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Blood, Vol. 93 No. 12 (June 15), 1999:
pp. 4096-4108
RAPID COMMUNICATION
Purified Photoproducts of Merocyanine 540 Trigger Cytochrome C Release
and Caspase 8-Dependent Apoptosis in Human Leukemia and Melanoma Cells
By
Shazib Pervaiz,
Mohamed A. Seyed,
Jayshreekumari L. Hirpara,
Marie-Véronique Clément, and
Kok W. Loh
From the Department of Physiology, National University of Singapore,
Singapore; and the Oncology Research Institute, NUMI, Singapore.
 |
ABSTRACT |
If the interplay between caspase proteases and mitochondria decide
the fate of the cell during apoptosis, they may constitute useful
molecular targets for novel drug design. We have shown that
photoactivated merocyanine 540 (pMC540) triggers caspase-mediated apoptosis in HL60 leukemia and M14 melanoma cells. Because pMC540 is a
mixture of photoproducts, we set out to purify the biologically active
component(s) from this mixture and to investigate their ability to
directly activate intracellular caspases and/or trigger mitochondrial
events associated with apoptosis. Two photoproducts, namely C1 and C2,
purified and characterized by mass spectroscopy and nuclear magnetic
resonance (NMR) analysis, effectively induced apoptosis in
HL60 and M14 cells. Interestingly, both C1 and C2 induced
non-receptor-dependent activation of caspase 8, which was responsible
for the downstream activation of caspase 3 and cell death. Both
compounds induced the release of cytochrome C from mitochondria of
tumor cells and from purified rat liver mitochondria; however,
different mechanisms were operative in cytochrome C translocation in
response to C1 or C2. C1-induced cytochrome C release was mediated by
the mitochondrial permeability transition (MPT) pore and accompanied by
a decrease in mitochondrial transmembrane potential
(  m), whereas cytochrome C release in response to C2
was independent of MPT pore opening. These findings do not exclude the
possibility that changes in mitochondrial m are
critical for apoptosis in some instances, but support the notion that
this may not be a universal step in the apoptotic process. Thus,
identification of two novel anticancer agents that directly activate
effector components of the apoptotic pathway could have potential
implications for the development of newer chemotherapeutic drugs.
© 1999 by The American Society of Hematology.
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INTRODUCTION |
AN ALTERNATIVE STRATEGY in cancer
management is to design treatments that directly activate effector
pathways involved in apoptosis. Thanks to a number of elegant studies,
the two most logical candidates to target would be the caspase family
of cysteine proteases and the mitochondria.1-19 The role of
caspases as orchestrators of apoptotic cell death has been well
established and elegantly reviewed by Thornberry and
Lazebnik.20 Each caspase is synthesized as an
inactive precursor molecule that which is activated in cells undergoing
apoptosis by proteolytic cleavage after specific aspartate residues.21 Once activated, caspases lead to proteolysis of a number of cellular substrates,22-25 culminating in
apoptotic collapse of the cell. Although caspase 3 (CPP32/apopain/Yama) is the most widely studied caspase in drug-induced
apoptosis,26-33 it is becoming clearer that there is a
molecular ordering of caspases in the apoptotic
program.6,34 Activation of an upstream initiator caspase,
such as caspase 8, leads to the subsequent induction of the effector
phase of apoptosis via processing of the executioner caspase, such as
caspase 3. Caspase 8 activation is classically observed upon triggering
of the CD95/CD95L system via induction of the death-inducing signaling
complex35; however, two recent reports demonstrate that
caspase 8 activation can also be induced by anticancer drugs in the
absence of cell surface death receptor-ligand interaction.4,6 Whereas receptor-dependent caspase 8 activation does not essentially require mitochondrial controlled
processes, drug-induced caspase 8 activation is completely abolished in
the absence of mitochondria,4 suggesting involvement of
mitochondrial-derived factors.
Early disruption of mitochondrial function is seen in most cases of
drug-induced apoptosis, and the various mechanisms that mediate
mitochondrial dysfunction involve changes in cellular redox state via
enhanced generation of reactive oxygen species (ROS) or a decrease in
their detoxification, depletion of NADPH, or depletion of
ATP.36-38 It is also well established that, after an
apoptotic stimulus, mitochondria release apoptogenic factors such as
apoptosis-inducing factor (AIF), cytochrome C (Cyt.C), and even
caspases.1,5,13,18,19,39 The translocation of these factors
to the cytosol together with additional factors, namely ATP and Apaf-3,
can proteolytically activate caspase 3,7 which, in turn,
activates cellular disassembly. Mitochondrial Cyt.C normally resides in
the intermembrane space, and its release into the cytosol during
apoptosis indicates an enhanced permeability of the outer mitochondrial
membrane. One of the mechanisms proposed for the translocation of Cyt.C
to the cytosol is a reduction in the mitochondrial transmembrane
potential ( m) that accompanies some forms of
apoptosis.40-43 This reduction in m is
thought to be mediated by the opening of the mitochondrial permeability transition (MPT) pore, a dynamic multiprotein complex located at the
contact site between the inner and the outer mitochondrial membranes.37,44-49 Opening of the MPT pore has been
implicated in cell death induced by ROS, hepatotoxins, calcium, and
anoxia, and inhibitors of MPT pore opening, such as cyclosporin A (CsA) and bongkrekic acid, have been shown to inhibit all signs of
apoptosis.37,41,44,50-52 Recent observations point to an
intricate cross-talk between caspases and mitochondria in the apoptotic
process. Several recombinant caspases (caspases 1, 2, 3, 4, and 6)
enhance the permeability of PT pore-containing liposomes, and inducers
of MPT pore opening, such as atractyloside, trigger the release of
caspases 2 and 9 from the mitochondria,13,44 which can
activate downstream caspases such as caspase 3, 6, and 7. Activated
caspases, in turn, can directly act on the mitochondria, thus engaging
in a self-amplifying feedback loop in which changes in mitochondrial
permeability lead to caspase activation and vice versa. Thus, if the
interplay between caspase proteases and mitochondria decides the fate
of the cell during apoptosis, they may constitute useful molecular
targets for novel drug design. In this regard, identifying anticancer agents with a predilection for mitochondrial membrane structures, including the MPT pore complex on the one hand and ability to directly
activate intracellular caspases on the other, would be an ideal
combination to induce effective apoptosis in tumor cells. By targeting
the effectors (caspases and mitochondria) of the apoptotic pathway, the
need for private signaling mechanisms could be bypassed, which could
significantly improve the response of tumor cells to chemotherapy.
