Hepatitis B Virus X Protein Induces Cell Death by Causing Loss of Mitochondrial Membrane Potential*

Yumiko Shirakata and Katsuro Koike {ddagger}

From the Department of Gene Research, The Cancer Institute (JFCR), Kami-Ikebukuro, Toshima-ku, Tokyo 170-8455, Japan

Received for publication, February 14, 2003
    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The hepatitis B virus X protein (HBx) has been implicated in the carcinogenicity of this virus as a causative factor by means of its transactivation function in development of hepatocellular carcinoma. However, we and others have recently reported that HBx is located in mitochondria and causes subsequent cell death (Takada, S., Shirakata, Y., Kaneniwa, N., and Koike, K. (1999) Oncogene 18, 6965–6973; Rahmani, Z., Huh, K. W., Lasher, R., and Siddiqui, A. (2000) J. Virol. 74, 2840–2846). In this study, we, therefore, examined the mechanism of HBx-related cell death. Using enhanced green fluorescent protein (EGFP) fusion constructs of HBx, the region required for its mitochondrial localization was mapped to amino acids (aa) 68–117, which is essential for cell death but inactive for transactivation function. In vitro binding analysis supported the notion that the recombinant HBx associates with isolated mitochondria through the region of aa 68–117 without causing redistribution of cytochrome c and apoptosisinducing factor (AIF). A cytochemical analysis revealed that mitochondrial membrane potential was decreased by HBx association with mitochondria, suggesting that HBx induces dysfunction of permeability transition pore (PTP) complex. Furthermore, PTP inhibitors, reactive oxygen species (ROS) scavengers and Bcl-xL, which are known to stabilize mitochondrial membrane potential, prevented HBx-induced cell death. Collectively, the present results suggest that location of HBx in mitochondria of hepatitis B virus-infected cells causes loss of mitochondrial membrane potential and subsequently induces mitochondria-dependent cell death.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Epidemiological studies show that chronic hepatitis B virus (HBV)1 infection is closely associated with development of hepatocellular carcinoma (3), and the X gene has been implicated in the carcinogenicity of this virus because of its ability to transform rodent cells (4). Moreover, the development of hepatocellular carcinoma in at least one strain of transgenic mouse has been shown (5). Although there was an absence of tumors in other X transgenic mice so far analyzed, some of these mice exhibited an increased susceptibility to chemical carcinogens (6, 7). These results suggest that HBV X protein (HBx) acts in cooperation with cellular oncogenic factors. However, pathogenesis and carcinogenesis by HBV infection are partly understood, because they occur over years of chronic infection and associate with hepatocyte death and regeneration.

It has been reported that HBx is essential for the viral replication (8, 9), in addition to its well documented transactivation ability to indirectly stimulate transcription (1012) and to activate several signal transduction pathways including mitogen-activated protein kinase, protein kinase C, and JAK/STAT pathways (1317). Although HBx transactivates a wide range of viral and cellular genes, including HBV itself, SV40, c-myc, and NF-{kappa}B, the mechanism of transactivation by HBx is still under debate (18). Because HBx is unable to directly bind any of the HBx-responsive elements of these genes, it has been proposed that HBx may stimulate transcription by interacting with several cellular sequence-specific transcription factors or with the basal transcriptional machinery (1921). From these observations, it has been suggested that HBx may facilitate hepatocarcinogenesis through transactivating cellular oncogenes and signaling cascades that stimulate cell proliferation.

In contrast to proliferative effects, several studies (1, 22, 23) showed that HBx induces cell death or sensitizes cells to apoptotic stimuli such as tumor necrosis factor {alpha}. Our previous report revealed that expression of X gene brought some of apoptotic features to cells, such as TUNEL-positive nucleus, the decreased mitochondrial membrane potential, and the membrane blebbing. We also reported that the ectopically expressed HBx is located in mitochondria (1). Recently, many reports have focused on mitochondria as an important player on apoptosis and programmed cell death, where many cell death signals reach to mitochondria. Mitochondria undergo major changes in membrane integrity before classical signs of cell death manifest. These changes concern both inner and outer mitochondrial membranes. Outer mitochondrial membrane permeabilization causes the redistribution of numerous apoptogenic factors such as cytochrome c, AIF, and Smac/DIABLO in the intermembrane space (24, 25). The redistribution of these apoptogenic factors is important step of cell death. For instance, cytochrome c release is essential for the generation of the apoptosome, the complex containing cytochrome c, Apaf-1, and caspase-9, and this apoptosome complex initiates the effector caspase activation (2628). On the other hand, the inner mitochondrial membrane permeabilization causes disruption of mitochondrial membrane potential ({Delta}{Psi}m). The {Delta}{Psi}m is indispensable for driving the ATP synthase, which phosphorylates ADP to ATP. Therefore the loss of mitochondrial membrane potential means the loss of ATP generation. In this paper, we show that the localization of HBx on mitochondria causes loss of mitochondrial membrane potential and subsequently causes cell death through a mitochondria-dependent pathway.


    EXPERIMENTAL PROCEDURES
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Reagents and Plasmids—The X expression plasmid pCMVX, pCMVX-dAvaI/AvaII, and pCMVX-{Delta}RsaI were described previously (12). The Bcl-xL cDNA was cloned into pCMV vector. EGFP or EBFP fusion constructs were cloned into pEGFP or pEBFP vector (Clontech), respectively. Substitution of pEGFP-X(68–117LA) in HBx pEGFP-X(68–117) was carried out by PCR based site-directed mutagenesis and confirmed by DNA sequencing. A {beta}-galactosidase expression plasmid, pCMV{beta}, was obtained from Clontech. Caspase inhibitors, z-VAD-fmk and z-DEVD-fmk were obtained from Enzyme Systems Products. Bongkrekic acid was obtained from Calbiochem, and cyclosporin A, 4,5-dihydroxy-1,3-benzenedisulfonic acid (Tiron), and N-acetyl-L-cysteine (NAC) were purchased from Sigma.

