Submitted Successfully!
Thank you for your contribution! You can also upload a video entry or images related to this topic.
Ver. Summary Created by Modification Content Size Created at Operation
1 -- 3834 2022-11-14 05:33:54 |
2 format correct + 5 word(s) 3839 2022-11-15 07:51:22 | |
3 format correct Meta information modification 3839 2022-11-16 07:21:02 |

Video Upload Options

Do you have a full video?


Are you sure to Delete?
If you have any further questions, please contact Encyclopedia Editorial Office.
Zhu, G.;  Zheng, X.;  Wang, Z.;  Xu, X. Acetylation. Encyclopedia. Available online: (accessed on 05 December 2023).
Zhu G,  Zheng X,  Wang Z,  Xu X. Acetylation. Encyclopedia. Available at: Accessed December 05, 2023.
Zhu, Guangrong, Xiangyang Zheng, Zhifeng Wang, Xingzhi Xu. "Acetylation" Encyclopedia, (accessed December 05, 2023).
Zhu, G.,  Zheng, X.,  Wang, Z., & Xu, X.(2022, November 14). Acetylation. In Encyclopedia.
Zhu, Guangrong, et al. "Acetylation." Encyclopedia. Web. 14 November, 2022.

Acetylation, also known as acetylation or acetylation, refers to the chemical reaction of adding an acetyl functional group to an organic compound. Conversely, the reaction in which the acetyl group is removed is called deacetylation or deacetylation. Acetylation of proteins is a post-translational modification.

DNA damage response post-translational modification lipid metabolites acetylation

1. Acetylation of Histones and Non-Histone Proteins

In eukaryotic cells, chromatin refers to a linear, complex structure composed of histone, non-histone, DNA, and a small amount of RNA and is located in the nucleus. Histones are alkaline proteins that bind to DNA in eukaryotic cells. They are composed of four core histone families (H2A, H2B, H3, H4) and a linker family (H1). Core histones are mainly responsible for binding to DNA to form nucleosomes, which represent the structural units of chromatin, while linker histones are responsible for binding with nucleosomes to form the next chromatin unit. Thus, histones play an important role in maintaining chromatin structure. Non-histone chromatin refers to other proteins that can bind to chromatin, including chromatin structural proteins, enzymes, and a small number of regulatory proteins. Although non-histones are less common in chromatin than histones, there a wide range of types, and they serve complex functions, such as regulating gene expression and maintaining chromatin structure. There are more basic amino acids in histones than acidic amino acids, but the ratio of basic to acidic amino acids in non-histones varies across situations and contexts.
Histone acetylation is a dynamic and reversible protein protein post-translational modification (PTM) and may be the most well-understood. Histone acetyltransferases (HATs, also referred to as lysine acetyltransferases) and histone deacetylases (HDACs, also known as lysine deacetylases) catalyze the addition or removal of acetyl groups, respectively, to lysine residues on both histone and non-histone proteins [1]. Acetylation occurs on amino-terminal proteins and on the O-junctions of serine and threonine, acetylation refers only to N ε-lysine acetylation (Kac) (defined as the deposition of an acetyl group onto the epsilon amino group of lysine; Figure 1) [2]. It was found that, by stable isotope tracking and acetylation proteomic analysis, more than 90% of acetylation modification on histone lysine is derived from the carbon of fatty acids [3]. Acetyl-CoA in mitochondria from fatty acid β-oxidation is converted to citrate, an intermediate of the tricarboxylic acid cycle. Citrate is exported out of mitochondria by citate transporter and subsequently cleaved by ATP-citrate lyase (ACLY) into acetyl-CoA [4].
Figure 1. Acetylation modification mechanism.
Acetylation of histones by HATs relaxes the structure of chromatin and increases DNA accessibility, which results in increased gene transcription. The effect of HDACs is the opposite, with deacetylation making the structure of chromatin more compact and inhibiting transcription. The histone code hypothesis suggests that histone modification can be explained by chromatin-binding proteins called “readers,” which distinguish between modified and unmodified nucleosomes and then decide on DNA transcription or other related downstream events [5]. At present, known domains with the ability to recognize or read acetylated lysine residues include YAF9, ENL, AF9, Taf14, and Sas5 YEATS domains, and tandem homologous domains (PHDs, also known as double PHD finger (DPF) domains) in plants [6]. In addition to histones, mass spectrometry has also revealed that acetylation occurs on non-histone proteins, some of which participate in protein folding, protein degradation, and chromatin structure adjustment [7].
The exact number of HATs in the human proteome remains unknown. However, the histone HATs that have been identified thus far include the P160, P300/CBP, TAFII230, MYST, GNAT, and PCAF families, with the current classification system primarily being based on homology with the original enzyme sequence found in yeast. In addition, a total of 13 non-histone HATs have also been identified, and these are roughly divided into three families: GCN5, P300, and MYST. These include histone acetyltransferase 1 (HAT1, also known as KAT1), α-tubulin N-acetyltransferase 1 (TAT1, also known as ATAT1) [8], and establishment of cohesion 1 homology 1 (ESCO1) and ESCO2, which all serve different functions. At present, non-histone HAT specificity is thought to be determined by the accessibility of lysine in their substrate proteins, specific subcellular localization, and interacting proteins. For example, with the exception of TAT1, HATs are primarily located in the cell nuclei. Most HAT substrates do not overlap, but some functionally similar HATs can acetylate the same sites: for example, CREB-binding proteins (CBP, also known as KAT3A) and p300 (also known as KAT3B) both acetylate histone H3Lys18 (H3K18) and H3K27.
HDACs, meanwhile, are divided into four categories based on their sequence similarity and degree of phylogenetic conservation. The class I HDACs include HDAC1, 2, 3, and 8; the class II HDACs include HDAC4, 5, 6, 7, 9, and 10 (being further subdivided into class IIa and IIb); the class III HDACs are known as the sirtuins and include SIRT1-7; while the fourth category currently includes HDAC11 only [9]. With the exception of class III HDACS, which are NAD+-dependent, the other HDAC classes are all zinc-dependent. Zinc-dependent HDACs contains a deacetylase domain which is highly conserved and are often referred to as classical HDACs. While class I and IV HDACs are localized in the nucleus, class IIb HDACs are distributed in the cytoplasm. Class IIa HDACs are mainly distributed in the nucleus but are exported to the cytoplasm when activated. The sirtuins are distributed in different locations in the cell as follows: cytoplasm (SIRT2), mitochondria (SIRT3, SIRT4, and SIRT5), nucleus (SIRT1 and SIRT6), and nucleolus (SIRT7).
It is worth noting that most non-histone deacetylases have limited or no deacetylase activity or primarily perform other types of acylation. For example, SIRT4 [10] removes the acyl group from hydroxymethyl glutaryl lysine, SIRT5 [11] acts as a demalonylase and a deglutarylase, and SIRT6 acts as a long-chain fatty acid deacylase. Class IIa HDACs lack obvious catalytic activity, primarily due to the alterations in the conserved amino acids found in their catalytic pocket. Recent studies have found that lymphoid augmenter factor 1 (LEF1) [12] and T cell-specific transcription factor 1 (TCF1, also known as Tcf7) [13] also exert HDAC activity and that these two transcription factors are involved in the regulation of the WNT signaling pathway [14]. The overall sequences of LEF1 and TCF1 are very different from HDAC8, but their functions are similar.
Recent studies have shown that acetylation can also occur through non-enzymatic mechanisms [15]. For example, lysine can be acylated by acyl-CoAs formed by the breakdown of fatty acids, and this mostly occurs through non-enzymatic mechanisms. Interestingly, some acyl-CoAs, such as glutaryl coenzyme A and succinyl coenzyme A, are derived from the carboxy cycle process but are more active than acetyl-CoA. The primary mechanism underpinning these modifications is acyl-CoA carboxyl group-induced intramolecular nucleophilic attack on CoA thioester bonds, which results in the formation of cyclic anhydride. This is more active than acyl-CoA and can produce non-enzymatic modifications more efficiently. Non-enzymatic acylation is thought to be affected by the cellular concentration of acyl-CoAs, the reactivity of acyl-CoA, local pH levels, and the number of lysine residues in proteins. These factors may differ between cell and tissue types. However, the specific regulatory mechanisms of this non-enzymatic modification remain unclear and require further investigation.