We have previously shown that photoactivation of lipophilic agent
merocyanine 540 generates a mixture of photoproducts (pMC540) that
selectively induce cell death in human leukemia, lymphoma, and a
variety of other tumor cell types in vitro and in
vivo.53-55 In a recent communication, we investigated the
mode of cell death triggered by pMC540 and showed that HL60 leukemia
and M14 melanoma cells underwent caspase-mediated cell death with all
the characteristic hallmarks of apoptosis.33 In the present
study, we isolated and purified three biologically active components
from the pMC540 mixture. Two of the purified compounds (C1 and C2)
triggered efficient activation of both an initiator caspase, caspase 8, and an executioner caspase, caspase 3, and induced caspase 8-dependent
apoptosis in HL60 leukemia and M14 melanoma cells. Both compounds were
able to induce Cyt.C release from mitochondria of tumor cells and from purified rat liver mitochondria; however, Cyt.C release in response to
C2 occurred independent of MPT pore opening and without a decrease in
mitochondrial m.
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MATERIALS AND METHODS |
Tumor cell lines.
The human promyelocytic leukemia cell line HL60 was obtained from ATCC
(Rockville, MD) and maintained in culture in RPMI 1640 supplemented
with 10% fetal bovine serum (FBS; GIBCO-BRL, Gaithersburg, MD). M14
human melanoma cell line was a generous gift from Dr Armando Bartolazzi
(Oncologia Clinica e Sperimentale, Rome, Italy) and cultured in
RPMI/5% FBS.
Purification of photoproducts from pMC540 mixture.
Photoactivation of MC540 was performed as described
previously.33 Briefly, 500 mg of MC540 at 1 mg/mL in 70%
aqueous ethanol was photoactivated by exposure to a bank of fluorescent
lamps (GE Cool White, 40W; General Electric, Cleveland,
OH) for 18 hours. After photoactivation, ethanol was removed by freeze
drying, and the mixture of photooxidation product was analyzed by
thin-layer chromatography (TLC) on aluminum sheets coated with
fluorescent silica gel 60F254 (Merck, Darmstadt,
Germany) by using the following solvent systems: (1) ethyl
acetate/hexane (8:2) and (2) CHCl3/methanol (6:4). UV/VIS
absorption spectra were obtained using a spectrophotometer (Biospec
1601, Shimadzu, Japan). Each fraction was then subjected to mass
spectrometry (MS) and proton (H1)- and carbon
(C13)-nuclear magnetic resonance (NMR)
(Bioscience Centre, National University of Singapore, Singapore).
Collected fractions were then resuspended in dimethyl sulfoxide (DMSO)
at 100 mg/mL and stored at  80°C protected from light.
Determination of caspases 8 and 3 activities.
Activation of intracellular caspase 8 or caspase 3 was assayed by the
ApoAlert Fluorescent Assay Kits (Clontech Lab Inc, Palo Alto, CA). HL60
and M14 cells (1 × 106 cells/mL) were
exposed to C1, C2, or C5 (50 to 150 µg/mL), washed twice with
1× phosphate-buffered saline (PBS), resuspended in 50 µL of
chilled cell lysis buffer (provided by the supplier), and incubated on
ice for 10 minutes. Fifty microliters of 2× reaction buffer
containing 10 mmol/L dithiothreitol (DTT) and 6 µL of
the substrate IETD-AFC (caspase 8) or DEVD-AFC (caspase 3) were added to each sample and incubated at 37°C for 30 minutes. Protease activity was determined by measuring the relative fluorescence intensity at 505 nm after excitation at 400 nm using a
spectrofluorimeter (Luminescence Spectrometer LS50B; Perkin Elmer,
Buckinghamshire, UK). Results are shown as the fold increase in
activity relative to untreated control cells.
Annexin V staining for detection of apoptotic cells.
Externalization of phosphatidylserine, an early marker of apoptosis,was
assessed by the ApoAlert-Annexin V Apoptosis Kit (Clontech Lab Inc).
Briefly, HL60 and M14 cells (1 × 106/mL) were exposed
to 100 µg/mL of C1 or C2 or C5 for 12 hours. Cells were then washed
twice with PBS + 1% FBS and resuspended in 200 µL of 1×
binding buffer (supplied by the vendor). Annexin V-fluorescein
isothiocyanate (FITC) (1 µg/mL) was then added to the cells and left
at room temperature for 15 minutes in the dark. After two washes with
PBS, cells were resuspended in 0.5 mL of PBS + 1% FBS and analyzed by
flow cytometry with the excitation wavelength at 488 nm and the
emission set at 525 nm (green). Flow cytometry data were analyzed by
the WINMDI software (University of Massachusetts, Amherst, MA).
Detection of cytosolic Cyt.C.
