Endotoxin (ET)-induced liver failure is characterized by parenchymal cell apoptosis and inflammation leading to liver cell necrosis. Members of the caspase family have been implicated in the signal transduction pathway of apoptosis. The aim of this study was to characterize ET-induced hepatic caspase activation and apoptosis and to investigate their effect on neutrophil-mediated liver injury. Treatment of C3Heb/FeJ mice with 700 mg/kg galactosamine (Gal) and 100 μg/kg Salmonella abortus equi ET increased caspase 3-like protease activity (Asp-Val-Glu-Asp-substrate) by 1730 ± 140% at 6 h. There was a parallel enhancement of apoptosis (assessed by DNA fragmentation ELISA and terminal deoxynucleotidyl transferase-mediated dUTP nick end labeling assay). In contrast, activity of caspase 1 (IL-1β-converting enzyme)-like proteases (Tyr-Val-Ala-Asp-substrate) did not change throughout the experiment. Caspase 3-like protease activity and apoptosis was not induced by Gal/ET in ET-resistant mice (C3H/HeJ). Furthermore, only murine TNF-α but not IL-1αβ increased caspase activity and apoptosis. Gal/ET caused neutrophil-dependent hepatocellular necrosis at 7 h (area of necrosis, 45 ± 3%). Delayed treatment with the caspase 3-like protease inhibitor Z-Val-Ala-Asp-CH2F (Z-VAD) (10 mg/kg at 3 h) attenuated apoptosis by 81 to 88% and prevented liver cell necrosis (≤5%). Z-VAD had no effect on the initial inflammatory response, including the sequestration of neutrophils in sinusoids. However, Z-VAD prevented neutrophil transmigration and necrosis. Our data indicate that activation of the caspase 3 subfamily of cysteine proteases is critical for the development of parenchymal cell apoptosis. In addition, excessive hepatocellular apoptosis can be an important signal for transmigration of primed neutrophils sequestered in sinusoids.

Liver dysfunction and failure are common problems during endotoxemia and sepsis (1). It is generally accepted that proinflammatory cytokines, especially TNF-α, are critical for the pathophysiology (2, 3, 4, 5). TNF-α has a wide variety of effects on leukocytes and liver cells that support inflammation; e.g., TNF-α is one of the mediators that induce up-regulation of the β2 integrin Mac-1 (CD11b/CD18) on neutrophils during endotoxemia in vivo (6); TNF-α also primes Kupffer cells (7) and neutrophils (8) for release of cytotoxic mediators. In addition, TNF-α activates the transcription factor NF-κB4 in endothelial cells and hepatocytes during endotoxemia (9), which leads to the transcriptional activation of a number of proinflammatory genes, e.g., chemokines (10, 11), nitric oxide synthase (12, 13), and adhesion molecules such as ICAM-1 (14, 15), VCAM-1 (16), and selectins (9, 15). Therefore, TNF-α alone or in combination with IL-1 and complement is responsible for neutrophil sequestration in hepatic sinusoids during endotoxemia and sepsis (14, 17). After transmigration, neutrophils attack parenchymal cells and cause severe liver cell necrosis (14, 16, 18). Thus, TNF-α is a critical early mediator for an acute inflammatory response in the liver during endotoxemia and sepsis.

TNF-α can act through binding to two different cell surface receptors, i.e., the 55-kDa TNF-R1 and the 75-kDa TNF-R2 (19, 20). Most of the proinflammatory effects of TNF-α are mediated through TNF-R1 (20). However, TNF-α can also induce apoptosis through the death domain of TNF-R1 (21). Recently, hepatocellular apoptosis has been characterized in various models of endotoxemia (22, 23, 24, 25), and it was confirmed that this effect was mediated in vivo through TNF-R1 (25, 26). In all eukaryotic cells, the intracellular signal transduction pathway leading to apoptosis involves the activation of a cascade of cysteine proteases (caspases/ICE proteases) (27, 28, 29). Currently, there are 10 human caspases identified (30) with equivalent enzymes in the mouse (31). The activation of caspase 3 (CPP32)-like proteases in liver cells was observed during the development of apoptosis after various insults, i.e., anti-Fas Ab in vivo (32), TGF-β1 (33), staurosporine (33), and hypoxia-reoxygenation (34) in vitro. Moreover, inhibitors of caspases are highly effective in preventing apoptotic cell death (32, 33, 34). This suggests that caspase inhibitors may allow blockage of apoptotic cell injury without affecting the TNF-α-induced proinflammatory signal transduction pathway.

