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Targeting epigenetic regulators for cancer therapy: mechanisms and advances in clinical trials

Yuan Cheng, Cai He, Manni Wang and 5 more

Signal Transduction and Targeted Therapy · 2019 · Review · CC BY

Abstract

Epigenetic alternations concern heritable yet reversible changes in histone or DNA modifications that regulate gene activity beyond the underlying sequence. Epigenetic dysregulation is often linked to human disease, notably cancer. With the development of various drugs targeting epigenetic regulators, epigenetic-targeted therapy has been applied in the treatment of hematological malignancies and has exhibited viable therapeutic potential for solid tumors in preclinical and clinical trials. In this review, we summarize the aberrant functions of enzymes in DNA methylation, histone acetylation and histone methylation during tumor progression and highlight the development of inhibitors of or drugs targeted at epigenetic enzymes.

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From PubMed Central, U.S. National Library of Medicine, as current on Oct 16, 2026. Paragraph numbers are ours, so an answer can point at one.

Introduction

1After the discovery of DNA and the double helix structure, classic genetics has long assumed that the sequences of DNA determine the phenotypes of cells. DNA is packaged as chromatin in cells, with nucleosomes being the fundamental repeating unit. Four core histones (H2A, H2B, H3, and H4) form an octamer and are then surrounded by a 147-base-pair (bp) segment of DNA. Nucleosomes are separated by 10–60 bp DNA. Researchers have gradually found organisms that share the same genetic information but have different phenotypes, such as somatic cells from the same individual that share a genome but function completely differently. The term epigenetics was first proposed and established in 1942 when Conrad Waddington tried to interpret the connection between genotype and phenotype. Later, Arthur Riggs and his group interpreted epigenetics as inherited differences in mitosis and meiosis, which could explain the changes in phenotypes. They were both trying to find the link between genotype and phenotype. Epigenetics is usually referred to as a genomic mechanism that reversibly influences gene expression without altering DNA sequences. Holliday assumed that epigenetics was also mitotically and/or meiotically heritable without DNA sequence change. Aberrant DNA methylation could be repaired via meiosis, but some patterns are still transmitted to offspring. This phenomenon covers a wide range of cellular activities, such as cell growth, differentiation, and disease development, and is heritable. Generally, epigenetic events involve DNA methylation, histone modification, the readout of these modifications, chromatin remodeling and the effects of noncoding RNA. The elements involved in different modification patterns can be divided into three roles, “writer,” “reader,” and “eraser”. The “writers” and “erasers” refer to enzymes that transfer or remove chemical groups to or from DNA or histones, respectively. “Readers” are proteins that can recognize the modified DNA or histones (Fig. 1). To coordinate multiple biological processes, the epigenome cooperates with other regulatory factors, such as transcription factors and noncoding RNAs, to regulate the expression or repression of the genome. Epigenetics can also be influenced by cellular signaling pathways and extracellular stimuli. These effects are temporary and yet long-standing. Given the importance of epigenetics in influencing cell functions, a better understanding of both normal and abnormal epigenetic processes can help to understand the development and potential treatment of different types of diseases, including cancer.

2The etiology of cancer is quite complicated and involves both environmental and hereditary influences. In cancer cells, the alteration of genomic information is usually detectable. Like genome instability and mutation, epigenome dysregulation is also pervasive in cancer (Fig. 2). Some of the alterations determine cell function and are involved in oncogenic transformation. However, by reversing these mutations by drugs or gene therapy, the phenotype of cancer can revert to normal. Holliday proposed a theory that epigenetic changes are responsible for tumorigenesis. The alteration of cellular methylation status by a specific methyltransferase might explain the differences in the probability of malignant transformation. In clinical settings, we noticed that although cancer patients share the same staging and grade, they present totally different outcomes. In tumor tissues, different tumor cells show various patterns of histone modification, genome-wide or in individual genes, indicating that epigenetic heterogeneity exists at a cellular level. Likewise, using molecular biomarkers is thought to be a potential method to divide patients into different groups. It is important to note that tumorigenesis is the consequence of the combined action of multiple epigenetic events. For example, the repression of tumor suppressor genes is usually caused by methylation of DNA CpG islands together with hypoacetylated and hypermethylated histones. During gene silencing, several hallmarks of epigenetic events have been identified, including histone H3 and H4 hypoacetylation, histone H3K9 methylation, and cytosine methylation.,

3Therefore, epigenetics enables us to investigate the potential mechanism underlying cancer phenotypes and provides potential therapy options. In this review, we focused and briefly expanded on three aspects of epigenetics in cancer: DNA methylation, histone acetylation and histone methylation. Finally, we summarized the current developments in epigenetic therapy for cancers.

DNA methylation

4The DNA methylation pattern in mammals follows certain rules. Germ cells usually go through a stepwise demethylation to ensure global repression and suitable gene regulation during embryonic development. After implantation, almost all CpGs experience de novo methylation except for those that are protected. Normal dynamic changes in DNA methylation and demethylation based on altered expression of enzymes have been known to be associated with aging., However, inappropriate methylation of DNA can result in multiple diseases, including inflammatory diseases, precancerous lesions, and cancer.– Of note, de novo methylation of DNA in cancer serves to prevent reactivation of repressed genes rather than inducing gene repression. Because researchers have found that over 90% of genes undergoing de novo methylation in cancer are already in a repressed status in normal cells. Nevertheless, aberrant DNA methylation is thought to serve as a hallmark in cancer development by inactivating gene transcription or repressing gene transcription and affecting chromatin stability.

5The precise mechanism by which DNA methylation affects chromatin structure unclear, but it is known that methyl-DNA is closely associated with a closed chromatin structure, which is relatively inactive. Hypermethylation of promoters and hypomethylation of global DNA are quite common in cancer. It is widely accepted that gene promoters, especially key tumor suppressor genes, are unmethylated in normal tissues and highly methylated in cancer tissues. P16, a tumor suppressor encoded by CDKN2A, has been found to gain de novo methylation in ~20% of different primary neoplasms. Mutations in important and well-studied tumor-suppressive genes, such as P53 and BRCA1, are frequently identified in multiple cancers.– Studies have found that the level of methylation is positively associated with tumor size. In support of this, a whole-genome methylation array analysis in breast cancer patients found significantly increased CpG methylation in FES, P2RX7, HSD17B12, and GSTM2 coincident with increasing tumor stage and size. After analysis of long-range epigenetic silencing at chromosome 2q14.2, methylation of EN1 and SCTR, the first well-studied example of coordinated epigenetic modification, was significantly increased in colorectal and prostate cancers., EN1 methylation has also been observed to be elevated by up to 60% in human salivary gland adenoid cystic carcinoma. Of note, only ~1% of normal samples exhibited EN1 CpG island hypermethylation. Therefore, the significant difference between cancer cells and normal cells makes EN1 a potential cancer marker in diagnosis. In human pancreatic cancer, the APC gene, encoding a regulator of cell junctions, is hypermethylated by DNMT overexpression. During an analysis of colorectal disease methylation patterns, researchers found several genes that showed significant changes between precancerous diseases and cancers, including RUNX3, NEUROG1, CACNA1G, SFRP2, IGF2 DMR0, hMLH1, and CDKN2A. In the human colon cancer cell line HCT116, hMLH1 and CDKN2A always bear genetic mutation and hypermethylation of one allele, and this leads to inactivation of key tumor suppressors. It is known that p16, p15, and pax6 are usually aberrantly methylated in bladder cancer and show enhanced methylation in cell culture.. Unlike gene promoter methylation, gene body methylation usually results in increased transcriptional activity. This process often occurs in CpG-poor areas and causes a base transition from C to T. The hypermethylation of specific CpG islands in cancer tissues is informative of mutations when the gene in normal tissues is unmethylated. One representative marker is glutathione S-transferase-π (GSTP1), which is still the most common alteration in human prostate cancer. Recently, DNA methylation in cancer has generally been associated with drug resistance and predicting response to treatment. For example, MGMT (O-6-methylguanine DNA methyltransferase) hypermethylation is still the best independent predictor of response to BCNU (carmustine) and temozolomide in gliomas because hypermethylation of MGMT makes tumor cells more sensitive to treatments and is associated with regression of tumor and prolonged overall survival., Similarly, MGMT is also a useful predictor of response to cyclophosphamide in diffuse large B-cell lymphoma (Table 1).

DNA methyltransferases (DNMTs)

6DNA methylation is a covalent modification of DNA and is one of the best-studied epigenetic markers. It plays an important role in normal cell physiology in a programmed manner. The best-known type of DNA methylation is methylation of cytosine (C) at the 5th position of its carbon ring (5-mC), especially at a C followed by a guanine (G), so-called CpG sites. Non-CpG methylation, such as methylation of CpA (adenine) and CpT (thymine), is not common and usually has restricted expression in mammals. CpG islands traverse ~60% of human promoters, and methylation at these sites results in obvious transcriptional regression. Meanwhile, among the ~28 million CpGs in the human genome in somatic cells, 60–80% are methylated in a symmetric manner and are frequently found in promoter regions., The process of DNA methylation is regulated by the DNA methyltransferase (DNMT) family via the transfer of a methyl group from S-adenosyl-L-methionine (SAM) to cytosine. There are five members of the DNMT family: DNMT1, DNMT2, DNMT3a, DNMT3b, and DNMT3L. DNMT1 is responsible for the maintenance of methyl-DNA, recognizes hemimethylated DNA strands and regenerates the fully methylated DNA state of DNA during cell division. In a recent study, DNMT1 with Stella, a factor essential for female fertility, was responsible for the establishment of the oocyte methylome during early embryo development. DNMT3a and DNMT3b are regarded as de novo methylation enzymes that target unmethylated CpG dinucleotides and establish new DNA methylation patterns, but they have nonoverlapping functions during different developmental stages., DNMT2 and DNMT3L are not regarded as catalytically active DNA methyltransferases. DNMT2 functions as an RNA methyltransferase, while DNMT3L contains a truncated inactive catalytic domain and acts as an accessory partner to stimulate the de novo methylation activity of DNMT3A. The DNA methyltransferase-like protein DNMT3L can modulate DNMT3a activity as a stimulatory factor.

7During aberrant DNA methylation, DNMTs play an important role. Compared with DNMT1 and DNMT3a, DNMT3b was significantly overexpressed in tumor tissues. Overexpression of DNMT1, DNMT3a, and DNMT3b has been observed in multiple cancers, including AML, CML, glioma, and breast, gastric, colorectal, hepatocellular, pancreatic, prostate, and lung cancers. In cervical cancer patients, DNMT1 was expressed in more than 70% of cancer cells, whereas only 16% of normal cells expressed DNMT1. The higher level of DNMT1 expression was also associated with worse prognosis. The expression of DNMT1, DNMT3a, and DNMT3b has been observed to be elevated in acute myeloid leukemia (AML) and various solid cancers. These three methyltransferases do not show significant changes in the chronic phase of chronic myeloid leukemia (CML), but they are significantly increased during progression to the acute phase in CML., Notably, downregulation of DNMTs can also lead to tumorigenesis (Table 1).

Methyl-CpG recognition proteins

8How DNA methylation leads to gene repression has been considered in many studies. Several hypotheses have been proposed. Three methyl-CpG binding domain protein (MeCP) families can read the established methylated DNA sequences and in turn recruit histone deacetylases, a group of enzymes responsible for repressive epigenetic modifications, to inhibit gene expression and maintain genome integrity., The first group is methyl-CpG binding domain (MBD) proteins, including MeCP2, MBD1, MBD2, and MBD4. MeCP1 is a complex containing MBD2, the histone deacetylase (HDAC) proteins HDAC1 and HDAC2, and the RbAp46 and RbAp48 proteins (also known as RBBP7 and RBBP4). MBD3 is unlike the other four family members and is not capable of binding to methylated DNA but instead binds to hydroxymethylated DNA. The zinc-finger and BTB domain-containing protein family is the second group and comprises three structurally different proteins, KAISO (ZBTB33), ZBTB4 and ZBTB38, which bind to methylated DNA via zinc-finger motifs. The third family includes two ubiquitin-like proteins with PHD and RING finger domains, UHRF1 and UHRF2, which recognize 5-mC via RING finger-associated (SRA) domains. On the other hand, methylation of DNA can also be a barrier for certain transcription factors to bind to promoter sites such as AP-2, c-Myc, CREB/ATF, E2F, and NF-kB.

9As for methyl-group binding proteins, many studies have investigated their roles in various cancers, but the mechanism underlying these alterations remains unclear. MBD proteins cooperate with other proteins to regulate gene transcription., However, the role of MBD1 and MBD2 has not been identified in human lung or colon cancer, with only limited mutations being detected. Furthermore, loss of MBD1 did not show any carcinogenic effect in MBD−/− mice. Compared with MBD1, MBD2 shows more effect on tumorigenesis. Deficiency of MBD2 strongly suppresses intestinal tumorigenesis in APCMin-background mice. A possible reason is that many important signaling pathways are downregulated in colorectal cancer, and loss of MBD2 leads to reexpression of these genes. Meanwhile, inhibition of MBD2 shows promising effects on suppression of the tumorigenesis of human lung cancer and colon cancer. Although MBD3 does not directly bind to methylated DNA, it regulates the methylation process via interactions with other proteins, such as MBD2 and HDAC. For example, application of an HDAC inhibitor in lung cancer cells upregulated p21 (also known as CDKN1A) and downregulated ErbB2, leading to inhibition of cancer cell growth. Silencing of MBD3 blocked the effects of an HDAC inhibitor. MBD3 and MBD2 form a complex, nucleosome remodeling and deacetylase (NuRD), which interacts with histone-demethylating enzymes to regulate gene expression in cancer. Mutation of MBD4 has been found in colorectal cancer, endometrial carcinoma and pancreatic cancer. Furthermore, this mutation unexpectedly affects the stability of the whole genome, not only CpG sites. Knockout of MBD4 indeed increased tumorigenesis in APCMin-background mice, which makes MBD4 a tumor suppressor. MBD4 is important in DNA damage repair, given the interaction between MBD4 and MMR. In contrast, the expression of MeCP2 and the UHRF family tends to promote tumor growth.– In the KAISO family, KAISO directly binds to p120ctn, a protein with an alternative location in some cancer cells, and they together regulate cell adhesion and motility., However, deficiency of ZBTB4 contributes to tumorigenesis (Table 1).

DNA-demethylating enzymes

10DNA methylation is a stable and highly conserved epigenetic modification of DNA in many organisms. However, loss of 5-mC and DNA demethylation have been identified in different biologic processes. For example, DNA demethylation is important for primordial germ cells (PGCs) to gain pluripotent ability., DNA demethylation is actively regulated by the TET protein family (ten-eleven translocation enzymes, TET1-3) via the removal of a methyl group from 5-mC. These three proteins differ from each other in terms of expression depending on the developmental stage and cell type. TETs oxidize 5-mC in an iterative manner and catalyze the conversion of 5-mC to 5-hydroxymethylcytosine (5-hmC), which is a key intermediate in the demethylation process. 5-hmC, as a relatively stable intermediate substrate, is less prone to further oxidation by TET proteins than 5-mC. However, overexpression of only TET1 and TET2 can cause a global decrease of 5-mC. Stepwise oxidation of 5-hmC by TET proteins can yield two products: 5-formylcytosine (5-fC) and 5-carboxylcytosine (5-caC). These two molecules can be excised by thymine-DNA glycosylase (TDG) and eventually be repaired to unmodified C. DNA demethylation or restoration of the unmodified cytosine can also occur passively through replication-dependent dilution of 5-mC.

11Disruption of normal DNA demethylation is thought to be associated with oncogenesis. TET proteins were initially associated with leukemia. Researchers have found that in a small number of AML patients, TET1 is fused to MLL via the chromosome translocation t(10;11)(q22;q23). Further studies found that TET2 was more widely expressed in different tissues than TET1 and TET3. Analyses revealed that mutation or deficiency of TET2 occurred in ~15% of patients with myeloid cancers, including myelodysplastic syndrome (MDS), myeloproliferative disorders, and AML. In patients with CML, mutation of TET2 has been detected in ~50% of patients. Although TET2 mutations have been found in several myeloid malignancies, their prognostic effect remains controversial. Based on the phenomenon that mutation of TET2 was elevated in patients whose disease transformed from chronic myeloid malignancy to AML, researchers considered that TET2 loss was important for cells to regain the ability to self-renew. The role of TET proteins has also been investigated in several solid tumors. Compared with surrounding normal tissues, 5-hmC is significantly reduced in human breast, liver, lung, pancreatic, and prostate cancers with reduced expression of TET family proteins. Deficiency of TET1 in prostate and breast cancer is associated with tumor cell invasion and breast xenograft tumor formation via the inhibition of the methylation of metalloproteinase (TIMP) family proteins 2 and 3. Loss of 5-hmC is an epigenetic hallmark of melanoma, and thus, introducing TET2 into melanoma cells results in suppression of tumor growth and increased survival in an animal model (Table 1).

Histone modification

12Histone modification can occur to the flexible tails as well as the core domain of histones, including those sites that are buried by DNA. In particular, the flexible histone tails are enriched with basic Lys/Arg and hydroxyl group-containing Ser/Thr/Tyr residues, thereby being hotspots for hallmark histone modifications. The tails extend from the surface of the nucleosome and are readily modulated by covalent posttranslational modification (PTM). PTMs modify histones by adding or removing chemical groups and regulate many biological processes via the activation or inactivation of genes. These processes mainly include acetylation and methylation of lysines (K) and arginines (R), phosphorylation of serines (S) and threonines (T), ubiquitylation, and sumoylation of lysines. In addition to those mentioned and discussed above, histone modifications also include citrullination, ADP-ribosylation, deamination, formylation, O-GlcNAcylation, propionylation, butyrylation, crotonylation, and proline isomerization at over 60 amino acid residues., In addition to conventional PTMs, novel PTM sites are also found outside of the N-terminal tails.

13Histone modifications at certain sites, such as promoters and enhancers, are thought to be largely invariant, whereas a small number of these sites remain dynamic. H3K4me1 and H3K27ac, two dynamic modifications, were identified to activate enhancers and regulate gene expression. H3K9ac and H3K9me3 are two common modifications at promoters., Appropriate histone modifications are important in gene expression and human biology; otherwise, alterations in PTMs may be associated with tumorigenesis. Analysis of cancer cells reveals that they exhibit aberrant histone modifications at individual genes or globally at the single-nuclei level., Understanding histone modification patterns in cancer cells can help us to predict and treat cancers. Thus far, most studies have focused on aberrant modifications within an individual site, such as H4K20me3 or H4K16ac, rather than enzymatic activity-associated abnormalities. Generally, alterations in histone modifications occur at an early stage and accumulate during tumorigenesis.

Histone acetylation (lysine)

14Histone acetylation occurs at multiple lysine residues at the N-terminus via the catalysis of histone acetyltransferases (HATs), also named lysine acetyltransferases (KATs). Histone acetylation regulates the compaction state of chromatin via multiple mechanisms, such as neutralizing the basic charge at unmodified lysine residues, and is associated with active transcription, especially at gene promoters and enhancers and the gene body; it also facilitates the recruitment of coregulators and RNA polymerase complexes to the locus., To date, HATs and histone deacetylases (HDACs) are the two of the best characterized groups of enzymes involved in histone PTMs. HATs transfer the acetyl groups from acetyl-CoA cofactors to lysine residues at histones, whereas the role of HDACs is the opposite, which makes histone acetylation a highly reversible process.

