1 Laboratory of Hepatobiology and Toxicology, Department of Pharmacology, 2 Department of Surgery, 3 Department of Radiation Oncology, and 4 Department of Cell Biology and Anatomy, University of North Carolina at Chapel Hill, Chapel Hill, North Carolina 27599
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ABSTRACT |
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Harvesting trauma to the
graft dramatically decreases survival after liver
transplantation. Since activated Kupffer cells play a role in
primary nonfunction, the purpose of this study was to test the
hypothesis that organ manipulation activates Kupffer cells. To mimic
what occurs with donor hepatectomy, livers from Sprague-Dawley rats
underwent dissection with or without gentle organ manipulation in
a standardized manner in situ. Perfused livers exhibited normal values
for O2 uptake (105 ± 5 µmol · g1 · h
1) measured
polarigraphically; however, 2 h after organ manipulation, values
increased significantly to 160 ± 8 µmol · g
1 · h
1 and
binding of pimonidazole, a hypoxia marker, increased about threefold
(P < 0.05). Moreover, Kupffer cells from manipulated livers
produced three- to fourfold more tumor necrosis factor-
and
PGE2, whereas intracellular calcium concentration increased twofold after lipopolysaccharide compared with unmanipulated controls (P < 0.05). Gadolinium chloride and glycine prevented both
activation of Kupffer cells and effects of organ manipulation.
Furthermore, indomethacin given 1 h before manipulation prevented
the hypermetabolic state, hypoxia, depletion of glycogen, and release
of PGE2 from Kupffer cells. These data indicate that gentle
organ manipulation during surgery activates Kupffer cells, leading to
metabolic changes dependent on PGE2 from Kupffer cells,
which most likely impairs liver function. Thus modulation of Kupffer
cell function before organ harvest could be beneficial in human liver
transplantation and surgery.
organ harvest; liver transplantation; hypoxia; primary nonfunction
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INTRODUCTION |
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THE CAUSE OF FAILURE OF
LIVER grafts is complex and includes many factors involving organ
retrieval, preservation, and transplantation. Important factors include
general condition and nutritional status of the donor, cold and warm
ischemic times, operative complications in the recipient,
immune status of the recipient, and the experience of the surgeon
(20, 33). Recently, gentle in situ liver
manipulation during organ harvest, which cannot be prevented with
standard harvesting techniques, has been shown to dramatically decrease survival after rat liver transplantation via mechanisms including hepatic injury with reperfusion (42). Interestingly,
gadolinium chloride (GdCl3), a rare earth metal, and
Kupffer cell toxicant, and glycine, a nontoxic amino acid, given to
donors before organ harvest totally prevented all effects of
manipulation (37, 39, 41, 42), suggesting a role for
Kupffer cells in mechanisms of harvest-related injury. Once activated,
Kupffer cells release toxic mediators such as proteases, tumor necrosis
factor- (TNF-
), and arachidonic acid derivatives (5, 8,
45), which could potentially impair liver function via
mechanisms including disturbances to the microcirculation, hypoxia,
increased oxygen consumption, and depletion of hepatic glycogen
reserves (12, 24, 25, 29). Hypoxia and increased
oxygen consumption (e.g., development of a hypermetabolic state) impair
graft survival after transplantation (25, 31). Moreover,
Fusaoka et al. (13) showed that activation of Kupffer
cells increases oxygen uptake of the liver after cold storage. This
effect is most likely due to Kupffer cell-derived PGE2,
which stimulates oxygen uptake in hepatic parenchymal cells and could
be involved in early dysfunction of a graft (13, 35). Therefore, the purpose of this study was to directly test the hypothesis that the operative trauma due to surgical manipulation in
situ of donor livers activates Kupffer cells before transplantation. Preliminary accounts of this work have been published elsewhere (40).
