Positive and negative feedback loops in the p53 and mRNA 3′ processing pathways

Abstract

Although the p53 network has been intensively studied, genetic analyses long hinted at the existence of components that remained elusive. Recent studies have shown regulation of p53 at the mRNA level mediated via both the 5′ and the 3′ untranslated regions and affecting the stability and translation efficiency of the p53 mRNA. Here, we provide evidence of a feedback loop between p53 and the poly(A)-specific ribonuclease (PARN), in which PARN deadenylase keeps p53 levels low in nonstress conditions by destabilizing p53 mRNA, and the UV-induced increase in p53 activates PARN deadenylase, regulating gene expression during DNA damage response in a transactivation-independent manner. This model is innovative because it provides insights into p53 function and the mechanisms behind the regulation of mRNA 3′ end processing in different cellular conditions.

Downstream signaling in the p53 pathway includes several cellular responses. The expression of a large number of genes involved in DNA repair, cell cycle arrest, and/or apoptosis is regulated by transactivating properties of p53. This occurs via specific DNA binding of the p53 protein to a p53 response element that is found either in promoters or introns of target genes (1). Transactivation-independent functions of p53 have also been described (2). For example, certain microRNAs (miRNAs) are regulated by p53, and these miRNAs cause dramatic changes in gene expression, offering an indirect p53-mediated control of gene expression at the posttranscriptional level (3). Recently, we showed that p53 can inhibit mRNA 3′ cleavage through its interaction with the cleavage stimulation factor 1 (CstF1) (4). CstF1 can also interact with poly(A)-specific ribonuclease (PARN) deadenylase, and the CstF1/PARN complex formation has a role in the regulation of gene expression by inhibition of mRNA 3′ cleavage and activation of deadenylation upon DNA damage (5). PARN, an mRNA decay enzyme, has been studied extensively in vitro at the biochemical levels but very little is known of its biological targets and its role in different cellular conditions. Recently, it has been shown that PARN regulates the expression of genes involved in mRNA metabolism, transcription, and cell motility in mouse myoblasts (6). Our studies indicate that the CstF/PARN complex can decrease the mRNA levels of housekeeping genes under DNA-damaging conditions and of genes involved in cell growth and differentiation under nonstress conditions (5).

Almost all eukaryotic mRNA precursors, with the exception of histones, undergo a cotranscriptional cleavage followed by polyadenylation at the 3′ end. This first round of polyadenylation is considered a default modification for most mRNAs and confers stability. In contrast, activation of deadenylation alters the length of poly(A) tails, affecting mRNA stability, transport, or translation initiation, and hence gene expression (7). Thus, mechanisms controlling deadenylation are highly regulated and play key roles in cellular responses, such as mRNA surveillance, DNA damage response (DDR), and tumor progression, as well as cell development and differentiation (5, 8–11). Deadenylation of mRNA is regulated by miRNAs, adenylate-uridylate–rich element (ARE) binding proteins, polyadenylation factors, and RNA binding (RB) factors that recognize cis-acting sequences in the target. About 12% of mammalian mRNAs bear important regulatory signal AREs in their 3′ untranslated regions (UTRs), which have been shown to play significant roles in mRNA stability regulation (12). PARN has been shown to be involved in ARE-mediated deadenylation and to promote tristetraprolin (TTP)-directed deadenylation in vitro (13). KH-type splicing regulatory protein recruits PARN to ARE-containing mRNAs to initiate the poly(A) tail shortening that is followed by exosome-mediated degradation (14). Interestingly, tumor suppressors, such as breast cancer type 1 susceptibility protein (BRCA1) and BRCA1-associated RING domain protein (BARD1), associated to the polyadenylation factor CstF1 have been shown to regulate deadenylation by functional interactions with PARN deadenylase (5).

