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Staurosporine induces apoptosis through both caspase-dependent and caspase-independent mechanisms


Sensitivity of tumor cells to anticancer therapy depends on the ability of the drug to induce apoptosis. However, multiple signaling pathways control this induction and thus determine this sensitivity. We report here that staurosporine, a well known inducer of apoptosis in a wide range of cell lines, displays distinct ability to trigger apoptosis in two different L1210 sublines (termed L1210/S and L1210/0). Staurosporine treatment resulted in an early cell death (within 3 h) in L1210/S cells, while in L1210/0 cells, death occurred only after 12 h. In both instances, death occurred by apoptosis. A broad spectrum caspase inhibitor, Z-VAD-fmk, blocked early apoptosis in L1210/S cells but did not confer any protection on late apoptosis in L1210/0 cells. Protection by Z-VAD-fmk observed in L1210/S cells was not lasting and unmasked a secondary process of cell death that also exhibited characteristics of apoptosis. Thus, staurosporine induces apoptotic cell death through at least two redundant parallel pathways. These two pathways normally coexist in L1210/S cells. However, the early cell death mechanism depending on caspase activation disguises the late caspase-independent apoptotic process. Staurosporine-induced apoptosis in L1210/0 cells develops only by the caspase-independent mechanism due to a general defect in caspase activation.


Apoptosis is a process essential to maintain homeostasis in multicellular organisms. It consists of three distinct phases: an initiation phase, triggered by a variety of physiological agents, which involves the activation of heterogeneous intracellular signaling pathways; a commitment phase, during which the cells become irreversibly committed to die (McCarthy et al., 1997; Brunet et al., 1998); and an execution phase, during which the characteristic morphological changes (membrane blebbing, cell shrinkage, condensation of chromatin) become obvious (Wyllie et al., 1980).

Genetic and biochemical data indicate that a family of cysteine proteases with aspartate specificity, known as caspases, play a pivotal role in the execution phase (Alnemri et al., 1996; Thornberry et al., 1997). Caspases are expressed as inactive proenzymes and become activated by proteolytic processing at internal aspartate residues when cells receive an apoptosis-inducing signal (Nicholson and Thornberry, 1997). At present, 14 mammalian caspase family members have been described. Some, including caspases-2, -8, -9 and -10, contain large prodomains and are initiators of cell death. Once activated, these initiator caspases in turn activate the executioner caspases such as caspases-3, -6 and -7 which carry small prodomains. These later promote apoptosis by cleaving critical cellular substrates. Among them, poly-(ADP-ribose)polymerase (PARP), a repair enzyme preferentially cleaved by caspases-3 and -7 (Lazebnik et al., 1994), and lamins, components of the nuclear lamina, preferentially cleaved by caspase-6 (Orth et al., 1996; Takahashi et al., 1996). This specific proteolysis induce the morphological and biochemical features of apoptosis (Wolf and Green, 1999). Therefore, caspases can activate each other and form an amplified protease cascade reminiscent of the protease cascade seen in clotting and complement activation.

At first, the results of numerous studies seemed to support the idea that caspases play general and essential roles in mammalian programmed cell death (Kumar and Lavin, 1996). Indeed, caspases were activated in virtually all programmed cell death with apoptotic morphology so far examined. Furthermore, inhibition of caspase activities by the use of broad-spectrum caspase inhibitors (Z-VAD-fmk or p35) resulted in the inhibition of apoptosis as assessed by the appearance of apoptotic nuclear morphology and oligonucleosomal DNA fragmentation, a biochemical hallmark of apoptosis. Overall, if most studies have established that inhibition of caspase activity blocks the morphological features of apoptosis-induced by a variety of stimuli and propounded the idea that caspases are critical determinants of apoptosis, these studies did not necessarily imply that the process of caspase activation is the sole determinant of life and death decisions in programmed cell death. In addition, the ability of these compounds to block the apoptotic process and allow cells to recover completely and resume cell growth and division remains controversial. Moreover, an increasing number of recently reported cases have shown that inhibition of caspase activities in mammalian apoptotic systems fails to prevent cell death (Lavoie et al., 1998; Miller et al., 1997; Xiang et al., 1996). In addition, it was suggested that certain cytoplasmic hallmarks of apoptosis may be triggered by enzymes other than caspases, but that nuclear events required caspase activity (McCarthy et al., 1997). Although such caspase-independent cell death was clearly demonstrated (Deas et al., 1998; Trapani et al., 1998; Tainton et al., 2000), the evidences for caspase-independent cell death were based on observations of cells in which caspase activities have been artificially inhibited by the irreversible tripeptide caspase inhibitor Z-VAD-fmk and therefore rely on its ability to block all known caspase activities so far. However, all caspases are certainly not yet identified, and it was not proven that this inhibitor did not affect other enzymes.

