BMB Reports 2024; 57(6): 299-304
Tumor antigen PRAME is a potential therapeutic target of p53 activation in melanoma cells
Yong-Kyu Lee1,#, Hyeon Ho Heo1,#, Nackhyoung Kim1, Ui-Hyun Park1, Hyesook Youn1, Eun-Yi Moon1, Eun-Joo Kim2 & Soo-Jong Um1,*
1Department of Integrative Bioscience and Biotechnology, Sejong University, Seoul 05006, 2Department of Molecular Biology, Dankook University, Cheonan 31116, Korea
Correspondence to: Tel: +82-2-3408-3641; Fax: +82-2-3408-4334; E-mail:
#These authors contributed equally to this work.
Received: December 26, 2023; Revised: March 12, 2024; Accepted: April 23, 2024; Published online: April 25, 2024.
© Korean Society for Biochemistry and Molecular Biology. All rights reserved.

cc This is an open-access article distributed under the terms of the Creative Commons Attribution Non-Commercial License ( which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.
Upregulation of PRAME (preferentially expressed antigen of melanoma) has been implicated in the progression of a variety of cancers, including melanoma. The tumor suppressor p53 is a transcriptional regulator that mediates cell cycle arrest and apoptosis in response to stress signals. Here, we report that PRAME is a novel repressive target of p53. This was supported by analysis of melanoma cell lines carrying wild-type p53 and human melanoma databases. mRNA expression of PRAME was down-regulated by p53 overexpression and activation using DNA-damaging agents, but upregulated by p53 depletion. We identified a p53-responsive element (p53RE) in the promoter region of PRAME. Luciferase and ChIP assays showed that p53 represses the transcriptional activity of the PRAME promoter and is recruited to the p53RE together with HDAC1 upon etoposide treatment. The functional significance of p53 activation-mediated PRAME downregulation was demonstrated by measuring colony formation and p27 expression in melanoma cells. These data suggest that p53 activation, which leads to PRAME downregulation, could be a therapeutic strategy in melanoma cells.
Keywords: Melanoma, p53 activation, PRAME, Repression, Transcription

Preferentially expressed antigen in melanoma (PRAME) was discovered as a melanoma-associated antigen recognized by cytotoxic T lymphocytes (CTL) (1). Like other cancer-testis antigens (CTAs), PRAME is not detected in normal tissues except germline tissues but is highly expressed in tumors of several histological types (2, 3). Elevated PRAME expression is associated with poor outcomes in neuroblastoma and breast cancer (4, 5). Recent immunohistochemical analysis revealed an increased expression of PRAME in primary and metastatic melanoma compared to normal nevi (6-8). The tumor promoting activity of PRAME was discovered via its depletion in xenograft mouse tumor models (9-11). Therefore, targeting PRAME, either by exploiting its immunogenic or oncogenic potential, has been proposed as a strategy for tumor therapy (12, 13). PRAME expression is upregulated by promoter DNA hypomethylation (14, 15) and is stimulated by cooperation with myeloid zinc finger 1 (16). However, little is known about how PRAME expression is downregulated in response to anti-tumor signaling.

Tumor suppressor protein p53, encoded by TP53, has been a focus of cancer research. Many stress signals, in particular DNA damage, promote p53 activation and stabilization via posttranslational modifications (PTMs), such as phosphorylation and acetylation (17). Acetylation of p53 is mediated by acetyltransferases including CBP/p300, and is counteracted by histone deacetylases (HDACs) and SIRT1 (18). As a nuclear transcription factor, p53 binds to the p53 response element (p53RE) comprising two 5’-RRRCWWGYYY-3’ decamers separated by a spacer of 0-13 base pairs (R = A/G, W = A/T, Y = C/T), and regulates the expression of a variety of target genes involved in cell cycle arrest, apoptosis, DNA repair, and senescence (19). Compared to the transactivation, p53-dependent transcriptional repression has proposed mechanisms: indirectly by p21 induction followed by activation of the DREAM repressive complex, or by binding directly to the target gene promoter (20, 21).