Extending those studies, here we are investigating the possibility that p53 might regulate not only mRNA 3′ cleavage (4) but also PARN-dependent deadenylation in different cellular conditions. As part of these studies, we also identified the mRNA targets of PARN in nonstress conditions. Our results provide evidence of a unique feedback loop between p53 and PARN, in which PARN deadenylase keeps p53 levels low in nonstress conditions by destabilizing p53 mRNA through its 3′ UTR, and the UV-induced increase in p53 activates PARN, representing a mechanism of gene expression regulation in a transactivation-independent manner.

Results

To investigate the role of p53 in deadenylation, we used a group of isogenic cell lines that express different levels of p53 (Fig. 1A and Fig. S1): the colon cancer HCT116 and p53-null HCT116 cell lines, the colon carcinoma RKO and RKO-E6 (low p53 levels) cell lines, and the mouse embryonic fibroblasts (MEFs) and p53-null MEFs. Nuclear extracts (NEs) from those cells were assayed for deadenylation activity using a radiolabeled L3(A₃₀) RNA substrate as described (5). As described in previous studies (5), Fig. 1A shows that deadenylation activity in NEs of RKO cells treated with control siRNA increased significantly after UV treatment. Interestingly, siRNA-mediated knockdown of p53 in RKO cells abolished the UV-induced activation of deadenylation, suggesting that p53 levels might activate deadenylation. Consistent with this, RKO-E6, p53-null HCT116, and p53-null MEFs cells did not show UV-induced activation of deadenylation, which was observed in RKO, HCT116, and MEFs cells. Importantly, only increasing amounts of either full-length His-p53 (Fig. 1 B and C) or the C-terminal fragment of p53 (Fig. 1C) can induce deadenylation in a reaction using a limited amount of His-PARN in a cell-free assay, suggesting that this is a transactivation-independent function of p53. However, neither the two p53 derivatives that lacked the C-terminal region of p53 nor GST alone had an effect on the deadenylation reaction (Fig. 1C). None of the His-p53 derivatives were able to deadenylate the substrate in the absence of His-PARN (Fig. 1C). To further analyze the role of p53 in deadenylation, we examined the physical association of p53 with PARN. Our pull-down (Fig. 2 A–C) and coimmunoprecipitation assays (Fig. 2D) indicate that p53 can form (a) protein complex(es) with PARN in NEs from RKO cells in nonstress conditions and after UV treatment. The results showed that the C-terminal domain of PARN (Fig. 2B), which has been described to interact with CstF1 and cap-binding protein 80 (CBP80) (5), and the C-terminal domain of p53 (Fig. 2C), which has been described to have regulatory functions in DDR (15), are important for the complex formation. Because samples were treated with RNase A, the observed interactions were probably not due to an RNA tethering effect. These results indicate that the same region of p53 required for binding PARN (Fig. 2B) is necessary for activating PARN deadenylase (Fig. 1C). Together these results indicate that p53 can interact with PARN to form (a) complex(es) and that p53 expression levels can activate PARN deadenylase and, therefore, might regulate gene expression.

Fig. 1.

The levels of deadenylation correlate with expression levels of p53. (A) Samples from cells expressing different levels of p53 show different levels of deadenylation. A representative deadenylation reaction from three independent assays is shown. siRNA-mediated knockdown of p53 abolishes UV-induced activation of deadenylation in RKO cells. NEs of the indicated cells treated with UV irradiation and allowed to recover for 2 h were analyzed for radiolabeled L3(A₃₀) deadenylation as described (4, 5). NEs from RKO cells treated with p53/control siRNA and UV irradiation were also analyzed. Positions of the polyadenylated RNA L3(A₃₀) and the L3 deadenylated product are indicated. Numbers beneath gel lanes indicate relative deadenylation (RD). RD was calculated as [L3 fragment/(L3 fragment + L3 (A₃₀)] × 100. Quantifications were done with ImageJ software (http://rsb.info.nih.gov/ij/). (B) p53 can activate PARN-dependent deadenylation in vitro. Deadenylation assays using different concentrations of His-PARN were performed in the presence of capped L3(A₃₀) RNA substrate as described (5) and increasing amounts of His-p53. Reactions were analyzed as in A. (C) The C-terminal domain of p53 activates PARN in vitro. Deadenylation assays were performed as in B with the addition of either full-length p53 (FL) or His-p53 derivatives. The truncated forms of p53 used in this assay were described before (4) and include p53 amino acids 1–293, p53 amino acids 94–293 (DNA binding domain), and p53 amino acids 94–393.

