Lysosomal enzymes promote mitochondrial oxidant production, cytochrome c release and apoptosis


  • Ming Zhao,

    1. Division of Pathology II, Faculty of Health Sciences, Linköping University, Sweden;
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  • Fernando Antunes,

    1. Division of Pathology II, Faculty of Health Sciences, Linköping University, Sweden;
    2. Grupo de Bioquímica e Biologia Teóricas – Instituto Bento da Rocha Cabral and Department of Chemistry and Biochemistry, Faculty of Sciences, University of Lisbon, Portugal;
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  • John W. Eaton,

    1. Division of Pathology II, Faculty of Health Sciences, Linköping University, Sweden;
    2. James Graham Brown Cancer Center, University of Louisville, Louisville, KY, USA
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  • Ulf T. Brunk

    1. Division of Pathology II, Faculty of Health Sciences, Linköping University, Sweden;
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M. Zhao, Division of Pathology II, Faculty of Health Sciences, Linköping University, SE-581 85 Linköping, Sweden. Fax: +46 13 22 15 29, Tel.: +46 13 22 15 15, E-mail:


Exposure of mammalian cells to oxidant stress causes early (iron catalysed) lysosomal rupture followed by apoptosis or necrosis. Enhanced intracellular production of reactive oxygen species (ROS), presumably of mitochondrial origin, is also observed when cells are exposed to nonoxidant pro-apoptotic agonists of cell death. We hypothesized that ROS generation in this latter case might promote the apoptotic cascade and could arise from effects of released lysosomal materials on mitochondria. Indeed, in intact cells (J774 macrophages, HeLa cells and AG1518 fibroblasts) the lysosomotropic detergent O-methyl-serine dodecylamide hydrochloride (MSDH) causes lysosomal rupture, enhanced intracellular ROS production, and apoptosis. Furthermore, in mixtures of rat liver lysosomes and mitochondria, selective rupture of lysosomes by MSDH promotes mitochondrial ROS production and cytochrome c release, whereas MSDH has no direct effect on ROS generation by purifed mitochondria. Intracellular lysosomal rupture is associated with the release of (among other constituents) cathepsins and activation of phospholipase A2 (PLA2). We find that addition of purified cathepsins B or D, or of PLA2, causes substantial increases in ROS generation by purified mitochondria. Furthermore, PLA2 − but not cathepsins B or D − causes rupture of semipurified lysosomes, suggesting an amplification mechanism. Thus, initiation of the apoptotic cascade by nonoxidant agonists may involve early release of lysosomal constituents (such as cathepsins B and D) and activation of PLA2, leading to enhanced mitochondrial oxidant production, further lysosomal rupture and, finally, mitochondrial cytochrome c release. Nonoxidant agonists of apoptosis may, thus, act through oxidant mechanisms.


arachidonic acid




horseradish peroxidase


lysosomal enzymes


lysosome-mitochondria enriched fraction


O-methyl-serine dodecylamide hydrochloride


phospholipase A2


reactive oxygen species

In the last two decades, the phenomenon of apoptosis has attracted great interest and many intricate molecular events underlying the process have been elucidated [1–8]. Several crucial steps are thought to involve mitochondrial release of pro-apoptotic factors, although the exact mechanisms involved in this release are less well understood.

In this regard, there is substantial evidence that, at least in some circumstances, the discharge into the cytosol of lysosomal constituents may be an early and, perhaps, initiating event in apoptosis, and that mitochondrial release of pro-apoptotic factors might be a consequence of earlier lysosomal destabilization [9–18]. In further, albeit indirect, support of this, it was recently found that activation of the pro-apoptotic tumour supressor protein, p53, also results in early lysosomal rupture, although through still unknown mechanisms [14].

In the case of simple oxidant-induced apoptosis, lysosomal rupture occurs in two sequential phases [19,20], where the second one includes activation of phospholipase A2 (PLA2) with production of free arachidonic acid (AA) [21,22]. Theoretically, released lysosomal enzymes, PLA2, and AA all might be capable of destabilizing mitochondrial membranes. Interestingly, over-expression of the anti-apoptotic protein, Bcl-2, abrogates the secondary phase of lysosomal rupture, the activation of PLA2, and the mitochondrial release of cytochrome c[19,21,22]. However, the precise mechanisms through which Bcl-2 mediates these effects are presently unknown.

