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Address correspondence to Normand Marceau, Centre de recherche HDQ (CHUQ), 9 McMahon, Québec, Canada, G1R 2J6. Tel.: (418) 691-5559. Fax: (418) 691-5439. E-mail: normand.marceau{at}crhdq.ulaval.ca
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Abstract |
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Key Words: keratin; Fas; Golgi; microtubules; hepatocyte
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Introduction |
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Apoptosis, a form of programmed cell death that generates a minimal inflammatory response, is required to maintain normal tissue homeostasis or to eliminate abnormal cells or cells damaged by various insults (Thompson, 1995; Peter and Krammer, 1998). Extensive in vitro studies on the role of three death-inducing members of the TNF family, TNF-, Fas ligand (FasL), and TNF-related apoptosis-inducing ligand (TRAIL), using a wide range of conventional cell lines as model systems have helped to elucidate the complex but interacting signaling pathways that underlie apoptosis (Ashkenazi and Dixit, 1998; Krammer, 1999). In particular, stimulation of Fas by FasL induces receptor trimerization at the surface membrane, which allows the recruitment of the adapter protein FADD which, with the initiator procaspase-8, forms the death-inducing signaling complex (Kischkel et al., 1995; Chinnaiyan et al., 1996). This triggers procaspase-8 proteolytic activation, which in turn initiates in type I cells a controlled proteolysis of proteins including effector procaspases like procaspase-3 and various downstream substrates (Caulin et al., 1997; Zheng et al., 1998; Hengartner, 2000). Fas stimulation can trigger a second caspase pathway that is mediated in type II cells by the release of cytochrome c from mitochondria and the subsequent activation of the initiator procaspase-9 (Hengartner, 2000; Kaufmann and Gores, 2000; Krammer, 2000). In contrast to FasL, TNF-
activates two receptors, TNF-R1 and TNF-R2 (Baker and Reddy, 1998; Ashkenazi and Dixit, 1999), which promote cell proliferation after TRADD-mediated binding to an appropriate member of the TRAF family. However, in cells sensitized with CHX or actinomycin D (Act D), oligomerized TNF-R1, but not TNF-R2 which lacks the death domain, can lead to apoptosis after binding to TRADD, which recruits FADD to activate the caspase pathway (Chinnaiyan et al., 1996; Hsu et al., 1996). Stimulation of TRAIL receptors, i.e., TRAIL-R1 and TRAIL-R2, can in the presence of CHX lead to apoptosis via a sequence of events that correspond to those triggered by Fas (Kischkel et al., 2000; Sprick et al., 2000). In various cell lines, CHX and Act D sensitize for Fas-induced apoptosis by downregulating the synthesis of c-FLIP, a labile protective protein homologous to caspase-8 but exhibiting an inactive catalytic site, so that the balance between cell survival and death can be modulated by the relative concentration of the death receptor and c-FLIP (Tschopp et al., 1998; Scaffidi et al., 1999; Fulda et al., 2000). However, cell survival can also be obtained through the action of external signals, like those generated by epidermal growth factor (EGF). In that case, protection against cell death results from the activation of the Akt pathway (Kulik et al., 1997; Gibson et al., 1999). The relative contribution of these pathways is particularly dependent on the cell type and the cellular context (Kaufmann and Gores, 2000; Tepper et al., 2000).
Hepatocytes are among the cell types that contain the highest level of Fas (Nagata, 1999). Accordingly, a single injection of the agonistic antibody of FasL Jo2 into mice is sufficient to induce massive hepatocyte apoptosis and rapid death of the animal (Ni et al., 1994; Lacronique et al., 1996). This high sensitivity to Fas-mediated apoptosis results from the fact that the Fas/FasL system provides an efficient means to exclude from the liver the hepatocytes that have been damaged by various insults (Lacronique et al., 1996; Nagata, 1999; Sodeman et al., 2000). Upon stimulation, Fas is targeted to the surface membrane through the Golgi-sorting compartment, and this transfer depends on functional microtubules (Feng and Kaplowitz, 2000). The current view is that this microtubule-dependent targeting of Fas provides an efficient mechanism to modulate the Fas density at the surface membrane and to avoid any spontaneous apoptosis that could result from an excess of cell surface Fas.
