The Chicken Anemia Virus-Derived Protein Apoptin Requires Activation of Caspases for Induction of Apoptosis in Human Tumor Cells

Abstract

The chicken anemia virus protein Apoptin has been shown to induce apoptosis in a large number of transformed and tumor cell lines, but not in primary cells. Whereas many other apoptotic stimuli (e.g., many chemotherapeutic agents and radiation) require functional p53 and are inhibited by Bcl-2, Apoptin acts independently of p53, and its activity is enhanced by Bcl-2. Here we study the involvement of caspases, an important component of the apoptotic machinery present in mammalian cells. Using a specific antibody, active caspase-3 was detected in cells expressing Apoptin and undergoing apoptosis. Although Apoptin activity was not affected by CrmA, p35 did inhibit Apoptin-induced apoptosis, as determined by nuclear morphology. Cells expressing both Apoptin and p35 showed only a slight change in nuclear morphology. However, in most of these cells, cytochrome c is still released and the mitochondria are not stained by CMX-Ros, indicating a drop in mitochondrial membrane potential. These results imply that although the final apoptotic events are blocked by p35, parts of the upstream apoptotic pathway that affect mitochondria are already activated by Apoptin. Taken together, these data show that the viral protein Apoptin employs cellular apoptotic factors for induction of apoptosis. Although activation of upstream caspases is not required, activation of caspase-3 and possibly also other downstream caspases is essential for rapid Apoptin-induced apoptosis.

Although many viruses encode apoptotic inhibitors, a number of viruses have been found to carry genes specifying apoptosis-inducing proteins (35, 39, 41). Apoptin, a 13.6-kDa protein encoded by the chicken anemia virus, is one such gene product. In cell culture, expression of Apoptin is sufficient to induce apoptosis (27). Interestingly, Apoptin only induces apoptosis in transformed or tumor-derived cells and not in normal diploid or primary cells of human or rodent origin (9; Y. Zhuang, unpublished results). In contrast to most chemotherapeutic agents, Apoptin induces apoptosis in cells lacking functional p53 or overexpressing Bcl-2 (47, 48). When cotransfected, Bcl-2 even enhances Apoptin activity (8, 10). In order to understand how Apoptin induces apoptosis, further insight into the involvement of known apoptotic effectors is required.

Several observations indicate that the mitochondria play an important role in the commitment to programmed cell death (13, 15, 19). Many apoptotic stimuli (e.g., Bax, oxidants, and high Ca²⁺) induce a loss of mitochondrial membrane integrity. Following a drop in the mitochondrial inner membrane potential (ΔΨ_m), it is thought that either a permeability transition pore opens or the outer mitochondrial membrane is physically disrupted. In either case, this results in release of proapoptotic molecules from the intermembrane space, such as procaspases (24, 36), apoptosis-inducing factor (37), and cytochrome c, which can act as a cofactor for caspase activation (22). Disruption of the mitochondrial membrane also leads to a drop in cellular ATP and production of reactive oxygen species, although this seems to occur relatively late in apoptosis (15). Antiapoptotic Bcl-2 family members, like Bcl-2 and Bcl-x_L, which block cytochrome c release from mitochondria and inhibit opening of the permeability transition pore, can completely rescue cells from cell death induced by many different stimuli (1, 31, 42). However, not all apoptotic stimuli are inhibited by Bcl-2. It has been proposed that there is also a mitochondrion-independent pathway, feeding directly into the caspase cascade, which is not inhibited by Bcl-2 (33).

