* Department of Pharmacology and Toxicology and
Department of Pathobiology and Diagnostic Investigation, National Food Safety and Toxicology Center and Institute for Environmental Toxicology, Michigan State University, East Lansing, Michigan 48824
Received February 18, 2003; accepted April 21, 2003
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ABSTRACT |
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Key Words: liver; inflammation; lipopolysaccharide; monocrotaline; coagulation system; fibrin deposition; heparin; warfarin; MCP-1.
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INTRODUCTION |
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Mild exposure to LPS is commonplace and episodic, varying with the lifestyle and health of an individual. Systemic LPS concentration can be enhanced during bacterial infection and by increased translocation of LPS from indigenous Gram-negative bacteria in the gastrointestinal tract into the portal circulation (Fink and Mythen, 1999; Ganey and Roth, 2001
). A variety of conditions, including disease, dietary alterations, trauma to the gastrointestinal tract, and alcohol consumption, can increase this translocation (reviewed in Roth et al., 1997
). Although such episodic exposure to smaller amounts of LPS is insufficient to cause tissue injury, a modest inflammatory response may result. This response includes the accumulation of neutrophils (polymorphonuclear leukocytes; PMNs) and the production of tumor necrosis factor-
(TNF-
) and other inflammatory mediators that have the potential to alter hepatocellular homeostasis (Michie et al., 1988
; Spitzer and Mayer, 1993
). This may cause tissues to become more susceptible to chemically induced injury. Indeed, LPS exposure augments the toxicity of a number of hepatotoxicants (reviewed in Ganey and Roth, 2001
).
Monocrotaline (MCT) is a pyrrolizidine alkaloid phytotoxin found in numerous plants of the Crotalaria genus worldwide (Mattocks, 1986; Stegelmeier et al., 1999
). It is a well-known hepatotoxicant (Mattocks, 1986
). Humans are exposed to this toxin through the accidental consumption of contaminated foodstuffs and the intentional ingestion of MCT and related pyrrolizidine alkaloids in alternative medicines (e.g., nong ji li, zi xiao rong, comfrey tea, and others; Roeder, 2000
; Stegelmeier et al., 1999
). At hepatotoxic doses of MCT, liver injury is characterized by centrilobular (CL) hepatocellular degeneration and necrosis, hemorrhage, and central venous and sinusoidal endothelial cell (SEC) injury (Copple et al., 2002a
; DeLeve et al., 1999
; Schoental and Head, 1955
; Yee et al., 2000
).
A noninjurious dose of LPS administered to rats in close temporal proximity to a nonhepatotoxic dose of MCT results in synergistic, acute liver injury, which becomes maximal 18 h after MCT administration (Yee et al., 2000). The resulting liver lesions are both CL and MZ, exhibiting characteristics similar to lesions associated with larger, toxic doses of MCT and LPS given separately. The nature of the MCT-like, CL and LPS-like, MZ lesions suggests that each agent enhances the injury of the other (Yee et al., 2000
). Inflammatory components such as Kupffer cells (KCs), TNF-
,, and PMNs are critical to the pathogenesis (Yee et al., 2003a
,b
). SEC injury and coagulation system activation also occur in MCT/LPS-cotreated animals before the onset of hepatic parenchymal cell (HPC) injury. Histopathological examination revealed pronounced congestion, hemorrhage, and fibrin deposition in the hepatic lesions (Yee et al., 2000
, 2003
). These results raise the possibility that the coagulation system has a causal role in hepatocellular necrosis in this model (Yee et al., 2003
).
Activation of the coagulation cascade occurs either intrinsically via surface-mediated reactions or extrinsically through a tissue factor (TF)-derived pathway. At the distal end of the cascade, factor Xa converts prothrombin into active thrombin. Thrombin, in turn, can convert circulating fibrinogen into insoluble fibrin clots (Bloom, 1990; Schultze and Roth, 1998
). It has been postulated that fibrin deposition in the liver leads to local sinusoidal hypoperfusion, which might contribute to HPC injury (Ba et al., 2000
; Copple et al., 2002a
; DeLeve et al., 1996
; Saetre et al., 2000
).
