Download Chemotherapeutic agents induce the expression

Carcinogenesis vol.34 no.1 pp.190–198, 2013
doi:10.1093/carcin/bgs318
Advance Access publication October 10, 2012
Chemotherapeutic agents induce the expression and activity of their clearing enzyme
CYP3A4 by activating p53
Ido Goldstein*,†, Noa Rivlin†, Or-yam Shoshana,
Osnat Ezra, Shalom Madar, Naomi Goldfinger
and Varda Rotter
The Department of Molecular Cell Biology, the Weizmann Institute of
Science, Rehovot, 76100 Israel
*
To whom correspondence should be addressed. Tel: +972 8 9342398; Fax:
+972 8 9342398;
Email: [email protected]
Cytochrome P450 (P450) enzymes are abundantly expressed in the
human liver where they hydroxylate organic substrates. In a microarray screen performed in human liver cells, we found a group of
eleven P450 genes whose expression was induced by p53 (CYP3A4,
CYP3A43, CYP3A5, CYP3A7, CYP4F2, CYP4F3, CYP4F11,
CYP4F12, CYP19A1, CYP21A2 and CYP24A1). The mode of regulation of four representative genes (CYP3A4, CYP3A7, CYP4F2 and
CYP4F3) was further characterized. The genes were induced in a
p53-dependent manner in HepG2 and Huh6 cells (both are cancerderived human liver cells) and in primary liver cells isolated from
human donors. Furthermore, p53 was found to bind to p53-responsive elements in the genes’ DNA-regulatory regions and to enhance
their transcription in a reporter gene assay. Importantly, when
p53 was activated following the administration of either of three
different anticancer chemotherapeutic agents (cisplatin, etoposide
or doxorubicin), it was able to induce CYP3A genes, which are
the main factors in systemic clearance of these agents. Finally, the
p53-dependent induction of P450 genes following either Nutlin or
chemotherapy treatment led to enhanced P450 enzymatic activity.
Thus, in addition to the well-established role of p53 at the tumor site,
our data unravels a novel function of hepatic p53 in inducing P450
enzymes and position p53 as a major factor in the hepatic response
to xenobiotic and metabolic signals. Importantly, this study reveals
a novel pathway for the induction of CYP3As by their substrates
through p53, warranting the need for careful consideration when
designing systemically administered chemotherapeutic regimens.
Introduction
Genes from the cytochrome P450 (P450) superfamily encode
enzymes that catalyze hydroxylation reactions. Each P450 enzyme
has different substrate specificities and, accordingly, different tissue distributions. Many P450 enzymes are abundantly expressed in
the liver where they hydroxylate a wide range of substrates wired to
many metabolic circuits and are involved in the metabolism of both
endogenous (e.g. lipids, steroids) and exogenous (e.g. drugs, toxins)
compounds. The P450 genes are divided into families and subfamilies
according to their homology, activity and substrate specificity (1,2).
The four members of the CYP3A subfamily (CYP3A4, CYP3A43,
CYP3A5 and CYP3A7) are liver-resident enzymes paramount to drug
metabolism. Particularly, CYP3A4, the most abundant P450 enzyme in
adult liver, is responsible for metabolizing more than 50% of clinically
used drugs. An important group of drugs heavily affected by CYP3A4 is
anticancer chemotherapeutics, a great number of clinically used chemotherapeutic agents serve as substrates for CYP3A4 (3). More often than
not, CYP3A4 facilitates the systemic clearance of these compounds,
sometimes leading to reduced efficiency or even treatment failure. In
Abbreviations: AMPK, adenosine monophosphate–activated protein kinase;
P450, cytochrome P450; p53REs, p53-responsive elements; mRNA, messenger RNA.
