediates (ROI) and 2) an elevation
of m. These events are followed by externalization of
phosphatidylserine (PS), disruption of m, and cell
death. The caspase inhibitor peptides, DEVD-CHO, Z-VAD.fmk, and
Boc-Asp.fmk, blocked Fas-induced PS externalization, disruption of
m, and cell death, suggesting that these events are
sequelae of caspase activation. By contrast, in the presence of caspase
inhibitors, ROI levels and m of Fas-stimulated cells
remained elevated. Because ROI levels and m are
regulated by the supply of reducing equivalents from the pentose
phosphate pathway (PPP), we studied the impact of transaldolase (TAL),
a key enzyme of the PPP, on Fas signaling. Overexpression of TAL
accelerated Fas-induced mitochondrial ROI production,
m elevation, activation of caspase-8 and caspase-3,
proteolysis of poly(A)DP-ribose polymerase, and PS externalization.
Additionally, suppression of TAL diminished these activities.
Therefore, by controlling the balance between mitochondrial ROI
production and metabolic supply of reducing equivalents through the
PPP, TAL regulates susceptibility to Fas-induced apoptosis. Early
increases in ROI levels and m as well as the dominant
effect of TAL expression on activation of caspase-8/Fas-associated
death domain-like IL-1ß-converting enzyme, the most upstream member
of the caspase cascade, suggest a pivotal role for redox signaling at
the initiation of Fas-mediated apoptosis. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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1/Mch5) recruited via its N-terminal death effector
domain to DISC (11, 12). Sequential activation of ICE/caspase-1,
caspase-3, and related cysteine proteases results in the proteolysis of
several cellular substrates, which, in turn, leads to the
characteristic morphologic and biochemical changes of apoptosis (4, 10). Nevertheless, cross-linking of the Fas receptor on different cell
types may lead to different outcomes. For example, Fas transduces an
activation signal and stimulates proliferation in freshly isolated PBL
(13, 14) or in certain tumor cell lines (15). The mechanisms of
processing biologically opposing signals through Fas stimulation have
not yet been determined. Reactive oxygen intermediates (ROIs) have long been considered toxic by-products of aerobic existence; however, evidence is now accumulating that controlled levels of ROIs modulate various aspects of cellular function and are necessary for signal transduction pathways, including those mediating apoptosis (16, 17, 18, 19, 20, 21, 22). Because apoptosis and Bcl-2 protection were demonstrated in very low oxygen pressure, ROI may not be absolutely required for PCD (23). Nevertheless, increased production of ROIs was demonstrated in TNF (24, 25, 26) and Fas-mediated cell death (27, 28, 29, 30, 31, 32). A cell may normally generate 1011 ROI molecules/day (33). ROI production during apoptosis may be controllable by increased synthesis of reducing equivalents (34). A normal reducing atmosphere, required for cellular integrity, is maintained by GSH, which protects the cell from damage by excess ROIs (35). Synthesis of GSH from its oxidized form, glutathione disulfide, depends on NADPH produced by the pentose phosphate pathway (PPP) (35). In fact, a fundamental function of PPP is to maintain glutathione in a reduced state and thereby protect sulfhydryl groups and cellular integrity from emerging oxygen radicals.
The PPP comprises two separate, oxidative and nonoxidative, phases.
Reactions in the oxidative phase are irreversible, whereas all
reactions in the nonoxidative phase are fully reversible. The two
phases are functionally connected. The nonoxidative phase converts
ribose 5-phosphate to glucose 6-phosphate for utilization by the
oxidative phase and thus indirectly contributes to generation of NADPH.
Different enzymes are rate limiting in the two phases. The oxidative
phase primarily depends on glucose 6-phosphate dehydrogenase (G6PD)
(36), whereas transaldolase (TAL) is the rate-limiting enzyme for the
nonoxidative phase (27, 37). TAL catalyzes the transfer of a 3-carbon
fragment, corresponding to dihydroxyacetone, to
D-glyceraldehyde 3-phosphate, D-erythrose
4-phosphate, and a variety of other acceptor aldehydes (38). TAL
expression and enzymatic activity are regulated in a tissue-specific
(37, 39, 40) and development-specific manner (41). TAL overexpression
lowers G6PD and 6-phosphogluconate dehydrogenase (6PGD) activities and
NADPH and GSH levels and renders the cell highly susceptible to
apoptosis induced by serum deprivation, hydrogen peroxide, nitric
oxide, TNF-, and anti-Fas mAb (27). When TAL levels are reduced,
G6PD and 6PGD activities and GSH levels are increased, and apoptosis is
inhibited. TAL activity profoundly impacts the balance between the two
branches of PPP and the ultimate output of NADPH and GSH (27). These
findings are consistent with an overwhelming influence of TAL-catalyzed
dihydroxyacetone transfer reactions on the distribution of flux between
the PPP and nucleotide metabolism, which determines the overall
propagation of biochemical signals in a metabolic network
(42).
