Department of Anatomy, Cell Biology, and Injury Sciences, University of Medicine and Dentistry of New Jersey, New Jersey Medical School, Newark, New Jersey 07103
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ABSTRACT |
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The present study investigated the effect of lipopolysaccharide (LPS; from Escherichia coli, 2 mg/kg body wt ip) on selected aspects of the antioxidant status in Kupffer and sinusoidal endothelial cells. Cells were isolated 18 h after the injection of saline or LPS. In fresh suspension cultures, cellular reduced glutathione (GSH) and H2O2 were determined by monochlorobimane, and 2',7'-dichlorofluorescein diacetate, respectively, using a fluorescence plate reader. LPS injection increased GSH content two- to threefold in Kupffer cells compared with cells from control rats. Cellular GSH content was higher in endothelial than Kupffer cells. However, LPS did not increase GSH content in endothelial cells. Addition of H2O2 (40-200 µM) to Kupffer or endothelial cells caused a transient decrease in GSH, which was more pronounced in cells from control rats (~45% drop) than in LPS-exposed cells (~25% drop). Depleted GSH levels were accompanied by a proportional increase in cellular H2O2. After inhibition of catalase by 3-amino-1,2,4-triazole, the presence of 0.2 mM H2O2 depleted GSH content by 75% and 40% in Kupffer cells from saline- or LPS-injected rats, respectively. The same treatments caused a similar 50% decrease in both activated and control endothelial cells. LPS decreased catalase activity by 45% in Kupffer cells, whereas it had no effect on catalase in endothelial cells. Glutathione reductase activity was not altered by LPS in either cell type. These data show that in activated Kupffer cells the elevated level of cellular glutathione plays an augmented role in the protection against reactive oxygen species, whereas the contribution of catalase to H2O2 detoxification is attenuated. In LPS-stimulated endothelial and Kupffer cells, the efficient maintenance of GSH is consistent with upregulated production of reducing power through the hexose phosphate shunt observed previously.
aminotriazol; antioxidants; glutathione reductase; endothelium; glucose-6-phosphate dehydrogenase; glutathione peroxidase; hepatocytes; hexose monophosphate shunt; inflammation; liver; lipopolysaccharide; macrophages; reactive oxygen species; superoxide anion
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INTRODUCTION |
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OXIDATIVE STRESS in endothelial, parenchymal, or Kupffer cells has several potential implications in the development of liver dysfunction during endotoxemia and sepsis. The pathophysiological consequences of the intra- or extracellular oxidative stress rendered on these cells depend on the functional balance of the pro- and antioxidant pathways (14, 16, 21). On the pro-oxidant side are the production and release of reactive oxygen species (ROS) from activated phagocytes (1, 15). These radicals target the phagocyte itself or, on their release, they may attack neighboring cells. Production of ROS by dysfunctional mitochondria or by xanthine oxidase may also contribute to lipopolysaccharide (LPS)-induced oxidative stress within these cells (11, 26, 42). On the antioxidant side are the cellular, enzymatic, and nonenzymatic defense systems that provide a powerful mean for the elimination of ROS (13, 14, 16, 21). Inflammatory stimuli may modulate both pro- and antioxidant pathways. Maintenance of the balance of these pathways within and among the cells of the hepatic microenvironment is required for normal hepatic function. Whereas the response of liver parenchymal cells to oxidative challenge has been well studied (14, 16, 21), the protective mechanisms against oxidative stress are only partially elucidated in hepatic sinusoidal cells (34, 39, 41, 44).
Previous studies demonstrated that glucose metabolism through the hexose monophosphate shunt (HMS) is potentially an important supporting pathway for either production or elimination of ROS in hepatic sinusoidal cells (35, 38). NADPH generated in the HMS is the primary reducing equivalent for the synthesis of superoxide anions or nitric oxide (24, 28, 31). After the dismutation of these compounds, the elimination of H2O2 is also dependent on HMS activity, as NADPH is required for the functions of glutathione peroxidase (via glutathione) and for the stability of catalase (9, 25, 32).
We showed previously that LPS in vivo caused elevated gene expression of glucose transporter GLUT-1, glucose-6-phosphate (G-6-P) dehydrogenase, superoxide dismutases, and selenium-dependent glutathione peroxidase (Se-GPX) in sinusoidal endothelial cells (34). This condition was accompanied by an elevated capacity of these cells to detoxify H2O2 (39). However, in Kupffer cells, although LPS increased cellular expression of GLUT-1 and G-6-P dehydrogenase, it failed to stimulate expression of superoxide dismutases or Se-GPX (34). Although glutathione peroxidase activity is high in Kupffer cells, it is unknown whether LPS stimulates alternative mechanisms that protect against self-inflicted oxidative stress in activated cells.