LPS activates the coagulation system primarily through the expression of TF on endothelial cells, monocytes, and PMNs (Bone, 1992; Polack et al., 1997
; Todoroki et al., 2000
). Thrombin has been implicated in the release of proinflammatory cytokines and other factors from macrophages, monocytes, and endothelial cells and is a weak PMN chemoattractant (Bizios et al., 1986
; Hoffman and Cooper, 1995
; Holland et al., 1998
; Moulin et al., 1996
). Accordingly, the coagulation system might be involved in the upregulation of proinflammatory mediators, in addition to forming fibrin clots. Indeed, at acutely toxic doses of LPS, an activated coagulation system is critical for liver pathogenesis, and it appears that interplay between thrombin and inflammatory factors is important (Copple et al., 2003
; Moulin et al., 2001
).
The degree to which the coagulation system has a role in the synergistic injury from the combination of noninjurious doses of MCT and LPS has not been explored. Accordingly, the present study was designed to investigate its role in MCT/LPS-induced liver injury and to explore the interdependence between the coagulation system and inflammatory factors that are critical for the toxic response.
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MATERIALS AND METHODS |
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Animals.
Male Sprague-Dawley rats (Crl:CD (SD)IGS BR, Charles River, Portage, MI) weighing 200300 g were used for all studies. Animals were allowed food (Rodent Chow/Tek 8640, Harlan Teklad, Madison, WI) and water ad libitum. They were housed no more than three to a cage on Aspen chip bedding (Northeastern Products Company, Warrenburg, NY) and were maintained on a 12-h light/dark cycle in a controlled temperature (1821°C) and humidity (55 ± 5%) environment for a period of 1 week before use. All procedures on animals followed the guidelines for humane treatment set by the American Association of Laboratory Animal Sciences and the University Laboratory Animal Research Unit at Michigan State University.
Treatment protocol.
MCT was dissolved in sterile saline minimally acidified by 0.2 M HCl. The pH was brought to 7 by addition of 2 M NaOH, and the volume was adjusted with sterile saline to the appropriate final concentration. Rats were given MCT (100 mg/kg) or an equivalent volume of sterile saline vehicle (Veh), intraperitoneally, followed 4 h later by LPS (7.4 x 106 EU/kg) or saline Veh via tail vein injection. LPS was administered 4 h after MCT to minimize interference with MCT bioactivation (Allen et al., 1972).
Treatment with HEP.
Rats were given HEP (2000 U/kg) or saline Veh intravenously 1.5 h after LPS administration. This treatment has been shown to inactivate the coagulation system and prevent injury from a hepatotoxic dose of LPS (Moulin et al., 1996). Rats were killed and liver injury was assessed 18 h after MCT treatment.
Treatment with WARF.
Rats were pretreated with WARF (7.5 mg/kg) or an equivalent volume of DMSO Veh intraperitoneally 34 and 10 h before MCT administration. This treatment has been shown to inactivate the coagulation system (Copple et al., 2002b). Rats were killed and liver injury was assessed 18 h after MCT treatment.
Assessment of hepatic injury and plasma TNF- and MCP-1 concentrations.
At 6, 12, or 18 h after MCT administration, rats were anesthetized with sodium pentobarbital (50 mg/kg, ip). A midline abdominal incision was made, blood was collected from the inferior vena cava into a syringe containing sodium citrate (0.38% final concentration), and animals were euthanized by exsanguination. HPC injury was evaluated by increases in the activities of ALT and AST in plasma. An ELISA kit was used to measure plasma HA concentration, a marker of hepatic SEC injury. Plasma TNF- and MCP-1 concentrations were determined with a rat TNF-
and a rat MCP-1 ELISA kit, respectively.
Assessment of plasma fibrinogen concentration and CINC-1.
Plasma fibrinogen concentration was evaluated with a BBL fibrometer (Becton, Dickson and Company, Hunt Valley, MD) and a fibrinogen diagnostic kit. Plasma fibrinogen concentration was determined in the HEP and WARF studies with MCT/LPS-cotreated animals, as well as in cotreated animals that underwent KC inactivation or TNF- or PMN depletion. In the KC inactivation study, GdCl3 (10 mg/kg) was administered to rats 24 h before LPS administration. In the TNF-
depletion studies, either PTX (100 mg/kg) or ATS (1 ml/rat) was administered 1 h before LPS administration. In the PMN depletion study, animals were pretreated with NAS 24 (1 ml/rat) and 8 (0.5 ml/rat) hours before LPS administration. These treatment regimens were effective in preventing KC activation, the LPS-induced increase in plasma TNF-
activity and hepatic PMN accumulation, respectively (Yee et al., 2003a
,b
). Administration of GdCl3, PTX, ATS, or NAS did not interfere with MCT bioactivation (Yee et al., 2003a
,b
). Plasma CINC-1 was measured using a rat GRO/CINC-1 ELISA kit.
