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Department of Pharmacological Sciences, School of Pharmacy (S.C., E.B., A.M., F.C., M.P.A.) and Center of Excellence for Neurodegenerative Diseases (S.C., F.C., M.P.A.), University of Milan, Milan, Italy; and Department of Ultrastructures, Istituto Superiore di Sanitá, Rome, Italy (P.M., W.M.)
Received November 25, 2002; accepted March 14, 2003.
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Abstract |
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 Top Abstract Materials and MethodsResultsDiscussionReferences |
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).
Multicellular organisms eliminate abnormal, damaged or unwanted cells
through apoptosis. Besides genetic mutations promoting unrestrained cellular
proliferation, the transformation of a normal to a neoplastic cell (and then
to more malignant phenotypes) is linked to the acquisition of genetic
resistance to induction of apoptosis
(Kleihues et al., 1995). For
this reason and because of their high invasiveness, these tumors are often
characterized by a poor prognosis and by unsatisfactory response to
pharmacological agents.
Several anticancer drugs are therapeutically effective in that they trigger
the apoptotic death of malignant cells via activation of different (in many
cases yet-to-be identified) mechanisms
(Kaufmann and Earnshaw, 2000).
Thus, the use of apoptosis-inducing molecules and the evaluation of their
mechanism(s) of action in appropriate in vitro models might not only disclose
new possible strategies for the pharmacological manipulation of cancer but may
also help understanding the intracellular pathways of apoptosis that are still
active in cancerous cells and could therefore be exploited in anticancer
therapy.
Purine-based molecules inducing apoptosis have been used for years as
anticancer agents (Plunkett and Saunders,
1991). Among these, the 2-chloro derivative of deoxy-adenosine
(2-CdA; Cladribine) has been employed for the pharmacological control of slow
growing leukemias and lymphomas (Bryson and
Sorkin, 1993) and has become the drug of choice in hairy cell
leukemia (Lauria et al.,
1997), even though its mechanism of action remains unclear. It is
worth noting that Cladribine has been extensively evaluated in blood tumors
but only very limited information is available regarding its possible use in
nonhematological tumors. In this respect, a single study in a human carcinoma
cell line (HepG2 cells) has been reported
(Graziadei et al., 1998).
All apoptotic pathways converge on a family of cysteine-aspartases, named
caspases (Earnshaw et al.,
1999), whose activity drives the biochemical events leading to
cellular disassembly and death. Caspases are present as inactive precursors.
"Initiator" caspases are thought to autoactivate themselves
proteolytically (see also below), whereas "effector" caspases are
activated by initiator caspases and are responsible for execution of cell
death (Earnshaw et al., 1999).
Two main pathways leading to the activation of initiator caspases have been
recognized: the "mitochondrial" and the
"death-receptor" pathways of cell death, involving caspase-9 and
caspase-8, respectively (Scaffidi et al.,
1998; Slee et al.,
1999). In several different experimental models, the main effector
caspase activated by caspase-9 and caspase-8 is caspase-3
(Earnshaw et al., 1999). The
role of other caspases is still a matter of debate. For instance, caspase 2,
known as a downstream "effector" caspase
(Paroni et al., 2001) has
recently been hypothesized to also play an upstream role in some cellular
model systems (Robertson et al.,
2002; Caballero-Benitez and
Moran, 2003; Mueller et al.,
2003).
The apoptotic effects of 2-CdA have been correlated to activation of
distinct caspase cascades. For instance, in MOLT-4 leukemia cells, 2-CdA
activates the death receptor/caspase-8 pathway through activation of the
Fas/Fas-L system (Nomura et al.,
2000). Conversely, in cell-free extracts, 2-chloro-deoxyATP (the
product of the intracellular phosphorylation/activation of 2-CdA) can activate
caspase-9 (and subsequently caspase-3) by participating to the formation of
the so-called "apoptosome"
(Leoni et al., 1998;
Genini et al., 2000a).
Moreover, in chronic lymphocytic leukemia cells, nucleotide derivatives can
damage mitochondria, leading to a drop of mitochondrial membrane potential and
release of cytochrome c and apoptosis inducing factor
(Genini et al., 2000b).
We have previously demonstrated that, upon their intracellular
phosphorylation/activation by two distinct kinases, 2-CdA and its
chemically-related adenosine analog 2-chloro adenosine (2-CA) can trigger
apoptosis of human astrocytoma cells (ADF cells;
Ceruti et al., 2000). Because
of the poor prognosis and unsatisfactorily pharmacological response that often
characterize this central nervous system tumor (see above), the identification
of new proapoptotic agents and the characterization of their mechanisms of
cell death are likely to disclose new pharmacological tools of potential
therapeutic importance. Moreover, adenosine analogs can be used as
experimental tools to dissect the apoptotic pathways that are still expressed
in cancerous astrocytoma cells. On this basis, the present study has been
aimed at analyzing the caspase cascade activated by 2-CdA and 2-CA in the same
experimental model.
