Toxicological Sciences 67, 322-328 (2002)
Copyright © 2002 by the Society of Toxicology
SYSTEMS TOXICOLOGY |
Mode of Cell Death after Acetaminophen Overdose in Mice: Apoptosis or Oncotic Necrosis?
* Department of Pharmacology and Toxicology, University of Arkansas for Medical Sciences, 4301 W. Markham St. (Mailslot 638), Little Rock, Arkansas 72205; and
Department of Pathology, University of Texas Health Science Center, Houston, Texas
Received October 25, 2001; accepted January 8, 2002
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ABSTRACT |
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Acetaminophen (AAP) overdose can cause severe liver injury and liver failure in experimental animals and humans. Recently, several authors proposed that apoptosis might be a major mechanism of cell death after AAP treatment. To address this controversial issue, we evaluated a detailed time course of liver injury after AAP (300 mg/kg) in fasted C3Heb/FeJ mice. Apoptotic hepatocytes were quantified in H&E-stained liver sections using morphologic criteria (cell shrinkage, chromatin condensation and margination, and apoptotic bodies). The number of apoptotic hepatocytes remained at baseline (0.2 ± 0.1 cells/10 high-power fields [HPF]) up to 2 h after AAP administration. However, between 3 and 24 h, apoptotic cell death increased significantly, e.g., 6.3 ± 0.8 cells/10 HPF at 6 h. Despite the increase in the number of hepatocytes meeting the morphological criteria of apoptosis, this cell fraction remained well below 1% of all parenchymal cells. No evidence for caspase-3 processing or increase in enzyme activity was detected at any time. These results were compared to the overall percent of necrotic cells in liver sections. Confluent areas of centrilobular necrosis were estimated to involve 40–60% of all hepatocytes between 3 and 24 h after AAP administration. These numbers correlated with the increase in plasma alanine aminotransferase activities, which reached a peak level of 5900 ± 1350 U/l at 24 h. A similar result was obtained with higher doses of AAP and with the use of fed animals. Thus, oncotic necrosis and not apoptosis is the principal mechanism of liver-cell death after AAP overdose in vivo.
Key Words: acetaminophen; liver failure; cell death; apoptosis; oncosis; necrosis; caspases.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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An overdose of acetaminophen (AAP) can cause severe centrilobular-cell injury and even liver failure in experimental animals and humans. Toxicity requires formation of a reactive metabolite, presumably N-acetylbenzoquinone imine (NAPQI) or an intermediate resembling it, in excess of the available glutathione (Mitchell et al., 1973
). After consumption of glutathione, NAPQI can covalently bind to a number of intracellular target proteins (Cohen and Khairallah, 1997; Pumford et al., 1997; Qiu et al., 1998), which leads to a variety of cellular dysfunctions, including mitochondrial damage, ATP depletion, and mitochondrial oxidant stress (Jaeschke, 1990; Meyers et al., 1988; Tirmenstein et al., 1990). Furthermore, accumulation of calcium in nuclei and fragmentation of DNA has been observed (Ray et al., 1993; Shen et al., 1991). Recently, peroxynitrite formation and protein nitration has been postulated to be an important factor in the pathophysiology (Hinson et al., 1998; Knight et al., 2001b; Gardner et al., 1998). Although the exact mechanism of AAP-induced cell injury is still not completely understood, it was always assumed that the mode of cell death was oncosis or oncotic necrosis (Cohen and Khairallah, 1997; Pumford et al., 1997). However, in recent years, an increasing number of authors have postulated that apoptosis plays a major role in AAP-induced liver failure (Ferret et al., 2001; Kanno et al., 2000; Ray et al., 1996; Zhang et al., 2000). Moreover, AAP induces apoptosis in other cell types (Ruppova et al., 1999; Wiger et al., 1997). For the liver, it was hypothesized that 40% or more of the hepatocytes actually die by apoptosis (Ray et al., 1996; Ray and Jena, 2000). Apparent experimental evidence for apoptotic cell death after AAP overdose in vivo included DNA fragmentation and DNA laddering (Ray et al., 1993), cleavage of poly(ADP-ribose)polymerase (PARP; Zhang et al., 2000), DNA strand breaks detected by terminal deoxynucleotidyl transferase-mediated dUTP nick end labeling (TUNEL) assay (Lawson et al., 1999), and morphological evidence for apoptosis of individual hepatocytes (electron and light microscopy; Ray et al., 1996; Ray and Jena, 2000). Although most of the biochemical parameters are characteristic for apoptotic cell death, none of these assays are specific for apoptosis and have been shown to be positive in oncotic cell death (Dong et al., 1997; Grasl-Kraupp et al., 1995; Gujral et al., 2001). In addition, caspases, which are critical signaling molecules in apoptosis, are not activated after AAP overdose (Lawson et al., 1999). In contrast, AAP toxicity prevents caspase activation after Fas receptor-induced apoptosis (Lawson et al., 1999). Thus, the mode of cell death induced by AAP is controversial. Because of the different therapeutic intervention points in apoptotic versus oncotic necrosis, it is important to clarify the mode of cell death in AAP hepatotoxicity. To address this issue, we analyzed and quantified hepatocellular injury during the first 24 h after AAP administration, using a combination of strict morphological criteria, TUNEL assay, and biochemical assays. |
MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Animals.
