Nitric oxide mediates hepatocyte injury
Jiang Huai
Wang,
H. Paul
Redmond,
Qiong
Di Wu, and
David
Bouchier-Hayes
The Royal College of Surgeons in Ireland, Department of Surgery,
Beaumont Hospital, Dublin 9, Ireland
 |
ABSTRACT |
The degree of acute hepatic failure after
severe trauma and sepsis is related to the extent of hepatocyte (HC)
damage and cell death resulting from either necrosis or apoptosis. We
have previously demonstrated that tumor necrosis factor-
(TNF-
)
and lipopolysaccharide (LPS) can directly lead to HC necrosis, but not
apoptosis. To date, the reactive oxygen intermediates (ROI) and nitric
oxide (NO) have been shown to play a potential role in the induction of
cell apoptosis. However, it is unknown whether ROI and NO are involved
in HC cell death. Therefore, in this study we tested the hypothesis
that NO and ROI exert different effects on HC cell death. TNF-
and
LPS alone failed to induce HC apoptosis but when combined with
antioxidants resulted in HC apoptosis and DNA fragmentation, which is
correlated with an increase in NO production. This effect was
attenuated by the NO synthase inhibitor NG-monomethyl-L-arginine
(L-NMMA). Moreover, the NO donor
sodium nitroprusside resulted in HC apoptosis and cell damage as
represented by hepatocellular enzyme release. Antioxidants inhibited
TNF-
- and LPS-mediated ROI generation and peroxynitrite formation in HC. TNF-
- and LPS-induced HC damage could be further reduced by the
combination of antioxidants and
L-NMMA. These results indicate that NO is involved in HC injury, primarily through the induction of HC
apoptosis.
apoptosis; reactive oxygen intermediates; antioxidant; lipopolysaccharide; tumor necrosis factor-
; peroxynitrite
 |
INTRODUCTION |
DURING THE MANAGEMENT of trauma and infectious
diseases, the systemic inflammatory response syndrome (SIRS) and
ensuing multiple organ dysfunction syndrome are major clinical
complications (9). In particular, despite current medical and surgical
advances, acute hepatic failure after severe trauma and sepsis is still associated with a high mortality rate (63). The degree of acute hepatic
failure is believed to be related to the extent of hepatocyte (HC)
damage and cell death resulting from either necrosis or apoptosis. We
have previously demonstrated that the proinflammatory mediators lipopolysaccharide (LPS) and tumor necrosis factor-
(TNF-
) can directly lead to HC damage as represented by hepatocellular enzyme release and cell necrosis, but they fail to induce HC apoptosis (57).
There is a growing body of evidence to implicate the reactive oxygen
intermediates (ROI) as potential mediators in the induction of
apoptosis in different types of eukaryotic cells (1, 11, 59, 61). We
have previously demonstrated that ROI are involved in human neutrophil
(61, 62) and endothelial cell apoptosis (58). More recently, nitric
oxide (NO), a free radical derived from a guanido nitrogen of
L-arginine, has also been
implicated as an inducer of apoptosis in a number of different cell
types, including macrophages (3), chondrocytes (8), pancreatic
-cells (31), and tumor cells (14). Furthermore, NO has been shown to
be cytotoxic and involved in cell killing by a necrotic process, either
by itself or through the formation of peroxynitrite with superoxide
anion (10, 25, 43). However, it is unknown whether ROI and NO are
involved in HC death.
In the present study, we tested the hypothesis that ROI and NO may
exert different effects on HC death. We report here that TNF-
and
LPS alone failed to induce HC apoptosis but resulted in increased NO
production and cell apoptosis when combined with antioxidants. This
effect is attenuated by the NO synthase inhibitor NG-monomethyl-L-arginine
(L-NMMA). Furthermore, the NO
donor sodium nitroprusside (SNP) induces not only HC apoptosis but also
HC damage as represented by hepatocellular enzyme release. Moreover, TNF-
and LPS-mediated hepatocellular enzyme release can be reduced by the combination of antioxidants and
L-NMMA. These results indicate that induction of NO results in HC apoptosis and, to a lesser degree,
HC damage.
 |
MATERIALS AND METHODS |
Reagents.
