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Chapter 3 - Mechanisms of Histone Modifications



Mechanisms of Histone Modifications
Zdenko Herceg1 and Rabih Murr1,2 1 Epigenetics Group. International Agency for Research on Cancer (IARC), Lyon, France 2 Friedrich Miescher Institute for Biomedical Research, Basel, Switzerland

The term “epigenetic” was first introduced by Conrad Waddington in 1942 to describe “The interactions of genes with their environment that bring the phenotype into being”. Currently, it includes all features such as chromatin and DNA modifications that are heritable and stable over rounds of cell division, but do not alter the nucleotide sequence within the underlying DNA [1]. Over the years, a wide variety of products and events have been lumped into epigenetics. These include paramutation, bookmarking, imprinting, gene silencing, X chromosome inactivation, position effect variegation, reprogramming, transvection, infection agents like prions, maternal conditioning, RNA interference, non coding RNA, small RNAs, DNA methylation and chromatin modifications. In this chapter, we will focus on epigenetic mechanisms involving histone modifications and recent development establishing a link between chromatin modifications (with an emphasis on acetylation and methylation) and cellular processes such as transcription and DNA repair.


Histone modifications
In all eukaryotes, chromatin is a highly condensed structure that forms the scaffold of fundamental nuclear processes such as transcription, replication and DNA repair [2]. Chromatin exists in at least two conceptually distinct functional forms: a condensed form during mitosis and meiosis that generally lacks DNA regulatory activity, called heterochromatin; and a looser decondensed form, which provides the environment for DNA regulatory processes, called euchromatin. Nucleosomes are the building blocks of chromatin and they represent two turns of genomic DNA (147 base pairs) wrapped around an octamer of two subunits of each of the core histones H2A, H2B, H3, and H4. The amino-terminal portion of the core histone proteins contains a flexible and highly basic tail region, which is conserved across various species and is subject to various post-translational modifications (Fig. 3.1). The structure of chromatin fulfils essential functions, not only by condensing and protecting DNA, but also in preserving genetic information and controlling gene expression [3]. However, given its compacted structure, chromatin hinders several important cellular processes including, transcription, replication, and the detection/repair of DNA breaks [4,5]. Therefore, chromatin

Handbook of Epigenetics: The New Molecular and Medical Genetics. DOI: 10.1016/B978-0-12-375709-8.00003-4 ? 2011 Elsevier Inc. All rights reserved.

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Figure 3.1 26

Schematic representation of a nucleosome and major histone modifications. Post-translational modifications of histones occur primarily on N-terminal tails of core histones (H2A, H2B, H3, and H4) and include acetylation, methylation, phosphorylation, and ubiquitination. Note that several lysines (e.g. Lys 9) can be either acetylated or methylated.

must be first made relaxed to allow access of cellular machineries to chromatin DNA. This leads to one of the most fundamental questions in biology – how is chromatin remodeled? A part of the answer resides in the fact that cells have evolved cellular mechanisms that alter the structure of chromatin. These activities include ATP-dependent nucleosome mobilization (chromatin remodeling) and post-translational histone modifications. Chromatin modifications can occur through covalent additions to histones. Histone amino-terminal tails are most frequently targets of modifications. There are at least 60 different residues on histones where modifications have been detected and it is likely that this number is underestimated; emergence of new techniques will, without any doubt, help identify new target residues and new modifications. There are, to date, at least eight different types of histone modification: acetylation, methylation, phosphorylation, ubiquitination, sumoylation, ADP ribosylation, deimination, and proline isomerization [6–8]. Traditionally, two mechanisms are thought to govern the function of these modifications. First, these different marks could affect the nucleosome-nucleosome or DNA-nucleosome interactions through the addition of physical entities or by changing histone charges. Second, different marks could represent a docking site for the recruitment of specific proteins which could result in different cellular outcomes. Additionally, numerous reports raised the possibility that all these modifications are combinatorial and interdependent and therefore may form the “histone code”, which means that combination of different modifications may result in distinct and consistent cellular outcomes [9,10]. The molecular mechanisms, the role, and the interdependence of these modifications will be discussed in the following paragraphs.