Remarkably, in apoptosis caused by a number of nonoxidative agents, there appears to be increased intracellular generation of reactive oxygen species (ROS), probably of mitochondrial origin [23–30]. Although the mechanisms responsible for enhanced mitochondrial ROS production during the process of apoptosis remain unknown, this phenomenon raises the possibility that internally generated ROS, like exogenously added oxidants, may act through a common pathway–lysosomal destabilization.

The present investigations were aimed at identifying intracellular events that might connect exposure of cells to nonoxidative agonists of apoptosis and intracellular ROS production. As mentioned above, there is abundant evidence that − at least in some circumstances − lysosomal rupture might be an early, perhaps even initiating, event in the apoptotic cascade. Therefore, in the present investigations we have used a synthetic lysosomotropic detergent, O-methyl-serine dodecylamide hydrochloride (MSDH) to specifically induce lysosomal rupture and ensuing apoptosis [12,31,32]. This was done in order to determine whether internal oxidative stress of mitochondrial origin might arise as a consequence of lysosomal rupture and act as an amplifying loop causing further lysosomal breach. Here, we present evidence that released lysosomal enzymes − both directly and through activation of PLA2 − may trigger enhanced mitochondrial production of superoxide and hydrogen peroxide, and cause the release of cytochrome c.

Materials and methods


Chemicals were from Sigma unless stated otherwise. RPMI 1640 medium, Hepes, foetal bovine serum, glutamine, penicillin, and streptomycin were from Gibco. BODIPY FL phallacidin and dihydroethidium (DHE) were from Molecular Probes. Monoclonal anti-cytochrome c Igs were from Pharmingen, and horseradish peroxidase (HRP)-conjugated goat anti-mouse Igs were from DAKO. Percoll was from Amersham Pharmacia Biotech.

Cell cultures

Human foreskin fibroblasts (AG-1518, passages 14–20; Coriell Institute, Camden, NJ, USA), J774 cells (a murine histiocytic lymphoma cell line), and human epithelial cells (HeLa) were cultured at 37 °C in humidified air with 5% CO2 in RPMI 1640 medium supplemented with 2 mm glutamine, 50 IU·mL−1 penicillin-G, 50 µg·mL−1 streptomycin, and 10% foetal bovine serum. Cells were subcultured once a week. Twenty-four hours before experiments, cells were trypsinized and seeded into 35-mm Petri dishes or 96-well plates (Costar, Cambridge, MA, USA) at a density of 10 000 cells per cm2.

Apoptosis assays

DNA fragmentation was assessed using propidium iodide staining of nuclear DNA, essentially as described by Nicoletti et al.[33]. Briefly, cell pellets from individual wells were gently resuspended in 1.5 mL of a hypotonic and membrane-disrupting solution of propidium iodide (50 µg·mL−1 in 0.1% sodium citrate with 0.1% Triton X-100) in 12 × 75 mm polypropylene tubes. The tubes were kept overnight in the dark at 4 °C before flow-cytometric analyses. The propidium iodide-induced red fluorescence of suspended individual nuclei was measured by flow cytofluorometry, using the FL3 channel. Nuclei with partly degraded DNA were counted, and their frequency was expressed as a percentage of the total number of nuclei analysed in at least 10 000 cells.

Actin staining

AG1518 fibroblasts were seeded in 35-mm Petri dishes and cultured for 24 h before being exposed to 30 µm MSDH in ordinary medium for 3 h. Cellular actin was stained with BODIPY FL phallacidin. Cells were fixed for 10 min in 4% formaldehyde in NaCl/Pi, permeabilized for 10 min in 0.3% Triton X-100 in phosphate-buffered saline (NaCl/Pi), and stained for 30 min with BODIPY FL phallacidin (final concentration 0.6 µg·mL−1) at 37 °C. After staining, cells were washed twice in NaCl/Pi, and visualized and documented (λEX 495 nm; λEM 520 nm) using a Nikon microphot-SA fluorescence microscope with a Hamamatsu ORCA-100 color digital camera and Adobe photoshop software.

Evaluation of oxidative stress

AG1518 fibroblasts, J774 and HeLa cells were seeded in 96-well plates and cultured for 24 h under standard conditions before being exposed to 30 µm MSDH and 10 µm DHE (in complete medium). Fluorescence intensity, indicating oxidation of DHE was assayed at various periods of time after addition of MSDH and DHE on a VICTOR 1420 (Wallac Sverige AB, Upplands Väsby, Sweden) fluorescent plate-reader (λEX 485 nm; λEM 620 nm). In some experiments, cells were observed and documented under green light excitation (λEX 546 nm; λEM 590 nm) using fluorescence microscopy as described above.