2. Acetylation in the DDR

2.1. Histone Acetylation in the DDR

Histone acetylation is well known to lead to changes in the structure of chromatin. Mechanistically, this occurs in two ways: on the one hand, the positive charge of lysine residues can be neutralized after acetylation, resulting in the weakening of the interaction between histone and DNA skeleton and the promotion of chromatin decompaction. This exposes more sites and enhances the accessibility of nucleosomal DNA. On the other hand, related chromatin remodeling complexes, such as SWI/SNF complexes, can also be recruited to the chromatin region to regulate chromatin structure. It was found that, in response to ionizing radiation (IR), nuclear ACLY is phosphorylated at S455 by ataxia telangiectasia mutated (ATM) and facilitates histone acetylation at double-strand breaks (DSBs), promoting HR-mediated repair by enabling BRCA1 recruitment while impairing 53BP1 recruitment [16]. This direct evidence demonstrates the link between histone acetylation and DNA damage response (DDR). Other studies have shown that the acetylation modification of histone H1K85 can mediate chromatin changes under the dynamic regulation of acetylase PCAF and deacetylase HDAC1 in response to DNA damage, thus ensuring genome stability [17]. However, compared with H1 histone, H2 histone acetylation has been studied more extensively in terms of DDR. When cells are stimulated by ionizing radiation (IR), H2AX lysine 36 (H2AX K36ac) can be acetylated by acetyltransferase CBP/P300 to recruit corresponding DNA repair proteins to DNA damage sites [18]. Furthermore, under ionizing radiation-induced stimulation, the DNA PKCs BRD domain can specifically recognize H2AX acetylated lysine 5 (K5ac), which can lead to the determination of cell fate via γH2AX [19]. Moreover, under IR stimulation, knockdown of the tumor suppressor gene ZNF668, known to be involved in breast cancer, will weaken the interaction between Tip60 and H2AX. This leads to reduced hyperacetylation of histone H2AX and the prevention of chromatin relaxation, resulting in a reduced recruitment of repair proteins to DNA damage sites, defective homologous recombination (HR) repair, and reduced cell survival rates. These examples fully illustrate the importance of H2 histone acetylation to the DDR and, thus, to cancer [20].
Histone H3 acetylation also serves an important regulatory role in the DDR. At the location of DNA damage, the histone deacetylases HDAC1 and HDAC2 maintain low H3K56 acetylation levels to ensure that the damaged DNA is repaired. After the repair is completed, the proteasome will degrade the acetylated core histones, and the newly formed core histones will be assembled into nucleosomes. This represents the completion of the process of coping with DNA damage and repair [21]. PhD bromo tandem domain containing trimer motif 66 (TRIM66) has also been found to recognize unmodified H3R2, H3K4, and acetylated H3K56, while TRIM66 can recruit SIRT6 to deacetylate H3K56, thereby initiating the DDR and maintaining genome stability [22]. In addition, after 24 h of stimulation of HaCaT cells with sodium arsenite (NaAsO2), arsenic has been reported to reduce histone H3K18 acetylation levels, affect the expression of xeroderma pigmentosa-related proteins (XPA, XPD, and XPF nucleotide excision repair (NER)-related genes), and further aggravate DNA damage. However, the use of histone deacetylase inhibitor trichostatin A (TSA) can inhibit the deacetylation of H3K18 in the promoter region of XPA, XPD, and XPF, increase the acetylation of H3K18, and promote the transcriptional expression of nucleotide excision repair (NER)-related genes. To a certain extent, it can inhibit arsenic-induced DNA damage [23].
In addition to H1, H2, and H3 acetylation, some studies also have been conducted on histone H4 acetylation. After doxorubicin treatment, cells in G0/G1 phase experience DNA damage but it can also promote the activation of chromatin kinase VRK1. VRK1 directly interacts with Tip60 and phosphorylates it, resulting in increased histone H4K16 acetylation, a marker of local chromatin relaxation. Inhibition of Tip60 expression by siRNA or its kinase inhibitor MG149 inhibited H4K16 acetylation, indicating VRK1-mediated phosphorylation of Tip60 increases its enzymatic activity. This work suggests that the dynamic remodeling of chromatin is closely related to the epigenetic modification of histone [24]. In addition, males absent of the first (MOF) proteins can regulate the level of H4K16 acetylation. MOF is responsible for maintaining sufficient levels of H4K16 acetylation in cells, which facilitates the generation of chromatin structures conducive to DNA repair. When MOF is absent in cells, the acetylation level of H4K16 is reduced and ultimately, the recruitment and cancellation of the corresponding signal proteins at the DNA damage site is impaired [25]. Furthermore, after DNA double-strand breaks induced by the HO endonuclease system, if the acetylation sites of newly synthesized histone H4 are mutated, the reassembly of chromatin structure is inhibited. Interestingly, the newly synthesized histone H4 acetylation mutation site changes, resulting in phosphorylated H2A (γ-H2AX) levels significantly decreasing around DSBs, indicating the critical role of chromatin assembly in DNA damage signaling [26].
In summary, Kac modification of histone tails can lead to the relaxation of chromatin structure, which is more conducive to the completion of DNA repair after DNA damage. Moreover, after DNA repair is completed, histones undergo deacetylation, and the chromatin structure becomes compact. Thus, this dynamic regulation by histone acetylation is critical to the maintenance of genome stability.