For determination of cytosolic Cyt.C, HL60 cells (30 × 106) were treated with the 100 µg/mL of C1 or C2 for 18 hours and cytosolic fractions were obtained. Briefly, cells were washed
twice with ice-cold PBS, pH 7.4, followed by centrifugation at
200g for 5 minutes. The cell pellet was then resuspended in 600 µL of extraction buffer, containing 200 mmol/L mannitol, 68 mmol/L
sucrose, 50 mmol/L PIPES-KOH, pH 7.4, 50 mmol/L KCl, 5 mmol/L EGTA,
2 mmol/L MgCl2, 1 mmol/L DTT, and protease
inhibitors (Complete Cocktail; Boehringer Mannheim, Mannheim,
Germany). After 30 minutes of incubation on ice, cells
were homogenized with a dounce homogenizer, the homogenate was spun at
14,000g for 15 minutes, and supernatants were removed and
stored at 80°C until analysis by gel electrophoresis for
Cyt.C. In a separate set of experiments, M14 cells (2 × 103) were grown on cover slips, exposed to 100 µg/mL of
C1 or C2, fixed with methanol:acetone (1:1 vol/vol), and incubated for
2 hours at 37°C with 1:150 dilution (in 3% bovine serum albumin [BSA]) of monoclonal anti-Cyt.C antibody (clone 7H8.2C12; Pharmingen, San Diego, CA). After three washes with 1× PBS + 1% FBS, cells were exposed to a 1:20 dilution of antimouse FITC-conjugated IgG (Pharmingen) for 1 hour, washed twice, and analyzed by confocal microscopy (NUMI Core Facility, NUS, Singapore). Cytosolic Cyt.C was
defined as diffuse cytoplasmic staining as compared with punctate mitochondrial Cyt.C staining obtained with nontreated control cells.
Preparation of rat liver mitochondria.
Mitochondria were isolated from rat liver (Albino rats, Wistar strain),
as described elsewhere.56 Briefly, liver cells were homogenized in 10 mL of buffer A (0.3 mol/L sucrose, 5 mmol/L TES, 0.2 mmol/L EGTA, pH 7.2, with KOH) and centrifuged at 2,000g for 10 minutes at 4°C. The supernatant (S1) was removed and the pellet was
resuspended in 10 mL of buffer A and centrifuged at 2,000g for
10 minutes at 4°C. The supernatant obtained (S2) was then mixed
with S1 and centrifuged at 8,000g for 10 minutes at 4°C.
The pellet was then resuspended in 1 mL of buffer A, loaded on top of a
percoll gradient (60%, 30%, 18%) prepared in buffer A, and
centrifuged at 8,000g for 10 minutes at 4°C. Mitochondria were then separated from nonmitochondrial membranes and nonfunctional organelles, collected at the 30%/60% interface, and washed with 10 vol of buffer A at 8,000g for 10 minutes at 4°C to wash off the percoll. Mitochondria were then resuspended in 2 mL of buffer A and
kept at 4°C with gentle stirring. All experiments with isolated mitochondria were performed within 4 hours of the preparation.
Mitochondrial swelling and release of Cyt.C.
Large amplitude mitochondrial swelling was determined spectroscopically
by the loss of absorbance at 540 nm, as described elsewhere.48 Diethyl pyrocarbonate (DEPC; 200 µmol/L;
Sigma, St Louis, MO) was used as a positive control to induce
mitochondrial swelling.46 Where indicated, 10 µmol/L CsA
(Sigma) was added to the mitochondria before the addition of DEPC, C1,
or C2,.
In a separate set of experiments, 0.5 mg of mitochondria was incubated
with 100 µg of C1 or C2 for 1 hour at 30°C in the presence and
absence of 10 µmol/L CsA, followed by centrifugation at
4,000g for 5 minutes. Control samples received an equal volume
of the carrier solvent. The resulting supernatants were used either for caspase activity assays or analyzed by Western blot analysis to detect
Cyt.C, as described below.
Determination of mitochondrial m by flow
cytometry.
Potential-sensitive probe 3, 3' dihexyloxacarbocyanine iodide
(DiOC6) was used to measure mitochondrial
m, as described elsewhere.44 Fifty
micrograms purified rat liver mitochondria was exposed to 100 µg/mL
of C1 or C2 for 1 hour at 25°C, followed by incubation for 15 minutes at 37°C with 40 nmol/L DiOC6. After two gentle
washes with 1× PBS, mitochondria were analyzed in an Epics
Profile (Coulter, Hialeah, FL) flow cytometer with the
excitation set at 488 nm. At least 10,000 events were collected per
sample and data were analyzed by the WINMDI software.
Western blotting for Cyt.C.
Fifty micrograms of cytosolic protein extracts or supernatants from
mitochondria treated with C1 or C2 was subjected to 12% sodium dodecyl
sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and transferred
to polyvinylidene difluoride (PVDF) membranes using a
Trans-blot SD semidry system (Bio-Rad Laboratories, Hercules, CA). Membranes were then blocked overnight with 5% dry
milk in TBST (50 mmol/L Tris/HCl, pH 7.4, 150 mmol/L NaCl, 0.1% Tween 20). Exposure of blocked membranes to primary anti-Cyt.C antibody (7H8.2C12) was accomplished at room temperature for 1 hour using an
antibody dilution of 1:5,000 made in TBS + 0.05% Tween 20 and 1% BSA.
After three washes with TBST, the membranes were exposed to 1:5,000
dilution of goat antimouse IgG-horseradish peroxidase (HRP) conjugate supplied as 0.8 mg/mL (Pierce, Rockford,
IL) for 1 hour and washed three times with TBST. Chemiluminescence was detected using the SuperSignal Substrate Western Blotting Kit (Pierce).
Cytotoxicity assays.
