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Caspase-2 is required for DNA damage-induced
expression of the CDK inhibitor p21WAF1/CIP1
Dennis Sohn, Wilfried Budach and Reiner U. Jänicke*
Laboratory of Molecular Radiooncology, Clinic and Policlinic for Radiation Therapy and
Radiooncology, University of Düsseldorf, Universitätsstrasse 1,
40225 Düsseldorf, Germany
*Corresponding author:
Reiner U. Jänicke, Laboratory of Molecular Radiooncology
University of Düsseldorf, Building 23.12
Universitätsstrasse 1, D-40225 Düsseldorf, Germany
Phone: +49-211-8115973, Fax +49-211-8115892, e-mail: janicke@uni-duesseldorf.de
Character count (excluding Materials/Methods and References): 39,372
Running title: Caspase-2-dependent p21 expression
Key words: apoptosis, cell cycle, translational control, 3’-UTR, p53, RAIDD
peer-00630274, version 1 - 8 Oct 2011
Author manuscript, published in "Cell Death and Differentiation (2011)"
DOI : 10.1038/cdd.2011.34
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ABSTRACT
Although caspase-2 represents the most conserved caspase across species and was the
second caspase identified, its precise function remains enigmatic. In several cell types we
show that knockdown of caspase-2 specifically impaired DNA damage-induced p21
expression, while overexpression of a caspase-2 mutant increased p21 levels. Caspase-2 did
not influence p21 mRNA transcription, and also various inhibitors targeting proteasomal or
non-proteasomal proteases including caspases could not restore p21 protein levels following
knockdown of caspase-2. As, however, silencing of caspase-2 only impaired exogenous
expression of p21 constructs containing 3’-UTR sequences, our results strongly indicate that
caspase-2 regulates p21 expression at the translational level. Intriguingly, unlike depletion of
caspase-2 that prevented p21 expression and thereby reverted the γ-irradiation-induced
senescent phenotype of wild-type HCT116 colon carcinoma cells into apoptosis, neither
knockdown of the caspase-2-interacting components RAIDD, RIP or DNA-PKcs was able to
mimic these processes. Together, our data suggest that this novel role of caspase-2 as a
translational regulator of p21 expression occurs not only independently of its enzymatic
activity, but does also not require known caspase-2-activating platforms.
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INTRODUCTION
Although caspases were initially identified as key apoptotic enzymes, they are now
known to also exert essential functions in diverse cellular processes including inflammation,
differentiation and proliferation.1 Among them, caspase-2 is not only the most conserved
caspase across species, but intriguingly also the most enigmatic, as it combines features of
both initiator and executioner caspases.2,3 It contains a long caspase activation and
recruitment domain (CARD), a characteristic trait of initiator caspases required for their
activation, while its cleavage specificity resembles that of effector caspases. Similar to
initiator caspases such as caspase-8, -9 and –10, caspase-2 is activated by dimerization via a
so-called “induced-proximity”-mechanism, although cleavage of caspase-2 was found to
stabilize the active protease.4,5 In contrast, as caspase-2 is a poor activator of effector
caspases, it surely lacks an essential feature of initiator caspases. Thus, not unexpectedly,
contradictory findings have been reported that place caspase-2 either upstream or
downstream of mitochondria, or even fail to assign any specific function to this enzyme in
apoptosis.6,7
In sharp contrast to the in utero or perinatal lethality of Casp88 and Casp9-knockout
mice,9,10 Casp2-knockout mice were found to be viable and develop normally, and
thymocytes from these animals show no obvious apoptosis defects compared to their wild-
type littermates.11,12 Female Casp2-deficient animals, however, display a slight increase in
their oocyte numbers probably due to an impaired apoptosis program,11 and mice lacking
caspase-2 show signs of premature ageing.13 Also other Casp2-deficient cell types such as
neurons or embryonic fibroblasts (MEFs) show a reduced or delayed apoptotic response
toward certain stimuli,6,7 indicating both redundant and non-redundant functions for caspase-
2 in apoptosis. In addition caspase-2 was reported to be involved in the regulation of DNA
damage responses and cell cycle progression and may even act as tumor suppressor.14,15
However, the underlying mechanisms are completely unresolved mainly due to the fact that
only few caspase-2-specific substrates have yet been identified.
Unlike caspase-8 and -9 that become activated in the death-inducing signaling
complex (DISC) and the apoptosome, activation of caspase-2 is achieved via formation of
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three functionally different multi-protein complexes. The first caspase-2-activating complex
identified was the so-called RAIDD-PIDDosome consisting of RAIDD [receptor-interacting
protein (RIP)-associated ICH/CED-3-homologous protein with death domain], PIDD (p53-
induced protein with death domain), and caspase-2.16 The RAIDD-PIDDosome is formed in
response to genotoxic stressors and induces apoptosis most likely via caspase-2-mediated
BID cleavage.17 However, caspase-2 was also found processed and capable of inducing
apoptosis in the absence of RAIDD or PIDD, suggesting the existence of additional caspase-
2-activating platforms.18 Thereby, caspase-2 was proposed to form a cytosolic complex with
RIP and TRAF-2 (TNF receptor-associated factor-2) that, however, does not induce
apoptosis, as it strongly activates NF-κB and p38.19 Surprisingly, this was independent of the
proteolytic activity of caspase-2, and instead required only the prodomain containing the
CARD oligomerization motif.
In contrast to these cytosolic complexes, a third caspase-2-activating complex formed
in the nucleus was implicated in the maintenance of a G2/M DNA damage checkpoint and in
DNA repair executed by the NHEJ pathway.14 Thereby, the DNA-dependent serine/threonine
protein kinase (DNA-PKcs) and PIDD form the backbone of the so-called DNA-PKcs-
PIDDosome that is constitutively present even in unstimulated cells. Following DNA
damage, caspase-2 is recruited and activated in this complex via the DNA-PKcs-mediated
phosphorylation of its Ser122 residue. Even though the identification of the DNA-PKcs-
PIDDosome resolved another enigma surrounding caspase-2, namely its constitutive nuclear
localization, it remains unanswered of how nuclear caspase-2 is integrated in the DNA
damage-induced G2/M checkpoint and NHEJ pathways.