Histone acetyltransferases

15HATs are predominantly located in the nucleus, but multiple lines of evidence have shown lysine acetylation in the cytoplasm, and their acetylation is associated with key cellular events. In addition, lysine acetylation found outside histones reminds us of the role of HATs in nonhistone PTMs. The first HAT was identified in yeast, and was named HAT1, and was then isolated from tetrahymena as HAT A by Allis and coworkers. In humans, HATs can be roughly divided into three groups: general control nondepressible 5 (GCN5)-related N-acetyl transferase (GNAT) (based on the protein Gcn5 found in yeast; including GCN5 and PCAF), MYST (based on the protein MOZ; including MOZ, MOF, TIP60, and HBO1), and p300/cAMP-responsive element-binding protein (CBP). Other HATs, including nuclear receptors and transcription factors, such as SRC1, MGEA5, ATF-2, and CLOCK, also harbor the ability to acetylate histones. Notably, a number of acetyltransferases also perform protein acetylation outside histones, such as TFIIB, MCM3AP, ESCO, and ARD1. Knockout of CBP/p300 is lethal for early embryonic mouse models., The acetyl group transfer strategies for each HAT subfamily are different. For the GCN5 and PCAF family, the protein crystal structure shows a conserved glutamate in the active site. Blockade of this amino leads to a significantly decreased acetylation function., Similarly, there is also a conserved glutamate plus a cysteine residue located at active sites of MYST family proteins. Unlike the other two families, the p300/CBP HAT subfamily has two other potential conserved residues, a tyrosine and a tryptophan. Generally, their catalytic mechanisms of acetyl group transfer can be divided into two groups. The GNAT family depends on a sequential ordered mechanism, whereas the members of the MYST family use a so-called ping-pong (i.e., double displacement) catalytic mechanism, which means that the acetyl groups are first transferred to a cysteine residue and then transferred to a lysine residue. In addition to differences in the acetyl transfer mechanism, HAT subfamilies, even different proteins in the same family, also have remarkable diversity in targeting sites.

16Appropriate acetylation within cells is important since upregulation or downregulation of HATs is associated with tumorigenesis or poor prognosis., Compared with solid tumors, the association between histone modifications and cancer has been widely investigated in hematological malignancies. Germline mutation of CBP results in Rubinstein-Taybi syndrome along with an increased predisposition to childhood malignancies. Meanwhile, loss of another family member, p300, has also been associated with hematological malignancies., Therefore, both CBP and p300 seem to function as tumor suppressors. During cancer development, the expression of HAT genes can be disrupted by chromosomal translocations, although these are rare events. Generation of the fused protein CBP-MOZ is the result of the t(8,16)(p11,p13) translocation in AML. Translocation of t(10;16)(q22;p13) leads to the CBP-MORF chimera. Similarly, p300-MOZ, MLL-CBP, and MLL-p300 (MLL, mixed lineage leukemia) have also been identified in hematological malignancies.– Generally, chromosomal rearrangements involving CBP are more common than those involving p300. Researchers have also investigated solid tumors, which are less mutated. The expression of translocated P300 in laryngeal squamous cell carcinoma (LSCC) tissue is much higher than that in adjacent normal tissue and is associated with advanced stage and poor prognosis. Missense point mutations in p300 are found in colorectal adenocarcinoma, gastric adenocarcinoma and breast cancer with quite low incidences., Rare inactivating mutations in CBP and PCAF have only been identified in cancer cell lines but not primary tumors. Based on these findings, we hypothesize that the differences between cell lines and primary tumors cannot be ignored. Amplified in breast cancer 1 (AIB1), also frequently called NCOA3 (nuclear receptor coactivator 3) or SRC3 (steroid receptor coactivator 3), is overexpressed in ~60% of human breast cancers, and increased levels of AIB1 are associated with tamoxifen resistance and decreased overall survival. Steroid receptor coactivator 1 (SRC1) is also associated with the chromosomal translocation t(2;2)(q35;p23), which results in PAX3–NCOA1 gene fusion in rhabdomyosarcoma without a consistent genetic abnormality during embryonic development (Table 2).

Acetyl-lysine recognition proteins

17The bromodomain (BRD) motif is an ~110-amino-acid conserved protein module and is regarded as the first and sole histone-binding module that contains a hydrophobic pocket to identify acetyl-lysine. The specificity of different BRDs depends on the sequences within the loops that form the hydrophobic pocket. Therefore, each BRD has a preference for different histones., In addition to their recognition of acetyl-lysine, BRDs are also capable of interacting with other chromatin molecules, such as plant homeodomain (PHD) finger motifs or another BRD. To date, 42 proteins containing bromodomains and 61 unique bromodomains have been discovered., Based on the sequence length and sequence identity of BRDs, the human BRD family can be divided into nine groups and one additional set of outliers, which has been well illustrated in published papers., Different BRD-containing proteins contain one to six BRDs. Intriguing, the most notable and well-studied bromodomain proteins are also HATs, such as PCAF, GCN5, and p300/CBP. Yaf9, ENL, AF9, Taf14, Sas5 (YEATS), and double PHD finger (DPF) have also been discovered to be acyl-lysine reader domains., Human MOZ and DPF2 are two proteins containing the DPF domain. Mutations in the YEATS and DPF domains are associated with cancer. For example, mutation of AF9 has been found in hematological malignancies, and ENL dysregulation leads to kidney cancer.,

18Another important family is the BRD and extraterminal domain (BET) protein family, including BRD2, BRD3, BRD4, and BRDt, and this family shares two conserved N-terminal bromodomains and a more divergent C-terminal recruitment domain., These bromodomain proteins are critical as mediators of gene transcriptional activity. Of note, bromodomains have also been found in some histone lysine methyltransferases, such as ASH1L and MLL. BRDs are promiscuous domains and have been discussed in other well-constructed papers., In this review, we focus on the role of BRDs in tumorigenesis.

19As histone acetylation “readers”, bromodomain proteins play important roles in tumorigenesis. BRD4 recruits the positive transcription elongation factor complex (P-TEFb), a validated target in chronic lymphocytic leukemia associated with c-Myc activity.– Chromosomal translocation of BRD4, via the t(15;19) translocation, results in the generation of the fusion protein BRD4-NUT (nuclear protein in testis), which is found in NUT midline carcinoma (NMC). Importantly, inhibition of BRD4-NUT induces differentiation of NMC cells. Moreover, BRD4 is required for the maintenance of AML with sustained expression of Myc (Table 2).

Histone deacetylases

20Histone deacetylases (HDACs) have recently attracted increasing attention. In humans, the genome encodes 18 HDACs. In contrast to the function of HATs, HDACs usually act as gene silencing mediators and repress transcription. Similarly, HDACs are expressed not only in the nucleus but also in the cytoplasm, and their substrates are also not limited to histones. Based on sequence similarity, HDACs can be divided into four classes: class I HDACs, yeast Rpd3-like proteins, are transcriptional corepressors and have a single deacetylase domain at the N-terminus and diversified C-terminal regions (HDAC1, HDAC2, HDAC3, and HDAC8); class II HDACs, yeast Hda1-like proteins, have a deacetylase domain at a C-terminal position (HDAC4, HDAC5, HDAC6, HDAC7, HDAC9, and HDAC10); class III HDACs are yeast silent information regulator 2 (Sir2)-like proteins (SIRT1, SIRT2, SIRT3, SIRT4, SIRT5, SIRT6, and SIRT7); and class IV involves one protein (HDAC11). The class IV protein shares sequence similarity with both class I and class II proteins., Classes I, II, and IV are included in the histone deacetylase family, whereas class III HDACs belong to the Sir2 regulator family. The catalytic mechanisms for these two families are different; classes I, II, and IV are Zn2+-dependent HDACs, whereas sir2-like proteins (sirtuins) are nicotinamide adenine dinucleotide (NAD+)-dependent HDACs and are also capable of mono-ADP-ribosyltransferase activity, another pattern of histone modification. Intriguingly, SIRT4 is thought to have more mono-ADP-ribosyltransferase activity than HDAC activity. SIR2 and SIRT6 seem to have equal levels of both mono-ADP-ribosyltransferase and HDAC activities., Moreover, after revealing the crystal structure of SIRT5, researchers found that SIRT5 is also a lysine desuccinylase and demalonylase. Therefore, the diversity of the sirtuin family makes them a group of multifunctional enzymes.

21So far, the major known recognition sites of each HDAC are different, and these largely remain to be uncovered. For example, HDAC3 is thought to deacetylate H4K8 and H4K12, but in an HDAC3-knockout HeLa cell line, the acetylation levels of H4K8 and H4K12, even the overall acetylation levels of H3 and H4, were comparable with those in wild-type cells. Nevertheless, HDAC1 or HDAC3 siRNA can indeed increase the acetylation levels of H3K9 and H3K18. Therefore, partially because of the functional complementation and diversity within HDAC families, especially in class I, II, and IV, it is difficult to identify the specific substrates of certain HDACs. However, the substrates of the sirtuin family are quite clear. It is notable that because SIRT4 and SIRT5 are only located in mitochondria, they have no effect on histones. However, nonhistone lysine acetylation is also prevalent, since more than 3600 acetylation sites on 1750 proteins have been identified. The tumor suppressor p53 and the cytoskeletal protein α-tubulin are two representative substrates of HDACs.– Notably, HDACs are also capable of regulating gene transcription by deacetylating other proteins that are responsible for epigenetic events, such as DNMTs, HATs, and HDACs., Another phenomenon is that some HDACs have to form a complex along with other components to function as transcriptional corepressors, which provides ideas and methods to design novel HDAC inhibitors. The Sin3, NuRD, and CoREST complexes are three complexes containing HDAC1 and HDAC2. Studies have found that purified HDAC1 or HDAC2 without associated components shows fairly weak deacetylation activity in vitro. HDAC3 interacts with the corepressors SMRT/NCoR to form the functional complexes, which significantly increases HDAC3 activity. NCoR also interacts with HDAC1, HDAC2 and the class II deacetylases HDAC4, HDAC5, and HDAC7, but usually not in the form of a complex., Deleted in breast cancer 1 (DBC1) and active regulator of SIRT1 (AROS) are two proteins that are able to bind to SIRT1, whereas their interactions present opposite functions. The DBC1/SIRT1 complex inhibits the deacetylation activity of SIRT1, whereas the combination of AROS and SIRT1 stimulates the activity of SIRT1.,

22HDACs not only are able to deacetylate histones and nonhistone proteins but also interact with other epigenetic-associated enzymes, which gives them a vital role in tumorigenesis., Alterations in HDACs in cancers usually result in aberrant deacetylation and inactivation of tumor suppressor genes. For example, hypoacetylation of the promoter of p21, a tumor suppressor encoded by CDKN1A, can be reversed by HDAC inhibitors, resulting in an antitumor effect. A screen of the mutations in several HATs and HDACs, such as CBP, PCAF, HDAC1, HDAC2, HDAC5, HDAC7, and SIRT1, in more than 180 cancer samples including primary tumors and cancer cells indicated that the expression profiles of HDAC1, HDAC5, HDAC7, and SIRT1 are distinctive for colorectal cancers and normal colorectal mucosa, and the expression profiles of HDAC4 and CBP are capable of distinguishing breast cancer tissue from normal tissues (Table 2).

Histone methylation (lysine and arginine)

23Similar to the process of histone acetylation, histone methylation also consists of three important components: “writers”, histone methyltransferases (HMTs), “readers”, histone methylation-recognizing proteins, and “erasers”, histone demethylases (HDMs). Methylation of histones occurs at arginine and lysine residues. Arginine and lysine both can be monomethylated or dimethylated, whereas lysine is also capable of being trimethylated. Histone methylation can either promote or inhibit gene expression, which depends on the specific situation. For example, lysine methylation at H3K9, H3K27, and H4K20 is generally associated with suppression of gene expression, whereas methylation of H3K4, H3K36, and H3K79 induces gene expression. Mutation of H3K27M (lysine 27 to methionine) and H3K36M are two important oncogenic events, and H3K27M and H3K36M serve as drivers of pediatric gliomas and sarcomas. H3K27M has been identified in more than 70% of diffuse intrinsic pontine gliomas (DIPGs) and 20% of pediatric glioblastomas, which results in a global reduction in the trimethylation of H3K27 (H3K27me3).– However, the H3K36M mutation impairs the differentiation of mesenchymal progenitor cells and generates undifferentiated sarcoma, leading to increased levels of H3K27me3 and global loss of H3K36 (me2 and me)., Meanwhile, depletion of H3K36 methyltransferases results in similar phenotypes to those seen with H3K36M mutation. To date, KMTs (lysine methyltransferases) have been better studied than arginine methyltransferases (PRMTs) due to their sequence of discovery, different prevalence and impact. Their targets are not limited to only histones, they also modify other key proteins, such as the tumor suppressor p53, TAF10, and Piwi proteins.–

Histone methyltransferases

24All KMTs contain a 130-amino-acid conserved domain, the SET (suppressor of variegation, enhancer of Zeste, trithorax) domain, except for DOT1L. The SET domain is responsible for the enzymatic activity of SET-containing KMTs. Instead of methylating lysine residues in histone tails, DOT1L methylates lysine in the globular core of the histone, and its catalytic domain is more similar to that of PRMTs., The enzymatic activity of KMTs results in the transfer of a methyl group from S-adenosylmethionine (SAM) to a the ε-amino group of a lysine residue. The first identified KMT was SUV39H1, which targets H3K9. Sequentially, more than 50 SET-containing proteins have been identified with proven or predicted lysine methylation potential. Of note, KMTs are highly specific enzymes, meaning that they are highly selective for lysine residues they can methylate and the specific methylation degree they can achieve. For example, SUV39H1 and SUV39H2 specifically methylate histone 3 at lysine 9 (H3K9), and DOT1L only methylates H3K79. Based on their structure and sequence around the SET domain, generally, KMTs can be divided into six groups, SUV39, SET1, SET2, EZH, SMYD, and RIZ (PRDM) (reviewed by Volkel and Angrand). The Pre-SET domain of the SUV39 family contains nine conserved cysteines that coordinate with three zinc ions to function. The SET1 family members share a similar Post-SET motif that contains three conserved cysteine residues. The SET2 family possesses an AWS motif that contains 7–9 cysteines. Their SET domain is located between the AWS motif and a Post-SET motif. The members of the enhancer of zeste homolog (EZH) family are the catalytic components of polycomb repressive complexes (PRCs), which are responsible for gene silencing. EZH proteins have no Post-SET motif but have 15 cysteines in front of the SET domain and show no methylated activity as isolated proteins. PRC2 shows lysine methylation activity through its catalytic components, EZH2 or its homolog EZH1. EZH2 can methylate not only histone H3 but also histone H1 at lysine 26. The SMYD family members, which are SET and MYND domain-containing proteins, possesses a MYND (myeloid-nervy-DEAF1) domain, a zinc-finger motif responsible for protein–protein interaction. The RIZ (PRDM) family is a large family containing a homolog of the SET domain, the PR domain. The PR and SET domains share 20–30% sequence identity and are both capable of inducing histone H3 methylation. However, most members of the RIZ family responsible for histone methylation are still unknown. So far, two of them have been proven to induce the methylation of histones: PRDM2 (RIZ1) is associated with H3K9 methylation; and Meisetz, the mouse homolog of PRDM9, trimethylates H3K4. Meanwhile, PRDM1 has been identified to interact with EHMT2, a member of the SUV39 family. PRDM6 acts as a transcription suppressor by interacting with class I HDACs and EHMT2 to induce cell proliferation and inhibit cell differentiation. Meanwhile, the recruitment of EHMT2 is based on the formation of a complex with PRDM1. Due to the lack of a characteristic sequence or structure flanking the SET domain, other SET-containing KMTs, such as SET7/9, SET8, SUV4-20H1, and SUV4-20H2, cannot be classified into these families. Notably, some KMTs contain more than one domain, which allows them to interact with other proteins, especially other epigenetic modifying proteins. SUV39H1 possesses a chromodomain that directly binds to nucleic acids and forms heterochromatin. MLL1 recognizes unmethylated DNA through its CpG-interacting CXXC domain. SETDB1 contains an MBD that interacts with methylated DNA. The Tudor domain in SETDB1 may potentially recognize the methylation of lysine residues. ASH1 is able to interact with CBP, a HAT, via a bromodomain within ADH1.

25Protein arginine methyltransferases (PRMTs) can be divided into two groups. Among the nine PRMTs, only PRMT5, PRMT7, and PRMT9 are type II PRMTs, and the other five PRMTs, except for PRMT2, are type I PRMTs. PRMT2 was identified by sequence homology but has not shown any catalytic activity during investigations, although PRMT2 acts as a strong coactivator for androgen receptor (AR), which is thought to be associated with arginine methylation. Both types of PRMTs first catalyze the formation of monomethylarginine as an intermediate. However, sequentially, type I PRMTs can form asymmetric dimethylarginine (ADMA, Rme2a), but type II PRMTs form symmetric dimethylarginine (SDMA, Rme2s). Rme2a means two methyl groups on one ω-amino group, whereas an Rme2s has one methyl group on each ω-amino group. PRMT1-PRMT8 were investigated by Herrmann and Fackelmayer, and FBXO11 was identified as PRMT9, which symmetrically dimethylates arginine residues.

26Most enzymes for histone methylation are substrate-specific proteins; therefore, alterations in the aberrant expression of enzymes are usually associated with specific histone residue mutations. One of the best-known examples of alterations in tumorigenesis is H3K4me3, which is associated with biphenotypic (mixed lineage) leukemia (MLL). The location of the MLL gene is where chromosomal translocations in AML and ALL usually occur. When the MLL gene is translocated, the catalytic SET domain is lost, which results in MLL translocation-generated fusion proteins, which recruit DOT1L. Maintenance of MLL-associated ALL depends on the methylation of H3K79 catalyzed by DOT1L. Therefore, DOT1L is usually associated with hematological malignancies rather than solid tumors. Alteration of the EZH2-induced methylation of H3K27 has been observed in multiple cancers, including various solid tumors (prostate, breast, kidney, bladder, and lung cancers) and hematological malignancies. Meanwhile, overexpression of EZH2 has been found in multiple cancers and is associated with poor prognosis. Different mechanisms have been proposed to describe the role of EZH2 in tumorigenesis (Table 3).

Methyl-histone recognition proteins

27“Readers” of histone methylation contain several specific domains recognizing lysine or arginine methylation, such as a chromodomain, the WD40 repeat, the MBT (malignant brain tumor) domain, the Tudor domain and the PHD (plant homeodomain) finger motif. Representative chromodomain-containing proteins in humans are HP1 and Chd1, which can recognize H3K9me and H3K27me, respectively., WDR5 is a protein containing WD40 repeats. In addition to H3K4me, WDR5 prefers to bind to H3K4me2 via a histone-methylating complex and is required for maintaining H3K4me3. Later, WDR5 was shown to directly read H3R2, a “WIN” motif of MLL1, as well as symmetrical H3R2 dimethylation through the WD40 domain. L3MBTLs are a group of proteins containing three MBT repeat domains. L3MBTL1 represses gene expression via monomethylation or dimethylation of H4K20 or H1BK26. BPTF, RAG2, PYGO, and the tumor suppressor ING2 are representative proteins containing PHD finger motifs. They are all able to recognize and bind to H3K4me3. Intriguingly, DNMT3L and BHC80 also possess a PHD finger motif, but they selectively bind to unmethylated H3K4., There are a number of proteins containing Tudor domains, with a representative protein being JMJD2A. JMJD2A is a histone demethylase that equally binds to H3K4me3 and H4K20me3 (Table 3).

Histone demethylases

28The identification of histone demethylases (HDMs or KDMs) has lagged behind that of HMTs. Thus far, KDMs can be classified into two groups. The amine-oxidase type lysine-specific demethylases (LSDs) and the highly conserved JumonjiC (JMJC) domain-containing histone demethylases. LSD1 and LSD2, also known as KDM1A and B, are flavin adenine dinucleotide (FAD)-dependent amine oxidases that can only demethylate monomethylated and dimethylated lysine residues. LSD1 has been identified to specifically activate androgen receptor (AR) target genes along with AR by demethylating H3K9. The human genome codes more than 30 JMJC-containing KDMs that are able to remove methyl groups from all three methyl-lysine states. JHDM1A was the first characterized JMJC domain-containing HDM and specifically demethylates H3K36me2 and H3K36me1. Not all JMJC domain-containing proteins are able to demethylate histone proteins, such as HIF1AN and the transmembrane phosphatidylserine receptor PTDSR. JMJC-containing HDMs can be divided into six families: the JHDM1, JHDM2 (JMJD1), JHMD3 (JMJD2), JARID, PHF, and UT families. Notably, not all of these families possess the ability of histone demethylation. However, some JMJC-containing proteins, including those that are not included in these six families, contain one or more methylated-histone-binding domains. Their potential to demethylate methyl-lysine or methyl-arginine must be investigated. In addition to demethylases for lysine residues, JMJD6 is the first described arginine demethylase and lysine hydroxylase. It can remove methyl groups from H3R2 and H4R3. Another kind of protein is peptidylarginine deiminases (PADs or PADIs) or protein-arginine deiminases, which are able to convert arginine and monomethylated arginine to citrulline.