To determine the sensitivity of tumor cell lines to C1, C2, and C5,
HL60 and M14 cells were treated with increasing concentrations of the
compounds (25 to 100 µg/mL) for 18 hours. HL60 cells were plated at 1 × 105/well in a 96-well plate and viability was
determined by the MTT assay. Ten microliters of 5 mg/mL
3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide (MTT)
was added to each well and incubated for 4 hours at 37°C. Elution
of the precipitate was performed with 100 µL of DMSO and 10 µL of
Tris-Glycine buffer. Cell viability was calculated from the absorption
values obtained at 570 nm using an automated enzyme-linked
immunosorbent assay (ELISA) reader. M14 cells were plated at 2 × 104/well in RPMI/5% FBS and allowed to attach overnight.
Medium was then removed and replaced with fresh medium containing 25 to
100 µg/mL of C1, C2, or C5. After 18 hours of incubation, wells were aspirated and 50 µL of 0.75% crystal violet solution containing 50%
ethanol, 0.25% NaCl, and 1.75% formaldehyde was added to each well
for 10 minutes. Cells were then washed with water and air-dried, and
the dye was eluted with PBS/1% SDS. Cell viability was measured by dye
absorbance at 590 nm on an automated ELISA reader.
| |
RESULTS AND DISCUSSION |
Purification of three active compounds from pMC540.
In a previous study, we showed that tumor cells undergo
caspase-mediated cell death upon exposure to a mixture of photoproducts generated upon photoactivation of MC540.33 Because pMC540
is a mixture of photoproducts and for it to have potential as a
clinical chemotherapeutic agent, the biologically active component(s)
in the mixture have to be identified. We report here the purification and characterization of three biologically active components from the
photoactivated mixture. The conditions used for photoactivation of
MC540 were identical to those described in our in vitro studies demonstrating caspase-dependent antitumor activity of
pMC540.33 After photoactivation in ethanol, the solvent was
evaporated and the mixture of photo-oxidation product was analyzed by
TLC on aluminum sheets coated with fluorescent silica gel. Three major fractions were identified. TLC solvent system 1 yielded two products of
Rf 0.83 and 0.65 termed here as C1 and C2, respectively,
and solvent system 2 yielded a third product at Rf 0.25 termed C5. Further analysis of the purified fractions by MS and
H1- and C13-NMR showed that C1 and C2 were pure
compounds with the structural formulae N,
N'-Dibutyl-thio-4,5-imidazolindion (molecular weight [MW] = 242) and N,
N'-Dibutyl-4,5-dihydro-5-hydroxy-5-ethoxy-4-oxo-2-thiouracil (MW = 316), respectively (Fig 1). These two
structures are similar to those previously described upon
photo-oxidation of MC540 in methanol and are known as merodantoin and
merocil.57 The third compound, named C5, was identified as
a highly photosensitive intermediate product with the proposed
structure shown in Fig 1. As determined by extraction after analysis on
silica-coated TLC plates, the majority of the pMC540 was composed of C5
(~60%), whereas the percentage yields for C1 and C2 were
approximately 16% each.
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| Fig 1.
Structural formulae for the three compounds determined by
MS and H1- and C13-NMR analysis of purified
fractions. C1, N, N'-Dibutyl-thio-4,5-imidazolindion (MW = 242); C2, N,
N'-Dibutyl-4,5-dihydro-5-hydroxy-5-ethoxy-4-oxo-2-thiouracil (MW
= 316); C5 was identified as a highly photosensitive intermediate
product with the proposed structure shown above.
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Having identified the major components of pMC540, we set out to
determine if the antitumor activity of pMC540 could be attributed to
one or more of these photoproducts. Because of our earlier findings
with pMC540, we were specifically interested in identifying the
compound(s) that could trigger apoptotic cell death in tumor cells by
directly activating intracellular caspases and/or trigger mitochondrial
events linked to the apoptotic process. Human promyelocytic leukemia
cell line HL60 was selected as a model, because our previous studies
had shown that human leukemia and lymphoma cells were specifically
sensitive to apoptosis induced by pMC540.53,55 In addition,
where indicated, similar experiments were performed with M14 human
melanoma cells used as a model solid tumor cell line. Although all
three compounds (C1, C2, and C5) were able to induce cell death in both
tumor cell lines after 18 hours of treatment, C1 and C2 had
significantly better activity than C5 (Fig
2A and B). Moreover, phenotypic analysis of HL60 cells after exposure
to 100 µg/mL of C1 or C2 for 12 hours showed that majority of the
cells (>70%) exhibited externalization of inner membrane phosphatidylserine (annexin V positive; Fig 2C), a characteristic morphological change associated with apoptotic cell
death.58 Similar to the results obtained with the cell
death assays, this morphological change was observed in significantly
lesser number (<30%) of C5-treated leukemia cells. These data show
that purified compounds C1 and C2 can efficiently kill tumor cells by
triggering the apoptotic pathway, as confirmed by PS externalization.
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| Fig 2.
Antitumor activity of C1, C2, and C5 against (A) HL60 and
(B) M14 cell lines. A total of 1 × 106 cells/mL were
exposed to increasing concentrations (25 to 100 µg/mL) of each
compound for 18 hours and cell death was determined by the MTT assay
(HL60) or crystal violet assay (M14) as described in Materials and
Methods. Data shown are the mean ± SD of three independent
experiments performed in triplicate. (C) To assess apoptotic phenotype,
HL60 cells (1 × 106) were exposed to 100 µg/mL of C1,
C2, or C5 for 12 hours and externalization of inner membrane PS was
detected by annexin V-FITC (1 µg/mL) staining and analysis by flow
cytometry. At least 10,000 events were analyzed by WINMDI software.
Arrows indicate log increase in fluorescence over untreated control
cells.