2.2. Non-Histone Acetylation in the DDR

Non-histone acetylation has also been found to participate in the DDR process. When inducing DNA damage in human cells, the acetylation of lysine 382 and phosphorylation of serine 392 in p53, a key DDR factor, can significantly enhance the interaction between p53 and MDC1 and promote the recruitment of these two proteins to DNA damage sites [27]. In response to DNA damage, N-acetyltransferase 10, NAT10 (also known as HALP), a member of the GNAT family, translocates to the nucleoplasm, promoting p53 acetylation at K120 with its acetyltransferase activity and proteosome-mediated degradation of MDM2 with its intrinsic E3 ligase activity, ultimately resulting in stabilizing p53 and p53-mediated cell cycle arrest and apoptosis [28]. Tip60-mediated acetylation, the DDR core kinase ATM also activates its kinase activity and subsequent checkpoint signaling upon DNA damage, while Tip60 inactivation sensitizes cells to ionizing radiation [29].
Werner syndrome is a rare autosomal recessive disease caused by mutations of the WRN gene. When the lysines K1127 and K1117 of WRN are mutated to arginine, cells may become sensitive to DNA-damaging agents such as mitomycin C and etoposide, indicating defective DNA repair. In fact, these two sites are critical for the recruitment of WRN to DNA damage sites [30].
Furthermore, the acetylation of proteins has also been found to be involved in the regulation of base excision repair (BER). For example, acetylation of depuridine/depyrimidine endonuclease 1 (APE1; also known as APEX1), an important regulator of BER, inhibits its interaction with XRCC1 when DNA damage occurs; resulting in decreased APE1 activity. However, SIRT1 can deacetylate APE1, recovering its function [31]. Additional acetylated non-histone proteins involved in the DDR are summarized in Table 1.

2.3. Roles of HATs and HDACs/SIRTs in DDR

Many HATs-mediated acetylations of histone and non-histone proteins directly or indirectly modulate DDR. KAT8 (hMOF), as a member of the histone acetyltransferase MYST family, can respond to DNA damage by catalyzing the acetylation of H4 at K16 (H4K16) and p53 at K120 [32][33]. KAT5 (TIP60) is also a member of the histone acetyltransferase MYST family [34], and its regulatory role in DNA damage signaling [35] has been reported. It was found that TIP60 can regulate the acetylation of a variety of histones (H2, H3, and H4) and non-histones (p53 [36] and ATM [37]). For example, TIP60 can acetylate the K15 position of H2A in response to DNA damage [38][39]. It has also been shown that Tip60 regulates the acetylation of H4 by forming a complex with transformation/transfer domain-associated protein (Trrap) and then promotes DNA damage repair by HR [40]. In addition, histone acetyltransferases KAT3A (CBP) and KAT3B (p300) are structurally similar and participate in the regulation of many functions in cells. In vitro enzyme activity test showed that CBP/p300 could acetylate all acetylation residue sites of histone H2A and H2B, among which K14, K18, and K56 of H3 and K5 and K8 of H4 were preferentially oxidized [41]. Moreover, CBP can also regulate the acetylation of non-histones in cancer. In human colon cancer cells, CBP can mediate the acetylation of K358 of DOT1L, which is positively correlated with the staging of colon cancer [42]. It was found that in vitro, acetyltransferase GCN5 can form a complex with its chaperone PCAF to acetylate multiple lysine residue sites on histone H3, such as H3K9, H3K14, H3K18, and H3K23 [43][44][45][46]. In addition, PCAF can acetylate H1K85 in response to DNA damage [47].
On the other hand, HDACs/SIRTs-mediated protein deacetylation also plays an important role in DDR. In mammals, SIRT1-7 has different subcellular localization, functions, and substrates and was initially identified as histone and non-histone protein deacetylases [48][49]. It was found that sirtuins can mediate the specific deacetylation of histone lysine residues to facilitate DNA repair. In mammals, H3K56 will undergo hyperacetylation upon inhibition of SIRT1 expression, leading to the instability of the S phase genome [50]. It has also been reported thatSIRT3, which is mainly located in mitochondria, will be transported to the nucleus to regulate the deacetylation of H4K16ac in response to DNA damage [51][52][53]. SIRT6 is an intranuclear deacetylase which plays an important role in regulating DDR signal and genome stability. Its histone substrates include H3K9, H3K18, and H3K56. H3K9 deacetylation mediated by SIRT6 can protect telomeres in mammalian cells. On the contrary, the lack of SIRT6 will lead to chromosome fusion due to telomere dysfunction [54]. Similarly, in the nucleus, SIRT7 can catalyze the deacetylation of H3K18ac at the late response to IR [55]. In addition, HDAC1/2-mediated deacetylation of H3K56 and H4K16 also plays an important role in chromatin regulation [56][57]. It was found that histone deacetylase was rapidly recruited to the DNA damage site, leading to histone deacetylation, thereby promoting non-homologous end joining (NHEJ) repair [56]. Several studies have found that the levels of H3K56ac, H4K16ac, and H4K91ac will increase after HDAC1 expression is inhibited, which is related to the decreased cell survival rate after treatment with DNA damaging reagents [56][57][58][59].
In conclusion, the dynamic regulation of both histone and non-histone acetylation and deacetylation serves an important function in the repair of DNA damage.
Table 1. Lipid metabolite associated PTMs in the DDR.

3. Acetylation in Cancer

3.1. HATs and Cancer

HATs are well known to be involved in tumorigenesis and cancer progression, with HAT activity being altered either by gene mutations or viral oncogenes in both blood and solid cancers. For example, the interaction between adenovirus SV40T antigen protein E1A and the co-activators p300 and CBP play a key role in cell transformation [74]. These HATs are then redistributed to the promoter regions of certain genes to promote cell growth, differentiation, and the transcriptional activation of specific genes [75]. Ambiguous p300 mutations can be found in solid stomach, rectum, breast, and prostate tumors [76][77][78].
Tip60 is another HAT that is closely associated with tumorigenesis [79][80][81] and may be involved in the regulation of DNA repair and the transcriptional activation of p53 and Myc [35][82]. Decreased expression of Tip60 results in low p53 acetylation levels and incomplete apoptosis signaling, which indicates transformation to a malignant tumor [83]. As a tumor suppressor protein, single allele deletion of human Tip60 is often found in head and neck tumors, breast cancer, and lymphoma [84]. In addition, Tip60 has also been found to inhibit Myc-mediated lymphoma formation in B-cell lymphoma [85].