29LSD1 (KDM1A) is one of the best-studied KDMs and has been found to be increased in multiple cancers. Inhibition of LSD1 leads to global H3K4 methylation and promotes differentiation of neuroblastoma cells. Unlike KDM1A, KDM1B is mostly involved in growing oocytes with a restricted expression pattern. Similar to the dual roles of LSD1, members of the KDM2 family can either promote tumor formation or inhibit tumorigenesis. Through dimethylating H3K36 in DUSP3 (dual specific phosphatase 3), KDM2A activates ERK1/2 expression in lung cancer cells. Knockout of KDM2B in breast cancer downregulates the tumor stem cell markers ALDH and CD44 via the repression of polycomb complexes. KDM2B is also overexpressed in pancreatic ductal adenocarcinoma (PDAC) and cooperates with KrasG12D to promote PDAC formation in mouse models. The LSD1 and KDM2 family possesses context-dependent tumor-promoting and -inhibiting functions, which might depend on the different features of various cancers and the specific substrates of the enzymes. Therefore, further studies should take the dual roles of these enzymes into consideration. KDM3A, induced by hypoxia and nutrient starvation within the tumor microenvironment, shows carcinogenic effects via the promotion of tumor cell migration and invasion. Inhibition of KDM3A downregulates tumor-associated angiogenesis and macrophage infiltration., KDM3C is required for MLL-AF9 leukemia maintenance and is mutated in patients with intracranial germline tumors., KDM4A, KDM4B, and KDM4C have shown increased expression in prostate cancer with decreased levels of H3K9me2/3 and increased levels of H3K9me1. H3K9me3 is thought to be a hallmark of heterochromatic areas of the genome. In addition, KDM4 family members were the first identified demethylases targeting trimethylated lysines. Aberrant expression of KDM4 family members might lead to instability of the genome and become involved in tumorigenesis. Members of the KDM6 family usually act as tumor suppressors and are thought to cause cell growth arrest. For example, the tumor suppressor proteins p16INK4A and p14ARF, encoded by the INK4A-ARF locus, are repressed by H3K27me3. When stimulated by oncogenic factors, KDM6B is recruited to the INK4A-ARF locus and activates the transcription of these two tumor suppressors. In colorectal cancer, KDM7C is required for the efficacy of oxaliplatin and doxorubicin and for the activation of p53 (Table 3).

Noncoding RNA

30Epigenetic related noncoding RNAs (ncRNAs) include microRNAs (miRNAs), small interfering RNA (siRNAs), Piwi-interacting RNA (piRNAs), and long noncoding RNAs (lncRNAs). MiRNAs, one of the most studied ncRNAs, are small RNAs between 19 and 22 nucleotides in length that play important roles in the regulation of gene expression by controlling mRNA translation. Intriguingly, the regions that miRNAs usually target are frequently associated with carcinogenesis. Generally, they can be divided into tumor-promoting and tumor-suppressing miRNAs. During tumorigenesis, oncogenic miRNAs such as miR-155, miR-21 and miR-17-92 are usually overexpressed, and tumor-suppressive miRNAs such as miR-15-16 are downregulated. There is another type of miRNA, cellular context-dependent miRNAs, functioning in tumorigenesis. For example, miR-146 has been shown to be overexpressed in multiple cancers, whereas a recent study has proven that miR-146 can reduce the expression of BRCA1., Meanwhile, the expression of proteins and enzymes is also regulated by certain miRNAs. MiR-101 directly represses EZH2, and abnormal downregulation of miR-101 has been observed in cancers., The expression of the miR-29 family is inversely correlated with that of DNMT3A and -3B in lung cancer tissues. Forced expression of miR-29 inhibits tumorigenesis by inducing reexpression of methylation-silenced tumor suppressor genes. LncRNAs are another large group of noncoding RNAs that play a vital role in tumorigenesis. Some lncRNAs are cancer type-specific, such as PCGEM1 in prostate cancer and HEIH in hepatocellular carcinoma., Many aberrant lncRNAs have been discovered in various cancers. Dysregulation of HOTAIR has been found in lung, pancreatic, and colorectal cancer.–

31Therefore, ncRNAs can either be directly involved in tumorigenesis or indirectly affect tumor development by participating in other epigenetic events.

Inhibitors and clinical trials

32Unlike genetic mutations, epigenetic alterations are reversible. Given the importance of epigenetic marks in tumorigenesis, the availability of corresponding inhibitors has attracted extensive attention. Meanwhile, epigenetic regulation of a gene usually requires more than one epigenetic event. Currently, there are six epigenetic drugs approved for clinical use by the FDA (Table 4).

Targeting DNA methylation

33Blockade of DNMTs is the most effective way to prevent aberrant DNA hypermethylation. However, until now, targeting of the methyltransferase enzymes still lacks specificity and even causes hypomethylation of the global genome. Complete deletion of DNMT1 in mice results in embryonic lethality. Knockout of DNMT1 in fibroblast cells causes aberrant expression of 10% of genes and p53-dependent death. Administration of DNA methylation inhibitors results in tumorigenesis in male Fischer rats. Regulation of DNA methylation is vital in cell survival and function, and in addition to the specificity needed and the side effect associated, it is hard to identify proper drugs.

34DNA methylation inhibitors can be divided into two groups: nucleoside analogs and nonnucleoside analogs. Nucleoside analogs have a modified cytosine ring and can be turned into nucleotides and incorporated into newly synthesized DNA or RNA. DNA methyltransferases are bound by covalent complexes with the analogs, which inhibits DNA methylation. 5-Azacitidine (5-Aza-CR) and 5-aza-2′-deoxycytidine (5-Aza-CdR) are currently the two most studied and promising demethylation agents. 5-Aza-CR and zebularine are ribonucleoside analogs that can be phosphorylated to be able to incorporate into RNA. However, they can also be incorporated into DNA via the ribonucleotide reductase pathway. 5-Azacitidine, an analog of cytidine, is an injectable suspension for the treatment of myelodysplastic syndromes (MDSs). It promotes cell differentiation, demethylation, and reexpression of inactivated genes. The 5-azacitidine side effects include fetal abnormalities and decreased male fertility, especially at high doses, but its analog, 6-azacytidine, does not show such effects. Notably, after treating the noninvasive breast cancer cell lines MCF-7 and ZR-75-1 with azacytidine, the cells gained invasive abilities due to the hypomethylation of several prometastasis genes. Decitabine (5-Aza-CdR) and 5-fluoro-2′-deoxycytidine (5-F-CdR) are deoxyribonucleoside analogs that are capable of incorporating into DNA following phosphorylation. Decitabine (5-aza-2′-deoxycytidine) inhibits DNA methylation in a dosage-dependent manner. It can reactivate silenced genes at low doses but gains cytotoxicity at high doses, while myelosuppression is the major side effect at all doses.. Dihydro-5-azacytidine (DHAC) is a biologically active and chemically stable analog of 5-azacitidine with decreased toxicity., Because of its hydrolytic stability, it may be administrated via prolonged i.v. infusion, potentially eliminating the acute toxicities caused by administration of 5-azacytidine. Zebularine is a potential oral DNA-demethylating drug with stability in acidic environments and in aqueous solutions. However, the near millimolar dose requirements and the limited bioavailability in rodents (<7%) and primates (<1%) leave zebularine far from clinical translation.

35Among the drugs discussed, 5-Aza-CR and 5-Aza-CdR have already been approved by the US Food and Drug Administration (FDA) for the treatment of certain subtypes of MDS and chronic myelomonocytic leukemia. Because of their intrinsic preference for newly synthetic DNA, they tend to affect dividing cells, i.e., cancer cells. Ongoing preclinical experiments and clinical trials are exploring their efficacy in solid tumors. The common side effects of these nucleoside-like analogs are mutagenic risk and genomic instability. Nonnucleoside analogs are capable of avoiding these side effects.

36Currently, many nonnucleoside analogs have been developed to prevent DNA from aberrant hypermethylation. These drugs are usually small molecular inhibitors and directly target catalytic sites rather than incorporating into DNA. Based on a three-dimensional model of DNMT1, RG108 was designed to block the activity of this enzyme and cause demethylation. Psammaplin is a group of natural extracts from the sponge Pseudoceratina purpurea and is capable of inhibiting both DNA methyltransferases and histone deacetylases with mild cytotoxicity. Similarly, EGCG (-epigallocatechin-3-gallate) is the major polyphenol from green tea and reversibly demethylates methyl-DNA, resulting in the reactivation of multiple key genes, including hMLH1, P16, and RA, in colon, esophageal, and prostate cancer cell lines. Both hydralazine and procainamide, two drugs associated with lupus-like autoimmune diseases, can inhibit DNA methylation and induce self-reactivity in cloned T-cell lines. They have promising tumor suppressor-reactivating and antitumor actions in breast cancer., Another strategy is developing antisense oligonucleotides to inhibit DNMT transcription. MG98 is a second-generation phosphorothioate antisense oligodeoxynucleotide that prevents DNMT1 mRNA translation effects but has no obvious antitumor effect. It has been under investigation in preclinical experiments and phase I/II clinical trials, especially in solid tumors., Of note, in a systemic analysis comparing nonnucleoside inhibitors with 5-Aza-CdR, the latter showed better efficacy in DNA demethylation inhibition.

37To date, hundreds of clinical trials have investigated the effects of anti-DNA methylation therapy for various cancers (Table 5).

Inhibitors of histone modifications

38Compared with DNA methylation, histone modifications have been investigated in broader areas of diseases, including solid tumors, hematological malignancies, and even many inflammatory diseases (such as viral infection, diabetes and inflammatory lung diseases). During the process of gene silencing, lysine deacetylation and demethylation of H3K4 rather than demethylation of H3K9 or cytosine methylation might be the primary causative event. Therefore, histone modification plays an essential role in the regulation of gene expression, which also makes it a promising target for disease treatment. Clinical trials targeting histone acetylation and histone methylation are listed in Table 6 and Table 7, respectively.

Inhibitors for HATs and BETs

39Generally, there are two strategies for preventing aberrant histone acetylation, including altering interactions within the active sites within HATs or using mimetic products of enzymatic substrates. To date, many inhibitors targeting BRD proteins have been investigated in clinical trials, whereas there are no clinical trials investigating inhibitors for HATs.

40Bisubstrate inhibitors are selective inhibitors for PCAF, p300, and TIP60. They mimic two substrates of HATs: the cofactor acetyl coenzyme A (Ac-CoA) and a peptide resembling the lysine substrate., However, due to their peptidic nature and size, they are not membrane-permeable and require the assistance of a delivery system. Based on inhibitory strategies for HATs, nonpeptide small molecular inhibitors have been developing as potential therapeutic agents. Several small molecule inhibitors are natural products, including garcinol, curcumin, and anacardic acid.– These natural HAT inhibitors lack selectivity between HATs and often have other targets. Therefore, structurally modified and synthetic compounds have been reported. Α-Methylene-g-butyrolactones are small molecular inhibitors of HATs with selectivity for either GCN5L2 or PCAF. Isothiazolone is another HAT inhibitor targeting p300 and PCAF. However, high reactivity towards thiolates limits the application of HAT inhibitors in biological systems. Other inhibitors of HATs, such as thiazide sulfonamide and C646, have been gradually identified and show promising effects in multiple cancers. Another strategy to inhibit HAT activity is to target protein–protein interactions between HATs and their interaction partners. This method is dependent on the function of the interactions rather than the acetylation activity of HATs. ICG-001 and PRI-724 are representatives of this kind of inhibitor. Appropriately applying HAT agonists is also important to correct aberrant acetylation during diseases. CTPB is derived from anacardic acid and selectively activates p300, resulting in gene transcription. TTK21 and SPV106 are two other agonists based on anacardic acid.

41Binding to BRDs and blocking acetylated lysine recognition is another mechanism that inhibits acetylation. JQ1 and I-BET762 are two representative inhibitors of the BET family. JQ1 is a cell-permeable small molecule and can competitively bind to BRD4 fusion oncoproteins, such as BRD4-NUT, resulting in cancer cell differentiation and apoptosis. Similarly, I-BET762 is also a synthetic mimic of and competes with BRD4. Other compounds, such as MS417, OTX-015, RVX-208, OXFBD, I-BET151, PFI-1, MS436, and XD14, are also BET inhibitors and have been well illustrated in other published papers. We will focus on the associations between these compounds and cancers. However, a number of non-BET proteins containing BRDs have attracted considerable attention. Many non-BET bromodomain inhibitors are based on a structure called the “WPF shelf” and a “gatekeeper” residue located at the start of the C helix. Several HATs have a BRD, such as Gcn5, PCFA, p300, and CBP. Inhibitors for CBP include MS2126, MS7972, ischemin, SGC-CBP30 and I-CBP112; optimized 1-(1H-indol-1-yl) ethanone derivatives have also shown promising results in inhibiting CBP and p300. BAZ2A/B bromodomain inhibitors include BAZ2-ICR and GSK2801. The quinolone-fused lactam LP99 was the first synthetic selective inhibitor for BRD7/9. I-BRD9 was identified by GlaxoSmithKline (GSK) and is a selective inhibitor of BRD9, which has more than 200-fold selectivity for BRD9 over BRD7 and 700-fold selectivity for BRD9 over BET family members. PFI-3 is a potential inhibitor of SMARCA4 and PB1 with a stronger affinity for the bromodomain of SMARCA4. However, Vangamudi et al. identified that the ATPase domain within SMARC4 bypassed the anticancer effects related to the bromodomain since PFI-3 did not inhibit cell proliferation. The BRPF1 (bromodomain and PHD finger-containing 1) protein is part of the BRPF family, which is a component of MYST family complexes. The inhibitors of BRPF1 include PFI-4, OF-1, and NI-57. 1,3-Dimethyl benzimidazolones were the first selective inhibitors of BRPF1. PFI-4 and OF-1 are two close analogs of 1,3-dimethyl benzimidazolone that have been identified by the Structural Genomics Consortium (SGC). Another BRPF1 inhibitor, NI-57, was discovered by the SGC based on a new quinolinone scaffold. Both NI-57 and OF-1 are thought to interact with BRPF1-3 as pan-BRPF bromodomain inhibitors. Based on the bromodomain contained within both TRIM24 (tripartite motif containing protein 24) and BRPF1, a dual inhibitor, IACS-9571, has been identified. Bromosporine is a panbromodomain inhibitor with good cellular activity, whereas in a recent study, researchers noticed that bromodomain inhibitors only targeted the BET family rather than other BRDs.

Inhibition of HDACs

42Given that multiple methods can regulate HDAC activity, the designation of HDAC inhibitors has its own advantages. In the 1970s, butyrate was found to induce the accumulation of acetylated histones in cancer cells, which is thought to be associated with the inhibition of deacetylation. Later, a natural extract, trichostatin A (TSA), was identified to inhibit the activity of partially purified HDACs and induce cancer cell differentiation and apoptosis. Gradually, more natural and synthetic compounds have been identified to inhibit histone deacetylation. A study reported that administration of HDAC inhibitors only regulates a small number of genes (1–2%) but induces an obvious and rapid decrease in c-Myc gene expression, which indicated that a restricted set of cellular genes was uniquely sensitive to regulation of histone acetylation. The combination of two HDAC inhibitors, SAHA and TSA, induced melanoma cell growth arrest by upregulating p21, p27 and NF-κB, and MG132 can enhance the effect of TSA. The inhibition of HDACs has been investigated in various cancers, with promising antitumor effects., Based on the characteristics of their chemical structures, HDAC inhibitors can be divided into five groups: short-chain fatty acids, hydroxamic acids, benzamides, cyclic peptides, and hybrid molecules. In addition to those included in the five groups, some new synthetic compounds also act as inhibitors of HDACs.

43The short-chain fatty acid group contains sodium butyrate, valproic acid (VPA), sodium phenylbutyrate, and AN-9 (pivaloyloxymethyl butyrate). The effective concentration of butyrate is usually at the micromolar level. The group of hydroxamic acids includes more than ten members and is the best-studied class. Structural analyses of TSA and suberoylanilide hydroxamic acid (SAHA) show that they are noncompetitive inhibitors of HDACs since they share significant homology with class I and class II HDACs, which makes them mimics of the lysine substrates. In addition, they chelate the active zinc ion in a bidentate manner, which is crucial for enzymatic activity. Hexamethylene bisacetamide (HMBA) is a representative of the hybrid polar compounds (HPCs), whereas second-generation HPCs, such as oxamflatin, SAHA, suberic bishydroxamic acid (SBHA), and m-carboxycinnamic acid bishydroxamide (CBHA), have shown better inhibition of HDACs and anticancer effects than first-generation agents. Oxamflatin, scriptaid, and amide are analogs of TSA and show anticancer effects.– Benzamide inhibitors (MS-275, MGCD0103, and CI-994) are well-studied and show promising effects in the treatment of diseases, especially cancers. They inhibit histone deacetylation via binding to catalytic zinc ions within HDACs through carbonyl and amino groups. Inhibition of HDACs by benzamide inhibitors is thought to be reversible, but the bond may become tight and pseudoirreversible in a time-dependent manner., However, benzamide inhibitors have less activity than members of the hydroxamate or cyclic peptide families, with an effective concentration around the micromolar range. Cyclic peptides can be further divided into two groups: cyclic tetrapeptide containing a 2-amino-8-oxo-9, 10-epoxy-decanoyl (AOE) moiety (HC-toxin, trapoxin) and cyclic peptides without the AOE moiety (apicidin and romidepsin). The epoxyketone group is essential for the inhibitors to bind to active zinc ions, but the epoxyketone-based bond is irreversible. Trapoxin is a fungal cyclic peptide and can irreversibly inhibit the activity of HDACs. Romidepsin, also known as FK228, most likely relies on one of the thiol groups to coordinate to the active site zinc ion. Garlic-associated derivatives, such as diallylsulfide and allylmercaptan, are capable of generating a thiol group that makes them potential inhibitors of HDACs. K-trap, an analog of trapoxin, and other derivatives, including 9-acyloxyapicidins and 9-hydroxy, have been under investigation. Depudecin is a natural epoxide derivative isolated from the fungus Alternaria brassicicola. Psammaplins is isolated from a marine sponge Pseudoceratina purpurea. These two natural extracts can inhibit the activity of HDACs.

44Early HDAC inhibitors were nonselective because of the high homology of the structure and catalytic mechanism of HDACs within each group. The first selective HDAC inhibitor was tubacin, which targets HDAC6 with increased tubulin acetylation but not histone acetylation. PCI-34051, a specific inhibitor of HDAC8, can induce caspase-dependent apoptosis in T-cell lymphoma but does not increase histone acetylation. Another benzamide inhibitor, SHI-1:2, shows HDAC1/HDAC2-specific inhibitory activity that is >100-fold more selective than that of other HDACs. New synthetic chemicals, such as SK7041 and splitomicin, selectively target class I HDACs and sir2-like family members, respectively. The same efforts have been made to develop inhibitors for sirtuins, the class III HDACs. Nicotinamide, a byproduct of the sirtuin enzyme reaction, is a widely used inhibitor of all sirtuins. Other compounds, such as cambinol, salermide, tenovin, EX-527, suramin, and AGK2, have also been reported as sirtuin inhibitors. Sirtuin inhibitors (such as nicotinamide) function via interactions with the NAD+ within the active site of sirtuins or through binding to acetyl-lysine.

45Of note, second-generation HDACs, including hydroxamic acids (vorinostat (SAHA), belinostat (PXD101), LAQ824, and panobinostat (LBH589)) and benzamides (entinostat (MS-275), tacedinaline (CI-994), and mocetinostat (MGCD0103)), are currently in clinical trials, and some of them have already been approved for disease treatment. The success of romidepsin in phase I clinical trials in cutaneous and peripheral T-cell lymphoma accelerated the development of HDAC inhibitors as anticancer drugs. In 2006, SAHA (vorinostat) was first approved by the US Food and Drug Administration (FDA) for the treatment of cancer, restricted to patients with cutaneous T-cell lymphoma (CTCL), as an HDAC inhibitor. Romidepsin (Istodax) was the second approved HDAC inhibitor, which was approved in 2009. Three members of the benzamide family have also shown clinical significance in anticancer drug development. Belinostat (Beleodaq, previously known as PXD101) was approved in 2014 by the US FDA and European Medicines Agency to treat peripheral T-cell lymphoma. Another HDAC inhibitor, panobinostat, is a nonselective HDAC (pan-HDAC). It has shown promising effects in anticancer treatments; therefore, the FDA accelerated its approval for the treatment of patients with multiple myeloma. Intriguing, as we mentioned before, truncating mutations in HDAC2 have been found in sporadic carcinomas and colorectal cancer and result in resistance to traditional HDAC inhibitors. Mutations in other HDACs also exist; therefore, screening of these mutations in cancer can improve the efficacy of HDAC inhibitors.