The significance of the glutathione redox cycle and catalase in the protection against toxic oxygen compounds is well known (9, 14, 16, 21, 32). Reduced glutathione (GSH) is consumed by peroxidases, and it also protects protein sulfhydryl groups against oxidation. Catalase is ubiquitous and plays a significant role in the elimination of H2O2, especially under conditions where H2O2 reaches high intracellular concentrations. Because our previous studies suggested that endotoxemia upregulates the HMS-dependent ROS-metabolizing pathways in a cell-specific fashion (34, 36, 39), in the present study we tested whether LPS in vivo causes alterations in H2O2 detoxification via glutathione and catalase in hepatic sinusoidal cells. Freshly isolated Kupffer and endothelial cells were also challenged in vitro to assess the functional status of antioxidant pathways.
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MATERIALS AND METHODS |
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Animals and LPS administration. Male Sprague-Dawley rats (300-340 g, Charles River, Wilmington, MA) were used in the study. LPS Escherichia coli 0111:B4 (Difco Laboratories, Detroit, MI) was dissolved in 0.3 ml 0.9% sterile, pyrogen-free saline and injected intraperitoneally (2 mg/kg body wt). Control animals were injected with the same volume of saline. Before the injection of LPS, animals were fasted overnight with free access to water. Cell isolation was initiated 18 h after the injection of LPS in anesthetized animals (ketamine and xylazine, 90 and 9 mg/kg body wt im, respectively). Because the cell isolation procedure required about 4 h, freshly isolated cells were analyzed 22 h after the injection of LPS or saline. The experiments were performed in accordance with the National Institutes of Health "Guide for the Care and Use of Laboratory Animals" [Publication no. (NIH) 85-23].
Cell preparation. Hepatic nonparenchymal cells were isolated with a modified version of the collagenase perfusion method, followed by subsequent Pronase digestion as described in detail previously (18, 36). Sinusoidal endothelial and Kupffer cells were separated by centrifugal elutriation (J2-21 centrifuge with a J6B elutriator rotor; Beckman, Columbia, MD). At constant 2,500 rpm, endothelial and Kupffer cells were eluted at 20 and 60 ml/min flow rates, respectively. The Kupffer cell fraction was subjected to gradient centrifugation (Histopaque 1077) to separate cells from contaminating neutrophils and parenchymal cells. Viability of isolated endothelial and Kupffer cells, as assessed by trypan blue exclusion, was greater than 95%. Purity of endothelial and Kupffer cell preparations was greater than 98% and 94%, respectively, as determined by morphology (Wright-Giemsa stain) or peroxidase stain.
Measurement of fluorescence and in vitro treatments. Freshly isolated endothelial and Kupffer cells were diluted to 4 × 106 and 2 × 106 cells/ml, respectively, in N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid (HEPES) bicarbonate buffer containing 1% albumin, 5 mM glucose, 0.5 mM glutamine, 0.5 mM arginine, 1 mM lactate, and 0.1 mM pyruvate at room temperature in Falcon 24-well cell culture plates (Becton- Dickinson Labware, Lincoln Park, NJ).
Cellular GSH was measured using 50 µM monochlorobimane (8) (MCLB; Molecular Probes, Eugene, OR). After 20-min preincubation in HEPES buffer, MCLB was added and cell-associated fluorescence was measured at 2-min intervals using 360 nm excitation and 460 nm emission filters by an FL500 Microplate Fluorescence Reader (Bio-Tek Instruments, Winooski, VT) at room temperature. MCLB fluorescence approached a plateau after 6-8 min of incubation. When cells were stimulated with freshly prepared H2O2, the oxidant was added 10 min before addition of MCLB unless otherwise noted. Parallel incubations in the absence of cells did not show notable changes in MCLB fluorescence in the presence of H2O2. In parallel incubations with MCLB determinations, cellular H2O2 was measured using 2',7'-dichlorofluorescein diacetate (4) (DCF; Molecular Probes). Cells were preincubated with HEPES buffer for 20 min, followed by incubation with 20 µM DCF or vehicle for 15 min. Cell-associated fluorescence was determined 10 min after the addition of the dye using 485 nm excitation and 530 nm emission filters. Parallel incubations in the absence of cells showed that H2O2 caused a measurable increase in fluorescence presumably by direct interactions between the dye and H2O2. Cell-associated fluorescence was corrected for cell-independent fluorescence, which was <15% of total in the presence of the highest H2O2 concentration used in the experiments. H2O2 treatment at the employed concentration and duration did not affect cell viability as determined by trypan blue exclusion. To inhibit catalase activity cells were preincubated for 15 min in the presence of 2 mM 3-amino-1,2,4-triazole (ATZ), a specific catalase inhibitor (6) or with vehicle. After ATZ treatment MCLB and DCF fluorescence was measured as previously described. All chemicals were purchased from Sigma Chemical unless otherwise indicated.Glutathione and enzyme assays. Freshly isolated cells were incubated for 20 min in HEPES buffer containing the nutrients described, and cellular glutathione and glutathione disulfide were determined using glutathione reductase and 2-vinylpyridine as described previously (45). In a separate series of experiments, freshly isolated hepatic cells were suspended in 1 ml of 50 mM tris(hydroxymethyl)aminomethane, 100 mM KCl, and 0.02% Triton X-100, pH 8.3. The buffer also contained the following proteinase inhibitors: 1 µg/ml leupeptin, 2 µg/ml antipain, 10 µg/ml benzamidine, 10 KU/ml aprotinin, 1 µg/ml chymostatin, and 1 µg/ml pepstatin. The suspension was sonicated (Sonic Disseminator 50; Fisher Scientific, Pittsburgh, PA) for 15 s at 50% energy output and at 4°C and then subjected to centrifugation at 14,000 g, 4°C, for 45 min. In samples of the supernatants the activity of catalase and glutathione reductase was measured by the spectrophotometric method (33) and through using 5,5'-dithio-bis(2-nitrobenzoic acid), respectively (45).