Most chemotherapeutic drugs have been shown to activate common apoptotic pathways in target cells (Dive and Hickman, 1991; Sen and D'Incalci, 1992; Hickman, 1992). However these drugs have widely disparate primary targets, suggesting that either the same pathway for apoptosis can be triggered by various stimuli or several parallel pathways may operate in the same cell (Fernandes and Cotter, 1993; McConkey et al., 1989; Alnemri and Litwack, 1990; Bertrand et al., 1994). Indeed, we have previously shown that at least two different apoptosis pathways could be initiated by two unrelated inducers (cisplatin and staurosporine), each leading to the activation of two distinct endonucleases (Ségal-Bendirdjian and Jacquemin-Sablon, 1995).

In keeping with this idea, we have now dissected the response to staurosporine of two cellular variants obtained from a L1210 cell line (L1210/S and L1210/0), in parallel. We showed that staurosporine-induced apoptosis operate through both caspase-dependent and caspase-independent pathway depending on the enzymatic equipment of the cell.

Results and Discussion

Confirmation of the common origin for L1210/S and L1210/0 cells

L1210 cells were originally developed in 1949 as a carcinogen-induced acute lymphoid leukemia in DBA mouse strain (Law et al., 1949). The cells were maintained by serial transplantation in mice and put in an in vitro culture system in 1966, and deposited in the American Type Culture Collection in 1979 (Moore et al., 1966). The L1210/0 subline was first described in 1977 (Burchenal et al., 1977) and obtained from Dr A Eastman (Darmouth, USA). The L1210/S subline, given by Dr S Cros (Toulouse, France), were obtained from ascitic fluid of female DBA/2 mice.

DNA fingerprints were performed on the two cell lines in order to confirm that these cell lines were truly genetical (Figure 1). As assumed, the DNA fingerprints profiles were very similar supporting a common tumor origin for these two cell lines.

Figure 1

DNA fingerprints of L1210 cells. 32P-labeled single-strand M13 DNA was hybridized with Southern blot of restriction endonuclease digests of NB4 (1), L1210/0 (2) and L1210/S (3) cells. Ten μg samples of DNA were digested with BstNI, HinfI, or HaeIII. NB4 DNA is used as unrelated DNA control

L1210/0 shows a delayed response to staurosporine-induced apoptosis

Staurosporine-induced apoptosis was investigated on both cell lines (L1210/S and L1210/0 cells) exposed for 3 h to the drug (5 μM). At this concentration, staurosporine induced a marked decrease (80%) in L1210/S cell viability (Figure 2A, lane 2 compared to the untreated cells, lane 1). This loss of viability was associated with DNA cleavage with a pattern of internucleosomal ladder characteristic of apoptosis. In contrast, in the same conditions (concentration and exposure), staurosporine induced neither significant loss of cell viability nor DNA fragmentation in L1210/0 cells (Figure 2A, lane 6 compared to lane 5). However, extended exposure of these cells for 12 h (Figure 2A, lane 10) induced a marked decrease in L1210/0 cell viability (80% at 12 h of exposure compared to 10% at 3 h of exposure) associated with an extensive DNA degradation into oligonucleosomal fragments. Thus, staurosporine-induce apoptosis was delayed in L1210/0 cells compared with L1210/S cells.