Histopathologic evaluation and morphometry.
Livers were fixed by immersion in 10% neutral buffered formalin for at least 3 days before being processed for histologic analysis. In addition, a portion (1 cm3) of the liver from the middle of the left lateral lobe was frozen in isopentane immersed in liquid nitrogen for immunohistochemical staining. Serial transverse sections from the left lateral liver lobe were processed for light microscopy. Paraffin-embedded sections were cut at 5 µm, stained with hematoxylin and eosin, and evaluated for lesion size and severity. Slides were coded, randomized, and evaluated by light microscopy.
Digitized color images of hematoxylin and eosin-stained liver sections were visualized with an Olympus AX-80T light microscope (Olympus Corp., Lake Success, NY) interfaced with a high-resolution CCD color camera (OLY-750, Olympus-America, Inc., Melville, NY) to quantify treatment-induced changes in liver morphology. Images were evaluated with Scion Image software (Scion Corporation, Frederick, MD) employing a 64-point lattice grid to determine (1) the total area of liver analyzed, (2) the area of CL lesion, (3) the area of MZ lesion, (4) the area of normal parenchyma, and (5) the area of nonparenchymal space. A lesion was defined as hepatic parenchymal cells with either swollen, eosinophilic cytoplasm and karyolytic or pyknotic nuclei (i.e., oncosis), or cells with shrunken cytoplasm and karyorrhexic nuclei or apoptotic bodies (i.e., apoptosis; Levin et al., 1999; Majno and Joris, 1995
). Nonparenchymal space was defined as nonparenchymal tissue, vessel lumen, and regions outside the perimeter of the liver section. The area of each object (category) of interest (i.e., lesion) was calculated from the following expression (Cruz-Orive, 1982
):
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Distance between points was 55 µm. Accordingly, the area represented by each point was 3025 µm2. One section from the liver of each animal in a treatment group was systematically scanned using adjacent, non-overlapping microscopic fields. The first image field analyzed in each section was chosen using a random number table (i.e., any image field between 1 and 10). Thereafter, every 10th field containing hepatic parenchymal cells was evaluated (minimum of 20 fields measured/section). The measured fields represented approximately 10% of the total area of each liver section. Percent lesion area was estimated based on the following formula:
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Enumeration of hepatic neutrophils.
Paraffin-embedded liver tissue (three serial liver sections per slide) was cut into 6 µm-thick slices. Paraffin was removed from the liver tissues with xylene before staining. PMNs within liver sections were stained with a rabbit anti-PMN Ig isolated from serum of rabbits immunized with rat PMNs as described by Hewett et al.(1992). After incubation with the primary antibody, tissue sections were incubated with biotinylated goat anti-rabbit IgG, avidin-conjugated alkaline phosphatase, and Vector Red substrate to stain PMNs. Hepatic PMN accumulation was assessed by averaging the numbers of PMNs enumerated in 30 randomly selected, high power fields (HPFs; x400) in each slide (i.e., 10 HPFs per liver section). The analyzed fields represented between 5 and 10% of the total area of each liver section. Analyzed fields were selected in an unbiased manner to cover the entire liver section. PMNs were identified by positive staining and cell morphology.
Immunohistochemistry.
Sections of frozen tissue (8 µm-thick) were fixed in 10% buffered formalin containing 2% acetic acid for 30 min at room temperature. This fixation protocol solubilizes all fibrinogen and fibrin species except for cross-linked fibrin. Hence, only cross-linked fibrin is stained in liver sections (Schnitt et al., 1993). Sections were incubated for 30 minutes with PBS containing 10% horse serum (i.e., blocking solution) and then with goat anti-rat fibrinogen (diluted 1:1000) in blocking solution overnight at 4°C. Next, the sections were incubated for 3 h in blocking solution with donkey anti-goat secondary antibody conjugated to Alexa 594 (1:1000). Sections were washed three times for 5 min each with PBS and visualized using fluorescence microscopy. No staining was observed in the controls in which the primary or secondary antibody was omitted from the staining protocol. All treatment groups compared morphologically were stained at the same time and evaluated on the same day.
Quantification of hepatic fibrin deposition.