Statistical analysis. Pairwise comparisons were analyzed by Student's t-test. Multiple comparisons were made by analysis of variance, followed by Tukey-Kramer test using a Statistical Analysis System program (SAS Institute). A P value < 0.05 was considered to be significant.
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RESULTS |
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In the first series of experiments we investigated the effect of LPS in vivo on the GSH levels in hepatic cells using the bimane method. The accompanying level of cellular H2O2 was also determined in parallel cell incubations (Fig. 1). Basal glutathione concentration was about threefold greater in endothelial than in Kupffer cells from control rats. LPS in vivo caused a two- to threefold increase in glutathione content in Kupffer cells, whereas it had no statistically significant increase in hepatic endothelial cells. The inset in Fig. 1 shows that the same changes were observed when a conventional assay was used for the determination of cellular glutathione (45). With use of the conventional assay we found that 96-98% of total glutathione was in the reduced form in nonstimulated cells. Furthermore, the baseline concentrations of glutathione were similar in endothelial and parenchymal cells (data not shown). Basal cellular H2O2 concentrations were also elevated in sinusoidal cells after LPS treatment. The increase was more marked in activated Kupffer cells than in endothelial cells (Fig. 1B).
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In the next series of experiments we tested the effect of H2O2 administration in vitro on the cellular contents of GSH and H2O2. Figure 2 depicts the time-dependent changes in GSH content in response to 0.2 mM H2O2 in Kupffer and endothelial cells. H2O2 administration caused a transient drop in GSH levels, which was more pronounced in cells from saline- than from LPS-treated animals. GSH content returned to normal levels faster in cells exposed to LPS in vivo than in cells from control rats (Fig. 2). Because cellular GSH reached the lowest concentration ~10 min after H2O2 challenge, GSH determinations were performed 10 min after H2O2 administration in subsequent experiments.
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Figure 3 compares the response of cellular GSH and H2O2 to exogenously administered H2O2 in resting and LPS-activated Kupffer cells. H2O2 challenge resulted in a concentration-dependent decrease in GSH (Fig. 3A) in both resting and activated cells. However, in LPS-treated cells, GSH levels remained markedly higher even in the presence of 1 mM extracellular H2O2 compared with nonstimulated cells in the absence of H2O2. As expected, the dose-dependent decrease in cellular GSH was accompanied by a dose-dependent increase in cellular H2O2 in these cells (Fig. 3B).
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To assess the contribution of catalase in the maintenance of cellular GSH and H2O2 detoxification, cells were treated with aminotriazol, a specific catalase inhibitor, before the in vitro challenges (Fig. 3). In the absence of H2O2, aminotriazol treatment caused a 30% decrease in cellular GSH in resting Kupffer cells, whereas it had no effect on LPS-treated cells (Fig. 3C). Combined treatment by aminotriazol and H2O2 caused a more pronounced decrease in GSH in resting than in LPS-treated Kupffer cells. (Fig. 3C).
After these treatments the changes in cellular contents of H2O2 and glutathione showed a proportional and inverse relationship (Fig. 3, B and D). After aminotriazol treatment the percent increase in H2O2 levels was less marked in LPS-stimulated than in resting Kupffer cells (Fig. 3, B and D).