chapter 3 Mechanisms of Histone Modifications

Proline Isomerization
Isomerization is defined as the transformation of a molecule into a different isomere. Isomerization of proteins was first described in 1968 [11] and it was shown to dramatically affect protein conformation by disrupting the secondary structure of polypeptides. It can adopt two distinct conformations: cis or trans. Isomerization occurs spontaneously, but enzymes called proline isomerases have evolved in order to accelerate switching between different conformations (cis-trans). The first evidence that histones can be isomerized was reported in 2006 [12] when Frp4 was identified as a histone isomerase of prolines 30 and 38 (P30 and P38) on histone H3 tail [Fig. 3.1]. The conformational status of P38 is necessary for the induction of lysine 36 of histone H3 (H3K36) methylation and its isomerization appears to inhibit the ability of Set2 to methylate H3K36.

Sumoylation consists in the addition of a “Small Ubiquitin-related MOdifier protein” (SUMO) of ~100 amino acids. Similar to ubiquitination, SUMO is always covalently attached to other proteins through the activities of members of an enzymatic cascade (E1-E2-E3). Histone sumoylation was first reported in 2003, when Shiio et al. found that H4 can be modified by SUMO and they suggested that this modification leads to the repression of transcriptional activity through the recruitment of HDACs and HP1 proteins [13]. Recently, it was demonstrated that all four core histones can be sumoylated in yeast [Fig. 3.1]. The putative sumoylation sites were identified as K6/7 and to a lesser extent K16/17 of H2B, K126 of H2A, and all four lysines in the N-terminal tail of H4. Histone sumoylation has a role in transcription repression by opposing other active marks such as acetylation and ubiquitination [14].

Ubiquitin is a 76 amino acid protein highly conserved in eukaryotes. Ubiquitination (or ubiquitylation) refers to the post-translational modification of the ?-amino group of a lysine residue by the covalent attachment of one (monoubiquitination) or more (polyubiquitination) ubiquitin monomers. Typically, polyubiquitination marks a protein to be degraded via the 26S proteasome, whereas monoubiquitination modifies protein function. Histone H2A was the first histone identified to be ubiquitinated [15]. Later on, H2B (K119, K120 (K123 in yeast) and K143), H3 and H1 were also reported to be ubiquitinated [16] [Fig. 3.1]. Histones appear to be mostly monoubiquitinated, although H2A and H2B may be polyubiquitinated [17,18]. As for non-histone proteins, histones ubiquitination consists of formation of an isopeptide bond between the C-terminus of ubiquitin and a lysine side chain of histone by sequential actions of E1 activating, E2 conjugating, and E3 ligase enzymes. E2 and E3 play a crucial role in specifying the protein to be ubiquitinated. E3-ligases mostly belong to HECT (Homologous to E6AP-C Terminus) or RING (Really Interesting New Gene) protein families. H2BK123 specific E2 in yeast, Rad6 was the first histone E2 to be identified at the beginning of the century [19]. Rad6 activity is combined to the RING finger E3 ligase, Bre1. Homologous of Rad6, Rhp6 in drosophila and HR6A/B in humans, as well as homologous of Bre1, Brl1 in Drosophila and RNF20 in humans were also shown to be involved in H2B ubiquitination [20–24]. Histone H2A ubiquitination is dependent on the Polycomb repressive complex 1 (PRC1); PRC2 set up the H3K27me3 marks that are recognized by the PRC1 complex which would ubiquitinate H2A and silence gene expression. At least two members of the PRC1 complex, RING1b (also known as Rnf2) and BMI1, were found to form a heterodimer that ubiquitinates H2A [25–29]. The PRC1 complex is formed of four core proteins that include the PcG proteins Polycomb, Polyhomeotic, Posterior sex combs (PsC) and RING (also known as sex comb extra), in addition to many other proteins