Cell Culture and DNA Transfection—HuH7 cells and HepG2 cells were maintained in DM-160AU supplemented with 10% fetal calf serum and 60 µg/ml kanamycin. Transfections were carried out using Tfx-20 (Promega) according to the manufacturer's instructions. Cells were cultured for 2 days after transfection and subjected to cytochemical staining or Western blot analysis unless stated.

Cell Death Assay—HuH7 cells were transfected with indicated plasmid along with the {beta}-galactosidase expression plasmid, pCMV{beta}, at a ratio of 5:1. The amount of cell death was determined by counting the {beta}-galactosidase-stained transfected cells with apoptotic morphology, such as membrane blebbing and/or round. Over 50 of {beta}-galactosidasestained cells were counted and calculated relative percent of cell death. Each experiment was repeated at least three times and obtained similar results. In situ staining of cells for {beta}-galactosidase activity were performed with {beta}-galactosidase staining kit (Invitrogen).

Luciferase Assay—HepG2 cells were plated on the day before transfection at a density of 6.8 x 105 cells per 60-mm-diameter dishes. 1 µg of pRSVL and 0.1 µg of pCMVX or HBx mutant constructs were cotransfected using Tfx-20 reagent. pCMV was used as carrier to adjust the total DNA to 2.75 µg. Two days after transfection, cells were harvested and lysed with lysis buffer (Promega). Luciferase assay was carried out with extracts according to the manufacturer's instructions.

Western Blot Analysis—Immunoblots were performed as described previously (1). The anti-His antibody (Santa Cruz Biotechnology), anti-cytochrome c antibody (PharMingen), anti-AIF antibody (Santa Cruz Biotechnology) has been used at dilution of 1:500. Reactive protein bands were visualized with the enhanced chemiluminescence (ECL; Amersham) system according to the manufacturer's instructions.

Microscopic Analysis—Mitochondria were visualized with MitoTracker (Red CMXRos) and 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolcarbocyanine iodide (JC-1). MitoTracker and JC-1 are obtained from Molecular Probes. Cells were incubated with MitoTracker (500 nM) or JC-1 (10 µg/ml) for 10 min at 37 °C, and then a specimen was analyzed using a Nikon microscope with CCD camera, which was operated by ARGUS FISH Imaging Software (Hamamatsu Photonics).

Mitochondria Preparation and Co-precipitation Assay—Before preparation a mouse (BALB/c, female, 6–8 weeks) was starved for 16 h. Isolation of mitochondria is followed by Shimizu et al. (29, 30) with slight modification. Briefly, the liver was flushed with phosphate-buffered saline and homogenized with a glass-Teflon Potter homogenizer in 15 ml of mitochondria isolation buffer (0.35 M mannitol, 10 mM Hepes, 0.1% bovine serum albumin, pH 7.2). Unbroken cells and nuclei were pelletted by centrifugation at 600 x g for 5 min at 4 °C. The supernatants were further centrifuged at 10,000 x g for 10 min at 4 °C to pellet the mitochondria. The mitochondria pellet was resuspended in 5 ml of mitochondria isolation buffer. A 30-µl aliquot of mouse liver mitochondria was incubated with the indicated recombinant protein in 30 µl of the same buffer for 60 min at 30 °C. At the end of incubation, the reaction mixture was centrifuged, and the pelleted mitochondria fraction was resuspended in 50 µl of 1 x SDS sample buffer, and 12 µl of 6 x SDS sample buffer was added to the resulting supernatant. The samples were boiled and subjected to Western blot analysis. Recombinant proteins were purified as follows. His-tagged fusion proteins were produced in Eschericha coli, BL21(DE3)pLysS, and purified on nickel-nitrilotriacetic acid-agarose beads (Qiagen) followed by high performance liquid chromatography purification. The protein concentration was determined by Bradford method (31).


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Membrane Blebbing Morphology of the HBx-expressing Cells—Our previous report (1) indicated that X gene expression brought about some apoptotic changes to cells such as TUNEL-positive nucleus, the decreased mitochondrial membrane potential, and the membrane blebbing. To characterize details of morphological changes, the membrane blebbing was examined using the HBx-expressing cells and the staining technique, because this protocol is easy to distinguish cells which are transfected with DNA and is able to handle many cells for estimating the frequency of dying cells. We carried out the time course analysis of HBx-induced cell death. HuH7 cells were transfected with an X gene expression plasmid, pCMVX, along with {beta}-galactosidase expression plasmid, pCMV{beta}. The {beta}-galactosidase staining was performed to visualize cells transfected with plasmids DNA. As shown in Fig. 1A, the HBx-expressing cells showed round and blebbing morphology that are easily distinguished from that of vector-transfected control cells. This type of morphological change is known to be characteristic of apoptosis (32). As summarized in Fig. 1B, the level of {beta}-galactosidase activity-positive cells with membrane blebbing reached a maximum in 3 days after transfection. The following experiments for cell death analysis were therefore carried out at 3 days after transfection.



