Caspase-9 plays a marginal role in serum starvation-induced apoptosis

Experimental Cell Research 302 (2005) 115 – 128 www.elsevier.com/locate/yexcr

Caspase-9 plays a marginal role in serum starvation-induced apoptosis Chantal J. Schamberger*, Christopher Gerner, Christa Cerni Institute of Cancer Research, Medical University of Vienna, 1090 Vienna, Austria Received 31 March 2004, revised version received 4 August 2004 Available online 25 September 2004

Abstract Serum withdrawal represents a potent trigger to induce caspase-dependent apoptosis in a series of cell culture models. In rat 423-cells, caspase-8 and caspase-3 were apparently sufficient to initiate and proceed apoptosis without involving the intrinsic amplification loop via caspase-9. To assess the reasons for this inactivity of an otherwise crucial initiator caspase, we examined the ability for apoptosome assembly in 423-cells. Caspase-9 and Apaf-1 were expressed and cytochrome c released from mitochondria upon serum withdrawal. Although functional apoptosomes were assembled successfully in vitro, caspase-9 processing was found essentially refrained during apoptosis in 423-cells. Cell fractionation experiments revealed that sequestration of caspase-9 to cytoskeletal structures in 423-cells contributed to the observed impairment of apoptosome formation. Altogether, these findings provide evidence that serum starvation-induced apoptosis may occur independently of the intrinsic pathway and that caspase-9 sequestration potentially represents a novel biological antiapoptotic strategy. D 2004 Elsevier Inc. All rights reserved. Keywords: Serum withdrawal; Starvation; Caspases; Apoptosome

Introduction Serum or trophic factor withdrawal can induce apoptosis. Cell culture systems sensitive for serum withdrawal can be subdivided in two groups by their rate of apoptotic response. A rapid time course of apoptosis within a few hours is described for undifferentiated PC-12 cells [1], AKR-2B mouse fibroblasts [2], Syrian hamster embryo cells [3], and myocardiacs [4], for example. A slow time course, which may take several days, was described for NGF-differentiated PC-12 cells [1], HUVEC [5] and myc-overexpressing Rat-1 cells [6], for example. Apoptosis induced by serum starvation is generally accepted as caspase dependent. These deathspecific cysteine proteases concert the destruction of initiated cells in a complex network [7]. Caspase-3 and caspase-6 were

* Corresponding author. Institute of Cancer Research, Medical University of Vienna, Borschkegasse 8a, 1090 Vienna, Austria. Fax: +49 1 4277 9651. E-mail address: [email protected] (C.J. Schamberger). 0014-4827/$ - see front matter D 2004 Elsevier Inc. All rights reserved. doi:10.1016/j.yexcr.2004.08.026

identified as executioner caspases after growth factor withdrawal [8,9]. The identity of involved initiator caspases after growth factor withdrawal remains controversial, depending on cell culture systems [9–11]. There are at least two fundamental pathways cross talking for a proper amplification of death signaling. The first one, the so-called extrinsic pathway leads to clustering of death receptors, followed by formation of the death-inducing signaling complex (DISC), and activation of initiator caspases-8 and/or -10. The second, induced upon treatment with cytotoxic drugs, like staurosporine (STS) [12–14], initiates release of cytochrome c and other apoptogenic factors from mitochondria [15]. This socalled intrinsic pathway of caspase activation is characterized by apoptosome assembly, a multimeric complex formed by Apaf-1, cytochrome c, and caspase-9. The cytochrome c initiates caspase-9 processing by formation of the apoptosome [16]. Impairment of one of this branches may confer resistance to or result in a delayed apoptotic response after the apoptotic trigger [17–20]. In this study, we investigated processing of initiator caspases after serum withdrawal in rat 423-cells. Despite the rapid course of apoptosis, the intrinsic caspase-signal-

116

C.J. Schamberger et al. / Experimental Cell Research 302 (2005) 115–128

ing cascade remained inactive, indicating a marginal role for caspase-9 in serum-starvation induced apoptosis. We investigated the possible reasons for the impairment of apoptosome assembly. Intriguingly, Apaf-1 and caspase-9 were functionally expressed and cytochrome c was released from mitochondria upon serum withdrawal in 423-cells as required for potential apoptosome assembly. Anyhow, apoptosome formation did not occur and was dispensable for apoptosis induction and processing. We found caspase-9 sequestered in 423-cells to cytoskeletal structures and will speculate whether this finding might represent a novel biological strategy to impair caspase-9 activation.

Materials and methods Chemicals and antibodies Antibodies directed against bid (FL-195), caspase-3 (H-277), caspase-9 p10 (F-7), and PKC-y (C-17) were supplied by Santa Cruz, caspase-8 (Ab-4), and cytochrome c (7H8.2C12) by NeoMarkers, bcl-2 (Clone 7) by Transduction Laboratories, fas (CH-11) by Upstate Biotechnology, caspase-9 (AAP-109) and Apaf-1 (AAP-300) by StressGene, caspase-3-p17 by New England Biolabs, and h-actin (AC-15) by Sigma. Caspase inhibitors (zVAD-FMK, zDQMD-FMK, zVEID-FMK, IETD-CHO, and LEHD-CHO), 4V-6-diamidine-2-phenyl indole (DAPI), MEK-1 inhibitor U0126, 5V-methyl thioadenosine (MTA), and staurosporine (STS) were purchased from Calbiochem. Mitochondrial dye JC-1, NP-40, Triton X-100, and Tween 20 were purchased from Sigma. CompleteR Protease Inhibitor Cocktail was purchased from Roche Diagnostics. Cell culture The nontransformed, G418-resistant embryonic rat cell line, 423, immortalized by Human Papilloma Virus type 11 DNA, was routinely cultivated in DMEM, supplemented with 10% fetal calf serum (FCS; Sigma), penicillin/ streptomycin (100 U/ml; Sigma), and G418 (200 Ag/ml; Gibco Invitrogen Coop.) at 378C in a humidified atmosphere containing 7.5% CO2. Bcl-2 supertransfected 423cell lines were generated by transfecting pBABEpuro-bcl-2 (kindly provided by B. Lqscher, Germany) together with pTK-hygro, for hygromycin-resistance, into the parental 423-cell line. Cells were cultured in G418-containing medium supplemented with Hygromycin (50 Ag/ml; Calbiochem). Individual clones were isolated and established into cell lines as described [21–23]. MR-6, a c-myc/ c-H-ras expressing, transformed rat cell line, established in our lab, was routinely cultivated in DMEM, supplemented with 10% FCS, Pen/Strep, and G418. HeLa cells were cultivated in RPMI-1640, supplemented with Pen/Strep

and 10% FCS in a humidified atmosphere containing 5% C02. Induction and quantification of apoptosis For induction of apoptosis, confluent 423-cells were washed twice with serum-free DMEM and further incubated in DMEM, 0.1% FCS. Apoptosis induction by staurosporine (250 nM) and anti-Fas antibody (200 ng/Al), respectively, was performed in the presence of 2.5% FCS in DMEM for 6 h. Apoptosis in exponentially growing MR-6 cells was induced by the addition of the MEK-1 inhibitor, U0126 (10 AM), under full serum condition. For determination of apoptosis rates, 423-cells were seeded in 24-well plates and grown to confluence. Cells were fixed with 3% formaldehyde at the indicated time points and stained with DAPI. Intact and apoptotic nuclei were counted in randomly chosen microscopic fields on a Nikon FX-35DX microscope. At least 400 cells were counted per time point. Apoptosis rates were calculated by combining the numbers of condensed and fragmented nuclei with the number of calculated cells detached in the course of the experiment. For determination of caspase activity, caspase-7 Activity Assay Kit (New England Biolabs) was used. Caspase-3 was precipitated with antibody against caspase-3 (H-277) and total caspase activity was determined from total lysate without immunoprecipitating a specific caspase. DNA ladders After induction of apoptosis, 423-cells were lysed in 50 mM Tris, pH 7.5; 10 mM EDTA with 0.5% laurylsarcosine followed by digestion with RNase A (50 Ag/ml) and proteinase K (50 Ag/ml). Genomic DNA was precipitated with ethanol and dissolved in 10 mM Tris, pH 8.0. Equal volumes were separated on a 2% TBE-agarose gel and stained with ethidium bromide. Isolation of mitochondrial, cytoplasmic, and total protein lysates For total protein lysates, cells were harvested by scraping, washed twice in ice-cold PBS, and lysed in RIPAII (500 mM NaCl; 50 mM Tris, pH 7.4; 0.1% SDS; 1% NP-40; 0.5% Na-DOC; 0.05% NaN3; and complete protease inhibitor mix) for 30 min on ice. Lysates were sonificated, centrifuged at 12,000  g. Supernatants were used as total cell lysates. For isolating mitochondria, cells were washed twice in ice-cold washing buffer (10 mM Tris pH 7.5; 200 mM sucrose) and resuspended (250 mM sucrose; 20 mM Tris, pH 7.5, 10 mM KCl; 1.5 mM MgCl2; 1 mM EDTA; 1 mM EGTA; 1 mM DTT; and complete protease inhibitor mix). After swelling on ice, cells were disrupted by six passages

