p53 exerts its tumor suppressor function mainly through transcriptional induction of target genes involved in several processes, including cell cycle checkpoints, apoptosis, and regulation of cell redox status. p53 antioxidant function is dependent on its transcriptional activity and proceeds by sequential induction of antioxidant and proapoptotic targets. However, none of the thus far renowned p53 targets have proved able to abolish on their own the intracellular reactive oxygen species (ROS) accumulation caused by p53 deficiency, therefore pointing to the existence of other prominent and yet unknown p53 antioxidant targets. Here, we show that TP53INP1 represents such a target. Indeed, TP53INP1 transcript induction on oxidative stress is strictly dependent on p53. Mouse embryonic fibroblasts (MEF) and splenocytes derived from TP53INP1-deficient (inp1−/−) mice accumulate intracellular ROS, whereas overexpression of TP53INP1 in p53-deficient MEFs rescues ROS levels to those of p53-proficient cells, indicating that TP53INP1 antioxidant function is p53 independent. Furthermore, accumulation of ROS in inp1−/− cells on oxidant challenge is associated with decreased expression of p53 targets p21/Cdkn1a, Sesn2, TAp73, Puma, and Bax. Mutation of p53 Ser58 (equivalent to human p53 Ser46) abrogates transcription of these genes, indicating that TP53INP1-mediated p53 Ser58 phosphorylation is implicated in this process. In addition, TP53INP1 deficiency results in an antioxidant (N-acetylcysteine)-sensitive acceleration of cell proliferation. Finally, TP53INP1 deficiency increases oxidative stress–related lymphoma incidence and decreases survival of p53+/− mice. In conclusion, our data show that TP53INP1 is a major actor of p53-driven oxidative stress response that possesses both a p53-independent intracellular ROS regulatory function and a p53-dependent transcription regulatory function. [Cancer Res 2009;69(1):219–26]

The TP53 gene, which encodes the p53 tumor suppressor, is the most frequently mutated gene in human cancers. p53-dependent response to cell injury involves key processes such as cell cycle arrest, DNA repair, replicative senescence, and apoptosis whenever damage overwhelms repair. Furthermore, p53 continuously protects cells from endogenous, highly mutagenic reactive oxygen species (ROS), including free radicals, produced by the cell itself as respiration products and second messengers during cell signaling. Although cytoplasmic and mitochondrial functions have been described for p53 (1, 2), its most relevant property seems to be its transcriptional activity over target genes that mediate tumor-suppressive functions. A good example of this is the recently reported physiologic antioxidant activity of p53, which is dependent on its DNA-binding domain (3). p53 response to elevated intracellular ROS concentration involves two waves of transcription induction. The first wave starts when cells endure low oxidant concentrations, in which case p53 induces transcription of genes with antioxidant and cell cycle regulatory properties, such as sestrins and p21, to favor the restoration of physiologic intracellular ROS levels (3, 4). When cells endure long-lasting and/or high concentrations of oxidant, p53 will induce a second wave of transcription of proapoptotic genes, such as those encoding Puma and Bax, to promote cell death (3, 5, 6). Interestingly, none of the above-mentioned p53 targets induced during the oxidative stress response is capable of fully restoring physiologic ROS levels in the absence of p53, suggesting that a yet unknown p53 target might be responsible for sensing and/or eliminating ROS surplus.

Tumor protein 53–induced nuclear protein 1 (TP53INP1; also known as TEAP, SIP, and p53DINP1) is a p53 target gene that encodes the TP53INP1 protein (79). Two isoforms of this protein exist, TP53INP1α and TP53INP1β (18 and 27 kDa, respectively), resulting from alternative splicing of the transcript and showing no known functional domain apart from a PEST motif characteristic of short half-life proteins (8, 9). TP53INP1 may act on a p53-positive feedback through its interaction with protein kinases HIPK2 and protein kinase Cδ (PKCδ; refs. 10, 11). These protein kinases enhance p53 protein stability and transcriptional activity through phosphorylation of p53 Ser46 (1012). TP53INP1 expression is induced by different cell stress agents (adriamycin, cisplatin, ethanol, and heat shock) and, most remarkably, by oxidants such as hydrogen peroxide (H2O2) or conditions promoting their formation, such as exposure to UV light or γ-rays (8, 9). In turn, TP53INP1 induces the transcriptional activation of p53 target promoters, such as those of CDKN1A and Bax genes, when expressed ectopically in cell lines (10). Furthermore, consistent with genotoxic stress-induced TP53INP1 transcriptional regulation by p53, p73, and E2F1 (which are transcription factors implicated in cell cycle control and tumor suppression), TP53INP1 overexpression provokes a G1 cell cycle arrest and apoptosis in vitro, suggesting a tumor suppressor activity (10, 13, 14). We recently showed that, compared with wild-type (wt), mice deficient for TP53INP1 present increased susceptibility to colon tumor development in a chronic inflammation setting and a greater production of ROS in inflamed colon (15). Moreover, the low blood ascorbate concentration observed in deficient mice indicates a chronic and systemic oxidative stress, suggesting an antioxidant function for TP53INP1. In this work, we address the question of the implication of TP53INP1 in the p53-dependent oxidative stress response and intracellular ROS regulation. This is achieved by the analysis of primary cells derived from mice deficient for TP53INP1 and p53 and by the generation and analysis of mice and cells that are deficient for both of these stress proteins.