Preparation of rat liver lysosome-mitochondria enriched fraction

Livers were removed from 160–200-g female Sprague–Dawley rats (starved overnight), homogenized in 0.3 m sucrose (1 : 9, w/v) and centrifuged at 500 g for 10 min. The supernatants were again centrifuged at 3500 g for 10 min, the pellets discarded, and the lysosome/mitochondria-containing supernatants centrifuged at 10 000 g for 10 min. The pellets were washed, suspended and re-centrifuged at 10 000 g for 10 min and finally resuspended in the sucrose solution to a protein concentration of ≈ 1.5 mg·mL−1. The resultant lysosome/mitochondria enriched fraction (LEF) was found to be stable (no release of lysosomal enzymes) for up to 4 h in the homogenization solution at 4 °C, while some release of lysosomal enzymes occurred within 2 h at 37 °C.

Preparation of a purified mitochondria fraction

Mitochondria were purified from rat liver using a combination of differential and Percoll gradient centrifugation [34,35]. All procedures were carried out at 4 °C. Briefly, fresh liver was minced and homogenized in M-SHE buffer (0.21 m mannitol, 0.07 m sucrose, 10 mm Hepes pH 7.4, 1 mm EDTA, 1 mm EGTA, 0.15 mm spermine, 0.75 mm spermidine). Unbroken cells and nuclei were pelleted at 500 g for 10 min. The supernatant was centrifuged at 10 000 g to pellet mitochondria and lysosomes which were resuspended and washed twice with M-SHE buffer. A 2-mL suspension was then layered onto 37.5 mL of Percoll solution (50% Percoll, 50% 2 × M-SHE) and centrifuged for 1 h at 50 000 g in a Ti-60 rotor. The brown mitochondrial band was collected, either by fractionating the gradient or by direct syringe aspiration. The purified mitochondria were pooled, diluted 10-fold with M-SHE buffer, again pelleted by centrifugation and, finally, resuspended in M-SHE buffer to a protein concentration of ≈ 1.5 mg·mL−1. The degree of lysosomal contamination of the purified mitochondria fraction was estimated by assaying β-galactosidase/protein and compared to that of LEF.

Enzymatic detection of lysosomal integrity and estimation of fraction purity

The integrity of lysosomes in the LEF preparation was assessed by assaying released β-galactosidase. LEF (200 µL) was incubated for 3 h at 37 °C with either PLA2 (0.2 U·mL−1), 30 µm MSDH, 2.5 µg·mL−1 cathepsin B, or 2.5 µg·mL−1 cathepsin D and then centrifuged at 14 000 g for 10 min. Stock solutions of the cathepsins were made up in NaCl/Pi pH 6.0, whereas MSDH and PLA2 were in NaCl/Pi pH 7.4. The supernatants were removed, and 1 mL distilled water with Triton X-100 (final concentration 0.1%) was added to the pellets to cause complete lysis of remaining intact lysosomes. Activities of β-galactosidase were measured as described previously [22] on the ruptured lysosomal pellet and on the supernatant. The results were expressed as percentage released over total β-galactosidase.

Mitochondrial generation of H2O2

Mitochondrial production of H2O2 was assayed essentially as described elsewhere [36]. Briefly, 1.33 U·mL−1 HRP, 0.066 mg·mL−1ρ-hydroxyphenylacetate, 0.013 mg·mL−1 superoxide dismutase, and 1 mg mitochondrial protein were added to 2.4 mL respiratory buffer (0.07 m sucrose, 0.23 m mannitol, 30 mm Tris/HCl, 4 mm MgCl2, 5 mm KH2PO4, 1 mm EDTA, 0.5% BSA, pH 7.4) in a spectrofluorophotometer cuvette at 37 °C. Succinate (final concentration 6.67 mm) and antimycin A (final concentration 0.83 µg·mL−1) were added, and H2O2-induced fluorescence recorded (λEX 320 nm; λEM 400 nm) during the first 10 min after mixing.

