3144
Research Article
Various p53 mutant proteins differently regulate the
Ras circuit to induce a cancer-related gene signature
Hilla Solomon1,*, Yosef Buganim1,*, Ira Kogan-Sakin1, Leslie Pomeraniec1, Yael Assia1, Shalom Madar1,
Ido Goldstein1, Ran Brosh1, Eyal Kalo1, Tsevi Beatus2, Naomi Goldfinger1 and Varda Rotter1,`
1
Department of Molecular Cell Biology, Weizmann Institute of Science, Rehovot 76100, Israel
Department of Materials and Interfaces, Weizmann Institute of Science, Rehovot 76100, Israel
2
*These authors contributed equally to this work
`
Author for correspondence ([email protected])
Journal of Cell Science
Accepted 13 February 2012
Journal of Cell Science 125, 3144–3152
2012. Published by The Company of Biologists Ltd
doi: 10.1242/jcs.099663
Summary
Concomitant expression of mutant p53 and oncogenic Ras, leading to cellular transformation, is well documented. However, the mechanisms
by which the various mutant p53 categories cooperate with Ras remain largely obscure. From this study we suggest that different mutant p53
categories cooperate with H-Ras in different ways to induce a unique expression pattern of a cancer-related gene signature (CGS). The DNAcontact p53 mutants (p53R248Q and p53R273H) exhibited the highest level of CGS expression by cooperating with NFkB. Furthermore, the Zn+2
region conformational p53 mutants (p53R175H and p53H179R) induced the CGS by elevating H-Ras activity. This elevation in H-Ras activity
stemmed from a perturbed function of the p53 transcription target gene, BTG2. By contrast, the L3 loop region conformational mutant
(p53G245S) did not affect CGS expression. Our findings were further corroborated in human tumor-derived cell lines expressing Ras and the
aforementioned mutated p53 proteins. These data might assist in future tailor-made therapy targeting the mutant p53–Ras axis in cancer.
Key words: Ras, Gain of function, Mutant p53
Introduction
The development of tumors is characterized by the accumulation
of mutations in tumor-suppressor genes and in proto-oncogenes.
Two of the most frequently mutated genes in human cancer are
the gene encoding the tumor-suppressor p53 (TP53) and the
proto-oncogene Ras (Bos, 1989; Hollstein et al., 1991).
p53 is a transcription factor that accumulates and is activated in
response to stress signals. Following its activation, p53 induces
processes such as cell cycle arrest, programmed cell death and
DNA repair, thereby, guarding the cell from malignancy (Levine,
1997). In most cancer cells the p53 pathway is defective, usually
because of genetic mutations within the p53 sequence (Guimaraes
and Hainaut, 2002). Most of the mutations in p53 are missense
mutations that reside within the p53 DNA binding domain (DBD),
causing an impaired binding of p53 to the DNA. The missense
mutations can be divided into two rudimentary sub-groups
according to their impact on the DBD folding. The first group
consists of DNA-contact mutations (e.g. R248, R273), which
affect amino acids that directly interact with DNA while retaining
wild-type p53 conformation. The second group includes p53
conformational mutations (e.g. R175, H179), which alter the
scaffold that orients the structure of the DNA-binding interface
(Cho et al., 1994). The observation that mutant p53 is accumulated
in tumors, raised the possibility that mutant p53 plays an active
role in the process of malignant transformation. Indeed, it is now
accepted that mutant p53 exerts its activity in either a dominantnegative manner, in which it inhibits the normal activity of the
wild-type p53, or by a gain-of-function mechanism, in which
mutant p53 undertakes new oncogenic activities. Several
mechanisms were suggested to account for mutant p53 gain of
function, such as transcriptional activation (e.g. MYC, MDR1,
NFkB2), transcriptional repression (e.g. ATF3, CD-95 and
MST1), unique interaction with specific DNA motives (MAR),
epigenetic modification and interactions with other proteins (e.g.
p63, p73, NFY) (Sigal and Rotter, 2000; Li and Prives, 2007;
Lozano, 2007; Weisz et al., 2007a; Brosh and Rotter, 2009;
Buganim and Rotter, 2009).
Another protein frequently manipulated by cancer cells is the
Ras proto-oncogene, which becomes hyperactive because of a
point mutation during tumor development. There are three pivotal
Ras genes in the human genome: HRAS, KRAS and NRAS. In
spite of their high similarity in sequence and function, H-Ras was
found to be a more potent mediator of cellular transformation in
fibroblast cells, whereas K-Ras and N-Ras are more active in
epithelial cells and hematopoetic cells, respectively (Maher et al.,
1995). Ras activation depends on its binding to GDP (the nonactive state) or GTP, which enables its active conformation
(Malumbres and Barbacid, 2003). Switching to the active state is
controlled by the guanine nucleotide exchange factor (RasGEF)
enzyme family, which facilitates the replacement of GDP by
GTP. Shifting to the non-active state is regulated by the GTPase
activating protein (RasGAP) enzyme family, which facilitates
GTP hydrolysis. When mutated, Ras is less sensitive to regulation
by RasGAPs and therefore remains constitutively active. Overactivated Ras causes uncontrolled proliferation, survival and
tumor aggressiveness by induction of signaling pathways such as
MAPK, PI3K and RALGDS (Downward, 2003).