3.2. HDACs and Cancer

HDACs function in the opposite manner to HATs, regulating transcription by removing acetyl groups from lysine residues of histone tails and other non-histone substrates. Thus, it stands to reason that they would also be involved in cancer. Functional experiments have indicated that type I HDACs are mainly responsible for regulating cell proliferation and apoptosis, while type II HDACs regulate cell metastasis and angiogenesis. For example, inhibiting HDAC1 and HDAC2 in vitro can inhibit colon cancer cell proliferation [86][87]. However, the inhibition of HDAC3 suppresses the proliferation of colon cancer cells more substantially [88]. In addition, inhibition of HDAC2 and HDAC3 is known to induce DDR and apoptosis after DNA damage.
Type II and IV HDACs are primarily localized in the cytoplasm and are mainly responsible for the deacetylation of non-histone proteins. Previous studies have shown that inhibiting HDAC4 reduces colon cancer cell proliferation and induces apoptosis [89][90][91]. Furthermore, while the inhibition of HDAC7 in endothelial cells does not affect cell growth and survival, it does inhibit cell metastasis and the formation of capillary-like structures in cancer [92][93]. Type II HDACs are mainly responsible for regulating angiogenesis, and inhibition of HDAC6 and HDAC10 results in decreased VEGFR1 and two expressions [94].
Type III HDACs, also known as the sirtuins, share no sequence homology with other deacetylases. However, they may also be involved in regulating the occurrence and development of tumors. Sirtuins induce the deacetylation of a range of protein substrates, including histones, but also mediate ADP ribosylation. Furthermore, overexpression of SIRT1, 2, 3, and 7 have been identified in many types of cancer [95][96][97]. For example, overexpression of SIRT1 is known to prevent apoptosis by regulating histone deacetylation, promoter methylation, and histone methylation and inhibiting the transcription of tumor suppressor genes. This ultimately promotes cancer cell growth by preventing cell senescence and differentiation, as well as the formation of tumor blood vessels by promoting the growth of endothelial cells and preventing their aging [98]. Interestingly, the expression of SIRT2 is absent in human glioma cells, and re-expression of SIRT2 can reduce the ability of colony-stimulating factor formation of cells [99]. This suggests that, in some cases, Sirtuins also serve as tumor suppressors. At present, whether Sirtuins function as oncogenes or as tumor suppressors remains controversial. However, it is clear that altered HDAC function plays a corresponding role to that of HATs in the process of tumorigenesis and development.

3.3. Summary

The extensive research conducted thus far on protein acetylation and the fact that abnormal acetylation is closely associated with cancer has laid the foundation for the discovery of many novel epigenetic drug targets. Certain HDAC inhibitors have been approved for cancer treatment, including romidepsin, panobinostat, and belinostat, while more are being tested in clinical trials. However, most research exploring the potential for HAT inhibitors to treat cancer is yet to enter clinical trials, leaving this treatment avenue in its primary stages. However, the identification of new HAT subtypes and improved characterization of their roles and functions have provided more potential treatment strategies. For example, curcumin, as a natural KATi, can inhibit the activity of p300/CBP and so suppress the proliferation of a variety of cancer cells, thereby achieving anti-inflammatory and anti-tumor effects [100]. Moreover, as a small molecule derived from anacardic acid, MG153 also acts as a potential p300/PCAF inhibitor and can suppress the proliferation of BCR-ABL-expressing cells, induce apoptosis, and resist DNA damage [101]. In addition, L002, a small molecule inhibitor of p300, has been shown to inhibit p300, PCAF, and GCN5 activity in leukemia, lymphoma, and breast cancer cell lines [102]. In conclusion, further research on both HDAC and HAT inhibitors will likely prove very fruitful when developing novel treatments for cancer [103].