Results show that, in these cells, induction of apoptosis by adenosine analogs occurs through activation of an atypical apoptotic cascade involving caspase-2 as an "initiator" caspase and "effector" caspase-3, with no concomitant involvement of either mitochondrial depolarization, release of cytochrome c, or activation of caspase-9 or caspase-8.
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Materials and Methods |
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TopAbstractMaterials and MethodsResultsDiscussionReferences |
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). Cells
were seeded on Petri dishes according to the experimental protocol (see below)
and treated with drugs 24 h after plating to allow them to attach to culture
dishes. Human promonocytic leukemia U937 cells were grown in RPMI 1640 + HEPES (Sigma, Italy) supplemented with 10% FBS, 2 mM glutamine, 100 u/ml penicillin, and 100 µg/ml streptomycin. Cells were seeded on Petri dishes according to the experimental protocol and pharmacological treatments were performed immediately.
Isolation and Activation of Peripheral Blood Lymphocytes. Human peripheral blood lymphocytes (PBL) from healthy donors were isolated from fresh heparinized blood through a Ficoll-Hypaque density gradient centrifugation and grown in standard conditions. In selected experiments, purified T cells were activated for 72 h with phytohemagglutinin (2 µg/ml; Roche, Milan, Italy) and interleukin-2 (60 IU/ml; Invitrogen, Milan, Italy).
Pharmacological Treatments 2-Chloro-2'-deoxy-adenosine (2-CdA; Sigma, Italy), Etoposide (Eto; Sigma, Milan, Italy) betulinic acid (BetA; Vinci-Biochem, Vinci, Italy) and the caspase inhibitors N-benzyloxycarbonyl-Val-Ala-DL-Asp-fluoromethylketone (zVAD-fmk; Vinci-Biochem), N-benzyloxycarbonyl-Asp-Glu-Val-Asp-fluoromethylketone (zDEVD-fmk), N-benzyloxycarbonyl-Leu-Glu-His-Asp-fluoromethylketone (zLEHD-fmk), N-benzyloxycarbonyl-Ile-Glu-Thr-Asp-fluoromethylketone (zIETD-fmk), and N-benzyloxycarbonyl-Val-Asp-Val-Ala-Asp-fluoromethylketone (zVDVAD-fmk) [all from Biomol (Trimital, Italy)] were dissolved in dimethyl sulfoxide (10-2M) and stored at -20°C as stock solutions. 2-Chloro-adenosine (2-CA) and 2-deoxyribose (dRib; both from Sigma) were stored powdered and dissolved immediately before experiments. Solutions of all drugs (10x) were prepared in sterile Hanks' balanced salt solution (Celbio, Milan, Italy). Sodium cyanide (NaCN) stock solution (38 mM) has been purchased from Fluka (Buchs, Switzerland) and Sigma. In selected experiments, zVAD-fmk was added to cultures 7 h after adenosine analogs (see Results); all analyses were performed immediately at the end of the incubation period (1–24 h).
CD95 Triggering. To activate the CD95/Fas pathway, 500 ng/ml of an
anti-human Fas IgM monoclonal antibody [anti-CD95/Fas-triggering monoclonal
antibody (
-Fas t, clone CH11; Upstate Biotechnology, Lake Placid, NY)]
was either added to resting or activated human lymphocytes (see below) or to
ADF cells for 48 h.
Western Blotting Analysis
Evaluation of Caspase Activation. Whole-cell lysates were prepared
and analyzed by immunoblotting, as described previously
(Abbracchio et al., 1997).
Sixty micrograms of proteins in each lane were size-fractionated by
SDS-polyacrylamide gel electrophoresis in 9 or 15% acrylamide gel for
caspase-9 and caspase-3, respectively, followed by electroblotting onto
nitrocellulose membrane. Membranes were then overnight-probed with mouse
monoclonal anti-caspase-3 antibody (1:750 in 3% nonfat dry milk in TBS;
Vinci-Biochem) or with rabbit anti-caspase-9 polyclonal antiserum (1:50 in 3%
nonfat dry milk in TBS; Vinci-Biochem), followed by corresponding
species-specific IgG antibodies conjugated to horseradish peroxidase (1:2000
for anti-mouse and 1:3500 for anti-rabbit antibodies, respectively; 1 h at
room temperature, in 3% nonfat dry milk in TBS; Sigma). Detection of proteins
was then performed by enhanced chemilumi-nescence (Amersham Biosciences,
Milan, Italy) and autoradiography.