Animals. TP53INP1-deficient (inp1−/−) mice were described elsewhere (15). TP53INP1- and p53-deficient [double knockout (DKO)] mice were obtained by inbreeding inp1−/− and p53−/− mice (16). All mice were kept within the animal facilities and according to the policies of the Laboratoire d'Exploration Fonctionnelle de Luminy (Marseille, France).

Cells. Primary mouse embryonic fibroblasts (MEF) from 14.5-d post-coitum wt, inp1−/−, p53−/−, and DKO embryos were prepared following a standard protocol (17). All experiments were performed using early-passage MEFs of at least two independent preparations per genotype. TP53INP1α-inducible MiaPaCa2 cell line was described elsewhere (18). Cells were cultured in DMEM and 10% (v/v) fetal bovine serum (FBS; Invitrogen) at 37°C, 5% CO2. Viral transduction of MEFs and transfection of MiaPaCa2 cells are described in Supplementary Materials and Methods.

Oxidative stress and intracellular ROS detection. MEFs were platted in triplicate in 35-mm culture dishes (BD Biosciences) and treated with 50 μmol/L H2O2 in DMEM without FBS at 37°C. After 1 h, medium was replaced by DMEM and 10% FBS with or without 5,000 units/mL catalase (Sigma), and cells were allowed to recover for the times indicated. For intracellular ROS detection, cells were further cultured for 15 min at 37°C in DMEM, 10% FBS, and 5 μmol/L dichlorofluorescein (DCF) diacetate (Sigma), which is oxidized in the presence of ROS into green fluorescent DCF, whose fluorescence was assessed by flow cytometry on a FACSCalibur cytometer (BD Biosciences). ROS accumulation rate is expressed as the fold changes of DCF-positive cells between mock- and H2O2-treated MEFs.

Electron spin resonance measurements. Assessment of extracellular release of free radicals from H2O2-pretreated MEFs was performed using electron spin resonance (ESR) technology and the spin trap 5-(diethoxyphosphoryl)-5-methyl-1-pyrroline N-oxide (DEPMPO). For more details, see Supplementary Materials and Methods.

Proliferation assays. One hundred thousand MEFs of each genotype were plated in triplicate in 35-mm dishes and cultured for up to 7 d in conventional medium with or without 10 mmol/L N-acetylcysteine (NAC; Sigma). Medium was renewed every 3 d. Cells were counted daily by trypan blue exclusion to establish growth curves. The number of population doublings (PD) was calculated using the formula PD = ln(Nf/Ni)/ln2, where Nf and Ni are final and initial cell numbers, respectively.

RNA extraction and quantitative reverse transcription-PCR analysis. Total RNA was extracted using Trizol (Invitrogen) and cDNAs were prepared using ImProm-II kit (Promega) following the manufacturer's instructions. Quantitative PCR was performed in a LightCycler (Roche) using the SYBR Premix Ex Taq (Takara Bio). For details of amplification, see Supplementary Materials and Methods.

Histologic analysis. p53−/−, p53+/−, and TP53INP1/p53 double-deficient mice were sacrificed when showing illness and autopsied. Tissues/tumors were collected, formalin fixed, and paraffin embedded. Sections were stained with H&E (Vector Laboratories) and mounted in Eukitt solution (Vector Laboratories) before diagnosis.

Statistics. All statistical analyses were performed using the StatView 5.0 software (SAS Institute, Inc.). Data on MEFs were analyzed using the nonparametric Mann-Whitney U test on triplicate measures. Mice survival was analyzed using the Kaplan-Meier test. All values were expressed as mean ± SE, with significance set at P < 0.05. ESR data were analyzed using two-way ANOVA over the entire incubation period. If a difference was found (P < 0.05), a Bonferroni test was carried out to test for differences among mean values of all groups at each time point.

TP53INP1 induction on oxidative stress is p53 dependent. To establish p53 dependence of TP53INP1 expression on oxidative stress, we derived fibroblasts (MEFs) from wt and p53-deficient (p53−/−) embryos, cultured them for 1 hour with or without H2O2, and allowed cells to recover for 12 hours. Then, we assessed TP53INP1 transcription levels by quantitative reverse transcription-PCR (qRT-PCR). Figure 1 shows the resulting expression data that confirm our previous observations showing strong TP53INP1 induction (>4-fold) in H2O2-treated wt cells (8). In contrast, TP53INP1 expression was severely compromised in p53−/− cells in the absence of stress (70% reduction) and on H2O2 challenge. These data show that induction of TP53INP1 on oxidative stress is totally dependent on p53, although a basal p53-independent TP53INP1 expression exists in unstressed and H2O2-treated MEFs.

