Departments of 1 Medicine and 3 Cell Biology and 2 Marion Bessin Liver Research Center, Albert Einstein College of Medicine, Bronx, New York 10461; 4 Departments of Medicine and of Biochemistry and Biophysics and Center for Gastrointestinal Biology and Disease, University of North Carolina, Chapel Hill, North Carolina 27599; and 5 IDUN Pharmaceuticals, Inc., La Jolla, California 92037
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ABSTRACT |
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Toxins convert
the hepatocellular response to tumor necrosis factor- (TNF-
)
stimulation from proliferation to cell death, suggesting that
hepatotoxins somehow sensitize hepatocytes to TNF-
toxicity. Because
nuclear factor-
B (NF-
B) activation confers resistance to TNF-
cytotoxicity in nonhepatic cells, the possibility that toxin-induced
sensitization to TNF-
killing results from inhibition of
NF-
B-dependent gene expression was examined in the RALA rat
hepatocyte cell line sensitized to TNF-
cytotoxicity by actinomycin
D (ActD). ActD did not affect TNF-
-induced hepatocyte NF-
B
activation but decreased NF-
B-dependent gene expression. Expression
of an I
B superrepressor rendered RALA hepatocytes sensitive to
TNF-
-induced apoptosis in the absence of ActD. Apoptosis was blocked
by caspase inhibitors, and TNF-
treatment led to activation of
caspase-2, caspase-3, and caspase-8 only when NF-
B activation was
blocked. Although apoptosis was blocked by the NF-
B-dependent factor
nitric oxide (NO), inhibition of endogenous NO production did not
sensitize cells to TNF-
-induced cytotoxicity. Thus NF-
B
activation is the critical intracellular signal that determines whether
TNF-
stimulates hepatocyte proliferation or apoptosis. Although
exogenous NO blocks RALA hepatocyte TNF-
cytotoxicity, endogenous
production of NO is not the mechanism by which NF-
B activation
inhibits this death pathway.
caspases; nitric oxide; inducible nitric oxide synthase; liver; hydrogen peroxide
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INTRODUCTION |
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TUMOR NECROSIS FACTOR- (TNF-
) activates a wide
array of cellular signaling pathways that result in divergent
biological responses depending on the physiological setting (30).
Although considerable interest has centered on responses in transformed cells because of the potential of TNF-
as an anticancer agent, TNF-
also regulates normal cell function. In vivo studies in the
liver have demonstrated that increased TNF-
expression modulates contrasting hepatocyte responses depending on the stimulus. During liver regeneration after partial hepatectomy, increased TNF-
production initiates or promotes hepatocyte proliferation (1, 36). In
contrast, in the setting of liver regrowth following toxin-induced
hepatocyte injury, TNF-
production triggers cell death rather than
proliferation. TNF-
expression is increased in animal models of
toxic liver injury (4, 8, 10, 21) and in humans during
alcohol-induced liver disease (26, 37). Although TNF-
production is
a normal response to tissue injury, TNF-
in the setting of
toxin-induced liver injury acts as a hepatocyte cytotoxin.
Investigations employing neutralizing antibodies or soluble receptors
to block biological activity of TNF-
have shown that TNF-
neutralization significantly reduces the liver damage (4, 11, 18) and
mortality (11) induced by a variety of toxins. These findings indicate
that the degree of cell death following toxin-induced liver injury
depends largely on the cytotoxic effects of TNF-
rather than on the
direct biochemical effects of the toxin or its metabolites. The
mechanism by which hepatotoxins sensitize hepatocytes to further injury
from TNF-
is unknown.
Studies in many cell systems, including hepatocytes, have shown that
inhibition of RNA or protein synthesis sensitizes cells to TNF-
toxicity, presumably by interfering with the upregulation of a
TNF-
-inducible protective factor (23). Liver toxins also interfere
with hepatocyte macromolecular synthesis, suggesting that they may
sensitize hepatocytes to TNF-
cytotoxicity by a similar mechanism.