C.J. Schamberger et al. / Experimental Cell Research 302 (2005) 115–128

through a 26-gauge needle applied to a 1-ml syringe and the heavy membrane fraction was separated by centrifugation at 750  g. Mitochondria were separated from cytoplasm by centrifugation at 10,000  g. Mitochondria were lysed in RIPAII buffer. SDS-PAGE and immunoblot procedure was performed as described in Ref. [24]. FACS analysis The 423-cells were seeded in 24-well plates and grown to confluence. Apoptosis was induced for up to 6 h. Staurosporine-treatment was performed under 0.1% FCS condition. Floating and attached cells were combined in DMEM, 2.5% FCS, and stained with 5 AM JC-1 [25,26] for 10 min in the dark. Samples were analyzed in a Becton Dickinson FACSCalibur system using CELLQuest software. In vitro apoptosome assembly The 423-cells were harvested, washed with PBS, and extracted as described [27]. Briefly, cells were resuspended in hypotonic buffer (20 mM HEPES-KOH, pH 7.5; 10 mM KCl; 1.5 mM MgCl2; 1 mM EDTA; 1 mM EGTA; 1 mM DTT; 100 AM PMSF; 10 Ag/ml leupeptin; and 2 Ag/ml aprotinin), incubated on ice for 15 min, and lysed by homogenization with 40 strokes of a Dounce homogenizator. Crude extracts were centrifuged for 30 min at 15, 000  g to remove intact cells, nuclei, and cell debris. The supernatant was used as cell-free extract for apoptosome assembly assays and stored at 808C. For the preparation of cell-free extracts in the presence of detergents, the nonionic detergents NP-40, Triton X-100, and Tween 20 were used at the concentration of 0.5% each. To induce cell shrinkage under full serum condition, 423cells were incubated with MTA (1 mM) 2 h prior to lysis under detergent-free condition. Fifty microliters of cell extract, 10 Al equine cytochrome c (5 Ag/Al), and 10 Al dATP (10 mM) were brought to a final volume of 100 Al. The mixture was incubated at 378C for up to 3 h and 10 Al aliquots was removed at each time point. Immunoprecipitated caspase-9 (F-7) from HeLa cell-free extract was mixed with 423-cell-free extract and incubated with cytochrome c for the indicated time points. The samples were analyzed by SDS-PAGE and immunoblotted with caspase-9 (AAP-109) antibody. Immunostaining The 423-cells were plated on coverslips and cultivated for at least 48 h before fixation. Cells were fixed and stained with anti-caspase-9 (AAP-109)/anti-rabbitCy3, phalloidin-FITC for actin staining, and anticytokeratin19/anti-mouse-FITC as described in Ref. [24], and digital images were taken on a Leica confocal microscope.

117

Results Kinetics of apoptosis after serum withdrawal Apoptosis in 423-cells was induced by reducing the serum concentration in the medium from 10% to 0.1%, which is further referred as serum withdrawal or starvation. Within 15 min cells shrinked, within 2 h cell blebbing occurred, and massive cell detachment was observed after 3 h (Fig. 1A). Apoptosis rate increased almost linear from initial 5% up to 45% within the first 7 h. After that time, apoptosis rate flattened and account for about 50% at 12 h and about 60% at 24 h after serum withdrawal (Fig. 1B). Due to the high fraction of apoptotic cells, all experiments were performed during the first 6 h after apoptosis induction. One hallmark of apoptosis is the formation of DNA ladders. Genomic DNA is cleaved by the nuclease CAD/ DFF40, after caspase-3 or -7-promoted release from its inhibitor ICAD/DFF45 [28–30]. We investigated these characteristics, low molecular ladders in the first 6 h after apoptosis induction. Laddering became detectable within 3 h after serum withdrawal and the amount increased until 6 h (Fig. 1C, left panel). Detachment-induced cell death in epithelials and trophic factor withdrawal-induced apoptosis in cerebellar granule cells were described to be associated with early activation of caspase-6 [8,31]. Therefore, we tested out the involvement of caspase-6 in our model system for serum-starvation induced apoptosis. We treated 423-cells during serum starvation with specific inhibitors for caspase-3 and caspase-6 to inhibit DNA ladder formation. Treatment with the general caspase inhibitor VAD served as control. Substrate specifities for DQMD (for caspase-3) and VEID (for caspase-6) were determined by Talanian et al. [32]. General caspase inhibitor VAD blocked DNA laddering, caspase-3 inhibitor, DQMD, displayed a reduction of DNA ladders, while the caspase-6-specific inhibitor, VEID, had no inhibitory effect in this short-time experiment (Fig. 1C, right panel). To examine the participation of caspase-7, we performed caspase activity assays with fluorochrome-coupled DEVD peptide and compared it to caspase-3 activity and caspase activity in total lysates (data not shown). Four hours after serum withdrawal, the increase of caspase activity was about fivefold in total lysate, about threefold for caspase-3-specific activity, but remained unaltered for caspase-7-specific activity. These results indicated that 423-cells undergo apoptosis in a rapid time course after serum withdrawal in a caspasedependent manner and identified caspase-3 as main executioner caspase. Activation of caspases after serum withdrawal To reveal a more detailed insight into the caspase cleavage cascade, we investigated the activation of initiator

118

C.J. Schamberger et al. / Experimental Cell Research 302 (2005) 115–128

Fig. 1. Induction of apoptosis in 423-cells by serum starvation. Confluent cultures of 423-cells were serum starved up to 6 h and analyzed as follows. (A) Uninduced and induced cells after 6 h were imaged by phase contrast (left panel) and by DAPI staining (right panel). A representative mixture of cells remaining in the layer, blebbing cells, and apoptotic bodies are shown in the lower panel. (B) Quantification of apoptosis rate by counting loss of intact nuclei during a time course up to 24 h. The error bars represent F SEM from at least three independent experiments per time point. (C) Generation of DNA ladders during a time course (left panel) and under the influence of specific caspase inhibitors (right panel). DNA was prepared from a combination of floating and attached cells. General caspase inhibitor VAD, caspase-6-specific inhibitor VEID, or caspase-3-specific inhibitor DQMD (40 AM each) were present during serum starvation for 6 h.

caspases-8 and Rapidly after cleaved, while first 6 h after

-9 and the executioner caspase-3 (Fig. 2A). serum withdrawal, we found caspase-8 caspase-9 remained unaffected within the apoptosis induction in 423-cells. Cleaved

caspase-3 was detectable as soon as 2 h after induction and further accumulated during the following 4 h. The participation of other initiator caspases, like caspase-2 or -12 cannot be excluded, but we focused on the observation, that

C.J. Schamberger et al. / Experimental Cell Research 302 (2005) 115–128

119

Fig. 2. Activation of caspases. (A) Processing of caspases during a time course in serum-starved 423-cells. Processing of caspase-8 and subsequently that of caspase-3 was evidenced by the detection of characteristic cleavage products. No processing of caspase-9 was detected. Actin was used as loading control. (B) Processing of caspase-8, but not caspase-9, induced by fas signaling and staurosporine treatment, respectively. Confluent 423-cells were treated with 200 ng/ml fas-antibody (anti-fas) or 250 nM staurosporine (STS) under serum-reduced condition (2.5% FCS; untreated) for 6 h. Lysate from serum-starved 423-cells was used as apoptotic control. PKC-y served as caspase-3 target. Arrows mark cleavage products. Actin was used as loading control. (C) Detection of rat caspase-9 processing. An apoptosis-prone c-myc/c-H-ras-transformed rat cell line was treated with MEK-1 inhibitor 5 AM U0126 up to 3 h. Processing of caspase-9 was evidenced already within 15 min by detection of the p10 subunit. Processing of caspase-8 generated intermediate fragments. Processing of caspase-3 followed as expected. Arrows mark cleavage products. Actin was used as loading control.