Figure 1.

TP53INP1 transcription on oxidative stress is p53 dependent. Histogram shows relative transcript expression of TP53INP1 quantified by qRT-PCR in wt and p53−/− MEFs incubated in serum-free DMEM and without (Mock) or with H2O2 (50 μmol/L) for 1 h and then cultured in conventional medium for 12 h. *, P < 0.005, compared with wt.

Figure 1.

TP53INP1 transcription on oxidative stress is p53 dependent. Histogram shows relative transcript expression of TP53INP1 quantified by qRT-PCR in wt and p53−/− MEFs incubated in serum-free DMEM and without (Mock) or with H2O2 (50 μmol/L) for 1 h and then cultured in conventional medium for 12 h. *, P < 0.005, compared with wt.

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TP53INP1 deficiency induces intracellular accumulation of ROS. To determine whether TP53INP1 possesses an antioxidant function, we cultured wt, TP53INP1-deficient (inp1−/−), p53−/−, and TP53INP1/p53 (DKO) MEFs with or without H2O2 for 1 hour and allowed cells to recover for 3 or 10 hours. Then, we assessed intracellular ROS content by flow cytometry using the DCF probe and determined the percentage of ROS-containing cells (RCC) within each population. In mock conditions, only inp1−/− cells showed a greater percentage of RCC than wt at any time point (Fig. 2A). H2O2 challenge increased the fraction of RCC in all genotypes tested, the highest increase being observed after 3 hours. inp1−/− cells showed more RCC than wt (2- and 1.7-fold at 3 and 10 hours, respectively) on H2O2 challenge. We found more RCC among p53−/− cells compared with wt after H2O2 challenge, consistent with previous reports (3). Double deficiency for TP53INP1 and p53 resulted in RCC increase 3 and 10 hours after oxidative stress compared with wt and p53−/− cells, with only a slight increase 3 hours after stress when compared with inp1−/− cells. Strikingly, there were more inp1−/− than p53−/− RCC with or without H2O2 treatment (3- and 1.5-fold, respectively), suggesting that TP53INP1 action over ROS could be, at least in part, independent of p53. To test this hypothesis, we performed retroviral transduction on E1A/rasV12-transformed p53−/− MEFs to introduce TP53INP1α or TP53INP1β alone or along with p53 and determined the ROS accumulation rate in these cells 3 hours after H2O2 treatment (Fig. 2B). As expected, restoration of p53 expression in p53−/− cells led to a drop of ROS accumulation on oxidant challenge compared with p53−/− cells transduced with empty vectors. Transduction of either TP53INP1α or TP53INP1β was able to drop ROS accumulation in p53−/− cells even below levels reached by p53 restoration alone, and cotransduction of p53 and TP53INP1 isoforms did not significantly improve TP53INP1 antioxidant effect. Therefore, once TP53INP1 is induced on oxidative stress, it seems to play its antioxidant function independently of p53.

Figure 2.

TP53INP1 deficiency causes intracellular ROS accumulation. A, wt, inp1−/−, p53−/−, and DKO MEFs were incubated in DMEM without FBS and without (Mock) or with H2O2 (50 μmol/L) for 1 h and then cultured in conventional medium for 3 and 10 h. Histogram shows percentage of DCF-positive wt, inp1−/−, p53−/−, and DKO RCCs determined by flow cytometry. B, E1A/rasV12-transformed p53−/− MEFs were transduced with empty MSCV-neo, MSCV-TP53INP1α, MSCV-TP53INP1β, empty pLPC, and pLPC-p53 retroviral vectors. Then, cells were submitted to H2O2 treatment as indicated above and allowed to recover for 3 h before DCF staining. Histogram shows ROS accumulation rate (fold changes of mock: H2O2-treated DCF-positive cells) in these cells. *, P < 0.05, compared with p53-restored p53−/− MEFs (white column). C, histograms show DCF mean fluorescence intensity (MFI) of splenocytes, thymocytes, and RBCs from wt and inp1−/− mice stained ex vivo immediately after sacrifice. D, left, dot plot showing side scatter (SSC), indicative of cell granularity, and forward scatter (FSC), indicative of cell size, which allows discrimination of myeloid and lymphoid compartments of the spleen for DCF fluorescence measurement; right, histogram shows DCF levels of spleen myeloid and lymphoid cells from wt and inp1−/− mice. Columns, mean of triplicates and are representative of three independent experiments; bars, SE. P < 0.05, compared with wt (*), inp1−/− (&), and p53−/− (#) MEFs.

Figure 2.