Caspases play a major role in the execution phase of apoptosis (7, 14, 28, 40) by cleaving a large number of proteins, which in turn leads to the typical morphology of apoptosis. Among these substrates are cytoskeletal and structural proteins, DNA repair enzymes, transcription factors, protein kinases, and proteins involved in cell cycle regulation (C. Stroh and K. Schulze-Osthoff, Editorial, Cell Death Differ. 5:997–1000, 1998). Also, some of the antiapoptotic Bcl-2 family members have been found to be cleaved by caspases (5, 6). Caspases all cleave after an aspartic acid residue. Specificity is largely determined by the tetrapeptide directly N terminal to the cleavage site (26). Caspases exist as inactive zymogens in the cell which become activated upon proteolytic cleavage by other caspases or by autocatalysis. Functionally, they can be divided into initiator (upstream) and effector (downstream) caspases. Different apoptotic signals activate different initiator caspases, in turn activating the effector caspases, resulting in a cascade of caspase activation. Cleavage of procaspases can be regulated by self-oligomerization (44), compartmentalization (24, 36), the availability of cofactors like cytochrome c (22), and the presence of cellular inhibitors (11, 32). It has been shown that caspases can activate cytosolic factors, e.g., Bid, which induce the release of cytochrome c from mitochondria, possibly acting as an amplification loop during apoptosis (2, 21, 23).

For viruses, blocking apoptosis is a way to circumvent the cellular defense mechanism against viral infection, and many of them have evolved their own caspase inhibitors, like CrmA from cowpox virus (30) and p35 from baculovirus (4). However, caspase inhibitors have also been found in mammals; for example, the IAP (inhibitor of apoptosis) family has both mammalian and viral homologs (11, 32, 34). Inhibition of caspase activation blocks the appearance of apoptotic morphology, illustrating the important role of caspases in the execution phase of apoptosis. However, blocking caspases does not necessarily lead to cell survival. In several cases, the apoptotic morphology is inhibited but clonogenicity is lost, and eventually the cells still die, albeit more slowly (16, 25). These results imply that the commitment to undergo programmed cell death is made upstream of the activation of the caspase cascade.

In this study, we used several inhibitors of caspases with different specificities to determine the involvement of caspases and mitochondrial factors in Apoptin-induced apoptosis. Apoptin-induced apoptosis exhibits remarkable specificity for transformed or tumorigenic cells. Therefore, determining how Apoptin interacts with the cell's normal apoptotic pathways is important. Evidence is provided that Apoptin can indeed utilize the cell's effector caspases yet remains independent of the initiator caspase pathways.

MATERIALS AND METHODS

Cell culture.

The human osteosarcoma cell line Saos-2, which lacks functional p53, was cultured in Dulbecco's modified Eagle's medium containing 10% fetal calf serum, penicillin, and streptomycin (Life Technologies, Rockville, Md.). One day prior to transfection, the cells were seeded in dishes containing uncoated glass slides. At the time of transfection, the cells were 30 to 50% confluent.

Plasmids and transfection.

The plasmids pCMV-VP3, encoding Apoptin, pCMV-p53, encoding p53, and pCMV-neo-Bam, the empty control plasmid, have been described previously (27, 47). The cDNAs for CrmA and p35 (kindly donated by D. J. Pickup and L. K. Miller, respectively) were each subcloned into the BamHI site of pCMV and confirmed by restriction enzyme analysis and sequencing, generating pCMV-CrmA and pCMV-p35. Expression of CrmA from pCMV-CrmA was shown by an in vitro transcription-translation assay (S. Olijslagers, unpublished results). Expression of p35 in Saos-2 cells transfected with pCMV-p35 was shown by Western blot analysis using a specific antibody against p35 (kindly donated by L. K. Miller).

phGFP-S65T (Clontech, Palo Alto, Calif.) was used to generate phGFP-VP3, fusing Apoptin to the C terminus of green fluorescent protein (GFP), under the control of a cytomegalovirus promoter. The plasmid was tested by sequencing, and expression of the fusion protein was confirmed by Western blot analysis. In transfection assays, GFP-Apoptin had the same localization and activity as wild-type Apoptin in tumor cells (A. van Zon, unpublished results). pcDNA3.1/ MycHis/LacZ (Invitrogen, Carlsbad, Calif.) encodes LacZ with both a myc tag and a His tag attached to the C terminus. All plasmids were purified with Jetstar maxiprep columns (Genomed, Bad Oeyenhausen, Germany). Saos-2 cells were transfected by the CaPO₄-method as described previously (43), with 5 to 6 μg of plasmid DNA per 6-cm-diameter dish or 3 μg per well of a 6-well plate or 3.5-cm-diameter dish. In cotransfections, the ratio of pCMV-VP3 to pCMV-CrmA or pCMV-p35 was always 1:2.