View larger version (39K):
[in this window]
[in a new window]
 
FIG. 1.
Morphological changes and cell death are caused by HBx expression. A, characteristic morphology of apoptosis, round and membrane blebbing phenotype are observed after HBx expression. HuH7 cells were transfected with an HBx expression plasmid pCMVX (b) or vector control pCMV (a) along with a {beta}-galactosidase expression plasmid, pCMV{beta}, at a ratio of 5:1. Three days after transfection, cells were stained for {beta}-galactosidase activity in situ. The arrowheads indicate cells with membrane blebbing and/or round morphology. Scale bar is 100 µM. The enlargements of the typical cells indicated by yellow arrows are shown (panels c and d). B, time course of HBx-induced cell death. HuH7 cells were transfected with pCMV (open bars) and pCMVX (filled bars) along with pCMV{beta}. After transfection and staining, at least 50 of the transfected ({beta}-galactosidase activity positive) cells were counted for estimation of the frequency of dying cells, such as morphologically different cells with round and/or blebbing.

 

Transactivation Function of HBx Is Not Required for Cell Death—As we previously reported, some regions of HBx, such as aa 61–69, 105–125, and 131–140 (Fig. 2A, indicated as black box) are responsible for transactivation function (12), and that mutants lacking one of these regions lose their transactivation ability, we, therefore, examined whether the transactivation function of HBx is required for HBx-induced cell death. The luciferase assay and the morphological characterization were carried out using two different transactivation-defective mutants, one is a deletion mutant, pCMVX-dAvaI/AvaII, which has an internal deletion from aa 5 to 62, and the other mutant is pCMVX-{Delta}RsaI that has C terminus truncation from aa 134 to 154. Consistent with previous report (12), these mutants were unable to activate the RSV promoter (Fig. 2C), whereas these two mutants still retain cell death activity comparable with that of wild type HBx (Fig. 2B). Furthermore, we analyzed the dose dependence of HBx expression plasmid being able to induce cell death or transactivation (Fig. 2, D and E). The 1:1 ratio of pCMVX to pCMV{beta} induced maximum transactivation activity, although cell death is barely observed in this condition. As to cell death induction, 10 times more HBx expression plasmid is required as compared with maximum transactivation induction. Results clearly indicate that the transactivation ability of HBx is not directly related with its cell death function.



View larger version (31K):
[in this window]
[in a new window]
 
FIG. 2.
Transactivation activity of HBx is not connected with cell death induction. A, schematic presentation of HBx-mutant. The regions necessary for transactivation are indicated as black boxes. These three parts are all required for transactivation activity; mutants lacking one of them lose transactivation ability. B, cell death inducibility was examined with these transactivation deficient mutants. The {beta}-galactosidase activity-positive cells with round and/or blebbing morphology were counted at 3 days after transfection. These two deletion mutants, pCMVX-dAvaI/AvaII (dAI/AII) and pCMVX-{Delta}RsaI ({Delta}RsaI), have similar cell death induction ability. C, transactivation ability of these constructs measured by luciferase assay. Cells were transfected with indicated plasmids along with pRSVL. Two days after transfection, luciferase assay was carried out. D, dose-dependent effect of HBx expression plasmid on cell death inducibility. The number indicated below is the ratio of plasmid pCMVX to plasmid pCMV{beta} (0.102 µg of pCMV{beta} DNA/2-well chamber slide). The total amount of plasmid DNA is adjusted with empty vector, pCMV. E, dose-dependent effect of HBx expression plasmid on transactivation ability. The number indicated below is the ratio of plasmid pCMVX to plasmid pRSVLuc (0.102 µg of pRSVLuc DNA/24-well plate). A 2-well chamber slide and a 24-well plate have the similar growth area, and therefore, the similar experimental condition was applied to compare of cell death inducing effect as shown in Fig. 2D.

 

A Region of HBx Required for Cell Death and Mitochondrial Localization—As reported previously (1) the mitochondrial localization of HBx caused abnormal aggregation of mitochondria, and as described above, that the transactivation function of HBx is not related to cell death induction, we tried to find out the possibility of whether HBx localization on mitochondria is prerequisite for cell death. We transfected an expression plasmid, pEGFP-HBx, that expresses an EGFP-HBx fusion protein, into HuH7 cells and stained with MitoTracker, a dye selectively stains mitochondria. As shown in Fig. 3, 1 day after transfection, EGFP-HBx protein exists all over the cytoplasm, and its location overlaps with the mitochondria. Two to three days after transfection, the localization pattern of EGFP-HBx protein tended to aggregate and gather at the periphery of the nucleus. MitoTracker staining revealed that mitochondria also tended to aggregate at the periphery of the nucleus. This mitochondrial aggregation is co-localized with EGFP-HBx protein. Three to four days after transfection, cells expressing EGFP-HBx protein often show abnormal morphology, round with the membrane blebbing as shown in Fig. 1A. Since EGFP-tagged HBx protein exhibited a similar behavior to that of HBx, we tried to map a region of HBx protein required for localization on mitochondria using plasmid constructs containing EGFP-fused HBx or a series of EGFP-fused HBx mutants (Fig. 4A).



View larger version (48K):
[in this window]
[in a new window]
 
FIG. 3.
Expression profile of EGFP fusion HBx. An expression plasmid, pEGFP-HBx, is transfected into HuH7 cells and stained with MitoTracker (middle panels) on the indicated day. Living cells were examined by fluorescent microscopy. The merged images are shown in the right panels. EGFP-HBx co-localizes with mitochondria.

 


View larger version (51K):
[in this window]
[in a new window]
 
FIG. 4.
EGFP fusion HBx constructs and their localization. A, schematic presentation of EGFP-HBx mutants. As shown in Fig. 2A the regions necessary for transactivation were indicated as black boxes. The region required for cell death inducibility is shown as a blue line. B, amino acid sequence of substitution mutant pEGFP-X(68/117LA). C, the region of HBx required for mitochondrial localization. Fluorescent microscopy of EGFP-HBx proteins is shown. The indicated plasmid was transfected into HuH7 cells, and 2 days after transfection, cells were stained with MitoTracker and monitored under a fluorescent microscopy. The merged images are shown in the right panels. D, cell death inducibility of these EGFP fusion HBx constructs were analyzed as shown in Fig. 1B. The indicated plasmids were transfected into HuH7 cells along with pCMV{beta}. The mutants, which failed to localize on mitochondria, lacked cell death inducibility.