Male C3HeB/FeJ mice (20–30 g body weight) were purchased from Jackson Laboratories (Bar Harbor, Maine). The animals had free access to food (certified rodent diet #5002C; PMI Feeds, Inc., Richmond, IN) and water. The experimental protocols followed the criteria of the University of Arkansas for Medical Sciences and the National Research Council for the care and use of laboratory animals in research.
Experimental protocol.
Animals were fasted overnight before the experiments. They were injected intraperitoneally with acetaminophen (300–500 mg AAP/kg) dissolved in phosphate buffered saline or vehicle (16 ml/kg). The animals were sacrificed at various time-points between 30 min and 24 h after AAP administration. As a positive control for apoptosis, mice were treated with 700 mg/kg galactosamine and 100 µg/kg endotoxin for 6 h (Jaeschke et al., 1998; Lawson et al., 1998). A blood sample was collected from the vena cava with a heparinized syringe. Samples of each liver were fixed in phosphate-buffered formalin for histological analysis, snap-frozen in liquid nitrogen, or immediately homogenized for caspase-3 activity measurements and Western blotting.
Analytical procedures.
Plasma was used for determination of alanine aminotransferase (ALT) activity with test kit DG 159-UV (Sigma Chemical, St. Louis, MO). Caspase-3 activities were determined as described in detail (Jaeschke et al., 1998). Briefly, a liver sample was homogenized in 25 mM HEPES buffer (pH 7.5) containing 5 mM EDTA, 2 mM DTT, and 0.1% CHAPS. After centrifugation at 14,000 g, the diluted supernatant was assayed for caspase activity using the synthetic fluorogenic substrate Ac-DEVD-MCA (Acetyl-Asp-Glu-Val-Asp-4-methylcoumaryl-7-amide; Peptide Institute, Osaka, Japan) for caspase 3 (CPP32) at concentrations of 50 µM. The samples were assayed in duplicate wells, with or without 10 µM pancaspase inhibitor Z-VAD-fmk (Z-Val-Ala-Asp-fluoromethylketone, Alexis Corp., San Diego, CA). The kinetics of the proteolytic cleavage of the substrate was monitored in a fluorescence microplate reader (Cytofluor 2350, Millipore, Bedford, MA) using an excitation wavelength of 360 nm and an emission wavelength of 460 nm. Caspase activity was calculated from the slope of the recorder trace and expressed in 
F/min/mg protein. All caspase activities are reported as ZVAD-inhibitable enzyme activities. Protein concentrations in the supernatant were assayed using the bicinchoninic acid kit (Sigma). Caspase-3 processing was evaluated by Western blot analysis as described (Bajt et al., 2000, 2001). Liver tissue was homogenized in 25 mM HEPES (pH 7.5) containing 5 mM EDTA, 2 mM DTT, 0.1% CHAPS, 1 mg/ml pepstatin, leupeptin, and aprotinin. Homogenates were centrifuged at 14,000 x g at 4°C for 20 min. Cytosolic extracts (50 µg per lane) were resolved by 4–20% SDS–polyacrylamide gel electrophoresis under reducing conditions. After transfer to polyvinylidine difluoride membranes (PVDF, Immobilin-P, Millipore, Bedford, MA), the membranes were first blocked with 5% milk overnight at 4°C followed by incubation with primary antibody for 2 h at room temperature. A goat anti-caspase 3 polyclonal IgG (Santa Cruz Biotechnology) was used as a primary antibody. The membranes were washed and then incubated with the secondary antibody anti-goat IgG-HRP (Santa Cruz Biotechnology). Proteins were visualized by enhanced chemiluminescence (Amersham Pharmacia Biotech, Inc., Piscataway, NJ) according to the manufacturer's instructions.