caspase-9 remained unaltered in 423-cells during serum starvation-induced apoptosis. To investigate the activation of extrinsic and intrinsic pathways selectively, we assessed apoptosis induction by other stimuli (Fig. 2B). We treated 423-cells with staurosporine, initiating the intrinsic, or fas-antibody, initiating the extrinsic pathway, for 6 h. Addition of anti-fas antibody led to caspase-8 and caspase-3 activation. Staurosporine, which is described to induce apoptosis by activating caspase-9 due to mitochondrial damage [33], but also via caspase independent mechanism [19], led to shrinkage and detachment of 423-cells. Caspase-8 was the only caspase found cleaved after staurosporine treatment. Detection of PKC-y and its cleavage fragments in this rat cell system [34–36] was applied to indicate caspase-3 activity. We detected a PKC-y fragment in fasantibody-treated and serum-starved cell lysates. Concomitantly, we determined apoptosis rate in these staurosporine

and anti-fas antibody treated 423-cells (Fig. 2B). The 423-cells were almost resistant against staurosporine treatment, while activating the death receptor pathway via fas-antibody add up to 17% apoptosis rate after 6 h under reduced serum condition of 2.5%. Based on this experiment, we concluded that only caspase-8, but not caspase-9, is activated under various apoptotic triggers in 423-cells. Detection of rat caspase-9 processing We tested out the sensitivity and specifity of the caspase-9 antibody to recognize cleavage fragments in rat cell lysates. For this purpose, we used another rat cellculture model for apoptosis, established in our lab. The cmyc/c-H-ras-transformed rat cell line, MR-6, is highly sensitive to MEK1 inhibition. MEK1 is part of one rassignaling transduction cascade, which is involved in

120

C.J. Schamberger et al. / Experimental Cell Research 302 (2005) 115–128

proliferation, differentiation, and survival [37]. Rapidly after addition of the MEK1 inhibitor U0126, MR-6 cells underwent apoptosis under full serum conditions (unpublished data). In this cell system, caspase-9 processing was detectable and followed the same kinetics as caspase-8. Caspase-3 cleavage became evident subsequently at 1 h after apoptosis induction (Fig. 2C). The fragmentation pattern of rat caspase-8, with fragments of approximately 42, 37, 30, and 15 kDa (Fig. 2C), differs from that of processed human caspase-8, with fragments of approximately 43, 26, 18, and 12 kDa [38]. Comparative sequence analysis of the human and rat caspase-8 sequences has resulted in that only the cleavage site between the large and the small subunit is conserved in rat caspase-8 (Table 1A). Additionally, there are two other putative caspase cleavage motives in the N-terminal DED domain of rat caspase-8, which may explain the differences in fragmentation pattern. In Table 1B, a schematic delineation of putative rat caspase-8 fragments with calculated molecular weight is shown. In contrast, the positions of cleavage sites in rat caspase-9 correspond to the positions in human caspase-9 [39] (Table 1A). Therefore, we exclude a detection problem of caspase-9 or caspase-8 processing in our rat cell system.

Involvement of caspase-8, but not caspase-9, upon serum withdrawal To support our findings, that caspase-9 played an ancillary role in serum starvation-induced apoptosis in 423-cells, we inhibited initiator caspases with their specific inhibitors. Apoptosis was induced in the absence or presence of the general caspase inhibitor VAD, caspase-3-specific inhibitor DQMD, caspase-8-specific inhibitor IETD, caspase-9-specific inhibitor LEHD, respectively, and both of them together. Survival rate was determined and quantified. A concentration-dependent decrease of apoptosis rate was observed for caspase-3, caspase-8, and by blocking all caspases. Inhibition of the initiator caspase-8 was almost as effective as blocking caspase-3. Caspase-9-specific inhibitor, LEHD, exhibited no survival benefit for 423-cells. Treatment with IETD and LEHD together induced no synergistic effect on apoptosis inhibition in 423-cells (Table 2). Overexpression of bcl-2 does not protect against serum starvation-induced apoptosis. As a second indirect approach to assess caspase-9 contribution, we tested the impact of bcl-2 on 423-cells survival. Bcl-2 protects cells by inhibiting cytochrome c release from mitochondria, preventing apoptosome assembly and caspase-9 processing [40]. We estab-

Table 1 (A) Comparative protein sequence analysis of human and rat caspases-8 and -9 and (B) schematic delineation of putative rat caspase-8 fragments with calculated molecular weight (Clone Manager Professional Suite)

Protein sequences of rat and mouse caspases were aligned against human caspase, and corresponding position of amino acids was calculated with Clone Manager Professional Suite in the algorithm of global-ref-alignment. Position of cleavage sites for human caspase-8 were taken from [38], that for human caspase-9 from [39]. Bold letters indicate sequence identity between the species. *Functional cleavage motif; **putative cleavage motif.

C.J. Schamberger et al. / Experimental Cell Research 302 (2005) 115–128

121

Table 2 Involvement of caspase-8, but not caspase-9, upon serum withdrawal Apoptosis rate after 6 h of serum starvation 44.1 F 7.2 Conc. (AM)

IETD

LEHD

IETD + LEHD

DQMD

VAD

20 40

33.9 F 6.3 23.9 F 5.8

40.1 F 3.4 38.4 F 5.3

29.2 F 2.9* 23.4 F 2.3*

26.5 F 4.9* 20.1 F 5.2*

27.9 F 3.8* 21.6 F 4.7*

Inhibition of

casp-8

casp-9

casp-8+9

casp-3

all caspases

Confluent 423-cells were treated with caspase-3-specific inhibitor DQMD, caspase-8-specific inhibitor IETD, caspase-9-specific inhibitor LEHD, general caspase inhibitor VAD, and a combination of IETD and LEHD (20 and 40 AM each) under serum-starved condition for 6 h. Apoptosis rate of 423-cells without caspase inhibition is indicated in the first row. Formaldehyde-fixed and DAPI-stained nuclei were counted and quantified. Data are means F SEM. * Significant decrease versus 423-apoptosis rate ( P b 0.05).

decreased after 2 h and correlated with loss of mitochondrial membrane potential (Fig. 3A, inlet). Translocation of pro- and antiapoptotic bcl-family members is a common event during apoptosis. Bid or tbid translocation from cytoplasm to mitochondria and the complex formation with bax at the mitochondrial outer membrane lead to pore opening and cytochrome c release [42]. We determined bcl-2 and bid location by cellular fractionation and Western blot analysis after serum withdrawal in 423-cells. As indicator for mitochondrial membrane pore opening, we examined cytochrome c release into the cytoplasm (Fig. 3B). Cytochrome c was found in the cytoplasm of apoptotic cells after 4 h, when about 20% of 423-cells lost mitochondrial membrane potential. Endogenous antiapoptotic bcl-2 was detected in the mitochondrial fraction under full serum and apoptotic conditions, with no remarkable changes during this 4-h period. We found full-length bid translocated to mitochondria after serum withdrawal, while the very weak signal of tbid remained undetectable (Fig. 3B). Translocation of fulllength bid to mitochondria was described recently as alternative bid mechanism in rodent cells [43]. These experiments revealed that mitochondria were damaged in 423-cells and cytochrome c was released into the cytoplasm for a possible apoptosome assembly.

lished bcl-2-overexpressing subclones with distinct bcl-2 levels (data not shown) and tested their resistance against serum starvation. As survival control, parental 423-cells were treated with the general caspase inhibitor VAD. Apoptosis rate after 4 h of those bcl-2 subclones, determined by counting intact nuclei, was almost identical to parental 423cells (Table 3). VAD-treatment decreased apoptosis rate significantly. The question arose if this overexpressed bcl-2 is functional active and able to protect mitochondria. Therefore, we examined changes in mitochondrial membrane potential by flow cytometric analysis of JC-1 complexes [41]. Bcl-2 overexpression prevented mitochondrial membrane disruption during serum starvation but did not prevent apoptosis. Mitochondria of those bcl-2 overexpressing cells were also resistant against staurosporine treatment (Table 3). These data indicate that apoptosis may proceed efficiently without the involvement of the intrinsic apoptosis pathway. Loss of mitochondrial membrane potential and cytochrome c release after serum withdrawal. We further examined changes in mitochondrial membrane potential during a time course after serum withdrawal by flow cytometric analysis of JC-1 complexes. The percentage of 423-cells with low mitochondrial membrane potential increased from a basal level of 5% up to 55% after 6 h (Fig. 3A). Disruption of mitochondria in the presence of processed caspase-8 occurred as a result of caspase-8 induced bid cleavage [41]. We examined the kinetics of bid cleavage in a time course during serum starvation by Western blot analysis. The level of bid in total protein lysate