Investigations in nonhepatic cells suggested that failure to upregulate
specific antioxidant defenses sensitized these cells to TNF-
toxicity (20, 34). However, prior studies have demonstrated that the
failure to upregulate hepatocellular antioxidant defenses such as
manganese superoxide dismutase (9) or glutathione (Xu, unpublished
data) cannot account for the sensitization of hepatocytes to
TNF-
-induced cell death. Recently it has been shown that blocking
the normal activation of the transcription factor nuclear factor-
B
(NF-
B) by TNF-
sensitizes fibroblasts, macrophages, and several
transformed cells to TNF-
cytotoxicity (2, 25, 31, 32). It has been
proposed that after TNF-
stimulation NF-
B activation is critical
in determining whether a cell enters into a pathway leading to survival
or to cell death (38). We hypothesized that toxins may therefore
sensitize hepatocytes to TNF-
toxicity by blocking the upregulation
of an NF-
B-dependent protective gene. These investigations assessed
the role that NF-
B activation plays in the sensitization of
hepatocytes to TNF-
cytotoxicity by actinomycin D (ActD).
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METHODS |
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Materials. All reagents were from Sigma (St. Louis, MO), unless otherwise indicated.
Cells and culture conditions.
The rat hepatocyte cell line RALA255-10G (6) was cultured in DMEM
(GIBCO BRL, Grand Island, NY) supplemented with 4% fetal bovine serum
(HyClone, Logan, UT), 2 mM glutamine, and antibiotics (GIBCO BRL).
These cells are conditionally transformed with a temperature-sensitive
T antigen. At the permissive temperature of 33°C they express T
antigen, remain undifferentiated, and proliferate. Culture of the cells
at the restrictive temperature of 37°C suppresses T antigen
expression, markedly slows growth, and allows differentiated gene
expression (6, 13). For these experiments, the cells were cultured at
33°C until confluent, trypsinized, and replated at 0.5 × 106 cells/dish on 35-mm plastic
dishes (Falcon, Becton Dickinson, Lincoln Park, NJ). After 24 h, the
medium was changed to DMEM supplemented with 2% fetal bovine serum,
glutamine, antibiotics, and 1 µM dexamethasone, and the cells were
placed at 37°C. After 3 days of culture at 37°C, the cells
received fresh serum-free medium containing dexamethasone. Medium was
supplemented with dexamethasone to optimize hepatocyte differentiation
as previously described (6). Cells were pretreated 24 h later with ActD
(15 ng/ml) for 30 min, and then mouse recombinant TNF- (R&D Systems, Minneapolis, MN) was added to some dishes, at a concentration of 10 ng/ml unless otherwise indicated.
[3H]thymidine incorporation. Cells were incubated with 2.5 µCi of [3H]thymidine (DuPont-NEN, Boston, MA) for 2 h. The medium was removed, and the cells were washed in PBS and then homogenized in 1 ml of 0.33 N NaOH; 0.3 ml of 40% TCA-1.2 M HCl was added, and the solution was centrifuged. The pellet was redissolved in 0.33 N NaOH, and the incorporated counts/min (cpm) were determined by scintillation counting. The DNA content was determined by means of bisbenzimidazole spectrofluorometry (24), and the amount of [3H]thymidine incorporation was calculated as cpm/mg of DNA.
MTT assay. The amount of cell death was determined by examining cell number with the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay (12). The cell culture medium was removed, and serum-free medium containing 1 mg/ml MTT was added to the cells. After a 1-h incubation, this medium was removed and the formazan product was solubilized in n-propanol. This solution was alkalinized with sodium hydroxide, and the absorbance at 560 nM was measured in a spectrophotometer. The percent cell survival was calculated by taking the optical density (OD) reading of cells given a particular treatment, dividing that number by the OD reading for the untreated, control cells, and then multiplying by 100. The accuracy of the MTT assay was validated by comparing it with the number of trypan blue-excluding cells recovered from trypsinized dishes.
