1 Center for Basic Research in Digestive Diseases, Mayo Clinic, Rochester, Minnesota 55905; and 2 Department of Biochemistry, Juntendo University School of Medicine, Bunkyo-ku, Tokyo 113, Japan
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ABSTRACT |
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We have previously demonstrated abrogation of bile salt-induced apoptosis by cathepsin B inhibitors. However, caspases have been strongly implicated in apoptosis, and the mechanistic interface between caspase and cathepsin B activation is unclear. Thus our aims were to determine the mechanistic relationship between caspases and cathepsin B in bile salt-induced apoptosis in a rat hepatoma cell line. Expression of cystatin A was used to inhibit cathepsin B, whereas Z-Val-Ala-Asp-fluoromethyl ketone (Z-VAD-FMK) was used to inhibit caspases. Cystatin A expression prevented cathepsin B activation and apoptosis during treatment with glycochenodeoxycholate (GCDC), a toxic bile salt. Caspase N-acetyl-Asp-Glu-Val-Asp-7-amino-4-methylcoumarin (DEVD-AMC) hydrolytic activity increased in both wild-type and cystatin A-transfected cells treated with GCDC, demonstrating caspase activation despite inhibition of cathepsin B. In contrast, Z-VAD-FMK blocked both DEVD-AMC hydrolytic activity and cathepsin B activity during GCDC treatment. Our data demonstrate that 1) bile salt-induced apoptosis can be inhibited by the cystatin A transgene and 2) caspase and cathepsin B activation are linked mechanistically with cathepsin B downstream of caspases.
caspase 3; cathepsin B; cholestasis; 4',6'-diamidino-2-phenylindole dihydrochloride
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INTRODUCTION |
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CHOLESTASIS, A PHYSIOLOGICAL impairment in bile flow, is a common feature in many human liver diseases (e.g., primary biliary cirrhosis, primary sclerosing cholangitis, chronic allograft rejection after liver transplantation, syndromatic paucity of intrahepatic bile ducts, Byler disease, and malignant and iatrogenic large bile duct strictures). Although the initial insult is to the bile ducts in many of these diseases, hepatocyte injury and progression of the liver disease are promoted by direct chemical damage to hepatocytes by toxic hydrophobic bile salts (10, 14). Indeed, the pathogenic effects of toxic bile salts have been well documented, especially hepatocellular death (21). The mechanisms leading to the induction of hepatocyte apoptosis by toxic bile salts remain unclear. These mechanisms are of clinical and scientific importance, because of the implications for developing future therapeutic approaches to treat cholestatic liver diseases. We have recently demonstrated (23) that cathepsin B is activated and translocated from the cytoplasm to the nucleus during bile salt-induced hepatocyte apoptosis; furthermore, the highly selective cathepsin B inhibitor CA-074 reduces apoptosis in this model of hepatocyte injury. These observations suggest that cathepsin B contributes to bile salt-induced hepatocyte apoptosis and may be a pharmacological target for the treatment of these liver diseases. However, cell fractionation studies are fraught with incomplete separation of cellular constituents and protease inhibitors may be nonselective. Therefore, additional complementary and more refined approaches are necessary to define the role of cathepsin B in bile salt-mediated apoptosis.
In addition to cathepsin B, other proteases, including calpains and
caspases, have been strongly implicated in hepatocyte apoptosis (11).
In particular, caspases, cytosolic cysteine proteases that cleave on
the carboxylic side of aspartic acid, have been implicated in
hepatocyte apoptosis by transforming growth factor- (TGF-
) and
activation of the Fas receptor (5, 7). These proteases are the human
homologs of ced-3, an essential gene
for apoptosis in Caenorhabditis
elegans (29). At least 10 mammalian
members of the caspase family have been cloned and characterized to
date. Caspases 2, 3, 6, 7, 8, and 10 have been implicated in apoptosis,
whereas caspases 1, 4, and 5 are thought to be involved in cytokine
activation (22, 25). Currently unknown, the role of caspases in bile
salt-mediated apoptosis and their relationship to cathepsin B
activation are important subjects for consideration.