Figure 2

Morphological and biochemical analysis of staurosporine-induced apoptosis in L1210/0 and L1210/S cells. Cells were treated at 37°C with staurosporine (5 μM) with or without the caspase inhibitor, Z-VAD-fmk (400 μM). Z-VAD-fmk was added to cell cultures 1 h prior the addition of staurosporine. A control experiment was performed using Z-Vad-fmk alone. (A) The percentage of survival was determined at 3 h (L1210/S and L1210/0) and 12 h (L1210/0) relative to the untreated controls by WST-1 assay (upper panel). Each point represents a mean of three independent experiments in triplicate. After treatment, cells were harvested, and DNA fragmentation was assayed as described in Materials and methods (lower panel). (B) Caspase-3 like enzyme activity was assayed by the measurement of DEVD-pNa hydrolysis by spectrophotometry, as described in Materials and methods (upper panel). Enzyme activities were measured as initial velocities and expressed as relative intensity/min/mg total protein. Crude lysates were prepared from L1210/S and L1210/0 cells and analysed by Western blotting (lower panel) with antibodies against PARP and lamin B. PARP (116 kDa) and Lamin B (60 kDa) are proteolytically cleaved during caspase-dependent apoptosis. PARP and Lamin B cleavages and the appearance of the respective 85 and 45 kDa proteolytic fragments are clearly shown in staurosporine L1210/S treated cells. Each lane contains the same amount of proteins (15 μg). (C) Assessment of apoptosis of L1210 cells after May-Grunwald-Giemsa staining (upper panel), by DAPI staining (middle panel), or by Annexin V staining (lower panel). L1210/S (a, b) and L1210 (c, d) cells were treated (b, d) or not (a, c) with 5 μM staurosporine for 3 h and 12 h, respectively (lower panel). After treatment, cells were labeled with Annexin-FITC and analysed by fluorescence-activated cell sorter. The percentages in each panel represent the number of cells exhibiting increase Annexin V binding. Ten thousand cells were analysed under each condition

Z-VAD-fmk does not prevent DNA fragmentation and apoptosis in staurosporine-treated L1210/0 cells

As most forms of apoptosis are mediated by the proteolytic actions of caspases (Cohen, 1997), we first examined the involvement of caspases to get insight into effectors of apoptosis that may be differentially involved in L1210/0 and L1210/S cells. Cells were treated with staurosporine (5 μM) in the presence of a potent cell-permeable caspase inhibitor, Z-VAD-fmk (200 μM), then analysed for apoptosis induction and DNA fragmentation. Z-VAD-fmk inhibits a broad spectrum of caspases (Fraser and Evan, 1996), and was shown to block various form of apoptosis by binding as pseudo-substrates to the catalytic site of caspases (Nicholson et al., 1995). Whereas Z-VAD-fmk protected cell death and DNA fragmentation in staurosporine-treated L1210/S cells (Figure 2A, lane 4), it did not provide any protection against a 12 h exposure of L1210/0 cells to staurosporine (Figure 2A, lane 12). Cell death that occurred in staurosporine-treated L1210/0 cells, either in the presence or in the absence of the inhibitor, was of the apoptotic type as indicated by the presence of both morphological (cell shrinkage and condensation of the chromatin) and biochemical features (DNA ladder and Annexin V staining) characteristic of apoptosis (Figure 2A, lane 12 and C). Interestingly, this result shows that phosphatidyl serine externalization on the plasma membrane can occur in the absence of caspase activation. Thus, in contrast with previous results (Coelho et al., 2000; Hirsch et al., 1997; Lemaire et al., 1998; Sarré and Bertrand, 1999; Kim et al., 2000), caspase inhibition did not convert staurosporine-induced cell death from apoptosis to necrosis. These results suggest that, whereas staurosporine-induced apoptosis and DNA fragmentation required caspase activation in L1210/S cells, this mechanism was not used for induction of DNA fragmentation and apoptosis in L1210/0 cells treated by staurosporine. Furthermore, the fact that staurosporine-induced apoptosis in L1210/0 cells proceeds through a caspase-independent pathway is not specific to the drug used, as caspase-independent apoptosis can also be induced in L1210/0 cells by other drugs (cisplatin, etoposide) despite their ability to activate caspases and induce apoptosis in L1210/S cells.

Defective activation of caspases in staurosporine-treated L1210/0 cells

To test the ability of staurosporine to activate caspases, cell lysates from both staurosporine-treated L1210/S and L1210/0 cells were prepared and analysed for caspase-3-like activity in an in vitro chromogenic assay using DEVD-pNa as substrate. The lysate from L1210/S cells treated with 5 μM staurosporine for 3 h showed more than 10-fold increase in caspase-3 like activity (Figure 2B, lane 2). In the presence of the inhibitor, no DEVDase activity was recorded (Figure 2B, lane 4). This result confirms that caspase 3-like proteases are activated following staurosporine-induced apoptosis of the L1210/S cells. In contrast, the lysates of L1210/0 cells treated for either 3 or 12 h with 5 μM staurosporine featured levels of caspase-3-like activity similar to those detected in cell lysates from control cells (Figure 2B, lanes 6 and 10, respectively).