Early studies in cancer research revealed that mutant p53
and Ras oncogene cooperate to induce cellular transformation
(Eliyahu et al., 1984; Parada et al., 1984). Later studies reported
Journal of Cell Science
Various roles of p53 mutants in cancer
several processes regulated by both p53 and Ras, such as cell
motility, proliferation and survival (Boiko et al., 2006; Song et al.,
2007; Xia and Land, 2007; Meylan et al., 2009). However,
hitherto, a mechanism underlying the cooperation between
the common forms of mutant p53 and activated Ras, has not
been addressed. Because p53 is commonly mutated during
carcinogenesis, and mutated forms of p53 further facilitate
transformation using a wide range of mechanisms (Buganim and
Rotter, 2009), we decided to focus on the role of the different
mutant p53 categories in the crosstalk with Ras.
Thus, the main motivation of this work was to identify the
specific molecular details that underlie the crosstalk between Ras
and the various mutant p53 categories.
Recently, we established an in vitro transformation model
and performed a wide genomic profiling to identify clusters of
genes that are related to the different steps of transformation
(Milyavsky et al., 2005; Buganim et al., 2010). In this analysis we
identified the cancer-related gene signature (CGS) that was of
particular interest because it was synergistically upregulated
when H-RasV12 and WTp53 inactivating peptide (GSE56) were
concomitantly expressed in the cells (Milyavsky et al., 2005;
Buganim et al., 2010). The CGS mainly consists of secreted
molecules that were shown to have pro-cancerous functions, and
at least parts of them were shown to be induced by activated HRas (Sternlicht et al., 1999; Dhawan and Richmond, 2002;
Mendes et al., 2005; Minn et al., 2005; Wang et al., 2006;
Apte and Voronov, 2008). The CGS consists of chemokines
[chemokine (C-X-C motif) ligand 1, 2 and 3 (CXCL1, CXCL2
and CXCL3)], interleukins (IL-1b, IL-6 and CSF2) and extracellular matrix (ECM)-related proteins [matrix metallopeptidase
3 (MMP-3), CLECSF2 and TREM-1]. For further investigation,
we decided to focus on representative genes of this signature,
mainly CXCL1, IL1B and MMP3, that represent the different subgroups of the CGS. In this study we show that the different p53
mutants utilize different mechanisms to induce the CGS.
Whereas the p53 DNA-contact mutations cooperate with the
NFkB pathway, the Zn+2 region conformational mutants augment
H-Ras activity. Notably, the L3 loop conformational mutant
p53G245S did not seem to affect the examined pathways.
These findings extend our knowledge of the mutant p53–Ras
axis in cancer and might be useful for rationale-based drug
design.
Results
Cells harboring the various mutant p53 categories exhibit
diverse CGS expression patterns
Previously, we reported a gene cluster, the CGS, encoding mainly
pro-cancerous secreted molecules, that is synergistically
upregulated by p53 inactivation and oncogenic H-RasV12
expression (Milyavsky et al., 2005; Buganim et al., 2010).
Because inactivation of p53 in human tumors occurs
predominantly by point mutations, and the fact that the various
p53 mutants exhibit a wide range of mechanisms toward
malignancy (Buganim and Rotter, 2009), we decided to
examine the effect of various p53 mutants on the regulation of
the CGS.
To that end, immortalized primary human embryonic lung
fibroblasts (WI-38 cells) overexpressing the H-RasV12 oncogene
(Ras) or its empty control vector (Con), were stably infected with
one of the following vectors: an shp53 vector [vector expressing
a short hairpin RNA (shRNA) against the wild-type p53 (TP53)
3145
sequence], an shCon control vector (vector expressing an shRNA
against the mouse Noxa coding sequence) or with a vector
expressing one of the five most frequent p53 mutant forms
(p53R175H, p53H179R, p53G245S, p53R248Q and p53R273H; Fig. 1A).
These p53 mutations represent three principal categories:
conformational mutations within the Zn2+ region (p53R175H,
p53H179R), conformational mutation within the L3 loop region
(p53G245S) and DNA-contact mutations (p53R248Q, p53R273H).