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MATERIALS AND METHODS |
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Experimental animals and treatment. Female Sprague-Dawley rats (200-230 g) were allowed free access to standard laboratory chow (Agway PROLAB RMH 3000, Syracuse, NY) and tap water. Some animals were given a single injection of GdCl3 (10 mg/kg) through the tail vein 24 h before surgery. This treatment destroys all large Kupffer cells (15). Other rats were fed chow diet containing 5% glycine for 5 days, which blunts the response of Kupffer cells to endotoxin (18). Furthermore, some rats were given indomethacin (3.0 mg/kg, in dimethyl sulfoxide) intragastrically 1 h before experiments (6).
Surgical procedures. After midline incision, minimal dissection of livers was performed in a standardized fashion during the first 12 min, including freeing the organ from ligaments. During the subsequent 13 min, livers were either left alone in controls or manipulated gently. To maintain standard conditions, gentle manipulation was carried out by the same surgeon touching, retracting, and moving the liver lobes in situ for a specified time interval. Care was taken to use the same number of manipulations in each experiment with similar pressures. Serum transaminases at the end of manipulation were identical regardless of pretreatment, validating the standardization of the technique (42).
Nonrecirculating hemoglobin-free liver perfusion. The perfusion technique has been described elsewhere (43). Briefly, the liver is perfused ex situ via the portal vein with oxygenated (95% O2-5% CO2) Krebs-Henseleit bicarbonate buffer (in mM: 118 NaCl, 25 NaHCO3, 1.2 KH2PO4, 1.2 MgSO4, 4.7 KCl, and 1.3 CaCl2) at pH 7.6. The oxygen concentration in the effluent perfusate was monitored continuously with a Teflon-shielded platinum electrode. The inflow oxygen concentration was maintained constant and measured before and after each experiment. Metabolic rates were calculated from influent-effluent concentration differences and the constant flow rate and expressed per gram of liver weight per hour (43).
Isolation and culture of Kupffer cells. Kupffer cells were isolated by collagenase digestion and differential centrifugation using Percoll (Pharmacia, Uppsala, Sweden) as described elsewhere (32). Immediately or 2 h after organ manipulation, livers were perfused in situ via the portal vein with Ca2+- and Mg2+-free Hanks' balanced salt solution containing collagenase IV (0.025%) (Sigma Chemical, St. Louis, MO) at 37°C at a flow rate of 26 ml/min. After digestion, livers were cut into small pieces in collagenase buffer. The suspension was filtered through nylon gauze, and the filtrate was centrifuged at 450 g for 10 min at 4°C. Cell pellets were resuspended in buffer, parenchymal cells were removed by centrifugation at 50 g for 3 min, and the nonparenchymal cell fraction was washed twice with buffer. Cells were centrifuged on a density cushion of Percoll at 1,000 g for 15 min. The Kupffer cell fraction was collected and washed with buffer again. Viability of cells determined by trypan blue exclusion was >90%. Cells were seeded onto 25-mm2 glass coverslips and cultured in DMEM (GIBCO Laboratories Life Technologies, Grand Island, NY) supplemented with 10% fetal bovine serum and antibiotics (100 U/ml of penicillin G, 100 µg/ml of streptomycin sulfate) at 37°C with 5% CO2. Non-adherent cells were removed after 1 h by replacing buffer and cells were cultured for 24 h before experiments. All adherent cells phagocytized latex beads, indicating that they were Kupffer cells.
Measurement of intracellular calcium.