Fig. 2.

p53 interacts with PARN to form a protein complex. (A) Immobilized His-PARN (Left) or His-p53 (Right) on nickel beads were incubated with NEs from untreated or UV-treated RKO cells. A representative pull-down reaction from three independent assays is shown. Equivalent amounts of the pulldowns (PD) and supernatants (SN) were analyzed and proteins were detected by immunoblotting with the indicated antibodies. His-p53 and His-PARN constructs were previously described (4). Twenty percent of the NE used in the pull-down reactions is shown as input. The basal level of the proteins was arbitrarily set at 1.0 in the first lane, and relative fold change of each protein level is shown below each lane. (B) C-terminal domain of PARN interacts with p53. Pull-down assays were performed as in A using full-length, N-terminal domain (NTD) or C-terminal domain (CTD) of His-PARN. (C) C-terminal domain of p53 interacts with PARN. Pull-down assays were performed as in A with full-length or the His-p53 derivatives described in Fig. 1D. (D) PARN and p53 coimmunoprecipitation from NEs of HeLa cells. A representative immunoprecipitation (IP) reaction from three independent assays is shown. The NEs were immunoprecipitated with anti-p53 and anti-PARN. Equivalent amounts of the SN and the pellets (IP) were resolved by SDS/PAGE and proteins were detected by immunoblotting using antibodies against PARN, p53, and topoisomerase II (Topo II) as described (4, 5). Twenty percent of the NE used in the IP reaction is shown as input.

Because PARN is involved in ARE-mediated deadenylation (13), promotes TTP-directed deadenylation (16), and decreases mRNA levels of ARE-containing genes under nonstress conditions (5), we decided to extend these studies and determine which mRNAs might be regulated by PARN using microarray assays. Nuclear RNA samples were isolated from HeLa cells treated with control or PARN siRNAs under nonstress conditions and analyzed by microarray. Pathway analysis of regulated genes using Ingenuity Systems applications indicated that the p53 signaling pathway is the most significantly affected by PARN knockdown in nonstress conditions (Fig. 3A and Tables S1 and S2), suggesting that PARN expression has a specific effect on the expression of genes associated with p53-mediated signaling in those conditions. In addition, the p53-related gene network was found to be the most significantly regulated by network analysis and transcription factor analysis (Fig. S2). We further confirmed the effect of PARN knockdown on the abundance of several transcripts in the p53 signaling pathway by quantitative (q) RT-PCR (Fig. S3). Although there was a good correlation between the change observed in the microarray and that determined by qRT-PCR, the qRT-PCR showed changes of a greater magnitude than the array. These results support our previous study that showed that PARN can promote deadenylation and mRNA instability of FBJ osteosarcoma oncogene and myelocytomatosis oncogene (c-myc), keeping their expression levels low under nonstress conditions (5). Because PARN is a deadenylase, we expected an increase in the steady-state levels of its targets by PARN knockdown. However, our results show both up- and down-regulation of transcripts, suggesting complex effects of PARN knockdown on the expression of these mRNAs. Importantly, consistent with the pathway analysis results, PARN knockdown resulted in a significant increase not only of p53 mRNA but also of p53 protein levels (compare lanes 1 and 2 in Fig. 3B Right), reaching expression levels similar to that observed after UV treatment (compare lanes 2 and 3 in Fig. 3B Right). After UV treatment, the changes in the levels of p53 mRNA and p53 protein were PARN-independent, indicating there is (are) other mechanism(s) involved in the regulation of p53 expression during DDR.