TP53INP1 deficiency causes intracellular ROS accumulation. A, wt, inp1−/−, p53−/−, and DKO MEFs were incubated in DMEM without FBS and without (Mock) or with H2O2 (50 μmol/L) for 1 h and then cultured in conventional medium for 3 and 10 h. Histogram shows percentage of DCF-positive wt, inp1−/−, p53−/−, and DKO RCCs determined by flow cytometry. B, E1A/rasV12-transformed p53−/− MEFs were transduced with empty MSCV-neo, MSCV-TP53INP1α, MSCV-TP53INP1β, empty pLPC, and pLPC-p53 retroviral vectors. Then, cells were submitted to H2O2 treatment as indicated above and allowed to recover for 3 h before DCF staining. Histogram shows ROS accumulation rate (fold changes of mock: H2O2-treated DCF-positive cells) in these cells. *, P < 0.05, compared with p53-restored p53−/− MEFs (white column). C, histograms show DCF mean fluorescence intensity (MFI) of splenocytes, thymocytes, and RBCs from wt and inp1−/− mice stained ex vivo immediately after sacrifice. D, left, dot plot showing side scatter (SSC), indicative of cell granularity, and forward scatter (FSC), indicative of cell size, which allows discrimination of myeloid and lymphoid compartments of the spleen for DCF fluorescence measurement; right, histogram shows DCF levels of spleen myeloid and lymphoid cells from wt and inp1−/− mice. Columns, mean of triplicates and are representative of three independent experiments; bars, SE. P < 0.05, compared with wt (*), inp1−/− (&), and p53−/− (#) MEFs.

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We determined the physiologic relevance of TP53INP1 antioxidant function by measuring ROS content in splenocytes, thymocytes, and RBCs from wt and inp1−/− mice (Fig. 2C). We observed a significant increase of intracellular ROS content in splenocytes but not in thymocytes or RBC from inp1−/− mice compared with wt. Splenocytes of both the myeloid and lymphoid compartments were affected to the same extent in inp1−/− mice (Fig. 2D). Altogether, these data show that TP53INP1 is a negative regulator of intracellular ROS content both under physiologic conditions in splenocytes and under oxidative stress in MEFs.

TP53INP1 deficiency affects the regulation of H2O2 content in MEFs. DCF is a general oxidant indicator rather than a specific marker for H2O2 (19). We determined TP53INP1 implication on intracellular H2O2 regulation by incubating H2O2-pretreated wt and inp1−/− MEFs with catalase (an H2O2 scavenger) before RCC assessment using DCF. Catalase treatment after H2O2 stress abolished RCC differences between inp1−/− and wt MEFs (Fig. 3A), indicating that TP53INP1 deficiency provokes H2O2 accumulation. Evidence that TP53INP1 deficiency triggers an abnormal extracellular release of H2O2-derived free radicals after H2O2 challenge was provided with spin trapping experiments in which the spin trap DEPMPO was added to H2O2-pretreated MEFs during recovery. Regardless of the genotype, no ESR signal was detected in medium samples collected up to 30 minutes after H2O2 removal (Fig. 3B,, trace a). From 30 minutes, addition of DEPMPO led to spin adducts formation, typical spectra of which are shown in Fig. 3B (traces b and d for inp1−/− and wt cells, respectively). Computer simulation of the signals afforded hyperfine coupling constants consistent with mixtures of DEPMPO/·OH as a superimposition of ∼64% trans (aN = 14.06 G, aHβ = 12.73 G, aP = 47.23 G) and 36% cis (aN = 14.06 G, aHβ = 14.06 G, aP = 47.29 G) diastereoisomers (20), and two DEPMPO/alkyl adducts [aN = 14.55 (14.34) G, aHβ = 21.47 (19.34) G, aP = 46.76 (47.14) G] accounting for 25% to 35% of the total signal (see Fig. 3B,, traces c and e). ESR signal was completely abolished when either catalase (Fig. 3B,, trace f) or the metal ion chelator deferoxamine (Fig. 3B,, trace g) was added to the medium from the beginning of the experiment. Figure 3C displays the time course of spin adduct formation in both genotypes, with a burst at 75 minutes following H2O2 removal and still detectable levels after 150 minutes. All along this 150-minute period, DEPMPO adducts detected in samples from wt cells were significantly lower than those from inp1−/− cells (P < 0.001, by two-way ANOVA).

Figure 3.

TP53INP1 deficiency affects antioxidant response to H2O2. A, histogram shows percentages of DCF-positive wt and inp1−/− MEFs after 1-h mock or H2O2 (50 μmol/L) treatment and 3-h recovery in culture medium with or without 5,000 units/mL catalase. B, DEPMPO (10 mmol/L) spin trapping of free radicals released from H2O2-pretreated wt or inp1−/− cells. Representative ESR spectra were recorded from inp1−/− cells 15 min (a) or 75 min (b) after removal of H2O2; c, computer simulation of b showing the participation of DEPMPO/·OH (DEPMPO-OH; 71%) and DEPMPO/alkyls (DEPMPO-R; 29%); d, same conditions as b for wt cells; e, computer simulation of d corresponding to a mixture of DEPMPO/·OH (68%) and DEPMPO/alkyls (32%); f and g, spectra from inp1−/− cells 75 min after H2O2 in the presence of catalase (5,000 units/mL) or deferoxamine (0.1 mmol/L), respectively. Instrumental settings: microwave power, 10 mW; modulation amplitude, 0.497 G; receiver gain, 3.2 × 105; time constant, 20.48 ms; and sweep rate, 2.86 G/s. C, comparative time course of DEPMPO/·OH (top), DEPMPO-alkyls (middle), and total (bottom) spin adduct formation in inp1−/− and wt cells. Two-way ANOVA: P < 0.001 versus wt followed by Bonferroni test (*, P < 0.05 versus wt). Columns, mean (n = 4–6); bars, SE.