Intracellular expression of proteins that are selective protease
inhibitors is a powerful technique to define the role of proteases in
cellular processes and is thought to avoid the nonspecificity associated with pharmacological protease inhibitors. The cystatins, also termed stefins, are a class of reversible, competitive inhibitors of cysteine proteases and are potent inhibitors of the cathepsins (2).
In particular, human cystatin A (an 11-kDa protein) efficiently inhibits cathepsin B with an inhibitor constant
(Ki) of 8 nM (1). The epidermis is the only tissue that maintains high-level expression of cystatin A, where it becomes incorporated into the keratin and may serve to protect against bacterial or viral proteases (17). Cystatin A is expressed at very low levels in the liver (17).
Although cystatin B, a related inhibitor, is relatively abundant in
hepatocytes, the
Ki of cystatin B
for cathepsin B is significantly higher (
73 nM) (1). Moreover,
unlike many of the other cystatins (cystatin C, D, S, SN, and SA),
cystatin A does not have a signal peptide for cellular secretion and
is, therefore, retained within the cell instead of being secreted (1).
Finally, overexpression of cystatin A has been shown to result in
reduced degradation of the cell cycle regulatory protein cyclin B in
NIH/3T3 fibroblasts and inhibition of virus-induced apoptosis in a
teleost cell line (3, 13). On the basis of this information, we
developed the hypothesis that expression of cystatin A in hepatocytes
would inhibit bile salt-induced apoptosis. Specifically, we aimed to
answer the following questions. 1)
Can cystatin A be efficiently expressed in hepatocytes, and if so, is
cathepsin B activity inhibited? 2)
Does expression of cystatin A reduce bile salt-induced apoptosis?
3) Are caspases also activated in
bile salt-mediated apoptosis? 4)
Does inhibition of cathepsin B with cystatin prevent caspase activation
or does inhibition of caspases with a selective inhibitor prevent
cathepsin B activation? We chose glycochenodeoxycholate (GCDC) as the
toxic bile salt for these studies, because it is a primary bile salt
whose concentrations increase during cholestasis (9, 10).
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EXPERIMENTAL PROCEDURES |
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Culture of McNtcp.24 cells. McNtcp.24 cells, a hepatocyte-derived cell line that stably expresses the sodium-dependent taurocholate cotransporting polypeptide and efficiently transports bile salts, were cultured in DMEM containing 10% fetal bovine serum and 10% horse serum (28). Cell culture medium was replaced every second day.
In vitro enzyme assays for measuring cathepsin B and caspase 3 activity in the presence of cystatin A. Purified cystatin A, purified cathepsin B, and recombinant caspase 3 were obtained from commercial sources and used as supplied. To measure cathepsin B activity, we performed experiments in 20 mM MOPS buffer, pH 6.0, containing 20 µM of the selective cathepsin B fluorogenic substrate Z-Arg-Arg-7-amino-4-methylcoumarin (Z-Arg-Arg-AMC), 0.5 µg/ml cathepsin B, and the desired concentration of cystatin A. To measure caspase 3-like activity, we performed experiments in 25 mM HEPES (pH 7.5), 10 mM dithiothreitol (DTT), 0.1% 3-[(3cholamidopropyl)dimethylammonio]-1-propane-sulfonate (CHAPS), 0.5 mM phenylmethylsulfonyl fluoride (PMSF), 100 U/ml Trasylol, 20 µM N-acetyl-Asp-Glu-Val-Asp-AMC (DEVD-AMC), 0.1 µg/ml caspase 3, and the desired concentration of cystatin A. Fluorescence was continuously monitored at room temperature using a fluorometer (Perkin-Elmer model LS 50, Norwalk, CT) equipped with a magnetic stirrer using excitation and emission wavelengths of 380 and 450 nm, respectively. We also verified that pretreatment of McNtcp.24 cells with 50 µM Z-VAD-fluoromethyl ketone (Z-VAD-FMK) for 30 min completely abolished DEVD-AMC hydrolytic activity in controls and GCDC-treated cells. Therefore, we used this concentration of Z-VAD-FMK in our experimental protocols.
