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

Article (PDF Available)inEuropean Journal of Biochemistry 270(18):3778-86 · October 2003with44 Reads
DOI: 10.1046/j.1432-1033.2003.03765.x · Source: PubMed
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.
Lysosomal enzymes promote mitochondrial oxidant production,
release and apoptosis
Ming Zhao
, Fernando Antunes
, John W. Eaton
and Ulf T. Brunk
Division of Pathology II, Faculty of Health Sciences, Linko
ping University, Sweden;
Grupo de Bioquı
mica e Biologia Teo
Instituto Bento da Rocha Cabral and Department of Chemistry and Biochemistry, Faculty of Sciences, University of Lisbon, Portugal;
James Graham Brown Cancer Center, University of Louisville, Louisville, KY, USA
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 mito-
chondria. Intracellular lysosomal rupture is associated with
the release of (among other constituents) cathepsins and
activation of phospholipase A2 (PLA2). We find that addi-
tion of purified cathepsins B or D, or of PLA2, causes
substantial increases in ROS generation by purified mito-
chondria. 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.
Keywords: apoptosis; cathepsins; lysosomes; lysosomotropic
detergents; oxidative stress.
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, lyso-
somal 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 intracel-
lular 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 evi-
dence that ) at least in some circumstances ) lysosomal
rupture might be an early, perhaps even initiating, event in
Correspondence to M. Zhao, Division of Pathology II,
Faculty of Health Sciences, Linko
ping University,
SE-581 85 Linko
ping, Sweden.
Fax: +46 13 22 15 29, Tel.: +46 13 22 15 15,
Abbreviations: AA, arachidonic acid; DHE, dihydroethidium; HRP,
horseradish peroxidase; LE, lysosomal enzymes; LEF, lysosome-
mitochondria enriched fraction; MSDH, O-methyl-serine
dodecylamide hydrochloride; PLA2, phospholipase A2;
ROS, reactive oxygen species.
(Received 28 April 2003, revised 11 July 2003,
accepted 24 July 2003)
Eur. J. Biochem. 270, 3778–3786 (2003) FEBS 2003 doi:10.1046/j.1432-1033.2003.03765.x
the apoptotic cascade. Therefore, in the present investiga-
tions 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%
in RPMI 1640 medium supplemented with 2 m
glutamine, 50 IUÆmL
penicillin-G, 50 lgÆmL
mycin, and 10% foetal bovine serum. Cells were subcul-
tured 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 cm
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 lgÆmL
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 cyto-
fluorometry, 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 l
ordinary medium for 3 h. Cellular actin was stained with
BODIPY FL phallacidin. Cells were fixed for 10 min in 4%
formaldehyde in NaCl/P
, permeabilized for 10 min in
0.3% Triton X-100 in phosphate-buffered saline (NaCl/P
and stained for 30 min with BODIPY FL phallacidin
(final concentration 0.6 lgÆmL
)at37C. After staining,
cells were washed twice in NaCl/P
, and visualized and
documented (k
495 nm; k
520 nm) using a Nikon
microphot-SA fluorescence microscope with a Hamamatsu
ORCA-100 color digital camera and Adobe
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 l
MSDH and
10 l
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 Va
sby, Sweden)
fluorescent plate-reader (k
485 nm; k
620 nm). In
some experiments, cells were observed and documented
under green light excitation (k
546 nm; k
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
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/mitochon-
dria-containing supernatants centrifuged at 10 000 g for
10 min. The pellets were washed, suspended and re-centri-
fuged at 10 000 g for 10 min and finally resuspended in
the sucrose solution to a protein concentration of
1.5 mgÆmL
. 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
Preparation of a purified mitochondria fraction
Mitochondria were purified from rat liver using a combi-
nation 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
mannitol, 0.07
sucrose, 10 m
Hepes pH 7.4,
EDTA, 1 m
EGTA, 0.15 m
spermine, 0.75 m
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,