The following reagents were used for the isolation and culture of rat
HC and assessment of HC cell death. DMEM, RPMI 1640, HBSS without
Ca2+ and
Mg2+, fetal calf serum (FCS),
penicillin, streptomycin sulfate, 0.05% trypsin-0.02% EDTA solution,
glutamine, and insulin were purchased from GIBCO BRL (Paisley,
Scotland, UK). LPS (Escherichia coli 055:B5), SDS, sodium chloride, sodium phosphate, calcium chloride, sodium bicarbonate, potassium chloride, sodium citrate, SNP, superoxide dismutase (SOD; 5,100 U/mg protein), catalase (CAT; 2,000 U/mg protein), glutathione (GSH), DMSO,
N-acetylcysteine (NAC), HEPES, EDTA,
Tris, Triton X-100, glucose, propidium iodide (PI), and collagenase
(type IV) were purchased from Sigma (St. Louis, MO). Recombinant human
TNF-
(2 × 107 U/mg) was
obtained from Genzyme (Cambridge, MA).
NG-monomethyl-L-arginine
(L-NMMA) was from ICN
(Cleveland, OH). RNase A and
X174
DNA/Hae III markers were obtained from
Boehringer Mannheim Biochemica (Mannheim, Germany) and Promega
(Madison, WI), respectively.
Isolation of rat HC.
Adult male pathogen-free Sprague-Dawley rats weighing 200-300 g
(obtained from Charles River Breeding Laboratories, Kent, UK) were
fasted overnight before experimentation and allowed water ad libitum.
Rat HC were isolated by a modification of the techniques of Seglen (52)
and Doolittle and Richter (19), and all of the procedures were
performed under sterile conditions. Briefly, the rats were anesthetized
with inhalation of halothane (May and Baker, Dagenham, UK), and then
the peritoneal cavity was opened to expose the portal vein. An 18-gauge
needle connected to a Masterflex perfusion pump (Cole-Parmer, Niles,
IL) with a three-way stopcock was introduced into the portal vein.
While the inferior vena cava was being cut, the liver perfusion was
begun at a constant speed of 30 ml/min with 500 ml of
Ca2+-free HEPES buffer solution
containing 160.8 mM sodium chloride, 3.15 mM potassium chloride, 0.7 mM
sodium phosphate, 33 mM HEPES, and 5% glucose maintained at 37°C.
This was immediately followed by perfusion of 300 ml
Ca2+-free HEPES buffer solution
supplemented with 0.025% collagenase type IV and 0.075% calcium
chloride at a constant speed of 15 ml/min at 37°C. After the
perfused liver was removed and dissociated, the cells, resuspended in
80 ml of cold Ca2+-free HEPES
buffer solution, were passed through a sterile 180-µm stainless steel
sieve (Endecotts, London, UK) into a 125-ml beaker. Rat HC were
separated from nonparenchymal cells by centrifugation at 50 g for 2 min, four times. HC pellets
were resuspended in complete RPMI 1640 medium containing 10% FCS,
penicillin (100 U/ml), streptomycin sulfate (100 µg/ml), 2 mM
glutamine, 28 mM sodium bicarbonate, 8 mM HEPES, and 0.16 U/ml insulin.
The remaining Kupffer cells were further eliminated by allowing Kupffer
cells to adhere to 60-mm dishes for 1 h at 37°C in 5%
CO2 condition. This procedure
yielded HC preparation with less than 5% Kupffer cell contamination as
determined by esterase staining and with a viability greater than 80%
according to the trypan blue exclusion. Before experimentation HC were
cultured for 12 h at 37°C in humidified 5%
CO2 conditions, and culture medium
was then replaced with fresh complete RPMI 1640.
Assessment of HC apoptosis.
For determination of apoptosis by flow cytometry and DNA gel
electrophoresis, HC were plated at 1 × 106 cells per 40-mm dish (Nunc;
GIBCO BRL) with different treatments at 37°C in humidified 5%
CO2 conditions. After 12- and 24-h
incubation, HC were detached by using a low concentration of
trypsin-EDTA at 0.025%-0.01% for detecting apoptosis. For detection
of apoptosis by ELISA, HC were plated at 1 × 105 cells/well in 24-well,
flat-bottom plates (Falcon, Lincoln Park, NJ) and incubated with
different treatments for 12 and 24 h at 37°C in humidified 5%
CO2 conditions.
Flow cytometry.