Fig. 3.

PARN deadenylase significantly affects the cellular expression of genes of the p53 pathway under nonstress conditions. (A) Pathway analysis of significantly regulated genes by PARN. Nuclear RNA samples isolated from HeLa cells, treated with siRNAs targeting PARN or control, were analyzed using the Human Gene 1.0 ST GeneChip (Affymetrix) array. Significant genes were selected by t test (P value < 0.05). Analysis of canonical pathways was conducted by using data from Ingenuity Systems (www.ingenuity.com). The bar graph shows significance of pathway for regulated genes. P values were calculated using the Fisher’s exact test, and the −log (P value) values are displayed. Only the top five pathways are shown. Data represent three independent experiments. Refer also to Fig. S2. (B) p53 mRNA and protein levels are affected by PARN expression. qRT-PCR and Western blot analysis of p53 expression after UV treatment using RNA or protein samples, respectively, from cells treated with control/PARN siRNA. A representative Western blot from three independent assays is shown. Topo II was used as loading control. The basal level of the proteins was arbitrarily set at 1.0 in the first lane, and relative fold change of each protein level is shown below each lane. (C) PARN regulates p53 mRNA half-life. mRNA decay rates for p53 and ACTIN, a non-PARN target gene, were determined by qRT-PCR at different time points following PARN/control siRNA- and Act-D treatment. The relative half-life of the p53 transcript was calculated from three independent samples. Errors represent the SD derived from three independent experiments. Western blot analysis of PARN expression after PARN/control siRNA and Act-D treatment is also shown. (D) PARN regulates p53 mRNA poly(A) tail length. Nuclear RNA from PARN/control siRNA-treated cells was reverse-transcribed using an oligo(dT)-anchor primer and amplified using an oligonucleotide that hybridizes within the 3′ UTR of p53 mRNA. The products were separated on a nondenaturing PAGE and detected by ethidium bromide staining. An RT-PCR product from a non-PARN target gene (ACTIN exon 3–4) was used as a loading control. A representative PAGE from three independent assays is shown. Molecular weight standard (MWS, 100-bp ladder from Promega) is also included. (E) PARN knockdown and UV treatment induce similar apoptotic responses in a p53-dependent manner. The DNA fragmentation was calculated from three independent samples.

To determine the effect of PARN expression on the stability of p53 mRNA, we compared mRNA decay rates of p53 transcript in cells treated with control- or PARN-siRNA. The half-life of the p53 transcript was analyzed by qRT-PCR of nuclear RNA samples taken at different time points from PARN/control siRNA- and actinomycin D (Act-D)-treated cells. Previous observations have shown a similar half-life for the p53 transcript (17). Our results indicate that PARN knockdown significantly stabilized the p53 transcript (Fig. 3C). Because PARN is a deadenylase, the stabilization of p53 mRNA by its knockdown might be due to changes in the poly(A) tail length. Importantly, as shown in Fig. 3D, siRNA-mediated knockdown of PARN elongated the poly(A) tail length of p53 mRNA (quantification is shown in Fig. S4). Together, our results indicate that the PARN deadenylase affects p53 expression by regulating poly(A) length and hence mRNA stability. The biological relevance of this observation is supported by the fact that PARN knockdown induces an apoptotic response similar to that observed after UV treatment in a p53-dependent manner (Fig. 3E). Our results indicate that UV treatment and PARN knockdown have a similar effect on cell death in cells expressing normal levels of p53 (HCT116 and RKO). RKO-E6 cells showed levels of apoptosis similar to those in RKO cells because the disruption of wild-type p53 function by E6 expression results in loss of p53-dependent DNA repair but not UV-induced apoptosis (18). Consistent with Ford and Hanawalt (19), p53-null HCT116 cells did not show an induction of apoptosis after either UV treatment or PARN knockdown, suggesting that the effect of PARN expression on cell death is p53-dependent. Together these results indicate that PARN plays a role in controlling p53 mRNA steady-state levels and p53 expression in nonstress conditions. Although many reports have been published about control of p53 protein expression and its effect on downstream pathways, very little is known of the mechanisms behind the control of p53 mRNA steady-state levels in different conditions (20).