Although apoptotic and necrotic hepatocytes were identified during endotoxemia (14, 15, 18, 22, 23, 24, 25, 35), the pathophysiologic significance of hepatocellular apoptosis for the later development of liver cell necrosis remains unclear. Previous interventions directed against TNF-α that protected against ET-induced liver injury, e.g., neutralizing Abs, TNF receptor knockout mice, and inhibition of TNF-α gene transcription and synthesis (2, 14, 25, 26, 36, 37), inhibited both the inflammatory and the apoptotic response and did not allow a conclusion regarding the relationship between apoptotic cell death and necrosis in the liver. The beneficial effects observed with selective blockade of adhesion molecules on neutrophils (35) and liver cells (14, 16) suggested a critical role for neutrophils in the development of severe necrosis. Thus, apoptosis under these conditions may be an epiphenomenon of limited relevance or could be an important signal for neutrophil transmigration. To address this critical question, we used the Gal/ET shock model with its extensively described inflammatory (14, 16, 18, 35, 37) and apoptotic (22, 24, 26) response to characterize the role of caspase activation in hepatocellular apoptosis and, by using caspase inhibitors to selectively prevent apoptosis, to study the relevance of apoptosis for the neutrophil-induced liver cell necrosis.

Male mice, strains C3Heb/FeJ (ET-sensitive) and C3H/HeJ (ET-resistant) (20–25 g body weight), were purchased from The Jackson Laboratory (Bar Harbor, ME). The animals had free access to food (certified rodent diet no. 5002C, PMI Feeds, Richmond, IN) and water. The experimental protocols followed the criteria of Pharmacia & Upjohn (Kalamazoo, MI) and of the National Research Council for the care and use of laboratory animals in research. Animals were treated i.p. with 700 mg/kg d-Gal (Sigma Chemical Co., St. Louis, MO) and 100 μg/kg Salmonella abortus equi ET (Sigma Chemical) dissolved in sterile PBS (pH 7.0). Some animals were treated with 3 × 10 mg/kg of the caspase inhibitor Z-VAD (Enzyme Systems Products, Dublin, CA); the drug was injected 3, 4.5, and 5.5 h after Gal/ET administration. Vehicle control animals received DMSO (1 ml/kg) at the same time. Other experiments included i.v. injection of murine recombinant TNF-α (15 μg/kg; specific activity, 4 × 108 U/mg) (Genzyme, Cambridge, MA), murine rIL-1α (13 μg/kg; specific activity, 8 × 106 U/mg) (Genzyme), or murine rIL-1β (23 μg/kg; specific activity, 1.5 × 106 U/mg) (Genzyme) in Gal-sensitized animals.

The animals (5–8 per treatment group) were killed by cervical dislocation various times after administration of Gal/ET or a cytokine (TNF-α, IL-1α, IL-1β). Blood was collected from the right ventricle into a heparinized syringe and centrifuged, and plasma was used for determination of ALT activity with Sigma test kit DG 159-UV. Pieces of the liver were immediately homogenized for caspase activity measurements; other parts of each liver were frozen in liquid nitrogen and stored at −80°C for analysis of DNA fragmentation, fixed in phosphate-buffered formalin for histologic analysis, or embedded in OCT embedding medium (Miles Diagnostic Division, Elkhart, IN), and snap-frozen in methylbutane cooled in liquid nitrogen for immunohistochemistry.

Freshly excised liver was homogenized in 25 mM HEPES buffer (pH 7.5) containing 5 mM EDTA, 2 mM DTT, and 0.1% CHAPS (3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate). After centrifugation at 14,000 × g, the diluted supernatant was assayed for caspase activity using synthetic fluorogenic substrates: Ac-DEVD-MCA (Ac-Asp-Glu-Val-Asp-MCA) (Peptide Institute, Inc., Osaka, Japan) for caspase 3 (CPP32)/caspase 7 (Mch3) and Ac-YVAD-MCA (Ac-Tyr-Val-Ala-Asp-MCA) (Peptide Institute) for caspase 1 (ICE) at concentrations of 50 μM. The kinetics of the proteolytic cleavage of the substrates was monitored in a fluorescence microplate reader (Fmax; Molecular Devices, Corp., Sunnyvale, CA) using an excitation wavelength of 360 nm and an emission wavelength of 460 nm. The fluorescence intensity was calibrated with standard concentrations of MCA, and the caspase activity was calculated from the slope of the recorder trace and expressed in picomols per minute per mg of protein. Protein concentrations in the supernatant were assayed using the bicinchoninic acid kit (Sigma). For the inhibitor studies in vitro, Ac-DEVD-Ald, Ac-YVAD-Ald (Peptide Institute), or CrmA (Kamiya Biomedical Co., Seattle, WA) were added to the supernatant (3.3 nM-10 μM) 15 min before adding the substrate Ac-DEVD-MCA.

Formalin-fixed portions of the liver were embedded in paraffin and 5-μm-thick sections were cut. Neutrophils were stained by the AS-D chloroacetate esterase technique as described in detail (17). Neutrophils were identified by positive staining and morphology and were counted in 50 high power fields (×400) using a Nikon Labophot microscope (Nikon, Melville, NY). Only those neutrophils present within sinusoids or extravasated into the tissue were counted; generally, there are no neutrophils present in large hepatic vessels, e.g., venules, at that time (18). The percentage of necrotic area was estimated by evaluating parallel sections stained with hematoxylin and eosin. The pathologist (A.F.) performing the histologic evaluation (polymorphonuclear leukocytes, area of necrosis) was blinded as to the treatment of animals.