Intracellular calcium concentration ([Ca2+]i)
was measured fluorometrically using the calcium indicator dye fura-2
and a microspectrofluorometer (PTI, South Brunswick, NJ) interfaced
with an inverted microscope (Diaphot, Nikon, Tokyo, Japan). Kupffer
cells were incubated in modified Hank's buffer (115 mmol/l NaCl, 5 µmol KCl, 0.3 mmol/l Na2HPO4, 0.4 mmol/l
KH2PO4, 5.6 mmol/l glucose, 0.8 mmol/l
MgSO4, 1.26 mmol/l CaCl2, and 15 mmol/l HEPES,
pH 7.4) containing 5 µmol/l fura 2- acetoxymethyl ester (Molecular
Probes, Eugene, OR) and 0.03% Pluronic F-127 (BASF, Wyandotte, MI) at
room temperature for 60 min. Coverslips plated with Kupffer cells were
rinsed and placed in chambers with buffer at room temperature. Changes
in fluorescence intensity of fura 2 at excitation wavelengths of 340 and 380 nm and emission at 510 nm were monitored in individual Kupffer
cells. Each value was corrected by subtracting the system dark noise
and autofluorescence, assessed by quenching fura 2 fluorescence with
Mn2+, as described previously (17).
[Ca2+]i was determined from the following
equation: [Ca2+]i = Kd[(R Rmin)/(Rmax
R)]/(Fo/Fs), where
Fo/Fs is the ratio of fluorescent
intensities evoked by 380 nm light from fura 2 pentapotassium salt
loaded in cells using a buffer containing 3 mmol/l EGTA and 1 µmol/l
ionomycin ([Ca2+]min) or 10 mmol/l
Ca2+ and 1 µmol/l ionomycin
([Ca2+]max). R is the ratio of fluorescent
intensities at excitation wavelengths of 340 and 380 nm, and
Rmax and Rmin are values of R at
[Ca2+]max and
[Ca2+]min, respectively. The values of these
constants were determined at the end of each experiment, and a
dissociation constant (Kd) of 135 nmol/l was
used (14).
Measurement of TNF- and
PGE2.
Isolated Kupffer cells were cultured for 24 h in 24-well culture
plates (Corning, Corning, NY) at a density of 5 × 105
cells/well in DMEM supplemented with 10% fetal bovine serum and antibiotics at 37°C in the presence of 5% CO2.
Subsequently, cells were incubated with fresh media containing
lipopolysaccharide (LPS) (100 ng/ml in 5% rat serum) for an additional
4 h. Samples were stored at
80°C until assay. TNF-
concentrations were determined in the culture media using an
enzyme-linked immunosorbent assay kit (Genzyme, Cambridge, MA).
Furthermore, supernatants were assayed for PGE2 by
competitive radioimmunoassay using 125I-labeled
PGE2 (Advanced Magnetics, Cambridge, MA).
Clinical chemistry. Tissue was homogenized, and glycogen was hydrolyzed and determined enzymatically (3). Furthermore, blood was collected before experiments for glycine determination in serum as described previously (3). Briefly, glycine was extracted and benzolated, and the resulting hippuric acid was extracted and dried. Subsequently, hippuric acid was determined spectrophotometrically at 458 nm (30).
Determination of reduced, protein-bound pimonidazole by ELISA and immunohistochemistry. Pimonidazole, a noninvasive 2-nitroimidazole marker for viable hypoxic cells (11), was given to donors intravenously to detect hypoxia in liver tissue 2 h after organ manipulation. Five minutes before tissue samples were collected, pimonidazole was given to donors and pimonidazole adduct accumulation was measured in tissue homogenates with a competitive ELISA procedure described previously (36) as modified for liver tissue (2). Protein levels in tissue homogenates were determined with the bicinchoninic acid assay using a commercially available kit (Pierce Chemical, Rockford, IL). Paraffin blocks of formalin-fixed liver tissue were sectioned at 6 µm, and pimonidazole was detected with a biotin-streptavidin-peroxidase indirect immunostaining method using diaminobenzidine as a chromogen as described previously (2). After the immunostaining procedure, a counterstain of hematoxylin was applied. A Universal Imaging Image-1/AT image acquisition and analysis system (Chester, PA) incorporating an Axioskop 50 microscope (Carl Zeiss, Thornwood, NY) was used to capture and analyze the immunostained tissue sections at ×100 magnification (1). Whereas results of ELISA give only the quantity of bound pimonidazole, immunohistochemical analysis for pimonidazole demonstrates patterns of adduct binding in the liver lobule.