Preparation of cystatin A expression vector.
Preparation of a plasmid vector encoding the cDNA for rat cystatin-
and for human cystatin A has previously been described (15, 16). The
cDNA for human cystatin A used in the current studies was originally
prepared in a pCR II plasmid (Invitrogen, San Diego, CA) and has been
submitted to the EMBL/Genbank databases (accession no. D88439). The
plasmid DNA was prepared from subcloned bacterial DNA using ampicillin
selection and purified with a commercial kit using the manufacturer's
suggested protocol (EndoFree Plasmid Maxi kit, Qiagen, Chatsworth, CA).
The DNA thus obtained was cut with
EcoR I and examined with agarose gel
electrophoresis to verify its identity.
Transient transfection of McNtcp.24 cells with cystatin A. Aliquots of McNtcp.24 cells (1.5 ml of 105 cells/ml) in DMEM containing 10% calf serum plus 10% fetal bovine serum were cultured in 35 × 10 mm petri dishes on collagen-coated 22-mm square glass coverslips. Twenty-four hours after plating, the medium was aspirated, and the cultured cells rinsed with 2 ml of Optimem I reduced serum medium (GIBCO-BRL, Grand Island, NY). For experiments evaluating the effect of de novo cystatin A expression on apoptosis, the cells were transfected by adding 1 ml of a mixture of Optimem I containing 8 mg lipofectamine (GIBCO-BRL) and 1 µg of the DNA plasmid to each culture dish. The cells were incubated in the above-described mixture for 5 h at 37°C in a 5% CO2-95% air incubator. Next, 1 ml of DMEM containing 20% fetal bovine serum plus 20% calf serum was added to the transfection medium in each culture dish. Twenty-four hours later, the medium was aspirated and replaced with 2 ml of DMEM containing 10% fetal bovine serum plus 10% calf serum. Transfection efficiency was established to be >80% by cotransfection with 0.5 µg of DNA of a pBK-CMV green fluorescent protein (GFP) expression vector along with the plasmid encoding cystatin A (0.5 µg of DNA). Transfection efficiency was quantitated by determining the number of cells expressing GFP using fluorescence microscopy (23). As assessed 48 h after transfection, viability and morphology of the McNtcp.24 cells were unaffected by transfection in the absence of DNA or transfection with the plasmid alone or the expression plasmid for cystatin A.
Immunoblot analysis for cystatin A. Cultured cells were scraped off the plates using a cell lifter in a hypotonic buffer (10 mM Tris, pH 7.4) containing protease inhibitors (Complete; EDTA-free protease inhibitor cocktail tablets; Boehringer Mannheim, Indianapolis, IN) and lysed by vortexing. After SDS-PAGE of cell proteins using a 18% polyacrylamide gel, proteins were transferred to a Nitrobind nitrocellulose membrane (Micron Separation, Westboro, MA) by electroblotting. The membrane was then rinsed briefly with 20 mM Tris and 0.5 M sodium chloride, pH 7.0 (TBS), blocked with 5% (wt/vol) skim milk in 20 mM Tris, 0.5 M sodium chloride, and 0.05% Tween 20, pH 7.0 (TTBS) for 30 min at room temperature to prevent nonspecific binding, and then incubated for 45 min with a 1:2,000 dilution of rabbit anti-cystatin A affinity-purified antibody solution. Membranes were washed three times in TTBS for 10 min each at room temperature and then incubated for 30 min at room temperature with a 1:4,000 dilution of horseradish peroxidase-conjugated goat anti-rabbit secondary antibody (Biosource International, Camarillo, CA). After three washes in TTBS for 10 min each at room temperature followed by a single wash in TBS for 10 min at room temperature, the blots were developed with an enhanced chemiluminescence substrate (Amersham, Arlington Heights, IL) and exposed to Kodak Biomax film. As a positive control for human cystatin A, cell lysates derived from a human keratinocyte cell line (generously provided by Dr. Mark Pittelkow, Mayo Clinic) were loaded into sample wells.