  1. Slaughter, M.J.; Shanle, E.K.; Khan, A.; Chua, K.F.; Hong, T.; Boxer, L.D.; Allis, C.D.; Josefowicz, S.Z.; Garcia, B.A.; Rothbart, S.B.; et al. HDAC inhibition results in widespread alteration of the histone acetylation landscape and BRD4 targeting to gene bodies. Cell Rep. 2021, 34, 108638.
  2. Luu, J.; Carabetta, V.J. Contribution of N(ε)-lysine Acetylation towards Regulation of Bacterial Pathogenesis. mSystems 2021, 6, e42221.
  3. Mcdonnell, E.; Crown, S.B.; Fox, D.B.; Kitir, B.; Ilkayeva, O.R.; Olsen, C.A.; Grimsrud, P.A.; Hirschey, M.D. Lipids Reprogram Metabolism to Become a Major Carbon Source for Histone Acetylation. Cell Rep. 2016, 17, 1463–1472.
  4. Sebastián, C.; Mostoslavsky, R. The Various Metabolic Sources of Histone Acetylation. Trends Endocrinol. Metab. 2017, 28, 85–87.
  5. Zhang, Y.; Sun, Z.; Jia, J.; Du, T.; Zhang, N.; Tang, Y.; Fang, Y.; Fang, D. Overview of Histone Modification. Adv. Exp. Med. Biol. 2021, 1283, 1–16.
  6. Gu, W.; Cheng, Y.; Wang, S.; Sun, T.; Li, Z. PHD Finger Protein 19 Promotes Cardiac Hypertrophy via Epigenetically Regulating SIRT2. Cardiovasc. Toxicol. 2021, 21, 451–461.
  7. Narita, T.; Weinert, B.T.; Choudhary, C. Functions and mechanisms of non-histone protein acetylation. Nat. Rev. Mol. Cell Biol. 2019, 20, 156–174.
  8. Li, L.; Jayabal, S.; Ghorbani, M.; Legault, L.M.; Mcgraw, S.; Watt, A.J.; Yang, X.J. ATAT1 regulates forebrain development and stress-induced tubulin hyperacetylation. Cell. Mol. Life Sci. 2019, 76, 3621–3640.
  9. Sivalingam, K.; Doke, M.; Khan, M.A.; Samikkannu, T. Influence of psychostimulants and opioids on epigenetic modification of class III histone deacetylase (HDAC)-sirtuins in glial cells. Sci. Rep. 2021, 11, 21335.
  10. Min, Z.; Gao, J.; Yu, Y. The Roles of Mitochondrial SIRT4 in Cellular Metabolism. Front. Endocrinol. 2018, 9, 783.
  11. Kumar, S.; Lombard, D.B. Functions of the sirtuin deacylase SIRT5 in normal physiology and pathobiology. Crit. Rev. Biochem. Mol. Biol. 2018, 53, 311–334.
  12. Xing, S.; Li, F.; Zeng, Z.; Zhao, Y.; Yu, S.; Shan, Q.; Li, Y.; Phillips, F.C.; Maina, P.K.; Qi, H.H.; et al. Tcf1 and Lef1 transcription factors establish CD8(+) T cell identity through intrinsic HDAC activity. Nat. Immunol. 2016, 17, 695–703.
  13. Li, F.; Zhao, X.; Zhang, Y.; Shao, P.; Ma, X.; Paradee, W.J.; Liu, C.; Wang, J.; Xue, H.H. T(FH) cells depend on Tcf1-intrinsic HDAC activity to suppress CTLA4 and guard B-cell help function. Proc. Natl. Acad. Sci. USA 2021, 118.
  14. Kim, C.; Jin, J.; Weyand, C.M.; Goronzy, J.J. The Transcription Factor TCF1 in T Cell Differentiation and Aging. Int. J. Mol. Sci. 2020, 21, 6497.
  15. Carrico, C.; Meyer, J.G.; He, W.; Gibson, B.W.; Verdin, E. The Mitochondrial Acylome Emerges: Proteomics, Regulation by Sirtuins, and Metabolic and Disease Implications. Cell Metab. 2018, 27, 497–512.
  16. Sivanand, S.; Rhoades, S.; Jiang, Q.; Lee, J.V.; Benci, J.; Zhang, J.; Yuan, S.; Viney, I.; Zhao, S.; Carrer, A.; et al. Nuclear Acetyl-CoA Production by ACLY Promotes Homologous Recombination. Mol. Cell 2017, 67, 252–265.
  17. Li, Y.; Li, Z.; Dong, L.; Tang, M.; Zhang, P.; Zhang, C.; Cao, Z.; Zhu, Q.; Chen, Y.; Wang, H.; et al. Histone H1 acetylation at lysine 85 regulates chromatin condensation and genome stability upon DNA damage. Nucleic Acids Res. 2018, 46, 7716–7730.
  18. Jiang, X.; Xu, Y.; Price, B.D. Acetylation of H2AX on lysine 36 plays a key role in the DNA double-strand break repair pathway. FEBS Lett. 2010, 584, 2926–2930.
  19. Wang, L.; Xie, L.; Ramachandran, S.; Lee, Y.; Yan, Z.; Zhou, L.; Krajewski, K.; Liu, F.; Zhu, C.; Chen, D.J.; et al. Non-canonical Bromodomain within DNA-PKcs Promotes DNA Damage Response and Radioresistance through Recognizing an IR-Induced Acetyl-Lysine on H2AX. Chem. Biol. 2015, 22, 849–861.
  20. Hu, R.; Wang, E.; Peng, G.; Dai, H.; Lin, S.Y. Zinc finger protein 668 interacts with Tip60 to promote H2AX acetylation after DNA damage. Cell Cycle 2013, 12, 2033–2041.
  21. Van, H.T.; Santos, M.A. Histone modifications and the DNA double-strand break response. Cell Cycle 2018, 17, 2399–2410.
  22. Chen, J.; Wang, Z.; Guo, X.; Li, F.; Wei, Q.; Chen, X.; Gong, D.; Xu, Y.; Chen, W.; Liu, Y.; et al. TRIM66 reads unmodified H3R2K4 and H3K56ac to respond to DNA damage in embryonic stem cells. Nat. Commun. 2019, 10, 4273.
  23. Zhang, A.L.; Chen, L.; Ma, L.; Ding, X.J.; Tang, S.F.; Zhang, A.H.; Li, J. Role of H3K18ac-regulated nucleotide excision repair-related genes in arsenic-induced DNA damage and repair of HaCaT cells. Hum. Exp. Toxicol. 2020, 39, 1168–1177.
  24. García-González, R.; Morejón-García, P.; Campillo-Marcos, I.; Salzano, M.; Lazo, P.A. VRK1 Phosphorylates Tip60/KAT5 and Is Required for H4K16 Acetylation in Response to DNA Damage. Cancers 2020, 12, 2986.
  25. Su, J.; Wang, F.; Cai, Y.; Jin, J. The Functional Analysis of Histone Acetyltransferase MOF in Tumorigenesis. Int. J. Mol. Sci. 2016, 17, 99.
  26. Ge, Z.; Nair, D.; Guan, X.; Rastogi, N.; Freitas, M.A.; Parthun, M.R. Sites of acetylation on newly synthesized histone H4 are required for chromatin assembly and DNA damage response signaling. Mol. Cell. Biol. 2013, 33, 3286–3298.
  27. Shahar, O.D.; Gabizon, R.; Feine, O.; Alhadeff, R.; Ganoth, A.; Argaman, L.; Shimshoni, E.; Friedler, A.; Goldberg, M. Acetylation of lysine 382 and phosphorylation of serine 392 in p53 modulate the interaction between p53 and MDC1 in vitro. PLoS ONE 2013, 8, e78472.
