panel II). These data indicate that AE-induced apoptosis requires caspase-8–mediated
TO INDUCTION OF CASPASE-8 ACTIVITY AND CELL DEATH
We next tested whether the suppression of CARPs contributes to the activation of caspase-8
in response to AE, as CARPs have previously been shown to regulate the activity of caspase-8
[McDonald and El-Deiry, 2004]. Treatment with AE for 1 h reduced the expression of CARP1
and 2 mRNAs, and this reduction in both mRNAs was maintained for up to 12 h of AE
exposure (Fig. 7A). By using actinomycin D (a transcriptional inhibitor) and cycloheximide (a
translational inhibitor), we observed that decreased CARP mRNA levels by AE was related to
reduced stabilization of the mRNAs. To extend the above observations, we tested whether the
ectopic expression of FLAG-tagged CARP1 or 2 leads to the suppression of caspase-8
activity and apoptosis. Expression levels of FLAG-CARP1 and -2 were confirmed by western
blotting using FLAG-specific antibody (Fig. 7B, panel I). As showed in panels II and III of
Fig. 7B, AE-induced increases of caspase-8 activity and apoptosis were inhibited in the
CARP1- or CARP2-transfected cells compared with control vector–transfected cells. Because
the overexpression of CARPs can contribute to the ubiquitin-mediated proteolysis of
caspase-8 [McDonald and El-Deiry, 2004], we then investigated whether CARP
overexpression affects the levels of caspase-8 protein. As expected, pro-caspase-8 protein
levels were reduced when FLAG-CARP1 or FLAG-CARP2 was transfected in cells (Fig. 7B,
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panel I), whereas pro-caspase-10 protein levels were partially affected. In contrast,
overexpression of FLAG-CARPs did not affect levels of pro-caspase-9 protein. Taken
together, these data demonstrate that the decrease of CARPs is required for caspase-8
activation and apoptosis induction in response to AE.
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DISCUSSION
In this study, we used the mutant p53 (R248L)–carrying cell line and the p53-null human lung
cancer cell lines H1299, Hep3B, and MG-63 to explore the mechanism of p53-independent
apoptosis induced by AE. Transient ectopic expression of wt p53, a p53- and p21-dependent
luciferase reporter assay, and quantitative analyses of DNA fragmentation and caspase-8
activity in the presence of Z-IETD-FMK confirmed the induction of apoptotic cell death by
AE through a caspase-8–dependent but p53-independent pathway. Furthermore, CARP
proteins appear to be a crucial regulator activation of caspase-8 in response to AE, since
overexpression of CARP 1 and 2 reduced AE-induced caspase-8 activation and apoptosis. AE
contains a quinine structure that is predicted to induce ROS production, which may play a role
in the induction of cancer cell apoptosis [Lee et al., 2006]. AE displays an affinity for nuclear
DNA, and high doses of AE disrupt chromatin structure and DNA template function in
susceptible cell lines [Mueller and Stopper, 1999]. The participation of ROS in cancer cell
apoptosis that is stimulated by chemotherapeutic agents through the induction of DNA
damage has been investigated for several decades [Lau et al., 2008]. Oxidative damage to
DNA is a result of the interaction of DNA with ROS. Consistent with previously reported
results [Lee et al., 2006], an increase in intracellular ROS levels was observed when cells
were induced to undergo apoptosis. ERK has been implicated in DNA damage–induced
p53-independent apoptosis, and its activation is regulated by ROS [Wang et al., 1998]. We
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found that AE caused an increase in ROS levels that was completely inhibited by co-treatment
with Z-IETD-FMK or CsA. The phosphorylation of ERK induced by AE was suppressed by
Z-IETD-FMK. Moreover, PD98058 and ERK siRNA significantly attenuated the AE-induced
expression of p21 and S-phase arrest, suggesting that ERK activation is correlated with
AE-mediated p21 induction and S-phase arrest. Our results are in agreement with a previous
report that activated ERK induces p21 expression in a p53-independent manner and that p21
up-regulation results in S-phase arrest [Zhu et al., 2004].
p21 inhibits the phosphorylation of Rb by the cyclin A–Cdk2 complex, which regulates the
transition of G1 to S-phase [Harper et al., 1993]. p21 also inhibits DNA synthesis and cell
growth by binding to PCNA and Cdk2 with its N- and C-terminal regions, respectively
[Rousseau et al., 1999]. The association of p21 with cyclin A–Cdk2 complexes was detected
12–48 h after AE treatment. Attenuating the activation of ERK with PD98059 or ERK siRNA
significantly inhibited the expression of p21 and partially inhibited apoptosis induced by AE.