Apoptotic cells were determined using an ApopTag in situ apoptosis peroxidase detection kit (terminal deoxynucleotidyl transferase-mediated dUTP nick end labeling; TUNEL assay) (Oncor, Gaithersburg, MD) and a MicroProbe staining station (Fisher Scientific, Pittsburgh, PA). Frozen liver samples were cryosectioned (6 μm; two sections/sample, 150 μm apart) using a Leitz 1720 cryostat (W. Nuhsbaum, McHenry, IL) and assayed for direct immunoperoxidase detection of digoxigenin-labeled genomic DNA. Staining was conducted according to the manufacturer’s kit protocol for fresh frozen tissue samples. DNase 1 treatment of additional sections, to nick all DNA, served as positive controls at a concentration of 1 μg/ml determined after a preliminary dilution experiment (Appligene DNase 1, Oncor). This was prepared and conducted according to manufacturer’s instructions. TUNEL-stained mouse liver tissue sections were quantitated for apoptotic cells using computer-assisted image analysis and Optimas software (Bioscan, Edmonds, WA). Visual images from TUNEL-stained slides were captured with a digital camera attached to a microscope with modifications of previously published methods (38). A color threshold was set (dark brown for diaminobenzidine) to match visual recognition of apoptotic cells. Ten random microscopic regions of interest (each covering a surface area of 0.55 mm2) were evaluated per tissue slide (two slides/sample) and for the number of apoptotic cells captured into computer memory and downloaded into Excel 4.0 (Microsoft Co, Redmond, WA) spreadsheets for statistical analysis.

For the cell death detection ELISA (Boehringer Mannheim, Indianapolis, IN), a 20% homogenate in 50 mM sodium phosphate buffer (120 mM NaCl, 10 mM EDTA) was prepared and centrifuged at 14,000 × g. Diluted supernatant was used for the ELISA. In this test, the kinetics of product generation (Vmax) is a measure of DNA fragmentation. The Vmax values obtained for untreated controls (100%) are compared with those in treated groups. The assay allows the specific quantitation of histone-associated DNA fragments (mono- and oligonucleosomes) in the cytoplasmic fraction of cell lysates and was used extensively to demonstrate apoptosis in the liver in vivo (22, 23, 26).

Animals (n = 4 per group) were anesthetized with a ketamine mixture (225 mg/kg ketamine; 11.4 mg/kg xylazine; 2.3 mg/kg acepromazine) i.m. The liver was perfused free of blood in an open system for 5 to 10 min using an oxygenated Ca2+-free Hanks’ buffer. A collagenase-supplemented (25 mg/100 ml buffer) Hanks’ buffer was used to digest the liver. When good digestion was obtained (∼10 min), the liver was removed, minced, and strained through a tissue sieve. Cells were then centrifuged at 50 × g for 3 min. The supernatant (nonparenchymal cells) was removed and saved. The pellet (parenchymal cells) was resuspended in Hanks’ buffer and spun at 50 × g for 3 min. The supernatant was combined with the supernatant from the first spin, and the pellet was resuspended. Cell fractions were then spun at 600 × g for 10 min. The supernatants were discarded, and the nonparenchymal pellet was resuspended in pronase buffer (200 mg/50 ml buffer) and stirred for 10 min to remove any hepatocytes in the suspension. This solution was then spun at 600 × g for 10 min, and the pellet was washed once. Both cell fractions were exposed to an ammonium chloride lysing solution for 10 min to lyse contaminating RBC. Cells were washed again, resuspended, and counted. Cell fractions were >98% pure and >95% viable as judged by trypan blue exclusion. Cell concentrations were adjusted to 4 × 106 cells/ml with either caspase assay buffer or 50 mM phosphate buffer (DNA fragmentation ELISA).

All data are given as mean ± SE. Statistical significance between the control group and a treated group was determined with the unpaired Student’s t test, or Wilcoxon rank sum test. Comparisons between multiple groups were performed with one way ANOVA followed by the Bonferroni t test. p < 0.05 was considered significant.

Administration of Gal/ET caused a time-dependent increase of hepatic caspase activity using Ac-DEVD-MCA as a substrate (Fig. 1,A). In contrast, no increased protease activity was detected with the caspase 1 substrate Ac-YVAD-MCA. These results suggest the activation of proapoptotic caspase 3-like proteases during endotoxemia in Gal-sensitized animals. Parallel to the increase of caspase 3-like protease activity, there was enhanced DNA fragmentation, indicating cells undergoing apoptosis at the same time (Fig. 1B). Neither Gal nor ET alone caused an increase in hepatic caspase activity or DNA fragmentation (data not shown). Apoptosis in the liver of Gal/ET-treated animals was confirmed using the TUNEL assay (Fig. 2). Quantitative analysis showed few apoptotic cells in controls (about 0.2% of hepatocytes) and no significant change up to 4 h after Gal/ET treatment (Fig. 3). However, at 5 and 6 h, the number of apoptotic cells increased dramatically reaching 13% at 6 h (65-fold increase). These data indicate that the increase of proapoptotic caspase activity correlated with the increase of apoptotic cell death in this model. Because the intact tissue did not allow us to distinguish which liver cell types had been affected, parenchymal cells (hepatocytes) and nonparenchymal cells (mixture of Kupffer cells and endothelial cells) were isolated 5.5 h after Gal-ET administration. Measurement of DEVD-MCA caspase activity and DNA fragmentation showed significant increases only in hepatocytes but not in the nonparenchymal cell fraction (Fig. 4).