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[30–32]. Ortholog of PRC1 and RING1b also exist in complexes distinct from PRC1 and these are the RING-associated factor (dRAF) in D. melanogaster and bCL6 corepressor (bCoR) in mammals. These PRC1-like complexes can also ubiquitinate H2A [26,33]. The addition of the ubiquitin moiety to histones is reversible through the activity of deubiquitinating enzymes that consist of ubiquitin C-terminal hydrolases and ubiquitin-specific processing proteases (UBPs). Sixteen UBPs have been identified so far in yeast. These UBPs differ by the length of their amino-terminal part that confers their specificity [34,35]. The best studied UBPs are UBP8 and UBP10 that are specific for H2B. UBP8 belongs to the SAGA complex [36–38] and orthologs were found in Drosophila (Nonstop) and in human (USP22) and they were both involved in H2B deubiquitination in the context of the SAGA complex [39–42]. UBP10 activity is SAGA-independent but SIR-dependent and its orthologs in higher eukaryotes are still to be defined [43–45]. Ubiquitinated H2A at K119 (uH2A) was shown to be important for transcriptional activation and several active genes were shown to contain a high percentage of uH2A [46,47,48]. Surprisingly, uH2A was also linked to transcription inhibition [48,49,50,51]. Similarly, ubiquitination of H2B was linked to both activation and inhibition of transcription [35,52]. The ambiguity on the role of the mono-ubiquitinated H2B (uH2B) in transcription regulation was mostly due to the lack of specific antibodies. However, the elaboration of a suitable anti-uH2B monoclonal antibody by using a branched peptide as an antigen partially clarified this matter and opted for a positive correlation between uH2B-K120 and gene expression. Indeed, this antibody was used in ChIP-Chip experiments on tiling arrays and the results showed a preferential association of uH2BK120 with the transcribed regions of highly expressed genes [53]. As this mark is not associated to distal gene promoters but rather to transcription start site (TSS) and further to gene bodies of active genes, it was suggested that it is linked to transcription elongation rather than initiation. Further proof of the correlation of uH2B with active transcription came from an elegant biochemical study performed in Tom Muir’s laboratory. In this study, the authors used two traceless orthogonal expressed protein ligation (EPL) reactions to chemically and specifically ubiquitinate H2BK120 incorporated into chemically defined nucleosomes. The results showed a direct activation of H3-K79 methylation by hDot1L, a mark related to gene activation [54]. Histone ubiquitination may affect other histone modifications. For example, histone deacetylase 6 (HDAC6) was shown to bind to ubiquitin through its zinc-finger domain. H3K4 and H3K79 methylation was shown to be dependent on Rad6-mediated H2BK123 ubiquitination [54–58]. The effect of ubiquitination on histone methylation can explain its role in both activation and inhibition of transcription. For instance, it has been proposed that ubiquitination of H2B occurs mostly in euchromatin leading to H3K4 and H3K79 methylation, which would prevent Sir proteins from association with active euchromatic regions, thereby restricting Sir proteins to heterochromatic regions to mediate silencing [59]. At the same time in euchromatin, the ubiquitination would activate the transcription by methylating H3K4 and by facilitating the transcriptional elongation [60,61].


ADP-ribosylation is a post-translational modification defined by the addition of an ADPribose moiety onto a protein using NAD? as a substrate. If the transfer takes place on an amino-acid acceptor, it is referred to as mono- or poly-(ADP-ribosyl)ation (PARation) and if it occurs on an acetyl group it is called O-acetyl-ADP-ribosylation. Mono(ADP-ribosyl) ation is mediated by ADP ribosyl transferases (ART) and the enzymes responsible for the PARation are the poly(ADP-ribose) polymerases (PARPs) [62,63]. All core histones and linker histone H1 are subjects to mono(ADP-ribosyl)ation either in response to genotoxic stress or in physiological conditions depending on the cell cycle stage, proliferation activity, and degree of terminal differentiation [64,65]. PARation can also be detected on the

