The p53 R181C mutation accumulates through impaired deacetylation by Sirt1 and facilitates tumor development.

Introduction
The p53 protein, known as the “guardian of the genome”, is a transcription factor that regulates cell fate by activating the expression of various target genes. It plays a crucial role in maintaining cellular homeostasis and genomic stability by participating in processes such as apoptosis and cell cycle arrest1,2. Mutations in p53 are common in tumor tissues, with a frequency of over 50%3, the mutations can lead to changes in the structure of the p53 protein, which may ultimately affect its ability to interact with MDM2 in specific regions4. Consequently, mutated p53 is incapable of initiating MDM2-mediated degradation, resulting in a decreased capacity to degrade mutated p53 itself. This widely-held concept has been widely regarded as the main mechanism responsible for the accumulation of mutated p53 protein in tumors5. The MDM2 variant can also facilitate the accumulation of mutp53 and promote tumor development by inhibiting the degradation of mutp53 protein mediated by MDM26. Additionally, members of the heat shock protein family, such as Hsp90, can interact with mutp53, inhibiting its ubiquitination and degradation7. p53 mutations have been found to be significant in the development and progression of tumors8. Most of them are missense mutations that occur within the DNA-binding domain9. These mutations cause the p53 protein to lose its tumor-suppressive function or acquire new pro-tumorigenic activities, contributing to tumor development, proliferation, migration, and drug resistance processes10. Furthermore, TP53 mutations in individuals with Li-Fraumeni syndrome can enhance oxidative metabolism through mitochondria in a cell-autonomous manner, thereby imparting survival and proliferative advantages to cancer cells11.
One of the earliest identified hotspot mutations associated with hereditary breast cancer is the mutation at residue R181, which is also associated with various cancers such as cervical cancer, colorectal cancer, and skin cancer12. There is an association between the TP53 R181C mutation in humans and increased fat metabolism and decreased body fat content in p53R178C mice13. These effects are believed to impact the development of cancer through the regulation of fat metabolism. Previous studies have reported that the interaction between R181 and the opposite-charged residue E180 is crucial for the DNA-binding cooperativity of p53, and impaired DNA-binding cooperativity reduces the tumor-suppressive activity of p5314. Additionally, post-translational modifications of proteins, such as acetylation, phosphorylation, and ubiquitination15–17, also play important roles in the stability and tumor-suppressive function of p535,18. However, there have been no reports on the protein translation level of p53R181C, and its protein accumulation and carcinogenic mechanisms remain poorly understood. As the p53R181C mutation is prevalent in various tumors within the population, it is crucial to gain a better understanding of the protein accumulation and carcinogenic mechanism associated with this mutation.
Sirtuin1 (Sirt1) is a conserved NAD+-dependent protein deacetylase that plays critical roles in various cellular events, including gene regulation, maintenance of genome stability, cell apoptosis, autophagy, proliferation, aging, and tumorigenesis19. Sirt1 also participates in the epigenetic regulation of tissue homeostasis and many diseases through deacetylation of histones and non-histone proteins20. Notably, p53 was the first Sirt1 non-histone protein target to be discovered21. Sirt1 deacetylates the K382 site of p53 in an NAD + -dependent manner, leading to the inhibition of its transcriptional activity22. This modulation of p53 activity by Sirt1 affects various pathways involved in cellular homeostasis and pathological conditions23. Thus, the Sirt1-p53 axis is considered to play a central role in tumorigenesis. However, no study has yet investigated the protein modification effects of Sirt1 on p53R181C, and the precise role and mechanism of Sirt1 in tumor development caused by p53R181C mutations remain unknown.
In this study, we found that the p53R181C mutant shows reduced binding to the deacetylase Sirt1, which leads to increased acetylation levels of the K382 site of p53R181C and inhibits its ubiquitin-mediated degradation by MDM2. As a result, there is an accumulation of p53R181C protein. Furthermore, the p53R181C mutation demonstrated “loss-of-function” of transcriptional regulating downstream tumor suppressor genes including p21, bax, and PUMA as well as “gain-of-function” of transcriptional regulating tumor promoting genes of PIK3CA, SHC1, SRC, and PAK4. This mutation is advantageous for cancer cells with genomic instability as it allows them to evade anti-cancer mechanisms including cell cycle arrest, cellular senescence, and apoptosis. In addition, the mutation promotes tumor initiation and progression. Our study provides insights into the molecular mechanism underlying the accumulation of the p53R181C mutant protein and the ways in which this mutation impairs its tumor suppressor function as well as facilitates tumor progression.