  28. Liu, X.; Tan, Y.; Zhang, C.; Zhang, Y.; Zhang, L.; Ren, P.; Deng, H.; Luo, J.; Ke, Y.; Du, X. NAT10 regulates p53 activation through acetylating p53 at K120 and ubiquitinating Mdm2. EMBO Rep. 2016, 17, 349–366.
  29. Sun, Y.; Jiang, X.; Chen, S.; Fernandes, N.; Price, B.D. A role for the Tip60 histone acetyltransferase in the acetylation and activation of ATM. Proc. Natl. Acad. Sci. USA 2005, 102, 13182–13187.
  30. Ghosh, D.; Bohr, V.A.; Karmakar, P. Acetylation of Werner protein at K1127 and K1117 is important for nuclear trafficking and DNA repair. DNA Repair 2019, 79, 22–31.
  31. Yamamori, T.; Dericco, J.; Naqvi, A.; Hoffman, T.A.; Mattagajasingh, I.; Kasuno, K.; Jung, S.B.; Kim, C.S.; Irani, K. SIRT1 deacetylates APE1 and regulates cellular base excision repair. Nucleic Acids Res. 2010, 38, 832–845.
  32. Dai, C.; Shi, D.; Gu, W. Negative regulation of the acetyltransferase TIP60-p53 interplay by UHRF1 (ubiquitin-like with PHD and RING finger domains 1). J. Biol. Chem. 2013, 288, 19581–19592.
  33. Li, X.; Corsa, C.A.; Pan, P.W.; Wu, L.; Ferguson, D.; Yu, X.; Min, J.; Dou, Y. MOF and H4 K16 acetylation play important roles in DNA damage repair by modulating recruitment of DNA damage repair protein Mdc1. Mol. Cell. Biol. 2010, 30, 5335–5347.
  34. Liu, R.; Gou, D.; Xiang, J.; Pan, X.; Gao, Q.; Zhou, P.; Liu, Y.; Hu, J.; Wang, K.; Tang, N. O-GlcNAc modified-TIP60/KAT5 is required for PCK1 deficiency-induced HCC metastasis. Oncogene 2021, 40, 6707–6719.
  35. Squatrito, M.; Gorrini, C.; Amati, B. Tip60 in DNA damage response and growth control: Many tricks in one HAT. Trends Cell Biol. 2006, 16, 433–442.
  36. Wang, P.; Bao, H.; Zhang, X.P.; Liu, F.; Wang, W. Regulation of Tip60-dependent p53 acetylation in cell fate decision. FEBS Lett. 2019, 593, 13–22.
  37. Jiang, X.; Sun, Y.; Chen, S.; Roy, K.; Price, B.D. The FATC domains of PIKK proteins are functionally equivalent and participate in the Tip60-dependent activation of DNA-PKcs and ATM. J. Biol. Chem. 2006, 281, 15741–15746.
  38. Jacquet, K.; Fradet-Turcotte, A.; Avvakumov, N.; Lambert, J.P.; Roques, C.; Pandita, R.K.; Paquet, E.; Herst, P.; Gingras, A.C.; Pandita, T.K.; et al. The TIP60 Complex Regulates Bivalent Chromatin Recognition by 53BP1 through Direct H4K20me Binding and H2AK15 Acetylation. Mol. Cell 2016, 62, 409–421.
  39. Ikura, T.; Tashiro, S.; Kakino, A.; Shima, H.; Jacob, N.; Amunugama, R.; Yoder, K.; Izumi, S.; Kuraoka, I.; Tanaka, K.; et al. DNA damage-dependent acetylation and ubiquitination of H2AX enhances chromatin dynamics. Mol. Cell. Biol. 2007, 27, 7028–7040.
  40. Murr, R.; Loizou, J.I.; Yang, Y.G.; Cuenin, C.; Li, H.; Wang, Z.Q.; Herceg, Z. Histone acetylation by Trrap-Tip60 modulates loading of repair proteins and repair of DNA double-strand breaks. Nat. Cell Biol. 2006, 8, 91–99.
  41. Schiltz, R.L.; Mizzen, C.A.; Vassilev, A.; Cook, R.G.; Allis, C.D.; Nakatani, Y. Overlapping but distinct patterns of histone acetylation by the human coactivators p300 and PCAF within nucleosomal substrates. J. Biol. Chem. 1999, 274, 1189–1192.
  42. Liu, C.; Yang, Q.; Zhu, Q.; Lu, X.; Li, M.; Hou, T.; Li, Z.; Tang, M.; Li, Y.; Wang, H.; et al. CBP mediated DOT1L acetylation confers DOT1L stability and promotes cancer metastasis. Theranostics 2020, 10, 1758–1776.
  43. Marmorstein, R.; Roth, S.Y. Histone acetyltransferases: Function, structure, and catalysis. Curr. Opin. Genet. Dev. 2001, 11, 155–161.
  44. Tjeertes, J.V.; Miller, K.M.; Jackson, S.P. Screen for DNA-damage-responsive histone modifications identifies H3K9Ac and H3K56Ac in human cells. EMBO J. 2009, 28, 1878–1889.
  45. Grant, P.A.; Eberharter, A.; John, S.; Cook, R.G.; Turner, B.M.; Workman, J.L. Expanded lysine acetylation specificity of Gcn5 in native complexes. J. Biol. Chem. 1999, 274, 5895–5900.
  46. Guo, R.; Chen, J.; Mitchell, D.L.; Johnson, D.G. GCN5 and E2F1 stimulate nucleotide excision repair by promoting H3K9 acetylation at sites of damage. Nucleic Acids Res. 2011, 39, 1390–1397.
  47. Aricthota, S.; Rana, P.P.; Haldar, D. Histone acetylation dynamics in repair of DNA double-strand breaks. Front. Genet. 2022, 13, 926577.
  48. Vaquero, A. The conserved role of sirtuins in chromatin regulation. Int. J. Dev. Biol. 2009, 53, 303–322.
  49. Jing, H.; Lin, H. Sirtuins in epigenetic regulation. Chem. Rev. 2015, 115, 2350–2375.
  50. Yuan, J.; Pu, M.; Zhang, Z.; Lou, Z. Histone H3-K56 acetylation is important for genomic stability in mammals. Cell Cycle 2009, 8, 1747–1753.
  51. Seto, E.; Yoshida, M. Erasers of histone acetylation: The histone deacetylase enzymes. Cold Spring Harb. Perspect. Biol. 2014, 6, a18713.
  52. Scher, M.B.; Vaquero, A.; Reinberg, D. SirT3 is a nuclear NAD+-dependent histone deacetylase that translocates to the mitochondria upon cellular stress. Genes Dev. 2007, 21, 920–928.
  53. Vaquero, A.; Sternglanz, R.; Reinberg, D. NAD+-dependent deacetylation of H4 lysine 16 by class III HDACs. Oncogene 2007, 26, 5505–5520.
  54. Michishita, E.; Mccord, R.A.; Berber, E.; Kioi, M.; Padilla-Nash, H.; Damian, M.; Cheung, P.; Kusumoto, R.; Kawahara, T.L.; Barrett, J.C.; et al. SIRT6 is a histone H3 lysine 9 deacetylase that modulates telomeric chromatin. Nature 2008, 452, 492–496.
  55. Barber, M.F.; Michishita-Kioi, E.; Xi, Y.; Tasselli, L.; Kioi, M.; Moqtaderi, Z.; Tennen, R.I.; Paredes, S.; Young, N.L.; Chen, K.; et al. SIRT7 links H3K18 deacetylation to maintenance of oncogenic transformation. Nature 2012, 487, 114–118.