Immunofluorescence assays and antibodies.

Two to 5 days after transfection, cells were fixed with 80% acetone for 10 min and kept at −20°C until further staining. For the antibody staining, the cells were first incubated with phosphate-buffered saline (PBS) plus 0.05% Tween 20 (PBS-Tween) plus 5% normal goat serum (NGS) for 30 min, incubated with the first antibody in PBS-Tween plus 5% NGS for 1 h, washed with PBS-Tween, and incubated with the second antibody in PBS-Tween plus 5% NGS for 1 h. Finally, the cells were washed with PBS-Tween and embedded in 90% glycerol–0.1 M Tris (pH 8.0) containing 2.3% Dabco (diazabicyclo-[2,2,2]-octane) to prevent quenching of the signal and 1 μg of DAPI (2,4-diamidino-2-phenylindole)/ml to stain the DNA. The cells were analyzed by fluorescence microscopy for expression of the transfected protein, and nuclear morphology indicating the apoptotic state of the cell was determined by DAPI staining. Staining with anti-cytochrome c was done in essentially the same way, except that fixation of the cells was done with 50% methanol–50% acetone for 5 min, incubation with the first antibody was done for 3 h instead of 1 h, and 5% NGS was replaced with 3% bovine serum albumin.

For propidium iodide (PI) exclusion, cells were first incubated with 5 μg of PI/ml in the medium for 20 min, washed with PBS, and then fixed with formaldehyde-methanol-acetone (sequential incubation with 1% formaldehyde for 10 min, 100% cold methanol for 5 min, and 80% cold acetone for 2 min). In the staining experiments with the mitochondrion-specific dye CMX-Ros, cells were washed with PBS, stained with 100 nM Mito Tracker Red CMX-Ros (Molecular Probes, Eugene, Oreg.) in PBS for 30 min, washed with PBS, and fixed with formaldehyde-methanol-acetone.

The mouse monoclonal antibody CVI-CAV-111.3 was used to detect Apoptin (supernatant was diluted 1:3) (9). A rabbit polyclonal antibody, RαVP3-C, raised against part of the C terminus of Apoptin, was used in double stainings (dilution, 1:200). p53 expression was detected with the mouse monoclonal antibody DO-1 (dilution, 1:100) (Santa Cruz Biotechnology, Santa Cruz, Calif.). The mouse monoclonal antibody 6H2.B4 was used to detect cytochrome c (dilution, 1:200), and a rabbit polyclonal antibody was used to detect active caspase-3 (dilution, 1:200) (both from Pharmingen, San Diego, Calif.). Second antibodies were conjugated either to fluorescein isothiocyanate or to rhodamine (dilution, 1:100) (Jackson ImmunoResearch Laboratories, West Grove, Pa.).

Subcellular fractionation and Western blot analysis.

Subcellular fractionation was performed essentially as described by Juin et al. (18). Saos-2 cells were grown in 15-cm-diameter dishes and transfected with FuGENE 6 (Roche Molecular Biochemicals, Indianapolis, Ind.) according to the manufacturer's protocol. The cells were washed with PBS and then scraped and centrifuged at 2,000 × g for 5 min. The cells were then resuspended in cell extraction buffer (300 mM sucrose, 10 mM HEPES [pH 7.4], 50 mM KCl, 5 mM EGTA, 5 mM MgCl₂, 1 mM dithiothreitol, 1 mM phenylmethylsulfonyl fluoride, 100 μM cytochalasin B), left on ice for 30 min, and homogenized by 50 strokes in a Dounce homogenizer. Unbroken cells and nuclei were pelleted by centrifugation for 5 min at 2,000 × g. Heavy membranes were removed from the resulting supernatant by centrifugation for 5 min at 13,000 × g. The resulting supernatant was the crude cytosolic fraction. All samples were frozen in liquid nitrogen and stored at −80°C.