Determination of caspase-9 function in vitro The reasons for apoptosome formation impairment may be multifaceted. Alternative splicing of caspase-9 [44–46]

Table 3 Overexpression of bcl2 protects mitochondria but does not prevent serum starvation-induced apoptosis 423

cl. 1

cl. 2

cl. 3

cl. 4

423 + VAD

Cell loss and apoptotic nuclei

Control 4h

7.6 F 1.3 31.6 F 3.5

6.1 F 2.8 31.4 F 3.1

3.3 F 1.9 27.5 F 3.5

5.8 F 2.2 23.9 F 5.2

2.5 F 1.1 31.6 F 3.3

2.3 F 0.5 10.3 F 2.1#

Reduced Dc

Control 2h 4h STS

4.9 13.5 22.5 24.9

F F F F

2.1 4.3* 4.1* 2.5*

2.2 0.7 1.2 1.7

F F F F

0.3 0.1 0.3 0.2

1.8 1.2 1.5 12.9

F F F F

0.6 0.3 0.5 2.1*

4.2 1.3 1.2 1.4

F F F F

1.3 0.2 0.3 0.2

1.8 1.6 1.4 3.5

F F F F

0.5 0.1 0.2 0.4*

n.d. n.d. 7.9 F 1.9# n.d.

Apoptosis rate of formaldehyd-fixed and DAPI-stained 423-cells and four bcl2-overexpressing subclones, respectively, revealed no significant differences, whereas mitochondria remained intact in bcl2-overexpressing clones only despite high apoptosis rate. Mitochondria of these subclones were even resistant against a 4 h staurosporine treatment (250 nM). Data are means F SEM; VAD general caspase inhibitor; STS: staurosporine; n.d. not determined. * Significant increase versus untreated control ( P b 0.05). # Significant decrease versus 423-apoptosis rate ( P b 0.05).

122

C.J. Schamberger et al. / Experimental Cell Research 302 (2005) 115–128

Fig. 3. Involvement of mitochondria in serum starvation-induced apoptosis. (A) Correlation of bid cleavage with breakdown of mitochondrial membrane potential. Cell death in 423-cells was induced by serum withdrawal. Time course of changes of mitochondrial membrane potential, determined by staining with JC-1 and followed by FACS analysis, is graphed. The error bars represent F SEM from at least three independent experiments. Western blot analysis of bid level in total cell lysates during a time course is shown in the inlet. (B) Cytochrome c release, bcl-2 location, and bid translocation. Mitochondrial and cytoplasmic fractions of untreated and serum-starved 423-cells after 4 h were analyzed for cytochrome c, bcl2, and bid levels. Bid translocation and cytochrome c release indicated involvement of mitochondria during serum-starved apoptosis.

and Apaf-1 [47], cellular inhibitor proteins like XIAP and heat shock proteins [45,48,49] were described as possible factors. First, we checked out caspase-9 mutations or splice variants in 423-cells. In rat cells, caspase-9 is alternatively spliced with a dominant negative function, as caspase-9CTD [27]. To screen for splicing products, we performed RT-PCR from RNA of growing and confluent 423-cells. We detected solely full-length mRNA and this cloned, fulllength caspase-9 carried no mutations compared to the published rat sequence (accession number NM_031632) (data not shown). Next, we performed an in vitro approach for caspase-9 processing [27] to test out the functionality of caspase-9 in 423-cells. Generally, apoptosome assembly assays are carried out in a detergent-free and hypotonic extraction buffer [50], further referred as cell-free extract. In vitro apoptosome assembly requires caspase-9 and Apaf-1, obtained from cell lysate. To induce apoptosome formation, cytochrome c and dATP are added to cell-free extracts.

After incubation, the reaction mixtures are analyzed by Western blotting. We established apoptosome assembly assays with extracts from HeLa cells to determine the required incubation conditions (Fig. 4). As soon as 10 min after cytochrome c, addition the small p10 subunit and after 30 min a further cleaved caspase-9 fragments at approximately 35 kDa were detectable. For comparison, an aliquot of cell-free extract from 423-cells was loaded on this Western blot, but beside the unspecific band, no caspase-9 immunoreactivity was detected (Fig. 4, lane 10). After this unexpected observation, we determined expression levels of two further caspases and Apaf-1 in cell-free extracts from untreated and serum-starved 423cells, respectively, and HeLa cells. Caspases-8 and -3 revealed a clear signal in Western blot analysis after this extraction method, with full-length procaspase as most prominent band. Cleaved caspase-3 fragment was detected in cell-free extract of serum-starved 423-cells, but caspase8 fragments remained undetectable (Fig. 5A). One possibility may be the recruitment of soluble, cytoplasmic procaspase-8 to the plasma membrane for processing and thus resulting in a detection limitation under this experimental condition. Also Apaf-1 was determined in cell-free extracts of 423-cells and HeLa. Surprisingly, the otherwise soluble cytoplasmic protein, caspase-9, could not be extracted under this detergent-free and hypotonic condition (Fig. 5A). Caspase-9 is sequestered in 423-cells Previous immunoblot data (Figs. 2A and B) of 423derived caspase-9 were obtained from detergent-containing lysates. Therefore, we asked for a possible caspase-9retention mechanism in 423-cells. As in vitro approach to reveal caspase-9, we harvested untreated 423-cells and extracted aliquots by cell disruption in the absence or presence of diverse detergents. Primary REC and HeLa were extracted as control cells without detergents. In both rat and human control cells, caspase-9 was clearly

Fig. 4. Caspase processing in cell-free extracts. HeLa cell-free extract was incubated with cytochrome c to induce apoptosome assembly for up to 1 h. The 423-cell-free extract is shown in the last lane (lane 10). Caspase-9 antibody detected procaspase-9, p10 subunit, and the 35 kDa intermediate cleavage product. An arrow marks the 35 kDa fragment.

C.J. Schamberger et al. / Experimental Cell Research 302 (2005) 115–128

123

Fig. 5. Availability of caspase-9 in 423-cells. (A) Caspase and Apaf-1 detection in cell-free extracts. Cell-free extracts from HeLa and confluent 423-cells under full serum and 4 h after serum withdrawal, respectively, were analyzed for full-length caspase-3, p17 subunit of caspase-3, caspase-8, caspase-9, and Apaf-1. In contrast to total cell lysates (Fig. 2A), rat caspase-9 remained undetectable. The unspecific band caused by caspase-9 antibody was used as loading control. (B) Dependence of caspase-9 detection on extraction conditions. Cell-free extracts of HeLa, REC, and 423-cells were extracted in cell-free buffer. Alternatively, cell-free extracts were prepared from 423-cells pretreated with 1 mM MTA. In addition, 423-cell aliquots were extracted in the presence of NP-40, Triton X100, or Tween 20 (w/o, no detergent). Caspase-9 and actin were detected by Western blot analysis. Intriguingly, caspase-9 and actin were detectable only in detergent-extracted and MTA-pretreated samples. The Ponceau S-stained membrane and the unspecific band, caused by caspase-9 antibody, are shown as loading control. (C) The 423-cell-free extract processed HeLa-derived caspase-9. Immunoprecipitated caspase-9 from HeLa cell-free extract was incubated with cytochrome c to induce apoptosome assembly for up to 3 h. IP was performed with monoclonal F-7 antibody and immunoblotting with rabbit polyclonal AAP-109. Cleavage products are marked by arrows. (D) Caspase-8 processing in 423-cell-free extracts. The 423-cell-free extract was incubated with cytochrome c for up to 2 h. Caspase-8 antibody detected procaspase-8 and two cleavage fragments, indicating functional assembly of the apoptosome. Cleavage products are marked by arrows.

detectable without adding detergents (Fig. 5B, lanes 1 and 7). Rat caspase-9 is 38 amino acids (Table 1A) or approximately 4 kDa heavier than the human protein, which resulted in a shift of the immunoreactive band in Western blot analysis. In the presence of all used, nonionic, mild detergents, caspase-9 was extracted from 423-cells, in the following order Triton X18 100 b NP40 b Tween 20 (Fig. 5B, lanes 3–5). As in vivo approach to solubilize caspase-9, we disrupted the cytoskeleton by a chemical substance before harvesting and extracting the cells in detergent-free cell-free extraction buffer. We choose 5V-methyl thioadenosine (MTA) to disrupt the cytoskeleton because shrinkage of 423-cells was faster and more effective than that induced

by cytochalasin B, an inhibitor of actin filament formation [51] (data not shown). MTA is known as FGF receptor inhibitor [52], as carboxymethyltransferases inhibitor, and apoptosis inducer [27]. In MTA-treated 423-cells, caspase-9 was extracted under detergent-free conditions (Fig. 5B, lane 6). Additionally, we found less actin solubilized in untreated 423-cells (Fig. 5B) compared to the detergent-treated cell-free extracts. Therefore, we used the unspecific band caused by caspase-9 antibody and the ponceau-stained membrane as protein loading control. As a conclusion, caspase-9 appeared to be tightly bound to yet uncharacterized structures in the cytoskeleton of 423-cells and therefore exhibited reduced availability for participation in apoptosis.