The opposite functions of tumor antigen PRAME and tumor suppressor p53 prompted us to investigate the role of p53 in the expression of PRAME. The PRAME level is downregulated in inverse proportion to p53 expression in melanoma cell lines. Here, we report the mechanism underlying p53-mediated PRAME repression in response to DNA damage. Because TP53 is rarely mutated in melanoma compared to a rate of ∼50% in other tumors, p53-activating agents may have therapeutic potential for melanoma via transcriptional repression of PRAME.


Inverse correlation between PRAME and p53 expression

To investigate the relationship between PRAME and p53, we evaluated protein levels in 15 cancer cell lines (Supplementary Table 1). In five melanoma cell lines, the PRAME protein levels are proportional to the metastatic and proliferative potentials of melanoma cells (16), and negatively correlated with p53 expression (Fig. 1A, top). Similar increases in PRAME mRNA levels were detected by RT-PCR (Fig. 1A, bottom). There was an inverse correlation between PRAME and p53 (R2 = 0.9351) (Fig. 1B). A similar correlation was observed in the other eight cell lines, albeit with smaller R2 values (Supplementary Fig. 1). Using the GSE4570 human dataset, we observed that the PRAME mRNA level is significantly increased in melanoma compared to normal samples (Fig. 1C), whereas p53 expression was increased in normal melanocyte compared with melanoma samples (Fig. 1D). Further assessment of the Pearson correlation coefficient in the GSE4570 dataset yielded an adjusted R2 value of 0.4626, suggesting a significant inverse correlation of p53 and PRAME expression in human melanoma (Fig. 1E).

p53 activation causes PRAME downregulation

To determine whether PRAME expression is regulated by p53, we first overexpressed p53 in HCT116 (p53−/−) and p53-null H1299 cells, and then measured the expression level of PRAME. As shown in Fig. 2A, PRAME protein expression in both cell lines was downregulated by p53 overexpression. In contrast, PRAME protein and mRNA levels was upregulated by depleting p53, accompanied with downregulation of p27, a negative target of PRAME (Fig. 2B) (16). To determine the effect of p53 activation on PRAME expression, A375P melanoma cells were treated with etoposide (ETO), a p53-inducing DNA-damaging agent. As shown in Fig. 2C, the protein and mRNA levels of PRAME decreased with increasing duration of ETO treatment. Likewise, ultraviolet C (UV-C) and methyl methane sulfonate (MMS) reduced the PRAME level in A375P cells (Fig. 2D). PRAME was also downregulated in the A375SM and TXM-13 melanoma cell lines upon ETO-induced p53 activation, as determined by visualizing acetylated p53 (Supplementary Fig. 2A). Also, similar assays were performed using p53wt and p53−/− HCT116 cells under two genotoxic conditions. Compared to p53wt HCT116 cells, no significant suppression of PRAME expression was observed in p53-null HCT116 cells, suggesting that PRAME repression by p53 activation is p53-dependent (Supplementary Fig. 2B). The response to MMS was less p53-dependent (data not shown). Together, these data suggest that PRAME expression is negatively regulated by p53 activation in melanoma cells.