Measurement of intracellular cathepsin B activity in single cells. Intracellular single-cell cathepsin B protease-like activity was measured using the fluorogenic protease substrate Z-Val-Leu-Lys-7-amino-4-chloromethlcoumarin (Z-Val-Leu-Lys-CMAC) and digitized video microscopy as we previously described and validated (23). Briefly, protease activity was measured by adding 50 µM Z-Val-Leu-Lys-CMAC to the medium and quantitating intracellular fluorescence over 20 min using digitized video microscopy and excitation and emission wavelengths of 380 and 460 nm, respectively. Total cellular fluorescence (number of pixels above background multiplied by the average fluorescence per cellular region) was quantitated over time using the Metafluor software imaging program (Universal Imaging, West Chester, PA). Cathepsin B activity was quantitated as change in cellular fluorescence per cell per minute over the linear range of cellular protease activity.
Measurement of caspase protease activity in cytosol.
Cells were cultured in 100-mm culture dishes (Becton-Dickinson Labware,
Lincoln Park, NJ) until confluent. At selected time points in the
various experimental paradigms, the medium was decanted and the cells
lifted in 1 ml of hypotonic buffer using a cell lifter. The hypotonic
buffer consisted of 25 mM HEPES, 5 mM
MgCl2, 1 mM EGTA, 0.5 mM PMSF, 2 µg/ml pepstatin, and 2 µg/ml leupeptin, pH of 7.5. The protease
inhibitors were added fresh. All subsequent steps were performed at
4°C. Cells were lysed by 10 strokes in a mechanized Potter-Elvehjem
homogenizer (Curtin Matheson Scientific, Houston, TX) using a Teflon
pestle. The homogenate was centrifuged for 45 min at 45,000 g in an ultracentrifuge (TL-100;
Beckman Instruments, Palo Alto, CA). Samples were frozen at
70°C. Protease activity was measured by adding 50 µl of
cytosol to 450 µl of assay buffer containing 25 mM HEPES (pH 7.5), 10 mM DTT, 0.1% CHAPS, 0.5 mM PMSF, 100 U/ml Trasylol, and 20 µM
N-acetyl-Tyr-Val-Ala-Asp-AMC (YVAD-AMC) or 20 µM DEVD-AMC. Fluorescence was monitored at room temperature using a fluorometer (model 450; Sequoia-Turner, Mountain View, CA) with excitation and emission wavelengths of 360 and 430 nm
for AMC.
Quantification of apoptosis. We quantitated apoptosis as previously described in detail by assessing the nuclear changes of apoptosis using the nuclear binding dye 4',6'-diamidino-2-phenylindole dihydrochloride (DAPI) and fluorescence microscopy (21).
Statistical analysis. All data represent at least three experiments using cells, tissue, or extract from a minimum of three separate isolations and are expressed as the means ± SE unless otherwise indicated. Differences between groups were compared using ANOVA for repeated measures and a post hoc Bonferroni test to compare for multiple comparisons. All statistical analyses were performed with the statistical software package InStat from GraphPAD (San Diego, CA).
Materials.
All chemicals used were of analytical grade purity. GCDC was >96%
pure as determined by the manufacturer through TLC and was used without
further preparation. GCDC, DMSO, bromphenol blue, Tris, EDTA, EGTA,
formaldehyde, Triton X-100, RNase, leupeptin, pepstatin, aprotinin,
-mercaptoethanol, Coomassie brilliant blue, and Z-Arg-Arg-AMC were
obtained from Sigma Chemical (St. Louis, MO). DEVD-AMC, DEVD-FMK, and
YVAD-AMC were purchased from Enzyme Systems (Dublin, CA). DAPI,
Z-Val-Leu-Lys-CMAC, and propidium iodide were obtained from Molecular
Probes (Eugene, OR). Proteinase K, collagenase type D, and PMSF were
obtained from Boehringer-Mannheim (Indianapolis, IN).