Microscopic determination of apoptosis. Phase-contrast and fluorescence microscopy were conducted as previously described (35). The relative number of apoptotic cells was determined by fluorescent costaining with acridine orange and ethidium bromide as previously employed (35). The percent of cells with apoptotic morphology (nuclear and cytoplasmic condensation, nuclear fragmentation, membrane blebbing, and apoptotic body formation) was determined by examining >400 cells/dish. Necrosis was excluded by the absence of ethidium bromide staining.
Electrophoretic mobility shift assays.
Nuclear proteins were isolated by the method of Schreiber et al. (28)
with slight modification as previously described (35). Electrophoretic
mobility shift assays were performed with the use of a commercially
supplied oligonucleotide for the NF-B consensus sequence (Santa Cruz
Biotechnology, Santa Cruz, CA). The DNA binding reaction was performed
at room temperature for 20 min in a 20-µl reaction mixture consisting
of 5 µg of nuclear extract, 50 µg/ml polydeoxyinosinic-deoxycytidylic acid, 10 mM Tris (pH 7.5), 100 mM
NaCl, 1 mM EDTA, 1 mM dithiothreitol (DTT), 1 mg/ml BSA, 10% glycerol,
and 25,000 cpm 32P-end-labeled
oligonucleotide. After incubation, the samples were resolved on a 4%
polyacrylamide gel, dried, and subjected to autoradiography. For
supershift assays, 8 µg of anti-p50 NF-
B, anti-p65 NF-
B, or
anti-Stat3 antibody (Santa Cruz Biotechnology) were added to the
reaction mixture, and the incubation time was extended for an
additional 20 min.
Transfections and luciferase assays.
RALA hepatocytes were transfected with NF-B-Luc, a luciferase
reporter gene driven by three NF-
B binding sites (16), using Lipofectamine (GIBCO-BRL). All treatments were initiated 48 h after the
time of transfection. To assay luciferase activity, cells were washed
in PBS, lysed with a buffer containing 1% Triton X-100 (Promega,
Madison, WI), scraped from the dish, and centrifuged, and the cell
extract was assayed for luciferase activity in a luminometer. All
luciferase values were normalized for extract protein concentration.
Adenovirus construction and infection.
A recombinant replication-deficient adenovirus, Ad5IB, was
constructed as previously described (19). This adenovirus contains an
I
B in which serines 32 and 36 are mutated to alanines, driven by the
cytomegalovirus promoter-enhancer. The presence of the mutant I
B
sequence packaged into the recombinant Ad5 virus (Ad5I
B) was
confirmed by PCR and by Western blotting. Ad5I
B was grown in 293 cells and purified by banding twice on CsCl gradients. Titers of viral
particles were determined by optical densitometry, and recombinant
virus was then stored in 10% (vol/vol) glycerol at
20°C. A
control virus, Ad5LacZ, which contains the Escherichia coli
-galactosidase gene, was also grown and
purified as described above.
Protein isolation and Western blot analysis. Cells were scraped in the medium and centrifuged. The cell pellet was resuspended in lysis buffer containing 10 mM HEPES (pH 7.4), 42 mM MgCl2, 1% Triton, 1 mM phenylmethylsulfonyl fluoride, 1 mM EDTA, 1 mM DTT, and 2 µg/ml pepstatin A, leupeptin, and aprotinin. The solution was then mixed at 4°C for 30 min. After centrifugation, the supernatant was collected and the protein concentration was determined by the Bio-Rad protein assay (Bio-Rad, Hercules, CA).