p53 binds to the PRAME promoter and recruits HDAC1

To investigate the mechanism by which p53 represses PRAME expression, we isolated a regulatory sequence of the human PRAME gene and subcloned it into the pGL2 basic luciferase reporter vector. In silico analysis revealed two putative p53 response elements (upstream RE1, −1835/−1808, and downstream RE2, +1511/+1543) at −3556 to +2444 of PRAME promoter (Supplementary Fig. 3A). HCT116p53wt and HCT116p53−/− cells were transfected with a luciferase reporter containing the −3556/+2444 region of the PRAME gene and the Flag-p53 vector. Luciferase assays indicated that the transcriptional activity of PRAME was significantly downregulated by p53 overexpression in both cell lines (Supplementary Fig. 3B). To identify the RE responsible for p53-dependent transcriptional repression, various truncation mutants were constructed and transfected with the Flag-p53 expression vector (Fig. 3A). Constructs containing −2667/+2444 and −1864/+2444 were transcriptionally repressed by p53, whereas other deletions lacking upstream RE1 did not respond to ectopically expressed p53 (Fig. 3B). Four additional mutants were generated in the promoter region of PRAME: mutRE, base substitution; ΔRE1 (RE1 deletion), −1806/+2444; −2667/+1, RE1-positive; −1778/+1, RE1-negative (Fig. 3C and Supplementary 3C). Compared to wild-type, mutRE and ΔRE1 were not repressed by p53 (Fig. 3D). No p53-mediated repression was observed upon deletion of RE1 (−1778/+1) (Supplementary 3C). Overall, these data suggest that the p53RE1 upstream of PRAME is responsible for p53-mediated repression.

To determine whether p53 binds to RE1 or RE2 of PRAME, we performed chromatin immunoprecipitation (ChIP) using primer sets covering RE1 and RE2 of PRAME, and p53RE of p21 as the positive control. Like the p53RE of p21, the p53RE1 of PRAME was occupied by Flag-tagged p53 (Supplementary 3D), which was verified by quantitative PCR following ChIP (Fig. 3E). Further, ETO treatment increased the occupancy of endogenous p53 in the RE1 of PRAME in A375P cells (Supplementary 3E). Subsequent quantification revealed that the enrichment of p53 and HDAC1 at p53RE1 is significantly enhanced in response to ETO treatment (Fig. 3F). As expected, the level of acetylated histone H3 at lysine 9 (H3K9ac), a substrate of HDAC1, was downregulated (Fig. 3F). These findings suggest that p53 represses the transcriptional expression of PRAME by binding p53RE1 in the PRAME promoter and recruiting HDAC1, leading to downregulation of H3K9ac, an active histone code.

PRAME loss mimics ETO treatment in proliferation of melanoma cells and p27 induction

As described previously, PRAME increases the proliferation of melanoma cells (16). To determine whether p53-dependent PRAME repression occurs at the cellular level, we evaluated the effect of PRAME knockdown and ETO treatment on the proliferation of melanoma cells. Colony formation was significantly decreased by PRAME depletion (Fig. 4A, B, left) and ETO treatment (Fig. 4A, B, right) in A375P and A375SM cells. Colony formation by p53-high/PRAME-low A375P cells was less efficient than by p53-low/PRAME-high A375SM cells. Other effects of PRAME loss and ETO treatment were investigated by monitoring expression of the cell cycle inhibitor p27 (also known as KIP1). As shown in Fig. 4C, the protein levels of p53 and PRAME were opposite in A375P and A375SM cells. p27 was upregulated by PRAME knockdown and ETO treatment in A375P cells. Taken together, these results suggest that ETO-induced p53 activation suppresses the proliferation of melanoma cells through the downregulation of PRAME.


High PRAME expression has been implicated in the progression of several tumors, including melanoma. However, no attempts have been made as yet to control the PRAME expression. Here we present evidence that p53 activation exerts transcriptional repression of PRAME: increased PRAME expression and downregulation upon p53 knockdown and overexpression, respectively; higher luciferase activity of the PRAME-luciferase reporter gene in p53-null cells than in p53wt cells. In addition, we demonstrated that PRAME repression by p53 activation is p53-dependent. Finally, we dissected the underlying mechanism and suggested that p53 binds to the p53 response element in the PRAME promoter and recruits histone deacetylase 1, leading to the reduced accumulation of acetylated histone H3 at lysine 9 (Fig. 4D). Our findings support that PRAME may be a potential therapeutic target of p53 activation in melanoma.