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majority of histone types. There seems to be some specificity in the activity of PARP proteins on histones; for example PARP1 seems to preferentially poly(ADP-ribosyl)ate the linker histone H1 whereas PARP2 prefers core histones. In response to single strand break (SSB), PARP1 and PARP2 poly(ADP-ribosyl)ate the C- and N-terminus of histones H1 and H2B leading to the relaxation of the chromatin structure facilitating the access of single strand break repair (SSBR)/base excision repair (BER) factors to the site of damage. This has been explained, at least in part, by the fact that PARation of histones leads to their removal from the chromatin. The removal of histones results in the opening of the chromatin structure (the same mechanism leads to transcriptional activation). Moreover, PAR is used for tagging the region affected by DNA damage allowing adequate response of the cell according to the extent of damage signaled by the presence of PAR moieties. On the other hand, recent studies indicated that PARP-dependent ribosylation in response to DNA damage may induce local chromatin condensation rather than relaxation. Indeed, PAR moieties are recognized by the macrodomain of the histone variant macroH2A1.1. This would lead to a transient condensation or looping, increase phosphorylation of H2AX at the sites of break and reduced recruitment of Ku70/80 leading to altered DNA damage response [66]. It is difficult to reconcile this last study with the rest of the literature on the role of PARP in DNA repair. The obvious explanation is that Poly(ADP-ribosyl)ation leads to a quick and transient compaction of the chromatin that would protect the DNA from additional damage and this is rapidly reversed allowing DNA repair to take place. This hypothesis is supported by the quick and transient nature of the PAR-macroH2A1.1 interaction-dependent chromatin condensation that is gradually lost after the reduction of Poly(ADP-ribosyl)ation levels. The role of mono- and poly(ADP-ribosyl)ation in DNA repair and transcription may be explained by their interaction with other chromatin modifications in the context of the “histone code”. For example, Mono-ADP-ribosylation on H4 seems to occur preferentially when H4 is hyperacetylated and mono-ADP-ribosylation of histone H1.3 on arginine 33 (R33) may reduce cyclic AMP-dependent phosphorylation of Serine 36 (S36) [67].


Protein phosphorylation represents the addition of a phosphate (PO4) group to a protein molecule. Phosphorylation is catalyzed by various specific protein kinases, whereas phosphatases mediate removal of the phosphate group. Histones can also get phosphorylated and the most studied sites of histone phosphorylation are the serine 10 of histone H3 (H3S10) that is deposited by the Aurora-B kinase during mitosis (Fig. 3.1) and S139 (129 in yeast) of H2A variant, H2AX, DNA damage-dependent phosphorylation by ATM and ATR. H2AX could be additionally phosphorylated on tyrosine 142. H4 (S1) and linker histone H1 (S18, S173, S189, T11, T138, and T155) were also shown to be phosphorylated by the CK2 and DNA-PK respectively.

Role of histone phosphorylation in transcription regulation
The relationship between histone phosphorylation and gene expression is far from being totally understood. The phosphorylation of H3S10 (H3S10P) was initially linked to chromosome condensation and segregation during mitosis and meiosis [68,69]. The role of this mark during mitosis was investigated in detail in a study performed in the laboratory of David Allis [70]. The authors followed the status of HP1-?, -? and -? during mitosis. These proteins are recruited via interaction with H3K9me3 and lead to heterochromatinization. However, during mitosis, these marks are ejected from chromatin even though H3K9me3 marks are preserved. The study demonstrated that the addition of the H3S10P mark is responsible for the eviction of these proteins. It was proposed that this would lead to better recruitment of players necessary for proper condensation and segregation of chromosomes, although this hypothesis awaits experimental confirmation. The role of H3S10P in chromatin condensation suggests that it should be involved in transcriptional repression;

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however, evidence has accumulated indicating that this mark has rather an important role in transcriptional activation of genes in various organisms. For instance, induction of heatshock genes in Drosophila is concomitant to a high increase of H3S10P [71,72]. On the other hand, dephosphorylation of H3S10 is carried out by a phosphatase called PP2A (Protein Phosphatase 2A) and results in the inhibition of transcription [72]. Additionally, H3S10P was shown to be important for the activation of NFKB-regulated genes but also “immediate early” genes such as c-fos and c-jun. This phosphorylation is proposed to induce the accumulation of phosphor-binding protein 14-3-3 [73]. Genome-wide ChIP-Chip analysis in budding yeast has shown that several kinases are not only present in the cytoplasm but also on the chromatin of specific genes [74], which suggests that the kinase signal transduction cascade could have direct effect on gene expression by phosphorylating the histones of specific genes or gene promoters [8].