Figure 3.

TP53INP1 deficiency affects antioxidant response to H2O2. A, histogram shows percentages of DCF-positive wt and inp1−/− MEFs after 1-h mock or H2O2 (50 μmol/L) treatment and 3-h recovery in culture medium with or without 5,000 units/mL catalase. B, DEPMPO (10 mmol/L) spin trapping of free radicals released from H2O2-pretreated wt or inp1−/− cells. Representative ESR spectra were recorded from inp1−/− cells 15 min (a) or 75 min (b) after removal of H2O2; c, computer simulation of b showing the participation of DEPMPO/·OH (DEPMPO-OH; 71%) and DEPMPO/alkyls (DEPMPO-R; 29%); d, same conditions as b for wt cells; e, computer simulation of d corresponding to a mixture of DEPMPO/·OH (68%) and DEPMPO/alkyls (32%); f and g, spectra from inp1−/− cells 75 min after H2O2 in the presence of catalase (5,000 units/mL) or deferoxamine (0.1 mmol/L), respectively. Instrumental settings: microwave power, 10 mW; modulation amplitude, 0.497 G; receiver gain, 3.2 × 105; time constant, 20.48 ms; and sweep rate, 2.86 G/s. C, comparative time course of DEPMPO/·OH (top), DEPMPO-alkyls (middle), and total (bottom) spin adduct formation in inp1−/− and wt cells. Two-way ANOVA: P < 0.001 versus wt followed by Bonferroni test (*, P < 0.05 versus wt). Columns, mean (n = 4–6); bars, SE.

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Increased proliferation is related to oxidative stress in TP53INP1-deficient cells. We determined whether TP53INP1 plays a physiologic role in cell proliferation by comparing growth rates of wt, inp1−/−, p53−/−, and DKO MEFs. Early-passage MEFs were seeded at equal number and counted daily for 7 days. inp1−/− MEFs proliferate more rapidly than wt, although slower than p53−/−, which proliferate slower compared with DKO (Fig. 4A). Consistently, the number of PDs of each genotype after 7 days confirmed the differences in growth rates shown by daily counts (Fig. 4B,, left). Importantly, addition of the antioxidant NAC during proliferation assays resulted in a restoration of inp1−/−, p53−/−, and DKO number of doublings back to that of wt (Fig. 4B , right), indicating that accelerated growth in deficient cells is related to oxidative stress. Nevertheless, DKO cells conserved a mild but significantly higher number of doublings than wt cells after NAC treatment, suggesting that TP53INP1 may have a redox-independent role in cell cycle regulation. Moreover, TP53INP1 restoration reinstated wt-like proliferation rates in inp1−/− but not in p53−/− E1A/rasV12-transformed MEFs (Supplementary Fig. S1), whereas DKO proliferation was only partially reestablished. In conclusion, these data show that TP53INP1 possesses a physiologic growth-inhibitory function related to its antioxidant properties.

Figure 4.

TP53INP1 and p53 repress cell proliferation through their antioxidant function. Proliferation of early-passage wt, inp1−/−, p53−/−, and DKO MEFs was assessed during 7 d. A, growth curves obtained from daily cell counts for each genotype. B, the histogram shows the number of MEF PDs after 7 d of culture in the presence or absence of 10 mmol/L NAC. Data in A and B are means of triplicates ± SE and are representative of three independent experiments. P < 0.05, compared with wt (*), inp1−/− (&), and p53−/− (#) MEFs.

Figure 4.

TP53INP1 and p53 repress cell proliferation through their antioxidant function. Proliferation of early-passage wt, inp1−/−, p53−/−, and DKO MEFs was assessed during 7 d. A, growth curves obtained from daily cell counts for each genotype. B, the histogram shows the number of MEF PDs after 7 d of culture in the presence or absence of 10 mmol/L NAC. Data in A and B are means of triplicates ± SE and are representative of three independent experiments. P < 0.05, compared with wt (*), inp1−/− (&), and p53−/− (#) MEFs.