Fibrin deposition in the liver was quantified by morphometrically analyzing the area of immunohistochemical staining for each liver section (Copple et al., 2002a). Fluorescent staining of liver sections was visualized using an Olympus AX-80T microscope (Olympus, Lake Success, NY). For morphometric analysis of fibrin deposition in a liver section, digital images of 5 randomly chosen x100 fields per tissue section were captured using a SPOT II camera and SPOT Advanced Software (Diagnostic Instruments, Sterling Heights, MI). Samples were coded so that the evaluator was not aware of the treatment, and the same exposure time was used for all captured images. Each digital image encompassed a total area of 1.4 mm2 and contained several CL, MZ, and periportal (PP) regions.
The area of immunohistochemical staining (number of pixels) within the CL, MZ, and PP regions was quantified using Scion Image software (Scion Corporation, Frederick, MD). For fibrin deposition quantification, a density slice from an inverted, gray-scale digital image of a liver section was used for analysis. A density slice allows analysis of pixels in a defined range of gray values (i.e., densities). The threshold was selected so that little positive staining was present in Veh/Veh/Veh-treated controls. The same threshold value was used to analyze digital images from all treatment groups. The area of positive staining was measured and divided by the total area of the image. Analysis of fibrin deposition in the CL and PP areas was conducted by drawing a 145 µm-diameter circle around the central vein or periportal region. The circumference of the circle is about 46 hepatocytes away from the central vein or vessels of the portal triad, and this area was arbitrarily defined as the CL and PP regions, respectively. The MZ region was defined as the center of area between the CL and PP regions using the same circle circumference, without having overlap of these arbitrary circles. The area of the circle was 16,512 µm2. The area of fibrin staining in each region was measured as described above and divided by the total area of the image. Results from the random fields analyzed for each liver section were averaged and counted as a replicate (i.e., each replicate representing a different rat).
Statistical analysis.
Results are expressed as mean ± SEM When variances were not homogeneous, data were log-transformed before analysis. Data expressed as percentages were transformed by arc sine square root prior to analysis. Data for single comparisons were analyzed by Students t-test or, when appropriate, Fishers exact test (Steele et al., 1997). Multiple comparisons of homogeneous data were analyzed by one-way or two-way analysis of variance (ANOVA), as appropriate, and group means were compared using Tukeys omega post hoc test (Steele et al., 1997
). The criterion for significance was p
0.05 for all comparisons.
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RESULTS |
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Plasma fibrinogen concentration was significantly decreased in rats treated with MCT/LPS. Both HEP and WARF prevented the activation of the coagulation system in MCT/LPS-coexposed animals (Figs. 1A and 1B, respectively), confirming their effectiveness.
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Effect of Anticoagulants on MCT/LPS-Induced Liver Lesions
MCT/LPS-cotreated control animals exhibited MCT-like, CL and LPS-like, MZ liver lesions as previously described by Yee et al.(2000). CL lesions consisted of moderate to marked hepatocellular degeneration and apoptotic and oncotic necrosis, hemorrhage, and loss of central vein intima. CL lesions also exhibited a moderate accumulation of PMNs and monocytes. MZ lesions comprised marked and more frequent, well-defined areas of hepatocellular coagulative necrosis accompanied by PMN and mononuclear cell infiltration. Pronounced congestion and hemorrhage were also present in these lesions. Livers from MCT/LPS-coexposed animals given either anticoagulant exhibited qualitatively similar CL and MZ lesions; however, both lesion types were smaller and considerably less frequent. The MZ lesions had a slightly greater reduction in size and frequency than the CL lesions (Table 1
). Significant decreases in the areas of CL (77% decrease) and MZ (87% decrease) lesions were found in the livers of MCT/LPS-cotreated animals that were treated with HEP. Similar decreases were found in CL (62%) and MZ (85%) lesions in livers from MCT/LPS-coexposed animals given WARF. No histologic evidence of injury was observed in animals given Veh or anticoagulant alone.
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DISCUSSION |
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Either HEP (Fig. 1A) or WARF (Fig. 1B
) treatment of MCT/LPS-cotreated animals prevented the activation of the coagulation system, confirming the effectiveness of these agents. Anticoagulant treatment caused a marked decrease in HPC injury but only a modest decrease in SEC injury 18 h after MCT administration (Figs. 2 and 3
). This suggests that the coagulation system only partially contributes to SEC injury and that additional factors are likely involved. Zonal analysis of liver sections revealed that anticoagulant treatment significantly reduced the areas of CL and MZ lesions (Table 1
). Accordingly, the coagulation system appears to contribute causally to MCT/LPS-induced liver injury.