Role of histone phosphorylation in DNA repair
Beside its role in chromosomal condensation and transcription, phosphorylation of histones, in particular phosphorylation of H2AX, has a role in DNA damage response and DNA repair. Rapid phosphorylation of H2AX, at serine 129 (?H2AX) by the PI3K kinases at double strand break (DSB) sites, is one of the first and most easily detectable DNA damage signaling posttranslational events. It anticorrelates to the phosphorylation of Tyrosine (Y) 142 of H2AX. Indeed, recent studies have shown that H2AX-Y142 is constitutively phosphorylated under physiological conditions by the activity of WSTF (Williams–Beuren syndrome transcription factor) and is dephosphorylated in response to DNA damage via the activity of Eya tyrosine phosphatase in correlation to the increase in serine phosphorylation. The kinase activity of WSTF as well as the phosphatase activity of Eya shown to be important for the early recruitment of phospho-ATM and MDC1 to sites of DNA damage, thus privileging DNA repair over apoptosis [75,76]. ?H2AX can be detected over kilobases (in yeast) or megabases (in mammalian cells) from sites of DSBs [77,78] and is required for the retention/ accumulation of repair proteins [79,80,81]. ?H2AX also plays a role in cohesion binding to a large region around DSB, an event thought to be important for sister chromatid cohesion in post-replicative repair [82,83]. Interestingly, ?H2AX is required for the recruitment of the NuA4 histone acetyl-transferase (HAT) complex (yeast homolog of mammalian TIP60) to sites of DNA DSBs induced by HO endonuclease [80]. The recruitment of this HAT complex to ?H2AX is mediated by Arp4 and leads to acetylation of chromatin surrounding the break site, thereby facilitating efficient repair of DNA damage [80]. As well as being a component of NuA4 HAT complex, Arp4 is a subunit of the ATP-dependent chromatin remodeling complex INO80/SWR1. It was shown that INO80/SWR1 is also recruited to ?H2AX around DNA breaks and its remodeling activity seems to be required for the repair of DNA DSBs [80,84,85]. Hence it would appear that cells can utilize the activities of both histone modifying and remodeling complexes in order to facilitate DNA repair. The precise role of ?H2AX in DSBs is still under debate. Originally, it was suggested that the phosphorylation of H2AX is essential for the recruitment of DNA repair enzymes [86] through their BRCT (BRCA1 COOH Terminal) domain [87]. However, a study by Celeste et al., changed our understanding on the role of ?H2AX by demonstrating that DNA repair proteins, including Brca1 and Nbs1, are recruited to DNA breaks even in the absence of ?H2AX. On the other hand, the presence of ?H2AX is essential for the formation of IRIF (Irradiation-Induced Foci) [79], indicating that the role of H2AX phosphorylation may be dispensable for the original recruitment of DNA repair factors but indispensable for the accumulation/retention of these factors at DNA break sites. DNA break-associated histone phosphorylation can also occur on H4S1 through the activity of Casein Kinase II (CK2) in response to DNA damage and this facilitates double strand break (DSB) repair via nonhomologous end joining (NHEJ) [88]. A study from C?té’s laboratory demonstrated that this phosphorylation coincides with a decrease in acetylation, suggesting that it occurs


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after H4 deacetylation and that these events may regulate chromatin restoration after repair is completed [89]. Finally, linker histone H1 is found to be phosphorylated by PI3K family member DNA-PKcs, and this phosphorylation is required for efficient DNA repair by NHEJ [90–92].