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TP53INP1 deficiency affects p53-dependent transcription on oxidative stress. Because expression of p53 target genes could be affected by TP53INP1 deficiency, we searched for modifications of the expression of well-known oxidative stress–induced p53 targets p21 (Cdkn1a), sestrin 2 (Sesn2), Puma, and Bax in inp1−/−, p53−/−, and DKO MEFs compared with wt, in mock- and H2O2-treated cells, using qRT-PCR. Although none of the expressions of these genes was modified in unstressed cells, we observed that their transcripts were drastically reduced in inp1−/−, p53−/−, and DKO cells compared with wt after H2O2 challenge (Fig. 5A). Similarly, oxidative stress–induced expression of p53 homologue TAp73 was abolished in the absence of TP53INP1 (Supplementary Fig. S2). Positive action of TP53INP1 on transcription on H2O2 challenge is not generalized to all p53 targets because transcription of cyclin G1 and Mdm2 was not affected in inp1−/− cells (Supplementary Fig. S3). Furthermore, we assessed the expression of the CD44 gene whose expression is repressed by p53 (21) and noticed that CD44 repression was released in inp1−/−, p53−/−, and DKO cells in the absence of stress (Fig. 5B). Interestingly, these cells failed to up-regulate CD44 expression on oxidative stress. Finally, unstressed inp1−/− splenocytes exhibit higher levels of CD44 expression than their wt counterparts (Supplementary Fig. S4). In conclusion, our data show that TP53INP1 deficiency significantly affects p53-dependent transcription on oxidant challenge.

Figure 5.

TP53INP1 deficiency alters expression of oxidative stress–induced p53 targets. Histograms show relative transcript expression of p21/Cdkn1a, Sesn2, Bax, and Puma in A, and of CD44 in B, as quantified by qRT-PCR in wt, inp1−/−, p53−/−, and DKO MEFs without stress and after H2O2 (50 μmol/L) treatment for the times indicated (* and & are P < 0.05 compared with wt and inp1−/− MEFs, respectively; note that Puma data are ×10−2). Histograms in C show relative transcript expression of p21/Cdkn1a, Sesn2, Bax, and Puma in TP53INP1α-inducible MiaPaCa2 cells transfected with pCAG 3.1 vectors encoding wt p53 or p53 S58A, which were previously treated for 24 h with ponasterone A (10 μmol/L) for TP53INP1α induction, or with a vehicle (DMSO). Bar colors in C indicate homologue to genotypes in A. *, P < 0.05. Target transcript expression was normalized by the corresponding cyclophilin or TBP values. Data are means of triplicates ± SE.

Figure 5.

TP53INP1 deficiency alters expression of oxidative stress–induced p53 targets. Histograms show relative transcript expression of p21/Cdkn1a, Sesn2, Bax, and Puma in A, and of CD44 in B, as quantified by qRT-PCR in wt, inp1−/−, p53−/−, and DKO MEFs without stress and after H2O2 (50 μmol/L) treatment for the times indicated (* and & are P < 0.05 compared with wt and inp1−/− MEFs, respectively; note that Puma data are ×10−2). Histograms in C show relative transcript expression of p21/Cdkn1a, Sesn2, Bax, and Puma in TP53INP1α-inducible MiaPaCa2 cells transfected with pCAG 3.1 vectors encoding wt p53 or p53 S58A, which were previously treated for 24 h with ponasterone A (10 μmol/L) for TP53INP1α induction, or with a vehicle (DMSO). Bar colors in C indicate homologue to genotypes in A. *, P < 0.05. Target transcript expression was normalized by the corresponding cyclophilin or TBP values. Data are means of triplicates ± SE.

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p53 Ser58 is essential for TP53INP1-mediated transcription of oxidative stress–induced p53 targets. Because TP53INP1 favors p53 Ser46 phosphorylation in humans (the homologue of p53 Ser58 in mice; refs. 10, 22), we determined whether Ser58 phosphorylation is necessary for TP53INP1 action on oxidative stress–related p53 targets. For this purpose, we introduced expression vectors encoding either a wt or a Ser58 to Ala mutated p53 (S58A) into a TP53INP1-inducible MiaPaCa2 cell line that lacks functional p53 and in which TP53INP1 expression can be triggered by ponasterone A (18). The absence of TP53INP1 expression in the absence of ponasterone A treatment was confirmed by qRT-PCR (Supplementary Fig. S5). We quantified p21, Sesn2, Bax, and Puma transcripts in these cells (Fig. 5C) and observed that, consistent with results in MEFs, TP53INP1 induction enhanced the transcription of these genes in MiaPaCa2 cells transfected with wt p53. In contrast, cells transfected with p53 S58A failed to induce transcription of the same genes independently of TP53INP1 presence. In conclusion, our data show that p53 Ser58 is crucial for the triggering of p21, Sesn2, Puma, and Bax transcription and for TP53INP1-dependent enhancement of this triggering.