Overexpression of p21 inhibits TRAIL death receptor DR4–dependent caspase-8 activation
[Xu and El-Deiry, 2000]. An analysis using chimeric caspase-8 with the transmembrane and
extracellular domains of the human CD8α chain did, however, demonstrate that caspase-8
oligomerization at the cell membrane is sufficient for its autoactivation and apoptosis
induction [Martin et al., 1998]. Several chemotherapeutic agents, such as N,N-dimethyl
phytosphingosine and curcumin, induce apoptosis through the cleavage of cytosolic Bid by
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activated caspase-8 [Anto et al., 2002; Kim et al., 2009]. Our results show that Z-IETD-FMK
inhibited the cleavage and activation of caspase-8 and of apoptotic cell death induced by AE.
The FaDu cell line has a homozygous deletion of the death receptor DR4 andisresistant to the
cytotoxic effects of TRAIL [Ozoren et al., 2000]. Moreover, FADD siRNA did not interfere
with the activation of caspase-8 in FaDu, Hep3B, or MG-63 cells treated with AE (data no
shown), suggesting that AE-induced caspase-8 activation and apoptosis do not occur through
the induction of the death receptor. The promoter region of caspase-8 contains an
E2F1-responsive element, and its transcriptional activity can be regulated by E2F1 [Afshar et
al., 2006]. Previous findings also show that radiation does not induce E2F1 activity, caspase-8
activity, or apoptotic cell death when cells express wt p53; however, inhibition of wt p53
function with HPV-16 E6 induces caspase-8 activity and apoptosis following radiation
[Afshar et al., 2006]. Taken together, these observations led us to speculate that the
caspase-8–dependent mitochondrial ROS production is critical for ERK activation and the
subsequent p21 induction and E2F1 expression. The formation of cyclin A–Cdk2–p21
complexes induced by activated ERK after treatment with AE was associated with the
induction of S-phase arrest, and E2F1 upregulation may contribute to caspase-8 activation.
The release of cytochrome c from the mitochondrial intermembrane space to the cytosol in
response to stress stimuli is a critical step for the activation of caspase-9 in apoptotic cell
death [Tsujimoto, 2003]. In cultured dopaminergic PC12 cells, caspase-9 activation via
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cytochrome c release in response to 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine results in
the activation of caspase-3 and -8 and the cleavage of Bid to tBid [Viswanath et al., 2001].
Our results show that either CsA or Z-IETD-FMK inhibited AE-induced loss of ∆ψm and
apoptotic cell death. Co-treatment with Z-IETD-FMK also completely blocked Bid cleavage,
the translocation of tBid to the mitochondria, Bax induction, caspase-9 activation, and
apoptosis. Moreover, experiments using Z-IETD-FMK showed that caspase-8 activity
appeared to be involved in the AE-induced release of cytochrome c, AIF, and Endo G from
the mitochondria and apoptosis. Our findings suggest that caspase-9 was activated directly by
caspase-8 through the release of cytochrome c.
CARPs act as ubiquitin protein ligases that bind to and regulate caspase-8 and -10.