HC apoptosis was assessed according to the percentage of cells with
hypodiploid DNA, using the PI staining technique as previously described (44). Briefly, after centrifugation the cell pellets were
gently resuspended in 1 ml of hypotonic fluorochrome solution (50 µg/ml PI, 3.4 mM sodium citrate, 1 mM Tris, 0.1 mM EDTA, 0.1% Triton
X-100) and incubated in the dark at 4°C overnight before they were
analyzed by a FACScan flow cytometer (Becton Dickinson, Mountain View,
CA). The forward scatter and side scatter of HC particles were
simultaneously measured. The PI fluorescence of individual nuclei with
an acquisition of fluorescence channel 2 (FL2) was plotted against
forward scatter, and the data were registered on a logarithmic scale.
The minimum number of 5,000 events was collected and analyzed on the
software Lysis II. Apoptotic HC nuclei were distinguished by their
hypodiploid DNA content from the diploid DNA content of normal HC
nuclei. The forward threshold was raised to exclude HC debris from
analysis. All measurements were performed under the same instrument
settings.
ELISA kit.
An ELISA kit (Boehringer Mannheim), which quantitatively detects
cytosolic histone-associated DNA fragments, was used to assess apoptosis in adhered HC (38, 39). DNA fragments were detected according
to the procedures described in the ELISA kit. Briefly, the cytosolic
fraction (13,000 g supernatant) of HC
was used as the antigen source in a sandwich ELISA with a primary
antihistone monoclonal antibody coated on the microtiter plate and a
second anti-DNA monoclonal antibody coupled to peroxidase. The
percentage of DNA fragmentation was calculated according to the
following formula
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DNA gel electrophoresis.
Gel electrophoresis for HC DNA fragmentation was carried out according
to a modified procedure for assaying DNA fragmentation in total genomic
DNA (41). Briefly, after centrifugation the cell pellets were
resuspended with 20 µl of lysis buffer (pH 8.0, 20 mM EDTA, 100 mM
Tris, and 0.8% N-lauroylsarcosine)
and 10 µl of RNase A (pH 4.8, 1 mg/ml containing 100 mM sodium
acetate and 0.3 mM EDTA) for 6 h at 37°C in a water bath. After
treatment with 10 µl of proteinase K (20 mg/ml) overnight at 50°C
in a water bath, the DNA preparations were added with 5 µl of loading
buffer (10 mM EDTA, 0.25% bromophenol blue, and 50% glycerol) and
electrophoresed on 1.5% agarose gel containing 0.3 µg/ml of ethidium
bromide in buffer containing 2 mM EDTA, 89 mM Tris, and 89 mM boric
acid, pH 8.0 (TBE buffer), for 3 h. An
Hae III digest of
X174-DNA was applied to each gel to provide size markers of 1.4, 1.1, 0.9, 0.6, and
0.3 kbp, respectively. Gels were photographed under ultraviolet transillumination.
Assay for hepatocellular enzyme release.
Aliquots of 500 µl of HC suspension (2 × 105 cells/ml) were seeded in
24-well, flat-bottom plates (Falcon) and incubated with 500 µl of
medium containing different treatments for 12 and 24 h at 37°C in
humidified 5% CO2 conditions.
Aspartate aminotransferase (AST) and lactate dehydrogenase (LDH)
activities in the supernatant of HC cultures were measured according to
the procedures of AST and LDH diagnostics kits (Sigma). The total
releasable AST and LDH activities were determined after different
incubation periods by lysis of the cells with the detergent Triton
X-100 (1% in HBSS) at 37°C for 30 min. Detection of AST and LDH
activity was performed on an ultraviolet/visible scanning spectrophotometer (CPU 8720; Philips, Holland) at 505 and 465 nm,
respectively. AST activity and LDH activity were measured as
Sigma-Frankel units per milliliter and Berger-Broida units per
milliliter, respectively. The results of AST and LDH release were
expressed as a percentage of the total releasable enzyme.
Assay for hepatocellular NO production.
HC were seeded in 24-well, flat-bottom plates (Falcon) at 1 × 105 cells/well and incubated with
different treatments at 37°C in humidified 5%
CO2 conditions. NO was measured
indirectly by determination of the concentration of the stable end
product, nitrite (24). Briefly, 100-µl aliquots were removed from HC
cultures and incubated with an equal volume of Griess reagent (1%
sulfanilamide, 0.1% N-1-naphthylethylene diamide
dihydrochloride in 2.5% phosphoric acid) for 10 min at 37°C.
Absolute values were determined using sodium nitrite as standard.