Next, we compared the expression levels of the CGS in the
established cell lines using three representative genes (CXCL1,
IL1B, MMP3). In agreement with our previous data, we found
that cells expressing the H-RasV12 oncogene along with p53
knock down (hereafter referred to as Ras/shp53 cells) upregulated
expression of the CGS both at the mRNA and protein levels
compared with their control counterparts (Con/shCon, Ras/
shCon, Con/shp53; Fig. 1B,C). Interestingly, the different p53
mutant forms had distinct expression patterns. Whereas the Zn2+
region conformational mutant (p53R175H, p53H179R) cells
upregulated the CGS to a similar level as observed in cells in
which p53 was knocked down, the DNA-contact mutant
(p53R248Q and p53R273H) cells upregulated the CGS to a much
higher extent. Interestingly, the L3 loop region conformational
mutant (p53G245S) cells exhibited only a minor effect on the CGS
expression (Fig. 1B,C).
To further strengthen this observation, we examined the
mutant-p53-dependent expression of the CGS in six humantumor-derived cell lines that endogenously express different
mutant p53 types. Accordingly, following mutant p53 knock
down, the representative CGS genes (CXCL1, IL1B, IL8) were
downregulated only in cells expressing the p53 DNA-contact
mutations (SW-620, SW-480, NCI-H322), whereas p53 knock
down hardly affected the CGS expression in cells expressing the
p53 conformational mutants (NCI-H23, Hs-578-T, SKBR-3;
supplementary material Fig. S1). This result supports our
hypothesis that the p53 conformational mutants give rise to
similar CGS levels as p53 knock down, whereas the p53 DNAcontact mutations further induce the CGS.
Altogether, these data demonstrate unique characteristics and
modes of action for each mutant p53 type in regulating the CGS.
H-Ras activity is differently regulated by the various
mutant p53 categories
Because CGS expression was dependent on H-RasV12, and in
order to reveal the mechanisms by which the various p53 mutants
induce the CGS, we examined whether H-Ras activity is directly
affected by the various p53 mutants. Using the Ras activity pulldown assay [Ras binding domain (RBD) assay] (Vojtek et al.,
1993) we have recently demonstrated that wild-type p53 inhibits
the activity of the H-RasV12 oncogene by reducing its GTPloading state, resulting in inhibition of CGS expression (Buganim
et al., 2010). To measure the activity levels of H-RasV12 in the
various mutant-p53-expressing cells, we performed the RBD
assay on established WI-38 cells. Importantly, all H-RasV12expressing cells exhibited comparable levels of H-Ras mRNA
and protein (Fig. 2A,B). Notably, cells expressing high CGS
levels (Ras/shp53, Ras/p53R175H) also exhibited high H-Ras–
GTP levels, whereas cells expressing low CGS levels (Ras/
shCon, Ras/p53G245S) exhibited low H-Ras–GTP levels
(Fig. 2B). Supporting this observation, we found that the
levels of activated H-RasV12 were similar in Ras/p53H179R,
Ras/p53R175H and Ras/shp53 cells (supplementary material Fig.
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Journal of Cell Science
Fig. 1. p53 knock down or expression of
different p53 mutants cooperates with HRasV12 to induce a CGS. H-RasV12 (Ras) or
a control empty vector (Con) were stably
introduced by retroviral infection into WI-38
cells. Then, the cells were stably infected
with the shRNA against p53 (shp53) or a
control vector (shCon), and the following
p53 mutants: p53R175H, p53H179R, p53G245S,
p53R248Q and p53R273H. (A) p53 and H-Ras
protein levels in the established WI-38 cell
lines were analyzed by western blotting.
GAPDH housekeeping protein was used as
loading control. (B) mRNA levels of
CXCL1, IL1B and MMP3, three
representative genes from the CGS, were
measured by QRT-PCR in the established
WI-38 cell lines. Error bars represent the
standard deviation of duplicates. The
experiments were performed at least three
times and a representative result is shown.
(C) CXCL1 and IL-1b protein levels were
measured by ELISA assay (left) and MMP3
protein levels were measured by
zymography assay (right).
S2). Interestingly, Ras/p53R248Q cells, which expressed the
highest levels of CGS, exhibited only moderate H-Ras–GTP
levels (Fig. 2B), suggesting that the DNA-contact p53 mutants
use a different mechanism to augment CGS levels. Together,
these results raise two interesting issues; the first one being the
mechanism by which the different p53 mutant forms regulate HRasV12 activity, and the second being the additional mechanism
that is responsible for the super-elevation of the CGS in p53
DNA-contact mutants. These two issues will be addressed in the
following results descriptions.
The Zn2+ region conformational mutants interact with
BTG2 and interfere with its capability to bind to H-RasV12
Recently, we suggested that the p53 target gene, BTG2, interacts
with H-RasV12 and mediates the p53-dependent CGS suppression
(Buganim et al., 2010). Additionally, it was shown that p53
mutants act in a dominant-negative manner (Shaulian et al.,
1992). In order to elucidate the mechanism by which the p53
Zn2+ region conformational mutants induce H-RasV12 activity,
we examined whether these p53 mutants could reduce BTG2
expression. Surprisingly, although Ras/shp53 cells reduced the
expression levels of BTG2, the five mutant p53 forms, hardly
affected its expression (Fig. 3A) although they accumulated in
the cells (Fig. 1A). The same phenomenon was observed when
we monitored the mRNA levels of a well-known p53 target,
p21WAF (also known as CDKN1A; supplementary material Fig.