Inhibitor of HMTs and HDMTs

46EPZ004777 was the first identified selective inhibitor of DOT1L and selectively kills MLL-translocated cells over those without MLL translocation. However, due to its poor pharmacokinetic properties, a second generation of EPZ004777, EPZ-5767, was developed with a cyclobutyl ring replacing the ribose moiety. EPZ-5767 also shows synergistic effects with cytarabine, daunorubicin, and the DNMT inhibitor azacitidine in treatments for ALL with MLL translocation. EPZ-5767, though still showing low oral bioavailability, has been investigated in clinical trials for the treatment of leukemia with MLL rearrangement. There are several inhibitors of EZH2. 3-Deazaneplanocin A (DZNep), a derivative of the antibiotic neplanocin-A, is one of the most studied compounds. In fact, DZNep is a SAH-hydrolase inhibitor and decreases EZH2 expression via upregulation of SAH, which leads to degradation of PRC2 in a feedback inhibition mechanism., Another kind of inhibitor is SAM competitive inhibitors. SAM is responsible for the methyl moiety of KMTs. EI1, a small molecular inhibitor of EZH2, inhibits EZH2 activity by directly binding to EZH2 and competing with SAM. GSK343 and GSK126 are two other SAM competitive inhibitors that have been investigated in clinical trials., EPZ005687, a potent inhibitor of EZH2, significantly reduces H3K27 methylation in lymphoma cells with point mutations at the Tyr641 and Ala677 residues of EZH2 without obvious effects on the proliferation of wild-type cells. EPZ-6438, which shows similar effects and superior oral bioavailability, was developed next. CPI-1205 is a novel inhibitor of EZH2 that belongs to the pyridone family.

47Tranylcypromine (TCP) is an approved drug for depression due to its ability to inhibit monoamine oxidase (MAO) activity. The structures of LSD enzymes and MAOs share many similarities. Therefore, the side effects of TCP as an HDMT inhibitor, including orthostatic hypotension, dizziness, and drowsiness, are mostly caused by targeting of MAO. Administration of TCP in MLL-AF9 leukemia promotes tumor cell differentiation and apoptosis. TCP is also capable of resensitizing non-acute promyelocytic leukemia (APL) AML cells to all-trans retinoic acid (ATRA) treatment via increasing H3K4me2 and the expression of myeloid-differentiation-associated genes. Several derivatives of TCP have been developed to achieve better bioavailability and selectivity, including OG-002, RN-1, SP2509, and GSK690.– Another LSD1 selective inhibitor, ORY-1001, can also promote the differentiation of leukemia cell lines, especially cells with translocations in MLL, and has good oral bioavailability. To date, three LSD1 inhibitors, including TCP, ORY-1001, and GSK2879552, have been under investigation in clinical trials for the treatment of cancer patients. Daminozide (N-(dimethylamino) succinamic acid, 160 Da), a plant growth regulator, selectively inhibits KDM2/7 by chelating the active site metal. Daminozide and siRNA can similarly downregulate KDM7 expression and then regulate tumor-repopulating cells via demethylation of H3K9. GSK-J1 was the first identified KDM6 inhibitor with restricted cellular permeability, which resulted from its highly polar structure. Its ethyl ester, GSK-J4, possesses an improved ability to enter cells. However, scientists have found that GSK-J1 shows compatible selectivity for the KDM6 and KDM5 families and that GSK-J4 is also a potential inhibitor for KDM5B and KDM4C. EPT-103182, a selective inhibitor of KDM5B, has shown promising results in terms of antiproliferative effects in hematological and solid cancer cells. KDM8 and JMJD6 share homology and can be inhibited by a broad spectrum inhibitor, NOG.

48Specific inhibitors usually have similar selectivity to closely related homologs within a group, and even across different groups, which needs to be taken into consideration when using compounds that are not highly selective.

Combined therapy

49Epigenetic regulation during tumorigenesis is complicated and involves multiple steps. Therefore, the combination of two or more therapies targeting various epigenetic events seems helpful. This combination synergistically inhibits the expression of tumor-growth-promoting genes and promotes the reexpression of tumor suppressor genes. 4SC-202 is a small molecular drug with dual effects that can inhibit HDAC1/2/3 and LSD1 with similar low micromolar potency. This drug is under clinical investigation. Other studies have administered two or more kinds of epigenetic drugs for anticancer therapy. Relevant clinical trials are listed in Table 8.

Conclusion

50Although more specific mechanisms need to be investigated, it is well accepted that epigenetic events are important in normal biological processes as well as in tumorigenesis and that the epigenetic status is usually widely altered during cancer initiation. This makes epigenome-targeted therapy a promising strategy for the treatment of cancer. Based on the complexity of cancer, epigenetic alterations have influenced multiple aspects in cancer, such as the expression of oncogenes and tumor suppressor genes and signal transduction, resulting in enhanced cancer growth, invasion and metastasis. Although epigenetic therapy has a rational and profound basis in theory, some problems remain to be discussed and solved. The first and most important is the problem of selectivity. Epigenetic events are ubiquitously distributed across normal and cancer cells. In fact, some cancers depend on certain epigenetic alterations and can be sensitive to this regulation, whereas under usual regulation, normal cells have the ability to compensate for these epigenetic changes. Therefore, the priority is to determine the most important epigenetic alterations for different cancers. The second problem extends from the first problem. Thus far, epigenetic therapy has obtained impressive results in hematological malignancies but not in solid tumors. The properties of hematological malignant cells and solid tumor cells are different. However, researchers have still investigated the appropriate strategies for solid tumors. Since epigenetic alterations have effects on the sensitivity of small molecule targeted therapy and chemotherapy or radiotherapy, epigenetic-targeted therapy seems to be an important adjunctive therapy. The combination of epigenetic therapy and immunotherapy has also been investigated in preclinical and clinical trials.

51Based on the achievements obtained, epigenetic-targeted therapy is a promising strategy for anticancer treatment. Epigenomes in cancer are related to many aspects during cancer initiation. A better understanding of the specific mechanisms underlying those alterations in different cancers is necessary. Meanwhile, optimized treatment options, including a variety of combinations, still remain to be discovered.