†
These authors contributed equally to this work.
some cases, substrates of CYP3A4 also serve as transcriptional inducers
of its gene resulting in increased activity and thus enhanced clearance
of the inducing drug (4). For example, tamoxifen, an estrogen-receptor
modulator used for treatment of breast cancer, was shown to induce
CYP3A4 leading to its own clearance (5). In addition to inducing its own
clearance, a drug that induces CYP3A4 may as a result induce the clearance of concomitantly administered drugs. This option is particularly
relevant to anticancer treatment regimens that often include a combination of drugs. Indeed, administration of tamoxifen combined with aromatase inhibitors (which are also CYP3A substrates) to breast cancer
patients led to a reduction in their plasma levels (6). Therefore, the efficiency of a given drug is highly dependent upon CYP3A activity levels
and the transcriptional regulation of CYP3A genes is a crucial determinant in drug administration regimens (4,7).
The CYP4F subfamily is also abundant in the liver where
it is involved in lipid metabolism. CYP4F enzymes catalyze
ɷ-hydroxylation of long fatty acids that, following two more reactions, can enter the more utilized pathway of fatty acid catabolism,
ɷ-oxidation. Consequently, these fatty acids can be either completely
oxidized to produce available energy, metabolized to succinate and
acetyl-CoA, or alternatively be excreted in the urine (8). In addition
to fatty acid metabolism, CYP4F enzymes take part in several metabolic pathways that are involved in both physiological and pathological aspects such as hypertension and inflammation (9). Similarly
to CYP3As, the CYP4F subfamily is subjected to transcriptional
induction in response to stimuli. For instance, activation of adenosine
monophosphate–activated protein kinase (AMPK), a master regulator
of metabolism, leads to induction of CYP4F2 expression (10).
The p53 protein is a sequence-specific transcription factor with
established roles in many biological processes. The classic property of
p53 is tumor suppression accomplished by regulation of various gene
networks leading to cell-cycle arrest, apoptosis etc (11,12). In recent
years, p53 was also found to regulate metabolic pathways such as glycolysis and oxidative phosphorylation (13,14). Accordingly, p53 was
shown to be phosphorylated and activated by the metabolic sensor
AMPK (15,16). AMPK accelerates adenosine triphosphate production by modulating metabolic pathways such as enhancing oxidative
phosphorylation and fatty acid oxidation (17). The activation of p53
by AMPK leads to activation of metabolism-related genes such as
GAMT, resulting in increased fatty acid oxidation (18). Also, p53 was
found to regulate two mouse P450 genes, Cyp7a1 and Cyp8b1, thus
affecting hepatic lipid metabolism (19).
In this study, we show that p53 is a regulator of a variety of P450
genes in cells of hepatic origin—11 members from five different P450
subfamilies were induced by p53. Focusing on genes from two P450
subfamilies—CYP3A and CYP4F, we demonstrate the ability of p53
to bind to DNA-regulatory elements in the genes and to promote their
transcription. Interestingly, we show that following a chemotherapeutic
stimulus, p53 induces the expression and enzymatic activity of CYP3As,
the main enzymes responsible for clearing chemotherapeutics. The
presented data points to a scenario in which p53 plays a systemic role in
response to chemotherapy in addition to its established role at the tumor
site. Considering these findings may help in improving the efficacy of
chemotherapeutic treatment regimens.
Materials and methods
Reagents, vectors and antibodies
Nutlin-3a was purchased from Alexis biochemicals (San Diego, CA), doxorubicin, etoposide and AICAR were purchased from Sigma. pRetroSuper vectors harboring sh-RNA against human p53 (sh-p53) or mouse noxa (sh-con)
were kindly provided by Dr Doron Ginsburg (Bar-Ilan University, Israel).
The p53 construct pC53-SN3 was kindly provided by Prof. Bert Vogelstein
(Johns Hopkins University School of Medicine, Baltimore, MD). Reporter
© The Author 2012. Published by Oxford University Press. All rights reserved. For Permissions, please email: [email protected]
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p53 induces P450 expression
gene vectors were purchased from New England Biolabs (Ipswich, MA). The
following antibodies were used: α-p53 (DO-1), α-p53 (CM1) (Novacastra
Laboratories, UK) α-p21 (Santa Cruz Biotechnology, sc-397) α-gapdh
(Chemicon, mab374), goat antimouse or goat antirabbit horseradish peroxidase–conjugated antibodies (Jackson Immunoresearch Laboratories).