For Western blotting, 30 μg of protein was loaded in each lane of a sodium dodecyl sulfide–15% polyacrylamide gel, separated by electrophoresis, and electroblotted onto Immobilon-P membranes (Millipore, Bedford, Mass.). The blots were then incubated with the monoclonal antibody 7H8.2C12 (Pharmingen) (1:1,000) to detect cytochrome c, and positive signals were visualized by enhanced chemiluminescence (Amersham, Piscataway, N.J.) according to the manufacturer's protocol.

RESULTS

Apoptin requires activation of downstream caspases for rapid induction of apoptosis.

The viral inhibitors CrmA and p35 show different specificities for the various caspases. In vitro studies of binding kinetics indicate that CrmA mainly represses upstream caspases, e.g., caspases 1 and 8, and has little effect on the more downstream caspases 3, 6, and 7 (12, 46). Comparable studies of p35 show that it is a more general caspase inhibitor which inhibits both upstream and downstream caspases (45). To determine which caspases are involved in Apoptin-induced apoptosis, plasmids encoding these inhibitors were cotransfected with Apoptin in Saos-2 cells, a human tumor cell line which lacks endogenous p53. Transfection with p53 was used as a positive control for apoptosis induction in these cells. Induction of apoptosis was scored by analysis of nuclear morphology by DAPI staining. Intact nuclei are stained evenly, but apoptotic nuclei are often fragmented and show irregular or weak DNA staining caused by condensation and fragmentation of the DNA (9, 38). Four days posttransfection, coexpression of CrmA did not inhibit Apoptin activity, whereas p53-induced apoptosis was inhibited by coexpression of CrmA by approximately 50% (Fig. 1). In contrast, in the presence of p35, Apoptin-induced apoptosis was almost completely abolished (Fig. 2a). Only a background level of apoptosis was observed, similar to that seen with overexpression of the nonapoptotic control protein Desmin or LacZ (reference 9;0 and data not shown), which is most likely caused by the transfection method. In parallel experiments, p35 also almost completely inhibited p53-induced apoptosis, as expected (Fig. 2b). Incubation for up to 4 days posttransfection with 50 μM zVAD-fmk, a synthetic, broad-spectrum caspase inhibitor with a relatively high affinity for upstream caspases (12), partially inhibited p53-induced apoptosis but had no effect on Apoptin (data not shown). Taken together, these results indicate that activation of downstream, but not upstream, caspases is required for rapid Apoptin-induced apoptosis.

FIG. 1.

CrmA inhibits p53-induced but not Apoptin-induced apoptosis. Saos-2 cells were transfected with plasmids encoding Apoptin or p53, together with a plasmid encoding CrmA or an empty control plasmid (neo). Four days after transfection, the cells were fixed, stained with antibodies recognizing Apoptin or p53, and analyzed by fluorescence microscopy. The percentage of apoptotic cells among those expressing Apoptin or p53 was determined by evaluating the DAPI staining. The values shown are the means of three independent experiments + standard deviations; at least 100 cells were scored per experiment.

FIG. 2.

p35 inhibits both Apoptin-induced and p53-induced apoptosis. Saos-2 cells were transfected with plasmids encoding Apoptin or p53, together with a plasmid encoding p35 or an empty control plasmid (neo). At several time points after transfection, cells were fixed, stained with specific antibodies, and analyzed by fluorescence microscopy. The percentage of apoptotic cells among those expressing Apoptin or p53 was determined by evaluating the DAPI staining. The values shown are the means of three independent experiments ± standard deviations; at least 100 cells were scored per experiment.