 

As shown in Fig. 4C, 2 days after transfection, a small truncation from the C terminus showed decreased affinity to mitochondria. Further truncation from the C terminus of HBx showed less association with mitochondria (compare Fig. 4C, panels c–e). Truncation of the C-terminal half, EGFP-X-(1–67) (Fig. 4C, panel f) completely lost its ability to localize to mitochondria. One mutant only containing the region of aa 68–117 fused to EGFP localized onto mitochondria (Fig. 4C, panel h), suggesting that aa 68–117 are sufficient for the localization onto mitochondria. We tried to obtain a minimally required region for localization on mitochondria, but further deletion of the aa 68–117 region turned out to be exhibiting diffused distribution in the cytoplasm besides mitochondria (data not shown). We then made substitution mutants instead of deletion mutants. Substitutions of basic or acidic amino acid residues in this region did not give any effect on HBx localization. We also substituted leucine to alanine in this region. One or two substitutions of leucine residues gave only a slight affect on its localization (data not shown). However, when residues Cys69, Leu71, Leu89, Leu100, Leu108, and Leu116 were substituted simultaneously with alanine (Fig. 4B), this mutation resulted in loss of mitochondrial localization (Fig. 4C, panel i), suggesting that hydrophobic residues in this region are important for localization of HBx on mitochondria.

These HBx plasmid constructs were co-transfected to HuH7 cells with {beta}-galactosidase expression plasmid, and then their cell death-inducing activity was examined. As shown in Fig. 4D, plasmids that expressed proteins locating in mitochondria, pEGFP-HBx, -X(1–117), -X(1–104), -X(1–87), and -X(68–117), were able to induce cell death. On the other hand, plasmids pEGFP-X(1–67), -X(118–154), and -X(68–117LA), which do not express proteins associating with mitochondria, were unable to induce cell death. The cell death inducibility of these proteins is not due to the level of protein expression. Because Western blot analysis showed that the truncation mutants, which are unable to induce cell death, tend to express more proteins in cells than EGFP-HBx (data not shown). Results clearly indicate that mitochondrial localization of HBx as well as its mutant proteins are important for their cell death-inducing activity.

Association of HBx with Isolated Mitochondria Is Unable to Release Cytochrome c and AIF—Then we prepared His-tagged recombinant HBx and its mutant proteins (Fig. 5A) and tested their ability to associate with isolated mouse mitochondria in vitro. After 1-h incubation, the reaction mixture was centrifuged, and the precipitated mitochondrial fraction was analyzed by Western blot with anti-His antibody (Fig. 5B). Recombinant HBx-related proteins that contain the aa 68–117 region (rHBx-(aa 1–154) and rX(68–117)) were co-sedimented with mitochondria; however, the mutant protein lacking this region, rX(1–67), was not co-sedimented. Data support a notion that the aa 68–117 region is important for association with mitochondria and is also consistent with the cytochemical data obtained with EGFP-fused HBx constructs (Fig. 4C). On the other hand, the supernatant was analyzed for redistribution of cytochrome c and AIF by Western blot (Fig. 5C). Although cytochrome c and AIF are released from isolated mitochondria after stimulation with CaCl2, as described previously (33), cytochrome c and AIF release from mitochondria are not significantly observed by incubation with recombinant HBx or any HBx mutant proteins. Data indicate that HBx is unable to directly induce cytochrome c and AIF release from isolated mitochondria. As it has been well characterized released cytochrome c, Apaf-1, procaspase-9, and ATP or dATP form a complex, named apoptosome (26), which activates caspase-3, and induces rapid apoptotic cell death. The lack of cytochrome c release suggests that HBx-induced cell death is caspase-3-independent pathway. To confirm this, HuH7 cells were transfected with pCMVX and pCMV{beta} and were incubated with caspase inhibitors, z-DEVD-fmk or z-VAD-fmk (Fig. 5D). A specific caspase-3 inhibitor, z-DEVD-fmk, did not prevent HBxinduced cell death, whereas a general caspase inhibitor, z-VAD-fmk, inhibited HBx-induced cell death in dose-dependent manner. Results clearly indicate that HBx induces cell death through a cytochrome c/caspase-3-independent pathway.



View larger version (54K):
[in this window]
[in a new window]
 
FIG. 5.
Localization of HBx on mitochondria does not relocate cytochrome c and AIF from mitochondria. A, purified recombinant HBx and mutant proteins. Three-hundred ng of each protein was separated on SDS-PAGE, and Coomassie staining was carried out. Lane 1, rHBx; lane 2, rX(1–67); lane 3, rX(68–117). B, co-precipitation experiments with isolated mitochondria and recombinant HBx or its mutants. Isolated mitochondria from mouse liver were incubated with the indicated recombinant protein at 30 °C for 1 h, and then the reaction mixture was centrifuged. The precipitated mitochondria fraction was analyzed by Western blot using anti-His antibody. Lanes 1, 5, and 9, control, mitochondria fraction without protein incubation; lanes 2, 6, and 10, incubated with a 0.1 µM concentration of the indicated recombinant protein and isolated mitochondria; lanes 3, 7, and 11, incubated with a 1 µM concentration of the indicated recombinant protein and isolated mitochondria; lanes 4, 8, and 12, incubated with a 1 µM concentration of the indicated recombinant protein without mitochondria. C, cytochrome c and AIF release from mitochondria. Supernatants were analyzed by Western blot using anti-cytochrome c and AIF antibodies. Lane 1, control, without protein incubation; lanes 2, 4, and 6, incubated with a 0.1 µM concentration of the indicated recombinant protein; lanes 3, 5, and 7, incubated with a 1 µM concentration of the indicated recombinant protein; lane 8, incubated with 150 µM CaCl2; lane 9, mitochondria fraction. Little cytochrome c and AIF were released after HBx association with mitochondria. D, effect of caspase inhibitor on HBx-induced cell death. After pCMVX and pCMV{beta} transfection, cells were incubated with z-VAD-fmk (lane 3, 10 µM; lane 4, 50 µM; lane 5, 100 µM) or z-DEVD-fmk (lane 6, 100 µM), and then cell death inducibility was analyzed 3 days after transfection. A caspase-3 inhibitor, z-DEVD-fmk, has no effect on HBx-induced cell death.