In the work reported here, we investigate the role of K8/K18 in regulating Fas-mediated apoptosis in mouse hepatocytes in primary culture and in vivo. The results show that Jo2 stimulation of Fas generates a higher apoptotic response in K8-null than in wild-type (WT) mouse hepatocytes that is directly associated with a higher and more rapid activation of its signaling pathway. In addition, analyses of Fas trafficking in hepatocytes, treated or non-treated with the microtubule-disrupting agent colchicine, demonstrate that the higher K8-null sensitivity to Jo2 is associated with increased Fas targeting to the surface membrane. This points to a major role of simple epithelium keratins in regulating Fas trafficking and thus apoptosis sensitivity.
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Results |
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K8/K18 interfere selectively with Fas-mediated cell death
Apoptosis can be induced by other members of the TNF receptor family, such as those activated by TNF- and TRAIL, respectively. We thus evaluated whether the protective role played by K8/K18 against Fas-mediated hepatocyte death was also applicable to those death receptors. As shown in Fig. 2
A, the addition of TNF-
to primary cultured hepatocytes induced a low level of apoptosis, but no significant difference was observed between WT and K8-null hepatocytes. Similarly, the addition of TRAIL yielded a low apoptotic response of WT hepatocytes on which the loss of K8/K18 had no influence (Fig. 2 A). This lack of response for hepatocytes in primary culture is consistent with previous findings obtained in various established cell lines (Jo et al., 2000).
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We next looked at the activation of caspase-3, a representative member of the Fas-signaling pathway, during the 24-h period after Jo2 stimulation of cultured hepatocytes. The increased sensitivity of K8-null hepatocytes to Fas-mediated apoptosis over that of WT hepatocytes was associated with a more prominent processing of caspase-3, as revealed by an increased generation of the p17 fragment (Fig. 2 B). The activation started at 2 h and reached a maximum already at 4 h in K8-null hepatocytes, whereas it started at 4 h and gradually increased during the following 24-h period in WT hepatocytes. Furthermore, assessment of the hepatocyte's sensitivity to Jo2 by DNA laddering showed a higher and more rapid DNA fragmentation in K8-null than in WT hepatocytes (Fig. 2 C). This corroborates with the data obtained by direct counting of fragmented nuclei.
Another pathway that can lead to apoptosis in hepatocytes is that activated by TGF-ß (Roberts et al., 2000). We thus assessed whether the loss of K8/K18 altered the response of hepatocytes to this multifunctional factor. Although its addition yielded a low level of apoptosis, it did not lead to a differential response between K8-null and WT hepatocytes, which further supports the high specificity of the interplay between K8/K18 and Fas-mediated apoptosis (not shown).
Cells can be protected from Fas-mediated apoptosis via the activation of survival pathways by growth factors like EGF (Gibson et al., 1999). Considering the major role of EGF in hepatocyte growth regulation, we asked whether it is also involved in the protective resistance provided by K8/K18 in these cells. Notably, the growth-promoting activity occurs via the activation of the ERK-signaling pathway, whereas the protective effect takes place through the activation of the Akt-signaling pathway (Gibson et al., 1999; Roberts et al., 2000). With regard to growth-promoting activity of EGF, we found that, although the ERK pathway was already primed in K8-null hepatocytes and little in WT hepatocytes, the addition of EGF led to essentially equivalent increases in ERK activation in both cell populations (Fig. 3 A). Comparable increases in Akt phosphorylation levels were observed in K8-null and WT hepatocytes after the addition of either 20 or 200 ng/dish EGF (Fig. 3 A), indicating that the loss of K8/K18 does not affect this EGF protection against Fas-mediated apoptosis. In line with these data on EGF-induced Akt activation, increasing the EGF concentration to 200 ng/dish was sufficient to decrease the Fas-induced apoptosis of K8-null hepatocytes to a level comparable to that of WT hepatocytes (Fig. 3 B).