S3A). To exclude the possibility that the mutant p53 constructs
retain some wild-type p53 activity, we introduced the three
representative mutant p53 constructs into a p53-null cell line
(H1299), and measured the mRNA levels of the p53 downstream
targets BTG2 and p21WAF. As expected, and in contrast to the
wild-type p53, all mutant p53 forms did not affect the mRNA
levels of the p53 targets (Fig. 3B; supplementary material Fig.
S3B). By contrast, when cells were treated with cisplatinum,
a well-known DNA-damage and p53-stabilizing agent, the
induction of the p53 targets was attenuated by the various p53
mutants, excepts of p53G245S (supplementary material Fig. S4),
indicating that the various mutant p53 forms can attenuate p53
activity by a dominant-negative mechanism following DNA
damage. Together, these results suggest that, in our system,
although cells express ectopic mutant p53 proteins, the
transcriptional activity of the endogenous wild-type p53, under
basal condition, remains mostly intact.
These observations suggest that the various mutant p53 forms
upregulated the CGS expression by different mechanism rather
than inhibiting the expression of BTG2 or suppressing wild-type
p53 transactivation capability. Because mutational analysis of
p53 revealed that the amino acids Leu22 and Trp23 are essential
Journal of Cell Science
Various roles of p53 mutants in cancer
Fig. 2. Cells expressing p53 Zn+2 region conformational mutants harbor
elevated levels of activated H-Ras. (A) mRNA levels of H-Ras in the
indicated WI-38 cell lines, measured by QRT-PCR. Error bars represent
standard deviation of duplicates. The experiments were performed at least
three times and a representative result is shown. (B) A pull-down assay of Ras
activity was carried out in the indicated WI-38 cell lines using beads that were
fused to the Ras binding domain (RBD) by glutathione S-transferase (GST)
(GST–RBD). Western blot analysis shows the protein levels of active Ras,
total Ras, p53, and b-tubulin as a loading control. (C) Quantification of the HRas–GTP bands, normalized to b-tubulin, analyzed by ImageJ software. NE,
not examined.
for transactivation (Lin et al., 1995; Roemer and MuellerLantzsch, 1996), we decided to mutate these amino acids (L22Q,
W23S) in the Zn+2 region conformational mutants and to measure
the CGS expression. Interestingly, similar induction of CGS was
observed in both the intact and in the mutated Zn+2 region
conformational mutants (Fig. 3C). This indicates that the
capability of the Zn+2 region conformational mutants to induce
the CGS expression by upregulating H-Ras activity is not
dependent on the transactivation activity of the p53 mutants or
the ability to reduce the expression of the wild-type p53-mediated
target, BTG2.
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In our recent paper we showed that p53 interacts with BTG2 and
ATF3 proteins (Buganim et al., 2010). This observation together
with the suggestion that mutant p53 can exert its oncogenic
function through protein–protein interactions (Buganim and
Rotter, 2009), raised the possibility that mutant p53 might
interact with BTG2, thus squelching its function. To this end, we
examined a possible interaction between BTG2 and the various
mutant p53 categories. By using GST–BTG2-fused beads (or
GST–Empty as a control), we pulled down and measured the levels
of mutant p53 proteins that were bound to BTG2 in the various
mutant-p53-expressing cells extracts. Interestingly, the Zn+2
region conformational mutants (p53R175H, p53H179R) that caused
the highest elevation in H-Ras activity (Fig. 2B) also bound most
strongly with BTG2 (Fig. 4A). The DNA-contact mutants
(p53R248Q, p53R273H) that moderately elevated H-Ras activity
exhibited a weak interaction with BTG2. As expected, the L3 loop
region conformational mutant (p53G245S) that does not affect HRas activity also did not interact with BTG2 (Fig. 4A). These
results might suggest that by binding to BTG2, the Zn+2 region
conformational mutants squelch its activity, which accounts for the
elevated H-Ras activity, and induction of the CGS.
To examine the hypothesis that the interaction of mutant p53
with BTG2 perturbs BTG2 function, we measured the ability of
H-Ras to interact with BTG2 in the different mutant-p53expressing cells. By using the GST–BTG2 beads, we pulled
down and measured the protein levels of H-Ras that were bound
to BTG2 in representative cells extracts. As expected, the control
cells Ras/shCon and Ras/shp53 exhibited strong BTG2–H-Ras
interaction. Importantly, this interaction was abolished in
cells expressing the Zn+2 region conformational mutants. Cells
expressing the DNA-contact mutants that exhibited a weak
interaction with BTG2 (Fig. 4A) exhibited a reduced binding of
BTG2 to H-Ras, whereas cells expressing the L3 loop region
conformational mutant demonstrated a similar BTG2–H-Ras
interaction as the control cells (Fig. 4B). These data suggest that
by binding to BTG2, the Zn+2 region conformational mutants
withdraw BTG2 from the BTG2–H-Ras equation and thus
augment H-Ras activity.