Absorbance was read at 550 nm on a microplate reader.
Assay for ROI generation and peroxynitrite formation in HC.
HC were plated at 1 × 106
cells per 40-mm dish (Nunc, GIBCO BRL) with different treatments at
37°C in humidified 5% CO2
conditions. After incubation for indicated time points, HC were
detached, using 0.025% trypsin-0.01% EDTA to detect ROI generation
and peroxynitrite formation.
The intracellular generation of ROI in HC was assessed with the use of
the fluorescent probe 2',7'-dichlorofluorescein diacetate (H2DCFDA) (Molecular Probes, Eugene, OR) as described
previously (23). Briefly, after experiments HC were washed twice and
resuspended in 200 µl PBS at 5 × 105 cells/ml. Cells were loaded
with 20 µM H2DCFDA and incubated in a 37°C water bath
for 10 min. The measurement of ROI generation was performed on a
FACScan flow cytometer (Becton Dickinson) for detecting the log of the
mean channel fluorescence intensity with an acquisition of
fluorescence channel 1 (FL1). The minimum number of 5,000 events
was collected and analyzed on the software Lysis II.
Peroxynitrite formation in HC was detected with the use of the
fluorescent probe dihydrorhodamine-123 (Molecular Probes) as described
previously (34, 64). Briefly, after experiments, 200 µl of HC
suspensions (5 × 105
cells/ml) were incubated with 20 µl of dihydrorhodamine-123 for 10 min at room temperature. The generation of peroxynitrite in HC was
detected based on the generation of the fluorescent product (rhodamine)
through the reaction of peroxynitrite with dihydrorhodamine-123, on a
FACScan flow cytometer (Becton Dickinson) for detecting the log of the
mean channel fluorescence intensity with an FL1. The minimum number of
5,000 events was collected and analyzed on the software Lysis II.
Statistical analysis.
All data are means ± SD. Statistical analysis was performed using
ANOVA. Differences were judged to be statistically significant when the
P value was less than 0.05.
 |
RESULTS |
LPS and TNF-
, in the presence of antioxidants,
result in HC apoptosis, which is abrogated by the NO synthase inhibitor
L-NMMA.
HC were treated with LPS and TNF-
in the presence or absence of
antioxidants (SOD, CAT, DMSO, NAC, and GSH) for 12 and 24 h. There was
no HC apoptosis found after 12 h incubation (data not shown). However,
after 24 h incubation as shown in Fig. 1, A and
C, TNF-
in combination with either
SOD, DMSO, NAC, or GSH resulted in a significant increase in HC
apoptosis (P < 0.05 vs. TNF-
alone), whereas TNF-
alone did not induce HC apoptosis. TNF-
plus
CAT failed to induce HC apoptosis. Similar results were found when HC
were treated with LPS alone and in combination with the antioxidants
(Fig. 1, B and
D). The addition of the NO synthase
inhibitor L-NMMA significantly
attenuated HC apoptosis induced by either TNF-
or LPS, in the
presence of SOD, DMSO, and GSH (P < 0.05 vs. LPS or TNF-
plus the antioxidant) (Fig. 2). These findings were further confirmed
by DNA gel electrophoresis. TNF-
in combination with either SOD,
DMSO, NAC, or GSH induced a typical DNA "ladder" pattern, which
indicates the occurrence of DNA fragmentation and thus apoptosis,
whereas TNF-
alone failed to cause DNA fragmentation.
L-NMMA prevented DNA
fragmentation induced by TNF-
plus the antioxidant (Fig.
3). Antioxidants alone did not cause HC
apoptosis (data not shown).

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Fig. 1.
Induction of hepatocyte (HC) apoptosis after exposure to either tumor
necrosis factor- (TNF- ) (A and
C) or lipopolysaccharide (LPS)
(B and
D) in the presence of antioxidants.
HC apoptosis (A and
B) and DNA fragments
(C and
D) were assessed as described in
MATERIALS AND METHODS after incubation
with either TNF- (25 ng/ml) or LPS (0.1 µg/ml) in presence or
absence of the antioxidants superoxide dismutase (SOD; 400 U/ml),
catalase (CAT; 2,000 U/ml), DMSO (0.5%),
N-acetylcysteine (NAC; 7.5 mM), and
glutathione (GSH; 7.5 mM) at 37°C in 5%
CO2 for 24 h. Data are means ± SD and are representative of 4 separate experiments.