Because Z-VAD-fmk has a broad range specificity, other caspase activities were directly assessed in the same in vitro assay using the chromogenic substrates, Ac-YVAD-pNa (caspase-1), Ac-VDVAD-pNA (caspase-2), Ac-LEVD-pNa (caspase-4), Ac-VEID-pNa (caspase-6), Ac-IETD-pNa (caspase-8), Ac-LEHD-pNa (caspase-9). Although these substrates were far from being very specific, they were used in order to compare caspase activation in L1210/S and L1210/0 cells. Thus, extracts from L1210/0 and L1210/S cells treated with 5 μM staurosporine, for 12 and 3 h respectively, were incubated with the chromogenic peptide substrates. Although treatment of L1210/S cells was accompanied by an increase in activities that cleaved all the substrates tested, excepted Ac-YVAD-pNa (caspase-1), no increase of either activity was detected in extracts prepared from L1210/0 cells (data not shown).

Moreover, no inhibition of caspase activities in extracts from staurosporine-treated L1210/S cells was observed after the addition of extracts obtained from L1210/0 cells, ruling out the possibility that L1210/0 cells contain an endogenous inhibitor of caspase activities that interferes with the assay (data not shown).

PARP is implicated in conserving the integrity of the genome because of its role in DNA repair. This enzyme has therefore been examined in apoptosis and was identified as a substrate for caspases-3 and -7 (Lazebnik et al., 1994; Orth et al., 1996; Takahashi et al., 1996). The cleavage of PARP from a 116-kDa to an 85-kDa form can be detected at an early stage in apoptosis. Cleavage of PARP was compared in L1210/S and L1210/0 staurosporine treated cells (Figure 2B). Whereas the 85-kDa form was readily detected in lysates from L1210/S treated cells, PARP remained intact in L1210/0-treated cells. As expected, Z-VAD-fmk inhibited cleavage of PARP in staurosporine-treated L1210/S cells.

In vitro, caspase-3 can process caspase-6, which is responsible for proteolysis of lamins whose cleavage is required for fragmentation of the nucleus into multiple apoptotic bodies (Lazebnik et al., 1995). Cleavage of lamin B was clearly detected in lysates prepared from L1210/S treated cells but not in lysates from L1210/0 treated cells (Figure 2B). This cleavage was inhibited by Z-VAD-fmk treatment.

Although staurosporine treatment resulted in apoptosis in both cell lines, Z-VAD-fmk only blocked death in L1210/S cells exposed for 3 h to staurosporine. Thus, staurosporine was able to induce apoptosis by two independent routes. One was fast (in L1210/S cells) and caspase-dependent, and the other was slow (in L1210/0 cells) and fully caspase-independent.

A slow caspase-independent pathway for apoptosis exists also in L1210/S cells

We wonder whether the slow apoptosis caspase-independent pathway evidenced in L1210/0 cells did exist also in L1210/S cells. To test this hypothesis, L1210/S cells were treated for 12 h with 5 μM staurosporine in the presence of 400 μM Z-VAD-fmk in order to block completely the rapid caspase-dependent apoptosis pathway. In the absence of Z-VAD-fmk, a 12-h exposure of L1210/S cells to staurosporine induced a massive cell death making all morphological and biochemical analyses impossible. In the presence of Z-VAD-fmk, at least 60% of cell death, associated with internucleosomal DNA fragmentation was observed (Figure 3A, lane 3). However, neither caspase-3 like activation, nor PARP or lamin B cleavages were observed during this cell death (Figure 3B, lane 3). As staurosporine-treated L1210/0 cells, staurosporine-treated L1210/S cells still exhibited morphological and biochemical features of apoptosis (Figure 3A, lane 3 and C, b) despite effective caspase inhibition. In this case also, caspase inhibition therefore did not convert staurosporine-induced cell death from apoptosis to necrosis. Thus, provided caspase activities were blocked, a 12-h treatment of L1210/S cells with staurosporine triggered caspase-independent cell apoptosis.