FEBS 2003 Lysosomes, apoptosis and mitochondria-mediated oxidative stress (Eur. J. Biochem. 270) 3779
resuspended in M-SHE buffer to a protein concentration
of 1.5 mgÆmL
. The degree of lysosomal contamination
of the purified mitochondria fraction was estimated by
assaying b-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 b-galactosidase. LEF
(200 lL) was incubated for 3 h at 37 C with either
PLA2 (0.2 UÆmL
), 30 l
MSDH, 2.5 lgÆmL
sin B, or 2.5 lgÆmL
cathepsin D and then centrifuged at
14 000 g for 10 min. Stock solutions of the cathepsins were
made up in NaCl/P
pH 6.0, whereas MSDH and PLA2
were in NaCl/P
pH 7.4. The supernatants were removed,
and 1 mL distilled water with Triton X-100 (final concen-
tration 0.1%) was added to the pellets to cause complete
lysis of remaining intact lysosomes. Activities of b-galac-
tosidase were measured as described previously [22] on the
ruptured lysosomal pellet and on the supernatant. The
results were expressed as percentage released over total
Mitochondrial generation of H
Mitochondrial production of H
was assayed essentially
as described elsewhere [36]. Briefly, 1.33 UÆmL
0.066 mgÆmL
q-hydroxyphenylacetate, 0.013 mgÆmL
superoxide dismutase, and 1 mg mitochondrial protein
were added to 2.4 mL respiratory buffer (0.07
mannitol, 30 m
Tris/HCl, 4 m
EDTA, 0.5% BSA, pH 7.4) in a spectro-
fluorophotometer cuvette at 37 C. Succinate (final concen-
tration 6.67 m
) and antimycin A (final concentration
0.83 lgÆmL
) were added, and H
-induced fluorescence
recorded (k
320 nm; k
400 nm) during the first 10 min
after mixing.
Western blotting for cytochrome
Two-hundred microlitres LEF, or purified mitochondria,
were incubated for 3 h at 37 C with either 30 l
MSDH, PLA2 (0.2 UÆmL
), 2.5 lgÆmL
cathepsin B, or
2.5 lgÆmL
cathepsin D. Stock solutions of the cathepsins
were made up in NaCl/P
pH 6.0, while MSDH and PLA2
were in NaCl/P
pH 7.4. Following centrifugation at
14 000 g for 10 min, the supernatants were separated by
Fig. 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 cellsÆcm
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 l
MSDH for 3 h. Actin
staining was then performed as described in
Materials and methods.
3780 M. Zhao et al. (Eur. J. Biochem. 270) FEBS 2003
SDS/PAGE (12% acrylamide) and transferred onto Immo-
bilon 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 mono-
clonal 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 chemilu-
minescence was detected by using enhanced chemilumines-
cence Western blotting reagents and hyperfilm according to
the manufacturer’s instructions (Amersham Pharmacia
Statistical analysis
All experiments were repeated at least three times. Values
are given as arithmetic mean ± SD. Significance
Fig. 2. MSDH induces intracellular ROS production. Cells were seeded into 96-well plates at a density of 10 000 cellsÆcm
. After 24 h, cells were
exposed simultaneously to 30 l
MSDH and 10 l
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 ¼ 3). Very similar results were obtained with HeLa and AG1518
cells under the same conditions although detectable oxidant generation occurred earlier.
FEBS 2003 Lysosomes, apoptosis and mitochondria-mediated oxidative stress (Eur. J. Biochem. 270) 3781
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 l
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).
Fig. 3. MSDH induces mitochondrial ROS production by rupturing lysosomes. Purified mitochondria (1.0 mg proteinÆmL
) or a lysosome/mito-
chondria-enriched fraction (1.0 mg proteinÆmL
) were incubated with either of MSDH (30 l
), PLA2 (0.2 UÆmL
), or cathepsin B or D
(12.5 lgÆmL
production, (B) cytochrome c release, and (C) lysosomal stability were assayed as described in Materials
and methods (n ¼ 3).
3782 M. Zhao et al. (Eur. J. Biochem. 270) FEBS 2003
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 kineti-
cally 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 expo-
sure to MSDH, when there were still no morphological
signs of apoptosis.
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
(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 mito-
chondrial production of H
(Fig. 3A), release of cyto-
chrome c (Fig. 3B), and lysosomal rupture (Fig. 3C).