The present study shows that Fas-induced disruption of
m, phosphatidylserine (PS) externalization, and cell
death are preceded by elevation of mitochondrial and cytosolic ROI
levels and hyperpolarization of m. The degree of TAL
expression can determine the amount of Fas-induced ROI production, PS
externalization, and cell death. Moreover, TAL is shown to influence 1)
activation of caspases, including caspase-8, and 2) changes in
m. Overexpression of TAL accelerated Fas-induced ROI
production, activation of caspases, PS externalization, and cell death
in Jurkat human T cells. In contrast, suppression of TAL activity
abrogated these effects and inhibited Fas-induced PCD. A key finding is
that increased ROI production serves as an early and defining step in
Fas-induced apoptosis.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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PBMC were isolated from heparinized venous blood on a
Ficoll-Hypaque gradient and resuspended in RPMI 1640 medium
supplemented with 10% FCS, 2 mM L-glutamine, 100 IU/ml
penicillin, and 100 µg/ml gentamicin. Cells (106/ml) were
prestimulated with 5 µg/ml Con A for 5–7 days; subsequently,
monocytes/macrophages were removed by adherence (43), and apoptosis was
induced with 1 µg/ml Fas Ab CH-11 (MBL, Watertown, MA) (44, 45, 46). To
measure Con A-induced cell proliferation, 105 PBLs were
incubated in each well of a microtiter plate using six parallel
samples. The plates were incubated at 37°C in a humidified atmosphere
with 5% CO2 for 72 h. The cultures were pulsed with
0.4 µCi of [3H]TdR for 8 h, and
[3H]TdR incorporation was measured as previously
described (43). Jurkat cell lines, producing increased (L26-3/1 and
L26-3/2D1) and suppressed (L18-3/1 and L18-3/1D9) levels of TAL, were
previously described (27). Twenty-four hours before assays, Jurkat
cells were fed fresh medium and seeded at a density of 2 x
105 cells/ml, and cell death was induced with 50 or 100
ng/ml anti-Fas mAb CH-11. Apoptosis was monitored by observing cell
shrinkage and nuclear fragmentation and was quantified by trypan blue
exclusion (46). DNA fragmentation during apoptosis was monitored by
agarose gel electrophoresis (27). Apoptosis was also measured by flow
cytometry after concurrent staining with fluorescein-conjugated annexin
V (annexin V-FITC; R&D Systems, Minneapolis, MN; FL-1) and propidium
iodide (FL-2) as previously described (47, 48). Staining with
phycoerythrin-conjugated annexin V (annexin V-PE; R&D Systems) was used
to monitor PS externalization (FL-2) in parallel with measurement of
ROI levels and m, using DHR, DCF, or
DiOC6 fluorescence, respectively (FL1, see below). Staining
with annexin V alone or in combination with DHR or DiOC6
was conducted in 10 mM HEPES (pH 7.4), 140 mM NaCl, and 2.5 mM
CaCl2. Using three-color fluorescence, Fas-induced changes
in mitochondrial ROI levels, m, and PS
externalization within CD4 and CD8 T cells were concurrently analyzed
by parallel staining with DHR or DiOC6 (FL-1), annexin V-PE
(FL-2), and Quantum Red-conjugated CD3, CD4, and CD8 mAbs (Sigma, St.
Louis, MO; FL-3). Quantum Red contains two covalently linked
fluorochromes, PE and Cy5. PE absorbs light energy at 488 nm and emits
at 670 nm, in the excitation range of Cy5, which acts as an acceptor
dye.
Flow cytometric analysis of ROI production, m,
and membrane integrity
The production of ROIs was estimated fluorometrically using
oxidation-sensitive fluorescent probes
5,6-carboxy-2',7'-dichlorofluorescein-diacetate (DCFH-DA), DHR, and
hydroethidine (HE; Molecular Probes, Eugene, OR) as previously
described (27, 49, 50). Following apoptosis assay, cells were washed
three times in 5 mM HEPES-buffered saline, pH 7.4, incubated in
HEPES-buffered saline with 0.1 µM DHR for 2 min, 1 µM DCFH-DA for
15 min, or 1 µM HE for 15 min, and samples were analyzed using a
Becton Dickinson FACStar Plus flow cytometer (Mountain View, CA)
equipped with an argon ion laser delivering 200 mW of power at 488 nm.