Evaluation of Cytochrome c Release from Mitochondria. At the end of the incubation period with the various pharmacological agents, culture supernatants were collected from petri dishes and adhering cells were detached using phosphate-buffered saline + 0.04% EDTA. The total cell suspension was centrifuged, washed in ice-cold phosphate-buffered saline, and the cytosolic and mitochondrial fractions were separated by means of the cytochrome c releasing apoptosis assay kit (BioVision; Vinci-Biochem), following the manufacturer's instructions. Briefly, cells were incubated on ice in cytosol extraction buffer mix (as supplied with the kit) containing 1 mM 1,4-dithio-DL-threitol and protease inhibitor cocktail, homogenized in an ice-cold tissue grinder (40 strokes in ice), and centrifuged at 700g for 10 min at 4°C. Pellets were discarded and supernatants were then centrifuged at 10,000g for 30 min at 4°C to obtain the cytosolic fraction (supernatants). Finally, pellets were resuspended in mitochondrial extraction buffer mix (as supplied with the kit) containing 1 mM 1,4-dithio-DL-threitol and protease inhibitor cock-tail, vortexed for 10 s, and saved as mitochondrial fraction.
Protein content in each fraction was determined according to the method of
Bradford (1976) and protein
samples for SDS-polyacrylamide gel electrophoresis were prepared as described
above. Twenty micrograms of proteins in each lane were size-fractionated in
15% acrylamide gel followed by electroblotting onto nitrocellulose membrane.
Membranes were then probed overnight with a monoclonal mouse anti-cytochrome
c antibody (supplied with the kit; 1:200 in 3% nonfat dry milk in
TBS), followed by IgG anti-mouse antibody conjugated to horseradish peroxidase
(Sigma; 1:2000), enhanced chemiluminescence detection of proteins, and
autoradiography. Densitometric analysis of the 12-kDa protein band detected
both in the cytosolic and in the mitochondrial fraction and corresponding to
cytochrome c was performed by means of the NIH Image program (ver.
1.52;
http://rsb.info.nih.gov/nih-image/Default.html).
Detection of Caspase Activity
Caspase activity was measured by means of a spectrophotometric assay kit
(CaspACE Assay System Colorimetric; Promega, Milan, Italy), following
manufacturer's instructions with some minor modifications. Briefly, at the end
of the incubation period, cells were collected in Cell Lysis Buffer (as
supplied by the manufacturer), exposed to repeated freeze/thawing cycles, and
incubated for 15 min on ice. Insoluble fraction was discarded by
centrifugation (5 min at 15,000 rpm) and the protein content in the
supernatant was determined according to the method of Bradford
(1976) and subsequently
adjusted to desired concentration in caspase assay buffer (as supplied by the
manufacturer). Several different protein concentrations have been tested, and
the best results have been obtained with a total amount of 50 to 60 µg in
each sample. The determination of caspase activity was carried out in a
96-well plate in the same buffer in a total volume of 100 µl, in the
presence of the corresponding tetrapeptide conjugated to paranitroaniline
(i.e., DEVD-pNA, LEHD-pNA IETD-pNA, and VDVAD-pNA in the case of caspase-3,
-9, -8, and -2, respectively; final concentration, 200 µM). Extracts were
incubated for 4 h at 37°C; at the end of the incubation period, released
pNA was measured in a spectrophotometer at 405 nm. The specificity of the
recorded absorbance was determined by adding the corresponding–CHO
tetrapeptide to the reaction mixture (final concentration, 20 µM); a 60 to
80% reduction of the signal was considered significant and specific. Each
condition was run in duplicate, and at least four independent experiments have
been performed.
Cytofluorimetric Analysis
Evaluation of Apoptosis. The percentage of apoptotic cells in the
total population (adhering + detached cells) was evaluated immediately at the
end of the incubation period by means of propidium iodide staining of DNA
followed by flow cytometric analysis, as described previously
(Ceruti et al., 2000).
Evaluation of Cell Death Receptors. Expression of CD95/Fas on the cell surface of activated human lymphocytes and ADF cells was verified by using monoclonal antibodies to human CD95 directly conjugated to (R)-phycoerythrin (BD Biosciences, San Jose, CA).