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 substan-
tial increases in mitochondrial production of H
(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
Fig. 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.
FEBS 2003 Lysosomes, apoptosis and mitochondria-mediated oxidative stress (Eur. J. Biochem. 270) 3783
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 cyto-
chrome 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 nonoxi-
dant 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 H
, 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-a [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 abun-
dance 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 exclu-
sively on upstream actions of cathepsins B and D, or
whether other lysosomal constituents might similarly desta-
bilize 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 destabiliza-
tion); (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, invol-
ving combined effects of caspases, lysosomal hydrolases and
mitochondrial ROS generation, may be of central import-
ance in the final execution of the apoptotic cascade wherein
a lysosomal/mitochondrial cross-talk may constitute an
amplifying loop.
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
ping University Hospital and is supported by The Common-
wealth of Kentucky Research Challenge Trust Fund.
1. Nicholson, D.W. (2000) From bench to clinic with apoptosis-
based therapeutic agents. Nature 407, 810–816.
2. Nicholson, D.W. (2001) Apoptosis. Baiting death inhibitors.
Nature 410, 33–34.
3. Savill, J. & Fadok, V. (2000) Corpse clearance defines the meaning
of cell death. Nature 407, 784–788.
4. Baumann, S., Krueger, A., Kirchhoff, S. & Krammer, P.H. (2002)
Regulation of T cell apoptosis during the immune response. Curr.
Mol. Med. 2, 257–272.
3784 M. Zhao et al. (Eur. J. Biochem. 270) FEBS 2003
5. Yuan, J. & Yankner, B.A. (2000) Apoptosis in the nervous system.
Nature 407, 802–809.
6. Kaufmann, S.H. & Hengartner, M.O. (2001) Programmed cell
death: alive and well in the new millennium. Trends Cell Biol. 11,
7. Hengartner, M.O. (2000) The biochemistry of apoptosis. Nature
407, 770–776.
8. Zakeri,Z.&Lockshin,R.A.(2002)Celldeathduringdevelop-
ment. J. Immunol. Methods 265, 3–20.
9. Zdolsek, J., Zhang, H., Roberg, K. & Brunk, U. (1993) H
mediated damage to lysosomal membranes of J-774 cells. Free
Radic. Res. Commun. 18, 71–85.
10. O
llinger, K. & Brunk, U.T. (1995) Cellular injury induced by
oxidative stress is mediated through lysosomal damage. Free
Radic. Biol. Med. 19, 565–574.
Photo-oxidative disruption of lysosomal membranes causes
apoptosis of cultured human fibroblasts. Free Radic. Biol. Med.
23, 616–626.
12. Li, W., Yuan, X., Nordgren, G., Dalen, H., Dubowchik, G.M.,
Firestone, R.A. & Brunk, U.T. (2000) Induction of cell death by
the lysosomotropic detergent MSDH. FEBS Lett. 470, 35–39.
13. Brunk, U.T., Neuzil, J. & Eaton, J.W. (2001) Lysosomal
involvement in apoptosis. Redox Report 6, 91–97.
14. Yuan, X.M., Li, W., Dalen, H., Lotem, J., Kama, R., Sachs, L. &
Brunk, U.T. (2002) Lysosomal destabilization in p53-induced
apoptosis. Proc. Natl Acad. Sci. USA 99, 6286–6291.
15. Ishisaka,R.,Utsumi,T.,Kanno,T.,Arita,K.,Katunuma,N.,
Akiyama, J. & Utsumi, K. (1999) Participation of a cathepsin
1-type protease in the activation of caspase-3. Cell Struct. Funct.
24, 465–470.
16. Katunuma, N., Matsui, A., Le, Q.T., Utsumi, K., Salvesen, G. &
Ohashi, A. (2001) Novel procaspase-3 activating cascade mediated
by lysoapoptases and its biological significances in apoptosis. Adv.
Enz. Reg. 41, 237–250.
17. Bursch, W. (2001) The autophagosomal-lysosomal compartment
in programmed cell death. Cell Death Differ. 8, 569–581.