Caspase-3 is a downstream caspase that plays a central role in the execution of programmed cell death. Therefore, the next step was to determine whether caspase-3 is activated in Apoptin-expressing cells. Saos-2 cells were transfected with Apoptin, fixed 5 days later, and stained with an antibody specific for active caspase-3. Among the Apoptin-expressing cells, active caspase-3 could be detected in all apoptotic cells but was observed in only 1% of the nonapoptotic cells containing Apoptin (Fig. 3). As expected, upon coexpression of p35, no active caspase-3 was seen (data not shown). These results show directly that caspase-3 is activated in Apoptin-induced cell death. Furthermore, the absence of active caspase-3 in nonapoptotic, Apoptin-positive cells indicates that Apoptin expression precedes caspase-3 activation.

FIG. 3.

Active caspase-3 is present in Apoptin-expressing apoptotic cells. A plasmid encoding Apoptin was transfected into Saos-2 cells. Four days later, the cells were fixed, stained with antibodies recognizing Apoptin or active caspase-3 (casp3), and analyzed by fluorescence microscopy in three independent experiments. Photographs were taken of representative cells: normal, nonapoptotic (a) and apoptotic (b) (magnification, ×1,000). Arrowheads point at Apoptin-positive cells.

Caspase inhibition does not prevent all aspects of Apoptin-induced apoptosis.

Whereas coexpression of p35 with either Apoptin or p53 diminished the number of Saos-2 cells showing the morphology typical of apoptosis at 5 days posttransfection (shrinkage of the nucleus, condensation and breakdown of DNA, and nuclear blebbing), some slight changes in nuclear morphology were still observed in 58% of the cells (Fig. 4). The nuclei of these cells no longer had smooth surfaces but appeared to be dented, and the DNA seemed to be slightly condensed. This was not observed in cells coexpressing p35 and LacZ (data not shown). Although p35 strongly inhibits Apoptin- or p53-induced apoptosis for up to 5 days posttransfection, it appears to be slowing down apoptosis rather than blocking it completely. Others have reported that caspase inhibition prevents certain aspects of apoptosis but does not prevent the cells from dying eventually (16, 25). It is possible that the cells coexpressing p35 and Apoptin with slightly irregular nuclei were dying already. To determine the condition of these cells, we next studied the integrity of the cell membrane by looking at PI exclusion. For this purpose, Saos-2 cells were cotransfected with plasmids encoding p35 and GFP-Apoptin, a fusion protein of GFP and Apoptin which behaves in the same way as wild-type Apoptin in human tumor cells (van Zon, unpublished). Three days posttransfection, the cells were analyzed for PI exclusion. Cells expressing GFP-Apoptin, either with or without p35, did not stain with PI, indicating that the cell membrane was still intact. Cells showing the slight morphological nuclear changes when coexpressing p35 also did not stain with PI (Fig. 5a). Only late apoptotic cells were positive for PI staining (Fig. 5b). These data show that the cells in which apoptosis is impaired by p35 still have an intact cell membrane, despite the irregular characteristics of their nuclei.

FIG. 4.

Coexpression of p35 with Apoptin slightly influences the nuclear morphology. Saos-2 cells were cotransfected with plasmids encoding Apoptin and p35. Five days later, the cells were fixed, stained with an antibody recognizing Apoptin and with DAPI, and analyzed by fluorescence microscopy. Four independent experiments were performed, in each of which at least 100 cells were scored; photographs were taken of representative cells. The arrowheads indicate two cells coexpressing Apoptin and p35 with slight changes in nuclear morphology (magnification, ×1,000).

FIG. 5.