 

Association of HBx with Mitochondria Degenerates Mitochondrial Membrane Potential—The lack of cytochrome c and AIF release from mitochondria suggested that alternative downstream targets are causally involved in cell death. As we reported previously (1) that decreased mitochondrial membrane potential was observed after expression of HBx, we next tested whether localization of HBx on mitochondria is directly relevant to decreased mitochondrial membrane potential. A cytochemical analysis was carried out to monitor mitochondrial membrane potential by a fluorescent dye, 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolcarbocyanine iodide (JC-1) staining, which is a reliable and sensitive method to analyze {Delta}{Psi} changes of mitochondria. JC-1 emits at 527 nm after excitation at 490 nm in monomeric form, and in the presence of a high {Delta}{Psi}, JC-1 forms so called J-aggregates that are associated with a large shift in emission to 590 nm (34, 35). To analyze mitochondrial membrane potential by JC-1, EBFP-fused HBx constructs were utilized to avoid interference of the green fluorescence with JC-1 staining. As shown in Fig. 6, expression of pEBFP-HBx and -X(68–117) (Fig. 6, b and d) (these proteins were localized on mitochondria), caused loss of mitochondrial membrane potential, whereas expression of pEBFP-X(1–67) (Fig. 6c) did not induce any change of mitochondrial membrane potential. Data indicate that association of HBx with mitochondria brings about decreased mitochondrial membrane potential.



View larger version (55K):
[in this window]
[in a new window]
 
FIG. 6.
Localization of HBx on mitochondria causes decrease of mitochondrial membrane potential. Mitochondrial membrane potential was analyzed by JC-1 dye, which is sensitive to {Delta}{Psi}m changes. The indicated EBFP fusion HBx constructs were transfected into HuH7 cells. Two days after transfection, cells were stained with JC-1 and then monitored under fluorescent microscope. Left panels, phase contrast images; middle panels, the expression of EBFP-X constructs; right panels, JC-1 staining. Arrows indicate cells transfected with plasmid.

 

Stabilization of {Delta}{Psi}m by PTP Inhibitors, ROS Scavengers, and Bcl-xL Rescues Cells from HBx-induced Cell Death—If decreased mitochondrial membrane potential is a prerequisite to HBx-induced cell death, stabilization of {Delta}{Psi}m is expected to prevent HBx-induced cell death. To assess this hypothesis, two PTP inhibitors, bongkrekic acid and cyclosporin A, which stabilize mitochondrial membrane potential, were subjected to analysis of HBx-induced cell death. After DNA transfection, these reagents were added to the culture medium, and cell death was monitored 3 days after transfection. As shown in Fig. 7, A and B, both PTP inhibitors prevented cells from HBx-induced cell death in a dose-dependent manner, suggesting that decreased mitochondrial membrane potential is a major cause of HBx-induced cell death. It has been reported that decreased mitochondrial membrane potential results in a variety of deleterious sequeale including generation of ROS and decreased production of ATP. As the generation of ROS has been shown to accelerate cell death events, we analyzed effects of ROS scavenger, NAC, and Tiron on HBx-induced cell death. As shown in Fig. 7, C and D, these ROS scavengers, NAC and Tiron, also clearly prevented cells from HBx-induced cell death in a dose-dependent manner. Data strongly suggest that HBx association with mitochondria opens the PTP complex and causes ROS generation, resulting in cell death.



View larger version (31K):
[in this window]
[in a new window]
 
FIG. 7.
HBx-induced cell death is rescued by PTP inhibitors, ROS scavenger and Bcl-xL. Cell death assays were performed as shown in Fig. 1B with two PTP inhibitors, bongkrekic acid (A) and cyclosporin A (B), and with two ROS scavengers, NAC (C) and Tiron (D). These inhibitors were added after DNA transfection, and cells were stained and assessed for cell death assay 3 days after transfection. A, the effect of bongkrekic acid (BA). Lane 1, control, cells transfected with pCMV, without bongkrekic acid incubation; lane 2, cells transfected with pCMV incubated with 50 µM bongkrekic acid; lanes 3–6, cells transfected with X expression plasmid, pCMVX; lane 3, 0 µM bongkrekic acid; lane 4, 10 µM bongkrekic acid; lane 5, 20 µM bongkrekic acid; lane 6, 50 µM bongkrekic acid incubation. B, the effect of cyclosporin A (CsA). Lane 1, control, cells transfected with pCMV, without cyclosporin A incubation; lane 2, cells transfected with pCMV incubated with 1.0 µM cyclosporin A; lanes 3–6, cells transfected with pCMVX; lane 3, without cyclosporin A; lane 4, 0.1 µM cyclosporin A; lane 5, 0.5 µM cyclosporin A; lane 6, 1.0 µM cyclosporin A incubation. C, the effect of NAC. Lane 1, control, cells transfected with pCMV, 0 mM NAC; lanes 2–5, cells transfected with pCMVX; lane 2, 0 mM NAC; lane 3, 1 mM NAC; lane 4, 5 mM NAC; lane 5, 10 mM NAC incubation. D, the effect of Tiron. Lane 1, control, cells transfected with pCMV, 0 mM Tiron; lanes 2–5, cells transfected with pCMVX. Lane 2, 0 mM Tiron; lane 3, 0.5 mM Tiron; lane 4, 1 mM Tiron; lane 5, 5 mM Tiron incubation. E, Bcl-xL prevents HBx-induced cell death. HBx and Bcl-xL expression plasmids were co-transfected into HuH7 cells. Lane 1, control, cells transfected with vector, pCMV, alone; lane 2, cells transfected with Bcl-xL expression plasmid, pCMV Bcl-xL; lanes 3–6, cells transfected with pCMVX; lane 3, without pCMV Bcl-xL; lane 4, co-transfected with a quarter amount of pCMV Bcl-xL; lane 5, co-transfected with a half-amount of pCMV Bcl-xL; lane 6, co-transfected with equal amount of pCMV Bcl-xL.