124

C.J. Schamberger et al. / Experimental Cell Research 302 (2005) 115–128

423-cell-free extract is capable to process caspase-9 So far, the question remained unsolved, whether apoptosome assembly occurred at all and whether Apaf-1 and caspase-9 were functionally active in 423-cells. To approach this problem, mixing experiment were performed with immunoprecipitated HeLa-derived caspase-9 and cell-free extracts from 423-cells (Fig. 5C). The 423-cell extract was able to process HeLa caspase-9 under those artificial conditions. As second approach, we wondered whether the sensitivity of this assay would be sufficient to observe cleavage of caspase substrates in vitro, considering that only traces of 423-derived caspase-9 were present in cell-free extracts (Fig. 5B, lane 2). The amplification loop from caspase-9 via caspase-3 to caspase-8 may be simulated in apoptosome assembly assay independently of its naturally occurrence after apoptotic stimuli in the cells [6,27]. We performed this assay with 423-cell-free extracts, containing

only traces of caspase-9, for a prolonged incubation time and were able to detect caspase-8 fragments (Fig. 5D). These experiment demonstrated that 423-cell-free extract was capable to form functional apoptosomes under artificial conditions. Caspase-9 is colocalized with cytokeratin in inclusion bodylike structures Because 423-derived caspase-9 is only soluble, if the cytoskeletal structure is disrupted for example after MTAtreatment (Fig. 5B, lane 6), and immunoblotting displayed that even lesser amounts of actin were soluble (Fig. 5B, lane 2), we hypothesized that caspase-9 is bound to cytoskeletal structures. Because co-immunostaining of actin and caspase-9 in 423-cells exhibited no colocalization (Fig. 6; left panel, merge), we tested out caspase-9 and cytokeratin filament network colocalization. We chose

Fig. 6. Caspase-9 colocalizes with cytokeratin in a punctuated pattern in 423-cells. Immunofluorescence with caspase-9 and cytokeratin-19 antibody and phalloidin for actin filament network was performed in the presence of 10% FCS. Merge of digital images of FITC and Cy3 signals display colocalization of caspase-9 with cytokeratin (right panel), but not actin (left panel).

C.J. Schamberger et al. / Experimental Cell Research 302 (2005) 115–128

cytokeratin-19 because it is rather highly expressed in 423cells [53]. Caspase-9 and cytokeratin-19 resulted in an unusual punctuated pattern (Fig. 6; left panel, merge). Typically, in immunofluorescence keratin network is similar to the actin filaments, whereas a punctuated pattern of cytokeratins is observed early upon apoptosis induction [54]. Further research is required if this caspase-9 retention property is an artefact of this cell line or if there is a deeper impact on apoptosis prevention.

Discussion Apoptosis induced by serum or trophic factor withdrawal leads to rapid apoptotic response in various cell culture model systems [6,55,56], but little is known about the role of caspase-9 thereby. In undifferentiated PC-12 cells, caspase-2 was described as main activator [6,57]. In myc-overexpressing Rat1 cells, apoptosis by growth factor withdrawal was dependent on caspase-8 [58], while the involvement of caspase-2 was excluded [59]. In mouse AKR2-B fibroblasts, ER-located caspase-12 [60] was found responsible for apoptosis. Additionally, no apoptosome assembly took place in this system because cytochrome c was retained in mitochondria during experimental time [9,60]. The participation of caspase-9 in serum starvation-induced apoptosis in different cell culture systems was proven indirectly by transfection experiments with bcl-family members. Whereas an ER-targeted bcl-2 variant, bcl2-cb, counteracted serum starvation-induced apoptosis most efficiently compared to bcl-2 and bcl-x, indicating an involvement of ER-located caspase-12 than one of caspase-9 [21,61–63]. In this study, we took advantage of an immortal rat embryonic cell line established in our lab [21]. This cell line, named 423, underwent apoptosis upon serum starvation, whereas about 50% underwent apoptosis in a caspasedependent manner within 6 h. We identified caspase-3 as main executioner and caspase-8 as main initiator caspase. The reasons why caspase-8 is activated in serum-starvation induced apoptosis are rather unclear. In myc-overexpressing cells, with a low kinetics of apoptosis, CD95 is up-regulated and propagates cell death [10]. Also in anoikis, a detachment-induced apoptosis, caspase-8 is rapidly activate after detachment mediated by FADD redistribution to the plasma membrane [64,65]. In serum-deprived 423-cells, the instantly detachment of the cells may induce anoikis-like cell death, perhaps via FADD recruitment to the DISC and disrupted integrin signaling. Caspase peptide inhibitor experiments revealed that blocking caspase-8 activity had almost the same rescuing effect as the application of a caspase-3-specific and general caspase inhibitor VAD, respectively (Table 2). Additionally, bcl-2 overexpressing subclones performed almost the same apoptosis rate as the parental cell-line after serum starvation; their mitochondria stayed intact (Table 3). Despite a missing amplification loop

125

via the intrinsic caspase cascade, 423-cells displayed a rapid course of apoptosis after serum deprivation. Therefore, we concluded that the intrinsic caspase-9 pathway seems to be dispensable for apoptosis after serum deprivation. Our main point of interest was how or why caspase-9 remained unprocessed in 423-cells. The reasons for impairment of apoptosome formation can be multifaceted. First, we checked for cytochrome c release from mitochondria. Disruption of mitochondria during apoptosis leads to release of proapoptotic factors [66], like cytochrome c, which is required for apoptosome assembly. Opening of mitochondrial membrane pores can among others be induced by BH3-only bcl-2-family members, like bax, bak, and bid [67]. We focused on bid because it is described as the link between extrinsic caspase activation and the involvement of mitochondria. Caspase-8 cleaves bid to truncated bid, tbid, which provokes mitochondrial disruption [41,67]. In a recent report, also full-length bid provoked disruption of mitochondria in rat cells [68]. Bid was cleaved in 423-cells following caspase-8 processing (Fig. 3A, inlet). We found translocation of full-length bid to mitochondria (Fig. 3B) after serum deprivation, loss of mitochondrial membrane potential (Table 2 and Fig. 3A), and cytochrome c release (Fig. 3B). Therefore, we excluded a lack of cytochrome c for the impairment of apoptosome assembly. Secondly, we checked for alternative splicing products [44,45,69] or mutations in 423-derived caspase-9. These alterations would allow apoptosome formation but impair caspase-9 processing and activity. We excluded caspase-9 alteration in due to the fact that just full-length mRNA was expressed and the cloned construct carried no mutation (data not shown). Thirdly, absent expression or alternative splicing of Apaf-1 may interfere with apoptosome assembly and may thus refer resistance against apoptosis in distinct tissues [70,71] and tumors [47,72,73]. Apaf-1 was expressed and functional in 423-cells (Fig. 5A), shown via apoptosome assembly in cell-free extracts (Figs. 5C and D). For the same reason, we excluded also inhibitory effects of cellular proteins like XIAP and heat shock proteins [48,49,74] in 423-cells. But finally, carrying out apoptosome assembly assays gave us an understanding of caspase-9 inactivation in 423-cells. We found, that in this cell line caspase-9 was bound to yet uncharacterized structures and therefore unavailable to participate in the amplification of caspase signaling after cytochrome c release. Binding of caspase-9 to these cellular structures was abrogated by detergent treatment, but also in cells treated with substances, that disrupt the intracellular filament network (Fig. 5B and data not shown). Additionally, a slight increase in caspase-9 level in comparison to untreated cells was detected in Western analysis at 1 and 2 h after serum deprivation (Fig. 2A, compare lane 1 to lanes 2 and 3). At these very early time points, 423-cells were shrunken and rounded up, which indicated disruption of cytoskeletal structures. In recent years, the localization of caspase-9 was a focus of interest. Beside its cytoplasmic distribution, recent reports