Cells coexpressing GFP-Apoptin and p35 are negative for PI. Plasmids encoding GFP-Apoptin and p35 were cotransfected into Saos-2 cells. Three days later, the cells were first stained with PI, then fixed, and finally stained with DAPI. The cells were analyzed by fluorescence microscopy in three independent experiments, and photographs were taken of representative cells. (a) Cell cotransfected with GFP-Apoptin and p35 with slight changes in nuclear morphology, negative for PI; (b) late apoptotic cell, transfected only with GFP-Apoptin, positive for PI (magnification, ×1,000). Arrowheads point at Apoptin-positive cells.

Apoptin induces cytochrome c release, which is not inhibited by p35.

Release of cytochrome c from mitochondria is a well-known event in apoptosis which is often required for activation of downstream caspases. Therefore, we investigated whether cytochrome c is released from mitochondria during Apoptin-induced apoptosis. Saos-2 cells were transfected with plasmids encoding Apoptin or with LacZ as a negative control. Expression of p53 was used as a positive control for apoptosis induction. Three days later, cytosolic extracts were prepared by subcellular fractionation and analyzed by Western blotting. In cells transfected with Apoptin or p53, the levels of cytochrome c were increased compared to those in cells transfected with LacZ (Fig. 6), which indicates that cytochrome c is released from mitochondria during Apoptin-induced apoptosis. In order to determine the status of the mitochondria on a single-cell level, we examined the release of cytochrome c from mitochondria in cells expressing Apoptin by immunofluorescence microscopy. Saos-2 cells were cotransfected with plasmids encoding Apoptin, fixed 5 days later, and stained with specific antibodies for cytochrome c and Apoptin. All Apoptin-expressing apoptotic cells had lost mitochondrial cytochrome c staining in all planes of focus (Fig. 7b), whereas almost all nonapoptotic Apoptin-positive cells had distinctly punctate mitochondrial staining (Fig. 7a). This indicates that although cytochrome c release takes place, it does not seem to be an early event in Apoptin-induced apoptosis. In contrast, for other apoptotic stimuli, it has been reported that cytochrome c release can be observed before any changes in nuclear morphology (3, 18). When p35 is coexpressed, 67% (mean of three experiments) of the Apoptin-expressing, nonapoptotic cells had lost mitochondrial cytochrome c staining (Fig. 7c). In these cells, cytochrome c had diffuse staining and seemed to be mainly present in the nucleus, unlike the diffuse localization in the cytoplasm or throughout the whole cell reported for other apoptotic stimuli (3, 18). These results indicate that inhibition of the final apoptotic events by p35 does not prevent the release of cytochrome c from the mitochondria, which may be an indirect effect of Apoptin expression.

FIG. 6.

Cytochrome c release in Apoptin-induced apoptosis. Saos-2 cells were transfected with plasmids encoding either LacZ (negative control), Apoptin, or p53 (positive control). Three days later, cytosolic extracts were prepared and analyzed by Western blotting for cytochrome c levels. Similar results were obtained in two independent experiments.

FIG. 7.

Cytochrome c release in Apoptin-induced apoptosis. Saos-2 cells were cotransfected with plasmids encoding Apoptin and p35 or with an empty control. Five days later, the cells were fixed and stained with an antibody recognizing Apoptin or cytochrome c and with DAPI. The cells were analyzed by fluorescence microscopy in four independent experiments; at least 100 cells were examined per experiment. Photographs were taken of representative cells. Cells transfected only with Apoptin which are nonapoptotic (a) or apoptotic (b) and a cell cotransfected with Apoptin and p35 with changed nuclear morphology (c) are shown. Arrowheads point at Apoptin-positive cells. Magnification, ×1,000.

Role of mitochondria in Apoptin-induced apoptosis.