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106.
Zhou Y, et al. Ginsenoside 20(S)-Rg3 inhibits the warburg effect via modulating DNMT3A/ MiR-532-3p/HK2 pathway in ovarian cancer cells. Cell Physiol. Biochem. 2018;45:2548-2559.
Not in our library10.1159/000488273
107.
Zhang Q, Feng Y, Liu P, Yang J. MiR-143 inhibits cell proliferation and invasion by targeting DNMT3A in gastric cancer. Tumour Biol. 2017;39:1010428317711312.
Not in our library10.1177/1010428317711312
108.
Gao Q, et al. Deletion of the de novo DNA methyltransferase Dnmt3a promotes lung tumor progression. Proc. Natl Acad. Sci. USA. 2011;108:18061-18066.
Not in our library10.1073/pnas.1114946108
109.
Garzon R, et al. MicroRNA-29b induces global DNA hypomethylation and tumor suppressor gene reexpression in acute myeloid leukemia by targeting directly DNMT3A and 3B and indirectly DNMT1. Blood. 2009;113:6411-6418.
Not in our library10.1182/blood-2008-07-170589
110.
Deivendran S, et al. Metastasis-associated protein 1 is an upstream regulator of DNMT3a and stimulator of insulin-growth factor binding protein-3 in breast cancer. Sci. Rep. 2017;7:44225.
Not in our library10.1038/srep44225
111.
Tang X, et al. Autocrine TGF-β1/miR-200s/miR-221/DNMT3B regulatory loop maintains CAF status to fuel breast cancer cell proliferation. Cancer Lett. 2019;452:79-89.
Not in our library10.1016/j.canlet.2019.02.044
112.
Chen Z, et al. miR-124 and miR-506 inhibit colorectal cancer progression by targeting DNMT3B and DNMT1. Oncotarget. 2015;6:38139-38150.
Not in our library10.18632/oncotarget.5709
113.
Kaempferol Modulates DNA Methylation and Downregulates DNMT3B in Bladder Cancer · 2017
Not checked
114.
Peres R, et al. Angiogenin contributes to bladder cancer tumorigenesis by DNMT3b-mediated MMP2 activation. Oncotarget. 2016;7:43109-43123.
Not in our library10.18632/oncotarget.10097
115.
Song N, et al. Lentivirus‑mediated knockdown of MeCP2 inhibits the growth of colorectal cancer cells in vitro. Mol. Med Rep. 2016;13:860-866.
Not in our library10.3892/mmr.2015.4612
116.
Ray BK, et al. Epigenetic regulation by Z-DNA silencer function controls cancer-associated ADAM-12 expression in breast cancer: cross-talk between MeCP2 and NF1 transcription factor family. Cancer Res. 2013;73:736-744.
Not in our library10.1158/0008-5472.can-12-2601
117.
Xu M, et al. MeCP2 suppresses LIN28A expression via binding to its methylated-CpG islands in pancreatic cancer cells. Oncotarget. 2016;7:14476-14485.
Not in our library10.18632/oncotarget.7507
118.
Yaqinuddin A, et al. Silencing of MBD1 and MeCP2 in prostate-cancer-derived PC3 cells produces differential gene expression profiles and cellular phenotypes. Biosci. Rep. 2008;28:319-326.
Not in our library10.1042/bsr20080032
119.
Zhang B, et al. MBD1 is an epigenetic regulator of KEAP1 in pancreatic cancer. Curr. Mol. Med. 2016;16:404-411.
Not in our library10.2174/1566524016666160316154150
120.
Pulukuri SM, Rao JS. CpG island promoter methylation and silencing of 14-3-3σ gene expression in LNCaP and Tramp-C1 prostate cancer cell lines is associated with methyl-CpG-binding protein MBD2. Oncogene. 2006;25:4559-4572.
Not in our library10.1038/sj.onc.1209462
121.
Martin V, et al. MBD2-mediated transcriptional repression of the p14ARF tumor suppressor gene in human colon cancer cells. Pathobiology. 2008;75:281-287.
Not in our library10.1159/000151708
122.
Pontes TB, et al. Reduced mRNA expression levels of MBD2 and MBD3 in gastric carcinogenesis. Tumour Biol. 2014;35:3447-3453.
Not in our library10.1007/s13277-013-1455-y
123.
Zhang Y, et al. Epigenetic silencing of RNF144A expression in breast cancer cells through promoter hypermethylation and MBD4. Cancer Med. 2018;7:1317-1325.
Not in our library10.1002/cam4.1324
124.
Abdel-Rahman WM, et al. Truncation of MBD4 predisposes to reciprocal chromosomal translocations and alters the response to therapeutic agents in colon cancer cells. DNA Repair. 2008;7:321-328.
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125.
Pierre CC, Hercules SM, Yates C, Daniel JM. Dancing from bottoms up—roles of the POZ-ZF transcription factor Kaiso in cancer. Biochim Biophys. Acta Rev. Cancer. 2019;1871:64-74.
Not in our library10.1016/j.bbcan.2018.10.005
126.
Wang H, et al. Kaiso, a transcriptional repressor, promotes cell migration and invasion of prostate cancer cells through regulation of miR-31 expression. Oncotarget. 2016;7:5677-5689.
Not in our library10.18632/oncotarget.6801
127.
Dai SD, et al. Kaiso is expressed in lung cancer: its expression and localization is affected by p120ctn. Lung Cancer. 2010;67:205-215.
Not in our library10.1016/j.lungcan.2009.06.013
128.
Bassey-Archibong BI, et al. Kaiso depletion attenuates the growth and survival of triple negative breast cancer cells. Cell Death Dis. 2017;8:e2689.
Not in our library10.1038/cddis.2017.92
129.
Roussel-Gervais A, et al. Loss of the methyl-CpG-binding protein ZBTB4 alters mitotic checkpoint, increases aneuploidy, and promotes tumorigenesis. Cancer Res. 2017;77:62-73.
Not in our library10.1158/0008-5472.can-16-1181
130.
Yu Y, et al. Tumor suppressive ZBTB4 inhibits cell growth by regulating cell cycle progression and apoptosis in Ewing sarcoma. Biomed. Pharmacother. 2018;100:108-115.
Not in our library10.1016/j.biopha.2018.01.132
131.
Kim K, et al. Induction of the transcriptional repressor ZBTB4 in prostate cancer cells by drug-induced targeting of microRNA-17-92/106b-25 clusters. Mol. Cancer Ther. 2012;11:1852-1862.
Not in our library10.1158/1535-7163.mct-12-0181
132.
Jing J, et al. The role of ZBTB38 in promoting migration and invasive growth of bladder cancer cells. Oncol. Rep. 2019;41:1980-1990.
Not in our library10.3892/or.2018.6937
133.
Saidi S, Popov Z, Janevska V, Panov S. Overexpression of UHRF1 gene correlates with the major clinicopathological parameters in urinary bladder cancer. Int Braz. J. Urol. 2017;43:224-229.
Not in our library10.1590/s1677-5538.ibju.2016.0126
134.
Jiao D, et al. UHRF1 promotes renal cell carcinoma progression through epigenetic regulation of TXNIP. Oncogene. 2019;38:5686-5699.
Not in our library10.1038/s41388-019-0822-6
135.
Kong X, et al. Defining UHRF1 domains that support maintenance of human colon cancer DNA methylation and oncogenic properties. Cancer Cell. 2019;35:633-648.e7.
Not in our library10.1016/j.ccell.2019.03.003
136.
Hu Q, et al. UHRF1 promotes aerobic glycolysis and proliferation via suppression of SIRT4 in pancreatic cancer. Cancer Lett. 2019;452:226-236.
Not in our library10.1016/j.canlet.2019.03.024
137.
Wei C, et al. Upregulation of UHRF1 promotes the progression of melanoma by inducing cell proliferation. Oncol. Rep. 2018;39:2553-2562.
Not in our library10.3892/or.2018.6356
138.
Gao SP, et al. UHRF1 promotes breast cancer progression by suppressing KLF17 expression by hypermethylating its promoter. Am. J. Cancer Res. 2017;7:1554-1565.
Not in our library
139.
Lu S, et al. Ubiquitin-like with PHD and ring finger domains 2 is a predictor of survival and a potential therapeutic target in colon cancer. Oncol. Rep. 2014;31:1802-1810.
Not in our library10.3892/or.2014.3035
140.
Zhang T, et al. UHRF2 decreases H3K9ac expression by interacting with it through the PHD and SRA/YDG domain in HepG2 hepatocellular carcinoma cells. Int J. Mol. Med. 2017;39:126-134.
Not in our library10.3892/ijmm.2016.2805
141.
Iguchi T, et al. Identification of UHRF2 as a negative regulator of epithelial-mesenchymal transition and its clinical significance in esophageal squamous cell carcinoma. Oncology. 2018;95:179-187.
Not in our library10.1159/000488860
142.
Jin C, et al. Loss of UHRF2 is associated with non-small cell lung carcinoma progression. J. Cancer. 2018;9:2994-3005.
Not in our library10.7150/jca.25876
143.
Si Y, et al. Fisetin decreases TET1 activity and CCNY/CDK16 promoter 5hmC levels to inhibit the proliferation and invasion of renal cancer stem cell. J. Cell Mol. Med. 2019;23:1095-1105.
Not in our library10.1111/jcmm.14010
144.
Huang H, et al. TET1 plays an essential oncogenic role in MLL-rearranged leukemia. Proc. Natl Acad. Sci. USA. 2013;110:11994-11999.
Not in our library10.1073/pnas.1310656110
145.
Jiang X, et al. Targeted inhibition of STAT/TET1 axis as a therapeutic strategy for acute myeloid leukemia. Nat. Commun. 2017;8:2099.
Not in our library10.1038/s41467-017-02290-w
146.
Chen LY, et al. TET1 reprograms the epithelial ovarian cancer epigenome and reveals casein kinase 2α as a therapeutic target. J. Pathol. 2019;248:363-376.
Not in our library10.1002/path.5238
147.
Filipczak PT, et al. p53-Suppressed oncogene TET1 prevents cellular aging in lung cancer. Cancer Res. 2019;79:1758-1768.
Not in our library10.1158/0008-5472.can-18-1234
148.
Decrease of 5-Hydroxymethylcytosine Is Associated with Progression of Hepatocellular Carcinoma through Downregulation of TET1 · 2013
No notice
149.
Guo H, et al. TET1 suppresses colon cancer proliferation by impairing á-catenin signal pathway. J. Cell Biochem. 2019;120:12559-12565.
Not in our library10.1002/jcb.28522
150.
Cimmino L, et al. Restoration of TET2 function blocks aberrant self-renewal and leukemia progression. Cell. 2017;170:1079-1095.e20.
Not in our library10.1016/j.cell.2017.07.032
151.
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Not in our library10.1038/ng.391
152.
Zhu X, Li S. TET2 inhibits tumorigenesis of breast cancer cells by regulating caspase-4. Sci. Rep. 2018;8:16167.
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153.
Huang Y, et al. Loss of nuclear localization of TET2 in colorectal cancer. Clin. Epigenetics. 2016;8:9.
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154.
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155.
Yang L, et al. Reduced expression of TET1, TET2, TET3 and TDG mRNAs are associated with poor prognosis of patients with early breast cancer. PLoS ONE. 2015;10:e0133896.
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156.
Ye Z, et al. TET3 inhibits TGF-β1-induced epithelial-mesenchymal transition by demethylating miR-30d precursor gene in ovarian cancer cells. J. Exp. Clin. Cancer Res. 2016;35:72.
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157.
Kouzarides T. Chromatin modifications and their function. Cell. 2007;128:693-705.
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158.
Audia JE, Campbell RM. Histone modifications and cancer. Cold Spring Harb. Perspect. Biol. 2016;8:a019521.
Not in our library10.1101/cshperspect.a019521
159.
Hawkins RD, et al. Dynamic chromatin states in human ES cells reveal potential regulatory sequences and genes involved in pluripotency. Cell Res. 2011;21:1393-1409.
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160.
Hon GC, Hawkins RD, Ren B. Predictive chromatin signatures in the mammalian genome. Hum. Mol. Genet. 2009;18:R195-R201.
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163.
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165.
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166.
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167.
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168.
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169.
Marmorstein R, Zhou MM. Writers and readers of histone acetylation: structure, mechanism, and inhibition. Cold Spring Harb. Perspect. Biol. 2014;6:a018762.
Not in our library10.1101/cshperspect.a018762
170.
Heery DM, Fischer PM. Pharmacological targeting of lysine acetyltransferases in human disease: a progress report. Drug Disco. Today. 2007;12:88-99.
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171.
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172.
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173.
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174.
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176.
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178.
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179.
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180.
Miller RW, Rubinstein JH. Tumors in Rubinstein-Taybi syndrome. Am. J. Med. Genet. 1995;56:112-115.
Not in our library10.1002/ajmg.1320560125
181.
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182.
Panagopoulos I, et al. Fusion of the MORF and CBP genes in acute myeloid leukemia with the t(10;16)(q22; p13). Hum. Mol. Genet. 2001;10:395-404.
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183.
Chaffanet M, et al. MOZ is fused to p300 in an acute monocytic leukemia with t(8;22). Genes Chromosomes Cancer. 2000;28:138-144.
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184.
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185.
Taki T, Sako M, Tsuchida M, Hayashi Y. The t(11;16)(q23;p13) translocation in myelodysplastic syndrome fuses the MLL gene to the CBP gene. Blood. 1997;89:3945-3950.
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186.
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Not in our library
187.
Gayther SA, et al. Mutations truncating the EP300 acetylase in human cancers. Nat. Genet. 2000;24:300-303.
Not in our library10.1038/73536
188.
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189.
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190.
Wachtel M, et al. Gene expression signatures identify rhabdomyosarcoma subtypes and detect a novel t(2;2)(q35; p23) translocation fusing PAX3 to NCOA1. Cancer Res. 2004;64:5539-5545.
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191.
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192.
Thompson M. Polybromo-1: the chromatin targeting subunit of the PBAF complex. Biochimie. 2009;91:309-319.
Not in our library10.1016/j.biochi.2008.10.019
193.
Zhang Q, et al. Biochemical profiling of histone binding selectivity of the yeast bromodomain family. PLoS ONE. 2010;5:e8903.
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194.
Sanchez R, Zhou MM. The role of human bromodomains in chromatin biology and gene transcription. Curr. Opin. Drug Disco. Devel. 2009;12:659-665.
Not in our library
195.
Filippakopoulos P, Knapp S. Targeting bromodomains: epigenetic readers of lysine acetylation. Nat. Rev. Drug Discov. 2014;13:337-356.
Not in our library10.1038/nrd4286
196.
Li Y, et al. AF9 YEATS domain links histone acetylation to DOT1L-mediated H3K79 methylation. Cell. 2014;159:558-571.
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197.
Morin RD, et al. Genetic landscapes of relapsed and refractory diffuse large B-cell lymphomas. Clin. Cancer Res. 2016;22:2290-2300.
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198.
Perlman EJ, et al. MLLT1 YEATS domain mutations in clinically distinctive Favourable Histology Wilms tumours. Nat. Commun. 2015;6:10013.
Not in our library10.1038/ncomms10013
199.
Chiang CM. Brd4 engagement from chromatin targeting to transcriptional regulation: selective contact with acetylated histone H3 and H4. F1000 Biol. Rep. 2009;1:98.
Not in our library10.3410/b1-98
200.
Zeng L, Zhou MM. Bromodomain: an acetyl-lysine binding domain. FEBS Lett. 2002;513:124-128.
Not in our library10.1016/s0014-5793(01)03309-9
201.
Pirrotta V. The necessity of chromatin: a view in perspective. Cold Spring Harb. Perspect. Biol. 2016;8:a019547.
Not in our library10.1101/cshperspect.a019547
202.
Yang Z, et al. Recruitment of P-TEFb for stimulation of transcriptional elongation by the bromodomain protein Brd4. Mol. Cell. 2005;19:535-545.
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203.
Phelps MA, et al. Clinical response and pharmacokinetics from a phase 1 study of an active dosing schedule of flavopiridol in relapsed chronic lymphocytic leukemia. Blood. 2009;113:2637-2645.
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204.
Rahl PB, et al. c-Myc regulates transcriptional pause release. Cell. 2010;141:432-445.
Not in our library10.1016/j.cell.2010.03.030
205.
French CA, et al. BRD-NUT oncoproteins: a family of closely related nuclear proteins that block epithelial differentiation and maintain the growth of carcinoma cells. Oncogene. 2008;27:2237-2242.
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206.
Zuber J, et al. RNAi screen identifies Brd4 as a therapeutic target in acute myeloid leukaemia. Nature. 2011;478:524-528.
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207.
de Ruijter AJ, et al. Histone deacetylases (HDACs): characterization of the classical HDAC family. Biochem J. 2003;370:737-749.
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208.
Verdin E, Dequiedt F, Kasler HG. Class II histone deacetylases: versatile regulators. Trends Genet. 2003;19:286-293.
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209.
Seto E, Yoshida M. Erasers of histone acetylation: the histone deacetylase enzymes. Cold Spring Harb. Perspect. Biol. 2014;6:a018713.
Not in our library10.1101/cshperspect.a018713
210.
Imai S, Armstrong CM, Kaeberlein M, Guarente L. Transcriptional silencing and longevity protein Sir2 is an NAD-dependent histone deacetylase. Nature. 2000;403:795-800.
Not in our library10.1038/35001622
211.
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212.
Frye RA. Characterization of five human cDNAs with homology to the yeast SIR2 gene: Sir2-like proteins (sirtuins) metabolize NAD and may have protein ADP-ribosyltransferase activity. Biochem Biophys. Res. Commun. 1999;260:273-279.
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213.
Du J, et al. Sirt5 is a NAD-dependent protein lysine demalonylase and desuccinylase. Science. 2011;334:806-809.
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214.
Johnson CA, et al. Human class I histone deacetylase complexes show enhanced catalytic activity in the presence of ATP and co-immunoprecipitate with the ATP-dependent chaperone protein Hsp70. J. Biol. Chem. 2002;277:9590-9597.
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215.
Zhang X, et al. Activation of the growth-differentiation factor 11 gene by the histone deacetylase (HDAC) inhibitor trichostatin A and repression by HDAC3. Mol. Cell Biol. 2004;24:5106-5118.
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216.
Hubbert C, et al. HDAC6 is a microtubule-associated deacetylase. Nature. 2002;417:455-458.
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217.
Luo J, et al. Deacetylation of p53 modulates its effect on cell growth and apoptosis. Nature. 2000;408:377-381.
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218.
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219.
Peng L, et al. SIRT1 negatively regulates the activities, functions, and protein levels of hMOF and TIP60. Mol. Cell Biol. 2012;32:2823-2836.
Not in our library10.1128/mcb.00496-12
220.
Ayer DE. Histone deacetylases: transcriptional repression with SINers and NuRDs. Trends Cell Biol. 1999;9:193-198.
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221.
The histone deacetylase-3 complex contains nuclear receptor corepressors · 2000
Not checked
222.
Watson PJ, Fairall L, Santos GM, Schwabe JW. Structure of HDAC3 bound to co-repressor and inositol tetraphosphate. Nature. 2012;481:335-340.
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223.
Kang H, et al. Peptide switch is essential for Sirt1 deacetylase activity. Mol. Cell. 2011;44:203-213.
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224.
Kim EJ, Kho JH, Kang MR, Um SJ. Active regulator of SIRT1 cooperates with SIRT1 and facilitates suppression of p53 activity. Mol. Cell. 2007;28:277-290.
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225.
Ocker M, Schneider-Stock R. Histone deacetylase inhibitors: signalling towards p21cip1/waf1. Int J. Biochem Cell Biol. 2007;39:1367-1374.
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226.
Differential expression of selected histone modifier genes in human solid cancers · 2006
No notice
227.
Xue L, et al. RNAi screening identifies HAT1 as a potential drug target in esophageal squamous cell carcinoma. Int J. Clin. Exp. Pathol. 2014;7:3898-3907.
Not in our library
228.
Fan P, et al. Overexpressed histone acetyltransferase 1 regulates cancer immunity by increasing programmed death-ligand 1 expression in pancreatic cancer. J. Exp. Clin. Cancer Res. 2019;38:47.
Not in our library10.1186/s13046-019-1044-z
229.
Miao BP, et al. Histone acetyltransferase 1 up regulates Bcl2L12 expression in nasopharyngeal cancer cells. Arch. Biochem Biophys. 2018;646:72-79.
Not in our library10.1016/j.abb.2018.03.040
230.
Jin X, Tian S, Li P. Histone acetyltransferase 1 promotes cell proliferation and induces cisplatin resistance in hepatocellular carcinoma. Oncol. Res. 2017;25:939-946.
Not in our library10.3727/096504016x14809827856524
231.
Zhang J, Liu M, Liu W, Wang W. Ras-ERK1/2 signalling promotes the development of osteosarcoma through regulation of H4K12ac through HAT1. Artif. Cells Nanomed. Biotechnol. 2019;47:1207-1215.
Not in our library10.1080/21691401.2019.1593857
232.
Han N, et al. HAT1 induces lung cancer cell apoptosis via up regulating Fas. Oncotarget. 2017;8:89970-89977.
Not in our library10.18632/oncotarget.21205
233.
Yin YW, et al. The histone acetyltransferase GCN5 expression is elevated and regulated by c-Myc and E2F1 transcription factors in human colon cancer. Gene Expr. 2015;16:187-196.
Not in our library10.3727/105221615x14399878166230
234.
Zhao L, Pang A, Li Y. Function of GCN5 in the TGF-β1-induced epithelial-to-mesenchymal transition in breast cancer. Oncol. Lett. 2018;16:3955-3963.
Not in our library10.3892/ol.2018.9134
235.
Zhao C, et al. C5a induces A549 cell proliferation of non-small cell lung cancer via GDF15 gene activation mediated by GCN5-dependent KLF5 acetylation. Oncogene. 2018;37:4821-4837.
Not in our library10.1038/s41388-018-0298-9
236.
Watts GS, et al. The acetyltransferase p300/CBP-associated factor is a p53 target gene in breast tumor cells. Neoplasia. 2004;6:187-194.
Not in our library10.1593/neo.03292
237.
Brasacchio D, et al. Down-regulation of a pro-apoptotic pathway regulated by PCAF/ADA3 in early stage gastric cancer. Cell Death Dis. 2018;9:442.
Not in our library10.1038/s41419-018-0470-8
238.
Yu C, et al. Phenethyl isothiocyanate inhibits androgen receptor-regulated transcriptional activity in prostate cancer cells through suppressing PCAF. Mol. Nutr. Food Res. 2013;57:1825-1833.
Not in our library10.1002/mnfr.201200642
239.