Because most of the regulatory elements involved in PARN-mediated regulation of mRNA stability are located in the 3′ UTR of the genes, we decided to determine whether the PARN-induced decrease of p53 mRNA levels under nonstressed conditions is through this region of p53. The firefly luciferase assay was used with constructs under the control of either the p53 3′ UTR or the vector 3′ UTR (Fig. 4A). A significant increase in firefly/Renilla ratio for the construct with the p53 3′ UTR relative to the control construct was detected in RKO/RKO-E6 and HCT116/HCT116 p53^−/− cells treated with PARN siRNA (Fig. 4B). RNA immunoprecipitation (RIP) assays using antibodies against PARN showed that p53 mRNA can form a complex with PARN in samples from cross-linked RKO cells (Fig. 4C and Fig. S5), indicating that PARN can regulate p53 mRNA stability by, most probably, an indirect association to the 3′ UTR. RIP assays also showed that PARN can form a complex with c-myc RNA, which is another target of PARN deadenylase (5). Recently, a G-quadruplex structure that protects the p53 mRNA from degradation upon stress by binding to heterogeneous nuclear ribonucleoprotein H/F has been described (21). This structure, which is located downstream of the 3′ cleavage site, was not included in our luciferase construct (Fig. 4A). Interestingly, the 3′ UTR of p53 mRNA also contains ARE that associates with ARE-binding proteins, such as wild-type p53-induced gene 1 (22) and human antigen R (23) and regulates p53 mRNA steady-state levels. Importantly, the replacement of the ARE sequence from the p53 3′ UTR (noARE construct) significantly increases the firefly/Renilla ratio compared with the WT p53 3′ UTR construct (Fig. 4D), showing that the AREs can decrease mRNA stability and hence expression of the luciferase-p53 3′ UTR construct. Interestingly, the siRNA-mediated knockdown of PARN only increases the expression ratio of firefly/Renilla luciferase from the constructs carrying the AREs but not from the constructs without the AREs (Fig. 4E), indicating that the AREs in the p53 3′ UTR are necessary for PARN-mediated regulation of p53 expression. Supporting this idea, our RIP assays indicate that PARN can form a complex with the luciferase mRNA carrying the 3′ UTR of p53 and this is abolished when AREs are replaced by other sequences (Fig. 4F). RKO/RKO-E6 and HCT116/HCT116 p53^−/− cells showed similar ratios for the expression of firefly/Renilla luciferase (Fig. 4 D and E) and for the RIP assays with different constructs (Fig. 4F), indicating that the PARN-mediated regulation of p53 expression and PARN binding to AREs are p53-independent (Fig. S6). This was confirmed using RNA pull-down assays with in vitro transcribed biotinylated RNAs, either with sequences of the p53 3′ UTR or ARE-replaced p53 3′ UTR, and NEs from RKO cells. Our results indicate that the RNA with the p53 3′ UTR pulled down PARN from the NEs, and this RNA–PARN interaction was lost in the absence of the ARE sequence (Fig. 4G). A weak nonspecific interaction of p53 with the 3′ UTR of its own RNA was detected (Fig. 4G and Fig. S7). Together, these results indicate that the AREs in the p53 3′ UTR are important for the PARN-mediated regulation of p53 mRNA steady-state levels and that this regulation is p53-independent.

Fig. 4.