Although originally identified as a cyclin-dependent kinase (CDK) inhibitor
mediating p53-induced cell cycle arrest, p21 is now known to also participate in diverse
biological processes including transcription, DNA repair, differentiation, senescence and
apoptosis.20-22 Thus, mechanisms are required that tightly control expression and functional
diversity of p21. This is achieved for instance by multiple phosphorylations that either result
in its stabilization or an increased degradation by the proteasome.23 In addition to this post-
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translational regulation, expression of p21 is also controlled at the transcriptional and post-
transcriptional level,24,25 giving the cell multiple opportunities to interfere with p21 function.
Here we uncovered an unprecedented role of caspase-2 as an essential translational
cofactor for DNA damage-induced p21 expression. This novel role that enables caspase-2 to
control the diverse activities of p21 and that does not depend on its activation in known
caspase-2-activating platforms was observed in different human cell types independently of
the kind of DNA damage applied. Thus, our findings add a new level of complexity to the
enigma surrounding the role(s) of caspase-2 and establish a novel apoptosis-independent
involvement of this protease in cellular DNA damage signaling pathways.
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RESULTS
Caspase-2 knockdown sensitizes wild-type HCT116 cells to
γ
IR-induced apoptosis by
reducing p21 protein levels.
Recently we reported that besides the induction of p53-dependent senescence, p21 is
also required for the simultaneous inhibition of caspase activation and apoptosis downstream
of mitochondria.26 This conclusion was drawn from our finding that only checkpoint-deficient
HCT116 cells (p21-/- and p53-/-) succumb to apoptosis upon exposure to ionizing radiation
(γIR), whereas similarly treated wild-type cells are driven into senescence. To verify that the
hypersensitivity of p21-deficient HCT116 cells is indeed due to the loss of p21 and not caused
primarily by the elevated levels of p53 and Puma (Fig. 1A),26 we silenced p21 expression in
wild-type cells by RNA interference. Knockdown of p21, but not a control siRNA reverted
the senescent phenotype of irradiated wild-type HCT116 cells and sensitized them to
apoptosis as evidenced by the processing and activation of caspase-9 and -3 (Fig. 1B,C).
More interestingly, following γIR exposure procaspase-2 levels decreased in a time-
dependent manner in both HCT116 lines regardless of whether or not they succumbed to
apoptosis (Fig. 1A,B), implying processing of caspase-2 (please see discussion for further
details). As in contrast, procaspase-9 and -3 are processed and activated exclusively in
apoptotic checkpoint-deficient cells, and because γIR induces activation of mitochondria even
in HCT116 wild-type cells that become senescent upon this treatment,26 our data imply an
important upstream role for caspase-2 in DNA damage signaling pathways.
To more closely examine this role, we compared the fate of irradiated HCT116 wild-
type cells in the presence and absence of caspase-2. Surprisingly, siRNA-mediated silencing
of caspase-2 reverted the senescent phenotype of irradiated wild-type cells and resulted in the
processing and activation of caspase-3 as well as in the death-associated release of the lactate-
dehydrogenase (LDH) (Fig. 2A-C). This phenomenon could not be explained by an elevated
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expression of p53 or Puma, because their γIR-induced protein levels did not differ
significantly in the absence or presence of caspase-2 (Fig. 2A). In contrast, wild-type cells
sensitized to γIR-induced apoptosis by knockdown of caspase-2 displayed a remarkable
strong decrease in p21 protein expression (Fig. 2A). This was also observed when individual
siRNAs were used that comprise the caspase-2 SMARTpool siRNA ruling out off-target
effects (Suppl. Fig. 1). On the other hand, p21 levels increased following overexpression of
caspase-2, however, only in unstressed cells and when a catalytically inactive caspase-2
mutant was used (Fig. 2D). This most likely reflects the facts that irradiation per se induces
the maximum level of p21 expression in HCT116 wild-type cells that cannot be further
enhanced by exogenous caspase-2 and that cells transfected with the wild-type enzyme
undergo massive apoptosis (not shown) explaining not only its weak detection, but also the
minor effect on p21 expression. Together, these results are consistent with our previous
observation that p21 protects HCT116 wild-type cells from γIR-induced apoptosis,26 and in
addition suggest that caspase-2 modulates DNA damage responses also independently of its
catalytic activity by positively regulating p21 expression.
Caspase-2 is commonly required for DNA damage-induced p21 expression.
To verify that the loss of caspase-2 sensitizes HCT116 wild-type cells toward γIR-
induced apoptosis via suppression of p21, we compared the effects of the caspase-2
knockdown on apoptotic responses of wild-type and p21-deficient HCT116 cells. Silencing of
caspase-2 increased the amount of γIR-induced caspase-3-like DEVDase activity in wild-type
cells to a comparable level detected in untransfected γIR-treated p21-/- cells (Fig. 3A).
Irradiated p21-/- cells on the other hand were not further radio-sensitized by this measure
(Fig. 3A), although a comparable caspase-2 knockdown was achieved as in wild-type cells
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(Fig. 3B). These data demonstrate that suppression of p21 is an integral part of the mechanism
by which caspase-2 deficiency increases the radio-sensitivity of HCT116 wild-type cells.