Results

The p53R181C mutation accumulates at the protein level and leads to an increase in K382 acetylation
The p53R181C mutant protein results from a substitution of the codon CGC that encodes arginine at position 181 in the p53 DNA binding domain with the codon TGC that encodes cysteine (Fig. S1A). To better understand the relationship between the p53R181C mutation and the development of tumors, we referred to the p53 online analysis website The TP53
DATABASE24, which provided information on the types and frequencies of human tumors associated with the p53R181C mutation. According to the reported data, breast cancer is the most common type of tumor associated with the p53R181C mutation, followed by uterine cancer (Fig. 1A). We used a mouse model carrying the p53R178C mutation, which corresponds to the human p53R181C mutation, to investigate the accumulation of the mutant protein. First, we extracted protein from the heart, liver, spleen, and kidney tissues of p53WT and p53R178C mice. Western blot analysis confirmed that the levels of p53 protein in the different tissues of p53R178C mice were significantly higher than in wild-type mice (Fig. 1B).
Post-translational modifications, such as phosphorylation, acetylation, glycosylation, and ubiquitination, play crucial roles in the stability and accumulation of p53. We investigated the post-translational modifications of both p53R181C and p53WT proteins via exogenous expression of Flag-tagged proteins, followed by immunoprecipitation and immunoblotting to detect specific modifications. The results revealed no difference between Flag-p53R181C and Flag-p53WT in the levels of detection of S15 phosphorylation (Fig. 1C), S20 phosphorylation (Fig. 1D), and O-GlcNAc glycosylation (Fig. 1E). The level of O-GlcNAc glycosylation was also similar between WT and p53R178C mouse embryonic fibroblast (MEF) cells (Fig. S1B).
Previous studies have shown that the deacetylase Sirt1 can impact the protein level of p53 by deacetylating the K382 site of p5325. We therefore examined the acetylation level of the K382 site in p53WT and p53R181C. Indeed, the acetylation level of Flag-p53R181C was higher than that of Flag-p53WT (Fig. 1F), specifically at the K382 site (Fig. 1G). Endogenous results revealed that REH cells (p53 R181C mutant) showed significant increased level of p53 acetylation at K382 than that in NALM6 cells (p53 wild-type) (Fig. 1H). Similar results were observed in MEF cells, where the acetylation level of endogenous p53R178C was also elevated (Fig. S1C). Additionally, the acetylation level of K382 in p53R178C was observed to be higher than in WT in MEF cells (Fig. 1I). Notably, when the K382 site was acetylation-inactivated, there was no longer any observable difference in acetylation levels between p53WT and p53R181C (Fig. 1J), indicating that the higher acetylation level of p53R181C is dependent on the K382 site. In MEF cells, the acetylation level of K382 in the p53R178C protein increased with the number of passages, surpassing that of p53WT (Fig. 1K). The knockout of p53 in HEK293 cells has been validated (Fig. S1D). Additionally, under stress conditions such as ionizing radiation (IR), the acetylation level of K382 in p53R178C was significantly higher than that in p53WT (Fig. 1L), consistent with the results in HEK293 p53 knockout (HEK293 p53KO) cells (Fig. S1E). In addition, we examined the acetylation levels of other hotspot mutations of p53, specifically focusing on K382. Our findings revealed no significant difference in acetylation levels compared to the p53WT in HEK293 p53KO cells, except that R181C showed high K382 acetylation (Fig. 1M).
Together, these findings suggest that the accumulation of p53R181C protein is associated with increased acetylation, and its higher acetylation level is dependent on the K382 site.

The p53R181C mutation reduces binding to Sirt1, resulting in an increase in acetylation level
The acetylation modification is a dynamic and reversible post-translational modification that is primarily enhanced by modifying acetyltransferase activity. The level of substrate acetylation modification can be decreased by deacetyltransferases15. Studies have shown that the acetyltransferase P300 can increase the acetylation level of the p53 K382 site26, while the deacetylase Sirt1 can decrease the acetylation level of the p53 K382 site21.
We further investigated the effect of P300 and Sirt1 on the acetylation levels at the p53R181C K382 site. Co-IP analysis of the interaction between endogenous P300 and Flag-p53WT or Flag-p53R181C indicated no difference in their interaction in HEK293 p53KO cells (Fig. S2A, B), suggesting that P300 is unaffected in its acetylation modification activity toward p53R181C. However, the interaction between Flag-p53R181C and endogenous Sirt1 was decreased in HEK293 p53KO cells (Fig. 2A, B) and H1299 cells (Fig. S2C, D). Endogenous co-immunoprecipitation revealed that the interaction between p53 and Sirt1 is significantly decreased in REH cells (p53 R181C mutant) than that in NALM6 cells (p53 wild-type) (Fig. 2C). Similarly, the interaction between endogenous p53R178C and Sirt1 in MEF cells was also reduced (Fig. S2E, F), indicating a decrease in the ability of Sirt1 to deacetylate the mutant p53 protein compared to p53WT.
To evaluate the effect of the weakened interaction between p53R181C and Sirt1 on the acetylation levels, specifically at the K382 site, we conducted experiments using HEK293 cells. We first knocked out Sirt1 in normal HEK293 cells and observed no difference in K382 acetylation levels between Flag-p53WT and Flag-p53R181C (Fig. 2D). We then transfected HEK293 p53KO cells with Flag-p53WT and Flag-p53R181C, and overexpressed Sirt1-WT or Sirt1-H363Y (Fig. S2G). The results showed that Sirt1-WT can downregulate the K382 acetylation level of Flag-p53WT, but was unable to reduce the K382 acetylation level of Flag-p53R181C. This suggests that Sirt1 is not effective in deacetylating the K382 site of Flag-p53R181C. On the other hand, Sirt1-H363Y did not have any impact on the K382 acetylation level of Flag-p53WT, indicating that Sirt1 deacetylates p53 through its classical H363 site27.
We further tested the deacetylation activity of endogenous Sirt1 towards Flag-p53WT and Flag-p53R181C using the Sirt1 inhibitor EX527. EX527 is a specific small molecule inhibitor of Sirt1 catalytic activity that significantly increases the acetylation level of p53 K382. The acetylation levels of K382 in Flag-p53WT and Flag-p53R181C were both increased in HEK293 p53KO cells, indicating that Sirt1 can still deacetylate Flag-p53R181C (Fig. 2E). Additionally, when we used the Sirt1-specific synthetic activator resveratrol in HEK293 p53KO cells, the K382 acetylation level of Flag-p53WT decreased, but the K382 acetylation level of Flag-p53R181C remained largely unchanged (Fig. 2F). This indicates a decreased deacetylation function of Sirt1 towards Flag-p53R181C. Moreover, we found that the increase in the acetylation level of the K382 site of p53R181C induced by H2O2 stimulation was not as significant as that of the wild type in H1299 cells which are p53-null (Fig. 2G). Importantly, we found that when the enzymatic activity of Sirt1 was either activated or inhibited, the mRNA levels of endogenous p53WT and p53R178C were not affected (Fig. 2H, I).
Taken together, our results indicate that the decreased interaction between p53R181C and the deacetylase Sirt1 results in Sirt1’s inability to effectively deacetylate p53R181C, thereby leading to an increase in the acetylation level at the K382 site.