PARN regulates p53 expression through ARE sequence present in the 3′ UTR of p53 mRNA. (A) Diagram of firefly luciferase reporter constructs with different 3′ UTR sequences from the p53 gene. Polyadenylation signals (PAS) are indicated. (B) Constructs carrying the p53 3′ UTR (p53) or not (vector) were transfected in cells treated with PARN or control siRNAs. The ratios of the firefly/Renilla values for the p53 construct relative to the vector construct are shown. The firefly/Renilla values were calculated from three independent samples. Errors represent the SD derived from three independent experiments. (C) PARN can interact with p53 and c-myc mRNAs under nonstress conditions. The extracts were immunoprecipitated with either anti-PARN or IgG antibodies. The endogenous nuclear RNA immunoprecipitated with the antibodies was quantified by qRT-PCR using primers specific for each gene. The qRT-PCR values were calculated from three independent samples. (D) Constructs carrying the p53 3′ UTR (p53) or ARE-replaced p53 3′ UTR (noARE) were transfected in cells. The firefly/Renilla values were as in A. (E) Luciferase assay done as in D using cells treated with control or PARN siRNA. The ratio of the firefly/Renilla values obtained for each construct in PARN knock-down cells relative to control siRNA-treated cells are shown. (F) ARE sequences present in the 3′ UTR are involved in the interaction of PARN with p53 mRNA under nonstress conditions. RIP analysis of samples cells transfected with luciferase constructs carrying either the p53 3′ UTR (p53) or ARE-replaced p53 3′ UTR (noARE) was performed as in C. The ratio of the fold change for p53/no-ARE RNA values obtained for each construct is shown. (G) PARN interacts with the p53 3′ UTR in an ARE-dependent manner. RNA pull-down experiments were performed using biotinylated RNA of the p53 3′ UTR or ARE-replaced p53 3′ UTR and NEs from RKO cells. *An overexposed film is included to show the weak nonspecific interaction of p53 with the 3′ UTR of its own mRNA. A representative pull-down reaction from three independent assays is shown.

Discussion

In this study we provide evidence of a unique feedback loop between p53 and PARN deadenylase, in which PARN keeps p53 levels low in nonstress conditions by destabilizing p53 mRNA, and the UV-induced increase in p53 activates PARN deadenylase regulating gene expression during DDR in a transactivation-independent manner. Several lines of evidence support this model. First, the C-terminal domain of p53 can activate PARN-dependent deadenylation in vitro and p53 expression levels correlate with levels of mRNA deadenylation (Fig. 1). Second, our results show the direct interaction of the C-terminal domain of p53 with the C-terminal domain of PARN and the existence of protein complexes of these factors in cellular NEs (Fig. 2). Third, PARN significantly affects the cellular expression of genes in the p53 pathway under nonstress conditions and the stability and poly(A) length of the p53 mRNA (Fig. 3 A–D). Fourth, PARN knockdown and UV treatment induce a similar increase in p53 expression and apoptotic responses (Fig. 3 B and E). Finally, PARN regulates p53 expression through the ARE sequence present in the 3′ UTR of p53 mRNA (Fig. 4). Taken together, our results provide insights into p53 function and the mechanisms behind the regulation of mRNA 3′ end processing in different cellular conditions.