126

C.J. Schamberger et al. / Experimental Cell Research 302 (2005) 115–128

showed that procaspase-9 [75,76] were dominantly located in the intermembrane space of mitochondria, whereas others excluded a mitochondrial location [77,78]. A very extensive localization study was done on rat brain tissue, where caspase-9 was found located predominantly in the nucleus and to a lesser extend in the cytoplasm [79]. In spite of this controversial about mitochondrial or nuclear localization, there were no doubts about a soluble cytoplasmic occurrence. In 423-cells, not only caspase-9 but also actin remained mainly insoluble without detergent treatment. This unusual property of actin substantiated our presumption of an involvement of filament networks in 423-cells to sequester a proapoptotic protein. Actin is described as one of the main components of intracellular aggregates (IA), insoluble protein structures, in tissue regions affected by neurodegenerative diseases. Other proposed components of these IA were caspases, while only caspase-3, but not caspase-8 or -9, was detected in those structures [79]. In 423-cells, caspase-9 was found associated with cytokeratin in inclusion bodylike structures (Fig. 6). Such structures were observed in MCF-7 [80] and A549 cells [81] after TRAILinduced apoptosis, but in contrast to the inhibiting effect determined in this study, caspase association to keratins acts as scaffold for further caspase processing. Taken together, two independent conclusions can be drawn from the presented results. Firstly, caspase-9 can be dispensable for serum starvation-induced apoptosis. Secondly, caspase-9 sequestration was identified as a novel cellular mechanism to impair apoptosome assembly. Further research is required to characterize this sequestering mechanism and to define its possible function as resistance mechanism against apoptosis induced by mitochondrial damage.

[2]

[3]

[4]

[5]

[6]

[7] [8]

[9]

[10]

[11]

[12]

[13]

[14]

Acknowledgments We express our deep sorrow about Christa Cerni, who deceased on 26th of December, 2003. The expression bsurprisinglyQ is dedicated to her. We thank J. Gotzmann, W. Mikulits, and G. Krupitza for providing antibodies against PARP, cytokeratin-19, and caspase-9. We thank I. Herbacek for excellent assistance at the FACS and generating statistics and G. Krupitza and I. Haberl for critical reading the manuscript. This work was supported by grants of the Institute of Cancer Research of the University of Vienna (to C.S.), the Bqrgermeisterfonds der Bundeshauptstadt Wien (Project 2167, to C.C.), and the Herzfelder’sche Familienstiftung (to C.G.).

References [1] L. Stefanis, D.S. Park, C.Y. Yan, S.E. Farinelli, C.M. Troy, M.L. Shelanski, L.A. Greene, Induction of CPP32-like activity in PC12 cells

[15]

[16]

[17]

[18]

[19]

[20]

by withdrawal of trophic support. Dissociation from apoptosis, J. Biol. Chem. 271 (1996) 30663 – 30671. A. Simm, V. Hoppe, A. Gazit, J. Hoppe, Platelet-derived growth factor isoforms prevent cell death during starvation of AKR-2B fibroblasts, J. Cell Physiol. 160 (1994) 295 – 302. S. Alexandre, C. Rast, G. Nguyen-Ba, P. Vasseur, Detection of apoptosis induced by topoisomerase inhibitors and serum deprivation in Syrian hamster embryo cells, Exp. Cell Res. 255 (2000) 30 – 39. W. Chao, Y. Shen, L. Li, A. Rosenzweig, Importance of FADD signaling in serum deprivation- and hypoxia-induced cardiomyocyte apoptosis, J. Biol. Chem. 277 (2002) 31639 – 31645. R. Harfouche, H.M. Hassessian, Y. Guo, V. Faivre, C.B. Srikant, G.D. Yancopoulos, S.N. Hussain, Mechanisms which mediate the antiapoptotic effects of angiopoietin-1 on endothelial cells, Microvasc. Res. 64 (2002) 135 – 147. G.I. Evan, A.H. Wyllie, C.S. Gilbert, T.D. Littlewood, H. Land, M. Brooks, C.M. Waters, L.Z. Penn, D.C. Hancock, Induction of apoptosis in fibroblasts by c-myc protein, Cell 69 (1992) 119 – 128. S.H. Kaufmann, M.O. Hengartner, Programmed cell death: alive and well in the new millennium, Trends Cell Biol. 11 (2001) 526 – 534. T.E. Allsopp, J. McLuckie, L.E. Kerr, M. Macleod, J. Sharkey, J.S. Kelly, Caspase 6 activity initiates caspase 3 activation in cerebellar granule cell apoptosis, Cell Death Differ. 7 (2000) 984 – 993. M. Kilic, R. Schafer, J. Hoppe, U. Kagerhuber, Formation of noncanonical high molecular weight caspase-3 and -6 complexes and activation of caspase-12 during serum starvation induced apoptosis in AKR-2B mouse fibroblasts, Cell Death Differ. 9 (2002) 125 – 137. A.O. Hueber, M. Zornig, D. Lyon, T. Suda, S. Nagata, G.I. Evan, Requirement for the CD95 receptor-ligand pathway in c-Myc-induced apoptosis, Science 278 (1997) 1305 – 1309. R. Haviv, L. Lindenboim, J. Yuan, R. Stein, Need for caspase-2 in apoptosis of growth-factor-deprived PC12 cells, J. Neurosci. Res. 52 (1998) 491 – 497. R. Bertrand, E. Solary, P. O’Connor, K.W. Kohn, Y. Pommier, Induction of a common pathway of apoptosis by staurosporine, Exp. Cell Res. 211 (1994) 314 – 321. J.E. Reynolds, J. Li, R.W. Craig, A. Eastman, BCL-2 and MCL-1 expression in Chinese hamster ovary cells inhibits intracellular acidification and apoptosis induced by staurosporine, Exp. Cell Res. 225 (1996) 430 – 436. M.D. Jacobsen, M. Weil, M.C. Raff, Role of Ced-3/ICE-family proteases in staurosporine-induced programmed cell death, J. Cell Biol. 133 (1996) 1041 – 1051. M. van Gurp, N. Festjens, G. van Loo, X. Saelens, P. Vandenabeele, Mitochondrial intermembrane proteins in cell death, Biochem. Biophys. Res. Commun. 304 (2003) 487 – 497. P. Li, D. Nijhawan, I. Budihardjo, S.M. Srinivasula, M. Ahmad, E.S. Alnemri, X. Wang, Cytochrome c and dATP-dependent formation of Apaf-1/caspase-9 complex initiates an apoptotic protease cascade, Cell 91 (1997) 479 – 489. M. Irmler, M. Thome, M. Hahne, P. Schneider, K. Hofmann, V. Steiner, J.L. Bodmer, M. Schroter, K. Burns, C. Mattmann, D. Rimoldi, L.E. French, J. Tschopp, Inhibition of death receptor signals by cellular FLIP, Nature 388 (1997) 190 – 195. K. Kuida, T.F. Haydar, C.Y. Kuan, Y. Gu, C. Taya, H. Karasuyama, M.S. Su, P. Rakic, R.A. Flavell, Reduced apoptosis and cytochrome cmediated caspase activation in mice lacking caspase 9, Cell 94 (1998) 325 – 337. C.A. Belmokhtar, J. Hillion, E. Segal-Bendirdjian, Staurosporine induces apoptosis through both caspase-dependent and caspaseindependent mechanisms, Oncogene 20 (2001) 3354 – 3362. E.E. Varfolomeev, M. Schuchmann, V. Luria, N. Chiannilkulchai, J.S. Beckmann, I.L. Mett, D. Rebrikov, V.M. Brodianski, O.C. Kemper, O. Kollet, T. Lapidot, D. Soffer, T. Sobe, K.B. Avraham, T. Goncharov, H. Holtmann, P. Lonai, D. Wallach, Targeted disruption of the mouse Caspase 8 gene ablates cell death induction by the

C.J. Schamberger et al. / Experimental Cell Research 302 (2005) 115–128

[21]

[22]

[23]

[24]

[25]

[26]

[27]

[28]

[29]

[30]

[31]

[32]

[33]

[34]

[35]

[36]

[37]