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C1 and C2 induce efficient activation of caspase 8 and caspase 3.
The critical role of intracellular caspases as mediators of all
apoptotic cell death has been well established and elegantly reviewed.20 Caspases are not only involved in cell death
triggered via cell surface death receptors, but their role in
drug-induced apoptosis has also been recently
highlighted.29-31,33,59 Whereas caspase 3 is the most
extensively studied and the central mediator common to a host of
apoptotic triggers, other upstream members of the caspase family, vis a
vis caspase 8 (FLICE), have also been shown to be critical for
effective apoptosis.4,15,60,61 In our earlier studies, we
had shown that tumor cells exposed to pMC540 exhibited efficient
activation of caspase 3 and that caspase 3 inhibition was able to block
pMC540-mediated apoptosis.33 Stimulated by these findings
and by our observation that the three purified components from pMC540
induced apoptosis in tumor cells, we next assessed the ability of the
three compounds to induce activation of intracellular caspase
proteases. Therefore, HL60 cells were exposed to 50 to 150 µg/mL of
C1, C2, or C5 for 12 hours and caspases 8 and 3 activities were
determined by fluorimetric assays using tetrapeptide substrates
IETD-AFC and DEVD-AFC, respectively. Our results showed that, within 12 hours of treatment, all three compounds (C1, C2, and C5) activated
caspases 8 and 3 in a dose-dependent manner, with the maximum activity
already obtained with 100 µg/mL for C1 and C2
(Fig 3A). However, C2 was the most
efficient (6.2× increase in activity), followed by C1 (5.1×
increase in activity) and C5 (2× increase in activity), as shown
in Fig 3A. Moreover, a kinetic analysis of caspase 8 activation upon
exposure to 100 µg/mL of C1, C2, or C5 for 4 to 18 hours showed that
C2 was the earliest (8 hours after treatment) and the most potent
(>5× increase in activity at 12 hours) activator of caspase 8 in leukemia cells (Fig 3B). C1 was also efficient in activating caspase
8; however, the peak activity (4× increase) was reached
significantly later (18 hours) than with C2 (Fig 3B). On the contrary,
C5 did not activate caspase 8 as efficiently as C1 or C2, with the
maximum activity (2× over the untreated cells) reached at 18 hours after drug treatment (Fig 3B). Similarly, caspase 3 activation
was induced by both C1 and C2, whereas C5-treated cells showed minimal
activation by 12 hours that did not significantly increase even after
18 hours of treatment (Fig 3C). Again, the activation kinetics for caspase 3 indicated that C2 was the most potent caspase 3 inducer, with
the peak activity (5.1× increase) at 18 hours after drug treatment, followed by C1 (3.5× increase), whereas C5 did not induce significant activation of caspase 3, as shown in Fig 3D. Similar
inductions and kinetics of caspases 8 and 3 activities were obtained
upon incubation of M14 cells with C1, C2, or C5 (data not shown). In
both cell lines, the kinetics of caspase 8 activation always preceded
caspase 3 activation, and the ability of each compound to activate
caspase proteases, particularly caspase 3, always correlated with the
efficiency of HL60 and M14 cell death induced by each purified
compound. Because we were interested in the compound(s) that
efficiently activated caspase proteases and effectively triggered
apoptosis, we subsequently focused our investigations on the mechanism
of action of compounds C1 and C2 only.
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| Fig 3.
Determination of (A) caspase 8 or (C) caspase 3 activation by a fluorimetric assay designed to detect cleavage of
tetrapeptide substrates. HL60 cells (1 × 106) were
treated with 50 to 150 µg/mL of C1, C2, or C5 for 12 hours and
lysates were analyzed for IETDase (caspase 8) or DEVDase (caspase 3)
activities as described in Materials and Methods. Data shown are the
mean ± SD of four independent experiments and are expressed as X
increase in activity over the untreated HL60 cells. Kinetics of (B)
caspase 8 or (D) caspase 3 activation in HL60 cells exposed to 100 µg/mL of C1 or C2 or 150 µg/mL of C5 for 4 to 18 hours. Data shown
are representative of three independent observations.
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Apoptosis and caspase 3 activation induced by C1 and C2 are caspase
8-dependent.