Ethylhexadecyldimethylammonium bromide was obtained from Eastman Kodak
(Rochester, NY). Precast polyacrylamide gels and Mark 12 wide-range
molecular weight markers for electrophoresis were obtained from Novex
(San Diego, CA). Nitrocellulose membranes were obtained from Micron
Separation. Secondary antibody (goat anti-rabbit) was obtained from
Biosource (Camarillo, CA). Restriction endonucleases
(Pst I,
EcoR I, and
Hind III) and PCR reagents
(MgCl2, Taq polymerase) were from GIBCOBRL.
The McNtcp.24 cell line was a generous gift from Dr. L. Agellon
(University of Alberta, Edmonton, Alberta, Canada).
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RESULTS |
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Does cystatin A inhibit cathepsin B and caspase 3 activity in vitro? Cystatin A is known to inhibit cathepsin B. However, as an inhibitor of cysteine proteases, cystatin could potentially inhibit caspase 3, a cysteine protease strongly implicated as a distal effector in many models of apoptosis. If cystatin A inhibits caspase 3, its lack of specificity as a protease inhibitor would preclude its use to delineate the role of cathepsin B in apoptosis. Therefore, we directly measured cathepsin B and caspase 3 activity in the presence of cystatin A (Fig. 1). As expected, cystatin A inhibits cathepsin B in a concentration-dependent manner. However, over the same concentration range that effectively inhibits cathepsin B, cystatin A had no effect on caspase 3 activity. Because cystatin A inhibits cathepsin B but not caspase 3, we continued with our experiments to determine if overexpression of cystatin A inhibited cathepsin B activity in bile salt-mediated hepatocyte apoptosis.
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Can cystatin A be expressed in hepatocytes and if so, is intracellular cathepsin B activity inhibited? We first wanted to establish that cystatin A was expressed at the level of the protein in transfected but not wild-type cells. Expression of cystatin A in transfected McNtcp.24 cells was determined by immunoblot analysis 48 h after transfection (Fig. 2). Cystatin A was not expressed in wild-type McNtcp.24 cells but was readily detected in cells transfected with the plasmid containing the cDNA for cystatin A. Having established the expression of cystatin A protein in transfected cells, we next sought to determine if the magnitude of cystatin A expression was sufficient to inhibit intracellular cathepsin B activity during treatment of McNtcp.24 cells with GCDC. Forty-eight hours after transfection of McNtcp.24 cells with the empty plasmid (pBK-CMV) or the plasmid containing the cDNA for cystatin A, cathepsin B activity was assessed after treatment of the cells with 50 µM GCDC for an additional 2 h (Fig. 3). Cystatin A expression reduced cathepsin B activity in untreated cells and prevented the increase in cathepsin B activity observed during treatment with GCDC. Although cathepsin B activity in cells treated with GCDC appeared to be reduced by transfection with the plasmid alone compared with untransfected cells, this difference was not statistically significant [279 ± 18% vs. 350 ± 20% of control, P = not significant (NS)]. Thus the efficiency of expression and the compartmentation of expressed cystatin A in transfected cells are sufficient to inhibit the increase in cathepsin B activity during treatment of McNtcp.24 cells with GCDC.
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Does expression of cystatin A reduce bile salt-induced apoptosis? We have previously demonstrated that >80% of McNtcp.24 cells can be transfected with lipofectamine, using expression of GFP as the marker for transfection (23). Because of this remarkably high transfection efficiency, we were able to measure rates of apoptosis in total cell populations rather than measuring apoptosis only in cells expressing a marker for transfection. Forty-eight hours after transfection of McNtcp.24 cells with the empty plasmid or the plasmid containing the cDNA for cystatin A, apoptosis was assessed after treatment of the cells with 50 µM GCDC for an additional 2 h (Fig. 4). In cells treated with GCDC, apoptosis was reduced by 64% in cells transfected with cystatin A compared with cells transfected with the empty plasmid (33 ± 1% vs. 12 ± 2%, P < 0.05). Although transfection with plasmid alone appeared to increase apoptosis compared with untransfected cells after treatment with GCDC, this difference was not statistically significant (33 ± 1% vs. 27 ± 4% of control, P = NS). Thus cystatin A not only inhibits cathepsin B activity but also reduces apoptosis during treatment of McNtcp.24 cells with toxic concentrations of GCDC. These observations suggest cathepsin B contributes to bile salt-induced apoptosis.