Our results imply that although p53 mutants do not interfere
with wild-type p53 transactivation, they can affect its targets by a
squelching mechanism.
The extensive elevation of CGS expression exerted by the
DNA-contact p53 mutants is mediated by the NFkB
transcription factor
The highest expression of the CGS was observed in cells
expressing H-RasV12 in conjunction with the p53 DNA-contact
mutants (p53R248Q and p53R273H) and was much higher than in p53
knock-down cells. This suggests that these p53 mutants possess
a gain-of-function activity in inducing the CGS expression.
However, these cells express only moderate levels of H-Ras–
GTP, suggesting that the mechanism of the CGS induction by these
p53 mutants does not exclusively involve H-Ras activation.
Therefore, we sought to elucidate the mechanism that underlies the
enhanced activity of the DNA-contact mutants in respect to the
CGS expression. Because both wild-type p53 and mutant p53 were
shown to be involved in the NFkB pathway (Mayo et al., 1997;
Dreyfus et al., 2005; Scian et al., 2005; Weisz et al., 2007b;
Fontemaggi et al., 2009; Meylan et al., 2009), and because we
recently demonstrated that NFkB is involved in the induction of
the CGS (Buganim et al., 2010) we postulated that NFkB might be
Journal of Cell Science
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Fig. 3. The function of the Zn+2 region conformational
mutants does not involve transcriptional regulation.
(A) mRNA levels of wild-type p53 target gene, BTG2, were
measured by QRT-PCR in the established WI-38 cell lines.
(B) The H1299 p53-null cell line was transiently transfected
with the following expressing vectors: control empty vector
(Con), wild-type p53 (p53 WT), p53R175H, p53G245S,
p53R248Q. Following 48 hours of transfection, cells were
collected and BTG2 mRNA levels were measured by QRTPCR. (C) The Zn+2 region conformational mutants were
additionally mutated in their transactivation domain (L22Q,
W23S). mRNA levels of CXCL1, IL1B and MMP3 were
measured by QRT-PCR in the established RasV12-expressing
WI-38 cell lines.
the factor that mediates the activity of the DNA-contact p53
mutants. To this end, we knocked down p65, a member of the
NFkB family, in the various WI-38 cell lines and measured its
effect on mutant p53-mediated CGS expression. The capacity of
the siRNA to knock down p65 levels was estimated by measuring
its protein levels (Fig. 5A; supplementary material Fig. S5A).
Notably, downregulation of p65 resulted in a substantial reduction
in the mRNA levels of the CGS independently of p53 status
(Fig. 5B; supplementary material Fig. S5B). Interestingly, we
found that the super-induction of the CGS exerted solely by the
DNA-contact p53 mutants (p53R248Q and p53R273H) was totally
abolished by the knock down of p65 (Fig. 5B; supplementary
material Fig. S5B). To further support the above results we
examined the expression of CGS in three different mutant p53
Fig. 4. p53 Zn+2 region conformational mutants induce the
expression of the CGS through interaction with BTG2 and
squelching its function. (A) A pull-down assay, detecting binding of
BTG2 to p53 mutants was carried out in the indicated RasV12-expressing
WI-38 cell lines, using GST–BTG2 beads and GST–Empty beads as a
control. Western blot analysis shows the protein levels of p53 (GAPDH
was used as a control). GelCode was used to visualize GST-fused
proteins. (B) A pull-down assay, detecting binding of BTG2 to H-Ras
was carried out in the indicated RasV12-expressing WI-38 cell lines,
using the same experimental set-up as in A. Western blot analysis shows
the protein levels of H-Ras (GAPDH was used as a control). Ponceau
staining of the blot was used to visualize GST-fused proteins.
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Various roles of p53 mutants in cancer
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Fig. 5. NFkB mediates elevated expression of the CGS exerted by
the p53 DNA-contact mutants. The indicated RasV12-expressing
WI-38 cell lines were transfected with either siRNA against p65 (sip65) or with a control siRNA (si-Con). Cell were collected 72 hours
post transfection. (A) p65 protein levels were measured by western
blot analysis. (B) CXCL1, IL1B and MMP3 mRNA levels were
measured by QRT-PCR. The number above each bar is the fold
repression in each individual cell line caused by p65 knock down.
bearing cancer cell lines (SW-620, SW-480, SKBR-3) in which
p53 was stably knocked down (supplementary material Fig. S5C).
Notably, silencing p65 in SW-620 (supplementary material Fig.