To determine whether caspase-2-dependent p21 expression is confined to γIR-induced
signaling, we treated cells with the topoisomerase-II inhibitor etoposide that also resulted in
an efficient processing of procaspase-2 (Fig. 3B). Consistently, the absence of caspase-2 in
etoposide-treated wild-type cells was also accompanied by a marked reduction of p21,
whereas expression of p53 and Puma remained unaffected (Fig. 3B). However, despite the
significant reduction in p21, only a marginal increase of DEVDase activity was observed in
etoposide-treated caspase-2-depleted wild-type cells when compared with similarly treated
untransfected cells or with cells transfected with a control siRNA (Fig. 3C). The failure of p21
to affect etoposide-induced apoptosis is most likely explained by our observation that wild-
type and p21-/- HCT116 cells display similar apoptosis sensitivities toward various
chemotherapeutic drugs including etoposide (not shown), suggesting that p21 is not sufficient
to protect the cells under those conditions.
Having established the consequences of a caspase-2 loss in wild-type and p21-/-
HCT116 cells exposed to γIR and etoposide, we analyzed whether this applies also to other
cell types. Therefore, MCF-7/Casp3 breast carcinoma cells that like HCT116 wild-type cells
also undergo senescence upon exposure to γIR,27 were transfected with control or caspase-2
siRNAs. Also in these cells, procaspase-2 was readily processed following γIR exposure (Fig.
4A) despite their apoptosis resistance toward this treatment.27 More importantly, they too
displayed an almost complete loss of the γIR-induced p21 expression following caspase-2
knockdown, whereas expression of p53 and PUMA remained unchanged (Fig. 4A). Similar
results were obtained with etoposide-treated MCF-7/Casp-3 cells (Fig. 4C) or when we
exposed caspase-2-depleted primary normal human dermal fibroblasts (NHDF) to γIR (Fig.
4E). In contrast to wild-type HCT116 cells, however, knockdown of caspase-2 could not alter
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their apoptosis susceptibilities (Fig. 4B, D and data not shown), suggesting that other
mechanisms besides p21 determine their response toward these treatments. Nevertheless, our
results demonstrate that expression of p21 commonly requires the presence of caspase-2.
Caspase-2 is essential for DNA damage-induced checkpoint control.
Next we analyzed the proliferative capability of HCT116 cells in the presence and
absence of caspase-2. Wild-type and p21-deficient HCT116 cells were transfected with
control or caspase-2 siRNA and incubated for four hours with BrdU two days following their
exposure to γIR in order to label cells actively synthesizing DNA. As expected, regardless of
the presence or absence of caspase-2, p21-deficient cells were unable to undergo γIR-induced
cell cycle arrest, as similar numbers of BrdU-positive cells were detected in irradiated and
non-irradiated samples (Fig. 5A). In contrast, untransfected or control siRNA-transfected
wild-type cells displayed significantly lower numbers of BrdU-stained cells post γIR (Fig.
5A), demonstrating that the observed cell cycle arrest of wild-type cells depends mainly on
the presence of p21. Consistently, diminishing p21 levels by the caspase-2 siRNA increased
the number of irradiated wild-type cells that are capable of incorporating BrdU (Fig. 5A).
Remarkably, similar to irradiated p21-deficient cells, irradiated wild-type cells
expressing the caspase-2 siRNA harbour extremely aberrant shaped nuclei that are
characteristic for cells lacking an important cell cycle checkpoint and hence, try to enter
mitosis despite an extensive damage to their DNA (Fig. 5B). As such an abnormal nucleus
structure was not observed in irradiated wild-type cells that were left untransfected or
transfected with the control siRNA (Fig. 5B), these data demonstrate that caspase-2 is crucial
for an orderly executed stress response pathway upon DNA damage.
Caspase-2 modulates p21 expression at the translational level.
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Several mechanisms could account for the observed decrease of p21 protein following
knockdown of caspase-2 including degradation by the proteasome, cleavage by proteases
including caspases or transcriptional repression of the CDKN1A gene encoding p21.20,23,28
Therefore, we first asked whether protease-mediated cleavage of p21 contributes to the
observed loss. However, irradiating caspase-2-depleted wild-type cells in the presence of the
pan-caspase inhibitor Q-VD-OPh could not prevent p21 decrease (Fig. 6A), demonstrating
that this event is not caused by caspase cleavage. In addition, Q-VD-OPh failed to inhibit
procaspase-2 processing in irradiated control cells, and was completely ineffective in reducing
γIR-induced p21 levels in untransfected wild-type cells, or in cells transfected with the control
siRNA (Fig. 6A). Similar results were obtained with z-VDVAD-fmk (not shown), another
caspase inhibitor preferentially targeting caspase-2. Based on the effectiveness of Q-VD-OPh
and z-VDVAD-fmk to completely suppress caspase-3 processing and activity (Suppl. Fig. 2,
and data not shown) and our finding that a catalytically inactive caspase-2 mutant
substantially increased p21 expression (Fig. 2D), these data strongly suggest that caspase-2
modulates p21 expression independently of its enzymatic activity.
Next we employed the proteasomal inhibitor MG-132 that due to its high cytotoxicity
in HCT116 cells was only applied for a maximum of eight hours post γIR, a time point clearly
sufficient to impair p21 expression in the absence of caspase-2 (Fig. 6B; compare lanes 2/6
with 10). Although MG-132 caused, as expected, a rapid accumulation of p53 and p21 even in
the absence of γIR, it was unable to restore p21 protein levels of non-irradiated and irradiated
caspase-2 knockdown cells to those observed in control siRNA-transfected or untransfected
cells (Fig. 6B, compare lanes 3/4 and 7/8 with 11/12). Similar results were obtained using
other inhibitors targeting either the proteasome (calpain inhibitor-I), calpains and cathepsins
(calpain inhibitor-II), or interfering with lysosomal acidification such as ammonium chloride
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(not shown). Thus, neither caspases nor any other proteases investigated are involved in the
reduction of p21 following caspase-2 knockdown.
Therefore, we analyzed whether p21 transcription is influenced by the presence or
absence of caspase-2. However, performing Real-Time-PCR analyses for p21 and Puma
mRNA, no significant caspase-2-dependent differences were observed in their control or γIR-
induced expression levels. Although both mRNA species were efficiently upregulated in wild-
type cells exposed to γIR, the knockdown of caspase-2 did not interfere with these processes
(Fig. 6C). Therefore, our data argue against a direct off-target effect of the caspase-2 siRNAs
on the p21 mRNA and additionally imply that caspase-2 controls p21 expression at the
translational level.