We were intrigued by our observation that two of the three compounds,
C1 and C2, efficiently activated caspase 8, an early protease in the
caspase cascade that has been better described in cell death induced
via the cell surface receptor CD95 (Fas/Apo1).61 Therefore,
we first addressed the questions of whether caspase 8 was critical in
the process of apoptosis induced by C1 and C2 and if activation of
caspase 3 was dependent on caspase 8 activation or if both caspases
were independently activated by C1 and/or C2. Using an aldehyde
tetrapeptide inhibitor of caspase 8, IETD-CHO (50 µmol/L), we showed
a significantly enhanced survival of HL60 cells after exposure to 100 µg/mL of C1 (from 30% to 70%) or 50 µg/mL of C2 (from 50% to
87%), as shown in Fig 4A. Furthermore, this enhanced survival was due to inhibition of apoptosis induced by C1
or C2, as shown by a complete inhibition of annexin V staining in the
presence of IETD-CHO as opposed to C1 (100 µg/mL) or C2 (50 µg/mL)
alone (Fig 4B). Furthermore, preincubation for 1 hour with IETD-CHO (50 µmol/L) completely inhibited the activation of caspase 3 induced by
12 hours of treatment with either C1 (100 µg/mL) or C2 (100 µg/mL),
as shown in Fig 4C. These data demonstrate that caspase 3 activation
and cell death induced upon exposure of tumor cells to C1 or C2 were
dependent on upstream activation of caspase 8. Whereas activation of
downstream executioner caspase 3 has been well documented with a
variety of anticancer drugs, the involvement of non-receptor-dependent
activation of caspase 8 in drug-induced tumor cell death is a somewhat
novel finding. There are two recent reports showing activation of
caspase 8 in response to commonly used chemotherapeutic drugs, such as
etoposide and doxorubicin, and another using apoptosis inducing agent
betulinic acid.4,6,62 Drug-induced caspase 8 activation
could be attributed to upregulation of the cell surface CD95/CD95L
system. In our present study, caspase 8 activation in response to C1 or
C2 was not secondary to induction of CD95-CD95L interaction in HL60
cells. C1 and C2 did not upregulate CD95 or CD95L and the antitumor
activity of C1 or C2 was not inhibited in the presence of anti-CD95 or anti-CD95L antibodies (data not shown). To further
corroborate these findings obtained in HL60 cells, we performed the
same series of experiments with C1 and C2 on M14 melanoma cells. M14
cells have been shown to lack both of the death-inducing receptors, CD95 and TNFR.63 Our results showed that, similar to HL60
cells, both C1 and C2 triggered caspase 8-dependent cell death in
CD95-deficient M14 cells (data not shown), thereby ruling out the
possibility that caspase 8 activation could be secondary to CD95
receptor aggregation. Thus, the activation of caspase 8 induced by C1
and C2 was due to a direct effect of the drugs and not via mechanisms involving cell surface death receptors. Our findings on caspase 8-dependent activation of caspase 3 by C1 and C2 are in agreement with
earlier reports that mature caspase 8 could by itself activate downstream effector caspases, including caspase 3.64 In the light of these findings, we contend that neither C1 nor C2 directly activate caspase 3, but the sequence of events involve activation of an
early caspase, caspase 8, that either directly or indirectly triggers
activation of the executioner caspase 3.
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| Fig 4.
Blocking caspase 8 activation enhances HL60 cell survival
by inhibiting downstream caspase 3 activation and cell death. (A) HL60
cells (1 × 106/mL) were incubated with C1 or C2 (100 µg/mL) in the presence or absence of IETD-CHO (50 µmol/L) for 18 hours and cell survival was determined by MTT assay. Data shown are the
mean ± SD of three independent experiments. (B) C1-and C2-treated
HL60 cells were assessed for apoptotic phenotype in the presence of
IETD-CHO (50 µmol/L) by annexin V-FITC staining by flow cytometry as
described in Materials and Methods. Data are shown as the percentage of
annexin V positive. (C) Cell lysates obtained from HL60 cells (1 × 106) after exposure to 100 µg/mL of C1 or C2 for 12 hours
in the presence or absence of IETD-CHO (50 µmol/L) were analyzed for
caspase 3 activation (DEVDase activity). Data are shown as X increase
in caspase 3 activity.
|
|
C1 and C2 induce translocation of mitochondrial Cyt.C.
Thus far, we have shown that compounds C1 and C2 trigger caspase
8-dependent apoptosis in HL60 and M14 cells; however, the intracellular
events linking activation of caspase 8 by C1 and C2 to downstream
caspase 3 activation and death remain to be elucidated. Caspase 8 has
been shown to activate downstream caspases, such as caspase 3, via
pathways that are either mitochondrial-dependent or
independent.4 However, engagement of the
mitochondrial-dependent pathway is more efficient than the
mitochondrial-independent pathway, because it can be activated by small
amounts of caspase 8.15 The mitochondrial-dependent pathway
relies on translocation of mitochondrial factors, such as AIF and
Cyt.C, which occurs during the early phase of apoptosis induced by a
variety of cell death triggers.1,19,65,66 Cyt.C, in concert
with other cytosolic factors, then causes the activation of executioner
caspases similar to caspase 3,19,65,67 leading to
downstream apoptotic events. Having shown that C1 and C2 can
efficiently activate caspase 8, we asked if C1 and/or C2 could trigger
translocation of mitochondrial Cyt.C to the cytosol of tumor cells.
Cytosolic release of Cyt.C was analyzed after exposure of HL60 leukemia
cells and M14 melanoma cells with 100 µg/mL of C1 or C2 for 12 hours.
Western blot analysis or confocal microscopy was used to detect
cytosolic Cyt.C in HL60 and M14 cells, respectively. Western blot
analysis of Cyt.C in HL60 cytosolic fractions showed that both C1 and
C2 induced translocation of Cyt.C to the cytosol (15-kD band), whereas
control cytosols from HL60 cells treated with the carrier solvent did
not contain Cyt.C (Fig 5A). Similarly,
analysis of M14 cells by confocal microscopy showed a punctate pattern
of staining for Cyt.C in untreated control cells, showing mitochondrial
localization; however, cells exposed to C1 and C2 for 12 hours
exhibited bright green diffuse cytoplasmic staining, indicating
translocation of Cyt.C to the cytosol (Fig 5B). Thus, C1 and C2 provoke
the cytosolic translocation of Cyt.C, which suggests that both
anticancer agents directly or indirectly induce mitochondrial events
associated with apoptotic cell death. We propose two possible
scenarios, one in which upstream caspase 8 activation induced by both
C1 and C2 acts on the mitochondria and induces Cyt.C release and a
second in which Cyt.C release could be a result of direct targeting of
the mitochondrial membrane structures by C1 and/or C2. We made an
attempt to dissect Cyt.C release from caspase 8 activation to decipher
the sequence of apoptotic events triggered by C1 and C2. M14 cells were
preincubated with caspase 8 inhibitor (IETD-CHO; 100 µmol/L) before
the addition of 100 µg/mL of C1 or C2 and analyzed by confocal
microscopy for cytosolic Cyt.C. Results showed that preincubation of
cells with IETD-CHO did not inhibit the cytosolic translocation of
Cyt.C (data not shown), suggesting a possible sequence of events in which C1 and C2 first trigger the release of mitochondrial Cyt.C, which
activates caspase 8, leading to downstream activation of executioner
caspase 3 and cell death.