Fluorescence emission from DCF (green) or DHR (green) was detected at a
wavelength of 530 ± 30 nm. Fluorescence emission from oxidized
HE, ethidium (red), was detected at a wavelength of 605 nm. Dead cells
and debris were excluded from the analysis by electronic gating of
forward and side scatter measurements. ROI levels, as determined by
fluorescence of control cells, served as a baseline for assessment of
increased ROI levels in response to Fas stimulation. While R123, the
fluorescent product of DHR oxidation, binds selectively to the inner
mitochondrial membrane, ethidium and DCF remain in the cytosol of
living cells (50). The m was estimated by staining
with 40 nm DiOC6 (Molecular Probes), a cationic lipophilic
dye (28, 51, 52), for 15 min at 37°C in the dark before flow
cytometry (excitation, 488 nm; emission, 525 nm recorded in FL-1). The
fluorescence of DiOC6 is oxidation independent and
correlates with m (52). DiOC6 staining
was complete after a 15-min incubation. DiOC6 fluorescence
was diminished 3- to 4-fold by 5 µM carbonyl cyanide
m-chlorophenylhydrazone (mClCCP) and 10-fold or
more by 50 µM mClCCP as previously described (53, 54).
m was also quantitated using a potential-dependent
J-aggregate-forming lipophilic cation,
5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolocarbocyanine
iodide (JC-1) (55). JC-1 selectively incorporates into mitochondria,
where it forms monomers (fluorescence in green, 527 nm) or aggregates,
at high transmembrane potentials (fluorescence in red, 590 nm) (55, 56). Cells were incubated with 1 µM JC-1 for 15 min at 37°C before
flow cytometry. Cotreatment with a protonophore, 5 µM mClCCP (Sigma),
for 15 min at 37°C resulted in decreased DHR, DiOC6, and
JC-1 fluorescence and served as a positive control for disruption of
mitochondrial transmembrane potential (52). Staining with 150 nM
bis-(1–3-dibutylbarbituric acid)trimethixine oxonol
(DiBAC4; excitation, 488 nm; emission, 525 nm) for 10 min
at room temperature was used to assess cell membrane potential (57).
TAL and G6PD activities and glutathione levels
TAL enzyme activity was tested in the presence of 3.2 mM
D-fructose 6-phosphate, 0.2 mM erythrose 4-phosphate, 0.1
mM NADH, and 10 µg of -glycerophosphate
dehydrogenase/triosephosphate isomerase at a 1:6 ratio at room
temperature by continuous absorbance reading at 340 nm for 6 min (58).
The enzyme assays were conducted in the activity range of 0.001–0.01
U/ml. G6PD was measured in the presence of 120 mM Tris (pH 7.7), 10 mM
MgCl2, 2 mM glucose 6-phosphate, 0.9 mM NADP, and 0.1 U/ml
6PGD (59). TAL activity levels correlated with changes in TAL
expression as determined by densitometry (model GS-700, Bio-Rad,
Hercules, CA). The total glutathione content was determined by the
enzymatic recycling procedure essentially as described by Tietze (60).
Cells (106) were resuspended in 50 µl of 4.5%
5-sulfosalicylic acid. The acid-precipitated protein was pelleted by
centrifugation at 4°C for 10 min at 15,000 x g. The
total protein content of each sample was determined using the Lowry
assay (61). The GSH content of the aliquot assayed was determined in
comparison to reference curves generated with known amounts of
GSH (27).
Caspase-3/CPP32 enzyme assay and protease inhibitors
CPP32 activity was measured by incubating cell extracts in 50 µl of 80 µM DEVD-7-amino-4-trifluoromethyl-coumarin (DEVD-AFC; Calbiochem, La Jolla, CA), 250 mM sucrose, 20 mM HEPES-KOH (pH 7.5), 50 mM KCl, 2.5 mM MgCl2, and 1 mM DTT for 15 min at 37°C (62, 63). The protein content of cell extracts was determined by the Lowry assay (61). Fluorescence (400–505 nm) after addition of 1 ml of ice-cold water was compared with a standard curve of AFC (Sigma). The specificity of the enzymatic reaction was tested by using caspase-3/CPP32 inhibitor DEVD-CHO and caspase-1/ICE inhibitor YVAD-CMK (Bachem, King of Prussia, PA) at a concentration range of 50–300 µM (62). Caspase inhibitors Z-Val-Ala-Asp(Ome).fmk (Z-VAD) and Boc-Asp.fmk (Boc-Asp) as well as noncaspase cysteine protease inhibitor, Z-Phe-Ala.fmk (Z-FA; Enzyme Systems Products (Livermore, CA) were tested at concentrations of 20, 50, and 300 µM (64).