The high acetylation of K382 in p53R181C inhibits ubiquitination-dependent degradation by MDM2
Ubiquitin-mediated proteasomal degradation is the main pathway for protein degradation17. Acetylation of the C-terminus lysine residues of p53 affects its ubiquitination and degradation28. MDM2 serves as an E3 ubiquitin ligase for p53 and tightly regulates its ubiquitination and degradation, maintaining low expression levels under normal conditions4. We found that the binding between Flag-p53R181C and endogenous MDM2 was reduced in HEK293 p53KO cells, as demonstrated by co-IP analysis (Fig. 3A, B), and decreased binding between endogenous p53R178C and MDM2 in MEF cells was also observed (Fig. S3A, B). Additionally, endogenous assays in NALM6 and REH cells demonstrated that the p53-MDM2 interaction is significantly attenuated in REH cells carrying p53-R181C mutant compared with wild-type p53 NALM6 cells (Fig. 3C). Consistently, in HEK293 p53KO cells, the ubiquitination level of Flag-p53R181C mediated by MDM2 was lower (Fig. 3D). Endogenous analysis of NALM6 and REH cells further showed that p53 ubiquitination is significantly lower in REH cells than in NALM6 cells (Fig. 3E). The similar results were also obtained in the in vitro ubiquitination experiments by in vitro purification of His-MDM2, GST-p53WT and GST-p53R181C (Fig. 3F). In addition, in vitro assays showed that the effect of MDM2 on the neddylated level of GST-p53R181C is consistent with that of GST-p53WT, indicating that the R181C mutation does not change the neddylation modification of MDM2 on p53 (Fig. S3C).
To confirm that ubiquitin-mediated protein degradation accounts for the difference in p53 mutant and WT protein levels, we compared the half-lives using cycloheximide to inhibit new protein synthesis. We observed that when protein synthesis was blocked, the half-life of p53R181C was longer compared to p53WT, indicating a more stable protein (Fig. S3D). Moreover, when proteasomal degradation was inhibited, the protein levels of both p53R181C and p53WT remained stable (Fig. S3E), suggesting that p53R181C could still be degraded through the ubiquitin-proteasome pathway.
The above results support that the mutant p53R181C protein is inefficiently targeted by MDM2-mediated protein degradation. We found that upon overexpression of MDM2, p53R181C could still be degraded, albeit at a slower rate (Fig. S3F). To further investigate the impact of acetylation at the K382 site on the binding of p53R181C to MDM2, we generated plasmids with mutations that either eliminated (K382R) or mimicked acetylation (K382Q) at this site in Flag-p53R181C. Co-IP experiments revealed that the high acetylation mimic state of the K382Q site not only hindered the interaction between Flag-p53R181C and MDM2 (Fig. 3G, H), but also impaired the interaction between Flag-p53WT and MDM2 in HEK293 p53KO cells (Fig. S3G, H). Ubiquitination detection experiments confirmed that the high acetylation mimic state of the p53WT K382Q site directly inhibited its ubiquitination and degradation in HEK293 p53KO cells (Fig. S3I).
We further investigated the impact of acetylation status at the K382 site on the ubiquitination and degradation of p53R181C. Increasing amounts of deacetylase myc-Sirt1 was co-transfected with Flag-p53WT or Flag-p53R181C in HEK293 p53KO cells. A gradual decrease in protein levels of p53 WT was apparent, which correlated with increased myc-Sirt1, consistent with deacetylation by Sirt1. However, p53R181C, which cannot be efficiently deacetylated by Sirt1, showed stable protein levels (Fig. S3J).
To further validate the attenuated deacetylase function of Sirt1 on p53R181C, which inhibits its ubiquitination and degradation, we knocked out Sirt1 in HEK293 cells and looked for an effect on ubiquitination levels. We observed a decrease in ubiquitination levels of both Flag-p53WT and Flag-p53R181C (Fig. 3I). When HEK293 p53KO cells were treated with the Sirt1 activator resveratrol, an increase in the ubiquitination levels of Flag-p53WT and Flag-p53R181C was observed, which was more obvious in Flag-p53WT (Fig. 3J). By contrast, Sirt1 inhibition with EX527 resulted in a decrease in ubiquitination levels of both proteins, which was more obvious in Flag-p53WT (Fig. 3K).
In summary, the above results indicate that the reduced interaction between p53R181C and the deacetylase Sirt1, along with the increased acetylation levels at K382, inhibits the binding of p53R181C to the E3 ubiquitin ligase MDM2. As a result, the ubiquitination and degradation of p53R181C are suppressed, leading to the stabilization and accumulation of p53R181C protein.