  56. Miller, K.M.; Tjeertes, J.V.; Coates, J.; Legube, G.; Polo, S.E.; Britton, S.; Jackson, S.P. Human HDAC1 and HDAC2 function in the DNA-damage response to promote DNA nonhomologous end-joining. Nat. Struct. Mol. Biol. 2010, 17, 1144–1151.
  57. Zhu, Q.; Battu, A.; Ray, A.; Wani, G.; Qian, J.; He, J.; Wang, Q.E.; Wani, A.A. Damaged DNA-binding protein down-regulates epigenetic mark H3K56Ac through histone deacetylase 1 and 2. Mutat. Res. 2015, 776, 16–23.
  58. Johnson, D.P.; Spitz, G.S.; Tharkar, S.; Quayle, S.N.; Shearstone, J.R.; Jones, S.; Mcdowell, M.E.; Wellman, H.; Tyler, J.K.; Cairns, B.R.; et al. HDAC1,2 inhibition impairs EZH2- and BBAP-mediated DNA repair to overcome chemoresistance in EZH2 gain-of-function mutant diffuse large B-cell lymphoma. Oncotarget 2015, 6, 4863–4887.
  59. Yang, Y.; Yang, C.; Li, T.; Yu, S.; Gan, T.; Hu, J.; Cui, J.; Zheng, X. The Deubiquitinase USP38 Promotes NHEJ Repair through Regulation of HDAC1 Activity and Regulates Cancer Cell Response to Genotoxic Insults. Cancer Res. 2020, 80, 719–731.
  60. Li, F.L.; Liu, J.P.; Bao, R.X.; Yan, G.; Feng, X.; Xu, Y.P.; Sun, Y.P.; Yan, W.; Ling, Z.Q.; Xiong, Y.; et al. Acetylation accumulates PFKFB3 in cytoplasm to promote glycolysis and protects cells from cisplatin-induced apoptosis. Nat. Commun. 2018, 9, 508.
  61. Lozada, E.M.; Andrysik, Z.; Yin, M.; Redilla, N.; Rice, K.; Stambrook, P.J. Acetylation and deacetylation of Cdc25A constitutes a novel mechanism for modulating Cdc25A functions with implications for cancer. Oncotarget 2016, 7, 20425–20439.
  62. Zhang, M.; Hu, C.; Moses, N.; Haakenson, J.; Xiang, S.; Quan, D.; Fang, B.; Yang, Z.; Bai, W.; Bepler, G.; et al. HDAC6 regulates DNA damage response via deacetylating MLH1. J. Biol. Chem. 2019, 294, 5813–5826.
  63. Zhang, L.; Li, D.Q. MORC2 regulates DNA damage response through a PARP1-dependent pathway. Nucleic Acids Res. 2019, 47, 8502–8520.
  64. Chen, G.; Luo, Y.; Warncke, K.; Sun, Y.; Yu, D.S.; Fu, H.; Behera, M.; Ramalingam, S.S.; Doetsch, P.W.; Duong, D.M.; et al. Acetylation regulates ribonucleotide reductase activity and cancer cell growth. Nat. Commun. 2019, 10, 3213.
  65. Liu, J.; Shangguan, Y.; Tang, D.; Dai, Y. Histone succinylation and its function on the nucleosome. J. Cell. Mol. Med. 2021, 25, 7101–7109.
  66. Liu, X.; Rong, F.; Tang, J.; Zhu, C.; Chen, X.; Jia, S.; Wang, Z.; Sun, X.; Deng, H.; Zha, H.; et al. Repression of p53 function by SIRT5-mediated desuccinylation at Lysine 120 in response to DNA damage. Cell Death Differ. 2022, 29, 722–736.
  67. Jing, Y.; Ding, D.; Tian, G.; Kwan, K.; Liu, Z.; Ishibashi, T.; Li, X.D. Semisynthesis of site-specifically succinylated histone reveals that succinylation regulates nucleosome unwrapping rate and DNA accessibility. Nucleic Acids Res. 2020, 48, 9538–9549.
  68. Chen, X.F.; Tian, M.X.; Sun, R.Q.; Zhang, M.L.; Zhou, L.S.; Jin, L.; Chen, L.L.; Zhou, W.J.; Duan, K.L.; Chen, Y.J.; et al. SIRT5 inhibits peroxisomal ACOX1 to prevent oxidative damage and is downregulated in liver cancer. EMBO Rep. 2018, 19, e45124.
  69. Shi, R.; Wang, Y.; Gao, Y.; Xu, X.; Mao, S.; Xiao, Y.; Song, S.; Wang, L.; Tian, B.; Zhao, Y.; et al. Succinylation at a key residue of FEN1 is involved in the DNA damage response to maintain genome stability. Am. J. Physiol. Cell Physiol. 2020, 319, C657–C666.
  70. Gao, X.; Bao, H.; Liu, L.; Zhu, W.; Zhang, L.; Yue, L. Systematic analysis of lysine acetylome and succinylome reveals the correlation between modification of H2A.X complexes and DNA damage response in breast cancer. Oncol. Rep. 2020, 43, 1819–1830.
  71. Fontana, G.A.; Hess, D.; Reinert, J.K.; Mattarocci, S.; Falquet, B.; Klein, D.; Shore, D.; Thomä, N.H.; Rass, U. Rif1 S-acylation mediates DNA double-strand break repair at the inner nuclear membrane. Nat. Commun. 2019, 10, 2535.
  72. Abbott, D.W.; Holt, J.T. Finkel-Biskis-Reilly mouse osteosarcoma virus v-fos inhibits the cellular response to ionizing radiation in a myristoylation-dependent manner. J. Biol. Chem. 1997, 272, 14005–14008.
  73. Jiang, G.; Li, C.; Lu, M.; Lu, K.; Li, H. Protein lysine crotonylation: Past, present, perspective. Cell. Death Dis. 2021, 12, 703.
  74. Pelka, P.; Ablack, J.N.; Torchia, J.; Turnell, A.S.; Grand, R.J.; Mymryk, J.S. Transcriptional control by adenovirus E1A conserved region 3 via p300/CBP. Nucleic Acids Res. 2009, 37, 1095–1106.
  75. Zhao, L.J.; Loewenstein, P.M.; Green, M. The adenoviral E1A N-terminal domain represses MYC transcription in human cancer cells by targeting both p300 and TRRAP and inhibiting MYC promoter acetylation of H3K18 and H4K16. Genes Cancer 2016, 7, 98–109.
  76. Peng, Y.; Wang, Y.; Tang, N.; Sun, D.; Lan, Y.; Yu, Z.; Zhao, X.; Feng, L.; Zhang, B.; Jin, L.; et al. Andrographolide inhibits breast cancer through suppressing COX-2 expression and angiogenesis via inactivation of p300 signaling and VEGF pathway. J. Exp. Clin. Cancer Res. 2018, 37, 248.
  77. Welti, J.; Sharp, A.; Brooks, N.; Yuan, W.; Mcnair, C.; Chand, S.N.; Pal, A.; Figueiredo, I.; Riisnaes, R.; Gurel, B.; et al. Targeting the p300/CBP Axis in Lethal Prostate Cancer. Cancer Discov. 2021, 11, 1118–1137.
  78. Gruber, M.; Ferrone, L.; Puhr, M.; Santer, F.R.; Furlan, T.; Eder, I.E.; Sampson, N.; Schäfer, G.; Handle, F.; Culig, Z. p300 is upregulated by docetaxel and is a target in chemoresistant prostate cancer. Endocr. Relat. Cancer 2020, 27, 187–198.
  79. Tan, K.N.; Avery, V.M.; Carrasco-Pozo, C. Metabolic Roles of Androgen Receptor and Tip60 in Androgen-Dependent Prostate Cancer. Int. J. Mol. Sci. 2020, 21, 6622.