* Statistically significant compared with TNF- alone or LPS
alone, P < 0.05.
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Fig. 2.
Attenuation of HC apoptosis by the nitric oxide (NO) synthase inhibitor
NG-monomethyl-L-arginine
(L-NMMA). HC were treated with
TNF- (25 ng/ml), LPS (0.1 µg/ml), SOD (400 U/ml), DMSO (0.5%),
GSH (7.5 mM), and L-NMMA (0.5 mM) in different combinations at 37°C in 5%
CO2 for 24 h. HC apoptosis
(A and
B) and DNA fragments
(C and
D) were assessed as described in
MATERIALS AND METHODS. Data are means ± SD and are representative of 4 separate experiments. Statistical
significances were compared with TNF- plus the antioxidant (SOD,
DMSO, GSH) (A and
C) or LPS plus the antioxidant (SOD,
DMSO, GSH) (B and
D).
@ P < 0.05.
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Fig. 3.
Gel electrophoresis for detection of HC DNA fragmentation. DNA gel
electrophoresis was performed as described in
MATERIALS AND METHODS. TNF- (25 ng/ml) plus either SOD (400 U/ml), DMSO (0.5%), NAC (7.5 mM), or GSH
(7.5 mM), but not TNF- alone or TNF- plus CAT (2,000 U/ml), induced the typical "ladder" pattern of DNA fragmentation.
L-NMMA (0.5 mM) prevented
TNF- plus either SOD, DMSO, or GSH-mediated DNA fragmentation.
Lanes 1-10 contained medium,
TNF- alone, TNF- + SOD, TNF- + CAT, TNF- + DMSO, TNF- + NAC, TNF- + GSH, TNF- + SOD + L-NMMA, TNF- + DMSO + L-NMMA, and TNF- + GSH + L-NMMA, respectively.
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Antioxidants and L-NMMA attenuate
HC damage induced by TNF-
.
As shown in Fig. 4, HC damage directly
induced by TNF-
was evident as elevations of both AST and LDH
activities, represented as the percentage of total releasable enzyme
after 12- and 24-h incubation. The addition of antioxidants, SOD, DMSO,
and GSH significantly reduced TNF-
-mediated hepatocellular enzyme
release (P < 0.05 vs. TNF-
alone). The combination of
L-NMMA and the antioxidants resulted in further reductions in AST and LDH release
(P < 0.05 vs. TNF-
plus the
antioxidant).

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Fig. 4.
Attenuation of HC damage by antioxidants and
L-NMMA as represented by
hepatocellular enzyme release. Aspartate aminotransferase (AST) and
lactate dehydrogenase (LDH) activities, represented as a percentage of
the total releasable enzyme, were measured as described in
MATERIALS AND METHODS after incubation
with TNF- (25 ng/ml), SOD (400 U/ml), DMSO (0.5%), GSH (7.5 mM),
and L-NMMA (0.5 mM) in different
combinations at 37°C in 5%
CO2 for 12 and 24 h. Data are
means ± SD from 3 separate experiments. Statistical significances
were compared with TNF- alone
(* P < 0.05) or with TNF-
plus antioxidant (SOD, DMSO, or GSH)
(@ P < 0.05).
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The combinations of antioxidants and
L-NMMA were also found to
protect against HC damage induced by LPS (data not shown).
The effect of LPS and TNF-
, with or without
antioxidants, on ROI generation, NO production, and peroxynitrite
formation in HC.
Exposure of HC to either LPS or TNF-
for 6 h led to a significant
increase in ROI generation in HC, whereas the antioxidants SOD, DMSO,
or GSH significantly inhibited LPS- and TNF-
-mediated ROI generation
(P < 0.05 vs. LPS or TNF-
alone)
(Fig. 5).

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Fig. 5.
Inhibitory effect of antioxidants on either TNF- - or LPS-mediated
reactive oxygen intermediate (ROI) generation in HC. ROI generation was
assessed as described in MATERIALS AND
METHODS after HC were incubated with TNF- (25 ng/ml), LPS (0.1 µg/ml), SOD (400 U/ml), DMSO (0.5%), and GSH (7.5 mM) at 37°C in 5% CO2 for 6 h. Data are expressed as mean channel fluorescence (MCF) for each cell
(MCF/cell). Results are means ± SD, representative of 4 separate
experiments. Statistical significances were compared with medium
(* P < 0.05) and with TNF-
alone or LPS alone (@ P < 0.05).