Analysis of Mitochondrial Membrane Potential. Changes in
mitochondrial membrane potential (
m) induced in the total cell
population by different pharmacological agents were analyzed by means of the
fluorescent dye
5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolcarbocyanine
iodide (JC-1; stock solution: 10-2M in dimethyl
sulfoxide; Molecular Probes, Societá Italiana Chimici, Rome, Italy), as
described previously (Ceruti et al.,
1997). J-aggregate fluorescence was recorded by flow cytometry in
the fluorescence channel 2 (FL2) and monomer fluorescence in the fluorescence
channel 1 FL1 necrotic fragments were electronically gated out on the basis of
morphological characteristics on the forward light scatter versus side light
scatter dot plot.
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Results |
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TopAbstractMaterials and MethodsResultsDiscussionReferences |
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).
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To investigate the role of caspases in this effect, we have started our
analysis from the most important "effector" caspase [i.e.,
caspase-3 (Earnshaw et al.,
1999)].
Because caspase activation is a relatively early event in the apoptotic
program, to evaluate in detail the time-dependence of apoptosis induction,
human astrocytoma cells were exposed for graded (1–24 h) periods of time
to 100 µM 2-CA or 2-CdA. At the end of the incubation period, both
caspase-3 activation and induction of apoptosis were evaluated. As shown in
Fig. 2, both adenosine analogs
induced a statistically significant activation of caspase-3 starting from a
10-h exposure, as evaluated by spectrophotometric analysis of yellow pNA
release caused by the cleavage of the synthetic tetrapeptide substrate
DEVD-pNA. Maximal caspase-3 activation was present at 20/22 h, when an 18-fold
increase of enzyme activity with respect to basal levels was detected.
Activation tended to decrease starting from 24 h. At any time point showing a
significant activation of the enzyme (10–24 h), an 80 to 92% reduction
of absorbance in the presence of DEVD-CHO with respect to corresponding
adenosine analog alone was detected (data not shown), therefore confirming
that pNA release was indeed specifically caused by the action of active
caspase-3 (see Materials and Methods for more detail). Activation of
caspase-3 preceded the appearance of nuclear signs of apoptosis, as evaluated
by cytofluorometric analysis of PI stained nuclei
(Fig. 2, line graphs),
therefore suggesting the existence of a temporal relationship between
caspase-3 activation and induction of cell death by adenosine analogs.
Activation of caspase-3 was also confirmed by immunoblotting analysis, with an
antibody recognizing both the inactive proenzyme (detected as a 32-kDa band)
and its active proteolytic fragments (detected as 17- and 12-kDa protein
bands; Fig. 3). In agreement
with the enzymatic activation data, active caspase-3 fragments were detected
starting from 10 h after beginning exposure of cells to either 2-CA or 2-CdA.
To support a role of caspase-3 in adenosine analog-induced apoptosis,
prevention of the intracellular phosphorylation/activation of 2-CA and 2-CdA
by specific inhibitors (Ceruti et al.,
2000) abolished both caspase-3 activation and induction of cell
death (Ceruti et al.,
2003).
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To further confirm the causal relationship between caspase-3 activation and induction of apoptosis, 2-CA- and 2-CdA-treated cultures were exposed to the pan-caspase inhibitor zVAD-fmk, and both caspase-3 activation and the percentage of nuclear apoptosis induced by the two analogs alone or in combination with this synthetic tetrapeptide were evaluated (Fig. 4). As described in Figs. 2 and 3, significant caspase-3 activation can be detected 10 h after addition of either 2-CA or 2-CdA. Hence, in this set of experiments, cells were exposed to either adenosine analog, and the pancaspase inhibitor (20–40 µM final concentration) was added 7 h later to inhibit the enzyme as soon as it was activated; both the extent of caspase-3 activation and the degree of apoptosis were determined after a total of 24 h in culture. As expected, 2-CA and 2-CdA significantly activated caspase-3 (Fig. 4A) and induced the appearance of an hypodiploid DNA peak, typical of apoptosis (Fig. 4, B and C). zVAD-fmk completely prevented both the activation of caspase-3 (Fig. 4A) and the appearance of the nuclear signs of apoptosis (Fig. 4, B and C) induced by either adenosine analog, therefore confirming the key role played by caspases in our experimental model. A partial, but statistically significant, protection against 2-CA and 2-CdA-induced apoptosis was already detected at a lower zVAD-fmk concentration (10 µM; data not shown). To further confirm the above results, when added to intact cells during the incubation with adenosine analogs, the specific caspase-3 inhibitor zDEVD-fmk was also able to prevent apoptosis (Fig. 9, see also below).