Western blot analysis
Forty micrograms of total cell lysate in 10 µl/well was separated by SDS-PAGE and electroblotted to nitrocellulose (65). For visualization of poly(ADP-ribose) polymerase (PARP), cells were lysed in 62.5 mM Tris-HCl (pH 6.8), 6 M urea, 10% glycerol, 2% SDS, 0.00125% bromophenol blue, and 5% 2-ME; sonicated for 15 s; and boiled for 5 min (66). Nitrocellulose strips were incubated in 100 mM Tris (pH 7.5), 0.9% NaCl, 0.1% Tween-20, and 5% skim milk with the primary Abs, anti-PARP mAb C-2-10 (66), anti-FLICE/Mch5/caspase-8 (mAb 5F7 directed to C-terminal amino acids 176–460 of human FLICE; Panvera, Madison, WI), anti-transaldolase Ab 170 (38), and actin mAb C4 (Boehringer Mannheim, Indianapolis, IN) at a 1000-fold dilution at room temperature overnight. After washing, the blots were incubated with biotinylated secondary Abs and subsequently with horseradish peroxidase-conjugated avidin (Jackson ImmunoResearch Laboratories, West Grove, PA). Between incubations, the strips were vigorously washed in 0.1% Tween-20, 100 mM Tris (pH 7.5), and 0.9% NaCl. The blots were developed with a substrate comprised of 1 mg/ml 4-chloronaphthol and 0.003% hydrogen peroxide.
Statistics
Alterations in cell survival, ROI levels, caspase-3 and PPP enzyme activities, and GSH levels were analyzed by Student’s t test. Changes were considered significant at p < 0.05.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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m precede PS
externalization during Fas-induced apoptosis
In accordance with previous observations (27, 28), Fas-induced
apoptosis was associated with increased ROI production (Fig. 1
A). To assess changes in
intracellular ROI levels we used oxidation-sensitive fluorescent probes
DHR, DCFH-DA (27, 50, 67), and HE (53). DHR is nonfluorescent,
uncharged, and readily taken up by cells, whereas R123, the product of
DHR oxidation, is fluorescent, is positively charged, and binds
selectively to the inner mitochondrial membrane of living cells (50).
The fluorescence of this dye is an indicator of mitochondrial ROI
production and membrane integrity. DCFH-DA is also readily taken up by
cells and, after deacetylation to DCFH, is oxidized to its fluorescent
derivative, DCF, and remains in the cytosol (50). HE is oxidized into
ethidium by ROIs and remains in the cytosol (53). Using all three
oxidation-dependent fluorescent probes, significant increases in ROI
levels were observed in Jurkat cells at 4°C as early as 20 min after
addition of Fas Ab CH-11 (Fig. 1A).
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A, DHR fluorescence increased in annexin V-negative
cells, suggesting that Fas-induced elevation of ROI levels in
mitochondria occurred before PS externalization. ROI production
continued to increase twofold or more over the baseline on a
logarithmic scale (Fig. 1C). DCF and ethidium fluorescence
were also increased in annexin V-negative cells, indicating elevated
levels of ROI in the cytosol upon Fas stimulation (Fig. 1D).
ROI levels remained elevated in annexin V-positive cells until they
underwent apoptotic shrinking as determined by forward angle light
scattering as a direct measure of particle size (data not shown). With
precipitous decline of cell viability and size, 12–24 h after Fas
stimulation DHR fluorescence returned toward baseline levels, in
correlation with a decrease in m (see below),
possibly reflecting leakage of ROI secondary to damage of mitochondrial
and cellular membranes (50, 51, 52).
Previous studies suggested that a decline of m may be
an early event in apoptosis, including Fas-dependent signaling (68).
Since an early increase in DHR fluorescence is dependent on the
integrity of a negatively charged inner mitochondrial membrane, we also
examined changes in m using the potentiometric dyes,
DiOC6 (51, 52) and JC-1 (55, 56). As shown in Fig. 1B, DiOC6 fluorescence was increased in annexin
V-negative cells as early as 20 min after stimulation with Fas Ab. Red
fluorescence of JC-1 (FL-2) was also increased in Fas-treated cells
(Fig. 1B). m of annexin V-negative cells
remained elevated for several hours after Fas stimulation (Fig. 1C). Similarly, increased DiOC6 staining in
annexin V-negative cells in response to Fas stimulation was noted by
fluorescence microscopy (Fig. 2).
DiOC6 staining was diminished in shrunken annexin
V-positive cells. Control cells stained with JC-1 showed green
fluorescence, while Fas-treated cells gained red JC-1 fluorescence
(Figs. 1B and 2), consistent with a higher
m (55, 56).