Fifty micrograms of protein were heated in 1× SDS gel loading buffer [50 mM Tris (pH 6.8), 100 mM DTT, 2% SDS, 0.1% bromphenol blue, and 10% glycerol] at 100°C for 2 min. The samples were subjected to 10% SDS-PAGE and subsequently transferred to a nitrocellulose membrane (Schleicher & Schuell, Keene, NH) in a transfer buffer containing 39 mM glycine, 48 mM Tris (pH 8.3), 0.037% SDS, and 15% methanol, using a Bio-Rad Trans-blot SD semidry transfer cell to which 50 mA was applied for 18 h. All membranes were stained with Ponceau red to ensure equivalent amounts of protein loading and electrophoretic transfer among samples. Blocking of the membranes was performed using a solution (TBS-T) of 5% dry milk, 10 mM Tris (pH 8.0), 0.15 M NaCl, and 0.05% Tween 20 for 1 h. A rabbit anti-caspase-1 polyclonal IgG (Santa Cruz Biotechnology), rabbit anti-caspase-2 polyclonal IgG (Santa Cruz Biotechnology), rabbit anti-caspase-3 polyclonal IgG (33), or rabbit caspase-8 polyclonal IgG (29) was used as primary antibody at 1:1,000, 1:2,000, 1:4,000, and 1:2,000 dilutions, respectively, in 5% dry milk-TBS-T for 2 h. A goat anti-rabbit IgG conjugated with horseradish peroxidase (GIBCO BRL) was used as a secondary antibody at a 1:10,000 dilution in 5% dry milk-TBS-T blocking solution for 1 h. After use of a chemiluminescence detection system (Supersignal Ultra, Pierce, Rockford, IL), the membranes were exposed to Reflection film (DuPont-NEN). For poly(ADP-ribose) polymerase (PARP) immunoblots, the protein isolation procedure was altered slightly. Cultured cells were washed twice in cold PBS, scraped, and centrifuged. The cell pellet was resuspended in 100 µl of lysis buffer composed of 62.5 mM Tris (pH 6.8), 2% SDS, 10% glycerol, 2%Statistical analysis. All numerical results are reported as means ± SE and represent data from a minimum of three independent experiments.
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RESULTS |
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Coadministration of toxin converts the RALA hepatocyte
TNF- response from proliferation to cytotoxicity.
The effects of treatment with TNF-
alone and in combination with the
toxin ActD were determined. Treatment with 10-50 ng/ml TNF-
for
24 h was nontoxic to RALA hepatocytes by microscopic examination and
MTT assay. TNF-
alone did stimulate a proliferative cellular
response, as determined by
[3H]thymidine
incorporation. A 24-h treatment with 10 ng/ml TNF-
increased
[3H]thymidine
incorporation to 237% of control levels (639 ± 61 cpm/µg DNA for
control vs. 1516 ± 191 cpm/µg DNA for TNF-
treated cells).
Cell number also increased slightly at 24 h despite the use of
confluent cultures (111 ± 2.0% of control). Treatment with ActD
alone caused 14 ± 3.1% cell death, whereas combined ActD-TNF-
administration increased cell death to 32 ± 2.5%. Cell death
resulted from apoptosis, as determined by light and fluorescence
microscopy (data not shown). Thus, consistent with in vivo studies,
stimulation with TNF-
alone elicited a proliferative response,
whereas TNF-
in combination with a toxin caused increased cell
death.
ActD does not affect NF-B activation but inhibits
NF-
B-regulated gene expression.
The finding that resistance to TNF-
cytotoxicity in nonhepatic cells
is mediated by NF-
B activation (2, 25, 31, 32) suggested that a
potential mechanism by which ActD could sensitize hepatocytes to
TNF-
-induced apoptosis is through the inhibition of expression of an
NF-
B-dependent protective gene. To examine whether ActD directly
interfered with NF-
B activation, the effects of ActD on
TNF-
-induced increases in NF-
B DNA binding were determined. DNA
gel shifts demonstrated that protein binding to an NF-
B consensus oligonucleotide was equivalent in cells treated with either TNF-
alone or ActD-TNF-
at a variety of time points (Fig.
1). The NF-
B complex
activated by TNF-
treatment of RALA hepatocytes was composed of
p50-p65 dimers, as determined by supershifts (Fig. 2).
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Inhibition of NF-B activation sensitizes RALA
hepatocytes to TNF-
cytotoxicity.