18. Salvesen, G.S. (2001) A lysosomal protease enters the death scene.
J. Clin. Invest. 107, 21–22.
19. Zhao, M., Eaton, J.W. & Brunk, U.T. (2000) Protection against
oxidant-mediated lysosomal rupture: a new anti-apoptotic activity
of Bcl-2? FEBS Lett. 485, 104–108.
20. Antunes, F., Cadenas, E. & Brunk, U.T. (2001) Apoptosis
induced by exposure to a low steady-state concentration of
is a consequence of lysosomal rupture. Biochem. J. 365,
21. Zhao, M., Eaton, J.W. & Brunk, U.T. (2001) Bcl-2 phosphory-
lation is required for inhibition of oxidative stress-induced lyso-
somal leak and ensuing apoptosis. FEBS Lett. 509, 405–412.
22. Zhao, M., Brunk, U.T. & Eaton, J.W. (2001) Delayed oxidant-
induced cell death involves activation of phospholipase A2. FEBS
Lett. 509, 399–404.
23. Obrador, E., Navarro, J., Mompo, J., Asensi, M., Pellicer, J.A. &
Estrela, J.M. (1998) Regulation of tumour cell sensitivity to TNF-
induced oxidative stress and cytotoxicity: role of glutathione.
Biofactors 8, 23–26.
24. Andrieu-Abadie, N., Gouaze, V., Salvayre, R. & Levade, T. (2001)
Ceramide in apoptosis signaling: relationship with oxidative stress.
Free Radic. Biol. Med. 31, 717–728.
25. Muller, J.M., Ziegler-Heitbrock, H.W.L. & Baeuerle, P.A. (1993)
Nuclear factor kappa B, a mediator of lipopolysaccharide effects.
Immunobiology 187, 233–256.
26. Albrecht, H., Tschopp, J. & Jongeneel, C.V. (1994) Bcl-2 protects
from oxidative damage and apoptotic cell death without inter-
fering with activation of NF-kappa B by TNF. FEBS Lett. 351,
27. Atabay, C., Cagnoli, C.M., Kharlamov, E., Ikonomovic, M.D. &
Manev, H. (1996) Removal of serum from primary cultures of
cerebellar granule neurons induces oxidative stress and DNA
fragmentation: protection with antioxidants and glutamate
receptor antagonists. J. Neurosci. Res. 43, 465–475.
28. Degli Esposti, M. & McLennan, H. (1998) Mitochondria
and cells produce reactive oxygen species in virtual anaerobiosis:
relevance to ceramide-induced apoptosis. FEBS Lett. 430,
29. Garcia-Ruiz, C., Colell, A., Mari, M., Morales, A. & Fernandez-
Checa, J.C. (1997) Direct effect of ceramide on the mitochondrial
electron transport chain leads to generation of reactive oxygen
species. Role of mitochondrial glutathione. J. Biol. Chem. 272,
30. Dobmeyer, T.S., Findhammer, S., Dobmeyer, J.M., Klein, S.A.,
Raffel, B., Hoelzer, D., Helm, E.B., Kabelitz, D. & Rossol, R.
(1997) Ex vivo induction of apoptosis in lymphocytes is mediated
by oxidative stress: role for lymphocyte loss in HIV infection. Free
Radic. Biol. Med. 22, 775–785.
31. Firestone, R.A., Pisano, J.M. & Bonney, R.J. (1979) Lysosomo-
tropic agents. 1. Synthesis and cytotoxic action of lysosomotropic
detergents. J. Med. Chem. 22, 1130–1133.
32. Wilson, P.D., Firestone, R.A. & Lenard, J. (1987) The role of
lysosomal enzymes in killing of mammalian cells by the lyso-
somotropic detergent N-dodecylimidazole. J. Cell Biol. 104,
33. Nicoletti, I., Migliorati, G., Pagliacci, M.C., Grignani, F. &
Riccardi, C. (1991) A rapid and simple method for measuring
thymocyte apoptosis by propidium iodide staining and flow
cytometry. J. Immunol. Methods 139, 271–279.