The same series of experiments was also performed on hepatic endothelial cells (Fig. 4). Addition of H2O2 in vitro resulted in a dose-dependent decrease in cellular GSH with an inverse increase in cellular H2O2. The relative decrease in intracellular GSH or increase in H2O2, however, was very similar in LPS-exposed and nonstimulated endothelial cells (Fig. 4). Aminotriazol, in the absence of exogenously administered H2O2, did not markedly alter cellular GSH or H2O2 (Fig. 4C). Furthermore, combined aminotriazol and H2O2 treatments caused similar percent changes in cellular GSH and H2O2 in endothelial cells from LPS- or saline-injected rats (Fig. 4, C and D).
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Figure 5 shows that the specific activity of catalase was at least an order of magnitude higher in parenchymal than in nonparenchymal cells (please note difference in scale of the ordinate in Fig. 5A). Basal catalase activity in endothelial cells was approximately one-half of that in Kupffer cells. LPS in vivo decreased catalase activity in Kupffer cells by ~45% (Fig. 5A), whereas it had no effect on catalase in endothelial cells. LPS also decreased catalase activity in parenchymal cells. The activity of glutathione reductase was very similar in parenchymal, Kupffer, and endothelial cells. LPS treatment did not alter glutathione reductase activity in these cells (Fig. 5B).
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DISCUSSION |
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The present study demonstrates that elevated cellular glutathione and an increased capacity to maintain glutathione in the reduced state play an important role in LPS-stimulated macrophages and endothelial cells in the detoxification of H2O2 and thus in the protection against oxidative stress. Catalase contributes to the detoxification of peroxides in these cells; however, its contribution is attenuated in activated macrophages. These findings are consistent with our previous studies indicating elevated supply of reducing power through the HMS in activated hepatic sinusoidal cells (35, 36, 38).
Alterations of glutathione metabolism in phagocytes after phagocytic stimuli are well studied. It has been shown that depletion of glutathione either by oxidation or secretion inhibits phagocytic activity and ROS production by neutrophils or macrophages (5, 30). Our study demonstrates for the first time that baseline GSH content is markedly elevated in hepatic macrophages on exposure to a mild dose of endotoxin in vivo.
The maintenance of glutathione in the reduced state is also more efficient in activated Kupffer cells as reflected by the findings that GSH depletion is less pronounced and recovery of GSH is faster after oxidative challenge. Fast recovery and maintenance of cellular GSH could be the result of elevated activity of catalase and glutathione reductase or of augmented supply of NADPH. Because cellular glutathione reductase activity was not altered by LPS treatment and catalase activity was decreased, it is plausible to suggest that the elevated glucose flux and NADPH supply through the HMS is the underlying supporting mechanism for the fast recovery of GHS levels in Kupffer cells (34-36).
Although LPS did not cause a statistically significant increase in glutathione levels in endothelial cells, the baseline glutathione concentration was markedly higher in endothelial than in Kupffer cells. Furthermore, recovery of GSH after H2O2 challenge was accelerated in LPS-stimulated endothelial cells compared with controls. This, together with the facts that glutathione reductase and catalase activity were not altered, suggests that the increased availability of NADPH produced in the HMS is the underlying mechanism for the faster recovery of GSH in LPS-activated endothelial cells as well (34-36, 38, 39).
Elevated GSH may protect cellular proteins or could also directly interact with ROS generated by activated Kupffer and endothelial cells. Whereas a dominant part of cellular glutathione is cytoplasmic, compartmentalization of glutathione in the mitochondria and around the nucleus and its role in the protection of histones were shown previously in hepatic parenchymal cells (2, 29). A similar role of glutathione can be assumed in hepatic macrophages and endothelial cells. The elevated capacity to maintain GSH in LPS-activated sinusoidal cells can be an important mechanism protecting the functional integrity of these cells during the native immune response or after exposure to toxins (7). Because the dominant portion of cellular glutathione is in the reduced form in these cells, the three- to fourfold elevated GSH measured by the bimane method indicates elevated cellular concentration of total glutathione as well. This suggests that LPS stimulates glutathione synthesis de novo in Kupffer cells (23, 30).
Inhibition of the ubiquitous glutathione reductase is deleterious to cells, as demonstrated by several independent investigations (19, 22, 40). However, no strong evidence indicates that glutathione reductase belongs to the acute phase response proteins in eukaryotic cells. Accordingly, in all the investigated cell types, the activity of glutathione reductase was very similar and was not altered by LPS. These findings indicate that cellular activity of glutathione reductase is adequate to support the glutathione redox cycle under a variety of conditions, and no increase in its activity is required for a more efficient maintenance of cellular glutathione in LPS-activated cells.