The p53R181C mutation gains transcriptional regulation of downstream tumor promoting genes
Mutant p53 often demonstrates novel function compared with wild-type p53, known as “gain of function”29–31. To further understand the mechanism underlying the gene expression changes of p53R178C, we performed chromatin immunoprecipitation followed by sequencing (ChIP-seq) to create a genomic profile of wild-type and mutant p53 binding. ChIP-seq was performed in p53-knockout HCT116 cells transfected with p53 wild-type or p53R181C mutation. The raw data of ChIP-seq were shared in public repository of National Genomics Data Center (https://ngdc.cncb.ac.cn/omix/view/OMIX011382). The KEGG pathway analysis and GO enrichment analysis were conducted to show the dysregulated signaling pathways (Fig. 4A-C). As a result, multiple tumor-related pathways were enriched, including renal cell carcinoma, non−small cell lung cancer, melanoma, gastric cancer, and so on. We therefore confirmed the tumor-related genes from these pathways of ChIP-seq, and found that p53R181C showed “gain-of-function” of transcriptional regulating tumor promoting genes of PIK3CA, SHC1, SRC and PAK4 according to the results of quantitative real-time PCR (Fig. 4D–G). In addition, we performed ChIP assay to confirm the results of ChIP-seq and found that p53R181C significantly bind with DNA promoter of tumor promoting genes of PIK3CA, SHC1, SRC, and PAK4, while wild-type p53 could not (Fig. 4H–K). ChIP-seq profile and the motifs of different peaks PIK3CA, SHC1, SRC, and PAK4 gene were shown (Fig. 4L, M). PIK3CA, SHC1, SRC, and PAK4 have been reported to contribute to the progression of different types of cancers32–35. Therefore, the p53R181C mutation showed “gain of function” of transcriptional regulation of downstream tumor promoting genes including PIK3CA, SHC1, SRC, and PAK4.

The p53R181C mutation impairs its transcriptional regulation of downstream tumor suppressor genes, leading to diminished anti-cancer function
The tumor suppressor protein p53 is one of the most effective natural cellular defenses against cancer. Upregulation of p21 results in cell cycle arrest or senescence, while upregulation of Bax and PUMA promotes apoptosis to prevent tumor development9.
Real-time fluorescence quantitative PCR analysis revealed a significant decrease in the levels of downstream anti-cancer target genes, including p21, bax, and PUMA, in p53R178C MEF cells (Fig. 5A). Western blot analysis showed a decrease in p21 protein expression in various tissues of p53R178C mice (Fig. 5B). We also extracted embryonic fibroblasts from p53WT and p53R178C mice and found that during cell passaging, the levels of p53R178C protein were significantly higher than those of p53WT (Fig. 5C). A significant decrease in p21 protein expression in fibroblasts of different passage numbers was found in p53R178C mice (Fig. 5D).
We also found that MEF cells expressing p53R178C showed higher levels of p53 compared to WT MEF cells after exposure to IR stimulation. Additionally, real-time fluorescence quantitative PCR analysis of mRNA extracted from embryonic fibroblasts of p53WT and p53R178C mice showed no difference in mRNA levels between them (Fig. 5E). These results indicate that the p53R181C mutation causes p53 accumulation at the protein level. We next investigated the consequences of the effect of p53R178C on cell fate. Compared to p53WT, the cell cycle arrest function was impaired in p53R178C MEF cells (Fig. 5F), and cellular senescence was delayed (Fig. 5G). The expression of PUMA and Bax proteins was decreased in p53R178C MEF cells, and under cellular stress conditions, their expression was not upregulated (Fig. 5H). Moreover, apoptosis function was reduced, even under IR stress conditions, resulting in significantly lower levels of apoptosis in p53R178C MEF cells compared to p53WT cells (Fig. 5I). Consistently, under stress conditions, Flag-p53R181C was also unable to induce apoptosis in p53KO HCT116 cells (Fig. 5J). The gating strategy of flow cytometry analysis is shown (Fig. S4A–C).
To examine the phosphorylation activation state of p53R181C under cellular stress conditions, we transfected Flag-p53WT and Flag-p53R181C into p53KO HEK293 cells and exposed them to IR. The upstream signaling pathway ATM-CHK2 of p53 was intact in response to DNA damage. Compared to Flag-p53WT, the phosphorylation of S15 in Flag-p53R181C was normal (Fig. S4D), but the phosphorylation of S20 was reduced (Fig. S4E). Consistent with the results from exogenous expression in HEK293 p53KO cells, the phosphorylation of S20 was reduced in p53R178C MEF cells (Fig. S4F), which corresponds to the weakened cell cycle arrest function mediated by p53R178C.
The above results indicate that the p53R181C mutation inhibits its anti-cancer function, which could thereby promote tumor occurrence and development.