The mutation frequency of p53 in melanoma is 10-20%, compared to about 50% in other tumors (22, 23). Intriguingly, p53 is frequently inactivated in melanoma harboring wild-type p53 (24, 25). Given the 10-20% mutation rate of p53 in melanoma, the next question is raised whether the PRAME mRNA expression is higher in p53-mutant melanoma samples. From the analysis of the TCGA human melanoma databases using the UALCAN and USCS Xena browser tools, we did not see a statistically significant difference in PRAME expression between patients carrying p53wt and mutant p53. This observation may be due to the small population of patients with a particular p53 mutation because of the low mutation rate of p53 in melanoma and the variety of p53 mutations, including nonsense, missense, deletion, and frameshift mutations. At present, it is not clear whether p53 mutations result in p53 loss or point mutations. It is well established that hotspot forms of p53 mutations, mostly accumulated in the DNA binding domain (DBD), alter the transcription of target genes and become oncogenic (26). For example, p53 DBD mutants (R273H, R175H) enhance the transcriptional activity of CXCL5 and CXCL8 (27), while suppressing the expression of the differentiation inhibitor Id2 (28) and the apoptosis receptor CD95 (29). Like Id2 and CD95, mutant p53 may inhibit PRAME expression. However, it is more plausible that mutant p53 does not affect PRAME repression or instead increases PRAME expression in cancer cells that require the oncogenic properties of PRAME. Further studies are needed to determine whether mutant p53, such as mutants in the DNA binding domain or transactivation domain, represses PRAME expression.


RNA interference

To deplete PRAME mRNA, we used siPRAME targeting exon 3 (sense strand, 5’-CAGUAGAGGUGCUCGUAGA (dTdT)-3’; antisense strand, 5’-UCUACGAGCACCUCUACUG (dTdT)-3’) (Bioneer, Korea). siRNA transfection was performed with Lipofectamine RNAiMAX (Invitrogen) according to the manufacturer’s instructions. Non-targeting scramble siRNA was also purchased from Bioneer (SN-1001) and used as a negative control. For p53 knockdown, shp53 pLKO.1 was purchased from Addgene (Watertown, MA, USA), (sense strand, 5’-CCGACTCCAGTGGTAATCTACTTCAAGAGAGTAGATTACCACTGGAGTCTTTTT-3’; antisense strand, 5’-AAAAAGACTCCAGTGGTAATCTACTCTCTTGAAGTAGATTACCACTGGAGTCGG-3’ with the p53 target region shown in bold type). As a negative control, the sh-luciferase (shLuc) sequence, subcloned into the pLKO.1 vector, was also purchased from Addgene.

Real-time reverse transcription-quantitative PCR (RT-qPCR)

Total RNA was extracted by standard methods using Isol-RNA Lysis Reagent (5 PRIME, Germany). cDNA synthesis was performed using M-MLV reverse transcriptase and random primers (Invitrogen). Semi-quantitative PCR was conducted with G-Taq (Labopass, Korea). For quantitation, real-time PCR reactions were performed using SYBR Green Real-time PCR Master Mix (TOYOBO, Japan) and CFX96 Real-Time PCR Detection System (Bio-Rad, CA, USA) with the following primer pairs (forward and reverse, respectively): PRAME, 5’-CGTTTGTGG GGTTCCATTC-3’ and 5’-GCTCCCTGGGCAGCAAC-3’; p21, 5’-CTGGAGACTCTCAGGGTC-3’ and 5’-TTAGGGCTTCCTCTTGGAGA-3’; and GAPDH, 5’-GACTCCACGACGTACTCA-3’ and 5’-GTGGATATTGTTGCCATC-3’. Murine primer sets are: p53, 5’-TGCTCACCCTGGCTAAAGTT-3’ and 5’-AATGTCTCCTGGCTCAGAGG-3’; p21, 5’-TGTCGCTGTCTTGCACTCTG-3’ and 5’-CGTGGGCACTTCAGGGTTTT-3’; Pcna, 5’-GGGGTGAAGTTTTCTGCAAGT-3’ and 5’-TCAGAGCAAACGTTAGGTGAAC-3’; Prame, 5’-TCTTCTGGGGGCCCTAAGTT-3’ and 5’-GGCTGGATTGCAGACTGACT-3’. All primer sequences were verified using Primer-Blast provided by NCBI. Results were normalized to GAPDH transcript levels as an internal standard.