The fluorescence method used for the simultaneous determination of cellular glutathione and H2O2 has the advantage of detecting two important determinants of the pro- and antioxidative balance. The inverse relationship between the two was consistent through the study. Namely, after the in vitro challenges, the more cellular H2O2 increased, the more GSH decreased. However, assuming that the H2O2-eliminating pathways were upregulated in LPS-stimulated cells, it could be expected that the same H2O2 challenge would cause a lower increase in cellular H2O2 content in LPS-activated than in nonstimulated cells. On the contrary, our findings showed higher H2O2 levels in LPS-stimulated cells after H2O2 challenge, especially in Kupffer cells. This can be the result of elevated basal cellular levels of H2O2 in LPS-exposed cells and/or the stimulation of superoxide and H2O2 production by H2O2 itself, consistent with previous observations (3, 4, 26). The latter suggestion is further supported by the fact that H2O2 administration increased ethidium bromide fluorescence, a superoxide indicator dye, in endothelial and Kupffer cells (data not shown).
Because of the differences between peroxidases and catalase in their Michaelis constant to H2O2, their contribution to H2O2 detoxification is also different (43). It has been proposed that glutathione peroxidases are responsible for the detoxification of H2O2 present at low concentration, whereas catalase comes into play when the glutathione peroxidase pathway is reaching saturation with the substrate. Previously, we showed an alleviated increase in cellular DCF fluorescence after H2O2 challenge in LPS-treated endothelial cells (39), which was not apparent in the present study. The explanation for this difference is the following. To assess the functional alterations in glutathione status and catalase we used a shorter incubation period (10 min) and higher concentrations of H2O2 (0.08-1 mM) in the present than in the previous study. Under this condition it is expected that the concentration of administered H2O2 and its immediate cellular effects will determine the cellular content of H2O2 (a quasi first-order kinetic). In our previous study we administered 0.008-0.2 mM H2O2 and determined its intracellular levels 20-40 min after the challenge. Under this condition the extra- and intracellular H2O2 concentrations approach steady state and cellular H2O2 concentration is primarily the function of the activity of H2O2-eliminating pathways. This assumption is supported by the fact that after H2O2 challenge the return of GSH concentration to baseline level required about 20 min in LPS-stimulated cells, whereas its recovery was slower in control cells.
It has been shown that the specific activity of catalase is higher in
parenchymal than in nonparenchymal cells (44). Our present study
confirms this and supports the view that catalase is a major
contributor to peroxide elimination in parenchymal cells. Studies on
red blood cells indicated that
H2O2
inactivates catalase via a suicidal mechanism after repeated
interactions between the enzyme and the substrate (10,
17). We suggest that the decreased catalase activity in
LPS-stimulated Kupffer and parenchymal cells is also the result of
inactivation by its own substrate. This suggestion is supported by the
facts that LPS-activated Kupffer cells contain elevated concentrations
of superoxide anion and
H2O2
(1, 35). Furthermore, catalase is more active in parenchymal cells.
Therefore, in these cells the LPS-induced elevated ROS production and
its elimination by catalase could cause inactivation of the enzyme.
Decreased catalase activity was demonstrated after LPS or tumor
necrosis factor- administration in whole rat liver tissue,
consistent with our present observations (27, 46). These data, however,
do not rule out the possibility that the gene expression or translation rate of catalase is decreased in LPS-exposed Kupffer and parenchymal cells.
Because NADPH protects catalase from inactivation (10, 17) the elevated HMS activity and thus efficient supply of NADPH on oxidative stress may provide some protection of catalase in LPS-stimulated Kupffer and endothelial cells. The significance of upregulated HMS as an antioxidant pathway in activated Kupffer and endothelial cells is further supported by recent findings indicating that G-6-P dehydrogenase-deficient eukaryotic cells become extremely sensitive to oxidative stress (25).
The attenuated catalase activity in activated Kupffer cells can explain that in the absence of exogenous H2O2 ATZ decreased GSH in nonstimulated cells, whereas it had no effect on activated cells (Fig. 3, C and D). Furthermore, decreased catalase activity in activated Kupffer cells is also consistent with the finding that after the combined treatment by ATZ and H2O2 the percent decrease in GSH or increase in H2O2 was less pronounced in LPS-treated than in resting Kupffer cells. In agreement with the unaltered activity of catalase in endothelial cells, the effect of ATZ, or the combined effects of ATZ and H2O2 on bimane and DCF fluorescence were very similar in LPS- and saline-treated cells.