Xie X, et al. Tip60 is associated with resistance to X-ray irradiation in prostate cancer. FEBS Open Bio. 2018;8:271-278.
Not in our library10.1002/2211-5463.12371
240.
Wang LT, et al. TIP60-dependent acetylation of the SPZ1-TWIST complex promotes epithelial-mesenchymal transition and metastasis in liver cancer. Oncogene. 2019;38:518-532.
Not in our library10.1038/s41388-018-0457-z
241.
Sakuraba K, et al. Down-regulation of Tip60 gene as a potential marker for the malignancy of colorectal cancer. Anticancer Res. 2009;29:3953-3955.
Not in our library
242.
McGuire A, et al. Quantifying Tip60 (Kat5) stratifies breast cancer. Sci. Rep. 2019;9:3819.
Not in our library10.1038/s41598-019-40221-5
243.
Yang Y, et al. Tat-interactive protein-60KDA (TIP60) regulates the tumorigenesis of lung cancer in vitro. J. Cancer. 2017;8:2277-2281.
Not in our library10.7150/jca.19677
244.
Jaganathan A, et al. Coactivator MYST1 regulates nuclear factor-κB and androgen receptor functions during proliferation of prostate cancer cells. Mol. Endocrinol. 2014;28:872-885.
Not in our library10.1210/me.2014-1055
245.
Hayashi Y, et al. NUP98-HBO1-fusion generates phenotypically and genetically relevant chronic myelomonocytic leukemia pathogenesis. Blood Adv. 2019;3:1047-1060.
Not in our library10.1182/bloodadvances.2018025007
246.
Quintela M, et al. HBO1 directs histone H4 specific acetylation, potentiating mechano-transduction pathways and membrane elasticity in ovarian cancer cells. Nanomedicine. 2019;17:254-265.
Not in our library10.1016/j.nano.2019.01.017
247.
Chen Z, et al. HBO1 promotes cell proliferation in bladder cancer via activation of Wnt/β-catenin signaling. Mol. Carcinog. 2018;57:12-21.
Not in our library10.1002/mc.22715
248.
Deguchi K, et al. MOZ-TIF2-induced acute myeloid leukemia requires the MOZ nucleosome binding motif and TIF2-mediated recruitment of CBP. Cancer Cell. 2003;3:259-271.
Not in our library10.1016/s1535-6108(03)00051-5
249.
Mohammadi K, et al. Investigation of the changes in the expression levels of MOZ gene in colorectal cancer tissues. J. Gastrointest. Oncol. 2019;10:68-73.
Not in our library10.21037/jgo.2018.09.12
250.
Yu L, et al. Identification of MYST3 as a novel epigenetic activator of ERα frequently amplified in breast cancer. Oncogene. 2017;36:2910-2918.
Not in our library10.1038/onc.2016.433
251.
Vizmanos JL, et al. t(10;16)(q22; p13) and MORF-CREBBP fusion is a recurrent event in acute myeloid leukemia. Genes Chromosomes Cancer. 2003;36:402-405.
Not in our library10.1002/gcc.10174
252.
Ono H, Basson MD, Ito H. P300 inhibition enhances gemcitabine-induced apoptosis of pancreatic cancer. Oncotarget. 2016;7:51301-51310.
Not in our library10.18632/oncotarget.10117
253.
Pattabiraman DR, et al. Interaction of c-Myb with p300 is required for the induction of acute myeloid leukemia (AML) by human AML oncogenes. Blood. 2014;123:2682-2690.
Not in our library10.1182/blood-2012-02-413187
254.
Hou X, et al. p300 promotes proliferation, migration, and invasion via inducing epithelial-mesenchymal transition in non-small cell lung cancer cells. BMC Cancer. 2018;18:641.
Not in our library10.1186/s12885-018-4559-3
255.
Xu B, et al. The LIM protein Ajuba recruits DBC1 and CBP/p300 to acetylate ERα and enhances ERα target gene expression in breast cancer cells. Nucleic Acids Res. 2019;47:2322-2335.
Not in our library10.1093/nar/gky1306
256.
Wang YM, et al. Histone acetyltransferase p300/CBP inhibitor C646 blocks the survival and invasion pathways of gastric cancer cell lines. Int J. Oncol. 2017;51:1860-1868.
Not in our library10.3892/ijo.2017.4176
257.
He H, et al. Selective p300 inhibitor C646 inhibited HPV E6-E7 genes, altered glucose metabolism and induced apoptosis in cervical cancer cells. Eur. J. Pharmacol. 2017;812:206-215.
Not in our library10.1016/j.ejphar.2017.06.005
258.
Huh JW, et al. Prognostic impact of p300 expression in patients with colorectal canceu. J. Surg. Oncol. 2013;108:374-377.
Not in our library10.1002/jso.23405
259.
Takeuchi A, et al. p300 mediates cellular resistance to doxorubicin in bladder cancer. Mol. Med Rep. 2012;5:173-176.
Not in our library10.3892/mmr.2011.593
260.
Inagaki Y, et al. Epigenetic regulation of proliferation and invasion in hepatocellular carcinoma cells by CBP/p300 histone acetyltransferase activity. Int J. Oncol. 2016;48:533-540.
Not in our library10.3892/ijo.2015.3288
261.
Díaz-Beyá M, et al. Acute myeloid leukemia with translocation (8;16)(p11; p13) and MYST3-CREBBP rearrangement harbors a distinctive microRNA signature targeting RET proto-oncogene. Leukemia. 2013;27:595-603.
Not in our library10.1038/leu.2012.278
262.
Jin L, et al. Therapeutic targeting of the CBP/p300 bromodomain blocks the growth of castration-resistant prostate cancer. Cancer Res. 2017;77:5564-5575.
Not in our library10.1158/0008-5472.can-17-0314
263.
Ding L, et al. CBP loss cooperates with PTEN haploinsufficiency to drive prostate cancer: implications for epigenetic therapy. Cancer Res. 2014;74:2050-2061.
Not in our library10.1158/0008-5472.can-13-1659
264.
Jia D, et al. Crebbp loss drives small cell lung cancer and increases sensitivity to HDAC inhibition. Cancer Discov. 2018;8:1422-1437.
Not in our library10.1158/2159-8290.cd-18-0385
265.
Pavón MA, et al. CKMT1 and NCOA1 expression as a predictor of clinical outcome in patients with advanced-stage head and neck squamous cell carcinoma. Head. Neck. 2016;38(Suppl 1):E1392-E1403.
Not in our library10.1002/hed.24232
266.
Qin L, et al. NCOA1 directly targets M-CSF1 expression to promote breast cancer metastasis. Cancer Res. 2014;74:3477-3488.
Not in our library10.1158/0008-5472.can-13-2639
267.
Ma YS, et al. High expression of miR-105–1 positively correlates with clinical prognosis of hepatocellular carcinoma by targeting oncogene NCOA1. Oncotarget. 2017;8:11896-11905.
Not in our library10.18632/oncotarget.14435
268.
Aikawa Y, et al. PU.1-mediated upregulation of CSF1R is crucial for leukemia stem cell potential induced by MOZ-TIF2. Nat. Med. 2010;16:580-585.
Not in our library10.1038/nm.2122
269.
Fancher AT, et al. Reconfiguring the AR-TIF2 protein-protein interaction HCS Assay in Prostate Cancer Cells and Characterizing the Hits from a LOPAC Screen. Assay. Drug Dev. Technol. 2016;14:453-477.
Not in our library10.1089/adt.2016.741
270.
O’Donnell KA, et al. A Sleeping Beauty mutagenesis screen reveals a tumor suppressor role for Ncoa2/Src-2 in liver cancer. Proc. Natl Acad. Sci. USA. 2012;109:E1377-E1386.
Not in our library10.1073/pnas.1115433109
271.
Yu J, et al. Disruption of NCOA2 by recurrent fusion with LACTB2 in colorectal cancer. Oncogene. 2016;35:187-195.
Not in our library10.1038/onc.2015.72
272.
Kumar S, et al. NCOA3-mediated upregulation of mucin expression via transcriptional and post-translational changes during the development of pancreatic cancer. Oncogene. 2015;34:4879-4889.
Not in our library10.1038/onc.2014.409
273.
Alkner S, et al. Prognostic and predictive importance of the estrogen receptor coactivator AIB1 in a randomized trial comparing adjuvant letrozole and tamoxifen therapy in postmenopausal breast cancer: the Danish cohort of BIG 1–98. Breast Cancer Res. Treat. 2017;166:481-490.
Not in our library10.1007/s10549-017-4416-0
274.
Zou JX, et al. ACTR/AIB1/SRC-3 and androgen receptor control prostate cancer cell proliferation and tumor growth through direct control of cell cycle genes. Prostate. 2006;66:1474-1486.
Not in our library10.1002/pros.20477
275.
Xie D, et al. Correlation of AIB1 overexpression with advanced clinical stage of human colorectal carcinoma. Hum. Pathol. 2005;36:777-783.
Not in our library10.1016/j.humpath.2005.05.007
276.
Liu WH, Chang LS. Arachidonic acid induces Fas and FasL upregulation in human leukemia U937 cells via Ca2+/ROS-mediated suppression of ERK/c-Fos pathway and activation of p38 MAPK/ATF-2 pathway. Toxicol. Lett. 2009;191:140-148.
Not in our library10.1016/j.toxlet.2009.08.016
277.
Desai S, Laskar S, Pandey BN. Autocrine IL-8 and VEGF mediate epithelial-mesenchymal transition and invasiveness via p38/JNK-ATF-2 signalling in A549 lung cancer cells. Cell Signal. 2013;25:1780-1791.
Not in our library10.1016/j.cellsig.2013.05.025
278.
Sun X, Lou L, Zhong K, Wan L. MicroRNA-451 regulates chemoresistance in renal cell carcinoma by targeting ATF-2 gene. Exp. Biol. Med. 2017;242:1299-1305.
Not in our library10.1177/1535370217701625
279.
RNA polymerase III transcription factor TFIIIC2 is overexpressed in ovarian tumors · 2000
Not checked
280.
Xiao L, et al. Induction of the CLOCK gene by E2-ERα signaling promotes the proliferation of breast cancer cells. PLoS ONE. 2014;9:e95878.
Not in our library10.1371/journal.pone.0095878
281.
Xu H, Wang Z, Mo G, Chen H. Association between circadian gene CLOCK and cisplatin resistance in ovarian cancer cells: a preliminary study. Oncol. Lett. 2018;15:8945-8950.
Not in our library10.3892/ol.2018.8488
282.
Xiang R, et al. Circadian clock gene Per2 downregulation in non‑small cell lung cancer is associated with tumour progression and metastasis. Oncol. Rep. 2018;40:3040-3048.
Not in our library10.3892/or.2018.6704
283.
Xie XW, et al. Expression of CIITA-related MHCII molecules in tumors linked to prognosis in hepatocellular carcinoma. Int J. Oncol. 2009;34:681-688.
Not in our library10.3892/ijo_00000238
284.
Truax AD, Thakkar M, Greer SF. Dysregulated recruitment of the histone methyltransferase EZH2 to the class II transactivator (CIITA) promoter IV in breast cancer cells. PLoS ONE. 2012;7:e36013.
Not in our library10.1371/journal.pone.0036013
285.
Satoh A, et al. Epigenetic inactivation of class II transactivator (CIITA) is associated with the absence of interferon-gamma-induced HLA-DR expression in colorectal and gastric cancer cells. Oncogene. 2004;23:8876-8886.
Not in our library10.1038/sj.onc.1208144
286.
Starska K, et al. Gene and protein expression of O-GlcNAc-cycling enzymes in human laryngeal cancer. Clin. Exp. Med. 2015;15:455-468.
Not in our library10.1007/s10238-014-0318-1
287.
Krześlak A, et al. Gene expression of O-GlcNAc cycling enzymes in human breast cancers. Clin. Exp. Med. 2012;12:61-65.
Not in our library10.1007/s10238-011-0138-5
288.
Fontanals-Cirera B, et al. Harnessing BET inhibitor sensitivity reveals AMIGO2 as a melanoma survival gene. Mol. Cell. 2017;68:731-744.e9.
Not in our library10.1016/j.molcel.2017.11.004
289.
Pérez-Pena J, et al. Mapping Bromodomains in breast cancer and association with clinical outcome. Sci. Rep. 2019;9:5734.
Not in our library10.1038/s41598-019-41934-3
290.
Handoko L, et al. JQ1 affects BRD2-dependent and independent transcription regulation without disrupting H4-hyperacetylated chromatin states. Epigenetics. 2018;13:410-431.
Not in our library10.1080/15592294.2018.1469891
291.
Sherman MH, et al. Stromal cues regulate the pancreatic cancer epigenome and metabolome. Proc. Natl Acad. Sci. USA. 2017;114:1129-1134.
Not in our library10.1073/pnas.1620164114
292.
Zhang Q, et al. By recruiting HDAC1, MORC2 suppresses p21 Waf1/Cip1 in gastric cancer. Oncotarget. 2015;6:16461-16470.
Not in our library10.18632/oncotarget.3889
293.
Dong ZY, Zhou YR, Wang LX. HDAC1 is indirectly involved in the epigenetic regulation of p38 MAPK that drive the lung cancer progression. Eur. Rev. Med. Pharm. Sci. 2018;22:5980-5986.
Not in our library10.26355/eurrev_201809_15932
294.
Zhang N, et al. SREBP1, targeted by miR-18a-5p, modulates epithelial-mesenchymal transition in breast cancer via forming a co-repressor complex with Snail and HDAC1/2. Cell Death Differ. 2019;26:843-859.
Not in our library10.1038/s41418-018-0158-8
295.
Li SZ, et al. Nemo-like kinase (NLK) primes colorectal cancer progression by releasing the E2F1 complex from HDAC1. Cancer Lett. 2018;431:43-53.
Not in our library10.1016/j.canlet.2018.05.032
296.
von BJ, et al. E-cadherin regulates metastasis of pancreatic cancer in vivo and is suppressed by a SNAIL/HDAC1/HDAC2 repressor complex. Gastroenterology. 2009;137:361-371.e1–5.
Not in our library10.1053/j.gastro.2009.04.004
297.
The p300/YY1/miR-500a-5p/HDAC2 signalling axis regulates cell proliferation in human colorectal cancer · 2019
No notice
298.
Zhu W, et al. FKBP3 promotes proliferation of non-small cell lung cancer cells through regulating Sp1/HDAC2/p27. Theranostics. 2017;7:3078-3089.
Not in our library10.7150/thno.18067
299.
Roy SS, et al. Significance of PELP1/HDAC2/miR-200 regulatory network in EMT and metastasis of breast cancer. Oncogene. 2014;33:3707-3716.
Not in our library10.1038/onc.2013.332
300.
Hulsurkar M, et al. Beta-adrenergic signaling promotes tumor angiogenesis and prostate cancer progression through HDAC2-mediated suppression of thrombospondin-1. Oncogene. 2017;36:1525-1536.
Not in our library10.1038/onc.2016.319
301.
Miao LJ, et al. Stat3 inhibits Beclin 1 expression through recruitment of HDAC3 in nonsmall cell lung cancer cells. Tumour Biol. 2014;35:7097-7103.
Not in our library10.1007/s13277-014-1961-6
302.
Chen G, et al. Long noncoding RNA SBF2-AS1 promotes colorectal cancer proliferation and invasion by inhibiting miR-619–5p activity and facilitating HDAC3 expression. J. Cell Physiol. 2019;234:18688-18696.
Not in our library10.1002/jcp.28509
303.
Bayat S, et al. Downregulation of HDAC2 and HDAC3 via oleuropein as a potent prevention and therapeutic agent in MCF-7 breast cancer cells. J. Cell Biochem. 2019;120:9172-9180.
Not in our library10.1002/jcb.28193
304.
Lucas J, et al. Upregulation of PD‑L1 expression by resveratrol and piceatannol in breast and colorectal cancer cells occurs via HDAC3/p300‑mediated NF‑κB signaling. Int J. Oncol. 2018;53:1469-1480.
Not in our library10.3892/ijo.2018.4512
305.
Kang Y, et al. HDAC8 and STAT3 repress BMF gene activity in colon cancer cells. Cell Death Dis. 2014;5:e1476.
Not in our library10.1038/cddis.2014.422
306.
Vanaja GR, Ramulu HG, Kalle AM. Overexpressed HDAC8 in cervical cancer cells shows functional redundancy of tubulin deacetylation with HDAC6. Cell Commun. Signal. 2018;16:20.
Not in our library10.1186/s12964-018-0231-4
307.
Wang ZT, et al. Histone deacetylase inhibitors suppress mutant p53 transcription via HDAC8/YY1 signals in triple negative breast cancer cells. Cell Signal. 2016;28:506-515.
Not in our library10.1016/j.cellsig.2016.02.006
308.
Wu S, et al. Suberoylanilide hydroxamic acid (SAHA) promotes the epithelial mesenchymal transition of triple negative breast cancer cells via HDAC8/FOXA1 signals. Biol. Chem. 2016;397:75-83.
Not in our library10.1515/hsz-2015-0215
309.
Halkidou K, et al. Nuclear accumulation of histone deacetylase 4 (HDAC4) coincides with the loss of androgen sensitivity in hormone refractory cancer of the prostate. Eur. Urol. 2004;45:382-389.
Not in our library10.1016/j.eururo.2003.10.005
310.
Wang B, et al. A dual role of miR-22 modulated by RelA/p65 in resensitizing fulvestrant-resistant breast cancer cells to fulvestrant by targeting FOXP1 and HDAC4 and constitutive acetylation of p53 at Lys382. Oncogenesis. 2018;7:54.
Not in our library10.1038/s41389-018-0063-5
311.
Wei JY, et al. Melatonin induces apoptosis of colorectal cancer cells through HDAC4 nuclear import mediated by CaMKII inactivation. J. Pineal Res. 2015;58:429-438.
Not in our library10.1111/jpi.12226
312.
Cao C, et al. HDAC5-LSD1 axis regulates antineoplastic effect of natural HDAC inhibitor sulforaphane in human breast cancer cells. Int. J. Cancer. 2018;143:1388-1401.
Not in our library10.1002/ijc.31419
313.
Liu C, et al. Hypermethylation of miRNA-589 promoter leads to upregulation of HDAC5 which promotes malignancy in non-small cell lung cancer. Int. J. Oncol. 2017;50:2079-2090.
Not in our library10.3892/ijo.2017.3967
314.
Sun S, et al. HDAC6 inhibitor TST strengthens the antiproliferative effects of PI3K/mTOR inhibitor BEZ235 in breast cancer cells via suppressing RTK activation. Cell Death Dis. 2018;9:929.
Not in our library10.1038/s41419-018-0931-0
315.
Won HR, et al. A452, an HDAC6-selective inhibitor, synergistically enhances the anticancer activity of chemotherapeutic agents in colorectal cancer cells. Mol. Carcinog. 2018;57:1383-1395.
Not in our library10.1002/mc.22852
316.
Wang Z, Hu P, Tang F, Xie C. HDAC6-mediated EGFR stabilization and activation restrict cell response to sorafenib in non-small cell lung cancer cells. Med. Oncol. 2016;33:50.
Not in our library10.1007/s12032-016-0765-5
317.
Seidel C, et al. 4-Hydroxybenzoic acid derivatives as HDAC6-specific inhibitors modulating microtubular structure and HSP90α chaperone activity against prostate cancer. Biochem. Pharmacol. 2016;99:31-52.
Not in our library10.1016/j.bcp.2015.11.005
318.
Witt AE, et al. Identification of a cancer stem cell-specific function for the histone deacetylases, HDAC1 and HDAC7, in breast and ovarian cancer. Oncogene. 2017;36:1707-1720.
Not in our library10.1038/onc.2016.337
319.
Wu MY, et al. MiR-34a regulates therapy resistance by targeting HDAC1 and HDAC7 in breast cancer. Cancer Lett. 2014;354:311-319.
Not in our library10.1016/j.canlet.2014.08.031
320.
Miremadi A, Oestergaard MZ, Pharoah PD, Caldas C. Cancer genetics of epigenetic genes. Hum. Mol. Genet. 2007;16:R28-R49.
Not in our library10.1093/hmg/ddm021
321.
Salgado E, et al. HDAC9 overexpression confers invasive and angiogenic potential to triple negative breast cancer cells via modulating microRNA-206. Biochem Biophys. Res. Commun. 2018;503:1087-1091.
Not in our library10.1016/j.bbrc.2018.06.120
322.
Okudela K, et al. Expression of HDAC9 in lung cancer-potential role in lung carcinogenesis. Int. J. Clin. Exp. Pathol. 2014;7:213-220.
Not in our library
323.
Yang Y, et al. HDAC10 promotes lung cancer proliferation via AKT phosphorylation. Oncotarget. 2016;7:59388-59401.
Not in our library10.18632/oncotarget.10673
324.
Islam MM, et al. HDAC10 as a potential therapeutic target in ovarian cancer. Gynecol. Oncol. 2017;144:613-620.
Not in our library10.1016/j.ygyno.2017.01.009
325.
Qin J, et al. Hypoxia-inducible factor 1 alpha promotes cancer stem cells-like properties in human ovarian cancer cells by upregulating SIRT1 expression. Sci. Rep. 2017;7:10592.
Not in our library10.1038/s41598-017-09244-8
326.
Santolla MF, et al. SIRT1 is involved in oncogenic signaling mediated by GPER in breast cancer. Cell Death Dis. 2015;6:e1834.
Not in our library10.1038/cddis.2015.201
327.
Sun L, et al. A SUMOylation-dependent pathway regulates SIRT1 transcription and lung cancer metastasis. J. Natl Cancer Inst. 2013;105:887-898.
Not in our library10.1093/jnci/djt118
328.
Farooqi AS, et al. Novel lysine-based thioureas as mechanism-based inhibitors of Sirtuin 2 (SIRT2) with anticancer activity in a colorectal cancer murine model. J. Med Chem. 2019;62:4131-4141.
Not in our library10.1021/acs.jmedchem.9b00191
329.
Hoffmann G, Breitenbücher F, Schuler M, Ehrenhofer-Murray AE. A novel sirtuin 2 (SIRT2) inhibitor with p53-dependent pro-apoptotic activity in non-small cell lung cancer. J. Biol. Chem. 2014;289:5208-5216.
Not in our library10.1074/jbc.m113.487736
330.
Li Y, et al. SIRT2 promotes the migration and invasion of gastric cancer through RAS/ERK/JNK/MMP-9 pathway by increasing PEPCK1-related metabolism. Neoplasia. 2018;20:745-756.
Not in our library10.1016/j.neo.2018.03.008
331.
Li Z, et al. SIRT2 inhibits non-small cell lung cancer cell growth through impairing Skp2-mediated p27 degradation. Oncotarget. 2016;7:18927-18939.
Not in our library10.18632/oncotarget.7816
332.
Fiskus W, et al. SIRT2 deacetylates and inhibits the peroxidase activity of peroxiredoxin-1 to sensitize breast cancer cells to oxidant stress-inducing agents. Cancer Res. 2016;76:5467-5478.
Not in our library10.1158/0008-5472.can-16-0126
333.
Damodaran S, et al. Dysregulation of Sirtuin 2 (SIRT2) and histone H3K18 acetylation pathways associates with adverse prostate cancer outcomes. BMC Cancer. 2017;17:874.
Not in our library10.1186/s12885-017-3853-9
334.
Xiong Y, et al. SIRT3 deacetylates and promotes degradation of P53 in PTEN-defective non-small cell lung cancer. J. Cancer Res. Clin. Oncol. 2018;144:189-198.
Not in our library10.1007/s00432-017-2537-9
335.
Wang S, et al. Programmed death ligand 1 promotes lymph node metastasis and glucose metabolism in cervical cancer by activating integrin β4/SNAI1/SIRT3 signaling pathway. Oncogene. 2018;37:4164-4180.
Not in our library10.1038/s41388-018-0252-x
336.
Wu Y, et al. SIRT3 aggravates metformin-induced energy stress and apoptosis in ovarian cancer cells. Exp. Cell Res. 2018;367:137-149.
Not in our library10.1016/j.yexcr.2018.03.030
337.
Quan Y, et al. SIRT3 inhibits prostate cancer by destabilizing oncoprotein c-MYC through regulation of the PI3K/Akt pathway. Oncotarget. 2015;6:26494-26507.
Not in our library10.18632/oncotarget.4764
338.
Lee DY, et al. Regulation of SIRT3 signal related metabolic reprogramming in gastric cancer by Helicobacter pylori oncoprotein CagA. Oncotarget. 2017;8:78365-78378.
Not in our library10.18632/oncotarget.18695
339.
Miyo M, et al. Tumour-suppressive function of SIRT4 in human colorectal cancer. Br. J. Cancer. 2015;113:492-499.
Not in our library10.1038/bjc.2015.226
340.
Sun H, et al. SIRT4 acts as a tumor suppressor in gastric cancer by inhibiting cell proliferation, migration, and invasion. Onco Targets Ther. 2018;11:3959-3968.
Not in our library10.2147/ott.s156143
341.
Xu L, et al. SIRT5 as a biomarker for response to anthracycline-taxane-based neoadjuvant chemotherapy in triple-negative breast cancer. Oncol. Rep. 2018;39:2315-2323.
Not in our library10.3892/or.2018.6319
342.
SIRT 5‐mediated deacetylation of LDHB promotes autophagy and tumorigenesis in colorectal cancer · 2018
No notice
343.
Lu W, Zuo Y, Feng Y, Zhang M. SIRT5 facilitates cancer cell growth and drug resistance in non-small cell lung cancer. Tumour Biol. 2014;35:10699-10705.
Not in our library10.1007/s13277-014-2372-4
344.
Tang Z, et al. CDK2 positively regulates aerobic glycolysis by suppressing SIRT5 in gastric cancer. Cancer Sci. 2018;109:2590-2598.
Not in our library10.1111/cas.13691
345.
Chen XF, et al. SIRT5 inhibits peroxisomal ACOX1 to prevent oxidative damage and is downregulated in liver cancer. EMBO Rep. 2018;19:e45124.
Not in our library10.15252/embr.201745124
346.
Liu Y, et al. Inhibition of SIRT6 in prostate cancer reduces cell viability and increases sensitivity to chemotherapeutics. Protein Cell. 2013;4:702-710.