FIGURE 1.

Time course of hepatic caspase activity (A) and DNA fragmentation (B) as an indicator for apoptosis after i.p. administration of 700 mg/kg Gal and 100 μg/kg S. abortus equi endotoxin (Gal/ET). Caspase activity was determined by measuring the proteolytic cleavage of the caspase 1 substrate Ac-YVAD-MCA and the caspase 3 substrate Ac-DEVD-MCA. 100% values are 30.6 ± 0.9 pmol MCA/min/mg protein (DEVD) and 118.2 ± 5.0 pmol MCA/min/mg protein (YVAD). DNA fragmentation was measured by ELISA; values are reported as % changes of Vmax values obtained from controls (=100%). Data represent means ± SE of six to eight animals per time point. n.d., not determined in tissue with severe hemorrhage because of interferences with the assay. *, p < 0.05 (compared with t = 0).

FIGURE 1.

Time course of hepatic caspase activity (A) and DNA fragmentation (B) as an indicator for apoptosis after i.p. administration of 700 mg/kg Gal and 100 μg/kg S. abortus equi endotoxin (Gal/ET). Caspase activity was determined by measuring the proteolytic cleavage of the caspase 1 substrate Ac-YVAD-MCA and the caspase 3 substrate Ac-DEVD-MCA. 100% values are 30.6 ± 0.9 pmol MCA/min/mg protein (DEVD) and 118.2 ± 5.0 pmol MCA/min/mg protein (YVAD). DNA fragmentation was measured by ELISA; values are reported as % changes of Vmax values obtained from controls (=100%). Data represent means ± SE of six to eight animals per time point. n.d., not determined in tissue with severe hemorrhage because of interferences with the assay. *, p < 0.05 (compared with t = 0).

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FIGURE 2.

Detection of hepatocellular DNA fragmentation using the TUNEL method as described in Materials and Methods. Arrows indicate apoptotic cells. A, Untreated control; B, liver section of an ET-sensitive mouse 6 h after treatment with Gal/ET; C, liver section of an ET-resistant mouse 6 h after Gal/ET treatment; D, a liver section that was pretreated with DNase 1 to nick all DNA, served as positive control. All samples: ×312.5.

FIGURE 2.

Detection of hepatocellular DNA fragmentation using the TUNEL method as described in Materials and Methods. Arrows indicate apoptotic cells. A, Untreated control; B, liver section of an ET-sensitive mouse 6 h after treatment with Gal/ET; C, liver section of an ET-resistant mouse 6 h after Gal/ET treatment; D, a liver section that was pretreated with DNase 1 to nick all DNA, served as positive control. All samples: ×312.5.

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FIGURE 3.

Quantitative analysis of apoptotic cells. TUNEL-stained mouse liver tissue sections were quantitated for apoptotic cells using computer-assisted image analysis as described in Materials and Methods. Cells were counted in 20 random fields (×125) of 2 slides from each liver and expressed as apoptotic cells per field. Data represent means ± SE of six to eight animals. *, p < 0.05 (compared with t = 0).

FIGURE 3.

Quantitative analysis of apoptotic cells. TUNEL-stained mouse liver tissue sections were quantitated for apoptotic cells using computer-assisted image analysis as described in Materials and Methods. Cells were counted in 20 random fields (×125) of 2 slides from each liver and expressed as apoptotic cells per field. Data represent means ± SE of six to eight animals. *, p < 0.05 (compared with t = 0).

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FIGURE 4.

Caspase activity and DNA fragmentation in parenchymal cells (hepatocytes) and nonparenchymal cells in untreated controls and 6 h after i.p. administration of 700 mg/kg Gal and 100 μg/kg S. abortus equi endotoxin (Gal/ET). Caspase activity was determined by measuring the proteolytic cleavage of the caspase 3 substrate Ac-DEVD-MCA. 100% value: 24.6 ± 4.5 pmol of MCA/min/mg protein (DEVD). DNA fragmentation was measured by ELISA; values are reported as percent changes of vmax obtained from controls (=100%). Data represent means ± SE of four animals per group. *, p < 0.05 (compared with control).

FIGURE 4.

Caspase activity and DNA fragmentation in parenchymal cells (hepatocytes) and nonparenchymal cells in untreated controls and 6 h after i.p. administration of 700 mg/kg Gal and 100 μg/kg S. abortus equi endotoxin (Gal/ET). Caspase activity was determined by measuring the proteolytic cleavage of the caspase 3 substrate Ac-DEVD-MCA. 100% value: 24.6 ± 4.5 pmol of MCA/min/mg protein (DEVD). DNA fragmentation was measured by ELISA; values are reported as percent changes of vmax obtained from controls (=100%). Data represent means ± SE of four animals per group. *, p < 0.05 (compared with control).