S5D), which endogenously express the DNA-contact mutant
p53R273H, reduced the expression of the representative CGS genes,
and abolished their increased expression cause by mutant p53R273H
(Fig. 6A). A complementary strategy to manipulate the NFkB
pathway would be to activate it by TNF-a. To this end, SW-480,
which harbors the DNA-contact mutant p53R273H, and SKBR-3,
which harbors the Zn+2 region conformational mutant p53R175H,
were treated with TNF-a, and the mRNA levels of the
representative CGS genes (IL8, CXCL1, CXCL2) were
examined. In agreement with the above-described results, a
substantial decrease in the level of CGS was observed in SW480 cells when mutant p53 was knocked down in basal conditions.
Following TNF-a treatment, the CGS expression levels were
induced. Interestingly, the fold reduction observed following
mutant p53R273H knock down, under basal conditions, were
diminished. This suggests that the mutant-p53R273H-dependent
induction of CGS expression under basal conditions is mediated by
NFkB. (Fig. 6B). By contrast, no major differences in the levels of
CGS were observed in SKBR-3 cells following mutant p53R175H
knock down under basal condition. Following TNF-a treatment,
CGS expression was induced in shp53 and shCon cells to similar
levels (Fig. 6C). Similarly to the SKBR-3 cells, when we
examined NCI-H23 and Hs-578-T cells, harboring p53
conformational mutants (p53M246I and p53V157F, respectively),
no substantial differences were observed in the levels of CGS
when comparing shCon and shp53, both in basal condition and
following TNF-a treatment (supplementary material Fig. S6).
These results indicate that NFkB is a major regulator of the CGS
expression mediated by the DNA-contact p53 mutants.
Discussion
Recently, by using an in vitro stepwise well-controlled
transformation model (Milyavsky et al., 2003) with a detailed
genomic profiling (Milyavsky et al., 2005), we were able to
uncover that loss of wild-type p53 synergizes with H-RasV12 to
induce a unique CGS (Buganim et al., 2010). This agrees with a
previous report suggesting that the combination of p53 loss and
Ras activation synergistically induces the expression of promalignant genes (McMurray et al., 2008). In our previous study,
we suggested that wild-type p53 represses the expression of CGS
by at least two mechanisms. On the one hand, p53 transactivates
its target BTG2, which binds to H-RasV12 and represses its
activity, and on the other hand, following p53 stabilization, the
p53 target ATF3 is activated, binds to the CGS gene promoters
and represses their expression (Buganim et al., 2010).
Here, we investigate the role of the different p53 mutants in the
induction of the CGS during tumor development.
When mutated, p53 either acts in a dominant-negative manner,
in which it inhibits wild-type p53 tumor-suppressive functions, or
acts in a gain-of-function manner, in which mutant p53 shows
additional oncogenic activities, supporting tumorigenesis (Brosh
and Rotter, 2009). Here we examined the mechanisms utilized by
the different p53 mutants, and found that the DNA-contact
mutants (p53R248Q and p53R273H), but not the p53 conformational
mutants (p53R175H and p53H179R), induce the CGS in a gain-offunction manner. This was demonstrated by the higher CGS
expression levels observed in cells expressing the p53 DNAcontact mutants, compared with the p53 knock down and p53
conformational mutants (Fig. 1B,C).
Supporting these data, the p53 DNA-contact mutants showed
higher CGS induction than p53 conformational mutants, when
endogenously expressed in human-cancer-derived cell lines
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Fig. 6. Endogenous p53 DNA-contact mutants, expressed in cancer cell
lines, cooperate with NFkB to induce the CGS. SW-620, SW-480 and
SKBR-3 cell lines were knocked down for the endogenously expressed
mutant p53. (A) SW-620 cells were transfected with either siRNA against p65
(si-p65) or with a control siRNA (si-Con). CXCL1, IL1B and MMP3 mRNA
levels were measured by QRT-PCR. (B,C) SW-480 and SKBR-3 cells were
treated with 20 ng/ml TNF-a for 24 hours, and mRNA levels of IL8, CXCL1
and CXCL2 were measured by QRT-PCR.
(supplementary material Fig. S1). These observations are
especially intriguing because they suggest that not all p53
mutants classes possess gain-of-function activity in CGS
induction.
Furthermore, we demonstrate that different types of mutant
p53 engage in different molecular pathways to facilitate the
cooperation with oncogenic H-RasV12 in inducing the CGS
(Fig. 7). The DNA-contact mutants (p53R248Q and p53R273H)
cooperate with the NFkB-dependent pathway to further induce
the CGS. Concordantly, it was demonstrated that mutant p53 can
exert a gain-of-function activity by stimulating NFkB activity
(Scian et al., 2005; Weisz et al., 2007b). Interestingly, we have
found that the expression of the different p53 mutants did not
reduce the expression levels of wild-type p53 gene targets,
including BTG2, which we recently found to mediate the p53dependent H-Ras activity inhibition. Nevertheless, we suggest
that the Zn+2 region conformational p53 mutants (p53R175H and
p53H179R) interact with BTG2 and disrupt its binding to H-Ras,
therefore enabling higher levels of active H-Ras and CGS. This
proposed mechanism is supported by the observation that
damaging the transactivation capability of mutant p53, by
mutating the L22, W23 codons, did not affect the expression of
the CGS, and agree with other studies that demonstrated protein–
protein interaction between mutant p53 and other proteins (e.g.
p63, p73, Mre11, NF-Y) (Buganim and Rotter, 2009) that leads to
mutant p53 gain of function. By the ability to interact with the
BTG2 protein, p53 mutants could interfere with p53 pathway,
and inhibit its tumor-suppressive functions.