Figure 3

Morphological and biochemical analysis of staurosporine-induced apoptosis in L1210/S cells in the presence of Z-VAD-fmk. Cells were treated for 12 h at 37°C with staurosporine (5 μM) with the caspase inhibitor, Z-VAD-fmk (400 μM). Z-VAD-fmk was administrated to the cells 1 h prior the addition of staurosporine. A control experiment was performed using Z-VAD-fmk alone. In the absence of Z-VAD-fmk, a 12-h exposure of L1210/S cells to staurosporine induced a massive cell death making all morphological and biochemical analyses impossible. (A) The percentage of survival (upper panel) and DNA fragmentation (lower panel) were determined at 12 h as described in Figure 2A. (B) Caspase-3-like enzyme activity (upper panel) and cleavage of PARP and lamin B (lower panel) were assayed as in Figure 2B. (C) Morphological assessment of apoptosis of L1210/S cells after May-Grunwald-Giemsa staining or by DAPI staining. L1210/S cells were treated (b) or not (a) with 5 μM staurosporine 12 h in the presence of 400 μM Z-VAD-fmk

RNase protection patterns of caspase transcripts

To determine whether a defect in caspase activities in apoptotic L1210/0 cells might result from defects in these proteases gene expression, a RNase protection assay with a multiprobe template for the expression of caspases -1, -2, -3, -6, -7, -8, -11, -12 was performed (Figure 4). This assay included the constitutive genes, glyceraldehyde-3-phosphate dehydrogenase (GAPDH) and the ribosomal L32 for relative quantitation of protected RNA hybrids. Expression of caspase transcripts were compared in L1210/0 and L1210/S cells. This analysis showed that the expression levels of caspases-6, and -7 were minimal in L1210/0 cells. However, no significant differences in the expression levels of capases-3 and -8 were seen in both cell lines. This experiment indicates that although some defect in caspase activities in L1210/0 cells could result from defect at a transcriptional level, they are not sufficient to explain the complete absence of caspase activities.

Figure 4

RNase protection assay (RPA). Total RNA was isolated from L1210/0 and L1210/S cell lines and hybridized with a multiprobe as described in Materials and methods before digestion by RNase. Protected fragments were separated on a denaturating acrylamide gel. Fragment assignment was determined by migration distance relative to unprotected standards. The expression levels of ribosomal L32 and cellular glyceraldehyde-3-phosphate deshydrogenase (GAPDH) serve as internal controls as well as RPA performed with yeast tRNA

Caspase 3 was expressed but not activated during staurosporine-induced apoptosis in L1210/0 cells

The above experiments showed that although caspase-3 and -8 transcripts were expressed in both L1210/0 and L1210/S cells, apoptotic L1210/0 cells were defective in any caspase activities. This might reflects either a failure to express these proteases at the protein level, or a failure to activate them during apoptosis. Moreover, in the case of a rapid, transient activation of caspases, their peaks of activity could have been missed. Therefore, the presence and the processing of caspases-3, -6, -7 and -8 was tested in both cell lines after staurosporine induction of apoptosis. Figure 5 showed that L1210/S cells expressed the proforms of caspases-3, -6, -7 and -8 of 32, 32, 35 and 55 kDa, respectively. Staurosporine treatment of these cells triggered maturation of caspases-3, -6 and -7 as demonstrated by the loss of the pro-caspases-6 and -7, and the appearance of the processed subunit of caspase-3 (17 kDa). Although L1210/S expressed the procaspase-8, its level remained unchanged, indicating that in this cell line staurosporine treatment did not activate this caspase.

Figure 5

Analysis of caspase processing. Equal amounts of extracts (15 μg) prepared from L1210/S and L1210/0 cells treated or not with 5 μM staurosporine for 3 and 12 h, respectively, were resolved by SDS–PAGE and probed with anti-caspases-3, -6, -7 and -8 antibodies as described under Materials and methods. Levels of procaspases-3, -6, -7 and -8 along with the processed caspase-3 and -8 subunits are shown. Activation of caspase-3 and -8 was characterized by a moderate reduction in the intensity of the proforms (32 and 55 kDa, respectively) and the appearance of a band at 17 kDa (for caspase-3) and 43 kDa (for caspase-8). Caspases-6 and -7 activation led to the disappearance of the 32 kDa band. A lysate prepared from untreated and staurosporine-treated Jurkat cells was included as a positive control for caspase-8 activation. In this cell line, staurosporine induced the processed subunit caspase-8

As expected from the RNase protection experiment neither caspase-6 nor caspase-7 were expressed in control or treated L1210/0 cells. Furthermore, although procaspase-3 and -8 were detected in L1210/0 control cells, no processing was induced following staurosporine-induced apoptosis.