As sequences in the 5’- and 3’-untranslated regions (UTR) are important regulators of
protein translation,29 wild-type cells were transfected with Flag-tagged p21 cDNA constructs
containing either 5’- or 3’-UTR sequences or both or with a construct that only consists of the
p21 coding region. Remarkably, although the two Flag-p21 proteins harbouring 3’-UTR
sequences were less well expressed than the p21 constructs that contain no UTRs or only the
5’-UTR, knockdown of capase-2 further compromised their expression in stressed and
unstressed cells (Fig. 6D). In contrast, expression of p21 constructs that lack both UTRs or
only include the 5’-UTR was not affected in either condition and remained unaltered even in
the absence of caspase-2. Obviously, when compared to the endogenous p21, expression of
the exogenously expressed Flag-p21-3’-UTR construct was less affected by the depletion of
caspase-2. This is most likely due to an important technical difference in the setup of these
two experiments. Whereas the endogenous p21 was only induced (e.g. by γ-IR) at a time point
at which caspase-2 was already depleted from the cells (48 hours post transfection with the
caspase-2 siRNA), the exogenous Flag-p21-UTR constructs were simultaneously
cotransfected with the caspase-2 siRNA, as two individual successive transfections proved to
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be extremely detrimental to the cells. As the siRNA-mediated knockdown of caspase-2
requires up to 48 hours, this cotransfection procedure denotes that transcription and
translation of the exogenous Flag-p21 constructs will be initially supported by the presence of
caspase-2. Thus, expression of exogenous Flag-p21 will only be impaired at a later time point
at which caspase-2 is sufficiently down regulated by the siRNA. Furthermore, together with
the observation that depletion of caspase-2 resulted also in a severe reduction of luciferase
activity when the p21 3’-UTR was cloned downstream of the luciferase cDNA42 (Fig. 6E),
our results provide strong evidence that caspase-2 controls p21 expression at the translational
level by an as yet unknown mechanism that involves 3’-UTR sequences of p21.
p53 critically determines the role of caspase-2 in DNA damage responses.
Caspase-2 can be recruited to three functionally different multi-protein complexes; the
RAIDD-PIDDosome,16 the DNA-PKcs-PIDDosome,14 and to a complex consisting of RIP
and TRAF2.19 To elucidate whether caspase-2 requires association with any of these
complexes in order to regulate p21 expression in wild-type cells, we silenced a specific
component of each using siRNAs. Although a sufficient knockdown was achieved for DNA-
PKcs, RIP and RAIDD (Fig. 7A), none of these prevented processing of procaspase-2 (Fig.
7B), or resulted in a decrease of p21 as it was observed following caspase-2 knockdown (Fig.
7A). Consistently, only irradiated wild-type cells transfected with caspase-2 siRNA showed
an increase in DEVDase activity (Fig. 7C), indicating that none of the known complexes are
involved in this novel function of caspase-2.
Intriguingly, a completely different picture emerged when we performed similar
experiments with cells lacking p53. In sharp contrast to the responses of irradiated wild-type
and p21-deficient HCT116 cells, caspase-2 knockdown rendered p53-deficient cells resistant
toward γIR-induced apoptosis and significantly inhibited DEVDase activity (Fig. 8B).
Moreover, siRNA-mediated inhibition of RAIDD expression, but not that of RIP or DNA-
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PKcs (Fig. 8A), resulted in a similar reduction of γIR-induced DEVDase activity in these cells
as observed following caspase-2 knockdown (Fig. 8B). Together with the observation that
only RAIDD deficiency, but not that of RIP or DNA-PKcs prevented processing of
procaspase-2 in p53-deficient cells (Fig. 8A), these data indicate that in the absence of p53,
caspase-2 signals apoptosis most likely via formation of the RAIDD-PIDDosome.
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DISCUSSION
We describe an unprecedented role of caspase-2 as an essential cofactor for the
efficient DNA damage-induced expression of the CDK inhibitor p21 that is evident in
different cell types independently of the kind of DNA damage applied. This function might
allow caspase-2 to participate in diverse signaling pathways modulated by p21 including cell
cycle control, gene transcription, DNA repair, differentiation, and apoptosis.20-22 Consistently,
we observed that the diminished p21 protein pool due to the knockdown of caspase-2 resulted
in an increased proliferation rate of irradiated HCT116 wild-type cells, a phenomenon
recently reported also with caspase-2-deficient MEFs.15
Our data also suggest that it is the p21-modulating capability, rather than its apoptosis-
inducing potential, that constitutes the primary function of caspase-2, at least within the
cellular context examined here. This assumption is based on the observation that knockdown
of caspase-2 was not sufficient to protect several p53-expressing cells from DNA damage-
induced apoptosis. Even the radio-sensitizing effect observed in caspase-2-depleted wild-type
HCT116 cells is probably not caused by a direct influence of caspase-2 on apoptosis
signaling, as it rather appears to be a consequence of p21 downregulation. As this
sensitization occurs only in cells susceptible to the anti-apoptotic function of p21 such as
irradiated HCT116 wild-type cells,26 our data are consistent with a previous report
demonstrating that p21 is not the only determinant in stress-induced p53 responses.30
Surprisingly, however, and in contrast to other reports demonstrating a p53 requirement for
caspase-2 activation and apoptosis induction,31,32 the pro-apoptotic function of caspase-2
unveiled in our study only upon loss of p53. Thus, our data suggest that caspase-2 is
additionally part of an apoptotic backup system that signals apoptosis only in the absence of a
functional p53 pathway. Although cell type and stimulus-dependent differences clearly
contribute to this controversial issue,31 our view is further supported by the observation that
knockdown of RAIDD, but not that of RIP or DNA-PKcs, successfully abrogated DNA
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damage-induced processing and signaling of caspase-2, however, only in p53-deficient
HCT116 cells. On the other hand, DNA damage-induced caspase-2 processing and
modulation of p21 expression was not affected in wild-type cells by knockdown of any of
these components suggesting the existence of additional caspase-2-activating platforms.