Most of p53 mutations reside within the DNA binding domain;
however, they can be divided into several categories according to
their structural conformation. Therefore, the differences in the
mode of action between the various mutant p53 categories could
be explained by their specific characteristics. For example,
mutant p53G245S, which in our system hardly elevates the
expression levels of the CGS, partially retains some of the wildtype p53 characteristics such as stabilization and DNA binding
affinity (Bullock and Fersht, 2001).
The three Ras family members, K-Ras, H-Ras and N-Ras, are
highly homologous (85% sequence identity), and share similar
functions (Downward, 2003). Moreover, in fibroblast cells, HRas is a more effective mediator of cellular transformation than
the other Ras oncoproteins (Maher et al., 1995). Therefore,
because our in vitro transformation model is based on human
primary fibroblast cells, H-RasV12 was used. Yet, human-cancerderived cell lines that harbor the K-Ras oncogene along with p53
DNA-contact mutants also showed higher CGS expression then
cells expressing p53 conformational mutants (supplementary
material Fig. S1). Therefore, the results obtained in a fibroblast in
vitro transformation model, exploiting the H-Ras oncogene, are
predictive also of K-Ras-oncogene-derived cancer.
In conclusion, we suggest that during the transformation
process, when p53 is mutated, the expression of a cancer-related
gene signature is induced, and the tumor-suppressive function of
p53 is overridden. The different p53 mutations can augment these
phenotypes by different means; either by cooperating with the
NFkB protein or by squelching the activity of BTG2.
Li-Fraumeni syndrome is a familial disorder associated with a
germline mutation in one of the p53 alleles. This predisposes the
patients to a high incidence of cancer (Malkin et al., 1990).
Because our model represents a situation whereby both wild-type
and mutant p53 are present in the cells, as in Li-Fraumeni
syndrome, it is tempting to speculate that high levels of mutant
p53 can overcome the tumor-suppressive effect of the wild-type
p53 protein, and thus, a p53 heterozygous state is sufficient
to drive the cells toward tumorigenesis. This hypothesis is
in accordance with Varley et al., demonstrate that 40% of
Li-Fraumeni syndrome patients did not undergo loss of
heterozygosity, and gave rise to tumors that were heterozygous
for p53 (Varley et al., 1997). Interestingly, it should be
mentioned that our established cells expressing the various p53
mutants (except of p53R245S) had a tumorigenic phenotype,
measured by tumor formation when injected into nude mice (data
not shown). Both the WI-38 cells grown in vitro and tumors
generated in vivo were heterozygous for p53, suggesting that
Various roles of p53 mutants in cancer
3151
Fig. 7. A schematic model describing the different
mechanisms utilized by the various p53 mutants in the
regulation of Ras circuit to induce the CGS. To induce the
CGS, p53 Zn+2 region conformational mutants (p53R175H
and p53H179R) mainly inhibit BTG2 function and therefore
elevate H-RasV12 activity, whereas p53 DNA-contact
mutants (p53R248Q and p53R273H) mainly cooperate with
NFkB. Mutant p53G245S does not affect any of the examined
pathways. Arrows indicate activation, whereas bars indicate
repression. Bold lines indicate stronger regulation.
Journal of Cell Science
these cells utilize other mechanisms for tumorigenesis, as in 40%
of Li-Fraumeni syndrome patients.
Uncovering the nature of the multiple pathways involved in
this important cellular junction and the capability to distinguish
between the p53 mutant categories might provide important tools
for tailor-made cancer therapy that is based on p53 status and its
distinct regulatory pathways.
Materials and Methods
Antibodies, compounds and plasmids
The following primary antibodies were used: anti-p53 (DO-1 and 1801) were
kindly provided by David Lane (Ninewells Hospital and Medical School, Dundee,
Scotland); anti-H-Ras, anti-p65 (Santa Cruz Biotechnology, Santa Cruz, CA); antiGAPDH (Chemicon, Billerica, MA); anti- b-tubulin (Sigma, St. Louis, MO).