The p53R181C mutation enhances the growth of tumor cells and tumors in nude mice
We next probed the role of p53R181C in tumor development. Proliferating cell nuclear antigen (PCNA) plays a role in facilitating DNA replication36 and can therefore serve as an indicator of cell proliferation37. Studies have shown that p21 can interact with PCNA, inhibiting its ability to activate DNA polymerase and thus impeding PCNA-dependent DNA replication38. Moreover, p21 can directly decrease the expression level of PCNA protein39. Consistently, western blot analysis revealed an increase in PCNA protein expression in p53R178C MEF cells of different generations (Fig. 6A). This increase corresponds to accelerated cell proliferation and is also linked to the downregulation of p21 protein expression in p53178C MEF cells.
We then utilized the RTCA technique to detect the proliferation of MEF cells expressing p53WT and p53R178C in order to explore the effects on tumor development. The results showed that p53R178C MEF cells proliferated faster (Fig. 6B), which is consistent with the findings from the CCK8 assay (Fig. 6C). Transfection of Flag-p53WT and Flag-p53R181C into p53KO HCT116 cells indicated that p53R181C promotes the proliferation of HCT116 cells (Fig. 6D).
We established a mouse mammary carcinoma cell line expressing p53R178C (Fig. S4G). Compared to normal 4T1 cells, p53R178C-4T1 proliferated significantly faster (Fig. 6E). To further explore the impact of p53R178C on tumor growth in vivo, we subcutaneously injected con-4T1 and p53R178C-4T1 cells into immunodeficient BALB/c nude mice and recorded the tumor growth rate. After 21 days, the p53R178C group showed a significantly faster tumor growth rate and higher tumor weight compared to the Con group (Fig. 6F). Immunohistochemistry (IHC) analysis of tumor samples revealed higher expression of the proliferation marker Ki67 in the p53R178C group compared to the Con group (Fig. 6G).
Taken together, these results indicate that p53R181C not only promotes tumor cell proliferation but also accelerates tumor growth in vivo.

Discussion
The human cellular genome is constantly subjected to various stressors that can lead to DNA damage, the fundamental cause of tumor development40,41. The transcription factor p53 plays a crucial role in the DNA damage checkpoint response, serving as a central regulatory factor. Germline p53 mutations are responsible for Li-Fraumeni syndrome, a condition that increases the risk of developing various early-onset cancers42. Here, we found that the p53R181C mutant shows reduced binding to the deacetylase Sirt1, which leads to increased K382 acetylation level and inhibits its ubiquitin-mediated degradation by MDM2. As a result, there is an accumulation of p53R181C protein. Furthermore, the p53R181C mutation demonstrated “loss-of-function” of transcriptional regulating p21, bax, and PUMA as well as “gain-of-function” of transcriptional regulating tumor promoting genes of PIK3CA, SHC1, SRC, and PAK4. This mutation is advantageous for cancer cells with genomic instability as it allows them to evade anti-cancer mechanisms including cell cycle arrest, cellular senescence, and apoptosis. In addition, the mutation promotes tumor initiation and progression.
The accumulation of mutant p53 (mutp53) is crucial for its role in tumor development43. Currently, the mechanisms underlying mutp53 accumulation remain unclear. It is generally believed that MDM2 cannot efficiently ubiquitinate and degrade mutp53, resulting in the accumulation of mutp53 protein. Missense p53 mutant variants often accumulate in tumors and drive progression through gain-of-function. Therefore, the accumulation of mutant p53 protein is an important target for anti-cancer drug development. Some tumor-specific mechanisms have been reported, such as HSP90 binding to mutp53 and inhibiting MDM2-mediated degradation, which leads to mutp53 accumulation7. Targeting HSP90 to reactivate MDM2-mediated degradation of mutp53 disrupts the stability of mutp53 and inhibits its gain-of-function in tumor development44. Additionally, various cancer-related stress stimuli, including DNA damage, oxidative and protein toxic stress, and metabolic stress, have been reported to promote mutp53 protein accumulation and gain-of-function in tumors through different mechanisms45,46.
In this study, we discovered that the accumulation of the p53 R181C mutant is not only influenced by ubiquitination modification but also regulated by acetylation. Previous literature has reported that acetylation of multiple lysine residues at the C-terminus of p53 affects its degradation through ubiquitination by the E3 ligase MDM247. Our experimental findings demonstrate that the acetylation status of a single lysine residue at the p53 K382 site impacts MDM2-mediated ubiquitination degradation of p53. This leads to the inability of p53 R181C to undergo proper degradation and results in protein accumulation.
This mechanistic study provides insights into the molecular regulatory mechanisms of post-translational modifications on the abnormal expression of p53 mutant variants. It also offers a new perspective for the development of anti-cancer drugs targeting mutant p53. Mutations in p53 are typically located in DNA-binding regions, and the mechanisms by which mutations accumulate at different sites vary. As a result, their acquired functional changes differ48. DNA contact mutations, such as R273H, lead to the substitution of critical amino acid residues that directly bind to p53 target genes. This substitution inhibits transcriptional activity but does not significantly affect protein conformation. On the other hand, conformational mutations, like R175H, generally cause more drastic changes in the folding and structure of the p53 protein. Point mutations in the DNA binding domain of p53 weaken its binding with the protein49, or specifically promote the stability of mutant p53 protein by binding with certain proteins50. The p53 Arg181 is also located in the DNA binding domain. Biochemical and structural studies have shown that when the positively charged arginine residue at position 181 is mutated to the uncharged cysteine residue, it affects the spatial conformation of the p53 protein due to the change in charge. This, in turn, influences its binding with DNA or proteins14. However, the specific structure of the binding between the p53 R181C mutation and Sirt1 is still unknown.
We found in this study that p53 R181C cannot activate p21 normally, leading to defective cell cycle arrest mediated by p53 R181C. This causes slower aging and accelerated proliferation, allowing cells with genomically unstable features to evade aging and continue to proliferate. Additionally, the H1 helix interaction of p53 is essential for the conformational activation of the pro-apoptotic gene Bax51. This study also found that the mutant p53 R181C loses its ability to induce cell apoptosis. Consequently, cells escape both cell cycle regulation and aging, as well as apoptosis, leading to the infinite proliferation of genomically unstable cells and ultimately resulting in tumor formation. In addition, the p53R181C mutation demonstrated “gain of function” of transcriptional regulation of downstream tumor promoting genes including PIK3CA, SHC1, SRC and PAK4, which contributes to the development of cancer. ChIP-seq analysis revealed that p53R181C binds to the promoters of genes implicated in cancer progression including PIK3CA, SHC1, SRC, and PAK4, despite the absence of the canonical p53 consensus binding motif. This suggests that p53R181C may interact with non-canonical DNA sequences or rely on indirect mechanisms such as chromatin remodeling or interactions with other co-factors to exert its oncogenic transcriptional activity, as was reported by previous study52. The potential clinical translation of these findings is multifaceted. Diagnostically, the p53R181C mutation and its associated gene expression signature could serve as biomarkers for early cancer detection and risk stratification. Therapeutically, the identification of Sirt1 and MDM2 as key regulators of p53R181C stability provides a rationale for developing targeted therapies. For instance, combining Sirt1 activators with MDM2 inhibitors may synergistically reduce p53R181C levels. Furthermore, these findings underscore the importance of personalized medicine approaches, where therapeutic decisions are based on the specific p53 mutation status of the tumor.
In summary, our research uncovers the molecular mechanisms responsible for the accumulation of p53 R181C and the impairment of its tumor suppressor function. Additionally, we identify key mechanisms through which p53 R181C facilitates tumor growth. Our study presents new possibilities for the development of effective therapeutic approaches aimed at treating cancers with p53 R181C, and establishes the theoretical basis for the development of precise cancer treatments targeting p53 R181C.