Protein methylation is a covalent modification that represents the addition of a methyl group from the donor S-adenosylmethionine (SAM) on carboxyl groups of glutamate, leucine, and isoprenylated cysteine, or on the side-chain nitrogen atoms of lysine, arginine, and histidine residues [93]. However, histone methylation occurs only on arginines and lysines. Arginines can be mono- or dimethylated whereas lysines can be mono-, di- or trimethylated [8]. Arginine methylation can be either symmetrical or asymmetrical. The enzymes responsible for histone methylation are grouped into three different classes: [1] the lysine-specific SET domain-containing histone methyltransferases (HMT) involved in methylation of lysines 4, 9, 27, and 36 of histone H3 and lysine 20 of histone H4; [2] non-SET domain-containing lysine methyltransferases involved in methylating lysine 79 of histone H3; and [3] arginine methyltransferases involved in methylating arginines 2, 17, and 26 of histone H3 as well as arginine 3 of histone H4 [Fig. 3.1]. Whereas most covalent histone modifications are reversible, until recently it was unknown whether methyl groups could be actively removed from histones. The first histone demethylase to be discovered was LSD1, which mainly demethylates H3K4 but could also demethylate H3K9, when it is present in a complex with the androgen receptor [94,95]. A flow of other related enzymes were subsequently discovered and classified into two families of histone lysine demethylases: JMD2 and JARID1 families. The JMD2 (Jumonji C (JmcC)-domain containing proteins) family includes JHDM3A (Jumonji C (JmjC)-domaincontaining histone demethylase 3A; also known as JMJD2A); JMJD2C/GASC1 [96], which can both demethylate H3K9 and H3K36 [97–99]; and JMJD2B [100] and JMJD2D [101] which demethylate H3K9, JHDM1 (JmjC domain-containing histone demethylase 1) (demethylates H3K36) and UTX [97,98,102–104]. JARID1 proteins include RBP2, PLU1, SMCY/Jarid1ds and SMCX [105,106]. Histone arginine methylation marks were shown to be reversible. The first report about arginine demethylation suggested that methylated arginines on histones H3 (R3) and H4 (R17) are converted into citrulline via the activity of the human “peptidylarginine deiminase 4” protein, Pad4; this process was called “demethylimination or deimination” because the methyl group is removed along with the imine group of arginine [107,108]. Pad4 can deiminate multiple arginine sites on histones H3 (R2, R8, R17, and R26) and H4 (R3) [107]. Beside its function as an intermediate in the histone demethylation process, deimination has been involved in the estrogen signaling pathway [109]. On the other hand, a direct histone arginine demethylase, namely JMJD6, was recently identified and was found to belong to the JMD2 family [110].


Role of histone methylation in transcription regulation
The methylation mark on histones could be related to activation, elongation, or repression of gene expression, For example, H3K4me, me2 and me3 have been found on active promoters and linked to transcription initiation and elongation [60,61,111–113] whereas HK36me2/me3 have been correlated to transcription elongation [10,60,114–116]. To obtain a more detailed picture on histone methylation distribution along genes, ChIP-Chip and ChIP-seq experiments were performed and showed that H3K4me3 peaks at 5’ ends and at promoter proximal regions of active genes, H3K4me2 peaks at active gene bodies whereas H3K4me is enriched at the 5’ end of active genes. On the other hand H3K36me2/3 marks are enriched in active gene bodies and mostly at the 3’ end of active genes [114,117–123]. The precise underlying mechanism of H3K4me-dependent transcription regulation is still not