MCT is nontoxic and must be bioactivated by cytochromes of the P450 3A family to its toxic metabolite, monocrotaline pyrrole (MCTP), in order to produce liver injury (Kasahara et al., 1997; Stegelmeier et al., 1999
; White and Mattocks, 1972
). This conversion happens rapidly, with peak MCTP production occurring within 2 h after MCT exposure (Allen et al., 1972
). Thus, HEP was given to MCT/LPS-cotreated animals after the bioactivation of MCT to MCTP was nearly complete (i.e., 5.5 h after MCT). A recent study by Copple et al.(2002b)
demonstrated that WARF does not interfere with MCT bioactivation. Therefore, it is unlikely that either anticoagulant reduced injury by interfering with MCT metabolism.
There are a variety of mechanisms by which the coagulation system might contribute to the liver injury of this model. Fujiwara et al.(1988) and others (Aria et al., 1996
; Ba et al., 2000
; Copple et al., 2002a
; DeLeve et al., 1996
; Saetre et al., 2000
) have postulated that fibrin clots in the liver cause local hypoperfusion and can thereby result in cellular injury. Consistent with this hypothesis, HEP substantially reduced fibrin deposition in CL and MZ regions of liver lobules (Fig. 4B
) from MCT/LPS-cotreated rats. Hence, it is possible that the reduction in hepatic fibrin deposition was responsible for the decrease in liver injury. However, in liver injury caused by a hepatotoxic dose of LPS, thrombin appears to be the critical factor, acting independently of fibrin clot formation (Hewett and Roth, 1995
; Moulin et al., 1996
, 2001
). Thrombin can contribute to liver injury by causing the stimulation and aggregation of platelets (Kito et al., 1985
; Shuman, 1986
; Sinha et al., 1983
; Wise et al., 1980
), by inducing PMN chemotaxis (Bizios et al., 1986
) and/or by enhancing MCP-1 release from liver stellate cells, monocytes, and endothelial cells (Colotta et al., 1994
; Marra et al., 1995
). Moreover, thrombins action on a protease-activated receptor present on KCs and SECs in liver might contribute to HPC injury, as it does after a large, hepatotoxic dose of LPS (Copple et al., 2003
; Moulin et al., 2001
). Indeed, activation of protease-activated receptor-1 by thrombin results in the release of various cytokines and growth factors, as well as the expression of adhesion molecules on endothelium that could contribute to inflammatory liver injury (Derian et al., 2002
). In MCT/LPS-cotreated animals, further study will be needed to define fully the mechanism by which an activated coagulation system promotes hepatotoxicity.
The relationship between the coagulation system and other inflammatory factors was explored. Although neither KC inactivation nor TNF- depletion prevented activation of the coagulation system, PMN depletion did (Table 3
). This suggests that PMNs promote coagulation system activation in this model. PMNs can affect the coagulation system through a number of mechanisms, including the expression of TF on their surfaces (Todoroki et al., 2000
) and the release of cathepsin G (Bray et al., 1987
). TF activates the coagulation system, leading to thrombin formation (Osterud and Rapaport, 1977
; Schultze and Roth, 1998
). In addition to its cytotoxic protease activity (Ho et al., 1996
), cathepsin G activates Factor X, which converts prothrombin to thrombin (Bray et al., 1987
; Goel and Diamond, 2001
; Plescia and Altieri, 1996
). Hence, the critical role of PMNs may derive both from their ability to release cytotoxic mediators and their participation in activating the coagulation system.
Although PMNs appear necessary for activation of the coagulation system in this model, they may not be sufficient. Another necessary factor may be damage to SECs, which is known to activate the coagulation system in other models (Copple et al., 2002a; Hirata et al., 1989
; Seto et al., 1998
). SEC injury occurs concurrently with activation of the coagulation system (Yee et al., 2003
), but further study will be needed to determine whether both PMN accumulation and SEC injury are required events and, if so, how they interact to promote coagulation system activation.