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Apoptosis by Adenosine Analogs Is Not Preceded by Changes of
m. In an attempt to characterize the apoptotic pathways
activated by adenosine analogs upstream of caspase-3, we focused on the
possible role of mitochondria, because this pathway has been involved in
2-CdA-induced apoptosis in other experimental models
(Genini et al., 2000b;
Marzo et al., 2001). Possible
changes in m have been evaluated by means of cytofluorometric
analysis after loading cells with the mitochondrial specific cationic dye JC-1
(Ceruti et al., 1997). In
healthy cells with polarized mitochondria, JC-1 accumulates in the
transmembrane space and forms the so-called "J-aggregates",
emitting orange fluorescence at 590 nm when excited at 490 nm and thereby
detectable by flow cytometry in fluorescence channel 2 (FL2; see
y-axis in Fig. 5
graphs). Drops in m result in disappearance of J-aggregates and
formation of JC-1 monomers that emit a greenish-yellow fluorescence at 530 nm
when excited at 490 nm, corresponding to fluorescence channel 1 (FL1, see
x-axis in Fig. 5
graphs). It is therefore possible to monitor changes in m by
looking at the number of cells showing fluorescence emission in the two
channels. In control untreated cultures, because of the equilibrium existing
between J-aggregates and monomers in healthy cells, the majority of cells
(96.25%) shows a high emission of fluorescence in both channels and is
therefore found in the upper right quadrant of the plot
(Fig. 5A). Only a very small
percentage of control untreated cells (3.75%) shows low emission of
fluorescence in FL2 and can therefore be found in the lower right (LR)
quadrant of the plot (Fig. 5A).
In initial experiments, we have evaluated the changes of m induced
by agents known to interfere with mitochondrial function, such as NaCN
(Rutter and Rizzuto, 2000),
dRib (Ceruti et al., 1997), and
the anticancer agent BetA (Fulda et al.,
1998), which have been used as positive controls. Also in human
astrocytoma cells, these agents induced dramatic drops in m
(Fig. 5A); for example, a 4-h
exposure to 3.8 mM NaCN resulted in a dramatic decrease of J-aggregate
fluorescence (y-axis; FL2), accompanied by a concomitant increase of
fluorescence in FL1, so that 80.75% of cells were now found in the lower right
quadrant of the plot (Fig. 5A).
In a similar way, marked increases of the percentage of cells with depolarized
mitochondria were detected with the other two depolarizing agents
(Fig. 5A). For all agents,
induction of mitochondrial depolarization was followed, a few hours later, by
induction of cell death (data not shown). These results demonstrate that: 1)
alterations of m can be induced in ADF cells and 2) with the above
method, changes of mitochondrial function induced by toxic stimuli can be
detected before appearance of cell death. We therefore evaluated
possible changes in m after exposure of cultures to adenosine
analogs. To check whether changes in m preceded or followed
caspase-3 activation, we exposed cultures to 2-CA and 2-CdA: 7 h showed no
significant activation; 10, 15, and 18 h showed partial activation; 20 and 22
h showed maximal activation, and 24 h showed declining activation (see
Fig. 2). As shown in
Fig. 5B, which reports the
percentage of cells in the upper or lower right quadrants of the m
analysis plot, significant alterations of m could be detected only
starting from an 18-h exposure to either adenosine analog, a quite late time
at which cells are already committed to apoptotic cell death. Therefore, we
conclude that the mitochondrial depolarization detected at quite late stages
of adenosine analog-induced apoptosis probably reflects a consequence, rather
than a cause, of cell death.
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Apoptosis Induced by Adenosine Analogs Does Not Involve Cytochrome
c Release from Mitochondria nor Caspase-9 Activation. Activation
of the mitochondrial pathway of cell death is coupled, sequentially, to a drop
in m, cytochrome c release, formation of the apoptosome,
stimulation of the "initiator" caspase-9, and activation of
effector caspases (e.g., caspase-3; Li et
al., 1997). Lack of early mitochondrial depolarization upon
exposure to adenosine analogs (Fig.
5B) would rule out a role for cytochrome c release and
caspase-9 in our experimental model. However, because a significant release of
cytochrome c can also be detected in the absence of changes of
m (Grubb et al.,
2001), specific experiments were done to investigate the
involvement of both cytochrome c and caspase-9 in the apoptosis
induced by adenosine analogs. The subcellular localization of cytochrome
c was evaluated by Western blotting analysis in the mitochondrial and
cytosolic fractions of control untreated cells and of cells exposed to either
2-CA or 2-CdA for various time periods (7, 10, 15, and 24 h). Cytochrome
c was easily detected in the mitochondria of both control and treated
cells (Fig. 6A); however, in no
case were we able to detect any accumulation of cytochrome c in the
cytoplasm of cells exposed to either 2-CA or 2-CdA. We have applied the same
methods to monitor cytochrome c release upon exposure to either NaCN,
dRib, or BetA, which induced mitochondrial depolarization in these cells
(Fig. 5A). Exposure to these
agents indeed resulted in marked release of cytochrome c from
mitochondria to the cytosolic fraction
(Fig. 6B), hence confirming the
suitability of these methods to follow changes of cytochrome c
subcellular localization in these cells.