Thus, the immunoblot evidence of caspase activation in staurosporine-induced apoptosis of L1210/S cells substantiates the measurements of caspase activities and the detection of caspase substrates cleavage (lamin B and PARP), indicating that the processed caspases were functionally active. On the other hand, the absence of caspase activation in apoptotic L1210/0 cells is consistent with the absence of both caspase activities and substrates cleavage indicating that, in this cell line, apoptosis occurred in a caspase-independent pathway.


We have previously shown that in response to drugs having different primary targets, the apoptotic cell death may operate through different signaling pathways (Ségal-Bendirdjian and Jacquemin-Sablon, 1995). The present results show that a same drug (staurosporine) is able to induce at least two different signaling pathways of cell death, depending on the enzymatic equipment of the cell (Figure 6). The first one is rapid (<3 h) and involves the activation of caspases, whereas the second is slow (>12 h) and caspase-independent. Despite the time differential and the difference in caspase involvement, both forms of staurosporine-induced cell death result in apoptotic cell death. These two pathways normally coexist in L1210/S cells, whereas only the slow caspase-independent pathway occurs in L1210/0 cells, due to a general defect in caspase activation. This defect has apparently developed spontaneously in this clone, and one may speculate that such a spontaneous acquisition might develop more commonly in tumor than recognized so far, leading to chemoresistant phenotypes. These two pathways for apoptosis were not specific to L1210/S cells since both coexist also in staurosporine-treated HL60 cells. However, only the pathway dependent on caspase activation existed in staurosporine-treated Jurkat cells (Belmokhtar et al., 2001, unpublished results).

Figure 6

Staurosporine induces two different signaling pathways of apoptosis. One is rapid and caspase-dependent and develops in L1210/S cells. The other is slower and caspase-independent and develops in L1210/0 cells, which are defective in general caspase activation, and also in L1210/S cells provided caspase activation has been inhibited by Z-VAD-fmk

Our study is in agreement with previous results showing that caspase activation is clearly important in determining the intrinsic resistance of cells to drugs (Los et al., 1997). The L1210 cellular model, reported here (Figure 6), provides a valuable tool to differentiate, in apoptosis, events that are clearly associated with caspase activation from those that are not, and to identify the factors that determine whether a cell enters a caspase-dependent or a caspase-independent mechanism. Our understanding of how cells can be manipulated to use one or the other mechanism should help to better exploit chemotherapeutic agents for trigering tumor cell death.

Materials and methods


Staurosporine was purchased from Roche (France). Stock solution (2.0 mM staurosporine in DMSO) were stored at −20°C. Electrophoresis grade agarose was from Amersham (Biotech, Orsay, France). Proteinase K and DNase-free RNase were obtained from Roche and the DNA molecular weight marker (123 base-pair ladder) from Life Technologies (Cergy-Pontoise, France). Polyvinylidene difluoride (PVDF) membranes (Immobilon-P) and the enhanced chemiluminescence (ECL) detection system were from Amersham. Benzoyloxycarbonyl-Val-Ala-Asp-fluoromethylketone (z-VAD-fmk) was from Bachem. It was diluted to 100 mM in DMSO and used at a final concentration of 200 or 400 μM.

Cell culture and media

The mouse leukemia L1210/0 cell line, originally isolated by Burchenal et al. (1977), was generously provided by Dr A Eastman (Department of Pharmacology, Dartmouth Medical School, Hanover, NH, USA) (Eastman and Bresnik, 1981). The L1210 cell line (called here L1210/S) was obtained from S Cros (Institut de Pharmacologie et de Biologie Structurale, Toulouse, France). L1210/0 cells were grown in suspension in Dulbecco's minimal essential medium (DMEM, life Technologies, Cergy-Pontoise, France) whereas L1210/S cells were grown in RPMI 1640 medium. Both medium were supplemented with 15% fetal calf serum (Life Technologies, Cergy-Pontoise, France), 2 mM glutamine, and antibiotics (streptomycin, 200 μg/ml; penicillin, 200 U/ml). Jurkat cells were grown as L1210/S cells. The acute promyelocytic leukemia cell line, NB4 was cultured as described previously (Lanotte et al., 1991). The cells were grown in a humidified 5% CO2 atmosphere.