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To further characterize the type of caspase involved, inhibitor studies were performed using liver homogenate of Gal/ET-treated animals (6 h). The inhibitor of caspase 3-like proteases, Ac-DEVD-Ald, dose dependently inhibited the enzyme activity in the liver (Fig. 5). An IC50 of 16.75 nM was calculated from these data. In contrast, the caspase 1 inhibitor Ac-YVAD-Ald or CrmA, an effective inhibitor of caspase 1 and the proapoptotic caspase 8 had no or very limited effects on Gal/ET-induced protease activity in the liver. The extrapolated IC50 for YVAD-Ald was 19.6 μM.

FIGURE 5.

Dose-dependent inhibition of caspase activity (Ac-DEVD-MCA) in liver homogenate derived from animals treated with 700 mg/kg Gal/100 μg/kg ET for 6 h. Inhibitors (CrmA, Ac-DEVD-Ald, or Ac-YVAD-Ald) were added to the supernatant 15 min before the substrate.

FIGURE 5.

Dose-dependent inhibition of caspase activity (Ac-DEVD-MCA) in liver homogenate derived from animals treated with 700 mg/kg Gal/100 μg/kg ET for 6 h. Inhibitors (CrmA, Ac-DEVD-Ald, or Ac-YVAD-Ald) were added to the supernatant 15 min before the substrate.

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To identify the critical mediators involved in the activation of caspases in vivo, ET-resistant animals, which do not generate cytokines in response to ET, were treated with Gal/ET. In contrast to the sensitive animals, ET-resistant animals did not show enhanced caspase 3-like protease activity or evidence for increased DNA fragmentation (Table I). Injection of TNF-α, IL-1α, or IL-1β in combination with Gal indicated that only Gal/TNF-α was able to induce an increase in caspase 3-like protease activity and apoptosis in ET-resistant (Table I) and in ET-sensitive animals (data not shown). These findings suggest that ET-induced proapoptotic caspase activation and apoptotic cell death in hepatocytes is mediated by TNF-α.

Table I.

Caspase activation and apoptosis in ET-resistant animalsa

DEVD (pmol/min/mg protein)YVAD (pmol/min/mg protein)DNA Fragmentation Vmax (%)
Controls 28.7 ± 3.0 86.2 ± 4.3 100 ± 5 
Gal/ET 25.5 ± 4.7 100.6 ± 5.5 78 ± 13 
Gal/TNF-α 190.3 ± 44.5* 92.1 ± 2.7 410 ± 5* 
Gal/IL-1α 34.1 ± 4.5 99.6 ± 6.6 88 ± 13 
Gal/IL-1β 34.6 ± 6.0 101.0 ± 4.1 83 ± 19 
DEVD (pmol/min/mg protein)YVAD (pmol/min/mg protein)DNA Fragmentation Vmax (%)
Controls 28.7 ± 3.0 86.2 ± 4.3 100 ± 5 
Gal/ET 25.5 ± 4.7 100.6 ± 5.5 78 ± 13 
Gal/TNF-α 190.3 ± 44.5* 92.1 ± 2.7 410 ± 5* 
Gal/IL-1α 34.1 ± 4.5 99.6 ± 6.6 88 ± 13 
Gal/IL-1β 34.6 ± 6.0 101.0 ± 4.1 83 ± 19 
a

Hepatic caspase activity and DNA fragmentation (% of control) as indicator for apoptosis were determined in C3H/HeJ (ET-resistant) mice 5 h after administration of 700 mg/kg Gal in combination with 100 μg/kg S. abortus equi ET (Gal/ET), 15 μg/kg murine recombinant TNF-α (Gal/TNF-α), 13 μg/kg murine rIL-1α (Gal/IL-1α), or 23 μg/kg murine rIL-1β (Gal/IL-1β). DNA fragmentation was measured by ELISA; values are reported as % changes of Vmax obtained from controls (=100%). Caspase activity was determined by measuring the proteolytic cleavage of the caspase 1 substrate Ac-YVAD-MCA and the caspase 3 substrate Ac-DEVD-MCA. Data represent means ± SE of five animals per group. *p < 0.05 (compared with controls).

Because we previously provided evidence that neutrophils are critical for ET-induced hepatocellular necrosis in this model, we tested the hypothesis that caspase activation and apoptosis are relevant for the inflammatory response. A large group of animals was treated with Gal/ET. After 3 h, i.e., at a time when TNF-α has been generated and TNF-α-mediated inflammatory responses were initiated (NF-κB activation, transcription of adhesion molecules) (2, 6, 9, 14, 16, 18), animals either were left untreated (disease control) or were injected with the caspase inhibitor Z-VAD in DMSO (10 mg/kg) or with DMSO alone (1 ml/kg). Z-VAD or vehicle treatment was repeated twice at 90-min intervals. Gal/ET administration caused severe liver injury at 7 h as indicated by a significant increase in plasma ALT activities and widespread hepatocellular necrosis (45% of hepatocytes) (Fig. 6,A). Z-VAD, in contrast to the vehicle DMSO, effectively inhibited liver injury. Necrotic cell injury was almost eliminated; i.e., only few individual hepatocytes could be identified as necrotic. In contrast, Z-VAD did not reduce the overall number of neutrophils in the liver (Fig. 6,B). However, virtually all neutrophils (>95%) remained localized within sinusoids. In the vehicle group, 30 to 35% of neutrophils were extravasated. To demonstrate the effect of Z-VAD treatment on caspase activity and apoptosis, the experiment was repeated, and the animals were killed at 6 h (before the onset of massive hemorrhage and necrosis). Z-VAD completely inhibited caspase 3-like protease activity in the liver but had no effect on the baseline caspase 1 activity (Fig. 7). Z-VAD treatment also significantly reduced the increase in apoptotic cell death as determined by two independent criteria, DNA fragmentation (81% inhibition) and TUNEL assay (88% reduction of apoptotic cells) (Table II).