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Finally, we assessed the presence and activation of caspase-9 in cells
treated with either 2-CA or 2-CdA. Evaluation of caspase-9 activation by the
same methods used to detect caspase-3 (i.e., cleavage of the specific
caspase-9 substrate LEHD-pNA and release of pNA;
Fig. 7A) revealed no
significant stimulation of enzyme activity. A small not statistically
significant stimulation of caspase-9 was detected after 20 h, a time when the
apoptotic process in these cells is already well established. We have applied
exactly the same methods to measure caspase-9 activity in another cell line
(U937 cells) that has been previously described to undergo caspase-9
activation and cell death upon exposure to 2-CdA
(Marzo et al., 2001). In these
cells, here used as a positive control for caspase-9 activation, both 2-CdA
and the pro-apoptotic agent Eto (Plo et
al., 2002) induced marked mitochondrial depolarization
(Fig. 7B) and significantly
increased caspase-9 activity with respect to control untreated cells
(Fig. 7C). To further rule out
a role for caspase-9 in adenosine analog-induced apoptosis of human
astrocytoma cells, Western blotting experiments with an antibody recognizing
both the inactive form of pro-caspase-9 and its active proteolytic fragment
revealed no activation of this caspase upon exposure to either 2-CA or 2-CdA
(data not shown). Globally, these results confirm that the lack of caspase-9
activity in human astrocytoma cells upon exposure to adenosine analogs is
actually caused by the inability of these agents to activate this pathway of
cell death.
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No Role for Caspase-8 in Adenosine Analog-Induced Apoptosis in Human
Astrocytoma Cells. In a similar way to caspase-9, a detailed (7–24
h) kinetic analysis of the activity of caspase-8, that is a part of the
"death receptor" pathway and has been demonstrated to be at the
basis of 2-CdA-induced cell death in leukemia cells
(Nomura et al., 2000), was
performed. To this purpose, cell extracts have been incubated with a specific
caspase-8 substrate, IETD-pNA, and enzyme activity has been evaluated as
described under Materials and Methods. A specific activation of the
enzyme (corresponding to a 1.8- to 2-fold increase over basal activity) has
been detected only starting from a rather late time period of incubation (18
h; Fig. 8), suggesting that its
activation is more a consequence than a cause of commitment to cell death,
hence ruling out a role for this caspase in triggering the apoptotic cascade
activated by adenosine analogs in astrocytoma cells. To rule out a role for
the "death" receptor pathway in initiating apoptosis in human
astrocytoma cells, activation of the Fas/Fas ligand pathway by exposure to
anti-FAS antibodies (a typical trigger of the caspase-8-dependent pathway;
Parlato et al., 2000) did not
result in any increase of the percentage of apoptotic cells (7.1 ±
1.8%) with respect to control untreated cells (6.6 ± 2.2). The same
concentration of anti-FAS antibodies induced highly significant apoptosis in
human phytohemagglutinin/interleukin-2–activated T lymphocytes, a
typical Fas-sensitive cell type (36 ± 3% after 48 h compared with 6.6
± 1.8% detected in resting cells, mean value of four different
experiments, p < 0.001, Student' s t test), whereas it
was ineffective in resting cells. The lack of effect of anti-Fas antibodies on
the human astrocytoma cells was not caused by the absence of surface
receptors, because both these cells and human T lymphocytes were CD95/Fas
positive when analyzed cytofluorimetrically with a specific antibody (data not
shown).
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Caspase-2 Is the Initiator Caspase Involved in Induction of Apoptosis by
Adenosine Analogs in Human Astrocytoma Cells. Although caspase-2
has never been reported to participate to induction of apoptosis by adenosine
analogs, we also determined the role of this caspase in our experimental model
by assessing whether zVDVAD-fmk, using a concentration (3 µM) that
selectively inhibits this caspase (Schotte
et al., 1999), could prevent induction of apoptosis by 2-CA or
2-CdA. In parallel, the effects induced by the selective caspase-8 inhibitor
zIETD-fmk, the caspase-9 inhibitor zLEHD-fmk, the caspase-3 inhibitor
zDEVD-fmk, and the pan-caspase inhibitor zVAD-fmk (all used at 3 µM) have
been assessed.