Western blotting for cytochrome c

Two-hundred microlitres LEF, or purified mitochondria, were incubated for 3 h at 37 °C with either 30 µm MSDH, PLA2 (0.2 U·mL−1), 2.5 µg·mL−1 cathepsin B, or 2.5 µg·mL−1 cathepsin D. Stock solutions of the cathepsins were made up in NaCl/Pi pH 6.0, while MSDH and PLA2 were in NaCl/Pi pH 7.4. Following centrifugation at 14 000 g for 10 min, the supernatants were separated by SDS/PAGE (12% acrylamide) and transferred onto Immobilon membranes (2 h; 200 mA). Membranes then were incubated at room temperature for 1 h in blocking buffer [5% low-fat milk powder in Tris-buffered saline (TBS)] and for another 2 h in dilution buffer (0.5% low-fat milk powder in TBS) containing a 1 : 400 dilution of a monoclonal anti-cytochrome c Ig. After washing in TBS with 0.06% Tween 20, Immobilon membranes were incubated for 1 h at room temperature in a buffer containing a 1 : 1500 dilution of peroxidase-conjugated secondary antibodies. After washing, peroxidase-dependent chemiluminescence was detected by using enhanced chemiluminescence Western blotting reagents and hyperfilm according to the manufacturer's instructions (Amersham Pharmacia Biotech).

Statistical analysis

All experiments were repeated at least three times. Values are given as arithmetic mean ± SD. Significance of differences between groups was determined using Student's two-tailed t-test. *P ≤ 0.05; **P ≤ 0.01; ***P ≤ 0.001.


Cultured cells exposed to the synthetic lysosomotropic detergent, MSDH, undergo lysosomal rupture and ensuing apoptosis or necrosis depending upon the extent of lysosomal destabilization [12]. In the present experiments, we induced apoptosis in fibroblasts, J774 cells, and HeLa cells by exposing them to 30 µm MSDH. After 8 h of MSDH exposure, nuclear propidium iodide staining and flow cytometry (to detect DNA fragmentation) revealed apoptotic nuclei appearing as a broad, hypodiploid DNA smear in front of a narrow peak of diploid DNA (Fig. 1A shows results in fibroblasts). At an early stage in this process, well before the appearance of frank apoptosis, fibroblasts showed significantly increased numbers of stress fibres (Fig. 1B).

Figure 1.

MSDH induces apoptosis and stress fibre formation in fibroblasts. (A) Cells were seeded into 35-mm Petri dishes at a density of 10 000 cells·cm−2. After 24 h, 30 µm MSDH was added to complete culture medium (2 mL), and cells were incubated for another 10 h under standard culture conditions. The Nicoletti technique for apoptotic nuclei was applied. One representative experiment out of three is shown. (B) Cells were seeded in 35-mm Petri dishes and kept for 24 h before being exposed to 30 µm MSDH for 3 h. Actin staining was then performed as described in Materials and methods.

Because oxidative stress has been reported to induce stress fibre formation [37], we suspected that the MSDH exposure might be causing increased intracellular generation of ROS. This latter was monitored by following changes in DHE-induced fluorescence. When oxidized, this compound intercalates into DNA and RNA, resulting in an increase in quantum yield. Fluorescence intensity was measured kinetically at indicated time points. Increased ROS production occurred after 1 h of MSDH-exposure in fibroblasts (AG1518) and epithelial cells (HeLa), but was significant only after 3 h in macrophages (J774) (Fig. 2A). Note that in fibroblasts and HeLa cells, and also in J774 cells (results not shown), the oxidation of DHE eventually reached a steady state consistent with only a transient production of ROS. Figure 2B shows DHE-exposed J774 cells after 3 h exposure to MSDH, when there were still no morphological signs of apoptosis.

Figure 2.

MSDH induces intracellular ROS production. Cells were seeded into 96-well plates at a density of 10 000 cells·cm−2. After 24 h, cells were exposed simultaneously to 30 µm MSDH and 10 µm DHE under otherwise standard culture conditions while control cells were exposed to DHE only. (A) Fluorescence intensity arising from oxidized dihydroethidium in J774, HeLa and AG1518 cells was measured at indicated time points. (B) J774 cells were visualized and photographed after 3 h exposure to MSDH (n = 3). Very similar results were obtained with HeLa and AG1518 cells under the same conditions although detectable oxidant generation occurred earlier.