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In contrast, TNF-
in combination with either SOD, DMSO, NAC, or GSH
resulted in a significant increase in NO level
(P < 0.05 vs. TNF-
alone) (Fig.
6A),
whereas TNF-
alone at different doses failed to augment NO
production in HC (Table 1). Similar results were also found in HC treated with LPS alone (Table 1) or in combination with these antioxidants (Fig.
6B).

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Fig. 6.
Augmentation of NO production in HC after treatment with either TNF-
(A) or LPS
(B) in presence of antioxidants. NO
production was measured after 24-h incubation with TNF- (25 ng/ml),
LPS (0.1 µg/ml), SOD (400 U/ml), CAT (2,000 U/ml), DMSO (0.5%), NAC
(7.5 mM), and GSH (7.5 mM) at 37°C in 5%
CO2 as described in
MATERIALS AND METHODS. Data are means ± SD, representative of 4 separate experiments. Each experiment was
carried out in triplicate. * Statistically significant difference
vs. TNF- alone or LPS alone (P < 0.05).
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The oxidation of dihydrorhodamine-123 by peroxynitrite has been
successfully utilized for the detection of peroxynitrite formation in
vitro (34, 64) and in vivo (55). The method is sensitive and specific,
since neither NO nor superoxide causes dihydrorhodamine-123 oxidation
(34, 55, 64). As shown in Fig. 7, TNF-
significantly increased peroxynitrite formation in HC. The antioxidants
SOD, DMSO, NAC, and GSH attenuated TNF-
-mediated peroxynitrite
formation. Similar results were found in HC treated with LPS alone or
in combination with antioxidants (data not shown).

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Fig. 7.
Attenuation of TNF- -mediated peroxynitrite formation in HC by
antioxidants. Peroxynitrite formation was detected after HC were
treated with TNF- (25 ng/ml), SOD (400 U/ml), CAT (2,000 U/ml), DMSO
(0.5%), NAC (7.5 mM), or GSH (7.5 mM) for 12 h at 37°C in 5%
CO2 as described in
MATERIALS AND METHODS. Data are
expressed as MCF/cell. Results are means ± SD, representative of 3 separate experiments. * Statistically significant difference vs.
TNF- alone (P < 0.05).
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The NO donor SNP induces HC apoptosis and cell damage.
The NO donor SNP was used in this study to further elucidate the effect
of NO on HC injury. Exposure of HC to 0.5 mM SNP for 24 h led to HC
apoptosis, as evidenced by flow cytometry and ELISA, whereas the
antioxidants SOD, DMSO, and GSH augmented SNP-induced HC apoptosis and
DNA fragmentation (P < 0.05 vs. SNP alone) (Fig. 8). In contrast, the addition of
antioxidants significantly reduced SNP-mediated HC damage as
represented by hepatocellular enzyme AST and LDH release
(P < 0.05 vs. SNP alone) (Fig.
9).

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Fig. 8.
Induction of HC apoptosis (A) and
DNA fragments (B) after exposure to
SNP. Isolated HC were incubated either with SNP alone (0.5 mM) or in
combination with SOD (400 U/ml), DMSO (0.5%), and GSH (7.5 mM) at
37°C in 5% CO2 for 24 h. HC
apoptosis and DNA fragments were assessed as described in
MATERIALS AND METHODS. Data are means ± SD, representative of 4 separate experiments.
* P < 0.05 vs. medium;
@ P < 0.05 vs. SNP alone.
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Fig. 9.
Induction of HC damage by SNP as represented by hepatocellular enzyme
release. AST (A) and LDH
(B) activities, represented as % of
total releasable enzyme, were measured as described in
MATERIALS AND METHODS after incubation
with SNP, either alone (0.5 mM) or in combination with SOD (400 U/ml),
DMSO (0.5%), or GSH (7.5 mM) at 37°C in 5%
CO2 for 12 h. Data are means ± SE from 3 separate experiments. Statistical significances were compared
with medium as control (* P < 0.05) or with SNP
alone (@ P < 0.05).