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clear. One possibility would be that histone modifying complexes or chromatin remodeling factors, such as Taf3 [124], could recognize and bind to the methylation mark through their PHD (Plant Homeo Domain), thus activating the transcription. H3K36 methylation marks are then recognized by the chromodomain of the Eaf3 subunit of The Rpd3S HDAC [120,125–128]. This would lead to the deacetylation of gene bodies and thus prevent, transcription initiation at cryptic sites of gene bodies. Methylation of H3K79 was also implicated in transcription activation and elongation [129]; however, this should be taken carefully, because it is based mainly on the fact that this mark was related to the activation of HOXA9 and that it limits the spreading of heterochromatin by preventing Sir2 and Sir3 from spreading into euchromatin. Moreover, detailed genome-wide study showed that while both H3K79me2 and H3K79me3 are enriched in gene bodies in yeast and Drosophila, only Drosophila H3K79me2 correlates with active transcription [122,130]. Three lysine methylation sites are connected to transcriptional repression: H3K9, H3K27, and H4K20. Very little is known regarding the repression functions of H4K20 methylation compared to a large number of studies on the two other repressive marks. Methylation of H3K9 is carried out by SUV39H1 and SUV39H2 in humans (its homolog Clr4, cryptic locus regulator 4, in Schizosaccharomyces pombe, and Su(var)3–9, suppressor of position-effect variegation, in Drosophila). These HMT have been shown to contain an SET domain. SET domain usually contains 130–140 amino acids and is a common feature of Trithorax (Thx) and Polycomb (PcG) group proteins which are involved in transcriptional activation and repression, respectively. Su(var)3–9 and its homologs were shown to be important for proper heterochromatin formation. These findings suggest a role for H3K9 methylation in gene silencing through correct heterochromatin folding [131,132]. It is now well established that HP1 recognizes methylated H3K9 through its chromodomain, contributing in part to the formation of heterochromatin. How are H3K9me2/3 and subsequent heterochromatinization initially targeted to DNA sequences? Two mechanisms could serve as the initial trigger for H3K9me: DNA-binding factors such as transcription factors or RNAi. Evidence for direct targeting of H3K9me by RNAi came first from studies on the core RNAi machinery which includes Dicer (Dcr), Argonaute (Ago) and RNAdependent RNA polymerase (RdRP) in S. pombe [133]. Later, several studies on different organisms demonstrated the role of RNAi in heterochromatin establishment [134–144]. The involvement of transcription factors such as Atf1, PCR1, and Taz1 in targeting heterochromatinization was also reported [145–149]. Although H3K9me was traditionally linked to repression, a recent study showed that H3K9me3 could be located in the gene bodies of active genes along with HP1 [150]. This observation led to the currently used model where H3K9me within the coding regions is activator whereas H3K9me in the promoters is repressive. H3K27 methylation has been implicated in the silencing of HOX gene expression. There are also indications that the same mark would be involved in the inactivation of the X chromosome and silencing during genomic imprinting. Interestingly, ChIP-Chip and ChIP-seq studies in ES cells indicated that some of the genes which are not expressed in ES cells have both repressive (H3K27me3) and active (H3K4me3) marks at their promoters, forming the so-called “bivalent domains”. Along differentiation, bivalent domain genes are resolved into monovalent by getting rid of one associated mark and thus get either stably activated or stably repressed. Therefore, these bivalent domains were thought to keep genes repressed at a certain developmental window but poised for activation in another subsequent developmental stage [111,151,152]. Arginine methylation was thought to be an activation mark as suggested by the fact that protein arginine methyltransferases are recruited to promoters by transcription factors [153]. One example of promoters regulated by histone arginine methylation is the pS2 promoter, a downstream target in the ER (Estrogen Receptor) pathway. Indeed, Metivier and colleagues showed that the transcription of this gene goes ON/OFF in a very controlled and specific fashion forming cycles of activation [154]. The activation of pS2 transcription correlated


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with the recruitment of protein arginine methyltransferase 1 (PRMT1) and Cofactor Associated Arginine Methyltransferase 1 (CARM1) recruitment. However, recent studies from the same group as well as another group found that these activation cycles could actually result from cycles of DNA methylation/demethylation and were probably not due to arginine methylation marks [155,156]. We now know that the arginine methylation effect on transcription could be either activating or repressive and depends on the type of arginine methyltransferase (RMT) involved [157]. For example, type I RMT, which include CARM1, PRMT1, and PRMT2 and generate monomethyl-arginine and asymmetric dimethyl-arginine derivatives, are involved in activation, while type II arginine methyltransferase PRMT5, which generates monomethyl-arginine and symmetric dimethyl-arginine derivatives, is involved in repression [158–164]. On the other hand, PRMT5 was shown to be associated with transcriptional repression. It associates with mSin3/HDAC and Brg1/hBrm and it is recruited to genes involved in control of cell proliferation (e.g. c-Myc target gene: cad and tumor suppressors: ST7 and NM23) in correlation with their repression [165,166].