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| Fig 5.
C1 and C2 trigger translocation of mitochondrial Cyt.C in
HL60 and M14 cells. (A) HL60 (30 × 106 cells) were
treated with 100 µg/mL of C1 or C2 for 12 hours and cytosolic
fractions were subjected to SDS-PAGE electrophoresis, transferred to
PVDF membrane, and subjected to Western blot analysis for Cyt.C as
described in Materials and Methods. Anti- -actin antibody was used
to assess equal loading of samples. (B) M14 cells (1 × 103) were grown on cover slips and treated with 100 µg/mL
of C1 or C2 for 12 hours, and Cyt.C localization was determined by
confocal microscopy using anti-Cyt.C as described in Materials and
Methods.
|
|
C1 reduces m and induces
MPT-dependent mitochondrial swelling, whereas C2 functions independent
of a reduction in
m.
Two major hypotheses have been experimentally suggested for the
mechanism of Cyt.C release from the mitochondria during apoptosis: one
that is dependent on a decrease in m secondary to
opening of the inner membrane MPT pore2 and the other that
contends that Cyt.C release is independent of a decrease in
m1. MPT pore opening results in volume
dysregulation of the mitochondria due to the hyperosmolality of the
matrix, causing the matrix space to expand. Inhibitors of MPT pore
opening, such as CsA and bongkrekic acid, appear to block apoptosis in
some systems,40,50,68 implying a general role. However,
some studies have provided evidence that Cyt.C release and caspase
activation can occur independently of any detectable loss in
mitochondrial m.1 Stimulated by our findings that both C1 and C2 triggered the release of mitochondrial Cyt.C in HL60 and M14 cells, we set out to investigate if this cytosolic translocation of Cyt.C was due to direct effect of the drugs
on mitochondrial membrane structures, specifically the MPT pore. To
accomplish that, we purified rat liver mitochondria and asked if C1
and/or C2 caused mitochondrial swelling secondary to MPT pore opening
and if Cyt.C release was dependent on reduction of mitochondrial
m. Freshly isolated mitochondria (0.5 mg) were exposed to 100 µg of C1 or C2 under conditions previously shown to be
conducive for swelling induced by activators of MPT, such as
Ca2+ and DEPC.46 As expected, the addition of
DEPC (0.2 mmol/L) to isolated mitochondria induced rapid swelling that
was measured by a decrease in absorbance at 540 nm using a double beam
spectrophotometer (Fig 6A). This swelling
was completely inhibited by prior addition of CsA (10 µmol/L), an MPT
pore inhibitor. Similar to DEPC, the addition of C1 to fresh
mitochondria resulted in a loss of absorbance (0.3 OD units) that was
completely inhibited by prior addition of CsA (Fig 6B). Unlike C1, C2
had no effect on mitochondrial swelling, as shown in Fig 6C. To further
provide impetus to these findings, we directly assessed the ability of
C1 and C2 to induce Cyt.C release from isolated mitochondria and to
determine if it was dependent on opening of the MPT pore and a decrease
in mitochondrial m. Purified mitochondria (0.5 mg)
were incubated with 100 µg/mL of C1 or C2 in the presence or absence
of MPT pore inhibitor CsA (10 µmol/L). Mitochondria were then gently
pelleted and supernatants were used for Western blot analysis of Cyt.C.
Our results showed that both C1 and C2 triggered the release of Cyt.C
from purified mitochondria; however, the presence of CsA was able to
inhibit Cyt.C release only from mitochondria treated with C1
(Fig 7A). Moreover, measurement of
mitochondrial m with DiOC6 after
incubation for 30 minutes with C1 or C2 showed that C1 induced a
decrease in mitochondrial m that was completely
inhibited by CsA, whereas C2 had no effect on m, as
shown in Fig 7B. These results suggest two different modes of Cyt.C
release for C1 and C2, whereby C1 directly targets the MPT pore and
induces mitochondrial swelling in a CsA inhibitable manner and
contrarily C2 triggers the release of Cyt.C from the mitochondria
independently of MPT pore opening.
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| Fig 6.
C1 induces mitochondrial swelling via induction of the
MPT pore, whereas C2 has no effect on the pore. Large amplitude
mitochondrial swelling was determined spectroscopically by monitoring
the loss of absorbance at 540 nm as described in Materials and Methods.
Mitochondria (0.5 mg) were treated with (A) 200 µmol/L DEPC as a
positive control to induce mitochondrial swelling or (B) C1 (100 µg) or (C) C2 (100 µg) in the absence or presence of
MPT pore inhibitor CsA (10 µmol/L).
|
|
View larger version (36K):
[in this window]
[in a new window]
| Fig 7.