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Discussion |
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K8/K18 do not interfere with the EGF-induced survival
Previous data have established that a EGF-induced protection normally occurs in type II cells, in which Fas-mediated cell death is dependent on mitochondria (Krammer, 2000). Hepatocytes resemble type II cells, and as shown in Fig. 7, the EGF-induced survival results from the activation of the Akt-signaling pathway, which in turn blocks the mitochondrial-dependent pathway at a step that regulates the initiator caspase-9 (Wennstrom and Downward, 1999; Krammer, 2000; Roberts et al., 2000). The present results show that the addition of a beneficial EGF dose activates the Akt-signaling pathway to a level that is essentially identical for both K8-null and WT hepatocytes, indicating that the EGF-induced survival against Fas-mediated apoptosis is essentially independent of the resistance provided by K8/K18. In the same way, the extent of Akt activation provided by a EGF dose of 200 ng/dish is high enough to reduce the Fas-mediated apoptosis of K8-null hepatocytes to the level obtained in WT hepatocytes. These findings are consistent with our immunolocalization results, suggesting that the resistance provided by K8/K18 takes place instead at a microtubule-dependent step that selectively affects Fas density at the hepatocyte surface.
K8/K18 modulate Fas density at the surface membrane
Our data show that both K8/K18 and Fas are localized at the surface membrane of WT hepatocytes. However, although the surface membrane is a major targeting site for the keratins in WT hepatocytes, a large fraction of Fas is present in the Golgi compartment before activation (Bennett et al., 1998; Sodeman et al., 2000). Since Fas activation implies a microtubule-dependent transfer from the Golgi-sorting compartment to the surface membrane, we propose that the role of K8/K18 in Fas-mediated apoptosis is to regulate Fas targeting to the surface membrane. This interpretation is supported by the observation that a loss of K8/K18 leads to an increased density of Fas at the membrane (Figs. 4 A and 5 A). Furthermore, as depicted in Fig. 7, recent data in other cell types suggest that the intensity of the Fas activation is dependent not only on its density at the surface but also on its degree of clustering, due to a receptor displacement in the surface membrane plane that is driven by an actin-dependent process (Parlato et al., 2000). Assuming that this association holds in hepatocytes, the data reported in Fig. 6 suggest that in K8-null hepatocytes, a depolymerization of microtubules can still allow the transfer of Fas to the cell surface. Although the nature of the interplay between Fas, K8/K18, microtubules and fibrillar actin is unclear, we believe that it involves the participation of plectin (Fig. 7), an integrator protein that is capable of mediating dynamic interactions between IFs, fibrillar actin, and microtubules (Herrmann and Aebi, 2000). In this regard, it is worth noticing that Fas-dependent activation of caspase-8 leads to a very early specific cleavage of plectin (Stegh et al., 2000), an event which may be part of the K8/K18 down-modulation process of Fas targeting.
Previous studies on apoptosis induced in cultured mouse hepatocytes by various ligands such as TNF- and TGF-ß have revealed a saturation of the death response at high dose (Sanchez et al., 1996; Senaldi et al., 1998). The present results demonstrate a similar saturation at Jo2 concentrations above 2.5 µg/ml. Jo2 stimulation of Fas-mediated apoptosis requires the trimerization of the receptor but, as discussed above, the resulting activation is also dependent on the receptor density and clustering. It is thus possible that at a high Jo2 dose the level of Fas trimerization becomes saturated in a manner that is independent of K8/K18 modulation of the Fas density at the surface.