Theoretically, the increased oxidant generation might arise from effects of released lysosomal enzymes (directly or by activation of PLA2) on mitochondrial ROS production or, alternatively, from direct effects of MSDH on the mitochondria. To discriminate between these possibilities, we added MSDH to purified rat liver mitochondria (4.5-fold purified from lysosomal contamination as compared to the LEF preparation, results not shown). Under these conditions, no changes in mitochondrial production of H2O2(Fig. 3A) or release of cytochrome c (Fig. 3B) took place. Because we previously observed that lysosomal contents cause activation of PLA2 in J774 cells [22], we also exposed mitochondria to that enzyme and found it to enhance mitochondrial production of ROS (Fig. 3A) and to release cytochrome c as well (Fig. 3B). These findings strongly suggest that MSDH affects mitochondria by first destabilizing lysosomes and causing the release of hydrolytic enzymes which, in turn, attack mitochondria or activate PLA2. Activated PLA2 may further promote this cascade of events, attacking both mitochondrial and lysosomal membranes and causing further lysosomal rupture. This supposed sequence of events was confirmed by adding MSDH to a lysosome/mitochondria-enriched rat liver fraction, where it was found to induce enhanced mitochondrial production of H2O2 (Fig. 3A), release of cytochrome c (Fig. 3B), and lysosomal rupture (Fig. 3C).

Figure 3.

MSDH induces mitochondrial ROS production by rupturing lysosomes. Purified mitochondria (1.0 mg protein·mL−1) or a lysosome/mitochondria-enriched fraction (1.0 mg protein·mL−1) were incubated with either of MSDH (30 µm), PLA2 (0.2 U·mL−1), or cathepsin B or D (12.5 µg·mL−1; pH 6.0) for 3 h. (A) H2O2 production, (B) cytochrome c release, and (C) lysosomal stability were assayed as described in Materials and methods (n = 3).

To test further the idea that released lysosomal hydrolases might enhance mitochondrial ROS production, release of cytochrome c, and activation of PLA2 (all of which may promote the apoptotic cascade), we tested the effects of two lysosomal cathepsins (cathepsin B, a cysteine protease, and cathepsin D, an aspartic protease) on purified mitochondria. Both proteases caused substantial increases in mitochondrial production of H2O2 (Fig. 3A) and release of cytochrome c (Fig. 3B). However, neither cathepsin B nor D caused detectable lysosomal rupture in LEF preparations (Fig. 3C), although, as expected, both MSDH and PLA2 did induce lysosomal rupture (Fig. 3C).

Thus, cathepsins B and D do not directly cause rupture of lysosomes in an LEF preparation. However, the possibility remains that the intracellular release of other lysosomal hydrolases may do so, or that lysosomal proteases might secondarily destabilize lysosomes through, for example, enhanced oxidative stress or activation of PLA2 following mitochondrial attack by cathepsins and PLA2. Indeed, low, steady-state oxidative stress has been shown to destabilize lysosomes [20] and relocation of lysosomal enzymes to the cytosol was earlier shown to activate PLA2 [22].


We previously suggested that oxidative stress-induced apoptosis might be initiated by iron-catalysed lysosomal rupture [9,10]. It has since been found that early release to the cytosol of lytic lysosomal enzymes may be characteristic of apoptosis caused by a variety of stimuli [10,12–14,19,21,22,38–40]. In these latter circumstances, it appears that relocation of lysosomal enzymes to the cytosol may, as in the case of oxidant-induced apoptosis, precede changes of mitochondrial membrane potential, release of cytochrome c, and all the morphological signs of apoptosis. These considerations raised the question of whether there might be some ROS-dependent mechanisms common to apoptosis caused by oxidants and that caused by nonoxidant agents.

In most cells, the predominant source of intracellular ROS generation is the mitochondrial electron transport chain which, even under normal conditions, may ‘leak’ 1–2% of all electrons as ROS [41–43] (although there is controversy regarding this estimate and the absolute percentage may well be lower [44]). Not only will exogenous oxidants, such as H2O2, directly induce apoptosis, but enhanced intracellular production of ROS occurs when cells are exposed to a number of pro-apoptotic agents including tumour necrosis factor-α[23], ceramide [24], growth factor withdrawal, HIV infection, and lipopolysaccharide [25–30]. In these cases it is unclear whether such oxidative stress is the cause or an effect of apoptosis.