Consistently, although caspase-2 was found by others to be associated with the RAIDD-
PIDDosome even in unstressed wild-type and p53-/- HCT116 cells, silencing of RAIDD and
PIDD also failed to prevent caspase-2 activation and apoptosis in 5-FU-treated HCT116 wild-
type cells.31 Thus, further studies are required to shed some light on this unresolved issue.
In this context, the most important questions remaining are: how does caspase-2 affect
expression of p21 and does this process require a processed and active caspase-2 enzyme? A
variety of mechanisms known to regulate p21 expression at the protein and DNA/RNA level
have been elucidated in recent years.20,23,24 However, post-translational regulation could be
excluded here, as various inhibitors targeting proteasomal or other proteases could not restore
p21 levels following caspase-2 knockdown. Also two different caspase inhibitors (Q-VD-
OPh, z-VDVAD-fmk) were despite their effectiveness in completely blocking γIR-induced
caspase-3 processing and activity in p53-deficient cells (Suppl. Fig. 2 and data not shown),
unable to prevent loss of p21. Combined with our finding that overexpression of a
catalytically inactive caspase-2 mutant was sufficient to increase p21 expression in unstressed
wild-type cells, these data strongly suggest that the enzymatic activity of caspase-2 is
dispensable for this event. A similar conclusion was reached with regard to its processing, as
impaired p21 expression as a consequence of caspase-2 depletion was already observed eight
hours post γIR, a time point at which the decrease of pro-caspase-2 was not even evident in
irradiated control cells. However, in regard to this, it is presently not entirely clear whether
the observed γIR-induced decrease of pro-caspase-2 reflects indeed its processing, as both
caspase inhibitory peptides neither compromised this process nor the concomitant generation
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of mature caspase-2 fragments (Suppl. Fig. 2). Using two different caspase-2 antibodies, the
mature large caspase-2 subunit was only generated in irradiated p53- and p21-deficient
HCT116 cells, whereas the appearance of a fragment probably resembling the processed
prodomain was also detected in wild-type cells. As this fragment might have also arisen from
an alternative splicing event,33 it remains possible that the decrease of caspase-2 in wild-type
cells does not reflect its processing, but rather a p53-mediated suppression as reported
recently.34 However, Real-Time PCR analyses revealed comparable amounts of caspase-2
mRNA transcripts in unstressed and irradiated wild-type and p53-deficient HCT116 cells (not
shown) clearly arguing against this possibility.
While therefore a direct intervention of caspase-2 with p53-mediated p21 transcription
seemed reasonable to assume, particularly in view of the observed interaction of p53 with
caspase-2 (not shown), irradiated caspase-2-depleted cells exhibited only a slight decrease in
p21 mRNA transcripts. Together with our finding that the lack of caspase-2 affects exogenous
expression only of those p21 constructs containing 3’-UTR sequences, these results clearly
argue against a direct effect of caspase-2 on p21 transcription. In addition, as expression of
these 3’-UTR-containing p21 constructs is even severely compromised in the presence of
caspase-2, we rather favour a model in which caspase-2 controls p21 expression at the
translational level involving 3’-UTR sequences. As caspase-2 does not harbour an obvious
RNA-binding domain, we would like to suggest an indirect mechanism by which caspase-2
affects p21-mRNA translation, perhaps via association with RNA-binding proteins or
modulation of related pathways. Thereby, we can surely exclude participation of certain
RNA-binding proteins such as members of the Hu family that were shown to interact and
stabilize p21 mRNA.35 Also CUGBP1 and calreticulin, two RNA-binding proteins that
promote and inhibit p21 translation are most likely not involved, as they compete with each
other for the binding to the same nucleotide sequence within the 5´-region of the p21
mRNA.36 Our results leave open, however, the possibility that caspase-2 modulates the
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function of a different class of RNA-binding proteins known to stimulate or inhibit mRNA
translation without interfering with their stabilities. For instance, binding of musashi or the
heterogeneous nuclear ribonucleoprotein K (hnRNP-K) to the 3´-UTR of the p21 mRNA was
shown to efficiently inhibit its translation.37,38 However, whether these or any other RNA-
binding proteins are involved in the caspase-2-modulated p21 expression remains to be
investigated.
Another possible mechanism by which caspase-2 may modulate p21 translation
involves repression of certain microRNAs known to prevent p21 mRNA translation without
interfering with its stability.39 Although there might exist an almost overwhelming number of
at least 100 microRNAs that theoretically could target the p21 mRNA (as predicted from
microRNA.org), we have analyzed the expression levels of miR-17, miR-20a, and miR-106b,
because these were not only shown to efficiently block p21 mRNA translation,40-42 but were
also found to be repressed by p53.43 However, expression of these microRNAs did not change
16 hours following exposure of HCT116 wild-type cells to γIR, and also remained unaltered
in the presence or absence of caspase-2 (not shown). In contrast, the p53-inducible miR-34a
that is involved in the induction of apoptosis and senescence44 was found upregulated
following γIR, however, in a caspase-2-independent manner (not shown). Although these
observations clearly argue against an involvement of these microRNAs in the caspase-2-
dependent expression of p21, the possibility remains that other microRNAs contribute to this
process.
In summary, we have uncovered an all new function for caspase-2 as an essential
cofactor for DNA damage-induced p21 expression. Certainly, this finding entails far-reaching
implications, as the translational regulation of p21 enables caspase-2 to participate in the
diverse signaling processes controlled by this CDK inhibitor. Now it will be challenging to
identify the components involved and to elucidate the mechanism of how exactly caspase-2
regulates p21 translation.