Biological significance of the K8/K18 IF and Fas interplay
Fas-mediated apoptosis is an efficient process by which damaged hepatocytes are excluded from the liver (Kanzler and Galle, 2000). The present results on the response of primary cultured K8-null versus WT hepatocytes to Fas, TNF-, or TRAIL show that K8/K18 IFs provide resistance only against Fas-mediated apoptosis. The significance of these in vitro findings is well supported by our complementary in vivo data demonstrating a large difference in ALT release between K8-null and WT hepatocytes in response to Jo2 injection. This is also consistent with the in vivo observation reported by others (Caulin et al., 2000) on the differential response of K8-null versus WT hepatocytes to the damaging agent concanavalin-A, a strong inducer of hepatitis in mice which has the ability to sensitize hepatocytes to Fas L and TNF-
mediated apoptosis (Kusters et al., 1997; Ksontini et al., 1998; Tagawa et al., 1998, 1997). At any rate, hepatocytes are the only epithelial cells where the IFs are made solely of K8/K18, and we propose that the close interplay between Fas and K8/K18 is linked to the hepatocyte differentiation program. Moreover, since the K8/K18 pair is present in all simple epithelial cells, we believe that this new information on their role in Fas-mediated apoptosis of hepatocytes applies to cells of other simple epithelium origins.
The K8-null mutation provides an extreme alteration of the usual K8/K18 IF network. We have shown previously that the reinsertion of the WT K8 gene into K8-null mice rescues the mechanical integrity of the hepatocyte surface membrane, implying that the null mutation exerts its action directly (Loranger et al., 1997). In the same way, the present data show that the transfer of a complete human K8 cDNA using a retroviral vector rescues the K8/K18 resistance to Fas-mediated apoptosis in cultured K8-null hepatocytes (Fig. 1 C). These results demonstrate the biological relevance of our model system. Moreover, in the light of the present findings on K8-null hepatocytes, the use of K18-null mice (Magin et al., 1998) should yield the same type of functional association between K8/K18 and Fas-mediated apoptosis. Previous analyses of mouse nonepithelial cell lines expressing normal or mutant human K8 and/or K18 cDNAs or mice carrying such transgenes have provided significant information on the contribution of the protein domains to resistance to various forms of stress (Ku et al., 1996, 2001; Oshima et al., 1996). Our next challenge therefore is to identify which of the particular K8 and K18 domains, and even which of the putative phosphoamino acid residues, are providing the protective resistance to Fas-mediated apoptosis of hepatocytes.
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Materials and methods |
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Mice
Details on the establishment, maintenance, and genotyping of the K8-deficient FVB/N mouse line were reported previously (Baribault et al., 1994; Loranger et al., 1997). The mice were housed in the Specific Pathogen Free Animal Facility at this research center. In particular, histological monitoring of mouse hepatic tissue for Helicobacter hepaticus revealed no pathological signs that are usually attributed to the pathogen (Fox et al., 1996; Ihrig et al., 1999), and PCR-based screening of the mouse feces confirmed the absence of the pathogen. The experiments were performed according to the rules of the Laval University Animal Care Committee.
Jo2 treatment of mice and hepatocyte damage assessment
The sensitivity of mice to a single i.p. injection of Jo2 depends on the strain (Sarraf et al., 1997; Hara et al., 2000). Accordingly, we first evaluated the response of K8-null and WT FVB/N mice to increasing Jo2 concentrations. A dose of 75 µg/kg body weight in 200 µl of saline solution resulted in a 90% WT mouse survival (see Results). Just before the injection, a blood sample was collected from the saphen vein with a nonheparinized capillary tube. At 24 h after injection, a blood sample was obtained by cardiac puncture while the mouse was under ether anaesthesia. The level of hepatocyte damage was assessed by the amount of ALT released in serum, based on the level of ALT activity measured with the Randox kit. In some experiments, mice received an i.p. injection of colchicine at a dose of 400 µg/kg body weight in 200 µl of saline solution 24 h before the challenge with a lethal dose (200 µg/kg) of Jo2. The proper colchicine dose was selected after assessment of the ALT activity in response to increasing doses (02000 µg/kg); at 400 µg/kg, no significant increase in ALT level was measured in the serum of either K8-null or WT mice before Jo2 injection.