Cytochrome c release can occur in the absence of disruption of ΔΨ_m (3). However, apoptosis is often accompanied by loss of ΔΨ_m. Therefore, we next investigated the status of the mitochondrial membrane potential in these cells. The dye Mito Tracker Red CMX-Ros is only taken up by actively respiring mitochondria with intact ΔΨ_m (19). Saos-2 cells were cotransfected with plasmids encoding p35 and GFP-Apoptin or with GFP as a control. Four days later, the cells were incubated with CMX-Ros, fixed, and examined by immunofluorescence microscopy. In the absence of p35, cells expressing GFP showed punctate staining with CMX-Ros in the cytoplasm (data not shown). In nonapoptotic cells expressing GFP-Apoptin, CMX-Ros still stained the mitochondria; hardly any cells that had lost the mitochondrial staining were observed (Fig. 8a). Apoptotic cells, either expressing GFP-Apoptin or not, did not show punctate staining with CMX-Ros, indicating a collapse in ΔΨ_m (Fig. 8b). This indicates that disruption of ΔΨ_m is not an early event in Apoptin-induced apoptosis but only appears at a late stage. In the presence of p35, 4 days posttransfection, approximately 50% of the nonapoptotic, GFP-Apoptin-expressing cells had lost the mitochondrial staining with CMX-Ros (Fig. 8c). One day later, this number had gone up to 80%, whereas in cells coexpressing GFP and p35, it was still less than 20% (data not shown). This result implies that, although the final apoptotic events are prevented by p35 (at least up to 5 days posttransfection), Apoptin is able to start processes leading to disruption of ΔΨ_m.

FIG. 8.

Staining of mitochondria with CMX-Ros. Saos-2 cells were cotransfected with plasmids encoding GFP-Apoptin and p35 or with an empty control. Four or 5 days later, the cells were first stained with CMX-Ros and then fixed and stained with DAPI. The cells were analyzed by fluorescence microscopy in three independent experiments, in each of which at least 100 cells were examined, and photographs were taken of representative cells. A nonapoptotic (a) and an apoptotic (b) cell transfected only with GFP-Apoptin and a cell coexpressing GFP-Apoptin and p35 with changed nuclear morphology (c) are shown. In the last cell, GFP-Apoptin is also present in the cytoplasm, which occurred more often in the presence of p35 than in its absence. Arrowheads point at Apoptin-positive cells. Magnification, ×1,000.

DISCUSSION

The activation of the caspase cascade plays a central role in most apoptotic pathways. Different apoptotic stimuli can activate different initiator caspases, in turn leading to the activation of downstream effector caspases. Here we studied the role of caspases in apoptosis induced by expression of Apoptin in human tumor cells. Apoptin-induced cell death is not affected by CrmA but is clearly inhibited by p35. The data presented here indicate that Apoptin does not require the activation of the upstream caspases 1 and 8 but that activation of one or more downstream caspases is necessary for the rapid induction of apoptotic cell death. Previously, it had been shown that p53-induced apoptosis can be inhibited by coexpression of p35 in insect cells (29); here we show that this is also the case in human cells.

In immunofluorescence studies, activation of caspase-3 could be demonstrated in Apoptin-expressing apoptotic cells. In contrast, in cells expressing Apoptin that still had a normal nuclear morphology, active caspase-3 was hardly ever observed. These results prove that caspase-3 is activated during Apoptin-induced cell death, but this activation seems to occur at a late stage. In a caspase-3-negative human breast cancer cell line, MCF7 (17, 20), Apoptin does not induce cell death in the same rapid way as in other tumor cell lines and the morphological changes typical of apoptosis are not observed (M. Noteborn, unpublished results). This again suggests that caspase-3 activation is necessary for rapid Apoptin-induced apoptosis. However, Apoptin-expressing MCF7 cells did not look entirely normal either: they displayed condensed DNA and appeared to be dying. Activation of other downstream caspases present in MCF7 cells, e.g., 6 and 7, could eventually lead to cell death. Similarly, the possibility that in Saos-2 cells other downstream caspases are activated by Apoptin in addition to caspase-3 cannot be excluded.