To assess whether the failure to increase NF-
B-regulated gene
expression sensitized hepatocytes to TNF-
toxicity, the effect of
blocking NF-
B activation during TNF-
stimulation was determined. NF-
B activation was blocked by infecting cells with an adenovirus expressing a mutant I
B (Ad5I
B). This mutant I
B lacks the
normal I
B phosphorylation sites so that it irreversibly binds
NF-
B, preventing its activation. Although TNF-
treatment led to
high levels of NF-
B activation in both uninfected cells and cells infected with a control adenovirus containing the
-galactosidase gene (Ad5LacZ), NF-
B DNA binding activity was low in cells infected with Ad5I
B (Fig. 3). Inhibition of
NF-
B activation in this fashion sensitized RALA hepatocytes to rapid
cell death from TNF-
treatment in the absence of ActD. Only 6 h
after TNF-
administration, there was 50 ± 1.9% cell death in
Ad5I
B-infected cells compared with only 5 ± 0.9% death in
Ad5LacZ-infected controls (Fig.
4). After 24 h of treatment, cell
survival was still markedly decreased in Ad5I
B-infected cells (Fig.
4). Fluorescent staining to determine the mode of cell death
demonstrated that Ad5I
B-infected cells underwent rapid apoptosis
following TNF-
treatment (Fig.
5). Treated cells were examined for the
presence of PARP cleavage as an additional indication of apoptosis.
PARP cleavage occurred after TNF-
treatment in Ad5I
B-infected
cells but not in uninfected or Ad5LacZ-infected control cells (Fig.
6). Inhibition of NF
B-dependent gene
expression was therefore sufficient by itself to sensitize RALA
hepatocytes to TNF-
cytotoxicity.
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NF-B inactivation during TNF-
stimulation results in caspase activation.
TNF-
-induced cytotoxicity occurs through the mechanism of caspase
activation (38). To determine whether NF-
B inactivation sensitized
cells to TNF-
-induced cell death in a caspase-dependent manner, the
ability of caspase inhibitors to block this cell death was examined.
TNF-
-induced cell death in Ad5I
B-infected cells was inhibited by
the conventional caspase inhibitors YVAD-CMK and DEVD-CHO, which target
but are not specific for caspase-1 and caspase-3, respectively (Table
1). Cell death was even more effectively
inhibited by the experimental caspase inhibitors IDN-1529, which has
additional activity against caspase-6 and caspase-8, and IDN-1965, a
specific inhibitor of caspase-6 and caspase-8. (Table 1).
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NO prevents TNF--induced cell death in the absence
of NF-
B activation.
As a transcriptional regulator, NF-
B presumably mediates
hepatocellular resistance to TNF-
toxicity by increasing expression of protective cellular genes. The gene product iNOS is TNF-
inducible and is regulated by NF-
B, and NO, the product of iNOS, has
been implicated as a protective factor against TNF-
toxicity in
several cell types, including hepatocytes (14, 22). Thus iNOS could be
the NF-
B-inducible protective gene mediating resistance to TNF-
toxicity. The ability of NO to replace the protective function of
NF-
B activation was therefore determined. Ad5I
B-infected cells
were pretreated for 18 or 2 h before TNF-
administration with the NO
donor SNAP. Treatment with 750 µM SNAP led to a 56.7 ± 10.8%
inhibition of cell death after an 18-h pretreatment and to an 85.0 ± 3.0% inhibition with a 2-h pretreatment. The protective effects
of SNAP were dose dependent (data not shown).
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DISCUSSION |
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Inhibition of NF-B activation sensitizes a variety of cell types to
cytotoxicity from TNF-
(2, 25, 31, 32). Increased expression of an
NF-
B-regulated gene(s) may serve as the central control point in
determining whether the cellular response to TNF-
is survival or
apoptotic cell death (38). This finding also suggests the possibility
that the ability of toxins to sensitize normal cells to TNF-
cytotoxicity may be mediated through an inhibition of either NF-
B
activation or upregulation of specific NF-
B-dependent cellular
genes.