The involvement of TNF- in coagulation system activation varies with different models of inflammation. After a large, hepatotoxic dose of LPS, TNF-
depletion prevents activation of the coagulation system (Hewett and Roth, 1995
). In that model, TNF-
is important for coagulation system activation through either stimulation of TF activity on endothelial cells (Bevilacqua et al., 1986
; Esmon, 2000
; Kirchhofer et al., 1994
) or through activation of PMNs (Goel and Diamond, 2001
; Klebanoff et al., 1986
; Todoroki et al., 2000
). By contrast, in the MCT/LPS-cotreatment model TNF-
does not appear to be necessary for activation of the coagulation system (Table 3
). Likewise, KC inactivation in this model failed to prevent coagulation system activation, a result consistent with the lack of effect of TNF-
depletion, since activated KCs are a major producer of TNF-
in the liver. Although both KCs and TNF-
are critical for the development of MCT/LPS-induced injury (Yee et al., 2003a
), their involvement is apparently not through activation of the coagulation system.
In MCT/LPS-cotreated rats, an early, pronounced increase in plasma TNF- concentration occurs, and this is followed by a more modest but sustained elevation that continues through 18 h (Yee et al., 2003a
,b
). Anticoagulant administration to MCT/LPS-cotreated animals did not affect this later, sustained phase of increased plasma TNF-
concentration (Figs. 5A,B
), suggesting that an activated coagulation system is not needed for sustained TNF-
release. Although it cannot be ruled out that the coagulation system influenced TNF-
formation at an earlier time, these results provide additional evidence that TNF-
generation and coagulation system activation are not interdependent. Previously, ATS treatment was shown to attenuate PMN accumulation at 18 h in livers of MCT/LPS treated rats (Yee et al., 2003b
). The observation that it also reduced plasma CINC-1 concentration (Table 4
) suggests that PMN influx occurs through a TNF-dependent mechanism involving CINC-1.
Anticoagulant administration reduced hepatic PMN accumulation (Figs. 6A,B). The coagulation system could affect PMN influx into liver tissue through the chemotactic activity of thrombin (Bizios et al., 1986
). PMN accumulation, however, was not completely eliminated by administration of anticoagulant, suggesting that thrombin may work in conjunction with other PMN chemoattractants (e.g., CINC-1; Luster et al., 1998
; Zhang et al., 1995
). In this regard, it is of interest that anticoagulant administration reduced but did not eliminate CINC-1 appearance in plasma. Overall, the results provide evidence of interplay between the coagulation system and PMNs in contributing to HPC injury in this model.
MCP-1 can attract and activate monocytes and induce the expression of TF on the surfaces of these cells (Gu et al., 1999; Luster, 1998
). Furthermore, MCP-1 has been implicated in the expression of intercellular adhesion molecule-1 on rat endothelial cells in vitro (Yamaguchi et al., 1998
). This adhesion molecule, which is present in hepatic SECs (Essani et al., 1995
), can interact with PMNs and prime them to release toxic products (Jaeschke et al., 1996
). Accordingly, MCP-1 may be an important chemokine in the development of MCT/LPS-induced liver injury. In this model, elevated plasma MCP-1 was observed early in LPS-treated animals, irrespective of MCT treatment (Table 2
). By 18 h, MCP-1 remained elevated in both groups of LPS-treated animals, but the increase was much greater in MCT/LPS-cotreated animals. The prolonged elevation in MCP-1 was unaffected by several pharmacologic manipulations, suggesting that neither KCs, PMNs, TNF-
, nor coagulation system activation are required for increased MCP-1 production after MCT/LPS-coadministration.
Since the coagulation system is not activated in Veh/LPS animals (Yee et al., 2003), and since the same level of MCP-1 was seen in MCT/LPS and Veh/LPS-treated animals at 6 h when coagulation activation occurs, it seems unlikely that MCP-1 contributes to the early activation of the coagulation system. However, MCP-1induced TF expression might contribute to liver injury at a later time by enhancing fibrin deposition (Falati et al., 2002
; Orvim et al., 1994
; Shebuski and Kilgore, 2001). Further study will be needed to determine the function of MCP-1 in this model. It is tempting to speculate, based on the sustained increase in MCP-1 at 18 h, that it may help prolong PMN adhesion (Yamaguchi et al., 1998
) and sustain monocyte accumulation in liver.
In summary, anticoagulant therapy reduced fibrin deposition in CL and MZ regions of liver lobules, prevented HPC injury, and caused a modest attenuation in SEC injury in MCT/LPS-cotreated animals. These results point to a critical role of the coagulation system in MCT/LPS-induced liver injury. Moreover, PMNs and the coagulation system appear to cooperate in inducing HPC injury in this model of synergistic hepatotoxicity.
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ACKNOWLEDGMENTS |
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NOTES |
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