Taken together, our present and previous studies indicate that Kupffer and endothelial cells respond to a sublethal dose of LPS with a characteristic pattern of changes in ROS-eliminating pathways. Endotoxemia upregulates glucose uptake and the HMS by elevated expression of GLUT-1 glucose transporter (34, 37) and G-6-P dehydrogenase (36), providing an elevated supply of NADPH in both cells (35, 38). The subsequent pathways, however, are regulated differently. In LPS-exposed Kupffer cells, NADPH oxidase becomes activated, resulting in elevated production of superoxide anions and thus H2O2 (1, 20, 35). Superoxide dismutases and Se-GPX are not altered (34), whereas catalase activity is decreased (Fig. 5). These changes show that the pro- and antioxidant balance moved toward the pro-oxidant side in activated Kupffer cells. However, the accompanying increase in cellular glutathione, the increased supply of NADPH through the HMS, and the high baseline activity of Se-GPX provide sufficient protection against ROS in these cells. In endothelial cells, LPS stimulates the expression of GLUT-1, G-6-P dehydrogenase, Cu-Zn-superoxide dismutase, and Se-GPX (34, 36, 37), which is accompanied by high cellular levels of glutathione and maintenance of catalase activity. This pattern indicates that the balance of the pro- and antioxidant pathways moved toward elevated ROS-detoxification in activated endothelial cells (39). Because these cells do not display an oxidative burst, the stimulated ROS-detoxifying pathways may represent sufficient protection against ROS, reaching the cells from the extracellular space or generated intracellularly.
This cell-characteristic response may represent the establishment of a new intercellular balance of the pro- and antioxidant pathways in the microenvironment of the hepatic sinusoids. Namely, the upregulated antioxidant status of sinusoidal endothelial cells could compensate for the upregulated pro-oxidant status of Kupffer cells and for accumulation of "pro-oxidant" neutrophils in the hepatic sinusoids. The maintenance of this functional intercellular balance may be an important mechanism in the protection of underlying parenchymal cells, supporting hepatic function during sepsis or endotoxemia.
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ACKNOWLEDGEMENTS |
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These studies were supported by the National Institute of General Medical Sciences Grants GM-48721 and GM-55005.
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FOOTNOTES |
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Address for reprint requests: Z. Spolarics, Univ. of Medicine and Dentistry of New Jersey, Dept. of Anatomy, Cell Biology, and Injury Sciences, 185 South Orange Ave., Newark, NJ 07103.
Received 9 May 1997; accepted in final form 27 August 1997.
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REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Bautista, A. P.,
K. Mészáros,
J. Bojta,
and
J. J. Spitzer.
Superoxide anion generation in the liver during the early stage of endotoxemia in rats.
J. Leukoc. Biol.
48:
123-128,
1990[Abstract].
2.
Bellomo, G.,
M. Vairetti,
L. Stivala,
F. Mirabelli,
P. Richelmi,
and
S. Orrenius.
Demonstration of nuclear compartmentalization of glutathione in hepatocytes.
Proc. Natl. Acad. Sci. USA
89:
4412-4416,
1992[Abstract].
3.
Bradley, J. R.,
D. R. Johnson,
and
J. S. Pober.
Endothelial activation by hydrogen peroxide. Selective increases of intercellular adhesion molecule-1 and major histocompatibility complex class I.
Am. J. Pathol.
142:
1598-1609,
1993[Abstract].
4.
Carter, W. O.,
P. K. Narayanan,
and
J. P. Robinson.
Intracellular hydrogen peroxide and superoxide anion detection in endothelial cells.
J. Leukoc. Biol.
55:
253-258,
1994[Abstract].
5.
Cohen, H. J.,
E. H. Tape,
J. Novak,
M. E. Chovaniek,
P. Liegey,
and
J. C. Whitin.
The role of glutathione reductase in maintaining human granulocyte function and sensitivity to exogenous H2O2.
Blood
69:
493-500,
1987[Abstract].
6.
Darr, D.,
and
I. Fridovich.
Irreversible inactivation of catalase by 3-amino-1,2,4-triazole.
Biochem. Pharmacol.
35:
36-42,
1986.
7.
DeLeve, L. D.,
X. Wang,
J. F. Kuhlenkamp,
and
N. Kaplowitz.
Toxicity of azathioprine and monocrotaline in murine sinusoidal endothelial cells and hepatocytes: the role of glutathione and relevance to hepatic venoocclusive disease.
Hepatology
23:
589-599,
1996[Medline].
8.
Fernández, J. C.,
and
N. Kaplowitz.
The use of monochlorobimane to determine hepatic GSH levels and synthesis.
Anal. Biochem.
190:
212-219,
1990[Medline].
9.
Gaetani, G. F.,
S. Galiano,
L. Canepa,
A. M. Ferraris,
and
H. N. Kirkman.
Catalase and glutathione peroxidase are equally active in detoxification of hydrogen peroxide in human erythrocytes.
Blood
73:
334-339,
1989[Abstract].
10.
Gaetani, G. F.,
H. N. Kirkman,
R. Mangerini,
and
A. M. Ferraris.
Importance of catalase in the disposal of hydrogen peroxide within human erythrocytes.
Blood
84:
325-330,
1994
11.