FIGURE 6.

Liver injury, as assessed by plasma ALT activities and histologically by the area of necrosis (A), and hepatic neutrophil sequestration (B) were evaluated in C3Heb/FeJ mice before (controls, C) and 7 h after the combined administration of 700 mg/kg Gal and 100 μg/kg S. abortus equi endotoxin (G/E). Some animals received 3 × 10 mg/kg of the caspase inhibitor Z-VAD; the drug was injected 3, 4.5, and 5.5 h after G/E administration. Vehicle control animals received DMSO (1 ml/kg) at the same time. Data represent means ± SE of eight animals per group. *, p < 0.05 (compared with control); #, p < 0.05 (ZVAD vs DMSO).

FIGURE 6.

Liver injury, as assessed by plasma ALT activities and histologically by the area of necrosis (A), and hepatic neutrophil sequestration (B) were evaluated in C3Heb/FeJ mice before (controls, C) and 7 h after the combined administration of 700 mg/kg Gal and 100 μg/kg S. abortus equi endotoxin (G/E). Some animals received 3 × 10 mg/kg of the caspase inhibitor Z-VAD; the drug was injected 3, 4.5, and 5.5 h after G/E administration. Vehicle control animals received DMSO (1 ml/kg) at the same time. Data represent means ± SE of eight animals per group. *, p < 0.05 (compared with control); #, p < 0.05 (ZVAD vs DMSO).

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FIGURE 7.

Increase in hepatic caspase activity before (controls, C) and 6 h after i.p. administration of 700 mg/kg Gal and 100 μg/kg S. abortus equi endotoxin (G/E). Caspase activity was determined by measuring the proteolytic cleavage of the caspase 1 substrate Ac-YVAD-MCA and the caspase 3 substrate Ac-DEVD-MCA. 100% values are 36.9 ± 3.1 pmol of MCA/min/mg protein (DEVD) and 72.5 ± 4.8 pmol of MCA/min/mg protein (YVAD). Some animals were treated with 3 × 10 mg/kg of the caspase inhibitor Z-VAD; the drug was injected 3, 4.5, and 5.5 h after G/E administration. Vehicle control animals received DMSO (1 ml/kg) at the same time. Data represent mean ± SE of six animals per group. *, p < 0.05 (compared with control); #, p < 0.05 (ZVAD vs DMSO).

FIGURE 7.

Increase in hepatic caspase activity before (controls, C) and 6 h after i.p. administration of 700 mg/kg Gal and 100 μg/kg S. abortus equi endotoxin (G/E). Caspase activity was determined by measuring the proteolytic cleavage of the caspase 1 substrate Ac-YVAD-MCA and the caspase 3 substrate Ac-DEVD-MCA. 100% values are 36.9 ± 3.1 pmol of MCA/min/mg protein (DEVD) and 72.5 ± 4.8 pmol of MCA/min/mg protein (YVAD). Some animals were treated with 3 × 10 mg/kg of the caspase inhibitor Z-VAD; the drug was injected 3, 4.5, and 5.5 h after G/E administration. Vehicle control animals received DMSO (1 ml/kg) at the same time. Data represent mean ± SE of six animals per group. *, p < 0.05 (compared with control); #, p < 0.05 (ZVAD vs DMSO).

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Table II.

Effect of the caspase inhibitor Z-VAD on Gal/ET-induced apoptosisa

DNA Fragmentation (Vmax (%))TUNEL Assay (Apoptotic Cells/Field)
Controls 100 ± 13 2.6 ± 0.3 
Gal/ET 489 ± 7* 165.5 ± 35.2* 
Gal/ET+ DMSO 515 ± 11* 197.8 ± 44.5* 
Gal/ET+ Z-VAD 177 ± 16*† 24.0 ± 1.2*† 
DNA Fragmentation (Vmax (%))TUNEL Assay (Apoptotic Cells/Field)
Controls 100 ± 13 2.6 ± 0.3 
Gal/ET 489 ± 7* 165.5 ± 35.2* 
Gal/ET+ DMSO 515 ± 11* 197.8 ± 44.5* 
Gal/ET+ Z-VAD 177 ± 16*† 24.0 ± 1.2*† 
a