Western blotting (WB)

For WB, cells were lysed in RIPA buffer (50 mM Tris-Cl pH 7.5, 150 mM NaCl, 0.4% Nonidet P-40, 1 mM EDTA, 1 mM PMSF, 1 mM Na3VO4, and 1 mM NaF) supplemented with a protease inhibitor cocktail (Roche, Switzerland). Proteins were separated by electrophoresis on 8-15% SDS gels, transferred to nitrocellulose, and incubated with the following primary antibodies: anti-PRAME (sc-137188, Santa Cruz Biotechnology, CA, USA; ab219650, Abcam, Cambridge, UK), anti-p53 (sc-6243; Santa Cruz Biotechnology), anti-acetyl-p53 (Lys 382) (#2525; Cell Signaling Technology, MA, USA), anti-p27 (sc-528; Santa Cruz Biotechnology), and anti-β-actin (sc-47778; Santa Cruz Biotechnology). Blots were next incubated with goat anti-mouse IgG-HRP (sc-2005; Santa Cruz Biotechnology) or goat anti-rabbit IgG-HRP (sc-2004; Santa Cruz Biotechnology).

Chromatin immunoprecipitation (ChIP)

ChIP analysis was conducted as described previously (16). Cross-linked sheared chromatin complexes were recovered by IP using IgG, anti-p53 (sc-6243; Santa Cruz Biotechnology), anti-H3K9Ac (ab10812; Abcam), anti-HDAC1 (05-614; Upstate Biotechnology, NY, USA), and anti-Flag M2 (F1804; Sigma-Aldrich). The cross-linking was reversed according to Millipore’s protocol. DNA pellets were recovered and analyzed by semi-quantitative PCR and RT-qPCR with the following primer pairs (forward and reverse, respectively): PRAME p53RE1, 5’-AGGAATGTCTTGCAGGGCTA-3’ and 5’-AAGGGCTCATGCTTACATGG-3’; PRAME p53RE2, 5’-CCCATCTGCTCCCCACC-3’ and 5’-CCAGGGCAAAATCTCACGA-3’, and p21 p53RE; 5’-TGGCTCTGATTGGCTTTCT-3’ and 5’-TGGGGTCTTTAGAGGTCTCC-3’.

Colony formation assay

A375P and A375SM cells were transfected with siRNA or treated with etoposide (ETO, 20 μM) for 2 days before seeding in 60 mm culture dishes at a density of 3,000 cells per dish. Ten days after plating, colony formation was monitored by crystal violet staining. Stained colonies were counted from three independent assays and relative colony counts were displayed.

Statistical analysis

Statistical analyses were conducted using SigmaPlot (version 10.0; Systat Software, USA) and MedCalc (version 22.021; MedCalc Software, Belgium). The correlation between PRAME and TP53 levels was assessed using Pearson correlation test and linear regression analysis. Student’s t-test was used to compare statistical significance between groups. Data are presented as mean ± standard deviation (SD) of at least three independent experiments. Protein band density was measured using ImageJ (version 1.52e, National Institutes of Health, USA). P-values < 0.05(*), 0.01(**), or 0.005(***) were considered statistically significant.


Some additional information on Materials and Methods can be found in the supplementary section.


This study was supported by the Basic Research Program through the National Research Foundation of Korea (NRF) funded by the Ministry of Science and ICT (2021R1A4A5033289), and also supported by Korea Basic Science Institute (National research Facilities and Equipment Center) grant funded by the Ministry of Education (2023R1A6C101A045).


The authors have no conflicting interests.