Garcia-Ruiz, C.,
A. Colell,
A. Morales,
N. Kaplowitz,
and
J. C. Fernadez-Checa.
Role of oxidative stress generated from the mitochondrial electron transport chain and mitochondrial glutathione status in loss of mitochondrial function and activation of transcription factor nuclear factor-kappa B: studies with isolated mitochondria and rat hepatocytes.
Mol. Pharmacol.
48:
825-834,
1995[Abstract].
12.
Griffith, O. W.
Determination of glutathione and glutathione disulfide using glutathione reductase and 2-vinylpiridine.
Anal. Biochem.
106:
201-212,
1980.
13.
Hamers, M. N.,
and
D. Roos.
Oxidative stress in human neutrophilic granulocytes: host defence and self-defence.
In: Oxidative Stress, edited by H. Sies. New York: Academic, 1985, p. 351-403.
14.
Inoue, M.
Protective mechanism against reactive oxygen species.
In: The Liver, Biology and Pathobiology, edited by I. M. Arias,
J. L. Boyer,
W. B. Jakoby,
N. Fausto,
D. Schacter,
and D. A. Shafritz. New York: Raven, 1994, p. p.443-459.
15.
Jaeschke, H.
Mechanism of oxidant stress-induced acute tissue injury.
Proc. Soc. Exp. Biol. Med.
209:
104-111,
1995[Abstract].
16.
Kaplowitz, N.,
and
H. Tsukamoto.
Oxidative stress and liver disease.
Prog. Liver Dis.
14:
131-159,
1996[Medline].
17.
Kirkman, H. N.,
S. Galiano,
and
G. F. Gaetani.
The function of catalase-bound NADPH.
J. Biol. Chem.
262:
660-666,
1987
18.
Knook, D. L.,
N. Blansjaar,
and
E. C. Sleyster.
Isolation and characterization of Kupffer and endothelial cells from rat liver.
Exp. Cell Res.
109:
317-329,
1977[Medline].
19.
Lopez-Barea, J.,
J. A. Barcena,
J. A. Bocanegra,
J. Florindo,
C. Garcia-Alfonso,
A. Lopez-Ruiz,
E. Martinez-Galisteo,
and
J. Peinado.
Structure, mechanism, functions, and regulatory properties of glutathione reductase.
In: Glutathione: Metabolism and Physiological Functions, edited by J. Vina. Boca Raton, FL: CRC, 1990, p. 105-116.
20.
McCloskey, T. W.,
J. A. Todaro,
and
D. L. Laskin.
Lipopolysaccharide treatment of rats alters antigen expression and oxidative metabolism in hepatic macrophages and endothelial cells.
Hepatology
16:
191-203,
1992[Medline].
21.
McCord, J. M.
Human disease, free radicals, and the oxidant/antioxidant balance.
Clin. Biochem.
26:
351-357,
1993[Medline].
22.
Meister, A.
Glutathione.
In: The Liver, Biology and Pathobiology, edited by I. M. Arias,
J. L. Boyer,
W. B. Jakoby,
N. Fausto,
D. Schacter,
and D. A. Shafritz. New York: Raven, 1994, p. 401-417.
23.
Miura, K.,
T. Ishii,
Y. Sugita,
and
S. Bannai.
Cystine uptake and glutathione level in endothelial cells exposed to oxidative stress.
Am. J. Physiol.
262 (Cell Physiol. 31):
C50-C58,
1992
24.
Moncada, S.,
and
E. A. Higgs.
Endogenous nitric oxide: physiology, pathology and clinical relevance.
Eur. J. Clin. Invest.
21:
361-374,
1991[Medline].
25.
Pandolfi, P. P.,
F. Sonati,
R. Rivi,
P. Mason,
F. Grosveld,
and
L. Luzatto.
Targeted distribution of the housekeeping gene encoding glucose 6-phosphate dehydrogenase (G6PD): G6PD is dispensable for pentose synthesis but essential for defense against oxidative stress.
EMBO J.
14:
5209-5215,
1995[Abstract].
26.
Phan, S. H.,
D. E. Gannon,
J. Varani,
U. S. Ryan,
and
P. A. Ward.
Xanthine oxidase activity in rat pulmonary artery endothelial cells and its alteration by activated neutrophils.
Am. J. Pathol.
134:
1201-1211,
1989[Abstract].
27.
Portoles, M. T.,
M. Catala,
A. Anton,
and
R. Pagini.
Hepatic response to oxidative stress induced by E. coli endotoxin: glutathione as an index of the acute phase during the endotoxic shock.
Mol. Cell. Biochem.
159:
115-121,
1996[Medline].
28.