In line with the data reported above, which rule out a role for caspase-8
and caspase-9 in our experimental model, neither zIETD-fmk nor zLEHD-fmk could
prevent the apoptosis induced by either 2-CA or 2-CdA
(Fig. 9), whereas a highly
significant reduction of adenosine analog-induced apoptosis was observed with
both the selective caspase-2 inhibitor zVDVAD-fmk, the selective caspase-3
inhibitor zDEVD-fmk, and, as expected, with the pan-caspase inhibitor zVAD-fmk
(Fig. 9). A full protection
against adenosine analogs-induced apoptosis was obtained at inhibitor
concentrations higher than 3 µM (data not shown); however, these are not
selective for any specific caspase
(Schotte et al., 1999).
To confirm the involvement of caspase-2, we evaluated the ability of 2-CdA and 2-CA to activate this caspase. By using a colorimetric assay similar to those used for the other caspases, we demonstrated statistically significant increases of caspase-2 activity at times (1–7 h after addition of either adenosine analog to cells; Fig. 10A) that precede the activation of effector caspase-3 (Fig. 2). Stimulation of caspase-2 by 2-CdA seemed to occur slightly earlier (already statistically significant at 1 h) and more robustly with respect to 2-CA (Fig. 10A). For both 2-CA and 2-CdA, caspase-2 activation started to decline at 10 h, a time at which effector caspase-3 is already highly activated in these cells (Fig. 2). Activation of caspase-2 by 2-CA or 2-CdA was specific, because it could be significantly reduced by treating cells with either adenosine analog in the presence of the selective caspase-2 inhibitor zVDVAD-fmk (Fig. 10B).
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Discussion |
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TopAbstractMaterials and MethodsResultsDiscussionReferences |
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), and, for the first time, shed light on the role of
caspases in such effects. Main findings of the present study are the following:
), neither zLEHD-fmk nor zIETD-fmk could prevent the
induction of cell death by either 2-CA or 2-CdA when used at low
concentrations that should selectively inhibit caspase-9 and caspase-8,
respectively. This finding is in line with the other data reported here that
rule out a role for caspase-9 and caspase-8 in our experimental model.
On this basis, we conclude that adenosine analogs can trigger the apoptosis
of astrocytoma cells by a cellular pathway that does not involve mitochondrial
depolarization, release of cytochrome c, or activation of either
caspase-9 or caspase-8 but requires activation of caspase-2 and effector
caspase-3. Our data are at variance with results obtained in other
experimental systems [i.e., Jurkat cells
(Robertson et al., 2002) or
transformed human fibroblasts (Lassus et
al., 2002)], where caspase-3 activation by caspase-2 has been
reported to involve cytochrome c release and caspase-9 activation.
The mechanistic basis for caspase-3 activation by caspase-2 in our
experimental model remains to be elucidated. This atypical pathway of cell
death may represent a novel biological target for the pharmacological
manipulation of tumor growth. Our results also underline the possibility that,
under specific cellular paradigms, caspase-2 may act as an
"initiator" caspase. Determining an emergent function for this
protease has been difficult, with results assigning this caspase a downstream
or feedback role (reviewed in Robertson et
al., 2002). Very recently, however, in caspase-9– blocked
testicular cancer cells, initiation of apoptosis by cisplatin has been
demonstrated to occur through activation of caspase-2 and caspase-3
(Mueller et al., 2003).
Moreover, in cerebellar granule cells, staurosporine-induced apoptosis has
been shown to require induction of caspase-2 before caspase-3
(Caballero-Benitez and Moran,
2003). The present data obtained with adenosine analogs in human
astrocytoma cells suggest that this apoptotic cascade may be recruited more
often than originally suspected.
As underlined above, our results are at variance from results obtained in
other experimental models. In MOLT-4 leukemic cells, 2-CdA–induced
apoptosis has been shown to crucially involve the Fas/Fas ligand/caspase-8
pathway (Nomura et al., 2000).
In other human leukemia cell lines, 2-CdA–induced cell death has been
shown to be always preceded by a loss of mitochondrial membrane potential
(Genini et al.,
2000a,b;
Marzo et al., 2001). These
differences may be attributable to the differential activation of specific
apoptotic programs in cells belonging to different lineages (immune cells in
the case of the above studies with respect to astrocytoma cells in the present
study) and/or to differential regulation of caspase activity. For instance, in
human leukemia cells, caspase-9 activation by 2-CdA can be prevented by
Bcl2 (Genini et al.,
2000b). No immunoreactivity to this protein can be detected in
human astrocytoma ADF cells, which instead express Bcl-XL at
relatively high level (Abbracchio et al.,
1997). Interestingly, Bcl-XL expression is up-regulated
by treatment of cells with adenosine analogs (S. Ceruti, R. Brambilla, and M.