The number of Kupffer cells was determined immunohistochemically as described elsewhere (42). Briefly, sections (6 µm) were cut on a rotary microtome and stained for ED1-positive cells using the DAKO Envision System and a primary anti-ED1 antibody (Biosource International, Camarillo, CA). Subsequently, the tissue was stained with hematoxylin. GdCl3 under these conditions decreased the number of ED1-positive Kupffer cells by ~82%, confirming earlier reports (15).Statistics. Values (means ± SE) for groups were compared using Fisher's exact test or analysis of variance (2-way ANOVA) with Student-Newman-Keuls post hoc test as appropriate. P < 0.05 was selected before the study as the criterion for significance.
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RESULTS |
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Gentle in situ organ manipulation causes a hypermetabolic state in
liver.
In pilot experiments, oxygen uptake was measured during perfusion
of isolated livers immediately and 2 h after manipulation. Nonmanipulated controls took up oxygen with values in the normal range
(105 ± 5 µmol · g1 · h
1).
Treatment with GdCl3, glycine, or indomethacin had no
effect on oxygen consumption under these conditions. Furthermore,
immediately after manipulation, values for hepatic oxygen uptake of
manipulated livers were not different from those of unmanipulated
controls; however, oxygen consumption increased to a maximum of
160 ± 8 µmol · g
1 · h
1
(P < 0.05) within 2 h after manipulation (Fig.
1). Thus all work was carried out at the
2-h time point in this study. The increase of oxygen consumption due to
in situ manipulation of liver was totally prevented with
GdCl3, glycine, and indomethacin given before surgery
(P < 0.05) (Fig. 1).
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Gentle in situ manipulation causes hypoxia in the liver.
Pimonidazole, a 2-nitroimidazole hypoxia marker, binds to
viable hypoxic liver cells in vivo (2, 11, 36). Binding of pimonidazole in nonmanipulated controls was 196 ± 28 pmol/mg
protein. Treatment with GdCl3, glycine, or indomethacin had
no effect on binding of pimonidazole under these conditions.
Furthermore, binding of pimonidazole was increased more than twofold
(P < 0.05) in livers 2 h after gentle
manipulation (Fig. 2); however, binding was not different from controls if GdCl3, dietary glycine,
or indomethacin was given before manipulation (Fig. 2). Binding was concentrated in oxygen-poor pericentral regions of the liver lobule after manipulation (data not shown).
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Hepatic glycogen is depleted after gentle organ manipulation.
Two hours after surgery without manipulation, hepatic glycogen levels
were ~4 mg/g. Treatment with GdCl3, glycine, or
indomethacin had no effect on liver glycogen under these conditions. In
contrast, gentle organ manipulation significantly depleted glycogen to
~25% of control values 2 h after surgery (Fig.
3). GdCl3, dietary glycine, and indomethacin given before surgery totally prevented the effect of
organ manipulation on hepatic glycogen levels (Fig. 3).
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Kupffer cells are activated after gentle organ manipulation.
LPS (100 ng/ml) increased [Ca2+]i in
Kupffer cells significantly from 89 ± 9 nM in nonmanipulated
controls to 182 ± 11 nM in cells from manipulated organs (Figs.
4 and
5); however, dietary glycine given
before organ manipulation blunted the increase of [Ca2+]i (Figs. 4 and 5). In contrast, this
phenomenon was not prevented by indomethacin (Figs. 4 and 5).
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Gentle manipulation of the liver increases TNF-
and PGE2 production from Kupffer cells.