Cell culture
All cell lines were cultured in a humidified incubator at 37°C and 5% CO2 in
Dulbecco's modified Eagle's medium supplemented with 10% fetal calf serum
and Pen/Strep solution (Biological industries, Beit-Haemek, Israel). HepG2
cells were kindly provided by Prof. Yehiel Zick; Hep3B and Huh6 cells were
kindly provided by Prof. Yosef Shaul (both from the Weizmann Institute of
Science, Israel).
Retroviral infection
Amphotropic and ecotropic Phoenix-packaging cells were transfected with
10 µg of DNA of the appropriate retroviral construct by a standard calcium
phosphate procedure. Culture supernatants were collected 48 h post transfection and filtered. Cells were infected with the filtered viral supernatants in
the presence of 4 µg/ml polybrene (Sigma) for 12 h, after which the medium
was changed. Fresh viral suspensions were added after a 24 h interval for an
additional 12 h. Infected cells were selected using Blasticidin (10 µg/ml) for
20 days.
Microarray
RNA from HepG2 cells treated with Nutlin (10 µM) or its solvent for 24 h was
collected (two separate biological replicates were used for each cell type and each
treatment). Complementary DNA samples were generated and labeled using
Affymetrix 1-cycle expression kit (Affymetrix, Santa Clara, CA) and hybridized
to ‘Human Gene 1.0 ST’ Affymetrix microarray using Affymetrix hybridization
kit materials according to manufacturer’s instruction. Data were processed by
applying the RMA algorithm and using the Partek software. The entirety of the
microarray results was deposited in GEO (accession #GSE30137).
Western blot
Cells lysed using tris lysis buffer TLB (10 mM tris pH7.5, 500 mM NaCl, 1% triton
X-100, 0.5% sodium deoxycholate, 0.1% sodium dodecyl sulfate, 1% protease
inhibitors). Lysate was centrifuged at 20,000g for 10min. protein concentration
was determined from the supernatant using BCA reagents (Pierce) and 50μg of
protein were electrophorated in sodium dodecyl sulfate–polyacrylamide gel electrophoresis gels. Gels were transferred to nitrocellulose membranes that were then
blocked using 5% milk in phosphate-buffered saline (0.05% tween) PBST. Then,
membranes were incubated for 1 h with a primary antibody followed by incubation
with secondary goat antimouse or goat antirabbit horseradish peroxidase–conjugated antibodies and developed using an ECL kit (Amersham Biosciences).
RNA isolation and quantitative real-time PCR
Total RNA was isolated using NucleoSpin kit (Macherey-Nagel) according
to the manufacturer’s protocol. A 2 µg aliquot of total RNA was reverse transcribed using Bio-RT (Bio Labs) and random hexamer primers. Quantitative
real-time PCR (QRT–PCR) was performed with an ABI-7300 instrument
(Applied Biosystems, Foster City, CA) using Platinum SYBR Green qPCR
SuperMix (Invitrogen, Carlsbad, CA). Gene values were normalized POLR2A.
QRT–PCR data are described in arbitrary units. The details of the primers used in this study are presented in Supplementary Table 1, available at
Carcinogenesis Online.
Isolation and culturing of primary human liver cells
Human liver cells were isolated as described previously (20). Informed consent was obtained from each donor and the study protocol conforms to the
ethical guidelines of the 1975 Declaration of Helsinki as reflected in a priori
approval by the Committee on Clinical Investigations (Institutional Review
Boards of Sheba Medical Center and Rabin Medical Center).