TP53INP1 deficiency increases lymphoma incidence and mortality in p53 heterozygous mice. Finally, we determined the effect of TP53INP1 deficiency on survival and tumor incidence in mice deficient for p53. As shown in Fig. 6A, TP53INP1 deficiency did not affect survival of homozygous p53-deficient mice (i.e., ∼50% of animals died by 5 months regardless of their TP53INP1 status). Tumor incidence and tumor type in p53−/− mice were not significantly altered by TP53INP1 deficiency (data not shown). Nevertheless, TP53INP1 deficiency affects p53 heterozygous mice fate (Fig. 6B). Indeed, whereas >80% of inp1+/+ and inp1+/− mice with a p53+/− background survived at 15 months, 50% of inp1−/− p53+/− animals had died by the same time (P < 0.001). Moreover, lymphoma penetrance was increased in inp1−/− p53+/− mice compared with inp1+/− p53+/− and inp1+/+ p53+/− animals (75%, 29%, and 0%, respectively; Fig. 5C). As TP53INP1 deficiency causes increased oxidative load in vivo, these observations are in accordance with previous data associating lymphoma incidence in p53-deficient animals with increased endogenous oxidative stress (3).

Figure 6.

TP53INP1 deficiency decreases survival of heterozygous p53-deficient mice. Cumulative survival of inp1+/+, inp1+/−, and inp1−/− mice on a p53-deficient homozygous (A) or heterozygous background (B). C, histogram shows number of cases of different tumor types within cohorts of inp1+/+, inp1+/−, and inp1−/− mice on a p53+/− background.

Figure 6.

TP53INP1 deficiency decreases survival of heterozygous p53-deficient mice. Cumulative survival of inp1+/+, inp1+/−, and inp1−/− mice on a p53-deficient homozygous (A) or heterozygous background (B). C, histogram shows number of cases of different tumor types within cohorts of inp1+/+, inp1+/−, and inp1−/− mice on a p53+/− background.

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In this work, we show that TP53INP1 is necessary for controlling intracellular ROS levels both in the absence of exogenous stress and after oxidant challenge, that its induction on oxidative stress is strictly dependent on p53, and that it enhances p53-dependent transcription on oxidative stress. This is the first report of TP53INP1 cell-intrinsic antioxidant function, and it sheds light on our previous observations of chronic oxidative stress in TP53INP1-deficient mice (15). We previously reported that inp1−/− mice are more sensitive than wt to experimentally induced colitis and colon carcinogenesis, associated with increased granulocyte infiltration and colon ROS release. Although in the case of colon inflammation increased ROS in inp1−/− colons could be attributed to a more aggressive response of ROS secreting granulocytes and thus to an indirect effect of TP53INP1 on ROS in vivo, data from primary fibroblasts clearly show that TP53INP1 plays an antioxidant function at the cell level.

Intracellular ROS accumulation observed in inp1−/− MEFs can result from either an increased production or a defective degradation of ROS. Although our experiments do not discriminate between these processes, we can take out some clues about TP53INP1 mechanism of action on regulation of cell ROS content. Abrogation by catalase of differences in RCCs between wt and inp1−/− cells indicates that H2O2 aberrantly accumulates in inp1−/− cells after oxidant challenge. Because TP53INP1 protein sequence does not suggest a potential catalytic activity, it is unlikely that it possesses an intrinsic capacity to produce or detoxify H2O2. Thus, it is possible that TP53INP1 enhances the activity of ROS regulatory enzymes. Indeed, such cofactor activity has already been described for TP53INP1 in the case of p53 regulation, in which TP53INP1 binds HIPK2 and PKCδ kinases and p53 itself to favor p53 phosphorylation (10, 11).

Two ways of H2O2 degradation can be foreseen in our experimental conditions that can be discriminated by the outcome of ESR. The first way is provided by the endogenous antioxidant defenses (i.e., catalase, glutathione peroxidases, and peroxiredoxins) that decompose H2O2 into water and molecular oxygen. Whenever this pathway is defective, excess H2O2 may engage into the Fenton reaction in the presence of metals, yielding HO· that will be trapped by DEPMPO to form ESR-detectable DEPMPO/·OH adducts. Here, we show that inp1−/− cells release more DEPMPO trappable HO· than wt after oxidant challenge (Fig. 3B and C) and that spin adduct formation is inhibited by the chelator deferoxamine (Fig. 3B,, trace g). Therefore, our data suggest that inp1−/− MEFs can either produce more de novo H2O2 or release greater amounts of H2O2 and metal catalysts than wt, or both, which ultimately lead to the observed extracellular DEPMPO/·OH buildup. Two mechanisms unrelated to direct trapping of HO· can yield DEPMPO/·OH: (a) metal-catalyzed nucleophilic addition of water occurring at the double bond of pyrroline N-oxides (20, 2325) and (b) reduction of the protonated form of DEPMPO/O2· by glutathione peroxidase (20, 23, 24). Because none of these two unspecific mechanisms can account for inhibitions by catalase and deferoxamine (Fig. 3B , traces f and g), a dominant pathway for DEPMPO/·OH formation in our study is likely to involve trapping of HO· formed by Fenton reactions with H2O2. This pathway can also explain the concomitant inhibition of secondary DEPMPO/alkyls, which may result from hydrogen abstraction by HO· from a variety of cell components. In further support, cis:trans ratios for DEPMPO/·OH seen here are rather in the range of those reported in vitro with Fenton reagents than of those observed when enzymatic reductive systems or nucleophilic synthesis is involved (20, 24). Therefore, the outcome of ESR assays strongly suggests that TP53INP1 deficiency leads to an improper function of the endogenous antioxidant machinery, directing H2O2 processing toward the Fenton reaction.