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Fas-induced cell death is associated with increased ROI production in peripheral blood T lymphocytes
Stimulation of freshly isolated PBL for up to 3 days with 50
ng/ml, 100 ng/ml, or 1 µg/ml CH-11 Fas Ab had no toxic effect but,
rather, increased cell survival (data not shown). These results are
consistent with data from other laboratories suggesting that
stimulation of the Fas receptor alone transduces activation rather than
death signals (13, 14). Incubation of PBL for 20 min on ice with 1
µg/ml Fas Ab increased m, as detected by DiOC6 and
JC-1 fluorescence (Fig. 3A).
Previous studies suggested that prestimulation of T lymphocytes with
mitogenic lectins increases susceptibility to apoptotic signaling
through the Fas receptor (13). We used the lectin Con A to trigger
polyclonal activation of T lymphocytes. Stimulation with Con A
dramatically elevated m. After incubation with Fas
Ab, m of Con A-prestimulated PBL was further
increased (Fig. 3A). The activation signals associated with
short term Fas or Con A stimulation did not significantly change
mitochondrial ROI levels as measured by DHR fluorescence (Fig. 3A). In contrast, prestimulation with Con A for 5–7 days
sensitized PBL to Fas-induced apoptosis (Fig. 3, B and
C). Fas stimulation of PBL that had been preincubated with
Con A for at least 5 days triggered a rapid increase in mitochondrial
ROI production and m. These changes occurred in
annexin V-negative cells (Fig. 3, B and C).
Three-color fluorescence staining with DHR or DiOC6 (FL-1),
annexin V-PE (FL-2), and Quantum Red-conjugated CD3, CD4, and CD8 mAbs
(FL-3) revealed that Fas-induced ROI production and elevation of
m occurred in both CD4+ and
CD8+ T cells (Fig. 3, B and C). With
decline of cell viability and size, 12–24 h after Fas stimulation DHR
fluorescence returned toward baseline levels in parallel with a
decrease in DiOC6 and JC-1 fluorescence (data not shown),
possibly reflecting leakage of ROI secondary to damage of mitochondrial
and cellular membranes (50, 51, 52). Thus, Fas-induced oxidative stress and
m elevation preceded PS externalization in both
Jurkat lymphoblastoid T cells and PBL (
Figs. 1–3).
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m, and PS externalization during Fas-induced
apoptosis
To assess the timing of ROI production with respect to caspase
activation, the effects of caspase inhibitors DEVD, Z-VAD, and Boc-Asp
(62) on Fas-induced cell death were investigated. Jurkat cells were
pretreated for 3 h with caspase inhibitors (62) before addition of
Fas Ab CH-11. Following pretreatment with 300 µM DEVD-CHO, PS
externalization did not occur, while ROI levels in Fas-stimulated cells
remained significantly elevated (up to twofold on a logarithmic scale)
compared with those in control cells (p <
0.001; Fig. 4,
A and B). An increase in m was
detectable in annexin V-negative cells as early as 20 min after
stimulation on ice with Fas Ab (Fig. 4C).
m diminished in annexin V-positive cells (Fig. 4C). DEVD pretreatment did not influence the Fas-induced
elevation in m (Fig. 4C). However, DEVD
blocked PS externalization and the drop in m was
confined to annexin V-positive Jurkat cells. In agreement with
earlier results (62, 69), DEVD completely blocked Fas-induced cell
death of Jurkat cells. Similar to DEVD-CHO, ZVAD.fmk (50 µM) and
Boc-Asp.fmk (50 µM) completely blocked PS externalization without
affecting Fas-induced elevation in m (Fig. 4D) or increases in ROI levels assessed by DHR, DCF, and
ethidium fluorescence (not shown). Pretreatment with the caspase-1
inhibitor YVAD-CMK or the cysteine protease inhibitor
Z-FA.fmk (up to 300 µM for 3 h) did not significantly inhibit
cell death (data not shown), in accordance with a dominant role of
caspase-8 and caspase-3 in Fas-dependent apoptosis (11).
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m elevation, caspase
activation, and cell death are regulated by transaldolase
Disruption of the mitochondrial membrane potential has been
proposed as the point of no return in apoptotic signaling (28, 54, 68).
Mitochondrial membrane permeability is subject to regulation by an
oxidation-reduction equilibrium of ROIs, pyridine nucleotides (NADH/NAD
and NADPH/NADP), and GSH levels (70). Since the overall NADPH output of
PPP, GSH levels, and sensitivity to apoptosis are regulated by TAL, we
investigated the effect of changes in TAL activity at various
checkpoints of Fas-induced cell death. Stimulation of Jurkat L26-3/4
and L26-3/2D1 cells with increased TAL activity and decreased NADPH,
NADH, and GSH levels (27) resulted in accelerated PS externalization
and cell death (Figs. 5A) as
well as increased mitochondrial ROI production compared with those of
control Jurkat cells (Fig. 5B). Increases in
DiOC6 fluorescence within annexin V-negative compartments
and the drop in DiOC6 fluorescence within annexin
V-positive compartments were accelerated in Fas-treated L26-3/4 and
L26-3/2D1 cells (Fig. 6A).