To evaluate the effect of organ manipulation on cytokine production by
Kupffer cells, LPS-induced TNF-
and PGE2 production was
measured in culture medium of isolated Kupffer cells. Isolated Kupffer
cells from manipulated livers produced ~9- to 30-fold more
PGE2 and TNF-
in the presence of LPS (100 ng/ml) than
did cells from minimally dissected livers (P < 0.05);
however, glycine given to rats before organ manipulation significantly
blunted these effects (Figs. 5 and 6). In
contrast, indomethacin had no effect on TNF-
production (Fig. 5),
whereas increased PGE2 production by Kupffer cells from
manipulated livers was totally prevented by indomethacin as expected
(Fig. 6; P < 0.05).
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DISCUSSION |
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Gentle in situ manipulation of liver activates Kupffer cells.
Primary nonfunction and dysfunction occur in 5-30% of human liver
transplantation cases leading to significant morbidity and mortality
(33); however, underlying mechanisms are largely unknown but most likely involve Kupffer cells, which play a role in the development of reperfusion injury and primary nonfunction
(24). Furthermore, the donor operation and surgical
technique most likely have an effect on outcome after transplantation.
Even laparotomy with mild abdominal exploration and preparation of the
portal vein alone impair the intrahepatic circulation (9,
21-23). This is important, because hypoxia can activate
Kupffer cells (26). Indeed, in a recent study, gentle in
situ liver manipulation during organ harvest rapidly disturbed
intrahepatic microcirculation and hypoxia developed rapidly as a result
of vasoconstriction caused by nerves to the liver (Fig.
7) (38). These immediate effects of organ manipulation increased injury to the liver upon reperfusion and decreased survival after liver transplantation dramatically via mechanisms involving Kupffer cells (39, 41, 42). Both denervation of the liver and inactivation of Kupffer cells with GdCl3 and dietary glycine prevented all
detrimental effects of organ manipulation. Thus it is possible that
stimulation of nerves to the liver may be involved in the development
of Kupffer cell-dependent injury on reperfusion as a result of in situ
liver manipulation during harvest for transplantation (39)
(Fig. 7). However, the exact underlying mechanisms by which liver
becomes predisposed for failure after manipulation still remain
unclear. Therefore, this study was designed to mimic what occurs during donor hepatectomy and to investigate its effects on Kupffer cells. Interestingly, oxygen consumption (Fig. 1) and hypoxia (Fig. 2) were
increased significantly, whereas hepatic glycogen was depleted (Fig.
3), 2 h after manipulation. Under these conditions, Kupffer cells,
the major source of eicosanoids and cytokines in the liver (26), were activated by manipulation reflected by
increased [Ca2+]i (Figs. 4, 5, and 7),
TNF- (Fig. 5), and PGE2 production (Figs. 6 and 7).
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PGE2 from activated Kupffer cells is
responsible for metabolic changes in liver.
How can this be explained? Activation of Kupffer cells by organ
manipulation increases [Ca2+]i in Kupffer
cells. It is well established that Ca2+ activates
phospholipases, leading to increased synthesis of TNF- and
PGE2 (4, 10). Qu et al. (35) have
shown that PGE2 from Kupffer cells stimulates oxygen uptake
in parenchymal cells, whereas TNF-
was without effect.
PGE2 acts on receptors in parenchymal cells to stimulate
mitochondrial respiration via second messenger systems most likely
involving cAMP. As a result, oxygen uptake increased, which can
partially be explained by enhanced demand of mitochondrial oxidative
phosphorylation for oxygen to compensate for reduced extramitochondrial
ATP production due to inhibition of glycolysis due to substrate
depletion. Indeed, in this study, manipulation increased oxygen uptake
(Fig. 1) at the time when Kupffer cells were activated (Figs. 4 and 5)
and production of PGE2 increased (Fig. 6). Whereas hypoxia
immediately after organ manipulation may be due to vasoconstriction
mediated by nerves to the liver (38), hypoxia concentrated
in pericentral areas measured 2 h after manipulation is most
likely due to a hypermetabolic state, which causes a steeper oxygen
gradient along the hepatic sinusoid (Fig. 2). Furthermore, depletion of
hepatic glycogen during manipulation can be explained by both hypoxia
and increased PGE2 production, which causes glycogenolysis
(16, 27, 29) (Fig. 3). To test the hypothesis that
PGE2 from Kupffer cells was responsible for changes after
organ manipulation, donors were pretreated with GdCl3, a
rare earth metal, and Kupffer cell toxicant (15), dietary
glycine, a nonessential amino acid that prevents activation of Kupffer
cells (19), or indomethacin, an inhibitor of
cyclooxygenase, which prevents PGE2 production
(35). Indeed, GdCl3 and glycine prevented the
hypermetabolic state (Fig. 1), hypoxia (Fig. 2), depletion of glycogen
(Fig. 3), activation of Kupffer cells (Figs. 4 and 5), and increased
PGE2 production (Fig. 6) in manipulated livers (Fig. 7).