Fig. 1. Negative correlation between PRAME and p53. (A) Negative correlation between p53 and PRAME expression in melanoma cell lines. Protein and mRNA levels were measured by Western blot and RT-PCR, respectively (n = 1). (B) Correlation coefficient between p53 and PRAME. The band densities of p53 and PRAME in Fig. 1A were quantified utilizing ImageJ software. (C, D) Comparison of p53 and PRAME expression. Using the GSE4570 data set, mRNA levels of p53 and PRAME were assessed in normal melanocytes (n = 2) and melanoma (n = 6). (E) Correlation of p53 and PRAME mRNA levels assessed by calculating the Pearson correlation coefficient using the GSE4570 data set. All Pearson correlation tests and linear regression analyses were performed using MedCalc software.
Fig. 2. Downregulation of PRAME by p53 activation. PRAME mRNA expression was assessed via RT-qPCR, and the data were displayed as means ± SD (n = 3, *P < 0.05; **P < 0.01; ***P < 0.005). Proteins were visualized by WB using indicated antibodies (n = 1). (A) Negative effect of p53 on PRAME expression. PRAME protein was visualized by WB in HCT116p53−/− and p53-null H1299 cells transfected with Flag-p53 expression vector (2 and 5 μg) (n = 3). (B) Effect of p53 knockdown on PRAME expression. HCT116 cells carrying wild-type p53 were transfected with a luciferase control or p53 shRNA. Twenty-four hours after transfection, the protein levels of PRAME and p27 were measured by WB (n = 1). PRAME mRNA expression was measured by RT-qPCR (n = 3, **P < 0.01). (C) Effect of ETO on PRAME expression. A375P cells were treated with 20 μM etoposide (ETO) for indicated times. Proteins were visualized by WB. PRAME band intensity was measured by ImageJ software and normalized to β-actin. The PRAME mRNA level was quantified by RT-qPCR and normalized to GAPDH (n = 3, *P < 0.05). (D) Effects of UV-C and MMS on PRAME expression. A375P cells were treated with 50 J/m2 UV-C or 2 mM methyl methane sulfonate (MMS). Proteins were visualized by WB using indicated antibodies (n = 3). PRAME mRNA levels were quantified using RT-qPCR and normalized to GAPDH (n = 3, *P < 0.05; ***P < 0.005).
Fig. 3. p53 binds to the PRAME promoter and recruits HDAC1. (A, B) Mapping of functional p53RE. Five deletion constructs carrying the luciferase gene (A) were introduced into HCT116p53−/− cells with the Flag-p53 vector for luciferase assays (B). Luciferase data are means ± SD (n = 3). *P < 0.05; **P < 0.01. (C) Nucleotide sequences of p53RE1 and RE2. Two mutants are displayed: mutRE, base substitution; ΔRE1, RE1 deletion. (D) Effect of RE1 mutation on p53-dependent transcriptional repression of the PRAME promoter. HCT116p53−/− cells were transfected with mutRE (left), ΔRE1 (right) with the Flag-p53 vector. Cell lysates were subjected to luciferase assays. Luciferase data are means ± SD (n = 3). **P < 0.01. (E) p53 binding to the RE1 of the PRAME promoter. H1299 cells were transfected with Flag or Flag-p53 for ChIP using an anti-Flag antibody. qPCR was performed using primer sets covering p53RE1 and RE2 of PRAME. The p53RE in p21 was used as the positive control. Quantification of ChIP-qPCR data; means ± SD (n = 3). **P < 0.01; ***P < 0.005. (F) HDAC1 recruitment to the RE1 of the PRAME promoter. A375 cells were treated with ETO and subjected to ChIP assay using antibodies against p53, histone deacetylase 1 (HDAC1), and acetylated histone H3 at lysine 9 (H3K9ac). Quantification was performed by qPCR using a primer set covering the RE1 of PRAME. Data are means ± SD (n = 3). *P < 0.05; ***P < 0.005.