Histology.
Formalin-fixed tissue samples were embedded in paraffin and 5-µm sections were cut. Replicate sections were either stained with hematoxylin and eosin (H&E) for evaluation of necrosis and apoptosis (Gujral et al., 2001) or stained with the terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling (TUNEL) assay (Roche Molecular Biochemicals, Indianapolis, IN). The numbers of apoptotic hepatocytes were counted in 10 high-power fields (x400 magnification) using a KF2 microscope (Carl Zeiss, Inc., Thornwood, NY). Preliminary counts in untreated control livers confirmed that this area contained an average of 1800 hepatocytes. Apoptotic cells were identified by morphological criteria (cell shrinkage, chromatin condensation and margination, apoptotic bodies) and by staining with the TUNEL assay. Cell necrosis was evaluated in replicate sections stained with hematoxylin and eosin. The percent of necrosis was estimated by evaluating the number of microscopic fields with necrosis compared to the entire histologic section. All histological evaluations were done in a blinded fashion by 2 investigators (A.F. and J.S.G.).
Statistics.
Data are given as mean ± SE. Comparisons between multiple groups were performed with 1-way ANOVA followed by Bonferroni t-test. If the data were not normally distributed, the Kruskal-Wallis Test (nonparametric ANOVA) followed by Dunn's Multiple Comparisons Test was performed; p < 0.05 was considered significant.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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C3Heb/FeJ mice treated with 300 mg/kg of acetaminophen developed substantial liver cell injury as indicated by the progressive increase of plasma ALT activities (Fig. 1
). To assess the predominant mode of cell death, sections of these livers were then stained with H&E (Fig. 2) and the TUNEL assay (Fig. 3). Apoptotic cells were identified by morphological criteria such as cell shrinkage, chromatin condensation and margination, and apoptotic bodies. Hepatocytes undergoing oncotic necrosis were identified using the following criteria: increased eosinophilia, cell swelling and lysis, loss of architecture, karyolysis, and karyorrhexis. In controls, apoptotic hepatocytes were rare (Figs. 2A and 3A). The sinusoids appeared normal and there was no congestion. After 3 h of acetaminophen, apoptotic hepatocytes were still very infrequent (Fig. 2B). Hepatocytes were eosinophilic with scattered foci of necrosis. Vacuolation and early stages of karyorrhexis were frequently observed. Sinusoids were focally congested with red blood cells. After 6 h, there were extensive areas of centrilobular oncotic necrosis but very few apoptotic cells (Fig. 2C). At both time points, a number of hepatocytes around the centrilobular areas stained positive with the TUNEL assay (Figs. 3B and 3C). However, based on morphological criteria, these cells were not apoptotic. As a positive control for apoptosis, mice were treated with 700 mg/kg galactosamine and 100 mg/kg endotoxin (Gal/ET) for 6 h. Sections of livers from these mice showed numerous apoptotic hepatocytes and apoptotic bodies (Fig. 2D). All apoptotic cells stained positive with the TUNEL assay (Fig. 3D), and all TUNEL-positive cells showed morphological characteristics of apoptosis. However, there was a clear difference between the cellular staining pattern of hepatocytes after Gal/ET compared to AAP. Whereas all cells after Gal/ET treatment had a very distinct nuclear staining (Fig. 3D), hepatocytes after AAP showed a diffuse staining in the cytosol and the nucleus.