Evaluation of apoptotic cell injury by DNA fragmentation and the TUNEL assay before (Controls) and 6 h after i.p. administration of 700 mg/kg Gal and 100 μg/kg S. abortus equi ET (Gal/ET). Some animals were treated with 3 × 10 mg/kg of the caspase inhibitor Z-VAD (Z-Val-Ala-Asp-CH2F); the drug was injected 3, 4.5, and 5.5 h after Gal/ET administration. Vehicle control animals received DMSO (1 ml/kg) at the same time. TUNEL-stained mouse liver tissue sections were quantitated for apoptotic cells using computer-assisted image analysis as described in Materials and Methods. Cells were counted in 20 random fields (×125) of 2 slides from each liver and expressed as apoptotic cells per field. DNA fragmentation was measured by ELISA; values are reported as % changes of Vmax obtained from controls (=100%).

Data represent means ± SE of six animals. *p < 0.05 (compared with controls); †p < 0.05 (compared with Gal/ET or Gal/ET + DMSO).

The principal objective of this investigation was to study the role of caspase activation in hepatocellular apoptosis and neutrophil-induced necrosis during ET-mediated liver injury in vivo. Our data demonstrated a substantial increase of caspase activity with DEVD-MCA, a substrate of caspase 3-like proteases.

However, no increase of activity was seen with YVAD-MCA, a substrate more specific for the caspase 1 subfamily. Furthermore, the enhanced caspase activity could be inhibited by DEVD-Ald but not by YVAD-Ald or CrmA. These data suggest that in the Gal/ET model in vivo, there is a predominant activation of the caspase 3-like subfamily of proteases. Caspase activation correlated with development of hepatocellular apoptosis in vivo. These results are consistent with critical involvement of caspase 3 subfamily members in apoptosis in a variety of cell types (27, 28, 29). The effect of Gal/ET treatment on hepatic caspase activity was not observed in ET-resistant animals that do not generate cytokines in response to ET. Furthermore, caspase activation could be induced with TNF-α but not with IL-1 in ET-resistant and -sensitive mice. These findings strongly indicate that Gal/ET-induced activation of caspase 3-like proteases in vivo and hepatocellular apoptosis is mediated by TNF-α. Other stimuli that induced caspase activation in hepatocytes, e.g., Fas Ab (32), TGF-β1 (33), and hypoxia-reoxygenation (34), showed increased protease activity with DEVD-MCA and YVAD-MCA as substrates. Inhibitors of caspase 1 (YVAD-Ald, YVAD-cmk) and caspase 3 (DEVD-Ald, Z-VAD) inhibited hepatocyte apoptosis in vitro (34, 39) and after Fas Ab administration in vivo (32, 39). On the basis of these data, Rouquet et al. (40) suggested that at least two distinct pathways of Fas signaling exist in hepatocytes. Activation of caspase 1- and caspase 3-like proteases are required, but these pathways involve different subclasses of serine proteases and can be selectively modulated by protein tyrosine kinase inhibitors (40). Interestingly, actinomycin/TNF-α-induced apoptosis in isolated hepatocytes and in vivo could be inhibited by YVAD-cmk (39). In contrast, our data indicated that Gal/TNF-α-mediated apoptosis correlated only with increased caspase 3-like activity and could only be inhibited with predominantly caspase 3 inhibitors, e.g., DEVD-Ald. This would suggest that there might also be multiple pathways for TNF-α signaling of apoptosis.

Hepatocellular apoptosis has been described in several models of ET shock (22, 23, 24, 25, 26); however, the importance of apoptosis for the overall injury and organ failure in these models remained unclear. Quantitative analysis of apoptotic and necrotic parenchymal cells showed that ∼13% of hepatocytes could be identified as undergoing apoptosis and only <10% of the cells were necrotic at 6 h after Gal/ET treatment. One h later, 38 to 45% of hepatocytes were necrotic. These data indicate that quantitively, apoptotic cell death could not explain the three- to fourfold higher number of necrotic cells 1 h later. Thus, apoptosis appears to be a trigger mechanism for the more severe attack of neutrophils. This hypothesis was investigated by the use of caspase inhibitors, which allowed the selective blockage of apoptosis. Administration of Z-VAD in vivo inhibited caspase 3-like activity (DEVD-MCA) but not caspase 1 activity (YVAD-MCA) and protected effectively against Fas-mediated apoptosis (32). Furthermore, Z-VAD blocked apoptosis (TGF-β1, staurosporine) but not necrosis (staurosporine) in isolated hepatocytes (41). Kinetic analysis in hepatocyte lysate showed that Z-VAD is substantially less potent than DEVD-Ald in inhibiting DEVD-AFC cleavage (33). However, similar inhibition curves indicated that the mechanism of action was the same; i.e., both inhibitors acted as suicide substrates for caspase 3 (33). YVAD-Ald acted as a suicide substrate for caspase 1 but was a competitive inhibitor for caspase 3 with a very high Ki of 12.6 μM (33). Our data agree with these findings. Whereas DEVD-Ald was a highly potent inhibitor (IC50 of 16.75 nM) for increased caspase activity in the liver homogenate, YVAD-Ald inhibited this activity with an IC50 of ∼19.6 μM. Because Z-VAD completely prevented the increase in hepatic caspase activity and apoptosis, it further supports the conclusion that caspase 3-like proteases are involved in hepatic parenchymal cell apoptosis in Gal/ET-treated animals.