Is TP53INP1 action on ROS dependent on p53 function? Although induction of TP53INP1 by oxidative stress is p53 dependent (Fig. 1), overexpression of TP53INP1 is sufficient to decrease ROS content in p53-deficient cells to levels comparable with p53-proficient cells (Fig. 2B). Therefore, TP53INP1 proceeds to its antioxidant function without direct participation of p53. Nevertheless, TP53INP1 seems to collaborate with p53 in transcription induction in response to oxidative stress. Indeed, we found a reduction of the expression of several p53 target genes in H2O2-pretreated inp1−/− MEFs compared with wt, of which some are cell cycle regulators (i.e., p21/Cdkn1a), some antioxidants (Sesn2), and some proapoptotic (i.e., TAp53, Puma, and Bax). Moreover, we report for the first time to our knowledge the dependence of the expression of these genes on TP53INP1-mediated p53 Ser58 phosphorylation. These results reinforce our previous report of TP53INP1 ability to enhance p53 transcriptional activity (10) and indicate that TP53INP1 is the missing piece in Sablina's picture of p53 antioxidant function (3). In this new picture, oxidative stress induces p53 expression, which in turn triggers expression of TP53INP1 that will then enhance, by a mechanism yet to be elucidated, the activity of the endogenous antioxidant machinery (Supplementary Fig. S6). In parallel, TP53INP1 interaction with HIPK2 and PKCδ will lead to p53 Ser58 phosphorylation, thus allowing transcription of p21 and Sesn2 and subsequent cell cycle arrest and ROS detoxification. As redox status is restored to physiologic one, TP53INP1 degradation is driven by its PEST domain, and p53 is subsequently degraded, shutting down the oxidative stress response. In conditions where oxidative stress persists, TP53INP1 enhances transcription of Puma and Bax to induce cell death.

Another interesting finding is the implication of TP53INP1 in repression of CD44 expression in unstressed MEFs and splenocytes, as it settles the notion of a transcription regulatory function of TP53INP1 in basal conditions without exogenous stresses as it has been described for p53 (21). Moreover, CD44 expression has been related to enhanced proliferation, which could provide a piece of an explanation for the enhanced proliferation without p21 expression alteration in inp1−/− MEFs.

About cell proliferation, we show for the first time here that p53 and TP53INP1 negatively regulate cell growth through their antioxidant properties. This is particularly remarkable because the action of these proteins at the G1 checkpoint has thus far been attributed exclusively to their transcriptional activity on cell cycle regulatory genes. In fact, antioxidant and transcriptional activities of these two proteins may act synergistically on cell cycle because low concentrations of oxidant are necessary for S-phase progression (26). In this way, TP53INP1 and p53 could control cell cycle progression by keeping ROS in check and, when necessary, by inducing G1 checkpoint.

Finally, TP53INP1 deficiency aggravates the outcome of p53 heterozygous mice as it enhances lymphomagenesis (Fig. 6), presumably promoted by chronic oxidative stress. These results are consistent with our previous report of TP53INP1 role in colon tumor suppression through ROS regulation (15). Although TP53INP1 deficiency alone is not sufficient to induce spontaneous tumors in mice, it seems to be a suitable “second hit” that allows tumor promotion after a first transforming event, such as chemical mutagenesis or loss of a p53 allele. Based on the data presented in this study, we hypothesize that loss of TP53INP1 favors malignant transformation by increasing genetic instability through ROS accumulation and by allowing uncontrolled proliferation of a damaged cell. Accordingly, we observed the loss of TP53INP1 during the progression of several cancers, particularly during pancreatic carcinogenesis, and we showed that loss of the TP53INP1 protein is caused by the action of protumoral miRNA-155 (18). Therefore, reestablishment of TP53INP1 expression seems as a promising therapeutic strategy for the treatment of cancers in which oxidative stress is recognized as a promoting factor, such as those related to chronic inflammation.

No potential conflicts of interest were disclosed.

Note: Supplementary data for this article are available at Cancer Research Online (http://cancerres.aacrjournals.org/).

C.E. Cano and J. Gommeaux contributed equally to this work.

Grant support: Institut National de la Santé et de la Recherche Médicale, Centre National de la Recherche Scientifique, Institut National du Cancer, and La Ligue Nationale Contre le Cancer. C.E. Cano and J. Gommeaux were supported by La Ligue Nationale Contre le Cancer and the Association pour la Recherche sur le Cancer.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

We thank S. Soddu for wt p53 and S58A expression vectors, G. Warcollier and F. Gianardi for animal care, and M.N. Lavaud for technical assistance in histologic analysis.

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Supplementary data