34. Hempel, S.L., Buettner, G.R., O’Malley, Y.Q., Wessels, D.A. &
Flaherty, D.M. (1999) Dihydrofluorescein diacetate is superior for
detecting intracellular oxidants: comparison with 2¢,7¢-dichloro-
dihydrofluorescein diacetate, 5 (and 6)-carboxy-2¢,7¢-dichlorodi-
hydrofluorescein diacetate, and dihydrorhodamine 123. Free
Radic. Biol. Med. 27, 146–159.
35. Gasnier, F., Rousson, R., Lerme, F., Vaganay, E., Louisot, P. &
Gateau-Roesch, O. (1993) Use of Percoll gradients for isolation of
human placenta mitochondria suitable for investigating outer
membrane proteins. Anal. Biochem. 212, 173–178.
36. Hyslop, P.A. & Sklar, L.A. (1984) A quantitative fluorimetric
assay for the determination of oxidant production by poly-
morphonuclear leukocytes: its use in the simultaneous uorimetric
assay of cellular activation processes. Anal. Biochem. 141, 280–
37. Huot, J., Houle, F., Marceau, F. & Landry, J. (1997) Oxidative
stress-induced actin reorganization mediated by the p38 mitogen-
activated protein kinase/heat shock protein 27 pathway in vas-
cular endothelial cells. Circ. Res. 80, 383–392.
38. Brunk, U.T. & Svensson, I. (1999) Oxidative stress, growth factor
starvation and Fas activation may all cause apoptosis through
lysosomal leak. Redox Report 4, 3–11.
39. Yuan, X.M., Li, W., Brunk, U.T., Dalen, H., Chang, Y.H. &
Sevanian, A. (2000) Lysosomal destabilization during macrophage
damage induced by cholesterol oxidation products. Free Radic.
Biol. Med. 28, 208–218.
40. Neuzil, J., Zhao, M., Ostermann, G., Sticha, M., Gellert, N.,
Weber, C., Eaton, J.W. & Brunk, U.T. (2002) Alpha-tocopheryl
succinate, an agent with in vivo anti-tumour activity, induces
apoptosis by causing lysosomal instability. Biochem. J. 362,
41. Boveris, A. & Chance, B. (1973) The mitochondrial generation of
hydrogen peroxide. General properties and effect of hyperbaric
oxygen. Biochem. J. 134, 707–716.
42. Chance, B., Sies, H. & Boveris, A. (1979) Hydroperoxide meta-
bolism in mammalian organs. Physiol. Rev. 59, 527–605.
FEBS 2003 Lysosomes, apoptosis and mitochondria-mediated oxidative stress (Eur. J. Biochem. 270) 3785
43. Liu, Y., Fiskum, G. & Schubert, D. (2002) Generation of reactive
oxygen species by the mitochondrial electron transport chain.
J. Neurochem. 80, 780–787.
44. Hansford, R.G., Hogue, B.A. & Mildaziene, V. (1997) Depen-
dence of H
formation by rat heart mitochondria on substrate
availability and donor age. J. Bioenerg. Biomembr. 29, 89–95.
45. Scorrano, L., Penzo, D., Petronilli, V., Pagano, F. & Bernardi, P.
(2001) Arachidonic acid causes cell death through the mitochon-
drial permeability transition. Implications for tumor necrosis
factor-alpha aopototic signaling. J. Biol. Chem. 276, 12035–12040.
46. Hancock, J.T., Desikan, R. & Neill, S.J. (2001) Does the redox
status of cytochrome C act as a fail-safe mechanism in the regu-
lation of programmed cell death? Free Radic. Biol. Med. 31, 697–
47. de Duve, C., de Barsy, T., Poole, B., Trouet, A., Tulkens, P. & Van
Hoof, F. (1974) Commentary. Lysosomotropic agents. Biochem.
Pharmacol. 23, 2495–2531.
48. Koshkin,V.,Wang,X.,Scherer,P.E.,Chan,C.B.&Wheeler,
M.B. (2003) Mitochondrial functional state in clonal pancreatic
beta-cells exposed to free fatty acids. J. Biol. Chem. 278, 19709
49. Roberg,K.&O
llinger, K. (1998) Oxidative stress causes reloca-
tion of the lysosomal enzyme cathepsin D with ensuing apoptosis
in neonatal rat cardiomyocytes. Am. J. Pathol. 152, 1151–1156.