C1-triggered Cyt.C release and decrease in mitochondrial
m is dependent on opening of the MPT pore. (A)
Purified rat liver mitochondria (0.5 mg) were exposed to 100 µg/mL of
C1 or C2 in the presence or absence of CsA (10 µmol/L) for 30 minutes
at 30°C. Mitochondria were then pelleted and supernatants were
subjected to SDS-PAGE electrophoresis and Western blot analysis for
Cyt.C as described in Materials and Methods. (B) Mitochondria were
treated with C1 and C2 in the presence or absence of CsA as above and
stained with membrane potential-sensitive dye DiOC6 (40 nmol/L) at 37°C for 30 minutes, washed, and analyzed by flow
cytometry for m.
|
|
The release of Cyt.C without a decrease in m seen
with C2 suggests that different events control permeability of the
inner and outer mitochondrial membranes. A possible mechanism for outer membrane disruption has recently been suggested that involves hyperpolarization,69 rather than hypopolarization of the
inner membrane usually associated with pore opening. It is also
possible that the release of Cyt.C triggered by C2 may involve
disruption of the outer mitochondrial membrane by a hitherto unknown
mechanism. Could it be a function of activated caspases directly
targeting the mitochondria leading to translocation of Cyt.C or better
yet a result of activation of caspases present within the mitochondria? In this regard, recent observations have shown that recombinant caspases (caspase 1, 2, 3, 4, and 6) enhance the permeability of MPT
pore-containing liposomes44 and that mitochondria contain pro-caspase 2, 3, and 9, which are liberated into the cytosol during
apoptosis.13,70 The possibility that other members of the
caspase family reside in the mitochondria and contribute to mitochondrial events during induction of the apoptotic pathway cannot
be ruled out. More interestingly, results obtained with C2 are
reminiscent of what has been shown with the mammalian cell death
protein Bax, which targets the mitochondria and causes release of Cyt.C
without MPT pore opening and mitochondrial swelling.71 Our
findings do not exclude the possibility that changes in mitochondrial m are critical for apoptosis in some instances, but
support the notion that this may not be a universal step in the
apoptotic pathway.
Antitumor activity of pMC540 could be reconstituted by simultaneous
exposure to C1, C2, and C5.
Having purified the reactive components from the photoactivated mixture
pMC540 and highlighted their mechanism(s) of induction of apoptosis, we
next asked if the antitumor activity observed in our earlier report
with pMC540 could be reconstituted by simultaneous exposure of tumor
cells to appropriate concentrations of C1, C2, and C5. As mentioned
earlier, the relative yields of C1, C2, and C5 from the pMC540 mixture
were approximately 16% each for C1 and C2 and 68% for C5. Keeping
these relative yields in mind, HL60 cells were treated with either 150 µg/mL of pMC540 or 25 µg/mL (16%) of C1 or C2 alone or 100 µg/mL
of C5 alone, or all possible combinations of the three purified
compounds (C1, C2, and C5). All three parameters, vis a vis activation
of caspases 8 and 3 (12 and 18 hours after treatment) and actual cell
death (18 hours), were determined. A summary of the results is
presented in Table 1. It is interesting to
note that, whereas all three purified compounds at the respective
concentrations were able to activate caspase 8, C2 (25 µg/mL) was the
most potent and the earliest caspase 8 activator (2.5× at 12 hours), and this activity correlated with downstream activation of
caspase 3 (1.8×) and the percentage of apoptosis assessed by the
MTT assay (28%). Moreover, the presence of C2 in the combination
experiments (C1+C2 or C2+C5) could explain most of the caspase 8 and 3 activation and tumor cell death observed in these experiments.
Furthermore, inclusion of all three compounds (C1+C2+C5) resulted in a
much enhanced activation of caspases 8 (6×) and 3 (7×) and
significantly increased cell death (50%). We make two conclusions from
these data: first, that a combination of purified compounds used at the
same ratio as recovered from the pMC540 mixture results in
reconstitution of the total pMC540 anti-tumor activity; and second,
that C2 is the most active of the three purified compounds. The later
deserves special mention due to the fact that C2 is a thiouracil
derivative and that one similar compound, 5-fluorouracil (5FU), is a
commonly used chemotherapeutic drug. A closer comparison of the
antitumor activities of C2 and 5FU is the focus of one of our ongoing
studies.
Conclusion.
Taking these findings together, we have gained insight into the
antitumor activity of novel compounds purified from a photoactivated mixture of MC540. In light of these findings, we propose a model for
the mode of action of C1 and C2 whereby non-receptor-dependent drug-induced activation of caspase 8 and release of Cyt.C from the
mitochondria lead to the activation of downstream executioner caspase 3 and tumor cell death. Whereas tumor cell apoptosis triggered by both C1
and C2 is caspase 8 dependent, the actual tumor cell death correlated
better with the amount of caspase 3 activation, further highlighting
the executioner role of caspase 3 in the apoptotic process. The release
of Cyt.C appears to be a universal event; however, we show that this
can occur in the absence of MPT pore opening and independently of a
decrease in m, as with C2. On the contrary, the mode
of Cyt.C release triggered by C1 seems to suggest a mechanism(s)
similar to caspase 8-mediated apoptosis involving cleavage of the
proapoptotic protein Bid upon ligation of the CD95
receptor.61,72,73 Our results seem to suggest that
drug-induced Bax-like activity is a more efficient activator of the
apoptotic pathway than drug-induced Bid-like activity. How does a
particular drug influence the recruitment of Bid-like activity or
Bax-like mechanism upon activation of caspase 8? Elucidation of this
mechanism(s) in drug-induced apoptosis is the focus of our ongoing investigations.
| |
ACKNOWLEDGMENT |
The authors thank Dr M. Kini and Dr Dong Hui (Bioscience Center, NUS,
Singapore) for their assistance with the purification of the
photoproducts and for discussions on the MS and NMR data and thank
Muneeza Jhonkar for help with manuscript preparation.
| |
FOOTNOTES |
Submitted February 16, 1999; accepted April 1, 1999.
Supported by Grant No. RP3970333 from the National University of Singapore.
The publication costs of this
article were defrayed in part by
page charge payment. This article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. section
1734 solely to indicate this fact.
Address reprint requests to Shazib Pervaiz, MD, PhD, Department of
Physiology, Faculty of Medicine, National University of Singapore,
10 Kent Ridge Crescent, Singapore 119260; e-mail:
phssp{at}leonis.nus.edu.sg.
| |
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ABSTRACT
INTRODUCTION