Methods

Chemicals and reagents
The following reagents and antibodies were purchased from Invitrogen: Dulbecco’s Modified Eagle’s Medium (DMEM), Roswell Park Memorial Institute (RPMI) 1640 medium, penicillin/streptomycin, fetal bovine serum (FBS), and trypsin. The antibodies used in this study include: anti-p53 (#sc-126, Santa Cruz, 1:1000), anti-p21 (#sc-6246, Santa Cruz, 1:1000), anti-Nedd8 (#sc-373741, Santa Cruz, 1:1000), anti-GFP (#A02375-50, Genscript, 1:2000), anti-Phospho-p53 (Ser20) (#9287S, Cell Signaling Technology, 1:1000), anti-Phospho-p53 (Ser15) (#9284S, Cell Signaling Technology, 1:1000), anti-Acetyl-p53 (Lys379) (#2570S, Cell Signaling Technology, 1:1000), anti-MDM2(#51541S, Cell Signaling Technology, 1:1000), anti-ATM(#2873S, Cell Signaling Technology, 1:1000), anti-Phospho-ATM (Ser1981) (#13050S, Cell Signaling Technology, 1:1000), anti-CHK2(#3440S, Cell Signaling Technology, 1:1000), anti-Phospho-CHK2 (Thr68) (#2661S, Cell Signaling Technology, 1:1000), anti-BCL2(#15071S, Cell Signaling Technology, 1:1000), anti-Phospho-Histone H2A.X (Ser139) (#9718S, Cell Signaling Technology, 1:1000), anti-acetylated-lysine (#9441S, Cell Signaling Technology, 1:1000), anti-HA (#3724, Cell Signaling Technology, 1:1000), anti-β-actin (AC004, ABclonal, 1:2000), anti-α-Tubulin (AC012, ABclonal, 1:2000), anti-Flag (#SG4110-16, Shanghai Genomics Technology, 1:2000), anti-myc (SG4110-18, Shanghai Genomics Technology, 1:2000), anti-GAPDH (A19056, ABclonal, 1:2000), anti-Bax (A19684, ABclonal, 1:1000), anti-PUMA (A3752, ABclonal, 1:1000), anti-p53(10442-1-AP, Proteintech, 1:1000), anti-Sirt1(#07-131, Millipore,1:1000). The antibodies were used according to the recommended dilution ratios. Anti-MDM2 (#51541S) to detect MDM2 isoform A1 from Cell Signaling Technology. Amphotericin B was purchased from Sigma. IR was generated using the Rad Source R1800Q irradiator (Rad Source Technologies, Buford, USA).