Along the same line, JC-1 fluorescence was enhanced in Fas-treated
L26-3/4 and L26-3/2D1 cells (Fig. 6B). Thus, increases in
m were larger and occurred earlier in cells with
increased TAL expression (Fig. 6, A and B). By
contrast, mitochondrial ROI production (Fig. 5C), changes in
m (Fig. 6), and cell death were inhibited in
Jurkat L18-3/1 and L18-3/1D9 cells with decreased TAL activity and
increased GSH content (Fig. 5A).
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and 5). A
drop in m, occurring in annexin V-positive cells
(Figs. 4C and 6A), was also eliminated by DEVD
(Fig. 4C), suggesting that activation of caspases was
required for disruption of m. In contrast,
accelerated mitochondrial ROI production (Fig. 5C) and early
increases in m (Fig. 4C) were not affected
by DEVD pretreatment. These results indicated that stimulation of
mitochondrial ROI production and increases in m
preceded or occurred independently from activation of caspases in
Fas-induced apoptosis.
Activation of cysteine proteases have been described as both a
result and a cause of oxidative stress. To test the effect of TAL
activity on activation of proteases through Fas, we quantitated
cleavage of DEVD-AFC, a substrate of caspase-3 (62, 63) and caspase-8
(11, 71), and monitored proteolysis of caspase-8/FLICE and PARP, a
signature substrate of caspase-3 (72) in Jurkat cells with altered
levels of TAL expression. In control Jurkat cells, significant
increases in DEVD-AFC cleavage activity were noted 2 h after
stimulation with 50 ng/ml Fas Ab at 37°C (p
< 0.01; Fig. 7). Proteolysis of FLICE,
isoforms caspase-8/a and caspase-8/b (73), required at least 1-h
stimulation with 50 ng/ml Fas Ab at 37°C. PARP cleavage was
detectable after 4-h Fas stimulation. Cleavage of DEVD-AFC (Fig. 7),
PARP (Fig. 8), and FLICE (Fig. 9) was accelerated in L26-3/4 and
L26-3/2D1 cells with increased TAL expression compared with that in
control Jurkat cells. By contrast, cleavage of DEVD-AFC, PARP, and
FLICE was inhibited in L18-3/1 and L18-3/1D9 cells with suppressed TAL
expression (
Figs. 7–9).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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m. The
latter event appears to be the point of no return in the effector phase
of PCD, as it leads to the mitochondrial release of apoptogenic
proteins, which, in turn, activate downstream caspases and
endonucleases (4, 10, 68, 75, 76, 77, 78). It has been proposed that
lymphocytes treated with dexamethasone or stimulated with Fas first
reduce their m by an unknown mechanism and then
hyperproduce ROI that serve as PCD effector molecules (53, 68). While a
disruption of m with the uncoupling of electron
transport can indeed lead to respiratory burst, a reverse scenario,
i.e., uncontrolled production of ROI initiating the decline of
m, is more consistent with the generation of
mitochondrial permeability transition (79). Mitochondrial function
depends on the integrity of its inner lipid bilayer. Agents that cause
permeability transition or pore formation are either pro-oxidants or
direct cross-linkers of SH groups (79).
We previously showed that Fas-mediated cell death was associated with
ROI production, and its rate correlated with intracellular GSH levels
as regulated by TAL expression (27). Similar findings were reported at
the same time by other laboratories (28, 29, 30, 31, 32). In the present studies,
ROI production was assessed using oxidation-sensitive fluorescent
probes. In living cells, R123, the fluorescent product of DHR
oxidation, binds selectively to the inner mitochondrial membrane,
whereas DCF, the fluorescent product of DCFH oxidation, and ethidium,
the fluorescent product of HE oxidation, remain in the cytosol (50).