Chromatin immunoprecipitation
Cells underwent cross-linking (1% formaldehyde, room temperature, 10 min)
followed by quenching (glycine 0.125 M). Cells were rinsed using cold phosphate-buffered saline, incubated with 20% trypsin (Gibco), again washed using
phosphate-buffered saline, scraped and centrifuged. Cells were lysed (5 mM
1,4-piperazinediethane-sulfonic acid, pH 8.0, 85 mM KCl, 0.5% NP-40, 1%
protease inhibitors) on ice for 20 min. Nuclei were collected by centrifugation
(4000 r.p.m.), resuspended in nuclear lysis buffer (50 mM Tris–Cl, pH 8.1,
10 mM ethylenediaminetetraacetic acid , 1% sodium dodecyl sulfate, 1% protease inhibitors) and incubated on ice for 10 min. Samples were sonicated to an
average DNA fragment length of 500 bp and then centrifuged (20,000g). The
chromatin solution was precleared by adding protein A beads (2 h, 4°C; Santa
Cruz Biotechnology). Immunoprecipitation of chromatin—rotation, 12 h,
4°C, 1 µg polyclonal α-p53 antibody—either CM1 or H47 was followed by
incubation with 30 µl protein A beads (2 h). Immunoprecipitates were consecutively washed with dilution buffer (100 mM Tris–Cl, pH 9.0, 500 mM LiCl,
1% NP-40, 1% deoxycholic acid, 1% protease inhibitors), TSE 150, TSE 500
and TE pH 8. Samples were treated with 10 µg RNase A (30 min), followed by
30 µg of proteinase K treatment (2 h, 50°C) and incubation at 65°C overnight.
DNA samples were extracted using QIAquick PCR Purification Kit (Qiagen).
QRT–PCR was performed as described above with each sample containing
2 µl of immunoprecipitated DNA; values were normalized for 1% input values.
Reporter gene assay
Cells were plated in 48-well plates in triplicates (5 × 104 cells/well) and
24 h later were transfected (lipofectamine 2000, Invitrogen, Carlsbad, CA)
with 450 ng/well of pCLuc-Basic2 harboring the p53-responsive elements
(p53REs) specified in Figure 3A in addition to 10 ng/well of pCMV-GLuc
and 10 ng/well of pC53-SN3. Luminescence values were estimated 48 h later
from 20 µl of culture medium. Values of the Cypridinia enzyme (encoded from
pCLuc-Basic2) were normalized to values of Gaussia luciferase (encoded
from pCMV-GLuc).
Cytochrome P450 activity assay
P450 3A activity was measured using a commercial kit according to the manufacturer’s instructions (Promega, Cat# V9001 (21)). Briefly, cells were plated
on 96-well plates (3 × 104 cells per well) and treated with P450-inducing compounds the next day. After 48 h, medium containing a P450-activated proluciferin was added to the cells for 1 h followed by quantification of luminescence.
Statistical analysis
All presented data are representative experiments from at least three repeats.
Columns marked with asterisks denote a significant (P < 0.05) elevation, measured by a student t-test compared with unmarked columns.
Results
p53 induces the expression of P450 genes in human liver cells
In order to explore hepatic-related p53-dependent gene networks,
we analyzed the transcriptome of hepatocyte-derived cells—HepG2.
These cells retain most of its original hepatic features and serve as
an excellent and extensively used in vitro model for both hepatocyte
research and p53 pathway analysis (10,15,22,23). We treated HepG2
cells with Nutlin, a compound that disrupts the interaction between
p53 and its negative regulator MDM2, thus stabilizing p53 protein
level, leading to its accumulation and to the activation of p53-dependent transcription (24). To avoid any p53-independent effects of Nutlin,
we used a control of HepG2 cells stably expressing short-hairpin
RNA targeting p53 (termed HepG2sh-p53), whereas the cells with unaffected p53 levels stably expressed a non-relevant short-hairpin RNA
(termed HepG2sh-con) as described previously (25). HepG2sh-con cells
treated with Nutlin showed a prominent increase in p53 protein levels
and in its downstream target gene—p21 as compared with untreated
cells. In contrast, HepG2sh-p53 cells exhibited lower p53 and p21 levels
compared with HepG2sh-con cells in the presence or absence of Nutlin
(Figure 1A). Therefore, this experimental setup is suitable for analyzing p53-dependent transcriptional patterns.