To inhibit caspase activities, cells were pretreated for 1 h at 37°C with the broad-spectrum caspase inhibitor, Z-VAD-fmk, before the addition of staurosporine.

Cell viability, morphologic and biochemical assessment of apoptosis

Cell viability was measured by the WST-1 colometric assay (Roche). Data (mean value of triplicates) were expressed as a per cent of untreated control cells. Apoptosis was monitored by cell morphology changes after May-Grünwald-Giemsa staining, typical condensed and fragmented nuclear morphology visible after staining with DAPI (diamino-2-phenyl-indol, Roche), and internucleosomal DNA fragmentation. For measurement of DNA fragmentation, DNA was isolated from 2×106 cells and separated by agarose gel electrophoresis as previously described (Ségal-Bendirdjian and Jacquemin-Sablon, 1995).

DNA fingerprints analysis

DNA from NB4, L1210/0 and L1210/S cells were purified as previously described (Ségal-Bendirdjian and Jacquemin-Sablon, 1995). These DNAs were digested with restriction enzyme (BstNI, HinfI, HaeIII), electrophoresed through a 20-cm long 1.1% agarose gel and transferred by blotting to a Hybond-N nylon membrane (Amersham, Life Science). The membrane was dried and baked at 80°C for 30 min and the DNA was crosslinked onto the membrane by UV radiation for 3 min. Prehybridization is carried out for 1 h at 60°C with a buffer containing 7% SDS, 1 mM EDTA (pH 8), 263 mM Na2HPO4 and 1% bovine serum albumin (fraction V). Hybridization is carried out at 60°C overnight with the addition of 32P-labeled single-strand M13 DNA probe. After hybridization, the membrane was washed in 2 SSC (Trisodium citrate, 30 mM, NaCl, 300 mM, pH 7.0), 0.1% SDS twice at room temperature and once at 60°C, and exposed to X-ray film.

Measurement of caspase activity

Caspase activities were assayed with synthetic substrates selective for different known caspases: YVAD-pNa (caspase-1), VDVAD-pNa (caspase-2), DEVD-pNa (caspase-3), LEVD-pNa (caspase-4), VEID-pNa (caspase-6), IETD-pNa (caspase-8), LEDH-pNa (caspase-9). Cells (2×106) were pelleted, washed with PBS, pH 7.2, and lysed in 50 mM Tris-HC1, pH 7.5, 0.03% NP-40, 1.0 mM DTT. Lysates were centrifuged at 14 000 r.p.m. for 15 min at 4°C. Total protein determination was done using the Bradford assay. Assays were set up in flat bottom 96 well plates containing 0.2 mM of the appropriate peptide substrate in a caspase reaction buffer (100 mM HEPES, pH 7.5, 10% sucrose, 0.1% CHAPS, 10 mM DTT) and 0.01 ml of protein extract (20–50 μg) in a total volume of 0.1 ml. Assays were incubated at 37°C. Release of pNa was detected by periodic readings of absorbance at 405 nm taken against a blank containing buffer and peptide alone (i.e., no extract) from 0–5 h to mark the linearity of the enzymatic reaction in time. Enzyme activities were measured as initial velocities and expressed as relative intensity/min/mg total protein within the linear range of the response.

Annexin V binding analysis

Staining for Annexin V-FITC binding was performed using Annexin V-FITC kit (Euromedex, Bender MedSystem). After washing once in PBS, the cells (7×105) were resuspended in binding buffer (10 mM HEPES/NaOH, pH 7.4, 140 mM Nacl, 2.5 mM Ca Cl2). Annexin V-FITC was added to a final concentration of 1 μg/ml and the cells were incubated at room temperature in the dark for 15 min. The cells were analysed with a Beckton-Dickinson flow cytometer (FASCalibur). Histograms of the change of the mean fluorescence intensity of the Annexin-FITC in control and treated cells were generated.