Our investigations examined the relationship of NF-B activation to
the toxin-induced sensitization of a hepatocyte cell line to TNF-
cytotoxicity because hepatocytes are known to be sensitized to TNF-
killing by toxins in vivo (4, 11, 18). Consistent with the hepatic
effects of TNF-
in vivo (1, 4, 11, 18, 35), the present in vitro
studies demonstrated that TNF-
alone stimulated RALA hepatocyte
proliferation in the absence of cell death, whereas the
coadministration of ActD and TNF-
led to cytotoxicity. This
conversion from a proliferative to a cytotoxic response was not the
result of toxin-mediated inhibition of NF-
B activation, since
NF-
B DNA binding was unaffected by the addition of ActD. ActD did
partially inhibit the upregulation of NF-
B-dependent gene
expression, potentially sensitizing RALA hepatocytes to TNF-
toxicity by preventing expression of a protective gene.
NF-B activation was essential for resistance to TNF-
toxicity,
since blocking activation with an I
B superrepressor rapidly sensitized RALA cells to TNF-
-induced cell death in the absence of
ActD. Previous in vivo investigations demonstrated that blocking the
NF-
B activation that normally occurs in rats following partial hepatectomy converts a hepatocellular proliferative response into one
of cell death, but the factor mediating this effect was not identified
(19). The present in vitro studies demonstrate that it is TNF-
that acts as a hepatocyte mitogen in the presence of NF-
B activation
but as a cytotoxin in the absence of NF-
B activation. Although our
data cannot completely exclude concurrent proliferative and toxic
effects, they demonstrate for the first time that selective inhibition
of NF-
B activation is sufficient to convert a cellular TNF-
response from proliferation to apoptosis. Despite the multitude of
complex cellular signaling events triggered by TNF-
stimulation, a
single transcription factor ultimately determines whether the cell has
the diametrically opposed responses of growth or cell death.
Previous studies in nonhepatic cells relating NF-B activation to
cellular resistance to TNF-
cytotoxicity did not address the
mechanism by which NF-
B inactivation led to cell death from TNF-
.
The present study demonstrates that the mechanism involves activation
of the caspase family of cysteine proteases that are the effectors of
apoptotic cell death (27). TNF-
-induced cell death mediated by
NF-
B inactivation was blocked by caspase inhibitors. In addition,
caspase-2, caspase-3, and caspase-8 were activated following TNF-
treatment in Ad5I
B-infected cells that underwent cell death but not
in resistant cells infected with Ad5LacZ. Thus TNF-
stimulation in
the absence of NF-
B activation led to caspase activation and
apoptosis. In contrast to a recent report in a mouse hepatocyte cell
line (3), infection with Ad5I
B in the absence of TNF-
administration did not result in caspase activation or cell death,
suggesting that constitutive NF-
B activation does not play a role in
preventing spontaneous hepatocellular apoptosis. The present data
suggest that TNF-
-induced signaling must trigger caspase activation
by a mechanism that is blocked when NF-
B activation upregulates a
protective cellular factor. This protective NF-
B-regulated gene
product may act directly on caspase-8, preventing its activation and
subsequent processing of downstream caspase-2 and caspase-3. Alternatively, TNF-
can directly activate caspase-2 through the adaptor protein RAIDD/CRADD, and NF-
B expression may block this initial activation of caspase-2 and the subsequent processing of
caspase-3 and caspase-8. Finally, an NF-
B-regulated gene product may
directly affect a cellular process upstream of caspases in the cell
death pathway, indirectly preventing caspase activation. For example,
recent investigations suggest that NF-
B may regulate the inhibitor
of apoptosis protein (IAP) family (7).