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ACKNOWLEDGEMENTS
We are grateful to B. Vogelstein and A. Villunger for the HCT116 cell lines and the caspase-2
plasmids and C. Disselhoff for excellent technical assistance. The luciferase cDNA constructs
with or without the p21-3’-UTR were generous gifts from X. He. This work was supported by
grants from the Deutsche Forschungsgemeinschaft (SFB 728) and the Forschungskommission
of the Heinrich-Heine-University of Düsseldorf.
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MATERIALS AND METHODS
Cell lines, reagents and antibodies. HCT116 wild-type cells and their checkpoint-deficient
variants (p53-/- and p21-/-) were maintained in McCoy’s 5A medium (PromoCell,
Heidelberg, Germany), whereas MCF-7/casp3 cells were cultured in RPMI 1640 (PAA
Laboratories, Linz, Austria) in the presence of 400 µg/ml neomycin. The primary normal
human dermal fibroblasts (NHDF) were obtained from PromoCell and were cultured in
DMEM low glucose supplied with 1% MEM nonessential amino acids solution and 0.1 mM
β-mercapto-ethanol. Only passages below P20 were used. All media were supplemented with
10% heat-inactivated fetal calf serum, 10 mM glutamine, 100 U/ml penicillin and 0.1 mg/ml
streptomycin (PAA Laboratories). The pan-caspase inhibitory peptide Q-VD-OPh (Q-Val-
Asp-CH2-O-Ph) as well as the caspase-2 inhibitory peptide z-VDVAD-fmk (z-Val-Asp-Val-
Ala-Asp-fluoromethyl-ketone) were from MP Biomedicals (Irvine, CA, USA). The
fluorometric caspase-3 substrate DEVD-AMC (N-acetyl-Asp-Glu-Val-Asp-
aminomethylcoumarin) was from Biomol (Hamburg, Germany). The polyclonal antibodies
against caspase-3 and caspase-9 were from R&D Systems (Wiesbaden, Germany) and from
Cell Signaling Technology (Danvers, MA, USA), respectively. The rat caspase-2 mAb was
from Alexis Biochemicals (Lausen, Switzerland). The p53 monoclonal Ab-6 antibody and the
polyclonal antibody for PUMA were from Calbiochem (Bad Soden, Germany), whereas the
mAbs recognizing p21 and RIP were from BD Biosciences (Heidelberg, Germany). The mAb
towards RAIDD was from MBL International (Woburn, MA, USA). From Santa Cruz
(Heidelberg, Germany) was the polyclonal antibody recognizing the catalytic subunit of
DNA-PK. The actin mAb, the nuclear stain 4’,6-diamidino-2-phenylindole (DAPI), the
topoisomerase II inhibitor etoposide, the two calpain inhibitors I (ALL-N) and II (ALL-M)
and the protease inhibitors PMSF, aprotinin, leupeptin and pepstatin were from Sigma-
Aldrich (Deisenhofen, Germany). From Enzo Life Sciences (Biozol; Eching, Germany) we
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obtained the proteasome inhibitor MG-132, and the peroxidase-labeled secondary antibodies
were from Promega GmbH (Mannheim, Germany).
Treatment of cells and measurement of cell death. Cells were exposed to γIR (routinely 20
Gy at 200 kV) using a Gulmay RS 225 X-ray system from IsodoseControl (Bochum,
Germany) or were treated with etoposide (50 µM) for the indicated times. Cell death was
assessed either microscopically or by determination of lactate dehydrogenase (LDH) activity
in supernatants of 105 cells according to the protocol of the manufacturer (Roche Molecular
Biochemicals, Mannheim, Germany). The values obtained are given in arbitrary units (AU).
Preparation of cell extracts and Western blotting. Total cell extracts were prepared in lysis
buffer (50 mM Tris/HCl pH 7.4, 150 mM NaCl, 1% NP-40) containing protease inhibitors as
described.26 Protein concentrations were determined with the BioRad protein assay, followed
by separation of the extracts in SDS-polyacrylamide gels and electroblotting onto
polyvinylidene difluoride membranes (Amersham Biosciences, Braunschweig, Germany).
Following antibody incubation, the proteins were visualized by enhanced chemiluminescent
staining using ECL reagents (Amersham Biosciences).
Fluorometric determination of caspase-3-like-activity (DEVDase assay). For the detection
of caspase-3-like activities, 50 μg of the cell extracts were incubated for 3 to 4 hours with 50
μM of the caspase-3 substrate DEVD-AMC in 200 μl buffer containing 50 mM HEPES (pH
7,4), 100 mM NaCl, 10% sucrose, 0.1% CHAPS and 10 μM DTT. The release of fluorogenic
AMC was measured at an excitation wavelength of 346 nm and an emission wavelength of
442 nm using an Infinite M200 microplate reader (Tecan, Langenfeld, Germany). The
detected fluorometric signal that directly correlates to the caspase activity in the cell extracts
is expressed in arbitrary units (AU).
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Plasmids and generation of the different p21-UTR constructs: The plasmids encoding Flag-
tagged wild-type caspase-2 or an inactive mutant (C320G) (pEF-3x-Flag-Casp2-WT and –
MUT) were generous gifts from A. Villunger (Innsbruck, Austria). The plasmids pCEP-
Waf145 and pMT5-Flag-21-WT46 were purchased from Addgene (Cambridge, MA, USA) and
served as templates for cloning of the different p21-UTR constructs that all contain a N-
terminal Flag epitop (see below). The open reading frame of Flag-p21 was amplified from the
pMT5-Flag-p21-WT plasmid using primers that harbour BamHI and EheI restriction sites at
the 5´- end and SacII and EcoRI sites at the 3´-end. Following digestion with BamHI and
EcoRI, the amplified sequence (~ 650 bp) was cloned into the pcDNA4-Myc-His-A vector
(Invitrogen) yielding the pcDNA4-Flag-p21 plasmid. The 5´-UTR of p21 (~ 80 bp) was
amplified from the pCEP-WAF1 plasmid using primers with a HindIII-site at the 5´-end and
an EheI-site at the 3´-end. After digestion with HindIII and EheI, this amplificate was cloned
into the pcDNA4-Flag-p21 plasmid resulting in the pcDNA4-5-UTR-Flag-p21 plasmid.