Fig. 4. PRAME knockdown and ETO treatment suppress melanoma cell proliferation and p27 induction. (A) Effects of PRAME depletion and ETO treatment. A375SM or A375P cells were transfected with siPRAME (left) or treated with 20 μM ETO (right) and subjected to colony formation assay (n = 3). Control siRNA (siCtrl) is an unspecific scrambled siRNA purchased from Bioneer (Korea). Colonies were fixed and stained with crystal violet. (B) The number of colonies was counted from three independent experiments and the relative numbers are shown. Data are mean ± SD (n = 3). *P < 0.05; **P < 0.01; ***P < 0.005. (C) Upregulation of p27 by PRAME depletion and ETO treatment. Cell lysates prepared as described above were subjected to WB using the indicated antibodies (n = 1). β-Actin was used as the internal control. (D) Hypothetical model for the p53-mediated PRAME downregulation through HDAC1 recruitment.
  1. Ikeda H, Lethé B and Lehmann F et al (1997) Characterization of an antigen that is recognized on a melanoma showing partial HLA Loss by CTL expressing an NK inhibitory receptor. Immunity 2, 199-208.
    Pubmed CrossRef
  2. Xu Y, Zou R, Wang J, Wang ZW and Zhu X (2020) The role of the cancer testis antigen PRAME in tumorigenesis and immunotherapy in human cancer. Cell Prolif 53, e12770.
    Pubmed KoreaMed CrossRef
  3. Kern CH, Yang M and Liu WS (2021) The PRAME family of cancer testis antigens is essential for germline development and gametogenesis. Biol Reprod 105, 290-304.
    Pubmed CrossRef
  4. Oberthuer A, Hero B, Spitz R, Berthold F and Fischer M (2004) The tumor-associated antigen PRAME is universally expressed in high-stage neuroblastoma and associated with poor outcome. Clin Cancer Res 10, 4307-4313.
    Pubmed CrossRef
  5. Epping MT, Hart AA, Glas AM, Krijgsman O and Bernards R (2008) PRAME expression and clinical outcome of breast cancer. Br J Cancer 99, 398-403.
    Pubmed KoreaMed CrossRef
  6. Cazzato G, Colagrande A and Ingravallo G et al (2022) PRAME immuno-expression in cutaneous sebaceous carcinoma: a single institutional experience. J Clin Med 11, 6936.
    Pubmed KoreaMed CrossRef
  7. Cazzato G, Cascardi E and Colagrande A et al (2022) PRAME immunoexpression in 275 cutaneous melanocytic lesions: a double institutional experience. Diagnostics (Basel) 12, 2197.
    Pubmed KoreaMed CrossRef
  8. Cascardi E, Cazzato G and Ingravallo G et al (2023) PReferentially Expressed Antigen in MElanoma (PRAME): preliminary communication on a translational tool able to early detect oral malignant melanoma (OMM). J Cancer 14, 628-633.
    Pubmed KoreaMed CrossRef
  9. Yan H, Zhao RM, Wang ZJ, Zhao FR and Wang SL (2015) Knockdown of PRAME enhances adriamycin-induced apoptosis in chronic myeloid leukemia cells. Eur Rev Med Pharmacol Sci 19, 4827-4834.
  10. Zhu H, Wang J and Yin J et al (2018) Downregulation of PRAME suppresses proliferation and promotes apoptosis in hepatocellular carcinoma through the activation of p53 mediated pathway. Cell Physiol Biochem 45, 1121-1135.
    Pubmed CrossRef
  11. Chen X, Jiang M and Zhou S et al (2023) PRAME promotes cervical cancer proliferation and migration via Wnt/β-catenin pathway regulation. Cancers (Basel) 15, 1801.
    Pubmed KoreaMed CrossRef