 

As it is known that anti-apoptotic proteins Bcl-2 and Bcl-xL stabilize mitochondrial membrane potential and protect cells from apoptosis, if HBx induces loss of mitochondrial membrane potential, excess expression of Bcl-2 and Bcl-xL protect cells from HBx-induced cell death. A Bcl-xL expression plasmid was co-transfected with HBx expression plasmid, and then the extent of cell death was analyzed. Fig. 7E clearly shows that expression of Bcl-xL protein prevents cells from dying under the conditions of HBx expression in dose-dependent manner, confirming that loss of mitochondrial membrane potential is principally related to HBx-induced cell death.

To further confirm that the above reagents preserve mitochondrial membrane potential under the conditions used, HuH7 cells are transfected with pEBFP-X and incubated with 50 µM bongkrekic acid (Fig. 8b). Although cells transfected with HBx expression plasmid showed decreased mitochondrial membrane potential (Fig. 8a), incubation with bongkrekic acid protected cells from degeneration of mitochondrial membrane potential. As shown Fig. 8b, some HBx-expressing cells treated with bongkrekic acid show high mitochondrial membrane potential even after their mitochondria are aggregated. These HBx-expressing cells with high {Delta}{Psi}m were not observed without bongkrekic acid incubation. The expression of Bcl-xL also exhibited a similar effect on stabilization of mitochondrial membrane potential (Fig. 8d). Data also strongly suggested that destabilization of mitochondrial membrane potential, which opens the PTP complex, is the main cause of HBx-induced cell death.



View larger version (64K):
[in this window]
[in a new window]
 
FIG. 8.
Bongkrekic acid (BA) and Bcl-xL prevent loss of {Delta}{Psi}m caused by HBx expression. HuH7 cells were transfected with pEBFP-X, and cells were cultured with (b) or without (a) 50 µM bongkrekic acid. Two days after transfection, cells were stained with JC-1 then monitored under fluorescent microscope. Cells were co-transfected with pEBFP-X and a Bcl-xL expression plasmid, pCMV-Bcl-xL (d), or pEBFP-X and a control vector, pCMV (c). Two days after transfection, cells were stained with JC-1 and monitored. Left panels, phase contrast (PC) images; middle panels, the expression of EBFP-X constructs; right panels, JC-1 staining. Arrows indicate cells expressed with pEGFP-X.

 


    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
First, the present results provided strong evidences that HBx-induced cell death is connected to localization of HBx on mitochondria. We mapped that aa 68–117 are sufficient for HBx localization on mitochondria and cell death induction. Our data clearly show that cell death induced by HBx is independent of its transactivation activity. Although several groups have pointed out that the pro-apoptotic effect of HBx correlates with its transactivation activity (23, 36, 37), our result indicates that the transactivation ability of HBx is not required for its cell death activity, suggesting that the mechanism of HBxinduced cell death is not due to transactivation of the cell death-promoting genes, such as bax.

Previously we reported that expression of HBx induced partial release of cytochrome c from aggregated mitochondria (1). Compared with pro-apoptotic proteins, such as Bax, cytochrome c release induced by HBx is a relatively late event. Although HBx localized on mitochondria right after its expression, cytochrome c release from aggregated mitochondria was only observed 3 days after transfection. Although cytochrome c release was observed from a part of aggregated mitochondria, the majority of mitochondria still preserve cytochrome c. Our analysis in this paper revealed that HBx association with isolated mitochondria did not cause redistribution of cytochrome c and AIF. These results suggested that HBx-induced cell death is dependent on mitochondria, but independent of caspase-3. Involvement of other caspase(s) remains to be analyzed.

In general, mitochondrial membrane permeabilization is a near-universal hallmark and a critical step of several apoptotic pathways (25). During the decision or commitment phase of apoptosis, mitochondrial membrane permeabilization differentially affects the outer and inner membranes of mitochondria. Outer mitochondrial membrane permeabilization becomes protein-permeable and then releases proteins that reside in the intermembrane space, such as cytochrome c and AIF. The redistribution of cytochrome c through the outer mitochondrial membrane is the critical event responsible for mitochondria-dependent caspase activation. The inner mitochondrial membrane permeabilization continues to retain matrix proteins, but dissipates the mitochondrial transmembrane potential. Our data show that HBx does not significantly release cytochrome c and AIF from the mitochondrial intermembrane space in vitro, although the association of HBx with mitochondria caused decreased mitochondrial membrane potential in vivo. These observations suggest that HBx only affects the inner mitochondrial membrane permeabilization but not the outer membrane permeabilization. Furthermore, we showed that PTP inhibitors, ROS scavengers and Bcl-xL, which stabilize mitochondrial membrane potential, rescued cells from HBx-induced cell death, strongly supporting the idea that decreased mitochondrial membrane potential, the inner membrane permeabilization, is the major cause of HBx-induced cell death. Although the temporary order of outer and inner mitochondrial membrane permeabilization, as well as their relative contribution to cell death, are still a matter of debate, the HBx-induced cell death shows that the inner membrane permeabilization of mitochondria is implicated in cell death.