Similarly, the 3´-UTR of p21 (~ 1500 bp) was amplified via PCR from the pCEP-WAF1
plasmid with primers including SacII at the 5´-end and NotI at the 3´-end and subsequently
cloned into pcDNA4-Flag-p21 following digestion with these two enzymes, thereby
generating the pcDNA4-Flag-p21-3-UTR plasmid. To finally produce a plasmid expressing
the full length p21 mRNA, the 3´-UTR was cut out of the pcDNA4-Flag-p21-3-UTR plasmid
with SacII and NotI and cloned into the pcDNA4-5-UTR-Flag-p21 plasmid. The generated
plasmid was termed pcDNA4-5-UTR-Flag-p21-3-UTR.
Transfection of siRNAs and plasmids. ON-TARGETplus SMARTpool siRNAs were
purchased from Dharmacon RNA technologies (Lafayette, CO, USA) and the knockdown was
performed according to the instructions of the manufacturer. 24-48 hours post-transfection,
cells were harvested and divided equally to receive either no treatment or exposure to γIR or
etoposide. At the indicated time points following DNA damage, cells were harvested and
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directly analyzed by Western blotting and by the fluorometric caspase substrate assay for a
successful knockdown of the target protein and DEVDase activity, respectively. For the
simultaneous transfection of plasmids and siRNAs, the Dharmafect DUO transfection reagent
was used according to the manufacturers protocol. Briefly, HCT116 wild-type cells seeded in
6-well plates were transfected with 2 μg plasmid and 200 nmol siRNA using 4 μl
Dharmacfect Duo in 2 ml antibiotics-free medium. One day after transfection, the cells were
trypsinized, divided equally into two wells and directly γ-irradiated or left untreated as a
control. One day after irradiation the cells were harvested and analysed by Western Blot.
Real-Time PCR. Total RNA was isolated using the RNeasy Kit (Qiagen, Hilden, Germany)
according to the protocol of the manufacturer. Reverse transcription was achieved with the
High Capacity cDNA Kit from Applied Biosystems (Darmstadt, Germany). Taqman gene
expression probes for human p21, Puma and actin mRNA (Applied Biosystems) were
employed to analyse their relative expression levels using the 7300 Real-Time PCR system
(Applied Biosystems). The actin mRNA served as an endogenous normalization control for
every sample. The fold induction of the analyzed RNAs was calculated via the 2^(Δ(ΔCt)-
method thereby normalizing all samples to the level of the analyzed RNA in untransfected
control HCT116 wild-type cells.
Luciferase Reporter Assay. Cells were transfected with a vector containing luciferase cDNA
with or without the p21-3’-UTR42 together with either caspase-2 or control siRNA. Two days
after transfection, cells were harvested and divided into two aliquots. Luciferase activities
were determined by using the Dual luciferase activity kit (Promega) and a Centro LB 960
luminometer (Berthold Technologie, Bad Wildbad, Germany). For normalization of the
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obtained activities, Real-Time-PCR was performed with a specific firefly luciferase mRNA
probe (Custom Taqman Assay from Applied Biosystems).
BrdU labeling. Cells actively synthesizing DNA were labeled with the BrdU (5-Bromo-2'-
deoxy-uridine) Labeling and Detection Kit I from Roche Molecular Biochemicals according
to the manufacturers protocol. In short, following siRNA transfection, cells were seeded on 15
mm coverslips and irradiated. Two days post γIR, cells were labeled for four hours with BrdU
before they were fixed, incubated with a monoclonal antibody recognizing BrdU followed by
incubation with a fluorescence-coupled antibody directed against the primary antibody. Total
DNA was stained with DAPI before the coverslips were mounted on glass slides.
Subsequently, 6-10 representative pictures were taken from every sample using a 20x
objective from the Zeiss Axio Observer A1 and the corresponding AxioVision Software (Carl
Zeiss MicroImaging GmbH, Göttingen, Germany). The number of cells containing green
(BrdU-positive) and blue (DAPI) nuclei were counted from all representative pictures taken.
Employing this method, 500 to 1,200 blue cells were analysed for every condition in each
single experiment.
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LEGENDS TO FIGURES
Figure 1. Loss of p21 radiosensitizes HCT116 wild-type cells towards γIR-induced
apoptosis. (A) Western blot analyses for the status of the indicated proteins in HCT116 wild-
type and p21-/- cells that were γ-irradiated and cultured for the indicated days. (B) HCT116
wild-type cells were either left untransfected or transfected with control and p21 siRNAs.
Following γ-irradiation, cells were cultured for the indicated days before the extracts were
subjected to Western blot analyses. For A and B, blots shown represent results from two and
three independent experiments, respectively. (C) Determination of DEVDase activities in
HCT116 wild-type cells transfected and treated as described in B and analyzed after the
indicated days. Arbitrary units (AU) shown are the mean from three independent experiments
+/-SD.
Figure 2. Modulation of p21 expression by caspase-2. (A) Western blot analyses for the
status of the indicated proteins in HCT116 wild-type cells that were either left untransfected
or transfected with the caspase-2 siRNA before they were exposed to γIR and cultured for the
indicated days. (B) Determination of DEVDase activities in HCT116 wild-type cells that were
either left untransfected or transfected with control or caspase-2 siRNAs before they were γ-
irradiated and cultured for the indicated days. Please see Fig. 3A and 7C for additional
independent results. (C) Cell death assessment of HCT116 wild-type cells that were either left
untransfected or transfected with control or caspase-2 siRNAs before γ-irradiation. The
release of LDH into their supernatants was determined five days post γIR. Arbitrary units
(AU) shown in B and C are the mean from two independent experiments +/-SD. (D) Western
blot analyses demonstrating the influence of overexpression of a Flag-tagged caspase-2 wild-
type protein and a catalytically inactive caspase-2 mutant on p21 expression in control and γ-
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irradiated wild-type cells. Blots shown in A and D are representative results out of two and
three independent experiments, respectively.