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DISCUSSION |
The abnormal or excessive interactions of the proinflammatory
mediators, such as exogenous LPS and endogenous TNF-
, with neutrophils and Kupffer cells are thought to play an important role in
hepatic dysfunction. The extent of HC damage and cell death, presumably
resulting from either necrosis or apoptosis, determines the degree of
acute hepatic failure. Evidence from our laboratory has demonstrated
that LPS and TNF-
, apart from their indirect effect on HC injury
through neutrophil and Kupffer cell activation, can also directly
result in HC damage and cell death, and that the form of cell death is
necrosis rather than apoptosis (57). Furthermore, the induction of HC
apoptosis has been shown to be responsible for the occurrence of
hepatic injury and to precede hepatic failure in experimental murine
shock models (39). However, the precise mechanisms involved in the
induction of HC apoptosis during SIRS remain to be elucidated. The
major finding in this in vitro study is that NO is involved in HC
injury, primarily through its role in the induction of HC apoptosis.
Since the discovery that murine macrophages could produce NO when
stimulated with interferon-
(IFN-
) and LPS (54) and the
subsequent observation that
L-arginine serves as the
substrate for macrophage NO production (29), a number of other cell
types, including endothelial cells (46), neutrophils (49), vascular smooth muscle cells (12), myocardial cells (21), Kupffer cells (5), and
HC (15, 22), have been shown to produce NO. It has been suggested that
different cell types may vary not only in the timing and quantity of NO
production but also in the signals that stimulate the target cell to
produce NO (5, 15, 22, 46, 48). In rat HC, Curran et al. (15, 16) and
Geller et al. (22) have shown that no single cytokine but rather
combinations of TNF-
, LPS, IFN-
, and interleukin-1
are
required for the induction of HC NO production in vitro, which
correlates with the expression of inducible NO synthase in HC (16, 22).
In contrast, other investigators have found that exposure of rat HC to
either TNF-
alone (45) or LPS alone (48) can induce a significant
increase in NO production and that endotoxin-induced NO production is
partially mediated by HC-derived TNF-
(48). In the present study, we
demonstrate that TNF-
and LPS alone failed to induce NO production
in HC. However, when TNF-
or LPS was combined with different
antioxidants such as SOD, DMSO, NAC, or GSH, NO production was
significantly augmented. The possible mechanism for this phenomenon is
that ROI, particularly superoxide anion, and NO can react rapidly to
form peroxynitrite anion through the near-diffusion-limited reaction,
which reduces the level of biologically active NO (8, 20). Therefore,
it is possible that TNF-
or LPS-mediated ROI generation lowered the
level of NO, possibly through peroxynitrite formation, as both of these oxygen species can be produced in HC by TNF-
or LPS. Moreover, antioxidants scavenging ROI inhibited peroxynitrite formation and thus
increased the NO level in TNF-
or LPS-stimulated HC cultures. This
finding is supported by the observation that oxygen radical scavengers
were found to elevate the NO level in interleukin-1-treated chondrocyte
cultures (8).
There are two distinct mechanisms involved in eukaryotic cell death,
apoptosis and necrosis, which can be distinguished by morphological and
biochemical criteria (13, 51). Apoptosis, characterized by cell
shrinkage, nuclear condensation, and DNA fragmentation, is a form of
programmed cell death that is genetically controlled and regulated by
signal transduction-coupled events. Apoptosis plays an important role
in controlled deletion of cells during metamorphosis, differentiation,
and normal cell turnover. However, recent accumulating evidence has
expanded the definition of apoptosis, and it is now recognized that
apoptosis is not always physiological (32). In endothelial cells, the
induction of apoptosis has been implicated in the development of
increased vascular permeability as a consequence of the loss of barrier
function, which may lead to significant organ dysfunction (1, 11, 58,
59). In different murine models of acute inflammatory liver failure,
hepatic damage induced in
D-galactosamine-sensitized mice
by endotoxin infection was found to be initiated by processes typical
of HC apoptosis (39).