We examined the possibility that if HBx and Bcl-xL proteins physically interact with the same mitochondrial protein using isolated mouse liver mitochondria, we could not see any competition between these proteins in their association with mitochondria (data not shown). This suggests that Bcl-xL is able to indirectly prevent HBx-induced cell death, such as stabilization of mitochondrial membrane potential. It has been shown that several viral and bacterial proteins regulate cell death at the mitochondrial level by targeting proteins to mitochondrial membranes. Rahmani et al. (2) reported that HBx interacts with one particular voltage-dependent anion channel (VDAC) isoform, VDAC-3, by a yeast two-hybrid system. The VDAC is one of the PTP complex proteins in the outer mitochondrial membrane. Our results suggested that HBx affects only inner membrane permeabilization, but it is possible to speculate that the interaction of HBx with VDAC-3 may cause inner membrane permeabilization.

Programmed cell death or apoptosis plays a major role in normal development and physiology as well as in many pathological states. Malfunctions of apoptosis have been implicated in many human diseases such as cancer, neurodegenarative disease, AIDS, and ischemic stroke (3843). It is plausible that during the stage of chronic HBV infection, HBx expression may induce or sensitize apoptosis in infected hepatocytes generating HBV pathogenesis such as inflammation and favor propagation of the viral particles. Such a mechanism has already been proposed for other virus infections (44, 45).


    FOOTNOTES
 
* This work was supported in part by a grant-in-aid from the Ministry of Education, Culture, Sports, Science and Technology and by a grant-in-aid from the Ministry of Health, Labor and Welfare, Japan (to K. K.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

{ddagger} To whom correspondence should be addressed. Tel.: 81-3-5394-3813; Fax: 81-3-5394-3902; E-mail: kkoike{at}jfcr.or.jp.

1 The abbreviations used are: HBV, hepatitis B virus; HBx, hepatitis B virus X protein; AIF, apoptosis-inducing factor; PTP, permeability transition pore; ROS, reactive oxygen species; VDAC, voltage-dependent anion channel; Tiron, 4,5-dihydroxy-1,3-benzenedisulfonic acid; TUNEL, terminal deoxynucleotidyltransferase-mediated dUTP nick end labeling; EGFP, enhanced green fluorescent protein; EBFP, enhanced blue fluorescent protein; z-VAD-fmk, z-Val-Ala-Asp-fluoromethylketone; z-DEVD-fmk, z-Asp-Glu-Val-Asp-fluoromethylketon; aa, amino acid(s); RSV, Rous sarcoma virus; NAC, N-acetyl-L-cysteine. Back


    ACKNOWLEDGMENTS
 
We thank M. Kobayashi for preparing and reading the manuscript.