Next, we analyzed the transcriptome of HepG2 cells under these
conditions using a gene expression microarray. Following an analysis
of variance and exclusion of genes with a fold change lower than 2 or
a P value higher than 0.05, we found a total of 341 annotated genes
whose expression was induced by p53 (25). Strikingly, out of this
group, 11 genes were members of the cytochrome P450 superfamily:
CYP3A4, CYP3A43, CYP3A5, CYP3A7, CYP4F2, CYP4F3, CYP4F11,
CYP4F12, CYP19A1, CYP21A2 and CYP24A1. In order to corroborate
our findings, we repeated the above-mentioned experimental setup and
performed a QRT–PCR analysis that completely reproduced the pattern
observed in the microarray analysis (Figure 1B).
In order to evaluate whether this pattern of expression is a more
general phenomenon, we evaluated the effect of p53 activation
on four representative genes from two different P450 subfamilies
(CYP3A4, CYP3A7, CYP4F2 and CYP4F3) in other hepatic-derived
cells. First, we treated Huh6sh-con cells (also from a hepatic origin
and harboring wild-type p53) with Nutlin, which resulted in both the
stabilization of p53 protein levels and the induction of messenger
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Fig. 1. The expression of various P450 genes is induced by p53 in HepG2 cells. (A) HepG2 cells were treated with the p53-activating agent Nutlin (10 µM) for
24 h, total protein was extracted and the protein levels of p53 and p21 were analyzed in a western blot analysis. (B) HepG2 cells were treated with the p53activating agent Nutlin (10 µM) for 24 h, total RNA was extracted and the mRNA levels of the indicated genes were analyzed in a QRT–PCR analysis. Results are
of a representative experiment from three experiments.
RNA (mRNA) levels of all four genes in comparison with the controls
(Figures 2A and 2B). In addition, we treated normal primary liver
cells isolated from human donors with Nutlin and observed both
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p53 stabilization and induction of CYP3A4, CYP3A7 and CYP4F2
(Figures 2C and 2D). We were not able to detect CYP4F3 mRNA in
these cells.
p53 induces P450 expression
Fig. 2. The expression of P450 genes is induced by p53 in liver-derived cells. (A) Huh6 cells were treated with the p53-activating agent Nutlin (25 µM) for
48 h, total protein was extracted and the protein level of p53 was analyzed in a western blot analysis. (B) Huh6 cells were treated with the p53-activating agent
Nutlin (25 µM) for 48 h, total RNA was extracted and the mRNA levels of the indicated genes were analyzed in a QRT–PCR analysis. (C) Primary human liver
cells were treated with the p53-activating agent Nutlin (10 µM) for 24 h, total protein was extracted and the protein level of p53 was analyzed in a western blot
analysis. (D) Primary human liver cells were treated with the p53-activating agent Nutlin (10 µM) for 24 h, total RNA was extracted and the mRNA levels of the
indicated genes were analyzed in a QRT–PCR analysis. Results are of a representative experiment from three experiments.
Taken together, the findings presented in Figures 1 and 2 show a
role for p53 in regulating the expression of P450 genes from a number of gene families in several liver-derived cells. Due to lack of validated commercial antibodies, we were unable to evaluate the protein
levels of P450 enzymes. However, the QRT–PCR analysis was performed with TaqMan probes for higher precision and specificity
of detection (Supplementary Table 1, available at Carcinogenesis
Online).
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Fig. 3. p53 binds to the DNA-regulatory elements of P450 genes and through them enhances their transcription. (A) The promoters of CYP3A4 and CYP3A7 and
the introns of CYP4F2 and CYP4F3 contain p53-responsive elements. The sequence of the indicated p53REs is detailed in the left-hand side of panel, the core
elements are underlined. p53REs are schematically depicted on the right-hand side of the panel; gray rectangles represent the p53RE with its location indicated
in base pair numbers relative to the transcription start site (the number indicated is of the first base in the p53RE). (B) p53 binds to the genes’ p53REs. HepG2
cells were treated with the p53-activating agent Nutlin (10 µM) for 24 h and then subjected to chromatin immunoprecipitation with two separate polyclonal
α-p53 antibodies. The immunoprecipitated DNA was then subjected to QRT–PCR analysis with primers amplifying the indicated p53RE regions. Samples are
normalized to 1% of total DNA. (C) p53 elevates gene expression through the p53REs of P450 genes. p53 was ectopically expressed in the p53-null cell line
Hep3B along with constructs harboring the indicated p53REs upstream to a minimal promoter and the luciferase gene (Cypridinia). Luminescence values were
measured as indication for transcription levels and normalized to a constitutively expressed luciferase gene (Gaussia). Results are of a representative experiment
from three experiments.