These data suggest that PGE2 from activated Kupffer cells
most likely mediates the metabolic changes (e.g., respiratory burst,
glycogenolysis) and hypoxia observed 2 h after organ manipulation
(Fig. 7).
Possible relationship between gentle organ manipulation and
viability of the graft.
The vulnerability of liver to hypoxic injury is greatly affected by
nutritional status. Thurman et al. (44) have shown that the hypermetabolic state induced by ethanol results in hepatic glycogen
depletion within a few hours. Such livers were much more susceptible to
anoxic injury (25, 44). In contrast, glycogen-rich livers
from fed animals are resistant to anoxic injury due to glycolytic ATP
formation utilizing endogenous glycogen as substrate (7,
28). Mitochondria typically supply the vast majority of ATP to
aerobic hepatocytes. However, in the first hours after liver
transplantation, glucose utilization by the graft is impaired until the
redox state of the mitochondria improves (31). Thus hepatic glycogen is essential to minimize reperfusion injury and to
improve survival after transplantation (25). It is likely that an increase of [Ca2+]i activates
cyclooxygenase and increases PGE2 production by Kupffer cells (Fig. 6), which causes a hypermetabolic state (Fig. 1), hypoxia
(Fig. 2) and depletion of glycogen (Fig. 3) after manipulation. Moreover, activated Kupffer cells release numerous inflammatory mediators, including oxygen radicals, TNF-, interleukins-1 and -6, prostaglandins, and nitric oxide (24), leading to injury and an increase of oxygen consumption in livers after transplantation (34). Because Kupffer cells are activated, oxygen
consumption and tissue hypoxia are dramatically increased and glycogen
is depleted during organ harvest, predisposing livers to primary nonfunction.
Conclusion and clinical implication. PGE2 from activated Kupffer cells causes a hypermetabolic state, hypoxia and depletion of hepatic glycogen in the donor due to manipulation during surgery, which is nearly inevitable. Because these changes are linked with reperfusion injury and primary graft nonfunction (12, 24, 25, 29, 34) and livers manipulated gently during harvest fail often after transplantation (37, 41, 42), modulation of Kupffer cell function with the nontoxic amino acid glycine before organ harvest could be beneficial in clinical liver transplantation.
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ACKNOWLEDGEMENTS |
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We thank Neha Mehta and Julie Vorobiov in the Center for
Gastrointestinal Biology and Disease, University of North Carolina at
Chapel Hill (supported by National Institute of Diabetes and Digestive
and Kidney Diseases Grant P30 DK-34987), for assistance with TNF-
and PGE2 measurements.
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FOOTNOTES |
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This work was supported, in part, by grants from the National Institute on Alcohol Abuse and Alcoholism, the National Cancer Institute, and the Deutsche Forschungsgemeinschaft.
Address for reprint requests and other correspondence: P. Schemmer, Dept. of Surgery, Univ. of Heidelberg, Im Neuenheimer Feld 110, 69120 Heidelberg, Germany (E-mail: Peter_Schemmer{at}med.uni-heidelberg.de).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 27 June 2000; accepted in final form 9 January 2001.
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