In cells in which Apoptin-induced apoptosis was inhibited by coexpression of p35, a slight change in nuclear morphology was observed. These cells still had intact cell membranes, as determined by PI exclusion. However, in most of these cells, the mitochondrial membrane integrity was disrupted: the mitochondria were no longer stained with CMX-Ros or antibody against cytochrome c. In almost all Apoptin-expressing cells that still had normal nuclear morphology (in the absence of p35), no loss of mitochondrial staining by either CMX-Ros or anti-cytochrome c could be detected, indicating that in these cells, the ΔΨ_m is not disrupted. Coexpression of p35 with LacZ did not have any effect on the nuclear morphology, and coexpression of p35 with GFP had only a minor effect on CMX-Ros staining (data not shown). Therefore, the effect on nuclear morphology and mitochondrial membrane integrity cannot be ascribed to overexpression of p35 in these cells but is primarily caused by overexpression of Apoptin. It had been reported earlier that inhibition of caspases does not prevent cytochrome c release caused by, e.g., UVB irradiation or staurosporine (3). Furthermore, in a number of cases, inhibition of caspases prevents certain morphological aspects of apoptosis, but eventually the cells still die (16, 25). Similarly, in cells coexpressing Apoptin and p35, the slight nuclear changes and loss of mitochondrial membrane integrity suggest that, in the end, these cells may not survive.

From the data obtained here, it cannot be concluded that mitochondria are crucial for Apoptin-induced apoptosis. If they are, it is a late event, which in the absence of p35 is practically only observed in apoptotic Apoptin-expressing cells and could be an effect rather than a cause of apoptosis induction. Furthermore, we have shown previously that Bcl-2, which is thought to inhibit apoptosis by preventing disruption of ΔΨ_m and release of cytochrome c, does not inhibit Apoptin-induced cell death in Saos-2 cells. Rather, overexpression of Bcl-2 even accelerates Apoptin activity (8, 10). A possible explanation could be that Apoptin expression leads to cleavage of Bcl-2 by activated caspases, resulting in a proapoptotic form of Bcl-2 (5). However, a mutant form of Bcl-2 which can no longer be cleaved by caspases can accelerate Apoptin activity to the same extent as wild-type Bcl-2 in Saos-2 cells (data not shown).

In this study, we have shown that cell death induction by Apoptin involves the activation of caspase-3 and possibly other downstream caspases, which also play an essential role in apoptosis induced by many other stimuli. Inactive procaspases are present in most cells, both normal, nontransformed cells and transformed or tumorigenic cells (7, 14, 28, 40). However, Apoptin only induces apoptosis in transformed or tumorigenic cells. We propose two possible explanations for these phenomena. First, Apoptin may be differentially modified in tumor cells so that it can exert its apoptotic activity. Second, the cellular decision machinery that decides whether to enter apoptosis may be different in nontransformed and transformed or tumorigenic cells. It is conceivable that expression or absence of expression of one or more genes in transformed or tumorigenic cells causes them to respond to certain stimuli by undergoing apoptosis. Previous data suggest that the difference in Apoptin activity is correlated with a difference in localization: in normal cells, it is found mainly in the cytoplasm, whereas in transformed or tumorigenic cells it is located predominantly in the nucleus (9). We hypothesize that once Apoptin is in the nucleus (either modified or not), it generates an apoptotic signal that eventually activates downstream caspases, leading to apoptosis.

In summary, we have shown that activation of caspases, especially caspase-3, is necessary for apoptosis induction by Apoptin. The role of mitochondria is still unclear, but they do not seem to be crucial for Apoptin activity. Studies of the modification of Apoptin, its binding factors, and its localization in normal versus tumor cells are in progress. Better understanding of the mechanism of Apoptin-induced apoptosis will be helpful in its use as an antitumor agent and may also provide information on the aspects of a cell that determine its transformed or tumorigenic state.