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Are caspases also activated in bile salt-mediated apoptosis? Members of the caspase family of proteases are cysteine proteases that have been strongly implicated in apoptosis (25). Therefore, we determined whether caspase activation also occurs in bile salt-mediated apoptosis. Cytosolic caspase activity was measured using two fluorogenic substrates, DEVD-AMC and YVAD-AMC. DEVD-AMC is a substrate for caspases 3, 7, and 8, whereas YVAD-AMC is an efficient substrate for caspases 1 and 4 (25). DEVD-AMC, but not YVAD-AMC, hydrolytic activity increased in cytosolic extracts obtained from McNtcp.24 cells treated with 50 µM GCDC (Fig. 5). These data suggest that specific members of the caspase family may be activated during bile salt-mediated hepatocyte apoptosis.
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Does inhibition of cathepsin B with cystatin prevent caspase activation or does inhibition of caspases with a selective inhibitor prevent cathepsin B activation? Having determined that caspases are activated in GCDC-mediated hepatocyte apoptosis, we next sought to determine if inhibition of cathepsin B by cystatin A expression would prevent caspase activation. Forty-eight hours after transfection of McNtcp.24 cells with the empty plasmid or the plasmid containing the cDNA for cystatin A, cells were treated with 50 µM GCDC for an additional 2 h. Total DEVD-AMC cleavage activity was virtually identical in the cytosol obtained from cells transfected with the empty plasmid vs. cystatin A after treatment with GCDC (Fig. 6). Thus inhibition of cathepsin B by cystatin A expression does not appear to prevent activation of the caspase family members recognizing DEVD-AMC. In contrast, the selective caspase inhibitor Z-VAD-FMK (50 µM) prevented activation of both caspases and cathepsin B (Fig. 7). The caspase inhibitor also reduced GCDC-mediated apoptosis by 53% after 2 h of treatment (17.2 ± 0.9% vs. 36.3 ± 2.1% for cells treated with Z-VAD-FMK + GCDC vs. GCDC, respectively) (Fig. 8). These data support the hypothesis that caspase activation and cathepsin B are linked mechanistically and that cathepsin B appears to be "downstream" of caspases in a linear proteolytic cascade.
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DISCUSSION |
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The original observations of this study relate to the protease effector mechanisms contributing to bile salt-induced apoptosis of hepatocytes. Our results directly demonstrate the following: 1) expression of cystatin A in McNtcp.24 cells reduced cathepsin B activity and apoptosis during treatment of the cells with GCDC; 2) caspase DEVD-AMC hydrolytic activity is increased in GCDC-mediated hepatocyte apoptosis; 3) inhibition of cathepsin B by expression of cystatin A does not prevent activation of DEVD-AMC hydrolytic activity during treatment of the cells with GCDC; and in contrast 4) inhibition of caspase activity with Z-VAD-FMK reduces activation of cathepsin B and apoptosis. These results provide additional data suggesting cathepsin B contributes to bile salt-induced apoptosis. Furthermore, these data begin to order the protease cascade occurring in bile salt-induced apoptosis; cathepsin B appears to be a downstream effector protease dependent on caspase activity for activation.