Ribonuclease protection assay (RPA)

Total cellular RNA was isolated from L1210 cells by using TRIzol (Life Technologies, France) according to the manufacturer's instructions. The expression of various caspase genes was measured by multi-probe RPA using the Pharmingen RiboQuant mAPO-1 kit. The assays were performed according to the manufacturer's protocol (Pharmingen, San Diego, CA, USA). Briefly, the multi-probe template was prepared by 32P incorporation in an in vitro reaction using T7 RNA polymerase. Template DNA was then eliminated by digestion with DNase free of RNase, followed by precipitation of labeled RNA. The probe (1 to 1.5×106 c.p.m. specific activity) was then combined with 20 μg of total cellular RNA and incubated at 90°C for 5 min followed by 56°C for 16 h to allow the hybridization to proceed. The hybridized RNA duplexes were then treated for 45 min with an RNase mixture consisting of RNase A and RNase T1 followed by proteinase K digestion. RNase resistant duplex RNA was then extracted with phenol once and precipitated by the addition of an equal volume of 4 M ammonium acetate and 2 volumes of ethanol. Then, the RNA pellet was solubilized and resolved on a 5% denaturing acrylamide gel, dried and subjected to autoradiography. The identity of the protected fragments were established using the undigested probes as markers.

Western blot analysis

Cleavage of caspases and the caspase substrates, poly (ADP-ribose) polymerase (PARP) and lamin B, was detected by immunoblotting. After washing twice with PBS, 2×106 cells were resuspended in 100 μl of RIPA buffer (Triton X-100 1%, deoxycholate 10%, Tris-Hcl 50 mM pH 7.5, NaCl 150 mM, SDS 0.1%, PMSF 0.1 mM, Leupeptin 10 μg/ml, Aprotinin 10 μg/ml) on ice. After 10 000 g centrifugation at 4°C for 15 min, the supernatants were harvested. Protein concentration in cellular extracts was determined using the bicinchoninic acid (BCA) protein assay reagent. For detection, samples (15 μg/lane) were fractionated on a 10 or 12% SDS–PAGE in a Tris-Glycine running buffer and blotted on polyvinylidene fluoride membranes (Roche). The loading homogeneity and transfer efficiency were checked by staining the membrane with red Ponceau and the gel with Coomassie blue. For caspase detection, the membranes were preblocked overnight at room temperature in PBS containing 5% non-fat milk powder. Afterwards, the blots were incubated 1 h at room temperature with the primary antibody diluted in PBS containing 0.5% non-fat milk powder. For caspase substrate detection, the membranes were blocked 2 h with 5% non fat dry milk powder in PBS and then immunoblotted overnight with the primary antibody. Mouse polyclonal anti-caspase-3 (Transduction Laboratories, Beckton Dickinson), anti-caspase-8 (Pharmingen, Beckton Dickinson), rabbit polyclonal anti-caspases-6 and -7 (Stressgen, Tebu), goat polyclonal anti-lamin B (Santa-Cruz, Tebu) and mouse polyclonal anti-PARP (Oncogene Research, France Biochem), were used as primary antibodies at 1 : 1000, 1 : 2000, 1 : 500, 1 : 500, 1 : 1000 and 1 : 400 dilutions, respectively. After washing, the membranes were then incubated with the respective peroxidase-conjugated secondary antibody for 1 h. Detection was performed using the chemiluminescence procedure (ECL, Amersham, les Ulis, France), according to the manufacturer's recommendations.


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We thank Dr A Eastman (Dartmouth, USA) and Dr S Cros (Toulouse, France) for generously providing the L1210/0 and L1210/S cell lines. We wish to aknowledge the assistance of Dr Macarena Robledo (INSERM EMI 003) who performed the flow cytometry analysis. Special acknowledgment is given to Dr M Lanotte (INSERM U496) for many helpful discussions and comments on this manuscript. This work was supported in part by grants from the Association pour la Recherche contre le Cancer (Villejuif, France) and from the Ligue Nationale contre le Cancer (Paris, France). CA Belmokhtar is supported by a Ligue Nationale contre le Cancer fellowship.

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Correspondence to Evelyne Ségal-Bendirdjian.

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Belmokhtar, C., Hillion, J. & Ségal-Bendirdjian, E. Staurosporine induces apoptosis through both caspase-dependent and caspase-independent mechanisms. Oncogene 20, 3354–3362 (2001).

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  • staurosporine
  • apoptosis
  • caspase
  • L1210

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