Hepatocyte isolation and culture
Hepatocytes were isolated according to a modified version of the two-step method with collagenase originally developed for rats (Seglen, 1976; Deschenes et al., 1980). Mice were anaesthetised with isoflurane, and their abdominal cavity was opened to cannulate the vena portalis and to section the vena cava. The liver was perfused at a flow rate of 5 ml/min at 37°C, first with a Ca2+-free Hepes, 25 mM buffer, pH 7.5, containing insulin (0.5 µg/ml) and EGTA (0.5 mM), and then with DME/F12 modified medium containing collagenase (0.2 U Wünsch/ml) and Ca2+ (5 mmol/liter). The yield of isolated hepatocytes was determined with a hemocytometer, and their viability evaluated with the standard Trypan blue exclusion assay. This isolation procedure yielded 67 x 107 hepatocytes/liver, with a viability of 9095%.
Hepatocytes were plated at a density of 1.2 x 105 cells/cm2 on fibronectin-coated dishes in DME/F12 modified medium supplemented with selenious acid (5 µg/l), insulin (5 mg/l), transferrin (5 mg/l), streptomycin (100 µg/ml), and penicillin (100 units/ml). After a 3-h attachment period, the culture medium was replaced by the same medium supplemented with dexamethasone (10-7 M) and EGF (20 ng/ml).
Assessment of apoptosis in culture
At 24 h after seeding, the medium was changed to DME/F12 supplemented with selenious acid (5 µg/liter), transferrin (5 mg/liter), dexamethasone (10-7 M), Matrigel (0,5 mg/ml), streptomycin (100 µg/ml), and penicillin (100 units/ml)). The apoptosis inducer (i.e., Jo2, TNF-, TRAIL, or TGF-ß) was added in the absence or presence of CHX or Act D, 24 h after the medium change. In some experiments, EGF (20 or 200 ng/dish), colchicine (1 µM), or
-lumicolchicine (1 µM) was added 24 h after seeding and then maintained throughout the experiment.
Apoptotic hepatocytes remained attached to the culture substratum and were readily detected by Acridine orange staining of the altered chromatin, as described previously (Guilhot et al., 1996). Hepatocytes were examined with a laser scanning confocal microscope (Bio-Rad MRC-1024) equipped with a Nikon Diaphot and a kryptonargon laser that emits light at 488 and 568 nm; the Acridine orange is excited by the 488-nm line. When one apoptotic hepatocyte exhibited multiple apoptotic bodies, these were still scored as one. At least 400 cells were counted in six random fields per dish, and each experiment was repeated in duplicate with at least three mice. The DNA ladder assay was performed as described (Feng and Kaplowitz, 2000).
Reinsertion of WT K8 gene into cultured K8-null hepatocytes
A complete human K8 cDNA was transferred into K8-null hepatocytes using a Moloney murine leukemia retroviral vector derived from MFGb2, known for providing a high-level gene expression in transduced cells (Riviere et al., 1995). A retroviral plasmid pGFP3 containing the Herpes simplex virus TK gene followed by an internal ribosomal entry site sequence and a enhanced green fluorescent protein cDNA, was provided by Dr. M. Caruso (Laval University, Quebec, Canada). To construct the pK8GFP retroviral plasmid, the NcoI-BamHI TK gene was replaced by the K8 cDNA, flanked by 5' NcoI and 3' BamHI restriction sites introduced by PCR. In these vectors, the internal ribosomal entry site allows cap-independent translation of the downstream gene, which leads to the translation of the two proteins from a single mRNA transcribed from the retroviral 5' long terminal repeat sequence (Martinez-Salas, 1999; Qiao et al., 2000). For generating the recombinant TK and K8 viruses, GP+E-86 packaging cells (Markowitz et al., 1988) were cotransfected by the calcium phosphate procedure with a puror plasmid and pGFP3 or pK8GFP, respectively. After 10 d of selection with 2 µg/ml puromycin, resistant clones were isolated, and virus supernatants were harvested and kept frozen at -80°C. The titration was performed on NIH-3T3 cells (Qiao et al., 2000), by determining the number of GFP-positive cells after infection at serial dilutions of virus. The titers obtained for the TK and K8 retroviral vectors were 3 x 106 and 1 x 107 infectious virus/ml, respectively. K8-null hepatocytes were infected 48 h after seeding at a ratio of 10 infectious virus per cell. The medium was changed 16 h after infection, and the apoptosis was induced with Jo2 (0.5 µg/ml) 48 h later, as described above. The gene transfer efficiency, as provided by the proportion of GFP-positve hepatocytes, was >80%.