Z-VAD treatment not only prevented caspase activation and apoptosis but also suppressed liver cell necrosis. Previous studies showed that parenchymal cell necrosis could be attenuated by antibodies against β2 integrins on neutrophils (35), ICAM-1 on sinusoidal endothelial cells and hepatocytes (14) as well as VCAM-1 on sinusoidal lining cells (16). Neutrophil transmigration in hepatic sinusoids has been identified as a critical step for neutrophil-mediated injury in this model (18). Antibodies to ICAM-1 or VCAM-1 blocked neutrophil extravasation and therefore prevented liver cell necrosis (14, 16). These data suggest that neutrophils are essential for the development of hepatocellular necrosis in this model. Thus, TNF-α initiates two separate responses in the liver, an apoptotic response and an inflammatory response. Treatment with Z-VAD was started at 3 h after Gal/ET administration. Since formation of TNF-α (2, 14), neutrophil sequestration in the hepatic vasculature (18, 35), activation of NF-κB (9, 37), mRNA formation of ICAM-1 (14, 15), VCAM-1 (16), and selectins (15), and even in part adhesion molecule protein synthesis (16, 42) already occurs before the 3-h time point, Z-VAD could have not affected the proinflammatory response in the liver. The only major events of the inflammatory response that occur after 3 h are neutrophil transmigration and the adherence-dependent cytotoxicity against hepatocytes. The fact that preventing apoptosis in hepatocytes suppresses neutrophil transmigration suggests that hepatocytes undergoing apoptosis represent a signal for neutrophil migration and attack on parenchymal cells. Generally, apoptosis is considered a physiologic way to remove unwanted cells without generating an inflammatory response (43, 44). In contrast, our data suggest that apoptosis may be able to aggravate an inflammatory response. How can these opposite viewpoints be reconciled? First of all, our data do not argue against the fact that single-cell apoptosis will not trigger an inflammatory response. Our data suggest that in the presence of activated and primed neutrophils in hepatic sinusoids, a large number of parenchymal cells undergoing apoptosis at the same time can represent a stimulus for these leukocytes to transmigrate and attack. Recently (45), it was reported that infection of hepatocytes with Listeria monocytogenes causes apoptosis and the generation of a neutrophil chemotactic factor. However, it was unclear whether the increased neutrophil chemotaxis was actually dependent on apoptosis or was more related to Listeria infection. Nevertheless, this observation is consistent with the presence of neutrophils around infected hepatocytes in vivo (45). Neutropenia experiments in this model indicate that a major function of neutrophils is the removal of infected hepatocytes undergoing apoptosis (45). In the Gal/ET model, triggering the transmigration of primed neutrophils may have a similar purpose, and this may be a general mechanism for removal of a large number of apoptotic cells. However, as demonstrated after Gal/ET treatment, the further activation of neutrophils in the liver vasculature bears the risk of additional damage to healthy tissue. Further studies are necessary to identify the nature of the chemotactic stimulus generated by apoptotic parenchymal cells.

In summary, a selective activation of caspase 3-like proteases was observed in the liver that correlated with the development of apoptosis in parenchymal cells after Gal/ET treatment. Caspase activation and apoptosis during endotoxemia in vivo was mediated by TNF-α. Injection of Z-VAD, an effective inhibitor of the caspase 3 subfamily, prevented caspase activation, apoptosis, and liver cell necrosis. Z-VAD did not affect neutrophil sequestration in sinusoids but inhibited transmigration. These data indicate that activation of the caspase 3 subfamily is critical for the development of parenchymal cell apoptosis. In addition, excessive hepatocellular apoptosis can be an important signal for transmigration of primed neutrophils sequestered in sinusoids. Thus, proapoptotic caspases may be a promising therapeutic target for ET- and sepsis-related liver failure.

1

The work was supported in part by National Institutes of Health Grant RO1 ES 06091.

4

Abbreviations used in this paper: NF-κB, nuclear factor κB; ET, endotoxin; Gal, galactosamine; ALT, alanine aminotransferase; TUNEL, terminal deoxynucleotidyl transferase-mediated dUTP nick end labeling; ICE, IL-1β-converting enzyme; Z-VAD, Z-Val-Ala-Asp-CH2F; Ac-DEVD-MCA, acetyl-Asp-Glu-Val-Asp-(4-methylcoumaryl-7-amide); Ac-YVAD-MCA, acetyl-Tyr-Val-Ala-Asp-(4-methylcoumaryl-7-amide); Ac-DEVD-Ald, acetyl-Asp-Glu-Val-Asp-CHO); Ac-YVAD-Ald, acetyl-Tyr-Val-Ala-Asp-CHO; CrmA, cowpox viral serpin cytokine response modifier A; Ac-YVAD-cmk, acetyl-Tyr-Val-Ala-Asp-chloromethylketone.

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