  12. Bose M (2023) Preferentially expressed antigen in melanoma is a multifaceted cancer testis antigen with diverse roles as a biomarker and therapeutic target. Int J Transl Med 3, 334-359.
  13. Cassalia F, Danese A and Tudurachi I et al (2024) PRAME updated: diagnostic, prognostic, and therapeutic role in skin cancer. Int J Mol Sci 25, 1582.
    Pubmed KoreaMed CrossRef
  14. Qian J, Zhu ZH and Lin J et al (2011) Hypomethylation of PRAME promoter is associated with poor prognosis in myelodysplastic syndrome. Br J Haematol 154, 153-155.
    Pubmed CrossRef
  15. Zhang W, Barger CJ and Eng KH et al (2016) PRAME expression and promoter hypomethylation in epithelial ovarian cancer. Oncotarget 7, 45352-45369.
    Pubmed KoreaMed CrossRef
  16. Lee YK, Park UH, Kim EJ, Hwang JT, Jeong JC and Um SJ (2017) Tumor antigen PRAME is up-regulated by MZF1 in cooperation with DNA hypomethylation in melanoma cells. Cancer Lett 403, 144-151.
    Pubmed CrossRef
  17. Dai C and Gu W (2010) p53 post-translational modification: deregulated in tumorigenesis. Mol Med 16, 528-536.
    Pubmed KoreaMed CrossRef
  18. Lee JY and Gu W (2013) SIRT1: regulator of p53 deacetylation. Genes Cancer 4, 112-117.
    Pubmed KoreaMed CrossRef
  19. Riley T, Sontag E, Chen P and Levine A (2008) Transcriptional control of human p53-regulated genes. Nat Rev Mol Cell Biol 9, 402-412.
    Pubmed CrossRef
  20. Ceribelli M, Alcalay M, Viganò MA and Mantovani R (2006) Repression of new p53 targets revealed by ChIP on chip experiments. Cell Cycle 5, 1102-1110.
    Pubmed CrossRef
  21. Rinn JL and Huarte M (2011) To repress or not to repress: this is the guardian's question. Trends Cell Biol 21, 344-353.
    Pubmed CrossRef
  22. Luca M, Lenzi R, Leejackson D, Gutman M, Fidler I and Bareli M (1993) p53 mutations are infrequent and do not correlate with the metastatic potential of human-melanoma cells. Int J Oncol 3, 19-22.
    Pubmed CrossRef
  23. Weiss J, Heine M and Arden KC et al (1995) Mutation and expression of TP53 in malignant melanomas. Recent Results Cancer Res 139, 137-154.
    Pubmed CrossRef
  24. Houben R, Hesbacher S and Schmid CP et al (2011) High-level expression of wild-type p53 in melanoma cells is frequently associated with inactivity in p53 reporter gene assays. PLoS One 6, e22096.
    Pubmed KoreaMed CrossRef
  25. Avery-Kiejda KA, Bowden NA and Croft AJ et al (2011) p53 in human melanoma fails to regulate target genes associated with apoptosis and the cell cycle and may contribute to proliferation. BMC Cancer 11, 203.
    Pubmed KoreaMed CrossRef
  26. Pfister NT and Prives C (2017) Transcriptional regulation by wild-type and cancer-related mutant forms of p53. Cold Spring Harb Perspect Med 7, a026054.
    Pubmed KoreaMed CrossRef
  27. Yeudall WA, Vaughan CA and Miyazaki H et al (2012) Gain-of-function mutant p53 upregulates CXC chemokines and enhances cell migration. Carcinogenesis. 33, 442-451.
    Pubmed CrossRef
  28. Yan W, Liu G, Scoumanne A and Chen X (2008) Suppression of inhibitor of differentiation 2, a target of mutant p53, is required for gain-of-function mutations. Cancer Res 68, 6789-6796.
    Pubmed KoreaMed CrossRef
  29. Zalcenstein A, Stambolsky P and Weisz L et al (2003) Mutant p53 gain of function: repression of CD95 (Fas/APO-1) gene expression by tumor-associated p53 mutants. Oncogene 22, 5667-5676.
    Pubmed CrossRef

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