Not in our library10.1007/s13238-013-3054-5
347.
Bauer I, et al. The NAD+-dependent histone deacetylase SIRT6 promotes cytokine production and migration in pancreatic cancer cells by regulating Ca2+ responses. J. Biol. Chem. 2012;287:40924-40937.
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348.
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349.
Min L, et al. Liver cancer initiation is controlled by AP-1 through SIRT6-dependent inhibition of survivin. Nat. Cell Biol. 2012;14:1203-1211.
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350.
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351.
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352.
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353.
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354.
SIRT7 antagonizes TGF-β signaling and inhibits breast cancer metastasis · 2017
No notice
355.
Jiang Y, Han Z, Wang Y, Hao W. Depletion of SIRT7 sensitizes human non-small cell lung cancer cells to gemcitabine therapy by inhibiting autophagy. Biochem Biophys. Res. Commun. 2018;506:266-271.
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356.
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357.
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358.
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359.
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360.
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361.
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362.
H3.3K27M mutant proteins reprogram epigenome by sequestering the PRC2 complex to poised enhancers · 2018
Not checked
363.
Chan KM, et al. A lesson learned from the H3.3K27M mutation found in pediatric glioma: a new approach to the study of the function of histone modifications in vivo. Cell Cycle. 2013;12:2546-2552.
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364.
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365.
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366.
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367.
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368.
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369.
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370.
Min J, et al. Structure of the catalytic domain of human DOT1L, a non-SET domain nucleosomal histone methyltransferase. Cell. 2003;112:711-723.
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371.
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372.
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373.
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374.
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375.
Kuzmichev A, Jenuwein T, Tempst P, Reinberg D. Different EZH2-containing complexes target methylation of histone H1 or nucleosomal histone H3. Mol. Cell. 2004;14:183-193.
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376.
Structure and Function of SET and MYND Domain-Containing Proteins · 2015
No notice
377.
Derunes C, et al. Characterization of the PR domain of RIZ1 histone methyltransferase. Biochem Biophys. Res. Commun. 2005;333:925-934.
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378.
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379.
Davis CA, et al. PRISM/PRDM6, a transcriptional repressor that promotes the proliferative gene program in smooth muscle cells. Mol. Cell Biol. 2006;26:2626-2636.
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380.
Gyory I, et al. PRDI-BF1 recruits the histone H3 methyltransferase G9a in transcriptional silencing. Nat. Immunol. 2004;5:299-308.
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381.
RNA-dependent stabilization of SUV39H1 at constitutive heterochromatin · 2017
No notice
382.
Hashimoto H, Vertino PM, Cheng X. Molecular coupling of DNA methylation and histone methylation. Epigenomics. 2010;2:657-669.
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383.
H3K14ac is linked to methylation of H3K9 by the triple Tudor domain of SETDB1 · 2017
Not checked
384.
Bantignies F, Goodman RH, Smolik SM. Functional interaction between the coactivator Drosophila CREB-binding protein and ASH1, a member of the trithorax group of chromatin modifiers. Mol. Cell Biol. 2000;20:9317-9330.
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385.
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386.
Meyer R, Wolf SS, Obendorf M. PRMT2, a member of the protein arginine methyltransferase family, is a coactivator of the androgen receptor. J. Steroid Biochem. Mol. Biol. 2007;107:1-14.
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387.
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388.
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389.
Slany RK. The molecular biology of mixed lineage leukemia. Haematologica. 2009;94:984-993.
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390.
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391.
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392.
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393.
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394.
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395.
Wysocka J, et al. WDR5 associates with histone H3 methylated at K4 and is essential for H3 K4 methylation and vertebrate development. Cell. 2005;121:859-872.
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396.
Shi X, et al. ING2 PHD domain links histone H3 lysine 4 methylation to active gene repression. Nature. 2006;442:96-99.
Not in our library10.1038/nature04835
397.
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398.
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399.
Trojer P, et al. L3MBTL1, a histone-methylation-dependent chromatin lock. Cell. 2007;129:915-928.
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400.
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401.
Lan F, et al. Recognition of unmethylated histone H3 lysine 4 links BHC80 to LSD1-mediated gene repression. Nature. 2007;448:718-722.
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402.
Ooi SK, et al. DNMT3L connects unmethylated lysine 4 of histone H3 to de novo methylation of DNA. Nature. 2007;448:714-717.
Not in our library10.1038/nature05987
403.
Lee J, Thompson JR, Botuyan MV, Mer G. Distinct binding modes specify the recognition of methylated histones H3K4 and H4K20 by JMJD2A-tudor. Nat. Struct. Mol. Biol. 2008;15:109-111.
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404.
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405.
Tsukada Y, et al. Histone demethylation by a family of JmjC domain-containing proteins. Nature. 2006;439:811-816.
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406.
Chang B, Chen Y, Zhao Y, Bruick RK. JMJD6 is a histone arginine demethylase. Science. 2007;318:444-447.
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407.
Wang Y, et al. Human PAD4 regulates histone arginine methylation levels via demethylimination. Science. 2004;306:279-283.
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408.
Schulte JH, et al. Lysine-specific demethylase 1 is strongly expressed in poorly differentiated neuroblastoma: implications for therapy. Cancer Res. 2009;69:2065-2071.
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409.
Ciccone DN, et al. KDM1B is a histone H3K4 demethylase required to establish maternal genomic imprints. Nature. 2009;461:415-418.
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410.
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411.
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412.
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413.
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414.
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415.
Wang J, et al. Epigenetic regulation of miR-302 by JMJD1C inhibits neural differentiation of human embryonic stem cells. J. Biol. Chem. 2014;289:2384-2395.
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416.
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417.
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418.
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419.
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420.
Lee KH, et al. PHF2 histone demethylase acts as a tumor suppressor in association with p53 in cancer. Oncogene. 2015;34:2897-2909.
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421.
Cai M, et al. Epigenetic silenced miR-125a-5p could be self-activated through targeting Suv39H1 in gastric cancer. J. Cell Mol. Med. 2018;22:4721-4731.
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422.
Yu T, et al. Metformin inhibits SUV39H1-mediated migration of prostate cancer cells. Oncogenesis. 2017;6:e324.
Not in our library10.1038/oncsis.2017.28
423.
Mo W, et al. mTOR inhibitors suppress homologous recombination repair and synergize with PARP inhibitors via regulating SUV39H1 in BRCA-proficient triple-negative breast cancer. Clin. Cancer Res. 2016;22:1699-1712.
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424.
Pandey M, et al. Involvement of EZH2, SUV39H1, G9a and associated molecules in pathogenesis of urethane induced mouse lung tumors: potential targets for cancer control. Toxicol. Appl. Pharmacol. 2014;280:296-304.
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425.
Kang MY, et al. Association of the SUV39H1 histone methyltransferase with the DNA methyltransferase 1 at mRNA expression level in primary colorectal cancer. Int J. Cancer. 2007;121:2192-2197.
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426.
Yang Z, et al. The KMT1A-GATA3-STAT3 circuit is a novel self-renewal signaling of human bladder cancer stem cells. Clin. Cancer Res. 2017;23:6673-6685.
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427.
Khanal P, et al. Prolyl isomerase Pin1 negatively regulates the stability of SUV39H1 to promote tumorigenesis in breast cancer. FASEB J. 2013;27:4606-4618.
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428.
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429.
Shuai W, et al. SUV39H2 promotes colorectal cancer proliferation and metastasis via tri-methylation of the SLIT1 promoter. Cancer Lett. 2018;422:56-69.
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430.
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431.
Nishikawaji T, et al. Oncogenic roles of the SETDB2 histone methyltransferase in gastric cancer. Oncotarget. 2016;7:67251-67265.
Not in our library10.18632/oncotarget.11625
432.
Wang YF, et al. G9a regulates breast cancer growth by modulating iron homeostasis through the repression of ferroxidase hephaestin. Nat. Commun. 2017;8:274.
Not in our library10.1038/s41467-017-00350-9
433.
Tian YF, et al. Preprogramming therapeutic response of PI3K/mTOR dual inhibitor via the regulation of EHMT2 and p27 in pancreatic cancer. Am. J. Cancer Res. 2018;8:1812-1822.
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434.
Cui J, et al. EHMT2 inhibitor BIX-01294 induces apoptosis through PMAIP1-USP9X-MCL1 axis in human bladder cancer cells. Cancer Cell Int. 2015;15:4.
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435.
Ezponda T, et al. The histone methyltransferase MMSET/WHSC1 activates TWIST1 to promote an epithelial-mesenchymal transition and invasive properties of prostate cancer. Oncogene. 2013;32:2882-2890.
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436.
Ryu TY, et al. SETDB1 regulates SMAD7 expression for breast cancer metastasis. BMB Rep. 2019;52:139-144.
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437.
Chen K, et al. Histone methyltransferase SETDB1 promotes the progression of colorectal cancer by inhibiting the expression of TP53. J. Cancer. 2017;8:3318-3330.
Not in our library10.7150/jca.20482
438.
Wong CM, et al. Up-regulation of histone methyltransferase SETDB1 by multiple mechanisms in hepatocellular carcinoma promotes cancer metastasis. Hepatology. 2016;63:474-487.
Not in our library10.1002/hep.28304
439.
Histone methyltransferase SETDB1 regulates liver cancer cell growth through methylation of p53 · 2015
No notice
440.
Wu PC, et al. H3K9 histone methyltransferase, KMT1E/SETDB1, cooperates with the SMAD2/3 pathway to suppress lung cancer metastasis. Cancer Res. 2014;74:7333-7343.
Not in our library10.1158/0008-5472.can-13-3572
441.
Zhu Q, et al. The Wnt-driven Mll1 epigenome regulates salivary gland and head and neck cancer. Cell Rep. 2019;26:415-428.e5.
Not in our library10.1016/j.celrep.2018.12.059
442.
Lu C, et al. The MLL1-H3K4me3 axis-mediated PD-L1 expression and pancreatic cancer immune evasion. J. Natl Cancer Inst. 2017;109:djw283.
Not in our library10.1093/jnci/djw283
443.
Yang Z, et al. Single-cell sequencing reveals variants in ARID1A, GPRC5A and MLL2 driving self-renewal of human bladder cancer stem cells. Eur. Urol. 2017;71:8-12.
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444.
Augert A, et al. Small cell lung cancer exhibits frequent inactivating mutations in the histone methyltransferase KMT2D/MLL2: CALGB 151111 (Alliance). J. Thorac. Oncol. 2017;12:704-713.
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445.
Chen J, et al. Pygo2 associates with MLL2 histone methyltransferase and GCN5 histone acetyltransferase complexes to augment Wnt target gene expression and breast cancer stem-like cell expansion. Mol. Cell Biol. 2010;30:5621-5635.
Not in our library10.1128/mcb.00465-10
446.
Xia M, et al. Downregulation of MLL3 in esophageal squamous cell carcinoma is required for the growth and metastasis of cancer cells. Tumour Biol. 2015;36:605-613.
Not in our library10.1007/s13277-014-2616-3
447.
Kim JH, et al. UTX and MLL4 coordinately regulate transcriptional programs for cell proliferation and invasiveness in breast cancer cells. Cancer Res. 2014;74:1705-1717.
Not in our library10.1158/0008-5472.can-13-1896
448.
Gallo M, et al. MLL5 orchestrates a cancer self-renewal state by repressing the histone variant H3.3 and globally reorganizing chromatin. Cancer Cell. 2015;28:715-729.
Not in our library10.1016/j.ccell.2015.10.005
449.
Li T, et al. SET1A cooperates with CUDR to promote liver cancer growth and hepatocyte-like stem cell malignant transformation epigenetically. Mol. Ther. 2016;24:261-275.
Not in our library10.1038/mt.2015.208
450.
Karakashev S, et al. CARM1-expressing ovarian cancer depends on the histone methyltransferase EZH2 activity. Nat. Commun. 2018;9:631.
Not in our library10.1038/s41467-018-03031-3
451.
Crea F, et al. An EZH2 polymorphism is associated with clinical outcome in metastatic colorectal cancer patients. Ann. Oncol. 2012;23:1207-1213.
Not in our library10.1093/annonc/mdr387
452.
Reijm EA, et al. High protein expression of EZH2 is related to unfavorable outcome to tamoxifen in metastatic breast cancer. Ann. Oncol. 2014;25:2185-2190.
Not in our library10.1093/annonc/mdu391
453.
Kanu N, et al. SETD2 loss-of-function promotes renal cancer branched evolution through replication stress and impaired DNA repair. Oncogene. 2015;34:5699-5708.
Not in our library10.1038/onc.2015.24
454.
Kim IK, et al. Acquired SETD2 mutation and impaired CREB1 activation confer cisplatin resistance in metastatic non-small cell lung cancer. Oncogene. 2019;38:180-193.
Not in our library10.1038/s41388-018-0429-3
455.
Li N, et al. AKT-mediated stabilization of histone methyltransferase WHSC1 promotes prostate cancer metastasis. J. Clin. Invest. 2017;127:1284-1302.
Not in our library10.1172/jci91144
456.
Li J, et al. MiR-2392 suppresses metastasis and epithelial-mesenchymal transition by targeting MAML3 and WHSC1 in gastric cancer. FASEB J. 2017;31:3774-3786.
Not in our library10.1096/fj.201601140rr
457.
Saloura V, et al. WHSC1L1-mediated EGFR mono-methylation enhances the cytoplasmic and nuclear oncogenic activity of EGFR in head and neck cancer. Sci. Rep. 2017;7:40664.
Not in our library10.1038/srep40664
458.
Chiou SH, et al. BLIMP1 induces transient metastatic heterogeneity in pancreatic cancer. Cancer Discov. 2017;7:1184-1199.
Not in our library10.1158/2159-8290.cd-17-0250
459.
Sciortino M, et al. Dysregulation of Blimp1 transcriptional repressor unleashes p130Cas/ErbB2 breast cancer invasion. Sci. Rep. 2017;7:1145.
Not in our library10.1038/s41598-017-01332-z
460.
Zhu Z, et al. Downregulation of PRDM1 promotes cellular invasion and lung cancer metastasis. Tumour Biol. 2017;39:1010428317695929.
Not in our library10.1177/1010428317695929
461.
Kang HB, et al. PRDM1, a tumor-suppressor gene, is induced by genkwadaphnin in human colon cancer SW620 cells. J. Cell Biochem. 2016;117:172-179.
Not in our library10.1002/jcb.25262
462.
Emterling A, Wallin A, Arbman G, Sun XF. Clinicopathological significance of microsatellite instability and mutated RIZ in colorectal cancer. Ann. Oncol. 2004;15:242-246.
Not in our library10.1093/annonc/mdh045
463.
Abbondanza C, et al. Identification of a functional estrogen-responsive enhancer element in the promoter 2 of PRDM2 gene in breast cancer cell lines. J. Cell Physiol. 2012;227:964-975.
Not in our library10.1002/jcp.22803
464.
Nanjundan M, et al. Amplification of MDS1/EVI1 and EVI1, located in the 3q26.2 amplicon, is associated with favorable patient prognosis in ovarian cancer. Cancer Res. 2007;67:3074-3084.
Not in our library10.1158/0008-5472.can-06-2366
465.
EVI1 promotes epithelial-to-mesenchymal transition, cancer stem cell features and chemo−/radioresistance in nasopharyngeal carcinoma · 2019
Not checked
466.
Taniguchi H, et al. Silencing PRDM14 expression by an innovative RNAi therapy inhibits stemness, tumorigenicity, and metastasis of breast cancer. Oncotarget. 2017;8:46856-46874.
Not in our library10.18632/oncotarget.16776
467.
Watanabe Y, et al. PRDM5 identified as a target of epigenetic silencing in colorectal and gastric cancer. Clin. Cancer Res. 2007;13:4786-4794.
Not in our library10.1158/1078-0432.ccr-07-0305
468.
Houle AA, et al. Aberrant PRDM9 expression impacts the pan-cancer genomic landscape. Genome Res. 2018;28:1611-1620.
Not in our library10.1101/gr.231696.117
469.
Ruark E, et al. Identification of nine new susceptibility loci for testicular cancer, including variants near DAZL and PRDM14. Nat. Genet. 2013;45:686-689.
Not in our library10.1038/ng.2635
470.
Moriya C, et al. Inhibition of PRDM14 expression in pancreatic cancer suppresses cancer stem-like properties and liver metastasis in mice. Carcinogenesis. 2017;38:638-648.
Not in our library10.1093/carcin/bgx040
471.
Takahata M, et al. SKI and MEL1 cooperate to inhibit transforming growth factor-beta signal in gastric cancer cells. J. Biol. Chem. 2009;284:3334-3344.
Not in our library10.1074/jbc.m808989200
472.
Reynoird N, et al. Coordination of stress signals by the lysine methyltransferase SMYD2 promotes pancreatic cancer. Genes Dev. 2016;30:772-785.
Not in our library10.1101/gad.275529.115
473.
Komatsu S, et al. Overexpression of SMYD2 contributes to malignant outcome in gastric cancer. Br. J. Cancer. 2015;112:357-364.
Not in our library10.1038/bjc.2014.543
474.
Sarris ME, et al. Smyd3 is a transcriptional potentiator of multiple cancer-promoting genes and required for liver and colon cancer development. Cancer Cell. 2016;29:354-366.
Not in our library10.1016/j.ccell.2016.01.013
475.
Cock-Rada AM, et al. SMYD3 promotes cancer invasion by epigenetic upregulation of the metalloproteinase MMP-9. Cancer Res. 2012;72:810-820.
Not in our library10.1158/0008-5472.can-11-1052
476.
Fenizia C, et al. SMYD3 promotes the epithelial-mesenchymal transition in breast cancer. Nucleic Acids Res. 2018;47:1278-1293.
Not in our library10.1093/nar/gky1221
477.
Hu L, Zhu YT, Qi C, Zhu YJ. Identification of Smyd4 as a potential tumor suppressor gene involved in breast cancer development. Cancer Res. 2009;69:4067-4072.
Not in our library10.1158/0008-5472.can-08-4097
478.
Kryczek I, et al. IL-22(+)CD4(+) T cells promote colorectal cancer stemness via STAT3 transcription factor activation and induction of the methyltransferase DOT1L. Immunity. 2014;40:772-784.
Not in our library10.1016/j.immuni.2014.03.010
479.
DOT1L cooperates with the c-Myc-p300 complex to epigenetically derepress CDH1 transcription factors in breast cancer progression · 2015
No notice
480.
Liu B, et al. MiR-502/SET8 regulatory circuit in pathobiology of breast cancer. Cancer Lett. 2016;376:259-267.
Not in our library10.1016/j.canlet.2016.04.008
481.
Yang S, et al. Association of miR-502-binding site single nucleotide polymorphism in the 3′-untranslated region of SET8 and TP53 codon 72 polymorphism with non-small cell lung cancer in Chinese population. Acta Biochim Biophys. Sin. 2014;46:149-154.
Not in our library10.1093/abbs/gmt138
482.
Tryndyak VP, Kovalchuk O, Pogribny IP. Loss of DNA methylation and histone H4 lysine 20 trimethylation in human breast cancer cells is associated with aberrant expression of DNA methyltransferase 1, Suv4–20h2 histone methyltransferase and methyl-binding proteins. Cancer Biol. Ther. 2006;5:65-70.
Not in our library10.4161/cbt.5.1.2288
483.
Martín-Morales L, et al. SETD6 dominant negative mutation in familial colorectal cancer type X. Hum. Mol. Genet. 2017;26:4481-4493.
Not in our library10.1093/hmg/ddx336
484.
Yao R, et al. Lysines 207 and 325 methylation of WDR5 catalyzed by SETD6 promotes breast cancer cell proliferation and migration. Oncol. Rep. 2018;40:3069-3077.
Not in our library10.3892/or.2018.6669
485.
Shen C, et al. SET7/9 regulates cancer cell proliferation by influencing β-catenin stability. FASEB J. 2015;29:4313-4323.
Not in our library10.1096/fj.15-273540
486.
Gu Y, et al. Opposite effects of SET7/9 on apoptosis of human acute myeloid leukemia cells and lung cancer cells. J. Cancer. 2017;8:2069-2078.
Not in our library10.7150/jca.19143
487.
Montenegro MF, et al. Tumor suppressor SET9 guides the epigenetic plasticity of breast cancer cells and serves as an early-stage biomarker for predicting metastasis. Oncogene. 2016;35:6143-6152.
Not in our library10.1038/onc.2016.154
488.
Liu LM, et al. Methylation of C/EBPα by PRMT1 inhibits its tumor suppressive function in breast cancer. Cancer Res. 2019;79:2865-2877.
Not in our library10.1158/0008-5472.can-18-3211
489.
PRMT1 Is a Novel Regulator of Epithelial-Mesenchymal-Transition in Non-small Cell Lung Cancer · 2015
No notice
490.