TNF receptors, Fas/Apo1, and DR3 and is lethal prenatally, Immunity 9 (1998) 267 – 276. C. Cerni, K. Patocka, G. Meneguzzi, Immortalization of primary rat embryo cells by human papillomavirus type 11 DNA is enhanced upon cotransfer of ras, Virology 177 (1990) 427 – 436. C. Cerni, K. Bousset, C. Seelos, H. Burkhardt, M. Henriksson, B. Luscher, Differential effects by Mad and Max on transformation by cellular and viral oncoproteins, Oncogene 11 (1995) 587 – 596. I. Simonitsch, D. Polgar, M. Hajek, P. Duchek, B. Skrzypek, S. Fassl, A. Lamprecht, G. Schmidt, G. Krupitza, C. Cerni, The cytoplasmic truncated receptor tyrosine kinase ALK homodimer immortalizes and cooperates with ras in cellular transformation, FASEB J. 15 (2001) 1416 – 1418. K.W. Sommer, C.J. Schamberger, G.E. Schmidt, S. Sasgary, C. Cerni, Inhibitor of apoptosis protein (IAP) survivin is upregulated by oncogenic c-HRas, Oncogene 22 (2003) 4266 – 4280. A. Cossarizza, M. Baccarani-Contri, G. Kalashnikova, C. Franceschi, A new method for the cytofluorimetric analysis of mitochondrial membrane potential using the J-aggregate forming lipophilic cation 5,5V,6,6V-tetrachloro-1,1V,3,3V-tetraethylbenzimidazolcarbocyanine iodide (JC-1), Biochem. Biophys. Res. Commun. 197 (1993) 40 – 45. M. Reers, S.T. Smiley, C. Mottola-Hartshorn, A. Chen, M. Lin, L.B. Chen, Mitochondrial membrane potential monitored by JC-1 dye, Methods Enzymol. 260 (1995) 406 – 417. E.A. Slee, M.T. Harte, R.M. Kluck, B.B. Wolf, C.A. Casiano, D.D. Newmeyer, H.G. Wang, J.C. Reed, D.W. Nicholson, E.S. Alnemri, D.R. Green, S.J. Martin, Ordering the cytochrome c-initiated caspase cascade: hierarchical activation of caspases-2, -3, -6, -7, -8, and -10 in a caspase-9-dependent manner, J. Cell Biol. 144 (1999) 281 – 292. X. Liu, H. Zou, C. Slaughter, X. Wang, DFF, a heterodimeric protein that functions downstream of caspase-3 to trigger DNA fragmentation during apoptosis, Cell 89 (1997) 175 – 184. M. Enari, H. Sakahira, H. Yokoyama, K. Okawa, A. Iwamatsu, S. Nagata, Acaspase-activated DNase that degrades DNA during apoptosis, and its inhibitor ICAD, Nature 391 (1998) 43 – 50. B.B. Wolf, M. Schuler, F. Echeverri, D.R. Green, Caspase-3 is the primary activator of apoptotic DNA fragmentation via DNA fragmentation factor-45/inhibitor of caspase-activated DNase inactivation, J. Biol. Chem. 274 (1999) 30651 – 30656. J. Grossmann, S. Mohr, E.G. Lapentina, C. Fiocchi, A.D. Levine, Sequential and rapid activation of select caspases during apoptosis of normal intestinal epithelial cells, Am. J. Physiol. 274 (1998) G1117 – G1124. R.V. Talanian, C. Quinlan, S. Trautz, M.C. Hackett, J.A. Mankovich, D. Banach, T. Ghayur, K.D. Brady, W.W. Wong, Substrate specificities of caspase family proteases, J. Biol. Chem. 272 (1997) 9677 – 9682. C.M. Wolf, A. Eastman, The temporal relationship between protein phosphatase, mitochondrial cytochrome c release, and caspase activation in apoptosis, Exp. Cell Res. 247 (1999) 505 – 513. Y. Emoto, Y. Manome, G. Meinhardt, H. Kisaki, S. Kharbanda, M. Robertson, T. Ghayur, W.W. Wong, R. Kamen, R. Weichselbaum, et al., Proteolytic activation of protein kinase C delta by an ICE-like protease in apoptotic cells, EMBO J. 14 (1995) 6148 – 6156. T. Ghayur, M. Hugunin, R.V. Talanian, S. Ratnofsky, C. Quinlan, Y. Emoto, P. Pandey, R. Datta, Y. Huang, S. Kharbanda, H. Allen, R. Kamen, W. Wong, D. Kufe, Proteolytic activation of protein kinase C delta by an ICE/CED 3-like protease induces characteristics of apoptosis, J. Exp. Med. 184 (1996) 2399 – 2404. I. Dal Pra, J.F. Whitfield, A. Chiarini, U. Armato, Changes in nuclear protein kinase C-delta holoenzyme, its catalytic fragments, and its activity in polyomavirus-transformed pyF111 rat fibroblasts while proliferating and following exposure to apoptogenic topoisomerase-II inhibitors, Exp. Cell Res. 249 (1999) 147 – 160. W. Kolch, Meaningful relationships: the regulation of the Ras/Raf/ MEK/ERK pathway by protein interactions, Biochem. J. 351 (Pt. 2) (2000) 289 – 305.

127

[38] J.P. Medema, C. Scaffidi, F.C. Kischkel, A. Shevchenko, M. Mann, P.H. Krammer, M.E. Peter, FLICE is activated by association with the CD95 death-inducing signaling complex (DISC), EMBO J. 16 (1997) 2794 – 2804. [39] S.M. Srinivasula, M. Ahmad, T. Fernandes-Alnemri, E.S. Alnemri, Autoactivation of procaspase-9 by Apaf-1-mediated oligomerization, Mol. Cell 1 (1998) 949 – 957. [40] S. Salvioli, A. Ardizzoni, C. Franceschi, A. Cossarizza, JC-1, but not DiOC6(3) or rhodamine 123, is a reliable fluorescent probe to assess delta psi changes in intact cells: implications for studies on mitochondrial functionality during apoptosis, FEBS Lett. 411 (1997) 77 – 82. [41] X. Luo, I. Budihardjo, H. Zou, C. Slaughter, X. Wang, Bid, a Bcl2 interacting protein, mediates cytochrome c release from mitochondria in response to activation of cell surface death receptors, Cell 94 (1998) 481 – 490. [42] Y. Tsujimoto, Cell death regulation by the Bcl-2 protein family in the mitochondria, J. Cell Physiol. 195 (2003) 158 – 167. [43] I. Iaccarino, D. Hancock, G. Evan, J. Downward, c-Myc induces cytochrome c release in Rat1 fibroblasts by increasing outer mitochondrial membrane permeability in a Bid-dependent manner, Cell Death Differ. 10 (2003) 599 – 608. [44] D.W. Seol, T.R. Billiar, A caspase-9 variant missing the catalytic site is an endogenous inhibitor of apoptosis, J. Biol. Chem. 274 (1999) 2072 – 2076. [45] J.M. Angelastro, N.Y. Moon, D.X. Liu, A.S. Yang, L.A. Greene, T.F. Franke, Characterization of a novel isoform of caspase-9 that inhibits apoptosis, J. Biol. Chem. 276 (2001) 12190 – 12200. [46] M.A. Benedict, Y. Hu, N. Inohara, G. Nunez, Expression and functional analysis of Apaf-1 isoforms. Extra Wd-40 repeat is required for cytochrome c binding and regulated activation of procaspase-9, J. Biol. Chem. 275 (2000) 8461 – 8468. [47] S.B. Bratton, G. Walker, S.M. Srinivasula, X.M. Sun, M. Butterworth, E.S. Alnemri, G.M. Cohen, Recruitment, activation and retention of caspases-9 and -3 by Apaf-1 apoptosome and associated XIAP complexes, EMBO J. 20 (2001) 998 – 1009. [48] H.M. Beere, B.B. Wolf, K. Cain, D.D. Mosser, A. Mahboubi, T. Kuwana, P. Tailor, R.I. Morimoto, G.M. Cohen, D.R. Green, Heatshock protein 70 inhibits apoptosis by preventing recruitment of procaspase-9 to the Apaf-1 apoptosome, Nat. Cell Biol. 2 (2000) 469 – 475. [49] P. Pandey, A. Saleh, A. Nakazawa, S. Kumar, S.M. Srinivasula, V. Kumar, R. Weichselbaum, C. Nalin, E.S. Alnemri, D. Kufe, S. Kharbanda, Negative regulation of cytochrome c-mediated oligomerization of Apaf-1 and activation of procaspase-9 by heat shock protein 90, EMBO J. 19 (2000) 4310 – 4322. [50] K. Maruyama, J.H. Hartwig, T.P. Stossel, Cytochalasin B and the structure of actin gels. II. Further evidence for the splitting of F-actin by cytochalasin B, Biochim. Biophys. Acta 626 (1980) 494 – 500. [51] P.A. Maher, Inhibition of the tyrosine kinase activity of the fibroblast growth factor receptor by the methyltransferase inhibitor 5V-methylthioadenosine, J. Biol. Chem. 268 (1993) 4244 – 4249. [52] S.H. Lee, Y.D. Cho, Induction of apoptosis in leukemia U937 cells by 5V-deoxy-5V-methylthioadenosine, a potent inhibitor of protein carboxylmethyltransferase, Exp. Cell Res. 240 (1998) 282 – 292. [53] C.J. Schamberger, C. Gerner, C. Cerni, bFGF rescues 423-cells from serum starvation-induced apoptosis downstream of activated caspase3, FEBS Lett. 573 (2004) 19–25. [54] M. van Engel, H.J. Kuijpers, F.C. Ramaekers, C.P. Reutelingsperger, B. Schutte, Plasma membrane alterations and cytoskeletal changes in apoptosis, Exp. Cell Res. 235 (1997) 421 – 430. [55] A. Simm, G. Bertsch, H. Frank, U. Zimmermann, J. Hoppe, Cell death of AKR-2B fibroblasts after serum removal: a process between apoptosis and necrosis, J. Cell Sci. 110 (1997) 819 – 828. [56] A. Batistatou, L.A. Greene, Internucleosomal DNA cleavage and neuronal cell survival/death, J. Cell Biol. 122 (1993) 523 – 532. [57] R. Haviv, L. Lindenboim, J. Yuan, R. Stein, Need for caspase-2 in