In the present investigations, NO inhibited cell death induced by
TNF- in the absence of NF-
B activation. NO is known to have both
pro- and antiapoptotic properties (reviewed in Ref. 15). NO has been
reported previously to inhibit ActD-TNF-
-induced cell death (14,
22). Kim et al. (22) demonstrated in rat primary hepatocytes that
ActD-TNF-
-induced cell death was blocked by NO administration. The
protective effect of NO in their study was dependent on a long
pretreatment time (12-18 h) and protein synthesis and was linked
to the induction of heat shock protein 70 expression (22). In our
investigations employing an identical SNAP concentration, the
protective effects of NO were independent of protein synthesis, present
even when NO was given 2 h after TNF-
administration, and not
reproduced by cGMP. These data suggest that the mechanism of NO
protection in our studies was posttranslational. NO has been reported
to directly inhibit caspase-3 activation by
S-nitrosylation (14). NO prevented
caspase-2 and caspase-3 activation and decreased caspase-8 activation
in RALA hepatocytes, although our studies do not prove a direct effect
of NO on these enzymes.
Inhibition of NO production did not sensitize RALA hepatocytes to cell
death from TNF- or ActD-TNF-
, demonstrating that endogenous NO
does not modulate hepatocellular sensitivity to TNF-
toxicity. These
data exclude iNOS as the NF-
B-dependent gene that mediates
hepatocellular resistance to TNF-
cytotoxicity. These findings
contrast with those of Kim et al. (22), who reported increased cell
death in ActD-TNF-
-treated primary hepatocytes with iNOS inhibition.
However, their studies actually demonstrated that additional
stimulation of hepatocyte iNOS by the cytokines interferon-
and
interleukin-1
was protective against ActD-TNF-
-induced death.
Their studies failed to show that iNOS inhibition caused cell death in
hepatocytes treated with TNF-
alone or worsened cell death in cells
treated with ActD-TNF-
. Hepatocyte iNOS expression and NO production
are stimulated to much higher levels by a combination of cytokines than
by TNF-
alone (17). Thus, although TNF-
alone may not stimulate
sufficient NO to protect against cell death, exposure of hepatocytes to
multiple cytokines in vivo may upregulate iNOS to sufficient levels to
be cytoprotective. Alternatively, NO that is released in large
quantities during injury by liver cells other than hepatocytes, such as
Kupffer cells, may still modulate hepatocyte TNF-
toxicity.
Consistent with this possibility are reports that inhibition of NO
produced predominantly by Kupffer cells in vivo during toxin-induced
liver injury increases hepatocyte cell death (5).
The identity of the NF-B-regulated gene product(s) that blocks
TNF-
-induced hepatocyte apoptosis remains to be determined. This
protective gene must express a protein that is specific to the TNF-
death pathway.
H2O2
or Cu each induced NF-
B activation and apoptotic cell death, and
H2O2-induced
apoptosis was blocked by caspase inhibitors (Czaja, unpublished data).
However, inhibition of NF-
B activation did not increase cell death
induced by
H2O2 or Cu or sensitize cells to death from lower concentrations of these
agents. Thus, although NF-
B activation is a common response to
environmental stresses that result in hepatocyte apoptosis, NF-
B is
not universally protective against all forms of caspase-dependent apoptosis.
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ACKNOWLEDGEMENTS |
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We thank Amelia Bobe and Anna Caponigro for secretarial assistance, Dr. Joseph L. Goldstein for providing the anti-caspase-3 antibody, and Dr. Janice Chou for providing the RALA255-10G cells.
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FOOTNOTES |
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This work was supported in part by National Institutes of Health Grants DK-34987, GM-41804 (to D. A. Brenner), and DK-44234 (to M. J. Czaja), a grant-in-aid from the American Heart Association, New York City Affiliate (to R. N. Kitsis), a Howard Hughes predoctoral fellowship award (to S. Bialik), and an Australian National Health and Medical Research Council research scholarship (to B. E. Jones).
R. N. Kitsis is the Charles and Tamara Krasne Faculty Scholar in Cardiovascular Research of the Albert Einstein College of Medicine.
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Address for reprint requests: M. J. Czaja, Marion Bessin Liver Research Center, Albert Einstein College of Medicine, 1300 Morris Park Ave., Bronx, NY 10461.
Received 13 February 1998; accepted in final form 7 July 1998.
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