    REFERENCES
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Takada, S., Shirakata, Y., Kaneniwa, N., and Koike, K. (1999) Oncogene 18, 6965–6973[CrossRef][Medline] [Order article via Infotrieve]
  2. Rahmani, Z., Huh, K. W., Lasher, R., and Siddiqui, A. (2000) J. Virol. 74, 2840–2846[Abstract/Free Full Text]
  3. Blum, H. E., Wieland, S., Walter, E., von Weizsacker, F., Offensperger, W-B., and Moradpour, D. (1998) in Hepatitis B Virus, Molecular Mechanisms in Disease and Novel Strategies for Therapy (Koshy, R., and Caselmann, W. H., eds) pp. 75–92, Imperial College Press, London
  4. Shirakata, Y., Kawada, M. Fujiki, Y., Sano, H., Oda, M., Yaginuma, K., Kobayashi, M., and Koike, K. (1989) Jpn. J. Cancer Res. 80, 617–621[Medline] [Order article via Infotrieve]
  5. Kim, C. M., Koike, K., Saito, I., Miyamura, T., and Jay, G. (1991) Nature 351, 317–320[CrossRef][Medline] [Order article via Infotrieve]
  6. Slagle, B. L., Lee, T. H., Medina, D., Finegold, M. J., and Butel, J. S. (1996) Mol. Carcinog. 15, 261–269[CrossRef][Medline] [Order article via Infotrieve]
  7. Madden, C. R., Finegold, M. J., and Slagle, B. L. (2001) J. Virol. 75, 3851–3858[Abstract/Free Full Text]
  8. Zoulim, F., Saputelli, J., and Seeger, C. (1994) J. Virol. 68, 2026–2030[Abstract]
  9. Bouchard, M. J., Wang, L. H., and Schneider, R. J. (2001) Science 294, 2376–2378[Abstract/Free Full Text]
  10. Twu, J. S., and Robinson, W. S. (1989) Proc. Natl. Acad. Sci. U. S. A. 86, 2046–2050[Abstract]
  11. Aufiero, B., and Schneider, R. J. (1990) EMBO J. 9, 497–504[Abstract]
  12. Arii, M., Takada, S., and Koike, K. (1992) Oncogene 7, 397–403[Medline] [Order article via Infotrieve]
  13. Kekule, A. S., Lauer, U., Weiss, L., Luber, B., and Hofschneider, P. H. (1993) Nature 361, 742–745[CrossRef][Medline] [Order article via Infotrieve]
  14. Luber, B., Lauer, U., Weiss, L., Hohne, M., Hofschneider, P. H., and Kekule, A. S. (1993) Res. Virol. 144, 311–321[Medline] [Order article via Infotrieve]
  15. Benn, J., and Schneider, R. J. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 10350–10354[Abstract/Free Full Text]
  16. Lee, Y. H., and Yun, Y. (1998) J. Biol. Chem. 273, 25510–25515[Abstract/Free Full Text]
  17. Waris, G., Huh, K. W., and Siddiqui, A. (2001) Mol. Cell. Biol. 21, 7721–7730[Abstract/Free Full Text]
  18. Yen, T. S. (1996) J. Biomed. Sci. 3, 20–30[Medline] [Order article via Infotrieve]
  19. Maguire, H. F., Hoeffler, J. P., and Siddiqui, A. (1991) Science 252, 842–844[Medline] [Order article via Infotrieve]
  20. Lin, Y., Nomura, T., Cheong, J., Dorjsuren, D., Iida, K., and Murakami, S. (1997) J. Biol. Chem. 272, 7132–7139[Abstract/Free Full Text]
  21. Shamay, M., Borak, O., Doitsh, G., Ben-Dor, I., and Shaul, Y. (2002) J. Biol. Chem. 277, 9982–9988[Abstract/Free Full Text]
  22. Su, F., and Schneider, R. J. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 8744–8749[Abstract/Free Full Text]
  23. Bergametti, F., Prigent, S., Luber, B., Benoit, A., Tiollais, P., Sarasin, A., and Transy, C. (1999) Oncogene 18, 2860–2871[CrossRef][Medline] [Order article via Infotrieve]
  24. Green, D. R., and Reed, J. C. (1998) Science 281, 1309–1312[Abstract/Free Full Text]
  25. Kroemer, G., and Reed, J. C. (2000) Nat. Med. 6, 513–519[CrossRef][Medline] [Order article via Infotrieve]
  26. Li, P., Nijhawan, D., Budihardjo, I., Srinivasula, S. M., Ahmad, M., Alnemri, E. S., and Wang, X. (1997) Cell 91, 479–489[Medline] [Order article via Infotrieve]
  27. Zou, H., Li, Y., Liu, X., and Wang, X. (1999) J. Biol. Chem. 274, 11549–11556[Abstract/Free Full Text]
  28. Wang, X. (2001) Genes Dev. 15, 2922–2933[Free Full Text]
  29. Shimizu, S., Eguchi, Y., Kamiike, W., Funahashi, Y., Mignon, A., Lacronique, V., Matsuda, H., and Tsujimoto, Y. (1998) Proc. Natl. Acad. Sci. U. S. A. 95, 1455–1459[Abstract/Free Full Text]
  30. Narita, M., Shimizu, S., Ito, T., Chittenden, T., Lutz, R. J., Matsuda, H., and Tsujimoto, Y. (1998) Proc. Natl. Acad. Sci. U. S. A. 95, 14681–14686[Abstract/Free Full Text]
  31. Bradford, M. M. (1976) Anal. Biochem. 72, 248–254[CrossRef][Medline] [Order article via Infotrieve]
  32. Bright, J., and Khar, A. (1994) Biosci. Rep. 14, 67–81[Medline] [Order article via Infotrieve]
  33. Andreyev, A., and Fiskum, G. (1999) Cell Death Differ. 6, 825–832[CrossRef][Medline] [Order article via Infotrieve]
  34. Reers, M., Smiley, S. T., Mottola-Hartshorn, C., Chen, A., Lin, M., and Chen, L. B. (1995) Methods Enzymol. 260, 406–417[Medline] [Order article via Infotrieve]
  35. Salvioli, S., Ardizzoni, A., Franceschi, C., and Cossarizza, A. (1997) FEBS Lett. 411, 77–82[CrossRef][Medline] [Order article via Infotrieve]
  36. Sirma, H., Giannini, C., Poussin, K., Paterlini, P., Kremsdorf, D., and Brechot, C. (1999) Oncogene 18, 4848–4859[CrossRef][Medline] [Order article via Infotrieve]
  37. Schuster, R., Gerlich, W. H., and Schaefer, S. (2000) Oncogene 19, 1173–1180[CrossRef][Medline] [Order article via Infotrieve]
  38. Dragunow, M., MacGibbon, G. A., Lawlor, P., Butterworth, N., Connor, B., Henderson, C., Walton, M., Woodgate, A., Hughes, P., and Faull, R. L. (1997) Rev. Neurosci. 8, 223–265[Medline] [Order article via Infotrieve]
  39. Mattson, M. P., Culmsee, C., and Yu, Z. F. (2000) Cell Tissue Res. 301, 173–187[CrossRef][Medline] [Order article via Infotrieve]
  40. Roshal, M., Zhu, Y., and Planelles, V. (2001) Apoptosis 6, 103–116[CrossRef][Medline] [Order article via Infotrieve]
  41. Orth, M., and Schapira, A. H. (2001) Am. J. Med. Genet. 106, 27–36[CrossRef][Medline] [Order article via Infotrieve]
  42. Johnstone, R. W., Ruefli, A. A., and Lowe, S. W. (2002) Cell 108, 153–164[Medline] [Order article via Infotrieve]
  43. Green, D. R., and Evan, G. I. (2002) Cancer Cell 1, 19–30[CrossRef][Medline] [Order article via Infotrieve]
  44. Collins, M. (1995) Am. J. Respir. Crit. Care Med. 152, S20–S24[Medline] [Order article via Infotrieve]
  45. Roulston, A., Marcellus, R. C., and Branton, P. E. (1999) Annu. Rev. Microbiol. 53, 577–628[CrossRef][Medline] [Order article via Infotrieve]