Thus, increased ROI levels in response to Fas stimulation were elevated
both inside and outside the mitochondria. In both PBL and Jurkat cells,
ROI production increased as early as 20 min after Fas stimulation
before activation of caspases, PS externalization, or a precipitous
decline of cell viability. The selective increase in DHR fluorescence
reflected increased mitochondrial ROI production at a time when
mitochondrial transmembrane potential and membrane integrity were
maintained (50). DiOC6 and JC-1 fluorescence were increased
as early as 20 min after stimulation with Fas Ab, suggesting that an
early increase in ROI production was accompanied by m
elevation, that is, a hyperpolarization of the inner mitochondrial
membrane. The dependence of elevated ROI levels on the maintenance of
m was underlined by the parallel inhibition of DHR,
DiOC6, and JC-1 fluorescence in the presence of the uncoupling agent
mClCCP. DiOC6 and JC-1 fluorescence decreased in annexin
V-positive cells with diminished size, possibly reflecting a disruption
of m (68). In correlation with Salvioli et al. (80),
the staining profile of DiOC6 was the most variable among
the potentiometric dyes. However, all three dyes, DiOC6,
rhodamine 123, and JC-1, consistently demonstrated an early elevation
of m/mean channel fluorescence in response to Fas
stimulation. Previous studies suggested that a decline of
m may be an early event in apoptosis, including
Fas-dependent signaling (68). Our data indicate that increased ROI
production and m elevation precede PS externalization
and a disruption of m. Similar findings of an early
increase in DHR fluorescence were recently suggested to represent a key
event in PCD that preceded cytochrome c release and
disruption of m (54). The release of cytochrome
c, beginning 2 h or later following Fas stimulation,
appears to be a consequence of mitochondrial membrane damage (81).
Activation of caspases has been considered a hallmark of PCD (4, 10, 74). An increased cleavage of DEVD-AFC, a substrate of caspase-3 (62, 63) and caspase-8 (11, 71), was demonstrated in Jurkat cells as early
as 2 h after stimulation with Fas Ab, in agreement with previous
findings (64). Pretreatment with the caspase inhibitors, DEVD, Z-VAD,
and Boc-Asp, completely abrogated Fas-induced PS externalization,
indicating that activation of caspase-3, caspase-8, and related
cysteine proteases was absolutely required for cell death (7, 8, 9, 74).
ROI levels were partially inhibited in DEVD-treated Jurkat cells,
suggesting that caspase-3 activation, perhaps through damage of
mitochondrial membrane integrity, contributes to ROI production and
serves as a positive feedback loop at later stages of the apoptotic
process. Nevertheless, ROI levels remained significantly elevated after
pretreatment with caspase inhibitors (up to twofold on a logarithmic
scale compared with control cells). This suggested that activation of
caspase-3 or caspase-8 was not required for increased ROI production
and m hyperpolarization. By contrast, DEVD, Z-VAD,
and Boc-Asp blocked PS externalization and the decline of
m in annexin V-positive Jurkat cells, suggesting that
disruption of m (1) was a relatively late event with
respect to ROI production and m hyperpolarization and
(2) depended on activation of caspase-3 and related proteases.
Disruption of m was previously suggested as a point
of no return in Fas-induced apoptosis of Jurkat cells, secondary to
activation of caspases (68, 74). The present data indicate that
increases in ROI levels and m precede disruption of
m during Fas-induced apoptosis of Jurkat cells.
Moreover, increased ROI production in addition to caspase activation
may be required to cause death of Fas-stimulated PBL.
Prestimulation with mitogenic lectins for 5 days was required to
sensitize peripheral blood T lymphocytes to apoptotic signaling through
the Fas receptor, in accordance with observations by others (13, 14, 82). Stimulation of freshly isolated PBL for up to 3 days with Fas Ab
was not toxic. However, incubation of PBL for as few as 20 min on ice
with 1 µg/ml Fas Ab or Con A increased m (detected
by DiOC6 and JC-1 fluorescence) without increasing ROI levels. By
contrast, prestimulation with Con A for 5–7 days sensitized PBL to
Fas-induced apoptosis. Fas stimulation of PBL after at least 5 days of
incubation with Con A triggered a rapid increase in ROI production and
m in annexin V-negative peripheral blood
CD4+ and CD8+ T lymphocytes. Thus, in two
different cell preparations (Jurkat lymphoblastoid T cells and PBL),
Fas-induced apoptosis was associated with increased mitochondrial ROI
levels and m elevation before PS externalization.
The Fas receptor is expressed on a variety of freshly isolated normal cells, among them PBL (4, 13, 14). Direct stimulation of PBL through Fas does not induce apoptosis but, rather, leads to cellular activation. To convert Fas-mediated signals from activation to apoptosis, 5 days of prestimulation with Con A were required. The process was accompanied by an early increase in mitochondrial ROI levels. This signaling conversion may mimic a state of antigenic and/or mitogenic overstimulation and depict a physiological process that eliminates unwanted cells. Sensitization to Fas-induced ROI production and apoptosis may be related to changes in metabolism, i.e., GSH depletion and accelerated DNA synthesis in activated T lymphocytes (21, 83). Interestingly, the PPP is the sole source of the NADPH needed for GSH synthesis and the ribose 5-phosphate used for synthesis of the DNA and other nucleic acids (35).