The study presented here shows an alternative mechanism to regulate the expression levels of p53 based on the control of the steady-state levels of p53 mRNA by PARN deadenylase under nonstress conditions (Fig. 5A). Supporting this, our previous work indicates that PARN has a role in decreasing the levels of short-lived mRNAs involved in the control of cell growth, DDR, and differentiation, keeping their expression levels low under nonstress conditions (5). Under stress conditions, the induction of p53 expression is associated with a decrease in the levels of total poly(A) mRNA (24). Because mRNA poly(A) tails are important for the regulation of mRNA stability, it is possible that these changes of poly(A) mRNA levels might represent another mechanism of p53-mediated control of gene expression. In fact, our studies indicate that an increase in the expression of p53 inhibits the mRNA 3′ cleavage step of polyadenylation (4) and induces PARN deadenylase activity (Fig. 1), suggesting that the p53 associated to the PARN/CstF/BARD1 complex might regulate gene expression by controlling the steady-state levels of mRNAs (Fig. 5B). Considering that the p53 pathway is tightly controlled in cells (reviewed in ref. 25), the p53-associated control of mRNA 3′ processing machinery could represent an indirect mechanism to repress target gene expression at the posttranscriptional level. The antiproliferative factor BTG2 represents another example of a general activator of mRNA deadenylation by its direct interaction with the Pop2–Caf1 and Ccr4 deadenylases (26). This model is consistent with the idea proposed by Singh et al. (10) that the interaction of the 3′ processing machinery and factors involved in the DDR/tumor suppression might result in cell-specific 3′ processing profiles.

Fig. 5.

Model for the regulation of expression of genes in the p53 pathway by PARN deadenylase-associated p53 in different cellular conditions. (A) PARN deadenylase decreases the stability of the p53 mRNA in nonstress conditions. The AREs in the 3′ UTR of the p53 mRNA have an important role in this regulatory process. (B) Under DNA damage conditions, p53 protein accumulates, allowing its association to and activation of PARN deadenylase resulting in the decrease levels of target mRNAs in the p53-dependent DDR pathway.

Control of deadenylation could represent a mechanism to regulate gene expression in different cellular conditions, such as development, stress treatment, or different metabolic conditions. Supporting this idea, recently it has been shown that PARN regulates the expression of genes involved in mRNA metabolism, transcription, and cell motility in mouse myoblasts, resulting in PARN-dependent regulation of cell motility and wound healing in those cells (6). Indeed, our Gene Ontology analysis also revealed significant down-regulation of genes involved in similar pathways (Table S3), such as structure morphogenesis, cell adhesion, cell migration, and so on. Consistently, our microarray data showed a decrease in the abundance of mRNA for several genes involved in cell motility, such as adenosine A2b receptor, ankyrin repeat-containing domain 54, and collagen alpha-2(I) chain in PARN knock-down cells. However, the p53 signaling pathway was not reported by Lee et al. (6), suggesting cell-specific functions of PARN. Like Lee et al. (6), we also observed a decrease in the steady-state levels of some transcripts by PARN knockdown. However, it is not clear whether this reflects the function of PARN per se or is the indirect consequence of PARN’s effect on genes involved in other mRNA metabolic pathways, such as transcription and RNA processing factors.

The characterization of the regulatory elements in the 3′ UTR of p53 and the factors involved in this PARN-dependent regulatory pathway may allow us to better understand the mechanisms that control p53 expression and to find alternative strategies for treating tumorigenesis and metastasis in various cancers.

Materials and Methods

Tissue Culture Methods and DNA Damaging Agents.

HeLa, RKO, RKO-E6, HCT116, HCT116 p53^−/−, MEF, and MEF p53^−/− cell lines were cultured and UV-treated as described (4, 5, 27).

Preparation of NEs.

After UV treatment, NEs were prepared from harvested cells essentially as described (4, 5).

Deadenylation Assays.

³²P-labeled L3(A₃₀) substrates were prepared and analyzed as in ref. 5. Protein concentrations of the extracts were equalized by Bradford assays (Bio-Rad) before use in deadenylation reactions.

Pull-Down Assays.

One microgram of His-PARN bound to Ni magnetic beads was incubated with 30 μL of NEs from RKO cells and then analyzed as described (4, 5). His-p53 and His-PARN constructs were previously described (4).

IP Analysis.

Total protein (100 μg) from the indicated NEs was immunoprecipitated with the polyclonal antibody against PARN (H-105; Santa Cruz Biotechnology) and p53 (SC-126; Santa Cruz Biotechnology) as described (4, 5).

PARN- and p53-siRNA Knockdown.