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Apoptosis and necrosis of hepatocytes were then quantitated in H&E and TUNEL-stained liver sections (Figs. 4 and 5
). Apoptotic hepatocytes were counted in 10 random HPF of the tissue sections, using morphological criteria and TUNEL staining. Controls had 0.15 ± 0.14 apoptotic hepatocytes in 10 HPF, which represents approximately 0.01% of all hepatocytes (Fig. 4). The number of apoptotic cells increased by 4-fold 1 h after AAP and by 45-fold at 6 h. Despite the large increase at 6 h, the number of apoptotic cells was 6.3 ± 0.8 per 10 HPF and constituted 0.35% of the total cells evaluated. In mice treated with Gal/ET, the liver sections showed 427 ± 28 apoptotic cells/10 HPF (23.7% of all hepatocytes). Estimation of necrosis in AAP-treated livers revealed a progressive increase from 0% at 0.5 h up to 67 ± 12% at 24 h (Fig. 5). These data correlated well with the plasma ALT values. Thus, on a quantitative basis, more than 99% of all dead or dying cells showed signs of oncotic necrosis, and less than 1% of these cells fulfilled the morphological criteria of apoptosis.
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Caspase-3 is one of the effector caspases in apoptosis, and its activation is an integral part of this mode of cell death. Therefore, we analyzed liver homogenates for caspase-3 processing by Western blotting (Fig. 6
). None of the treatment groups showed any processing of the procaspase-3 (p32) to its active fragment (p11). Consistent with these data, no significant increase in caspase-3 activity was found in any AAP-treated group compared to baseline values (0.9 ± 0.9 F/min/mg protein). As a positive control, samples from mice treated with Gal/ET for 6 h were used. As expected, these livers showed significant processing of the proenzyme to its active fragment (Fig. 6) and a 375-fold increase in caspase-3 activity (338 ± 47 F/min/mg protein).
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To evaluate whether the mode of cell death might change with higher doses of AAP, animals were treated with 300, 400, and 500 mg/kg AAP and killed after 6 h. All groups of animals had severe liver injury as indicated by plasma ALT values of 2500–5000 U/l and extensive necrosis (300 mg/kg: 35 ± 5%; 400 mg/kg: 45 ± 3%; 500 mg/kg: 43 ± 5%). However, the number of apoptotic cells significantly declined with increasing doses of AAP (300 mg/kg: 4.0 ± 0.4 cells/10 HPF; 400 mg/kg: 1.8 ± 0.5; 500 mg/kg: 0.5 ± 0.5).
Recently, it was reported that fed mice developed liver injury and apoptosis 12 h after injection with 300 mg/kg AAP (Zhang et al., 2000). Therefore, we repeated our experiment with fed mice and evaluated apoptosis and necrosis 12 h after AAP injection. Consistent with the feeding-dependent diurnal rhythm of liver glutathione (Jaeschke and Wendel, 1985), fed mice, which received 300 mg/kg AAP in the evening, had extensive liver injury (ALT: 6270 ± 1395 IU/l; n = 3). In contrast, injection of AAP during the peak levels of hepatic glutathione in the morning did not result in any relevant liver injury (ALT: 140 ± 20 IU/l). Despite the difference in liver injury, there was no evidence of any significant changes in hepatic caspase-3 activity compared to controls or between the 2 AAP-treated groups (1.1 ± 0.6 F/min/mg protein). Quantitation of apoptotic hepatocytes showed a slightly higher number of apoptotic cells in the AAP-treated group with injury (3.5 ± 0.9 cells/10 HPF) compared to the one without injury (0.5 ± 0.5). However, even the elevated numbers are in the range observed with starved mice (Fig. 4
).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The aim of this investigation was to evaluate the mode of cell death in the liver during AAP overdose. Our approach was to use the standard morphological criteria of apoptosis, i.e., cell shrinkage, chromatin condensation and margination, and apoptotic body formation (Kerr et al., 1972
). Using these criteria, we demonstrated a significant increase in the number of cells after AAP that met the definition of apoptosis. However, on a quantitative basis, the total number of necrotic cells was approximately 2 orders of magnitude higher. This suggests that 99% of all dying or dead cells were undergoing an oncotic process characterized by cell swelling and lysis, vacuolation, karyorrhexis, and karyolysis. Further support for this conclusion comes from the observation that the damaged cells form a confluent area (Fig. 2C), which is characteristic for oncotic necrosis. On the other hand, apoptosis is usually a single-cell event, even when a large number of cells are involved (Fig. 2D). Some investigators show a transmission electron micrograph to confirm the apoptotic morphology of an injured cell (Ray et al., 1996; Ray and Jena, 2000). However, apoptosis, in contrast to oncotic necrosis, occurs also in livers from untreated animals. The number of apoptotic cells in control livers ranges from 0.01% (Fig. 4) to 0.3% (Gujral et al., 2001). This means that between 10,000 and 300,000 hepatocytes per gram of liver undergo apoptosis at any given time. Thus, showing an electron micrograph of a single apoptotic cell (Ray et al., 1996; Ray and Jena, 2000), without quantitative assessment of the overall number of apoptotic cells, calls into question the validity of the conclusion that apoptosis is the major mode of cell death after AAP overdose.