50. Roberg, K., Johansson, U. & O
llinger, K. (1999) Lysosomal
release of cathepsin D precedes relocation of cytochrome c and
loss of mitochondrial transmembrane potential during apoptosis
induced by oxidative stress. Free Radic. Biol. Med. 27, 1228–1237.
51. Roberg, K. (2001) Relocalization of cathepsin D and cytochrome
c early in apoptosis revealed by immunoelectron microscopy. Lab.
Invest. 81, 149–158.
52. Roberg, K., Kagedal, K. & O
llinger, K. (2002) Microinjection of
cathepsin D induces caspase-dependent apoptosis in fibroblasts.
Am. J. Pathol. 161, 89–96.
53. Stoka, V., Turk, B., Schendel, S.L., Kim, T.H., Cirman, T.,
Snipas, S.J., Ellerby, L.M., Bredesen, D., Freeze, H., Abraham-
V. & Salvesen, G.S. (2001) Lysosomal protease pathways to
apoptosis. Cleavage of bid, not pro-caspases, is the most likely
route. J. Biol. Chem. 276, 3149–3157.
54. Zhou, Q. & Salvesen, G.S. (1997) Activation of pro-caspase-7 by
serine proteases includes a non-canonical specificity. Biochem.
J. 324, 361–364.
55. Katunuma, N., Matsui, A., Kakegawa, T., Murata, E., Asao, T. &
Ohba, Y. (1999) Study of the functional share of lysosomal
cathepsins by the development of specific inhibitors. Adv. Enzyme
Regul. 39, 247–260.
56. Guicciardi, M.E., Deussing, J., Miyoshi, H., Bronk, S.F., Svingen,
P.A., Peters, C., Kaufmann, S.H. & Gores, G.J. (2000) Cathepsin
B contributes to TNF-alpha-mediated hepatocyte apoptosis by
promoting mitochondrial release of cytochrome c. J. Clin. Invest.
106, 1127–1137.
57. Cuvillier, O., Edsall, L. & Spiegel, S. (2000) Involvement of
sphingosine in mitochondria-dependent Fas-induced apoptosis of
type II Jurkat T cells. J. Biol. Chem. 275, 15691–15700.
58. Werneburg, N.W., Guicciardi, M.E., Bronk, S.F. & Gores, G.J.
(2002) Tumor necrosis factor-a-associated lysosomal permeabili-
zation is cathepsin B dependent. Am. J. Physiol. Gastrointest. Liver
Physiol. 283, 947–956.
59. Ka
gedal, K., Zhao, M., Svensson, I. & Brunk, U.T. (2001)
Sphingosine-induced apoptosis is dependent on lysosomal pro-
teases. Biochem. J. 359, 335–343.
3786 M. Zhao et al. (Eur. J. Biochem. 270) FEBS 2003
    • "Lysosomal membrane permeabilization per se triggers intracellular formation of ROS, a process which can be mediated by the action of lysosomal proteases, such as cathepsins B and D, which leak into the cytosol [208]. These enzymes affect mitochondria inducing further cytochrome c release and activation of caspase-mediated cell death [208, 228]. The involvement of lysosomes and their iron content in radiation-induced cell death is supported by the observation that cells are significantly protected from radiation damage if exposed to iron chelators [163, 165] . "
    [Show abstract] [Hide abstract] ABSTRACT: There is significant evidence that, in living systems, free radicals and other reactive oxygen and nitrogen species play a double role, because they can cause oxidative damage and tissue dysfunction and serve as molecular signals activating stress responses that are beneficial to the organism. Mitochondria have been thought to both play a major role in tissue oxidative damage and dysfunction and provide protection against excessive tissue dysfunction through several mechanisms, including stimulation of opening of permeability transition pores. Until recently, the functional significance of ROS sources different from mitochondria has received lesser attention. However, the most recent data, besides confirming the mitochondrial role in tissue oxidative stress and protection, show interplay between mitochondria and other ROS cellular sources, so that activation of one can lead to activation of other sources. Thus, it is currently accepted that in various conditions all cellular sources of ROS provide significant contribution to processes that oxidatively damage tissues and assure their survival, through mechanisms such as autophagy and apoptosis.