Plasmid construction and viral infection
The FLAG-p53WT, MYC-Sirt1WT, MYC-Sirt1H363Y, and HA-Ub plasmids available in the laboratory were confirmed by sequencing. All plasmids were confirmed by sequencing (GenScript, Nanjing, China). The His-MDM2, GST-p53WT, and GST-p53R181C plasmids were purchased from Miaoling Bio (Wuhan, China). The FLAG-p53R181C and various point mutants were constructed using the QuickChange site-directed mutagenesis kit. Lentivirus infection was performed using control and p53R178C-expressing lentiviruses purchased from GeneChem (Shanghai, China). Stable overexpressing cell lines were selected using puromycin and confirmed by Western blot analysis.

Construction of TP53-knockout HEK293 cells
TP53-knockout HEK293 cells (HEK293 TP53-KO) were generated in-house using CRISPR/Cas9-mediated genome editing as previously described. The CRISPR Cas9-p53 plasmid was created by attaching the designed sgRNA to the CRISPR Cas9 vector. The sgRNA sequence used was:
Oligo-p53-Primer F: CACCCCATGAGCGCTGCTCAGATAGCG;
Oligo-p53-Primer R: AAACCGCTATCTGAGCAGCGCTCATGG.
Absence of p53 protein expression was confirmed by Western blot with anti-p53 DO-1 antibody (Santa Cruz, #sc-126). Cells were maintained in DMEM high-glucose supplemented with 10% fetal bovine serum and 1% penicillin-streptomycin and routinely tested negative for mycoplasma contamination.

Cell culture and transfection
HEK293 cells were cultured in DMEM supplemented with 10% fetal bovine serum (FBS). HCT116, H1299, and 4T-1 cells were cultured in RPMI 1640 medium supplemented with 10% FBS. All cells were obtained from the Shanghai Institute of Cell Biology, Chinese Academy of Sciences. MEF cells were derived from 13.5-day-old mouse embryos and cultured in DMEM supplemented with 15% FBS. For cell transfection, cells were seeded at approximately 80% confluence and transfected with BIOBEST (polyplus-transfection, USA) following the manufacturer’s instructions. After transfection, cells were cultured for 24–48 h before analysis.

Western blot analysis
Cells were lysed on ice for 30 min in IP lysis buffer containing 50 mM Tris-HCl, 1467 mM EDTA, 1% Triton X-100, 150 mM NaCl, and 1.25% C24H39NaO4. The lysates were centrifuged at 14,000 rpm, 4 °C for 20 min, and the supernatants were collected. Protein concentration was determined using Coomassie Brilliant Blue G-250 staining. 30–50 μg of protein was separated by 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and transferred to a polyvinylidene fluoride membrane (Millipore). The membrane was blocked with 5% bovine serum albumin (BSA) in Tween 20 Tris-buffered saline (TBST) for 1 h at room temperature. Then, the membrane was incubated with the primary antibody overnight at 4 °C. Afterward, the membrane was washed three times with TBST for 10 min each and incubated with a secondary antibody for 1 h at room temperature. Finally, the membrane was developed and saved.

Co-IP and IP analyses
For Co-IP, the lysates were incubated with the primary antibody at 4 °C for 3 h. Then, A/G-sepharose beads (Santa Cruz) were added and the mixture was incubated overnight at 4 °C. The beads were then centrifuged at 700 × g, 4 °C for 5 min, washed three times with pre-chilled phosphate-buffered saline (PBS), and the supernatant was discarded. Next, the beads were boiled in 2x sample buffer for 10 min, and the resulting supernatant was subjected to SDS-PAGE.

Cycloheximide (CHX) chase assay
Cells were seeded in 6-well plates and transfected with the indicated plasmids (Flag-p53 WT or Flag-p53 R181C) using Lipofectamine 3000 (Invitrogen) according to the manufacturer’s instructions. Twenty-four hours post-transfection, the medium was replaced with fresh complete medium containing 20 µM cycloheximide (CHX, Selleck, S7418) to block de novo protein synthesis. Cells were harvested at 0, 0.5, 1, 2 and 3 h after CHX addition. At each time point, cells were washed once with ice-cold PBS, lysed in IP lysis buffer (50 mM Tris-HCl, 1467 mM EDTA, 1% Triton X-100, 150 mM NaCl, 1.25% C24H39NaO4) supplemented with protease inhibitors (Protease Inhibitor Cocktail, HY-K0010), and clarified by centrifugation at 13,000 × g for 20 min at 4 °C. Equal amounts of protein (20 µg) were resolved by SDS-PAGE, transferred to PVDF membranes, and immunoblotted with anti-Flag (Shanghai Genomics Technology, #SG4110-16) and anti-β-actin (Abclonal, AC004) antibodies. Band intensities were quantified using ImageJ and normalized to β-actin. Protein half-lives were calculated by fitting the data to a one-phase exponential decay model (GraphPad Prism).

In vitro ubiquitination assay
To determine the direct effects of MDM2 on the ubiquitination of p53WT and p53R181C, His-MDM2, GST-p53WT and GST-p53R181C plasmids were expressed in Escherichia coli BL21 and purified with anti-His beads or anti-GST beads in vitro, respectively. These purified proteins were added to the reaction mixture containing Mg-ATP, Ubiquitin, Ubiquitin-Activating Enzyme(E1), UbcH5a (E2), and Ub E3 Ligase Buffer which all provided by the E3 Ligase Auto-Ubiquitination Assay Kit (Abcam, ab139469). After incubation at 37°C for 1 h, the reaction was then terminated with 2×SDS loading buffer and subjected to western blot with anti-ubiquitin antibody.