siRNA-mediated attenuation of CARP expression significantly sensitizes cells to
chemotherapy-induced apoptosis and inhibits colony formation by cancer cells [McDonald
and El-Deiry, 2004]. In addition, treatment with TNF-α/cycloheximide or TRAIL leads to
enhanced cleavage of caspase-8 and -10 after treatment with CARP1 or CARP2 siRNA. One
study using a caspase-3 inhibitor or a combination of caspase-8 and -10 inhibitors and the
proteasome inhibitor MG132 demonstrated that cleavage of CARPs was caspase-dependent
during death receptor–mediated apoptosis [McDonald and El-Deiry, 2004]. CARPs are also
overexpressed in a variety of human cancers and cancer cell lines, suggesting that CARPs
could contribute to development of tumors [McDonald and El-Deiry, 2004]. However,
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resistance to chemotherapy-induced or death receptor–mediated apoptosis has not been
observed in cells overexpressing CARP1 or CARP2. AE treatment of FaDu, Hep3B, and
MG-32 cells led to the rapid decrease of both CARP1 and CARP2 mRNAs. Overexpression of
either CARP1 or CARP2 blocked AE-induced caspase-8 activity and apoptosis as well as
decreased the levels of pro-caspase-8 protein. Unexpectedly, pro-caspase-10 protein levels
only slightly decreased in CARP1- or CARP2-overexpressing cells. The sequences of
caspase-8 and -10 are highly similar [Fernandes-Alnemri et al., 1996]. The involvement of
caspase-10 in apoptosis signaling by death receptors has also been reported [Wang et al.,
2001]. However, in vitro experiments showed that overexpression of caspase-10 is able to
induce the process of caspase-3 cleavage and apoptosis [Vincenz and Dixit, 1997].
Caspase-8–deficient cell lines were shown to be resistant toward Apo2L/TRAIL–induced
apoptosis, implying that caspase-8 plays a critical role in death receptor–mediated signaling,
whereas caspase-10 is not important for this function [Bodmer et al., 2000]. Interestingly,
studies using a series of caspase-10–specific antibodies demonstrated that apoptosis signaling
by death receptors involves not only caspase-8 but also caspase-10 [Kischkel et al., 2001]. If
AE treatment induced apoptosis by increasing the activities of both caspase-8 and -10, the
addition of an inhibitor of either caspase-8 or -10 would be predicted to result in decreased
apoptosis. We found that apoptotic induction by AE was inhibited by caspase-8 inhibitor, but
not by caspase-10 inhibitor. Furthermore, caspase-10 cleavage and activation were not
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detected in AE-treated cells (data no shown). These findings indicate that caspase-8 activation
is an important event in induction of apoptosis by AE. These observations together with our
findings suggest that the function and regulation of both caspase-8 and -10 are independent of
apoptosis induction. This raised the possibility that the two proteins have distinct and perhaps
complementary functions.
In conclusion, we present a novel p53-independnet mechanism of AE-induced apoptosis of
cancer cells involving decreased stability of CARP mRNAs and the subsequent induction of
ERK and caspase-8–mediated mitochondrial death pathways. However, the molecular
mechanisms of the diminished CARP mRNA stability in AE-treated cells remain to be further
explored.
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ACKNOWLEDGMENTS
M.-L. Lin was supported by a grant (CMU98-C-06) from China Medical University, Taiwan.
S.-S. Chen was supported by grants from the Taichung Veterans General Hospital and Central
Taiwan University of Science and Technology (TCVGH-CTUST987715), Taiwan.
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FIGURE LEGENDS
Fig. 1. AE induces apoptotic cell death in a p53-independent manner. (A) The effect of AE on
cell viability. FaDu, Hep3B, MG-32, or H1299 cells were plated in 24-well plates and treated
with either DMSO (vehicle control) or the indicated concentrations of AE for 36 h (panel I).
After treatment, cell viability was determined by flow-cytometric analysis of PI uptake. The
values presented are the mean ± standard error from three independent experiments. *p < 0.05,
significantly different from vehicle-treated cells. The IC50 value of AE in these cells is
indicated (panel II). (B) p53 is not involved in the induction of apoptotic cell death or of
caspase-3 activity by AE. Panel I: levels of p53 proteins determined by western blot analysis
with p53 antibodies (left) and levels of p53 transcripts determined by RT-PCR with specific
primers (right) in cells stably expressing the empty control vector (-), GFP shRNA, or p53
shRNA. β-Actin was used as an internal control for sample loading. Panels II and III: after
treatment of the empty control vector (-), GFP shRNA, and p53 shRNA cells with vehicle, AE
(60 µM), AE (60 µM) plus Ac-DEVD-CMK (10 µM), or AE (60 µM) plus Z-VAD-FMK (15
µM) for 36 h, caspase-3 activity and DNA fragmentation were determined using flow
cytometry and cell death–detection ELISA, respectively. The values presented in panels I and
II are the mean ± standard error from three independent experiments.