128

[58]

[59]

[60]

[61]

[62]

[63]

[64] [65]

[66] [67]

[68]

[69]

C.J. Schamberger et al. / Experimental Cell Research 302 (2005) 115–128 apoptosis of growth-factor-deprived PC12 cells, J. Neurosci. Res. 52 (1998) 491 – 497. A.O. Hueber, M. Zornig, D. Lyon, T. Suda, S. Nagata, G.I. Evan, Requirement for the CD95 receptor-ligand pathway in c-Myc-induced apoptosis, Science 278 (1997) 1305 – 1309. T. Nakagawa, H. Zhu, N. Morishima, E. Li, J. Xu, B.A. Yankner, J. Yuan, Caspase-12 mediates endoplasmic-reticulum-specific apoptosis and cytotoxicity by amyloid-beta, Nature 403 (2000) 98 – 103. W. Zhu, A. Cowie, G.W. Wasfy, L.Z. Penn, B. Leber, D.W. Andrews, Bcl-2 mutants with restricted subcellular location reveal spatially distinct pathways for apoptosis in different cell types, EMBO J. 15 (1996) 4130 – 4141. S.T. Lee, K.P. Hoeflich, G.W. Wasfy, J.R. Woodgett, B. Leber, D.W. Andrews, D.W. Hedley, L.Z. Penn, Bcl-2 targeted to the endoplasmic reticulum can inhibit apoptosis induced by Myc but not etoposide in Rat-1 fibroblasts, Oncogene 18 (1999) 3520 – 3528. J. Hacki, L. Egger, L. Monney, S. Conus, T. Rosse, I. Fellay, C. Borner, Apoptotic crosstalk between the endoplasmic reticulum and mitochondria controlled by Bcl-2, Oncogene 19 (2000) 2286 – 2295. M.G. Annis, N. Zamzami, W. Zhu, L.Z. Penn, G. Kroemer, B. Leber, D.W. Andrews, Endoplasmic reticulum localized Bcl-2 prevents apoptosis when redistribution of cytochrome c is a late event, Oncogene 20 (2001) 1939 – 1952. S.M. Frisch, Evidence for a function of death-receptor-related, deathdomain containing proteins in anoikis, Curr. Biol. 9 (1999) 1047 – 1049. M. Rytomaa, L.M. Martins, J. Downward, Involvement of FADD and caspase-8 signalling in detachment-induced apoptosis, Curr. Biol. 9 (1999) 1043 – 1046. S. Cory, D.C. Huang, J.M. Adams, The Bcl-2 family: roles in cell survival and oncogenesis, Oncogene 22 (2003) 8590 – 8607. H. Li, H. Zhu, C.J. Xu, J. Yuan, Cleavage of BID by caspase 8 mediates the mitochondrial damage in the Fas pathway of apoptosis, Cell 94 (1998) 491 – 501. S.M. Srinivasula, M. Ahmad, Y. Guo, Y. Zhan, Y. Lazebnik, T. Fernandes-Alnemri, E.S. Alnemri, Identification of an endogenous dominant-negative short isoform of caspase-9 that can regulate apoptosis, Cancer Res. 59 (1999) 999 – 1002. C.A. Belmokhtar, J. Hillion, C. Dudognon, S. Fiorentino, M. Flexor, M. Lanotte, E. Segal-Bendirdjian, Apoptosome-independent pathway for apoptosis. Biochemical analysis of APAF-1 defects and biological outcomes, J. Biol. Chem. 278 (2003) 29571 – 29580.

[70] D. Sanchis, M. Mayorga, M. Ballester, J.X. Comella, Lack of Apaf-1 expression confers resistance to cytochrome c-driven apoptosis in cardiomyocytes, Cell Death Differ. 10 (2003) 977 – 986. [71] L. Jia, S.M. Srinivasula, F.T. Liu, A.C. Newland, T. FernandesAlnemri, E.S. Alnemri, S.M. Kelsey, Apaf-1 protein deficiency confers resistance to cytochrome c-dependent apoptosis in human leukemic cells, Blood 98 (2001) 414 – 421. [72] J.R. Liu, A.W. Opipari, L. Tan, Y. Jiang, Y. Zhang, H. Tang, G. Nunez, Dysfunctional apoptosome activation in ovarian cancer: implications for chemoresistance, Cancer Res. 62 (2002) 924 – 931. [73] M.S. Soengas, P. Capodieci, D. Polsky, J. Mora, M. Esteller, X. OpitzAraya, R. McCombie, J.G. Herman, W.L. Gerald, Y.A. Lazebnik, C. Cordon-Cardo, S.W. Lowe, Inactivation of the apoptosis effector Apaf-1 in malignant melanoma, Nature 409 (2001) 207 – 211. [74] S.A. Susin, H.K. Lorenzo, N. Zamzami, I. Marzo, C. Brenner, N. Larochette, M.C. Prevost, P.M. Alzari, G. Kroemer, Mitochondrial release of caspase-2 and -9 during the apoptotic process, J. Exp. Med. 189 (1999) 381 – 394. [75] P. Costantini, J.M. Bruey, M. Castedo, D. Metivier, M. Loeffler, S.A. Susin, L. Ravagnan, N. Zamzami, C. Garrido, G. Kroemer, Preprocessed caspase-9 contained in mitochondria participates in apoptosis, Cell Death Differ. 9 (2002) 82 – 88. [76] G. van Loo, X. Saelens, F. Matthijssens, P. Schotte, R. Beyaert, W. Declercq, P. Vandenabeele, Caspases are not localized in mitochondria during life or death, Cell Death Differ. 9 (2002) 1207 – 1211. [77] M. Potokar, I. Milisav, M. Kreft, M. Stenovec, R. Zorec, Apoptosis triggered redistribution of caspase-9 from cytoplasm to mitochondria, FEBS Lett. 544 (2003) 153 – 159. [78] S. Shimohama, H. Tanino, S. Fujimoto, Differential subcellular localization of caspase family proteins in the adult rat brain, Neurosci. Lett. 315 (2001) 125 – 128. [79] S.T. Suhr, M.C. Senut, J.P. Whitelegge, K.F. Faull, D.B. Cuizon, F.H. Gage, Identities of sequestered proteins in aggregates from cells with induced polyglutamine expression, J. Cell Biol. 153 (2001) 283 – 294. [80] M. MacFarlane, W. Merrison, D. Dinsdale, G.M. Cohen, Active caspases and cleaved cytokeratins are sequestered into cytoplasmic inclusions in TRAIL-induced apoptosis, J. Cell Biol. 148 (2000) 1239 – 1254. [81] D. Dinsdale, J.C. Lee, G. Dewson, G.M. Cohen, M.E. Peter, Intermediate filaments control the intracellular distribution of caspases during apoptosis, Am. J. Pathol. 164 (2004) 395 – 407.