In vitro neddylation assay
His-MDM2, GST-p53WT, and GST-p53R181C plasmids were expressed in Escherichia coli BL21 and purified with anti-His beads or anti-GST beads in vitro, respectively. These purified proteins were added to the reaction mixture containing Mg-ATP, Nedd8, E1 (APPBP1-UBA3), E2 (Ubc12). Samples were incubated at 30 °C for 1 h, and then terminated with 2×SDS loading buffer and subjected to western blot with anti-Nedd8 antibody.

Sequencing and computational analysis
ChIP-seq was performed in duplicate biological replicates on p53-knockout HCT116 cells transiently transfected with either Flagp53-WT or Flagp53-R181C plasmids using Lipofectamine 3000; 24 h post-transfection, cells were cross-linked with 1% formaldehyde for 10 min, quenched with 125 mM glycine, lysed in 1% SDS buffer, chromatin was sheared to 200–500 bp fragments with a Bioruptor Pico (30 s on /30 s off, 15 cycles), and 100 µg of chromatin was immunoprecipitated overnight at 4 °C with 5 µg anti-Flag antibody or IgG control followed by capture on protein G magnetic beads, stringent washing, and elution at 65 °C; after reverse cross-linking and proteinase K treatment, DNA was purified with the Zymo ChIP DNA Clean & Concentrator kit, libraries were prepared from 50 ng DNA using the MGIEasy DNA Library Prep Kit. ChIP-seq was performed by Wuhan Kangce Technology Co., Ltd (Wuhan, China). All ChIP-Seq samples were sequenced on the MGISEQ-T7. After obtaining the raw sequencing data, the raw data was first filtered using fastp (version 0.23.1, Shifu Chen 2018) software to obtain high-quality sequencing data, the sequencing data was compared with the reference genome of the project species, the comparison results were called by Macs2 for genome-wide peak calls, the binding preference of proteins on the genome was studied, and the binding site motif was analyzed.

Quantitative real-time PCR
Total RNA was isolated using Trizol reagent (Invitrogen, USA), and 1 μg of RNA was subjected to cDNA synthesis using HiScript 4 RT SuperMix for qPCR (Vazyme, R423-01). Quantitative real-time PCR (qPCR) was performed using 2* ChamQ Universal SYBR qPCR Master Mix (Vazyme, Q711-03) following the manufacturer’s instructions. The expression levels of the indicated mRNAs were calculated using the 2˗ΔΔCt method and were normalized to the internal control actin.

ChIP experiments
ChIP experiments were performed according to the manufacturer’s instructions (Cell Signaling Technology, 9003S). Briefly, cells were cross-linked with 1% formaldehyde for 10 min at 37 C, then treated with glycine for 5 min. Lysis buffer was added and the cells were incubated for 20 min on ice, then sonication was performed to cleave the chromatin into fragments. Next, primary antibodies and protein magnetic beads were added and incubated overnight at 4 °C to pull down the target protein-DNA fragment complexes. Then, the bead-antibody-DNA fragment complexes were collected and reverse cross-linked. After DNA purification, the immunoselected DNA fragments were used in real time-PCR analysis to detect the target gene.

Flow cytometry
Cells were fixed in 70% ethanol at 4 °C for 1 h, centrifuged at 1200 rpm, 4°C for 5 min, and washed once with PBS. Subsequently, the cells were stained with either 10 μg/ml propidium iodide (PI) staining solution or a combination of 5 μg/ml PI and 5 μg/ml Annexin FITC. The staining was performed at room temperature in the dark, with a duration of 30 min for the PI staining and 15 min for the Annexin FITC staining. Following staining, data acquisition was performed using a BD C6 flow cytometer and analysis was conducted using flow Jo software.

In vivo tumor implantation
Control or p53R178C-expressing 4T-1 cells were harvested after trypsinization and washed once with PBS. Then, 5.0 × 10^5 cells were resuspended in 100 μl of PBS and injected subcutaneously into immune-deficient nude mice (8 weeks old, n = 4) in the axillary area. The survival status of the mice was monitored, and the tumor size was measured and recorded every two days using calipers. After 21 days, the tumors were excised, weighed, and the tumor volume was calculated using the formula (length × width^2/π). All animal experiments were approved by the Institutional Animal Care and Use Committee of China Medical University.

Immunohistochemistry
Tumor tissues were fixed using 4% paraformaldehyde, dehydrated in a 30% sucrose solution, embedded, and frozen sectioned. The sections were then washed twice with PBS and permeabilized with 0.3% Triton-X100 (in PBS) at room temperature for 20 min. To block non-specific staining, goat serum was applied and left at room temperature for 1 h. The sections were then incubated overnight at 4 °C with primary antibodies. The following day, the sections were washed three times with PBS, incubated with fluorescent secondary antibodies at room temperature, and stained with DAPI to visualize the cell nuclei. Finally, the sections were dried.

Quantification and statistical analysis
SPSS 20.0 (SPSS Inc., Chicago IL, USA) was used for statistical analysis. Statistical comparisons between two groups were carried out using the unpaired Student’s t test or the Mann–Whiney U-test. Data are presented as mean ± standard error of the mean. P value < 0.05 was considered as statistically significant.

Supplementary information