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p53 induces P450 expression
p53 binds to p53REs in the DNA-regulatory elements of P450
genes, thereby inducing their transcription
It is well established that p53 induces gene expression by binding
to DNA-regulatory elements termed p53REs in its target genes (26).
For that reason, we scanned the sequence of the genes’ introns, exons
and promoters (10 kb upstream of the genes’ transcription start site—
TSS) for p53REs using the p53MH algorithm (27). Indeed, the distal
promoters of CYP3A4 and CYP3A7 each contained a putative p53RE
and so did the third intron of CYP4F2 and CYP4F3 (Figure 3A). The
p53REs of CYP4F2 and CYP4F3 are completely identical, agreeing
with the notion that the two genes have originated from a gene-duplication event (2). Likewise, the p53REs of CYP3A4 and CYP3A7 are
strikingly similar with only one nucleotide differing between them.
To determine whether these putative p53REs are indeed occupied by p53 in a cellular context, we performed a chromatin immunoprecipitation experiment in which HepG2sh-con cells were treated
with either Nutlin or its solvent as a control and their chromatin was
immunoprecipitated with two different α-p53 polyclonal antibodies
(H47 and CM1). p53 was able to bind all p53REs following Nutlin
treatment as deduced by the enrichment of these DNA segments in
p53-precipitated DNA (Figure 3B). The binding of p53 to the P450
p53REs was similar in its extent to the binding of p53 to the known
p53RE of p21 (Figure 3B). Furthermore, no binding was observed
in the negative control in which primers amplifying a non-relevant
genomic area were used (data not shown).
In order to show that the aforementioned p53REs are functional
(i.e. able to induce transcription in a p53-dependent manner), we
performed a reporter gene assay in which the p53REs were cloned
upstream to a minimal promoter followed by a luciferase reporter
gene. This construct was co-transfected with either a p53-harboring
vector or a control vector to the hepatic-derived, p53-null cell line,
Hep3B. All three p53REs significantly enhanced the expression of
luciferase in the presence of exogenous p53, similarly to the positive control (a construct harboring the consensus sequence of p53RE)
(Figure 3C). Collectively, the findings presented in Figure 3 uncover
the mode of regulation of the p53-dependent induction of P450 genes,
namely, binding to p53REs in DNA-regulatory elements and the subsequent promotion of transcription.
p53 is activated by AMPK to induce the lipid-metabolizing
gene CYP4F2
The stabilization and activation of p53 by Nutlin is due to disruption of the MDM2-p53 complex, which results in degradation of p53.
We were interested in finding more specific p53-activating signals
that promote a more selective induction of gene expression. p53 is
activated following a metabolic stress through an AMPK-dependent
phosphorylation (15,16). Moreover, CYP4F2 is also induced by
AMPK (10). However, the transcriptional activator responsible for
this induction is unknown.
In order to link between these two observations and our findings,
we treated HepG2 cells with AICAR (an AMPK activator) and examined the level of CYP4F2 in HepG2sh-con and HepG2sh-p53. As expected,
AICAR treatment resulted in an elevation in both p53 protein levels
and its phosphorylation, indicating increased activity (Figure 4A).
Interestingly, AICAR was able to fully induce the expression of
CYP4F2 only in HepG2sh-con cells, whereas in HepG2sh-p53 cells the
induction was much less pronounced (Figure 4B). This finding, coupled with published data, points to a scenario in which AMPK is activated, leads to the phosphorylation of p53 (resulting in its activation)
that, in turn, induces the expression of CYP4F2 potentially leading to
fatty acid oxidation (8). This observation agrees with previous findings linking p53 to lipid metabolism (13,18,19,25,28) and suggests a
mechanism for AMPK-stimulated transcription of CYP4F2.