We hypothesized that released lysosomal enzymes or PLA2 directly or indirectly activated by such enzymes [22] might attack mitochondria and induce not only release of cytochrome c, but also enhanced formation of ROS. Released arachidonic acid may further exaggerate this process [45]. These ROS of mitochondrial origin could promote further lysosomal rupture but could also have the secondary effect of maintaining any cytochrome c released by the mitochondria in the oxidized form (although we should note that the cellular cytoplasm contains an abundance of reducing agents which could counteract this). Cytochrome c is involved in the activation of caspase-9 [7,46] and is considered a key component of the apoptotic cascade. Ordinarily, any cytochrome c released from mitochondria in oxidized form would rapidly be reduced by the reductive cytosolic milieu. However, it has been proposed that cytochrome c needs to remain oxidized in order to promote apoptosis [46], and the oxidizing equivalents generated by mitochondria may have precisely this effect.

MSDH is a lysosomotropic detergent that rapidly induces specific lysosomal rupture and therefore is a very useful tool for detailed kinetic studies of the consequences of lysosomal rupture. The pKa of MSDH is 5.8–5.9 [31,32], allowing it to accumulate in charged form intralysosomally (pH ≈ 4.5) due to proton trapping [47], while its accumulation in the cytosol (pH ≈ 7.2) is negligible. In protonated, charged form MSDH acts as a much stronger detergent than when uncharged, further targeting the action of this agent to the lysosomal compartment [31].

We previously reported that released lysosomal enzymes activate PLA2 causing further lysosomal fragmentation [22]. The new data presented here confirm and extend those findings and show that relocated lysosomal enzymes work in concert with activated PLA2, causing the release of cytochrome c, enhanced mitochondrial formation of ROS, and promoting further lysosomal degradation. With regard to the mechanisms involved in enhanced mitochondrial ROS production, one particularly likely possibility is that of generation of free fatty acids. At least in pancreatic beta cell mitochondria, free fatty acids have been shown to increase ROS generation, perhaps through electron leak involving complex I of the respiratory chain [48]. Whether the progressive lysosomal destabilization is dependent exclusively on upstream actions of cathepsins B and D, or whether other lysosomal constituents might similarly destabilize mitochondria and lysosomes is not yet clear.

Our present understanding concerning the involvement of lysosomes in apoptosis is summarized in Fig. 4. As shown, the initiation of apoptosis by exogenous oxidants, and by at least some other agonists, may involve early lysosomal rupture. The release of lysosomal enzymes (LE) into the cell cytoplasm may set off a cascade of intracellular degradative events. These LE may: (a) attack mitochondria directly, inducing release of cytochrome c; (b) directly and/or indirectly cause enhanced formation of mitochondrial ROS (and further oxidant-induced lysosomal destabilization); (c) activate lytic pro-enzymes, such as PLA2, which in turn would attack both mitochondria and lysosomes; (d) activate Bid and/or other pro-apoptotic proteins; and (e) directly activate pro-caspases. Notably, this sequence of early events (except for cytochrome c release) may be relatively independent of the classical apoptotic cascade involving caspase activation. In many circumstances, this ‘lysosomal-mitochondrial axis’ apoptotic pathway, involving combined effects of caspases, lysosomal hydrolases and mitochondrial ROS generation, may be of central importance in the final execution of the apoptotic cascade wherein a lysosomal/mitochondrial cross-talk may constitute an amplifying loop.

Figure 4.

The lysosomal/mitochondrial axis theory of apoptosis. Both the internal and external pathways may involve lysosomal rupture. Released lysosomal enzymes (LE) may: (a) attack mitochondria directly, inducing oxidative stress and release of cytochrome c (this study and [12,20–22,49–52]); (b) activate lytic pro-enzymes, such as PLA2, which may attack both mitochondria or lysosomes (this study and [22]); (c) activate Bid [53]; (d) directly activate caspases [15,16,54,55]. It is also possible that released lysosomal enzymes backfire on still intact lysosomes, causing further rupture. Caspase 8 may somehow induce lysosomal rupture [56,57] or the activation of death receptors may cause production of sphingosine [58], which is a lysosomotropic detergent [59]; while p53 causes lysosomal labilization by unknown mechanisms [14]. Other mechanisms may also be involved in lysosomal labilization in relation to apoptosis.


We thank G. Dubowchik (Bristol-Myers Squibb; Pharmaceutical Research Institute) for the kind gift of MSDH. This study was supported by a grant from the Swedish Cancer Foundation (grant no. 4296). JWE was the recipient of a visiting professorship from the Linköping University Hospital and is supported by The Commonwealth of Kentucky Research Challenge Trust Fund.