Immunofluorescence microscopy and FACS® analysis
Cultured hepatocytes or liver tissue sections were fixed according to procedures that varied with the nature of the antigen examined. For multiple labeling, the cells were rinsed twice in PBS, followed by an incubation in 2% paraformaldehyde in PBS for 10 min at room temperature and a 10 min treatment with 0.1% Triton X-100 at room temperature. After two consecutive washes with PBS, the fixed cells were incubated in a blocking solution (10% goat serum in PBS) for 30 min followed by two consecutive washes with PBS. The labeling was performed with PE-labeled anti-Fas Jo2 (5 µg/ml), antimannosidase II (1:100), or anti-K8 (TROMA-1) antibody, followed by 60-min incubations at room temperature with a ALEXA 350tagged goat antirabbit immunoglobulin antibody or an FITC-labeled goat antirat immunoglobin, respectively. In the case of tubulin, cultured hepatocytes were rinsed in PBS at 37°C followed by a 10-min 37°C incubation in a fixation buffer that contained 0.1 M Pipes, pH 6.75, 4% PEG-6000, 1 mM EGTA, 1 mM MgSO4, 1% Triton X-100, 2% paraformaldehyde (Caron et al., 1985). Cells were transferred to -20°C methanol for 5 min. After two successive 5-min washes in PBS at room temperature, hepatocytes were incubated overnight at 4°C with the first antibody (i.e., directed against tubulin), followed by a 60-min incubation at room temperature with Alexa 488tagged rabbit antimouse immunoglobulin antibody. Images were captured on an Eclipse TE300 inverted microscope (Nikon) with a MicroMAXTM interline transfer charge-coupled device camera (Princeton Instruments) controlled by a Metamorph digital imaging software, or collected with the laser scanning confocal microscope using the laser line 488 nm (Alexa 488; FITC) or 568 nm (PE; Texas red) for fluorochrome excitation and a high numerical aperture (1.4 NA, 60x) oil immersion objective. The density of Fas on the surface of freshly isolated hepatocytes was analyzed by FACS® as described in detail by Feng and Kaplowitz (2000). In brief, live cells were washed twice, incubated in proper buffer with 2 µg/ml PE-labeled anti-Fas Jo2 on ice for 20 min, washed three times, and then analyzed with a FACS® (Beckman Coulter) using the laser line at 488 nm. The relative mean fluorescence was calculated with EXPOTM Software, v2.0 (Applied Cytometry Systems/Beckman Coulter).
Western blotting
Total proteins were extracted with 300 µl/35 mm plastic petri dish of preheated at 90°C in 2x SDS-PAGE sample buffer (4x Tris HCl[0,5 M]/SDS[0,4%], pH 6.8, 20% glycerol, 4% SDS, 2% mercaptoethanol, and 1% Bromophenol blue), as described (Ausubel, 1994). Proteins (10 µg) were separated by PAGE according to Laemmli (1970) and then electrophoretically transferred onto a PVDF membrane. The blots were incubated with the primary antibody and then the second antibody conjugated with horseradish peroxidase. The staining was revealed with the SuperSignal West Pico kit (Pierce Chemical Co.).
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Footnotes |
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Acknowledgments |
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This work was supported by a grant from Canadian Institutes of Health Research and a grant from the The Cancer Research Society.
Submitted: 22 February 2001
Revised: 13 July 2001
Accepted: 13 July 2001
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References |
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