Anticancer chemotherapeutic agents activate p53 to induce their
own clearing enzymes.
DNA damage is another kind of p53-activating signal. Due to their
DNA-damaging attributes, many of the clinically used anticancer
Fig. 4. AICAR, an AMPK activator, induces the expression of CYP4F2 in
a p53-dependent manner. (A) HepG2 cells were treated with the AMPKactivating agent AICAR (500 µM) for 24 h, total protein was extracted and
the protein levels of total p53 and ser-15 phosphorylated p53 were analyzed
in a western blot analysis. (B) HepG2 cells were treated with the AMPKactivating agent AICAR (500 µM) for 24 h, total RNA was extracted and the
mRNA levels of CYP4F2 was analyzed in a QRT–PCR analysis. Results are
of a representative experiment from three experiments.
chemotherapeutic compounds lead to activation of p53 and the subsequent transcriptional activation of p53 target genes (29). In addition, many anticancer chemotherapeutic compounds are metabolized
and thus inactivated by the CYP3A subfamily of enzymes. In order
to assess the p53-dependent induction of CYP3A genes following chemotherapeutics, we treated HepG2 cells with either of three
widely used genotoxic chemotherapeutic drugs—doxorubicin (Dox),
etoposide (Eto) and cisplatin (Cis). Cells were treated with concentrations reflecting the peak plasma concentrations of each agent as
reported in clinical measurements (30–33). These compounds are
known to activate p53 (29) and to be metabolized by CYP3A4 (3). As
expected, all three chemotherapeutics led to phosphorylation of p53
at serine 15 (an indicator of its activation), Dox or Cis treatment also
resulted in marked elevation in p53 protein levels (Figure 5A). These
drugs induced the expression of the CYP3A4 and CYP3A7 genes in a
p53-dependent manner (Figure 5B). In order to assess the functional
effect of this induction, we examined the activity of the CYP3A4
enzyme following treatment with either rifampicin (Rif, a wellknown inducer of CYP3A4), Nutlin, Dox or Eto (we did not examine
the effect of Cis due to its relatively low effect on CYP3A4 mRNA
expression). Remarkably, Nutlin-treated HepG2sh-con cells showed a
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Fig. 5. p53 is activated by chemotherapeutic agents to induce CYP3A expression and activity. (A) HepG2 cells were treated with chemotherapeutic agents (Dox
0.4 µg/ml, Eto 100 µM, Cis 5 µg/ml) for 24 h, total protein was extracted and the protein levels of total p53 and ser-15 phosphorylated p53 were analyzed in a
western blot analysis. (B) HepG2 cells were treated with chemotherapeutic agents (Dox 0.4 µg/ml, Eto 100 µM, Cis 5 µg/ml) for 24 h, total RNA was extracted
and the mRNA levels of the indicated genes were analyzed in a QRT–PCR analysis. (C) HepG2 cells were treated with rifampicin (50 µM), Nutlin (10 µM) or
chemotherapeutic agents (Dox 0.2 µg/ml, Eto 20 µM) for 48 h. CYP3A4 activity was measured and normalized to cell quantity. Asterisks denote a significant
(P < 0.05) elevation, measured by a student t-test compared with both untreated cells and treated HepG2sh-p53 cells. Results are of a representative experiment
from three experiments.
significant increase in CYP3A4 activity, whereas HepG2sh-p53 cells
did not. Similarly, HepG2sh-con treated with either Dox or Eto showed
a significant increase in CYP3A4 activity, this increase was milder in
HepG2sh-p53 cells (Figure 5C). Thus, following its activation, p53 is
able to significantly induce CYP3A enzymatic activity in liver cells
activity.
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Discussion
In this report, we show that p53 regulates several P450 genes
involved in various metabolic pathways. p53 induces the activity
of CYP3A4, which is the main drug-metabolizing enzyme in adult
liver (3). p53 regulates CYP4Fs that metabolize lipids and affects
p53 induces P450 expression
physiological pathways such as inflammation and hypertension (8).
p53 regulates the expression of CYP19A1, an enzyme that converts
androgen to estrogen. CYP21A2, another enzyme induced by p53,
catalyses a crucial step in glucocorticoid synthesis. Finally, the p53induced CYP24A1 enzyme hydroxylates and thus inactivates vitamin
D3 and is considered the main catabolic enzyme of vitamin D (1).