DEVD-AFC cleavage activity and proteolysis of caspase-8 and PARP were accelerated in cells with increased TAL expression and were inhibited in cells with suppressed TAL expression. Proteolytic activation of caspase-8 (FLICE/MACH) appears to be the first step in the death cascade at the cytosolic face of the Fas receptor (12). Cleavage of caspase-8 is initiated by its recruitment to DISC (11, 12, 74). Alternatively, caspase-8 can be activated by dimerization (71, 84). The dominant effect of TAL expression on cleavage of caspase-8 is consistent with a redox control mechanisms at the pinnacle of the death cascade. The redox mechanisms influencing assembly of DISC, oligomerization, and/or cleavage of FLICE are critical questions to be investigated.
TAL regulates key checkpoints in Fas-mediated apoptosis: mitochondrial
ROI formation, activation of caspase-8 and caspase-3,
m, PS externalization, and, ultimately, cell death,
through regulation of the PPP, where the enzyme controls the levels of
intracellular reducing equivalents. TAL overexpression in Jurkat or H9
human T cells down-regulates G6PD and 6PGD activities and decreases
NADPH, NADH, and GSH levels. Alternatively, decreased TAL expression
up-regulates G6PD and 6PGD activities and increases GSH (27, 85). TAL
overexpression increases sensitivity while suppression of TAL
diminishes sensitivity to Fas-induced cell death. In cells
overexpressing TAL, glucose 6-phosphate is profoundly depleted (27)
which may be directly responsible for the diminished G6PD activities
and NADPH/GSH levels and increased sensitivity to apoptosis.
Alternatively, increased G6PD activities and GSH levels of cells with
suppressed TAL activity may result from increased availability of
glucose 6-phosphate for generation of NADPH. Of note, glucose transport
can be lost within minutes of the stimulation of Jurkat cells with Fas
Ab (86), thereby blocking the cell’s ability to combat oxidative
stress and diminishing the survival of Fas-stimulated cells. The
results indicate that TAL is a dominant regulator of glucose
utilization and propagation of apoptotic signals in Fas-stimulated T
cells. The impact of transaldolase on Fas-induced mitochondrial ROI
production and changes in m as reported here implies
a central role for this enzyme in the PPP and propagation of
biochemical signals dependent on NADPH and GSH production (27, 42).
These findings are compatible with a pivotal role we describe for ROIs
in Fas signaling.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Andras Perl, State University of New York Health Science Center, 750 East Adams St., Syracuse, NY 13210. E-mail:
3 Abbreviations used in this paper: PCD, programmed cell death; DISC, death-inducing signaling complex; ICE, IL-1ß-converting enzyme; FLICE, Fas-associated death domain-like ICE; ROI, reactive oxygen intermediate; GSH, reduced glutathione; PPP, pentose phosphate pathway; G6PD, glucose 6-phosphate dehydrogenase; TAL, transaldolase; 6PGD, 6-phosphogluconate dehydrogenase; m, mitochondrial transmembrane potential; PS, phosphatidylserine; annexin V-FITC, fluorescein-conjugated annexin V; annexin V-PE, phycoerythrin-conjugated annexin V; DHR, dihydrorhodamine 123; DCF, 5,6-carboxy-2',7'-dichlorofluorescein; DiOC6, 3,3'-dihexyloxacarbocyanine iodide; DCFH-DA; DCF-diacetate; HE, hydroethidine; mClCCP, carbonyl cyanide m-chlorophenylhydrazone; JC-1, 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolocarbocyanine iodide; DEVD, Asp-Glu-Val-Asp; AFC, 7-amino-4-trifluoromethyl-coumarin; YVAD, Tyr-Val-Ala-Asp; Z-VAD, Z-Val-Ala-Asp; Z-FA, Z-Phe-Ala; PARP, poly(ADP-ribose) polymerase.
Received for publication June 24, 1998. Accepted for publication October 8, 1998.
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m) in Fas-stimulated Jurkat
cells. Cells were analyzed after exposure to 50 ng/ml Fas Ab for 20 min
(on ice) and 1 h (at 37°C). Dead cells and debris were gated out
by forward (FSC) and side (SSC) scatter measurements. A,
PS externalization and ROI production were concurrently monitored by
staining with annexin V-PE (FL-2) and DHR (FL-1), respectively (dot
plot, left column). The Fas-induced increase in ROI
levels is shown by overlay of DHR fluorescence of annexin V-negative
populations (histograms, right column). Open curves
correspond to control cells, while shaded curves represent Fas-treated
cells. The x-axis shows the log FL-1 fluorescence
intensity; the y-axis indicates the cell number. Values
over curves indicate the mean channel of DHR fluorescence of control (0
min) and Fas-treated cells (20 min and 1 h). B,
Left column, DiOC6 fluorescence (FL-1) of
annexin V-PE-negative cells. The mean channel of histograms and the
percentage of cells with increased 