P. Abbracchio, unpublished data). It may well be that, in our experimental
model, this antiapoptotic protein prevents adenosine analogs from promoting
the formation of the apoptosome (and thus from activating procaspase-9), hence
shifting the apoptotic program to other intracellular pathways. The lack of
pro- and antiapoptotic proteins in a simplified cell-free system as in that
used by Leoni and coworkers
(1998) may explain the
different outcome of their study (i.e., activation of caspase-9 by
2-chlorodeoxyATP upstream of caspase-3) with respect to the present
results.
The present data may have important implications for the pharmacological
manipulations of drug-resistant central nervous system tumors. Interestingly,
untransformed astroglial cells and neurons are relatively resistant to 2-CdA-
and 2-CA–induced apoptosis (Ceruti et al.,
1997;
2000), which is quite important
in view of a possible exploitation of adenosine-analog induced apoptosis in
human astrocytoma/glioblastoma tumors. Moreover, the demonstration that, in
human astrocytoma cells, adenosine analogs can induce apoptosis by activating
an alternative pathway of cell death has intriguing implications in
understanding which apoptosis pathways are still expressed by other neoplastic
cells. In this respect, Chandra et al.
(2002) have recently selected
three 2-CdA–resistant leukemic cell lines. Acquisition of resistance to
the apoptosis induced by 2-CdA was accompanied by decreased susceptibility of
mitochondria to undergo permeability transition and lack of cytochrome
c release. Our demonstration that adenosine analogs can also induce
apoptosis by bypassing the mitochondrion would predict that these compounds
could also act as effective anticancer agents in other kinds of human
tumors.
|
Acknowledgements |
|---|
|
Footnotes |
|---|
Part of the present data has been presented at the following meetings: International Symposium "Purine 2000" (Madrid, Spain; published as an abstract in Drug Dev Res 50:90, 2000); "XXX Congress of the Italian Pharmacological Society" (Genova, Italy; published as an abstract in Pharmacol Res 43:129, 2001); "3rd Meeting of the Federation of the European Pharmacological Societies (EPHAR)" (Lyon, France; published as an abstract in Fundam Clin Pharmacol 15:147, 2001).
S.C. and E.B. contributed equally to this work.
ABBREVIATIONS: 2-CdA, 2-chloro-2'-deoxy-adenosine;
m, mitochondrial transmembrane potential; 2-CA,
2-chloro-adenosine; ANOVA, analysis of variance; BetA, betulinic acid;
DEVD-pNA, N-acetyl-Asp-Glu-Val-Asp-paranitroaniline; dRib,
2-deoxyribose; Eto, etoposide; FBS, fetal bovine serum; IETD-pNA,
N-acetyl-Ile-Glu-Thr-Asp-paranitroaniline; JC-1,
5,5',6,6'-tetrachloro-1,1',3,3'-tetraethyl-benzimidazol-carbocyanine
iodide; LEHD-pNA, N-acetyl-Leu-Glu-His-Asp-paranitroaniline; NaCN,
sodium cyanide; PI, propidium iodide; TBS, Tris-buffered saline; zDEVD-fmk,
N-benzyloxycarbonyl-Asp-Glu-Val-Asp-fluoromethylketone; zIETD-fmk,
N-benzyloxycarbonyl-Ile-Glu-Thr-Asp-fluoromethylketone; zLEHD-fmk,
N-benzyloxycarbonyl-Leu-Glu-His-Asp-fluoromethylketone; zVAD-fmk,
N-benzyloxycarbonyl-Val-Ala-DL-Asp-fluoromethylketone;
zVDVAD-fmk, N-benzyloxy
carbonyl-Val-Asp-Val-Ala-Asp-fluoromethylketone; zVDVAD-pNA,
N-benzyloxycarbonyl-Val-Asp-Val-Ala-Asp-paranitroaniline.
Address correspondence to: Prof. Maria P. Abbracchio-Department of Pharmacological Sciences, University of Milan and Center of Excellence for Neurodegenerative Diseases (CEND)-Via Balzaretti, 9-20133 Milan, Italy. E-mail: mariapia.abbracchio{at}unimi.it
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,
p < 0.05 with respect to corresponding control, one-way ANOVA
(Scheffé F test). At the same time points, caspase-3 activity
was determined in parallel samples by evaluating cleavage of the specific
caspase-3 substrate DEVD-pNA and release of pNA at 405 nm (histogram graphs).
Mean control value was 7.8 ± 0.5 pmol of pNA/µg of protein. The
specificity of the release was determined by adding the reversible caspase-3
inhibitor DEVD-CHO during the assay in vitro (see Results). Data
represent the mean ± S.E. of at least four independent experiments, run
in triplicate. *, p < 0.05 with respect to corresponding control,
one-way ANOVA (Scheffe F test).