A ladder pattern of DNA fragments on an agarose gel created by multiples of 180-bp fragments (Nagata, 2000) and leakage of DNA/histone fragments into the cytosol (Leist et al., 1995) are characteristic features of apoptotic cells. Furthermore, DNA strand breaks, as indicated by the TUNEL assay, also occur in apoptotic cells. However, none of these assays is absolutely specific for an apoptotic cell (Dong et al., 1997; Grasl-Kraupp et al., 1995; Gujral et al., 2001). DNA ladders (Dong et al., 1997) and DNA/histone fragments in the cytosol Gujral et al., 2001; Lawson et al., 1998) have been observed in oncotic cells. Moreover, both apoptotic and oncotic cells can be TUNEL-positive (Grasl-Kraupp et al., 1995; Gujral et al., 2001). However, as clearly demonstrated in Figure 3, the staining pattern shows a distinct nuclear staining in apoptotic hepatocytes from Gal/ET-treated livers compared to generalized staining of the entire cell during an oncotic process induced by AAP toxicity. Furthermore, a clear difference in the size of DNA fragments released into the plasma of animals was also found after Fas receptor-mediated apoptosis compared to AAP-induced necrosis (Jahr et al., 2001). A spectrum of multiples of 180-bp fragments dominates after Fas receptor-induced apoptosis (Jahr et al., 2001), which is characteristic for caspase-activated endonucleases (Nagata, 2000). However, 6 h after AAP administration, the overall concentration of DNA in plasma is almost 10-fold higher compared to Fas-induced apoptosis. In addition, the majority of DNA fragments are >10,000 bp (Jahr et al., 2001), which is characteristic for karyolysis during oncotic necrosis. Thus, DNA fragmentation and positive TUNEL staining during Fas or TNF receptor-mediated apoptosis and AAP-induced cell injury appear to be the result of different processes. These findings further question if apoptotic cell death is relevant for the pathophysiology of AAP-induced liver failure.
Extensive investigations into the signaling mechanisms of receptor-mediated apoptosis demonstrated a critical role of the caspase cascade of proteases (Cohen, 1997). Proteolytic processing of constitutively present procaspases leads to the formation of the active enzymes (Cohen, 1997). Caspase-3 is one of the prominent downstream effector caspases. The processing of this enzyme and the increased enzyme activity are easily detectable in the liver during TNF-
(Jaeschke et al., 1998Jaeschke et al., 2000) and Fas receptor-mediated hepatocellular apoptosis in vivo (Bajt et al., 2000, 2001). Furthermore, pancaspase inhibitors effectively prevent processing of this enzyme and protect against receptor-mediated apoptosis (Jaeschke et al., 1998; 2000; Lawson et al., 1999). In striking contrast to these findings, caspase-3 is not processed during the first 24 h after AAP administration (Fig. 6), there is no increase in caspase-3 activity, and pancaspase inhibitors do not prevent AAP-induced liver injury (Lawson et al., 1999). In fact, AAP treatment actually inhibits Fas receptor-mediated apoptosis (Lawson et al., 1999), presumably by causing mitochondrial injury and interrupting the vital signal transduction through the mitochondria (Knight et al., 2001a). Thus, AAP not only fails to induce apoptotic signaling mechanisms in hepatocytes by itself, but AAP even prevents the execution of the intracellular signaling cascade of Fas-receptor-mediated apoptosis.