Among the inducers of cell apoptosis and the mechanisms that permit the
cell to die in this stereotypical fashion, both ROI and NO have been
implicated. In the present study, stimulation with TNF-
and LPS
failed to induce HC apoptosis. However, attenuation of ROI with
antioxidants such as SOD, DMSO, NAC, or GSH resulted in the induction
of apoptosis by either TNF-
or LPS, which correlated with an
elevation in NO production, indicating the involvement of endogenous NO
in HC apoptosis. This notion was further confirmed in experiments in
which the addition of the NO synthase inhibitor L-NMMA prevented apoptosis and
DNA fragmentation in TNF-
- or LPS-treated HC in the presence of
antioxidants. NO also appeared to cause HC damage, as demonstrated by
evidence that L-NMMA resulted in further
reductions in TNF-
-mediated hepatocellular AST and LDH release. To
further confirm these novel findings, we therefore used the NO donor
SNP to examine the effect of exogenous NO on HC cell death. We found
that SNP at 0.5 mM was responsible for HC apoptosis and hepatocellular
enzyme release. The augmentation of SNP-induced HC apoptosis and DNA
fragmentation by antioxidants indicates that NO is a primary inducer of
HC apoptosis, as peroxynitrite formation through ROI and NO is blocked
by antioxidants. On the other hand, the reduction of SNP-induced
hepatocellular enzyme release by antioxidants indicates that HC damage
is possibly mediated by NO through peroxynitrite formation.
NO has been implicated in a variety of diverse cellular functions and
biological responses. Although much of the interest in NO has focused
on its role as a signaling or effector molecule in the cardiovascular,
nervous, and immune systems (18, 28, 35), whether NO exerts a
detrimental or protective effect on SIRS is still controversial.
Consistent with our findings, Billiar et al. (5), Curran et al. (15),
and Stadler et al. (53) have shown that NO biosynthesis in HC in vitro
is associated with the suppression of hepatic protein synthesis and the
inhibition of mitochondrial function. Other investigators also reported
that addition of the NO synthase inhibitor
L-NMMA to HC was responsible for
reversal of endotoxin-induced inhibition of cell growth, protein synthesis, and mitochondrial function (36, 37). Furthermore, NO exerts
cytotoxic properties by participating in Kupffer cell and
neutrophil-mediated HC injury (6, 42). Interestingly, in vivo studies
suggest that NO may protect against hepatic damage during endotoxemia
(7, 26, 27). In contrast, data from Laskin et al. (37) and Ma et al.
(40) indicate that NO may contribute to hepatic injury with structural
alterations after acute endotoxemia or hepatic
ischemia-reperfusion in rats. In addition to the role of NO in
hepatic dysfunction during SIRS, an expanding body of literature has
revealed that overproduction of NO may be involved in the
pathophysiology of endotoxemic and septic shock, as well as
ischemia-reperfusion injury, which is associated with tissue
injury and eventual organ failure (30, 33, 49, 56).
ROI, a series of potent oxidants, have been shown to be responsible for
hepatic damage either through a direct effect or through effector
cell-mediated HC injury (4, 17, 60). More recently, ROI have been
implicated as potential mediators in the induction of apoptosis in
different cell types (1, 11, 59, 61). In the present study we found
that exposure of HC to either TNF-
or LPS led to ROI generation with
an elevation in hepatocellular enzyme release. This result is
consistent with other observations in which LPS or TNF-
incubated
with HC induced increased superoxide anion release (60), diminished
intracellular antioxidant levels (2, 47, 60), and accumulated lipid
peroxidation (47, 60). However, in this study TNF-
- or LPS-mediated
ROI generation did not induce HC apoptosis, and ROI seemed to lower the
level of biologically active NO through the pathway of peroxynitrite formation.
In summary, we present two novel findings.
1) NO is the potential inducer of HC
apoptosis. Endogenously derived NO generated by the proinflammatory
mediators TNF-
or LPS in the presence of antioxidants or exogenous
NO supplied by the NO donor SNP induces typical apoptotic changes in
HC. Pharmacological inhibition of NO production by the NO synthase
inhibitor L-NMMA ameliorates HC
apoptosis. 2) TNF-
- or
LPS-stimulated ROI generation is responsible for HC damage and appears
to prevent endogenously produced NO from inducing apoptosis, possibly
through peroxynitrite formation. Further studies are required to
elucidate the possible mechanisms by which NO leads to HC apoptosis.
 |
ACKNOWLEDGEMENTS |
This work was supported by a grant from The Royal College of
Surgeons in Ireland.
 |
FOOTNOTES |
This study was presented in preliminary form at the 16th Annual Meeting
of the Surgical Infection Society, 24 April 1996, Milwaukee, WI.
Present address and address for reprint requests: J. H. Wang, Surgical
Research Laboratory, Clinical Sciences Bldg., Cork Univ. Hospital,
Wilton, Cork, Ireland.
Received 11 November 1997; accepted in final form 19 June 1998.
 |
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