Mathioudaki K, et al. The PRMT1 gene expression pattern in colon cancer. Br. J. Cancer. 2008;99:2094-2099.
Not in our library10.1038/sj.bjc.6604807
491.
Lin Z, et al. Overexpressing PRMT1 inhibits proliferation and invasion in pancreatic cancer by inverse correlation of ZEB1. IUBMB Life. 2018;70:1032-1039.
Not in our library10.1002/iub.1917
492.
Zhong J, et al. PRMT2β, a C-terminal splice variant of PRMT2, inhibits the growth of breast cancer cells. Oncol. Rep. 2017;38:1303-1311.
Not in our library10.3892/or.2017.5786
493.
Al-Dhaheri M., Wu J., Skliris G. P., Li J., Higashimato K., Wang Y., White K. P., Lambert P., Zhu Y., Murphy L., Xu W. CARM1 Is an Important Determinant of ER -Dependent Breast Cancer Cell Differentiation and Proliferation in Breast Cancer Cells. Cancer Research. 2011;71(6):2118-2128.
Not in our library10.1158/0008-5472.can-10-2426
494.
Zhong XY, et al. CARM1 methylates GAPDH to regulate glucose metabolism and is suppressed in liver cancer. Cell Rep. 2018;24:3207-3223.
Not in our library10.1016/j.celrep.2018.08.066
495.
Wang YP, et al. Arginine methylation of MDH1 by CARM1 inhibits glutamine metabolism and suppresses pancreatic cancer. Mol. Cell. 2016;64:673-687.
Not in our library10.1016/j.molcel.2016.09.028
496.
Rengasamy M, et al. The PRMT5/WDR77 complex regulates alternative splicing through ZNF326 in breast cancer. Nucleic Acids Res. 2017;45:11106-11120.
Not in our library10.1093/nar/gkx727
497.
Fu T, Lv X, Kong Q, Yuan C. A novel SHARPIN-PRMT5-H3R2me1 axis is essential for lung cancer cell invasion. Oncotarget. 2017;8:54809-54820.
Not in our library10.18632/oncotarget.18957
498.
Demetriadou C, et al. NAA40 contributes to colorectal cancer growth by controlling PRMT5 expression. Cell Death Dis. 2019;10:236.
Not in our library10.1038/s41419-019-1487-3
499.
Huang S, et al. CAPG enhances breast cancer metastasis by competing with PRMT5 to modulate STC-1 transcription. Theranostics. 2018;8:2549-2564.
Not in our library10.7150/thno.22523
500.
Almeida-Rios D, et al. Histone methyltransferase PRMT6 plays an oncogenic role of in prostate cancer. Oncotarget. 2016;7:53018-53028.
Not in our library10.18632/oncotarget.10061
501.
Okuno K, et al. Asymmetric dimethylation at histone H3 arginine 2 by PRMT6 in gastric cancer progression. Carcinogenesis. 2019;40:15-26.
Not in our library10.1093/carcin/bgy147
502.
Chan LH, et al. PRMT6 regulates RAS/RAF binding and MEK/ERK-mediated cancer stemness activities in hepatocellular carcinoma through CRAF methylation. Cell Rep. 2018;25:690-701.e8.
Not in our library10.1016/j.celrep.2018.09.053
503.
Cheng D, et al. PRMT7 contributes to the metastasis phenotype in human non-small-cell lung cancer cells possibly through the interaction with HSPA5 and EEF2. Onco. Targets Ther. 2018;11:4869-4876.
Not in our library10.2147/ott.s166412
504.
Yao R, et al. PRMT7 induces epithelial-to-mesenchymal transition and promotes metastasis in breast cancer. Cancer Res. 2014;74:5656-5667.
Not in our library10.1158/0008-5472.can-14-0800
505.
Hernandez SJ, Dolivo DM, Dominko T. PRMT8 demonstrates variant-specific expression in cancer cells and correlates with patient survival in breast, ovarian and gastric cancer. Oncol. Lett. 2017;13:1983-1989.
Not in our library10.3892/ol.2017.5671
506.
Bagger SO, et al. Aggressiveness of non-EMT breast cancer cells relies on FBXO11 activity. Mol. Cancer. 2018;17:171.
Not in our library10.1186/s12943-018-0918-6
507.
Lee YH, Ann DK. Bi-phasic expression of Heterochromatin Protein 1 (HP1) during breast cancer progression: potential roles of HP1 and chromatin structure in tumorigenesis. J. Nat. Sci. 2015;1:e127.
Not in our library
508.
Shenoy TR, et al. CHD1 loss sensitizes prostate cancer to DNA damaging therapy by promoting error-prone double-strand break repair. Ann. Oncol. 2017;28:1495-1507.
Not in our library10.1093/annonc/mdx165
509.
Rodrigues LU, et al. Coordinate loss of MAP3K7 and CHD1 promotes aggressive prostate cancer. Cancer Res. 2015;75:1021-1034.
Not in our library10.1158/0008-5472.can-14-1596
510.
Pan L, Tang Z, Pan L, Tang R. miR-3666 inhibits lung cancer cell proliferation, migration and invasion by targeting BPTF. Biochem Cell Biol. 2018;97:415-422.
Not in our library10.1139/bcb-2018-0301
511.
Zhao X, et al. BPTF promotes hepatocellular carcinoma growth by modulating hTERT signaling and cancer stem cell traits. Redox Biol. 2019;20:427-441.
Not in our library10.1016/j.redox.2018.10.018
512.
Zeng H, et al. Physical activity and breast cancer survival: an epigenetic link through reduced methylation of a tumor suppressor gene L3MBTL1. Breast Cancer Res. Treat. 2012;133:127-135.
Not in our library10.1007/s10549-011-1716-7
513.
Kumamoto K, et al. ING2 is upregulated in colon cancer and increases invasion by enhanced MMP13 expression. Int J. Cancer. 2009;125:1306-1315.
Not in our library10.1002/ijc.24437
514.
Okano T, et al. Alterations in novel candidate tumor suppressor genes, ING1 and ING2 in human lung cancer. Oncol. Rep. 2006;15:545-549.
Not in our library
515.
Li Y, et al. RNA splicing of the BHC80 gene contributes to neuroendocrine prostate cancer progression. Eur. Urol. 2019;76:157-166.
Not in our library10.1016/j.eururo.2019.03.011
516.
Li LL, et al. JMJD2A contributes to breast cancer progression through transcriptional repression of the tumor suppressor ARHI. Breast Cancer Res. 2014;16:R56.
Not in our library10.1186/bcr3667
517.
Kim TD, et al. The JMJD2A demethylase regulates apoptosis and proliferation in colon cancer cells. J. Cell Biochem. 2012;113:1368-1376.
Not in our library10.1002/jcb.24009
518.
Kauffman EC, et al. Role of androgen receptor and associated lysine-demethylase coregulators, LSD1 and JMJD2A, in localized and advanced human bladder cancer. Mol. Carcinog. 2011;50:931-944.
Not in our library10.1002/mc.20758
519.
Cao C, et al. Functional interaction of histone deacetylase 5 (HDAC5) and lysine-specific demethylase 1 (LSD1) promotes breast cancer progression. Oncogene. 2017;36:133-145.
Not in our library10.1038/onc.2016.186
520.
Stewart CA, Byers LA. Altering the course of small cell lung cancer: targeting cancer stem cells via LSD1 inhibition. Cancer Cell. 2015;28:4-6.
Not in our library10.1016/j.ccell.2015.06.011
521.
Liang Y, et al. LSD1-mediated epigenetic reprogramming drives CENPE expression and prostate cancer progression. Cancer Res. 2017;77:5479-5490.
Not in our library10.1158/0008-5472.can-17-0496
522.
Wang Y, et al. LSD1 is a subunit of the NuRD complex and targets the metastasis programs in breast cancer. Cell. 2009;138:660-672.
Not in our library10.1016/j.cell.2009.05.050
523.
Chen L, et al. Functional characterization of lysine-specific demethylase 2 (LSD2/KDM1B) in breast cancer progression. Oncotarget. 2017;8:81737-81753.
Not in our library10.18632/oncotarget.19387
524.
Chen JY, et al. Lysine demethylase KDM2A inhibits TET2 to promote DNA methylation and silencing of tumor suppressor genes in breast cancer. Oncogenesis. 2017;6:e369.
Not in our library10.1038/oncsis.2017.71
525.
Kong Y, et al. RUNX3-mediated up-regulation of miR-29b suppresses the proliferation and migration of gastric cancer cells by targeting KDM2A. Cancer Lett. 2016;381:138-148.
Not in our library10.1016/j.canlet.2016.07.038
526.
Dhar SS, et al. Transcriptional repression of histone deacetylase 3 by the histone demethylase KDM2A is coupled to tumorigenicity of lung cancer cells. J. Biol. Chem. 2014;289:7483-7496.
Not in our library10.1074/jbc.m113.521625
527.
Zacharopoulou N, et al. The epigenetic factor KDM2B regulates cell adhesion, small rho GTPases, actin cytoskeleton and migration in prostate cancer cells. Biochim Biophys. Acta Mol. Cell Res. 2018;1865:587-597.
Not in our library10.1016/j.bbamcr.2018.01.009
528.
Kottakis F, et al. NDY1/KDM2B functions as a master regulator of polycomb complexes and controls self-renewal of breast cancer stem cells. Cancer Res. 2014;74:3935-3946.
Not in our library10.1158/0008-5472.can-13-2733
529.
Peng K, et al. Histone demethylase JMJD1A promotes colorectal cancer growth and metastasis by enhancing Wnt/β-catenin signaling. J. Biol. Chem. 2018;293:10606-10619.
Not in our library10.1074/jbc.ra118.001730
530.
Ramadoss S, et al. Lysine-specific demethylase KDM3A regulates ovarian cancer stemness and chemoresistance. Oncogene. 2017;36:6508.
Not in our library10.1038/onc.2017.331
531.
Wade MA, et al. The histone demethylase enzyme KDM3A is a key estrogen receptor regulator in breast cancer. Nucleic Acids Res. 2015;43:196-207.
Not in our library10.1093/nar/gku1298
532.
Cai Y, Fu X, Deng Y. Histone demethylase JMJD1C regulates esophageal cancer proliferation Via YAP1 signaling. Am. J. Cancer Res. 2017;7:115-124.
Not in our library
533.
Chen C, et al. Downregulation of histone demethylase JMJD1C inhibits colorectal cancer metastasis through targeting ATF2. Am. J. Cancer Res. 2018;8:852-865.
Not in our library
534.
An J, et al. HistoneH3 demethylase JMJD2A promotes growth of liver cancer cells through up-regulating miR372. Oncotarget. 2017;8:49093-49109.
Not in our library10.18632/oncotarget.17095
535.
Wang W, et al. KDM4B-regulated unfolded protein response as a therapeutic vulnerability in PTEN-deficient breast cancer. J. Exp. Med. 2018;215:2833-2849.
Not in our library10.1084/jem.20180439
536.
Zhao L, et al. JMJD2B promotes epithelial-mesenchymal transition by cooperating with β-catenin and enhances gastric cancer metastasis. Clin. Cancer Res. 2013;19:6419-6429.
Not in our library10.1158/1078-0432.ccr-13-0254
537.
Fu LN, et al. Role of JMJD2B in colon cancer cell survival under glucose-deprived conditions and the underlying mechanisms. Oncogene. 2018;37:389-402.
Not in our library10.1038/onc.2017.345
538.
Luo W, et al. Histone demethylase JMJD2C is a coactivator for hypoxia-inducible factor 1 that is required for breast cancer progression. Proc. Natl Acad. Sci. USA. 2012;109:E3367-E3376.
Not in our library10.1073/pnas.1217394109
539.
An Y, et al. circZMYM2 competed endogenously with miR-335-5p to regulate JMJD2C in pancreatic cancer. Cell Physiol. Biochem. 2018;51:2224-2236.
Not in our library10.1159/000495868
540.
Peng K, et al. Histone demethylase JMJD2D interacts with β-catenin to induce transcription and activate colorectal cancer cell proliferation and tumor growth in mice. Gastroenterology. 2019;156:1112-1126.
Not in our library10.1053/j.gastro.2018.11.036
541.
Yang GJ, et al. Selective inhibition of lysine-specific demethylase 5A (KDM5A) using a Rhodium(III) complex for triple-negative breast cancer therapy. Angew. Chem. Int Ed. Engl. 2018;57:13091-13095.
Not in our library10.1002/anie.201807305
542.
Uemura M, et al. Jumonji domain containing 1A is a novel prognostic marker for colorectal cancer: in vivo identification from hypoxic tumor cells. Clin. Cancer Res. 2010;16:4636-4646.
Not in our library10.1158/1078-0432.ccr-10-0407
543.
Overexpression of the JmjC histone demethylase KDM5B in human carcinogenesis: involvement in the proliferation of cancer cells through the E2F/RB pathway · 2010
No notice
544.
Li N, Jiang D. Jumonji domain containing 2C promotes cell migration and invasion through modulating CUL4A expression in lung cancer. Biomed. Pharmacother. 2017;89:305-315.
Not in our library10.1016/j.biopha.2017.02.014
545.
Lin H, et al. KDM5c inhibits multidrug resistance of colon cancer cell line by down-regulating ABCC1. Biomed. Pharmacother. 2018;107:1205-1209.
Not in our library10.1016/j.biopha.2018.08.041
546.
Shen X, et al. KDM5D inhibit epithelial-mesenchymal transition of gastric cancer through demethylation in the promoter of Cul4A in male. J. Cell Biochem. 2019;120:12247-12258.
Not in our library10.1002/jcb.27308
547.
Komura K, et al. Resistance to docetaxel in prostate cancer is associated with androgen receptor activation and loss of KDM5D expression. Proc. Natl Acad. Sci. USA. 2016;113:6259-6264.
Not in our library10.1073/pnas.1600420113
548.
Zhu XX, et al. Jarid2 is essential for the maintenance of tumor initiating cells in bladder cancer. Oncotarget. 2017;8:24483-24490.
Not in our library10.18632/oncotarget.15522
549.
Tange S, et al. JARID2 is involved in transforming growth factor-beta-induced epithelial-mesenchymal transition of lung and colon cancer cell lines. PLoS ONE. 2014;9:e115684.
Not in our library10.1371/journal.pone.0115684
550.
Paccez JD, et al. Dihydroartemisinin inhibits prostate cancer via JARID2/miR-7/miR-34a-dependent downregulation of Axl. Oncogenesis. 2019;8:14.
Not in our library10.1038/s41389-019-0122-6
551.
Ler LD, et al. Loss of tumor suppressor KDM6A amplifies PRC2-regulated transcriptional repression in bladder cancer and can be targeted through inhibition of EZH2. Sci. Transl. Med. 2017;9:eaai8312.
Not in our library10.1126/scitranslmed.aai8312
552.
Andricovich J, et al. Loss of KDM6A activates super-enhancers to induce gender-specific squamous-like pancreatic cancer and confers sensitivity to BET inhibitors. Cancer Cell. 2018;33:512-526.e8.
Not in our library10.1016/j.ccell.2018.02.003
553.
Mo J, et al. Multifunctional nanoparticles for co-delivery of paclitaxel and carboplatin against ovarian cancer by inactivating the JMJD3-HER2 axis. Nanoscale. 2017;9:13142-13152.
Not in our library10.1039/c7nr04473a
554.
Wang W, et al. KDM6B counteracts EZH2-mediated suppression of IGFBP5 to confer resistance to PI3K/AKT inhibitor treatment in breast cancer. Mol. Cancer Ther. 2018;17:1973-1983.
Not in our library10.1158/1535-7163.mct-17-0802
555.
Ahn J, et al. Target sequencing and CRISPR/Cas editing reveal simultaneous loss of UTX and UTY in urothelial bladder cancer. Oncotarget. 2016;7:63252-63260.
Not in our library10.18632/oncotarget.11207
556.
Lee KH, et al. Histone demethylase KDM7A controls androgen receptor activity and tumor growth in prostate cancer. Int J. Cancer. 2018;143:2849-2861.
Not in our library10.1002/ijc.31843
557.
Björkman M, et al. Systematic knockdown of epigenetic enzymes identifies a novel histone demethylase PHF8 overexpressed in prostate cancer with an impact on cell proliferation, migration and invasion. Oncogene. 2012;31:3444-3456.
Not in our library10.1038/onc.2011.512
558.
Li S, et al. Histone demethylase PHF8 promotes progression and metastasis of gastric cancer. Am. J. Cancer Res. 2017;7:448-461.
Not in our library10.1158/0008-5472.can-16-2350
559.
Shen Y, Pan X, Zhao H. The histone demethylase PHF8 is an oncogenic protein in human non-small cell lung cancer. Biochem Biophys. Res. Commun. 2014;451:119-125.
Not in our library10.1016/j.bbrc.2014.07.076
560.
Zhao Z, et al. Overexpression of histone demethylase JMJD5 promotes metastasis and indicates a poor prognosis in breast cancer. Int J. Clin. Exp. Pathol. 2015;8:10325-10334.
Not in our library
561.
Hsia DA, et al. KDM8, a H3K36me2 histone demethylase that acts in the cyclin A1 coding region to regulate cancer cell proliferation. Proc. Natl Acad. Sci. USA. 2010;107:9671-9676.
Not in our library10.1073/pnas.1000401107
562.
Abu-Jamous B, Buffa FM, Harris AL, Nandi AK. In vitro downregulated hypoxia transcriptome is associated with poor prognosis in breast cancer. Mol. Cancer. 2017;16:105.
Not in our library10.1186/s12943-017-0673-0
563.
Liu Y, et al. JMJD6 regulates histone H2A.X phosphorylation and promotes autophagy in triple-negative breast cancer cells via a novel tyrosine kinase activity. Oncogene. 2019;38:980-997.
Not in our library10.1038/s41388-018-0466-y
564.
Lee YF, et al. JMJD6 is a driver of cellular proliferation and motility and a marker of poor prognosis in breast cancer. Breast Cancer Res. 2012;14:R85.
Not in our library10.1186/bcr3200
565.
Zhang Z, Yang Y, Zhang X. MiR-770 inhibits tumorigenesis and EMT by targeting JMJD6 and regulating WNT/β-catenin pathway in non-small cell lung cancer. Life Sci. 2017;188:163-171.
Not in our library10.1016/j.lfs.2017.09.002
566.
Chang X, et al. Investigating the pathogenic role of PADI4 in oesophageal cancer. Int J. Biol. Sci. 2011;7:769-781.
Not in our library10.7150/ijbs.7.769
567.
Calin GA, et al. Human microRNA genes are frequently located at fragile sites and genomic regions involved in cancers. Proc. Natl Acad. Sci. USA. 2004;101:2999-3004.
Not in our library10.1073/pnas.0307323101
568.
Kasinski AL, Slack FJ. Epigenetics and genetics. MicroRNAs en route to the clinic: progress in validating and targeting microRNAs for cancer therapy. Nat. Rev. Cancer. 2011;11:849-864.
Not in our library10.1038/nrc3166
569.
Garcia AI, et al. Down-regulation of BRCA1 expression by miR-146a and miR-146b-5p in triple negative sporadic breast cancers. EMBO Mol. Med. 2011;3:279-290.
Not in our library10.1002/emmm.201100136
570.
Friedman JM, et al. The putative tumor suppressor microRNA-101 modulates the cancer epigenome by repressing the polycomb group protein EZH2. Cancer Res. 2009;69:2623-2629.
Not in our library10.1158/0008-5472.can-08-3114
571.
Varambally S, et al. Genomic loss of microRNA-101 leads to overexpression of histone methyltransferase EZH2 in cancer. Science. 2008;322:1695-1699.
Not in our library10.1126/science.1165395
572.
Fabbri M, et al. MicroRNA-29 family reverts aberrant methylation in lung cancer by targeting DNA methyltransferases 3 A and 3B. Proc. Natl Acad. Sci. USA. 2007;104:15805-15810.
Not in our library10.1073/pnas.0707628104
573.
Yang L, et al. lncRNA-dependent mechanisms of androgen-receptor-regulated gene activation programs. Nature. 2013;500:598-602.
Not in our library10.1038/nature12451
574.
He Y, et al. Long noncoding RNAs: Novel insights into hepatocelluar carcinoma. Cancer Lett. 2014;344:20-27.
Not in our library10.1016/j.canlet.2013.10.021
575.
Kim K, et al. HOTAIR is a negative prognostic factor and exhibits pro-oncogenic activity in pancreatic cancer. Oncogene. 2013;32:1616-1625.
Not in our library10.1038/onc.2012.193
576.
Kogo R, et al. Long noncoding RNA HOTAIR regulates polycomb-dependent chromatin modification and is associated with poor prognosis in colorectal cancers. Cancer Res. 2011;71:6320-6326.
Not in our library10.1158/0008-5472.can-11-1021
577.
Nakagawa T, et al. Large noncoding RNA HOTAIR enhances aggressive biological behavior and is associated with short disease-free survival in human non-small cell lung cancer. Biochem Biophys. Res. Commun. 2013;436:319-324.
Not in our library10.1016/j.bbrc.2013.05.101
578.
Gius D, et al. Distinct effects on gene expression of chemical and genetic manipulation of the cancer epigenome revealed by a multimodality approach. Cancer Cell. 2004;6:361-371.
Not in our library10.1016/j.ccr.2004.08.029
579.
Li E, Bestor TH, Jaenisch R. Targeted mutation of the DNA methyltransferase gene results in embryonic lethality. Cell. 1992;69:915-926.
Not in our library10.1016/0092-8674(92)90611-f
580.
Jackson-Grusby L, et al. Loss of genomic methylation causes p53-dependent apoptosis and epigenetic deregulation. Nat. Genet. 2001;27:31-39.
Not in our library10.1038/83730
581.
Carr BI, et al. The tumorigenicity of 5-azacytidine in the male Fischer rat. Carcinogenesis. 1984;5:1583-1590.
Not in our library10.1093/carcin/5.12.1583
582.
Christman JK. 5-Azacytidine and 5-aza-2′-deoxycytidine as inhibitors of DNA methylation: mechanistic studies and their implications for cancer therapy. Oncogene. 2002;21:5483-5495.
Not in our library10.1038/sj.onc.1205699
583.
Jones PA, Laird PW. Cancer epigenetics comes of age. Nat. Genet. 1999;21:163-167.
Not in our library10.1038/5947
584.
Rosen MB, House HS, Francis BM, Chernoff N. Teratogenicity of 5-azacytidine in the Sprague-Dawley rat. J. Toxicol. Environ. Health. 1990;29:201-210.
Not in our library10.1080/15287399009531383
585.
Doerksen T, Trasler JM. Developmental exposure of male germ cells to 5-azacytidine results in abnormal preimplantation development in rats. Biol. Reprod. 1996;55:1155-1162.
Not in our library10.1095/biolreprod55.5.1155
586.
Chik F, Szyf M. Effects of specific DNMT gene depletion on cancer cell transformation and breast cancer cell invasion; toward selective DNMT inhibitors. Carcinogenesis. 2011;32:224-232.
Not in our library10.1093/carcin/bgq221
587.
Issa JP, et al. Phase 1 study of low-dose prolonged exposure schedules of the hypomethylating agent 5-aza-2′-deoxycytidine (decitabine) in hematopoietic malignancies. Blood. 2004;103:1635-1640.
Not in our library10.1182/blood-2003-03-0687
588.
Beisler JA, Abbasi MM, Kelley JA, Driscoll JS. Synthesis and antitumor activity of dihydro-5-azacytidine, a hydrolytically stable analogue of 5-azacytidine. J. Med. Chem. 1977;20:806-812.
Not in our library10.1021/jm00216a013
589.
Presant CA, Coulter D, Valeriote F, Vietti TJ. Contrasting cytotoxicity kinetics of 5-azacytidine and dihydro-5-azacytidine hydrochloride in L1210 leukemia in mice. J. Natl Cancer Inst. 1981;66:1151-1154.
Not in our library10.1093/jnci/66.6.1151
590.
Curt GA, et al. A phase I and pharmacokinetic study of dihydro-5-azacytidine (NSC 264880). Cancer Res. 1985;45:3359-3363.
Not in our library
591.
Cheng JC, et al. Inhibition of DNA methylation and reactivation of silenced genes by zebularine. J. Natl Cancer Inst. 2003;95:399-409.
Not in our library10.1093/jnci/95.5.399
592.
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593.
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596.
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599.
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619.
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626.
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628.
Singh TR, Shankar S, Srivastava RK. HDAC inhibitors enhance the apoptosis-inducing potential of TRAIL in breast carcinoma. Oncogene. 2005;24:4609-4623.
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629.
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A class of hybrid polar inducers of transformed cell differentiation inhibits histone deacetylases · 1998
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631.
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633.
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636.
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643.
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659.
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664.
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Not in our library10.1038/nature13688

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