Apoptosis and oncosis are not completely independent processes (Lemasters, 1999). In particular, when a large number of cells are undergoing apoptosis, the process may switch to "secondary necrosis," which then may become indistinguishable from oncotic necrosis (Levin et al., 1999). However, as shown with the Fas antibody model, such a mechanism starts out with all the characteristic features of apoptosis, i.e., apoptotic morphology, caspase activation, and DNA fragmentation without cell content release (Bajt et al., 2000). Only at later stages does secondary necrosis with cell lysis occur. However, many of the apoptotic features such as caspase activation are still detectable at this time (Bajt et al., 2000). In contrast, in our detailed time course evaluation of AAP-induced liver injury, we could not find, at any time, evidence for relevant morphological or biochemical changes characteristic of apoptosis. The observations made with AAP overdose, especially when compared to the secondary necrosis seen after Fas receptor-induced apoptosis (Bajt et al., 2000), clearly suggest that this cell injury is the result of an oncotic process and not secondary necrosis.
In summary, our data showed the time-dependent development of severe hepatocellular injury after various doses of AAP. Based on morphological evaluation, we found that the number of apoptotic hepatocytes was very limited and never exceeded 0.35% of all injured cells. These findings were corroborated by lack of caspase-3 processing and the absence of any increase in caspase-3 enzyme activities in these livers. In contrast, extensive, confluent centrilobular oncotic necrosis of up to 60% of all hepatocytes was observed. Therefore, we conclude that oncotic necrosis and not apoptosis is the predominant mode of cell death during AAP overdose in mice.
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ACKNOWLEDGMENTS |
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This investigation was supported in part by National Institutes of Health grants ES0906 and AA12916.
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NOTES |
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1 To whom correspondence should be addressed. Fax: (501) 686-8970. E-mail: jaeschkehartmutw{at}uams.edu.
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  M. L. Bajt, H.-M. Yan, A. Farhood, and H. Jaeschke Plasminogen Activator Inhibitor-1 Limits Liver Injury and Facilitates Regeneration after Acetaminophen Overdose Toxicol. Sci., August 1, 2008; 104(2): 419 - 427. [Abstract] [Full Text] [PDF]
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 B. Hu and L. M. Colletti Stem cell factor and c-kit are involved in hepatic recovery after acetaminophen-induced liver injury in mice Am J Physiol Gastrointest Liver Physiol, July 1, 2008; 295(1): G45 - G53. [Abstract] [Full Text] [PDF]
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 B. Saberi, M. Shinohara, M. D. Ybanez, N. Hanawa, W. A. Gaarde, N. Kaplowitz, and D. Han Regulation of H2O2-induced necrosis by PKC and AMP-activated kinase signaling in primary cultured hepatocytes Am J Physiol Cell Physiol, July 1, 2008; 295(1): C50 - C63. [Abstract] [Full Text] [PDF]
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 N. Hanawa, M. Shinohara, B. Saberi, W. A. Gaarde, D. Han, and N. Kaplowitz Role of JNK Translocation to Mitochondria Leading to Inhibition of Mitochondria Bioenergetics in Acetaminophen-induced Liver Injury J. Biol. Chem., May 16, 2008; 283(20): 13565 - 13577. [Abstract] [Full Text] [PDF]
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 M. L. Bajt, A. Farhood, J. J. Lemasters, and H. Jaeschke Mitochondrial Bax Translocation Accelerates DNA Fragmentation and Cell Necrosis in a Murine Model of Acetaminophen Hepatotoxicity J. Pharmacol. Exp. Ther., January 1, 2008; 324(1): 8 - 14. [Abstract] [Full Text] [PDF]