Plasmids
The pRetroSuper-p53 shRNA-Blast, the pRetroSuper-shmNOXA-Blast and the
pBabe-H-RasV12-Hygro constructs were kindly provided by Doron Ginsberg (BarIlan University, Israel). GST-BTG2 expression plasmids were constructed by
cloning the BTG2 open reading frame (ORF) into the pGEX-GP1/3 vector that was
kindly provided by Elior Peles (The Weizmann Institute, Israel). The various p53
vectors were constructed by sub-cloning the wild-type p53 ORF from the PLXSNneo-p53 construct into the PWZL-Blast construct using BamHI restriction. Sitedirected mutagenesis (Santa Cruz Biotechnology), was conducted on the pWZLblast-p53 construct according to the manufacturer’s instructions using specific
primers to create the specific mutations. Supplementary material Table S2 shows
all primer sequences.
Compounds
Dimethyl sulfoxide (DMSO) (Sigma), cisplatinum (Pharmachemie, Petach Tikva,
Israel), TNF-a (R&D systems, Minneapolis, MN), GelCode Blue staining reagent
(Thermo Scientific, MA, USA), Ponceau (Sigma).
Cell culture, transfections and retroviral infections
The immortalized primary human embryonic lung fibroblasts (WI-38) were
established and maintained as described previously (Milyavsky et al., 2003). The
colon carcinoma cell lines, SW-480 and SW-620, were maintained in DMEM
supplemented with 10% fetal calf serum and antibiotics. The breast
adenocarcinoma cell line, SKBR-3, was maintained in McCoy’s medium
supplemented with 10% fetal calf serum, 2 mM L-glutamine, 1 mM sodium
pyruvate and antibiotics. The non-small lung carcinoma cell lines H1299, NCIH322, NCI-H23 and breast cell line Hs-578-T were maintained in RPMI-1640
supplemented with 10% fetal calf serum and 2 mM L-glutamine. The infection
procedures using ecotropic Phoenix-packaging cells were described previously
(Milyavsky et al., 2003). The transfections into H1299 cells were conducted using
Lipofectamine 2000 reagent (Invitrogen, Carlsbad, CA) according to the
manufacturer’s instructions. Knock down of p65 was conducted by transfection
with specific oligonucleotides (Dharmacon, Lafayette, CO) using DharmaFECT3
reagent (Dharmacon). The analysis was performed 72 hours after transfection.
RNA isolation and quantitative real time-PCR
Total RNA was isolated using a NucleoSpin kit (Macherey-Nagel, Düren,
Germany) according to the manufacturer’s protocol. A 2 mg aliquot of the total
RNA was reverse transcribed using Bio-RT (Bio Lab, Jerusalem, Israel) and
random hexamer primers. QRT-PCR was performed with an ABI 7300 instrument
(Applied Biosystems, Carlsbad, CA) using Platinum SYBR Green qPCR SuperMix
(Invitrogen). Specific primers were designed for the following genes: CXCL1,
IL1B, MMP3, CXCL2, IL8, BTG2, p21WAF, H-Ras and p65. cDNA levels were
normalized to GAPDH (for primer sequences see supplementary material Table
S1).
CXCL1 and IL-1b enzyme-linked immunosorbent assay and MMP3
zymography assay
Cells (26105) were grown on six-well plates with serum-free MEM for 72 hours.
Cell-conditioned media were collected and centrifuged at 20,800 g, at 4 ˚C. CXCL1
and IL-1b proteins were detected by enzyme-linked immunosorbent assay
(ELISA) using the Human GRO-a/CXCL1 or IL-1b immunoassay kits (R&D
systems), according to the manufacturer’s instructions. For determination of
MMP3 protein levels, equal volumes of cell media were separated on gelatincontaining polyacrylamide gels. Then, gels were incubated overnight in
developing buffer containing 0.01% Brij (Sigma), at 37 ˚C, to activate the MMP
enzymes. Gels were then stained with Coomassie Blue solution (0.1–0.5%
Coomassie Brilliant Blue R250 in 5% acetic acid and 10% methanol) to detect
areas of catalytic activity.
Ras activity pull-down assay, BTG2 binding assay and western blotting
The glutathione S-transferase (GST)–Ras binding domain (RBD; GST–RBD) and
the GST–BTG2 fused beads were prepared as described previously (Frangioni and
Neel, 1993). Cells were lysed in cold RBD lysis buffer. Equal amounts of cell
extracts were incubated with the GST–RBD, GST–BTG2 or GST–Empty beads,
overnight. The proteins were eluted with sample buffer, separated on SDSpolyacrylamide gels, and analyzed. Western blotting was conducted as described
by Milyavsky et al. (Milyavsky et al., 2003).
Funding
This research was supported by a Center of Excellence grant from
Flight Attendant Medical Research Institute, Yad Abraham Center
for Cancer Diagnosis and Therapy [grant number 7034640901]; and
the European Community EC FP7-INFLACARE [number 223151,
grant number 7104370402]. This publication reflects the authors’
views and not necessarily those of the European Community. The EC
is not liable for any use that may be made of the information
contained herein. V. R. is the incumbent of the Norman and Helen
Asher Professorial Chair Cancer Research at the Weizmann Institute.
Supplementary material available online at
http://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.099663/-/DC1
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