Bile salt-induced apoptosis is a tissue- and disease-specific model of
apoptosis. Hepatocytes, cholangiocytes, and ileal enterocytes are the
only cells that can transport bile salts intracellularly. In
cholestatic liver diseases associated with high serum concentrations of
bile salts, these toxins can be retained specifically within hepatocytes leading to apoptosis. In the current study we used the rat
hepatoma McNtcp.24 cell line because human, mouse, and rat hepatocyte
primary cultures dedifferentiate after isolation and rapidly lose the
ability to efficiently transport bile salts. This rapid
dedifferentiation hinders the use of primary hepatocyte cultures for
transient transfection studies that often require 48 h for maximum
protein expression. The McNtcp.24 cell line also has the advantage of a
high transfection efficiency. Indeed, this cell line allowed us to
determine if cystatin A inhibited GCDC-mediated apoptosis. For a cell
line to be an appropriate model of bile salt-apoptosis, it would need
to be a hepatocyte-derived cell line that transports bile salts with
kinetics similar to primary hepatocytes and undergoes apoptosis. The
McNtcp.24 cells derived from the McA-RH7777 rat hepatoma cell line
would appear to fulfill these criteria. First, these cells transport
bile salts into and out of the cell at rates virtually identical to
primary hepatocytes (28). Second, this cell line undergoes Fas ligand-,
tumor necrosis factor- (TNF-
)-, and staurosporine-induced
apoptosis, demonstrating that the apoptotic pathways are intact
(unpublished data). The concentrations of bile salts that induce
apoptosis in this cell line are also identical to those that induce
apoptosis in primary cultures of rat or mouse hepatocytes (23).
Although extrapolation of data obtained with cell lines to fully
differentiated cells in vivo has potential pitfalls, based on the
information available the McNtcp.24 cell line would appear to be an
acceptable model for bile salt-induced apoptosis.
We have recently demonstrated that cathepsin B is activated and
translocated to the nucleus during bile salt-induced apoptosis in
primary cultures of rat hepatocytes and in vivo during extrahepatic cholestasis by bile duct ligation. Furthermore, inhibition of cathepsin
B with protease inhibitors blocks bile salt-induced apoptosis (23). In
our current study, we have extended and strengthened these observations
by demonstrating that expression of cystatin A, a potent inhibitor of
cathepsin B, reduces cathepsin B activity and apoptosis during
treatment of rat hepatoma cells with GCDC. These results are bolstered
by the observations that cystatin A does not block caspase 3, a
protease strongly implicated in other models of apoptosis. Numerous
other reports have also implicated cathepsins in various models of
apoptosis. Cathepsin B antisense cDNA attenuates hepatocyte apoptosis
by bile salts (23). Cathepsin D antisense cDNA blocks Fas-,
interferon--, and TNF-
-induced apoptosis in HeLa cells (6).
Cathepsin D may activate cathepsin B through proteolysis of
procathepsin B and may enhance cathepsin B activity by degrading
cystatin A (1, 24). Pharmacological inhibition of cathepsin B blocks
apoptosis induced by p53 and cytotoxic agents (18). Cathepsin B
expression is increased in apoptosis-related postlactation regression
of the mammary gland and involution of the prostate after castration
(12). A redistribution of cathepsin B from vesicles to the cytosol has
been identified in neuronal death after global ischemia (27).
Finally, apoptosis during palatine development is associated with the
appearance of cathepsin B in the nucleus (20). These numerous,
independent observations suggest that cathepsins can participate in the
effector processes of apoptosis.
We have demonstrated that inhibition of caspases blocked activation of
cathepsin B, while inhibition of cathepsin B did not block caspase
activation. Although the specificity of the pharmacological caspase
inhibitor Z-VAD-FMK is unknown, we have recently verified these
observations using transfection with CrmA, a highly selective caspase
inhibitor (unpublished observations). These data suggest that cathepsin
B is downstream of caspases in a proteolytic cascade. The mechanisms by
which caspases lead to cathepsin B activation are likely indirect, as
cathepsin B is not converted from the proform to the active protease by
cleavage at aspartate residues (24). Furthermore, cathepsin B is
predominantly localized to acidic vesicles and caspases in the cytosol,
thereby limiting the access of cathepsin B to caspases (24). It is
possible that caspases may modulate the permeability, function, or
localization of acidic vesicles, promoting cathepsin B activation,
release from acidic vesicles, or localization with apoptotic
substrates. Indeed, Monney et al. (19) have demonstrated that
alkalinization of acidic vesicles attenuates TNF--induced apoptosis.