Fig. 2. p53 activity is not involved in the induction of cell death or of p21 activity. (A) A
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luciferase reporter driven by three p53-binding sites was co-transfected with vector alone
(lanes 1 and 2), wt p53 (lanes 3 and 4), mutant p53 (R248W) (lanes 5 and 6), or mutant p53
(R175H) (lanes 7 and 8) into stably p53 shRNA–expressing cells. (B) As in (A), except that
the reporter was driven by the p21 promoter. At 12 h post-transfection, cells were treated with
vehicle or AE (60 µM) for 24 h, and then luciferase activity was determined. (C and D). p53
shRNA–expressing stable cells were transfected with vector alone (lanes 1 and 2), wt p53
(lanes 3 and 4), mutant p53 (R248W) (lanes 5 and 6), or mutant p53 (R175H) (lanes 7 and 8)
for 12 h. Cells were then treated with either vehicle or AE (60 µM) for 24 h. Cell viability and
protein expression were determined using the MTT assay and western blot analysis,
respectively. γ-Tubulin was used as an internal control for sample loading in panel IV.
Fig. 3. Involvement of cyclin A–Cdk2–p21–E2F1 complexes in AE-induced S-phase cell
cycle arrest. (A) Effects of AE on cyclin protein expression and the levels of cyclin A, p21,
and E2F1 bound to Cdk2. Panel I: after 36 h of treatment of the GFP shRNA– or p53
shRNA–expressing cells with vehicle or AE (60 µM), the levels of cyclin A, cyclin B1, cyclin
D, cyclin E, Cdk2, p-Cdk2 (Thr14/Thr15), E2F1, and PCNA in the cell lysates were analyzed
using specific antibodies. γ-Tubulin was used as an internal control for sample loading. Panel
II: co-immunoprecipitation of cyclin A, Cdk2, and p21 was performed using lysates prepared
from AE (60 µM)–treated p53 shRNA–expressing stable cells at 36 h. The antibody used for
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co-immunoprecipitation is indicated at the top. The proteins in the immunoprecipitated
complexes were detected using western blotting with specific antibodies. Mouse normal IgG
was used as a control for antibody specificity. (B) At 12 h after transfection of p53
shRNA–expressing cells with control or cyclin A siRNA, cells were treated with either
vehicle or AE (60 µM) for 24 h. After WB was used to examine the cyclin A and Cdk2 levels,
the percentage of cells in S-phase was analyzed by flow-cytometric analysis of PI-stained
cells.
Fig. 4. Activation of ERK is involved in the induction of p21 and E2F1 expression, S-phase
arrest, and apoptosis. At 12 h after transfection with ERK siRNA (pKD-MAPK1/Erk) or 2 h
after treatment with PD98059, cells were treated with either vehicle or AE (60 µM) for 24 h.
(A) The levels of ERK, p-ERK (Tyr202/204), p21, and E2F1 in the cell lysates were then
determined by western blotting using specific antibodies. γ-Tubulin was used as an internal
control for sample loading. (B) The cell cycle profile and fraction of apoptotic cells (sub-G1
phase population) were determined using flow cytometry. The values presented are the mean
± standard error from three independent experiments.
Fig. 5. Changes in Bcl-XL and Bax expression are involved in AE-induced apoptosis. (A) The
effect of AE on the expression of Bcl-2 family proteins. The levels of Bcl-XL, Bcl-2, Bax, Bak,
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Bid, and tBid in cell lysates prepared after 36 h of treatment with vehicle or AE (60 µM) were
analyzed using specific antibodies (panels I and II). γ-Tubulin was used as an internal control
for sample loading. (B) Ectopic expression of Bcl-XL suppresses AE-induced apoptosis. At 12
for sample loading. (B) Ectopic expression of Bcl-XL suppresses AE-induced apoptosis. At 12