    Full-text · Article · Jul 2016
    • "The treated TIB-75 cells also experienced additional mitochondrial dysfunction in the form of increased mitochondrial ROS production. This dysfunction was likely triggered by the MMP which is known to stimulate ROS generation, which in turn can elicit more lysosomal damage [62]. This vicious cycle is likely to continue in the TIB-75 cells given their reduced ROS scavenging capacity. "
    [Show abstract] [Hide abstract] ABSTRACT: Background: Recent studies have shown that low density lipoproteins reconstituted with the natural omega 3 fatty acid docosahexaenoic acid (LDL-DHA) is selectively cytotoxic to liver cancer cells over normal hepatocytes. To date, little known about the subcellular events which transpire following LDL-DHA treatment. Methods: Herein, murine noncancer and cancer liver cells, TIB-73 and TIB-75 respectively, were investigated utilizing confocal microscopy, flow cytometry and viability assays to demonstrate differential actions of LDL-DHA nanoparticles in normal versus malignant cells. Results: Our studies first showed that basal levels of oxidative stress are significantly higher in the malignant TIB-75 cells compared to the normal TIB-73 cells. As such, upon entry of LDL-DHA into the malignant TIB-75 cells, DHA is rapidly oxidized precipitating global and lysosomal lipid peroxidation along with increased lysosomal permeability. This leakage of lysosomal contents and lipid peroxidation products trigger subsequent mitochondrial dysfunction and nuclear injury. The cascade of LDL-DHA mediated lipid peroxidation and organelle damage was partially reversed by the administration of the antioxidant, N-acetylcysteine, or the iron-chelator, deferoxamine. LDL-DHA treatment in the normal TIB-73 cells was well tolerated and did not elicit any cell or organelle injury. Conclusion: These studies have shown that LDL-DHA is selectively cytotoxic to liver cancer cells and that increased levels of ROS and iron catalyzed reactions promote the peroxidation DHA which lead to organelle dysfunction and ultimately the demise of the cancer cell. General significance: LDL-DHA selectively disrupts lysosomal, mitochondrial and nuclear function in cancer cells as a novel pathway for eliminating cancer cells.
    Article · Jul 2016
    • "Due to the fact that many of molecules under degradation within lysosomes contain iron in their composition, this organelle is particularly sensible to oxidative stress, especially those caused by H 2 O 2 . The lysosomal compartments, under oxidative stress, can in turn release hydrolytic enzymes with high degradation potential (proteases, lipases, nucleases, glycosidases, phospholipases, phosphatases and sulfatases), which can also act promoting mitochondrial ROS generation, constituting a cycle of oxidative damage amplification [75][76][77]. In addition, lysosomal proteases such as those belong to the cathepsins family are involved in apoptosis via different pathways, including the activation of caspases or the release of pro-apoptotic factors from the mitochondria [78][82, 83]. "
    [Show abstract] [Hide abstract] ABSTRACT: Several plant-derived compounds have been screened by antioxidant assays, but many of these results are questionable, since they do not evaluate the pharmacologic parameters. In fact, the development of better antioxidants stills a great challenge. In vitro cell-based assays have been employed to assess the antioxidant effect of various compounds at subcellular level. Cell-based assays can also reveal compounds able to enhance the antioxidant pathways, but without direct radical scavenging action (which could not be detected by traditional assays). These methodologies are general of easy implementation and reproducible making them suitable for the early stages of drug discovery. Hydrogen peroxide, a nonradical derivative of oxygen, can be employed as an oxidative agent in these assays due its biochemical properties (presence in all biological systems, solubility) and capacity of induction cell death. Truthfully, if their limitations are understood (such as difference on cell metabolism when in in vitro conditions), these cell-based assays can provide useful information about the pathways involved in the protective effects of phytochemicals against cell death induced by oxidative stress, which can be exploited to develop new therapeutic approaches.
    Full-text · Article · May 2016
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