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I. Manov, Y. Bashenko, A. Eliaz-Wolkowicz, M. Mizrahi, O. Liran, and T. C. Iancu High-Dose Acetaminophen Inhibits the Lethal Effect of Doxorubicin in HepG2 Cells: The Role of P-glycoprotein and Mitogen-Activated Protein Kinase p44/42 Pathway J. Pharmacol. Exp. Ther., September 1, 2007; 322(3): 1013 - 1022. [Abstract] [Full Text] [PDF]
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 A. N. Heinloth, G. A. Boorman, J. F. Foley, N. D. Flagler, and R. S. Paules Gene Expression Analysis Offers Unique Advantages to Histopathology in Liver Biopsy Evaluations Toxicol Pathol, February 1, 2007; 35(2): 276 - 283. [Abstract] [Full Text] [PDF]
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 J.-H. Zhu, X. Zhang, J. P. McClung, and X. G. Lei Impact of Cu,Zn-Superoxide Dismutase and Se-Dependent Glutathione Peroxidase-1 Knockouts on Acetaminophen-Induced Cell Death and Related Signaling in Murine Liver Experimental Biology and Medicine, December 1, 2006; 231(11): 1726 - 1732. [Abstract] [Full Text] [PDF]
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M. L. Bajt, C. Cover, J. J. Lemasters, and H. Jaeschke Nuclear Translocation of Endonuclease G and Apoptosis-Inducing Factor during Acetaminophen-Induced Liver Cell Injury Toxicol. Sci., November 1, 2006; 94(1): 217 - 225. [Abstract] [Full Text] [PDF]
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H. Jaeschke and M. L. Bajt Intracellular Signaling Mechanisms of Acetaminophen-Induced Liver Cell Death Toxicol. Sci., January 1, 2006; 89(1): 31 - 41. [Abstract] [Full Text] [PDF]
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C. Cover, A. Mansouri, T. R. Knight, M. L. Bajt, J. J. Lemasters, D. Pessayre, and H. Jaeschke Peroxynitrite-Induced Mitochondrial and Endonuclease-Mediated Nuclear DNA Damage in Acetaminophen Hepatotoxicity J. Pharmacol. Exp. Ther., November 1, 2005; 315(2): 879 - 887. [Abstract] [Full Text] [PDF]
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H. Jaeschke Comments on "Glycogen Synthase Kinase-3 Mediates Acetaminophen-Induced Apoptosis in Human Hepatoma Cells" J. Pharmacol. Exp. Ther., September 1, 2005; 314(3): 1401 - 1402. [Full Text] [PDF]
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G. E. N. Kass Response: Comments on "Glycogen Synthase Kinase-3 Mediates Acetaminophen-Induced Apoptosis in Human Hepatoma Cells" J. Pharmacol. Exp. Ther., September 1, 2005; 314(3): 1403 - 1404. [Full Text] [PDF]
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P. Macanas-Pirard, N.-S. Yaacob, P. C. Lee, J. C. Holder, R. H. Hinton, and G. E. N. Kass Glycogen Synthase Kinase-3 Mediates Acetaminophen-Induced Apoptosis in Human Hepatoma Cells J. Pharmacol. Exp. Ther., May 1, 2005; 313(2): 780 - 789. [Abstract] [Full Text] [PDF]
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E. J. Martin and P.-G. Forkert 1,1-Dichloroethylene-Induced Mitochondrial Damage Precedes Apoptotic Cell Death of Bronchiolar Epithelial Cells in Murine Lung J. Pharmacol. Exp. Ther., April 1, 2005; 313(1): 95 - 103. [Abstract] [Full Text] [PDF]
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C. Cover, P. Fickert, T. R. Knight, A. Fuchsbichler, A. Farhood, M. Trauner, and H. Jaeschke Pathophysiological Role of Poly(ADP-Ribose) Polymerase (PARP) Activation during Acetaminophen-Induced Liver Cell Necrosis in Mice Toxicol. Sci., March 1, 2005; 84(1): 201 - 208. [Abstract] [Full Text] [PDF]
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A. B. Reid, R. C. Kurten, S. S. McCullough, R. W. Brock, and J. A. Hinson Mechanisms of Acetaminophen-Induced Hepatotoxicity: Role of Oxidative Stress and Mitochondrial Permeability Transition in Freshly Isolated Mouse Hepatocytes J. Pharmacol. Exp. Ther., February 1, 2005; 312(2): 509 - 516. [Abstract] [Full Text] [PDF]
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E. J. Martin and P.-G. Forkert Evidence That 1,1-Dichloroethylene Induces Apoptotic Cell Death in Murine Liver J. Pharmacol. Exp. Ther., July 1, 2004; 310(1): 33 - 42. [Abstract] [Full Text] [PDF]
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