The siRNAs specific for human p53, PARN, and the control RNA duplex were synthesized by Dharmacon RNA Technologies. siRNA and UV treatments were as described (4, 5).

RNA Purification and Microarray Analysis.

Nuclear RNA was purified from HeLa cells using the RNeasy kit (Qiagen) following manufacturer’s protocol. The RNA concentrations of the RNA samples obtained under different conditions were equalized. Equivalent amounts of purified RNA were used in microarray analysis. The GeneChip Human Gene 1.0 ST (Affymetrix) expression array was used. Microarray data were normalized using the Robust Multichip Average method.

Plasmid Constructs.

Luciferase vector pEZX-MT01 with TP53 miTarget microRNA 3′ UTR target clones (product ID HmiT054283) was purchased from GeneCopoeia. Mutations in the ARE sequence of p53 3′ UTR were introduced with the QuikChange Lightning Site-Directed Mutagenesis Kit (Agilent Technologies) and the following primers 5′-GGGTCAATTTCC GTTCGCGAATTCTGTTCTGATCTGCTTTTTCTTTGAGACTGGG-3′ and 5′-CCCAGTCT CAAAGAAAAAGCAGATCAGAACAGAATTCGCGAACGGAAATTGACCC-3′, following the manufacturer’s instructions. Plasmids were sequenced to confirm the presence of the mutation. Twenty-four micrograms of the different luciferase constructs were transfected into cells using Lipofectamine 2000 reagent (Invitrogen).

Luciferase Assay.

Cells were cotransfected with the luciferase constructs indicated and either siRNA-targeting PARN or control siRNA. Forty-eight hours after transfection cells were harvested and a dual luciferase assay was performed using a Luc-pair miR Luciferase kit from GeneCopoeia following manufacturer’s instructions.

RIP Assays.

IP of nuclear RNA–protein complexes was performed as described (28) using antibodies against PARN (H-105) or control rabbit IgG (Sigma).

RT-qPCR Assays.

As described before (4, 5), equivalent amounts (2 μg) of purified RNA were used as a template to synthesize cDNA using random hexamer primers, oligo-d(T) primers, and GoScript Reverse Transcriptase (Promega). Relative levels were calculated using the ΔCτ method.

Cell Death ELISA Assay.

Fragmentation of DNA after induction of apoptosis was determined by photometric enzyme immunoassay (Cell Death Detection ELISA^PLUS; Roche Applied Science) as recommended by the manufacturer.

RNA Isolation and qRT-PCR Analysis of mRNA Half-Lives.

Control and PARN knockdown RKO cells (see above) were treated with Act-D (8 µg/mL) for 30 min before the beginning of the time course. Nuclear RNA was purified at different time points using RNeasy Mini Kit (QIAGEN) according to the manufacturer’s directions.

RNA Pull-Down.

Biotin-labeled RNAs were in vitro transcribed with the biotin RNA labeling mix (Roche) and T7 RNA polymerase (Promega) following manufacturer’s instructions. Biotinylated RNA was incubated with 1 mg of NEs and then analyzed as described (17).

RACE-poly(A) Test Assays.

Nuclear RNA from RKO cells treated with PARN/control siRNA was reverse-transcribed using oligo (dT)-anchor primer (5′-GGGGATCCGCGGTTTTTTTTTT-3′) and GoScript Reverse Transcriptase (Promega). One microliter of each cDNA was used for PCR amplification by GoTaq PCR mix (Promega) using p53 3′ UTR-specific primer [5′-CTGCATTTTCACCCCACCCTTCC-3′ located 90 bp upstream of the poly(A) site] and oligo(dT)-anchor primer.

RNA Electrophoretic Mobility Shift Assay (REMSA) Supershift.

NE fractions (10 μg) were incubated with 4 μg of the indicated antibody for 1 h on ice before the addition of the radiolabeled RNA as described elsewhere (17).