  80. Ravichandran, P.; Davis, S.A.; Vashishtha, H.; Gucwa, A.L.; Ginsburg, D.S. Nuclear Localization Is Not Required for Tip60 Tumor Suppressor Activity in Breast and Lung Cancer Cells. DNA Cell Biol. 2020, 39, 2077–2084.
  81. Judes, G.; Dubois, L.; Rifaï, K.; Idrissou, M.; Mishellany, F.; Pajon, A.; Besse, S.; Daures, M.; Degoul, F.; Bignon, Y.J.; et al. TIP60: An actor in acetylation of H3K4 and tumor development in breast cancer. Epigenomics 2018, 10, 1415–1430.
  82. Sun, Y.; Jiang, X.; Price, B.D. Tip60: Connecting chromatin to DNA damage signaling. Cell Cycle 2010, 9, 930–936.
  83. Avvakumov, N.; Côté, J. The MYST family of histone acetyltransferases and their intimate links to cancer. Oncogene 2007, 26, 5395–5407.
  84. Gorrini, C.; Squatrito, M.; Luise, C.; Syed, N.; Perna, D.; Wark, L.; Martinato, F.; Sardella, D.; Verrecchia, A.; Bennett, S.; et al. Tip60 is a haplo-insufficient tumour suppressor required for an oncogene-induced DNA damage response. Nature 2007, 448, 1063–1067.
  85. Fisher, J.B.; Kim, M.S.; Blinka, S.; Ge, Z.D.; Wan, T.; Duris, C.; Christian, D.; Twaroski, K.; North, P.; Auchampach, J.; et al. Stress-induced cell-cycle activation in Tip60 haploinsufficient adult cardiomyocytes. PLoS ONE 2012, 7, e31569.
  86. Liu, L.; Qiu, S.; Liu, Y.; Liu, Z.; Zheng, Y.; Su, X.; Chen, B.; Chen, H. Chidamide and 5-flurouracil show a synergistic antitumor effect on human colon cancer xenografts in nude mice. Neoplasma 2016, 63, 193–200.
  87. Kiweler, N.; Schwarz, H.; Nguyen, A.; Matschos, S.; Mullins, C.; Piée-Staffa, A.; Brachetti, C.; Roos, W.P.; Schneider, G.; Linnebacher, M.; et al. The epigenetic modifier HDAC2 and the checkpoint kinase ATM determine the responses of microsatellite instable colorectal cancer cells to 5-fluorouracil. Cell Biol. Toxicol. 2022; advance online publication.
  88. Spurling, C.C.; Godman, C.A.; Noonan, E.J.; Rasmussen, T.P.; Rosenberg, D.W.; Giardina, C. HDAC3 overexpression and colon cancer cell proliferation and differentiation. Mol. Carcinog. 2008, 47, 137–147.
  89. Wu, G.; Yu, W.; Zhang, M.; Yin, R.; Wu, Y.; Liu, Q. MicroRNA-145-3p suppresses proliferation and promotes apotosis and autophagy of osteosarcoma cell by targeting HDAC4. Artif. Cells Nanomed. Biotechnol. 2018, 46, 579–586.
  90. Lee, B.S.; Kim, Y.S.; Kim, H.J.; Kim, D.H.; Won, H.R.; Kim, Y.S.; Kim, C.H. HDAC4 degradation by combined TRAIL and valproic acid treatment induces apoptotic cell death of TRAIL-resistant head and neck cancer cells. Sci. Rep. 2018, 8, 12520.
  91. Li, X.H.; Huang, M.L.; Wang, S.M.; Wang, Q. Selective inhibition of bicyclic tetrapeptide histone deacetylase inhibitor on HDAC4 and K562 leukemia cell. Asian Pac. J. Cancer Prev. 2013, 14, 7095–7100.
  92. Nie, X.; Jia, W.; Li, X.; Pan, X.; Yin, R.; Liu, N.; Su, Z. FBXW7 induces apoptosis in glioblastoma cells by regulating HDAC7. Cell Biol. Int. 2021, 45, 2150–2158.
  93. Zhu, C.; Chen, Q.; Xie, Z.; Ai, J.; Tong, L.; Ding, J.; Geng, M. The role of histone deacetylase 7 (HDAC7) in cancer cell proliferation: Regulation on c-Myc. J. Mol. Med. 2011, 89, 279–289.
  94. Park, J.H.; Kim, S.H.; Choi, M.C.; Lee, J.; Oh, D.Y.; Im, S.A.; Bang, Y.J.; Kim, T.Y. Class II histone deacetylases play pivotal roles in heat shock protein 90-mediated proteasomal degradation of vascular endothelial growth factor receptors. Biochem. Biophys. Res. Commun. 2008, 368, 318–322.
  95. Schmid, N.; Dietrich, K.G.; Forne, I.; Burges, A.; Szymanska, M.; Meidan, R.; Mayr, D.; Mayerhofer, A. Sirtuin 1 and Sirtuin 3 in Granulosa Cell Tumors. Int. J. Mol. Sci. 2021, 22, 2047.
  96. Ceballos, M.P.; Angel, A.; Delprato, C.B.; Livore, V.I.; Ferretti, A.C.; Lucci, A.; Comanzo, C.G.; Alvarez, M.L.; Quiroga, A.D.; Mottino, A.D.; et al. Sirtuin 1 and 2 inhibitors enhance the inhibitory effect of sorafenib in hepatocellular carcinoma cells. Eur. J. Pharmacol. 2021, 892, 173736.
  97. Tan, J.; Liu, Y.; Maimaiti, Y.; Wang, C.; Yan, Y.; Zhou, J.; Ruan, S.; Huang, T. Combination of SIRT1 and Src overexpression suggests poor prognosis in luminal breast cancer. Onco. Targets Ther. 2018, 11, 2051–2061.
  98. Huffman, D.M.; Grizzle, W.E.; Bamman, M.M.; Kim, J.S.; Eltoum, I.A.; Elgavish, A.; Nagy, T.R. SIRT1 is significantly elevated in mouse and human prostate cancer. Cancer Res. 2007, 67, 6612–6618.
  99. Saunders, L.R.; Verdin, E. Sirtuins: Critical regulators at the crossroads between cancer and aging. Oncogene 2007, 26, 5489–5504.
  100. Marcu, M.G.; Jung, Y.J.; Lee, S.; Chung, E.J.; Lee, M.J.; Trepel, J.; Neckers, L. Curcumin is an inhibitor of p300 histone acetylatransferase. Med. Chem. 2006, 2, 169–174.
  101. Kusio-Kobialka, M.; Dudka-Ruszkowska, W.; Ghizzoni, M.; Dekker, F.J.; Piwocka, K. Inhibition of PCAF by anacardic acid derivative leads to apoptosis and breaks resistance to DNA damage in BCR-ABL-expressing cells. Anticancer Agents Med. Chem. 2013, 13, 762–767.
  102. Yang, H.; Pinello, C.E.; Luo, J.; Li, D.; Wang, Y.; Zhao, L.Y.; Jahn, S.C.; Saldanha, S.A.; Chase, P.; Planck, J.; et al. Small-molecule inhibitors of acetyltransferase p300 identified by high-throughput screening are potent anticancer agents. Mol. Cancer Ther. 2013, 12, 610–620.
  103. Di Martile, M.; Del, B.D.; Trisciuoglio, D. The multifaceted role of lysine acetylation in cancer: Prognostic biomarker and therapeutic target. Oncotarget 2016, 7, 55789–55810.
Contributors MDPI registered users' name will be linked to their SciProfiles pages. To register with us, please refer to : , , ,
View Times: 587
Revisions: 3 times (View History)
Update Date: 16 Nov 2022