Generation of ceramide by acidic sphingomyelinase present in acidic
vesicles has also been implicated as an effector mechanism mediating
apoptosis (26). This activation of acidic sphingomyelinase also appears to be dependent on caspases (4). These data, along with our observations, suggest that acidic vesicles and their enzyme
constituents are involved in apoptosis after activation of caspases,
probably the proximal signaling caspases, caspases 2, 8, and 10. Indeed, all of these caspases along with caspases 3 and 7 could
contribute to the observed increase in DEVD-AMC hydrolytic activity in
our studies. Further studies will be required to elucidate which
specific caspases are activated in this model of apoptosis and how
their activity promotes activation of cathepsin B.
Although Z-VAD-FMK prevented the increase in cathepsin B activity, it did not completely block GCDC-mediated apoptosis. Likewise, although Z-VAD-FMK completely abolished DEVD-AMC hydrolytic activity, it also did not completely attenuate apoptosis. There are several potential interpretations of these data. First, Z-VAD-FMK may have inherent cellular toxicity in the context of exposure to GCDC. Second, GCDC may also activate a cathepsin B- and caspase-independent proapoptotic pathway. Finally, our enzymatic assays may not be sensitive enough to detect biologically important, but small, increases in either cathepsin B or caspase activity. The first interpretation of these data is difficult to exclude but would appear to be unlikely based on experiences in the literature. Z-VAD-FMK has been widely used as an "industry standard" for generic inhibition of caspases, and no toxicity has been reported. It is difficult to envision how GCDC would unmask or result in Z-VAD-FMK cytotoxicity. However, we cannot distinguish between the other two possibilities. GCDC may well initiate a cathepsin B-independent and or caspase-insensitive apoptosis pathway that becomes manifest in the presence of enzyme inhibitors. On the other hand, the amount of activated cathepsin B or caspase necessary to initiate an amplified apoptosis cascade is unknown. Knockout mice for cathepsin B and individual caspases will be required to completely answer these questions.
The long-term objective of our studies is to delineate the mechanisms causing hepatocellular injury during cholestasis. Our current working hypothesis is that during cholestasis the retention and accumulation of toxic bile salts within hepatocytes trigger the apoptotic machinery of the cell leading to cell death. The data in the current study in combination with our previous observations suggest that toxic bile salts cause cholestasis, in part, by activating a protease cascade. Caspases appear to be initially activated with subsequent activation of cathepsin B. Fraser and Evan (8) have developed a schema classifying the proteases participating in apoptosis as initiator, amplifier, and machinery proteases. On the basis of this schema, caspases would appear to serve as initiator and amplifier proteases in bile salt cytotoxicity, wheras cathepsin B would be a machinery protease. Indeed, the relative nondiscriminatory endopeptidase activity of cathepsin B would make it an attractive degradative protease. Finally, we believe our observations have potential therapeutic implications. We speculate that pharmacological inhibition of either caspases or cathepsin B, especially if the inhibitor were specific and selectively targeted to the liver, could conceivably have a salutary effect on cholestatic liver injury by blocking hepatocyte apoptosis.
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ACKNOWLEDGEMENTS |
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We gratefully acknowledge the secretarial assistance of Sara Erickson and the expert technical services of Steven Bronk. We thank Dr. Luis Agellon (University of Alberta, Edmonton, Alberta, Canada) for generously providing the McNtcp.24 cell line.
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FOOTNOTES |
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This work was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant DK-41876 and by grants from the Gainey Foundation (St. Paul, MN) and the Mayo Foundation (Rochester, MN).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Address for reprint requests: G. J. Gores, Mayo Medical School, Clinic, and Foundation, 200 First St., SW, Rochester, MN 55905.
Received 23 February 1998; accepted in final form 4 June 1998.
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