Figure 3. DNA damage-induced p21 expression requires caspase-2. (A) Determination of
DEVDase activities in HCT116 wild-type and p21-/- cells that were either left untransfected
or transfected with the indicated siRNAs before they were γ-irradiated and cultured for two
days. (B) Western blot analyses for the status of the indicated proteins in HCT116 wild-type
and p21-/- cells that were either left untransfected or transfected with control or caspase-2
siRNAs before they were exposed to γIR or etoposide. Cells were harvested 36 hours
(etoposide) or 48 hours (γIR) post treatment. Blots shown represent results from two
independent experiments. (C) Determination of DEVDase activities in HCT116 wild-type
cells that were either left untransfected or transfected with the control or caspase-2 siRNAs
before they were exposed to etoposide. Arbitrary units (AU) shown in A and C are the mean
from three independent experiments each +/-SD.
Figure 4. Caspase-2 is commonly required for p21 expression. (A, C, E) Western blot
analyses for the status of the indicated proteins in MCF-7/Casp3 cells and NHDFs that were
either left untransfected or transfected with control or caspase-2 siRNAs before γ-irradiation
(A, E) or etoposide treatment (C). Cells were harvested two (E) to three (A) days post γIR and
24 hours following etoposide. Blots shown represent results from three independent
experiments. (B and D) Determination of DEVDase activities in MCF-7/Casp3 cells that were
transfected and treated as described in A and C. Arbitrary units (AU) are the mean from three
independent experiments +/-SD.
Figure 5. Caspase-2-deficient HCT116 wild-type cells exhibit an increased proliferation
rate following γ-irradiation. (A) Determination of the percentage of BrdU-positive HCT116
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wild-type and p21-/- cells that were transfected with control or caspase-2 siRNAs before γ-
irradiation. BrdU and DAPI stainings were performed two days post γIR. Data shown are the
mean of two independent experiments +/-SD. (B) Representative micrographs of HCT116
wild-type and p21-/- cells that were transfected and treated as described in A. Please note that
an increased number of irradiated wild-type cells display an aberrant nuclear structure only
following knockdown of caspase-2. This phenomenon can be also observed in irradiated p21-
deficient cells, however, regardless of whether or not they were transfected with the caspase-2
siRNA.
Figure 6. Caspase-2 modulates p21 expression at the translational level. (A and B)
HCT116 wild-type cells were either left untransfected or transfected with control or caspase-2
siRNAs before they were γ-irradiated in the presence or absence of the pan-caspase-inhibitor
Q-VD-OPh (A) or after an 1 hour preincubation with the proteasomal inhibitor MG-132 (B).
Cells were harvested 2 days (A) or 8 hours (B) post γIR and their cellular extracts were
analysed by Western blotting for the expression of the indicated proteins. Blots shown
represent results from two (A) and three (B) independent experiments, respectively. (C) Real-
Time-PCR for the determination of the expression levels of the p21 and Puma mRNAs in
HCT116 wild-type cells that were either left untransfected or transfected with control or
caspase-2 siRNAs before γ-irradiation. Total RNA was isolated 4 hours post γIR and analyzed
with transcript-specific probes from Applied Biosystems. Data shown are the mean of three
independent experiments +/-SD. (D) Western blot analyses demonstrating the influence of a
caspase-2 knockdown on expression of the indicated Flag-tagged p21 constructs containing
either 5’- or 3’-UTRs or both in unstressed and irradiated HCT116 wild-type cells. As a
control a plasmid encoding only the Flag-p21 open reading frame was used. Representative
blots out of three independent experiments are shown. (E) Luciferase reporter assay showing
the influence of a caspase-2 knockdown on expression of luciferase cDNA constructs with or
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without the p21-3’-UTR. In each sample, the obtained luciferase activity was normalized to
the amount of luciferase mRNA. Shown is the percentage of luciferase activities obtained
from caspase-2 siRNA-treated cells compared to control siRNA-treated cells. Values are
derived from three independent experiments (+/- SD).
Figure 7. Caspase-2 modulates p21 expression independently of known caspase-2-
activating complexes. (A) Western blot analyses for the status of the indicated proteins in
HCT116 wild-type cells that were either left untransfected or transfected with the indicated
siRNAs before γ-irradiation. Cells were analyzed 8 hours post γIR. Blots shown represent
results from three independent experiments. The asterisk denotes a band of unknown origin
that was still detectable following DNA-PKcs knockdown serving as an additional loading
control. (B) To demonstrate caspase-2 processing in these cells, an event not evident 8 hours
post γIR (see A), HCT116 wild-type cells were treated as in A, but were analyzed for caspase-
2 processing three days post γIR. Similar results were obtained two days post γIR (data not
shown). (C) Determination of DEVDase activities in HCT116 wild-type cells that were
treated as described in A. Cells were analyzed 3 days post γIR. Arbitrary units (AU) shown
are the mean from three independent experiments +/-SD.
Figure 8. RAIDD and caspase-2 are required for γ-IR-induced apoptosis in p53-deficient
cells. (A) Western blot analyses for the status of the indicated proteins in HCT116 p53-/- cells
that were either left untransfected or transfected with the indicated siRNAs before they were
γ-irradiated. Cells were analyzed 3 days post γIR. Blots shown represent results from three
independent experiments. The asterisk denotes a band of unknown origin (see also figure
legend 7A). (B) Determination of DEVDase activities in HCT116 p53-/- cells that were
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treated as described in A. Cells were analyzed 3 days post γIR. Arbitrary units (AU) shown
are the mean from three independent experiments +/-SD.
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