In addition to our findings, p53 was shown previously to reduce the
expression of two mouse P450 genes—Cyp7a1 and Cyp8b1 (19).
The regulation of a variety of P450 genes by p53 establishes a novel
and direct role for p53 in aspects of lipid, steroid, vitamin and drug
metabolism and may help in promoting our understanding of their
associated pathologies.
Until now, delineation of molecular mechanisms for the induction of CYP3A genes was concentrated on nuclear receptors (3). The
results described above unravel another route for the induction of
CYP3As, through activation of p53, its binding to p53REs and the
subsequent transcriptional enhancement of CYP3As. Of note, the
functional p53REs in CYP3A4 is located upstream of its conventional
promoter (Figure 3A). This finding should be considered when studying CYP3A4 induction because many of the studies exploring CYP3A4
induction patterns use a cloned promoter with an approximate length
of 1 kb, which lack the p53RE (e.g. (34,35)). Therefore, some drugs
that may not induce CYP3A4 in these assays would be a ‘false negative’ and would actually induce CYP3A4 through its p53RE in an
endogenous context.
Our findings show that p53 responds to a chemotherapeutic signal
or to Nutlin to transcriptionally induce CYP3A genes in liver cells.
This molecular observation may bear clinical relevance considering
the widespread use of anticancer chemotherapeutic drugs and the
optional future use of Nutlin as an anticancer compound (24,36).
Both Nutlin and genotoxic chemotherapy are administered systematically and thus reach the liver and may trigger hepatic p53 to induce
CYP3A genes. Consequently, the elevated activity level of CYP3A
enzymes may lead to reduced efficacy of the drugs. Similar cases of a
drug inducing its own clearance leading to undertreatment have been
reported in other cases of CYP3A induction (4,5). Moreover, cases of
drug–drug interactions in which a drug induces CYP3A4, leading to
the clearance of co-administered drugs were reported (6). Because
most anticancer treatment regimens are composed of several concomitantly administered drugs (in most cases, at least one of which
is a p53-activating drug), such drug–drug interactions should be carefully considered. A combined administration of a p53-inducing drug
with a second drug that is catalyzed by CYP3A enzymes could prove
to be counterproductive—not only would the first drug’s efficacy be
curtailed by induced CYP3A activity but so would the efficacy of the
second drug.
It is worth mentioning that the examination of P450 induction in
rodent models is problematic because the transcriptional response of
P450 genes varies greatly between species (3) and therefore studies
regarding P450 induction were traditionally done in human cell lines.
Accordingly, treating mice with etoposide or Nutlin did not lead to
CYP3A induction (data not shown).
Our finding linking between chemotherapy treatment and p53 in
the liver rather than in a specific tumor site sheds light on the systemic
roles of p53 in response to chemotherapy. p53 may initiate a hepatic
process leading to chemotherapy clearance potentially resulting in
reduced efficacy. However, this option warrants further investigation.
In conclusion, in addition to the well-established role of p53 in preventing tumorigenesis, we describe here a novel function of p53 in
liver-specific molecular pathways relating to chemotherapy clearance.
The regulation of chemotherapy-clearing enzymes by p53 may influence drug administration regimens and carefully considering it could
prove beneficiary and improve drug efficacy.
Supplementary material
Supplementary Table 1 can be found at http://carcin.oxfordjournals.org/‍
Funding
This work was supported by the Flight Attendant Medical Research
Institute Center of Excellence (ECFP6 grant LSHC-CT-2004–503576);
Yad Abraham Center for Cancer Diagnosis and Therapy. VR is the
incumbent of the Norman and Helen Asher Professorial Chair Cancer
Research at the Weizmann Institute.
Conflict of Interest Statement: None declared.
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Received April 17, 2012; revised September 24, 2012; accepted September
29, 2012