Robinson, J. M.,
and
J. A. Badwey.
The NADPH oxidase complex of phagocytic leukocytes: a biochemical and cytochemical view.
Histochem. Cell Biol.
103:
163-180,
1995[Medline].
29.
Romero, F. J.,
and
D. Galaris.
Compartmentalization of cellular glutathione in mitochondrial and cytosolic pools.
In: Glutathione: Metabolism and Physiological Functions, edited by J. Vina. Boca Raton, FL: CRC, 1990, p. 29-38.
30.
Rouzer, C. A.,
W. A. Scott,
O. W. Griffith,
A. L. Hamill,
and
Z. A. Cohm.
Glutathione metabolism in resting and phagocytizing peritoneal macrophages.
J. Biol. Chem.
257:
2002-2008,
1982
31.
Shatwell, K. P.,
and
A. W. Segal.
NADPH oxidase.
Int. J. Biochem.
28:
1191-1195,
1996.
32.
Sies, H. Oxidative stress: from basic research to clinical
applications. Am. J. Med.
91, Suppl. 3C: 31S-38S, 1991.
33.
Smith, I. K.,
T. L. Vierheller,
and
C. A. Thorne.
Assay of glutathione reductase in crude tissue homogenates using 5,5'-dithio-bis(2-nitrobenzoic acid).
Anal. Biochem.
175:
408-413,
1988[Medline].
34.
Spolarics, Z.
Endotoxin stimulates the gene expression of reactive oxygen eliminating pathways in rat hepatic endothelial and Kupffer cells.
Am. J. Physiol.
270. (Gastrointest. Liver Physiol. 33):
G660-G666,
1996
35.
Spolarics, Z.,
A. P. Bautista,
and
J. J. Spitzer.
Primed pentose cycle activity supports production and elimination of superoxide anion in Kupffer cells from rats treated with endotoxin in vivo.
Biochim. Biophys. Acta
1179:
134-140,
1993[Medline].
36.
Spolarics, Z.,
and
L. Navarro.
Endotoxin stimulates the expression of glucose-6-phosphate dehydrogenase in Kupffer and hepatic endothelial cells.
J. Leukoc. Biol.
56:
453-457,
1994[Abstract].
37.
Spolarics, Z.,
P. H. Pekala,
G. J. Bagby,
and
J. J. Spitzer.
Brief endotoxemia markedly increases expression of GLUT-1 glucose transporter in Kupffer, hepatic endothelial and parenchymal cells.
Biochem. Biophys. Res. Commun.
193:
1211-1215,
1993[Medline].
38.
Spolarics, Z.,
and
J. J. Spitzer.
Augmented glucose use and pentose cycle activity in hepatic endothelial cells after in vivo endotoxemia.
Hepatology
17:
615-620,
1993[Medline].
39.
Spolarics, Z.,
D. S. Stein,
and
Z. C. Garcia.
Endotoxin stimulates hydrogen peroxide detoxifying activity in rat hepatic endothelial cells.
Hepatology
24:
691-696,
1996[Medline].
40.
Starke, P. E.,
and
J. L. Farber.
Endogenous defense against the cytotoxicity of hydrogen peroxide in cultured rat hepatocytes.
J. Biol. Chem.
260:
86-92,
1985
41.
Steinberg, P.,
H. Schramm,
L. Schladt,
L. W. Robertson,
H. Thomas,
and
F. Oesch.
The distribution, induction and isoenzyme profile of glutathione S-transferase and glutathione peroxidase in isolated rat liver parenchymal, Kupffer and endothelial cells.
Biochem. J.
264:
737-744,
1989[Medline].
42.
Taylor, D. E.,
A. J. Ghio,
and
C. A. Pantadosi.
Reactive oxygen species produced by liver mitochondria of rats in sepsis.
Arch. Biochem. Biophys.
316:
70-76,
1995[Medline].
43.
Toussaint, O.,
A. Houbion,
and
J. Remacle.
Relationship between the critical level of oxidative stresses and the glutathione peroxidase activity.
Toxicology
81:
89-101,
1993[Medline].
44.
Van Berkel, T. J. C.
Difference spectra, catalase- and peroxidase activities of isolated parenchymal and non-parenchymal cells from rat liver.
Biochem. Biophys. Res. Commun.
61:
204-209,
1974[Medline].
45.
Worthington, C. C.
Catalase.
In: Worthington Enzyme Manual: Enzymes and Related Biochemicals, edited by C.C. Worthington. Freehold, NJ: Worthington Biochemical, 1988, p. 72-74.
46.
Yasmineh, W. G.,
J. L. Parkin,
J. I. Caspers,
and
A. Theologides.
Tumor necrosis factor/cachectin decreases catalase activity of rat liver.
Cancer Res.
51:
3990-3995,
1991[Abstract].