Role of histone methylation in DNA repair
The role of histone methylation in the DNA damage response and DNA repair is less clear than the role of histone acetylation and phosphorylation; however, the involvement of lysine methylation, in processes other than transcriptional regulation, has recently received considerable attention. Methylation of H4 by Set9 histone lysine methyltransferase functions to localize Crb2, a DNA damage sensor and checkpoint protein in Schizosaccharomyces pombe, to sites of DNA damage hence increasing cellular survival following genotoxic stress [167]. Crb2 recruitment to DNA repair foci is dependent on the recognition of methylated H4K20 via the double tudor domains of Crb2 [168]. Subsequently, ionizing radiation-induced DNA damage generates nuclear foci at sites of DNA repair, which contain methylated H4K20 and the cell-cycle checkpoint protein Crb2 [167]. Similarly, the mammalian homolog of Crb2, 53BP1, also binds methylated H3 at sites of DNA DSBs [168,169]. Interestingly, Crb2 and 53BP1 do not recognize the trimethyl form of K20, which may indicate a different role of this modification in response to DNA damage.


Acetylation describes a reaction that introduces an acetyl functional group into an organic compound. Both histones and non-histone proteins can be acetylated. Histone acetylation consists in the transfer of an acetyl group from acetyl-CoA to the lysine ?-amino groups on the N-terminal tails of histones. This enzymatic activity is catalyzed by enzymes called histone acetyltransferases (HATs). The acetyl-CoA recognizes a specific domain within HATs called AT domain, Arg/Gln-X-X-Gly-X-gly/Ala. HAT enzymes often exist in multisubunit complexes that count one HAT catalytic subunit, adapter proteins, several other molecules of unknown function and, in many cases, a large scaffold protein called TRRAP. Acetylation can occur on specific lysines in all four histones (H3, H4, H2B, and H2A) [Fig. 3.1]. Hyperacetylation of histones is considered as a hallmark of transcriptionally active regions. Studies also revealed that the role of acetylation is not exclusively related to transcription, but can affect other DNA-based cellular processes such as DNA repair and replication.

Classification of HATs
Two classifications can be used to separate HATs.

First Classification
This divides HAT complexes into two big classes based on their suspected cellular origin and functions: A-type and B-type HATs. A-type HATs are nuclear enzymes that catalyze acetylation on already deposited histones in the context of the chromatin. B-type HATs are cytoplasmic

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enzymes and thought to be responsible for the acetylation of newly synthesized histones leading to their transport from the cytoplasm to the nucleus where they are deposited onto newly replicated DNA [170,171].

Second Classification
Instead of classifying HATs based on their cellular localization, modern classification uses structural criteria such as the presence or absence of chromodomains, bromodomains, and zinc finger domains. This classification separates the HATs into two major families: Gcn5related acetyltransferases (GNATs) and the MYST (for ‘MOZ, Ybf2/Sas3, Sas2, and Tip60)related HATs. To these families, we can add p300/CBP HATs, the general transcription factor HATs which include the TFIID subunit TAFII250, and the nuclear hormone-related HATs: SRC1 and ACTR (SRC3). The classification of these different families is not based on functional criteria. Due to space restriction, only complexes based on the second classification will be detailed in the following.

GNAT superfamily
All the GNAT superfamily members share structural and sequence similarity to Gcn5. This superfamily is characterized by four regions with different degrees of conservation (labeled A to D) spanning over 100 residues. These regions were first defined by the comparison between Gcn5 and B-type Hat1. Motif A, also called AT domain, contains an Arg/GlnX-XGly-X-Gly/Ala sequence and is shared with other HAT families. It is the most highly conserved and is important for acetyl-CoA recognition and binding. Tridimensional structure of this motif is highly conserved in all 15 GNAT proteins crystallized so far [172]. The C motif is found in most of the GNAT family acetyltransferases but not in the majority of known HATs. The GNAT superfamily contains over 10,000 members distributed in all kingdoms of life including histone acetyltransferases (HATs) but also nonhistone AT (see http://supfam.mrc-lmb.cam.ac.uk/SUPERFAMILY). The most relevant HATs of this family are: Gcn5, PCAF, Hat1, Elp3, and Hpa2.


MYST superfamily
The MYST family was named after its founding members: MOZ, Ybf2/Sas3, Sas2, and Tip60 [173]. These proteins are grouped together on the basis of their close sequence similarities and their possession of a particular acetyltransferase homology region (part of motif A of the GNAT superfamily) that binds Acetyl-CoA [174], in addition to a zinc finger domain called C2HC (C-X2-C-X13-H-X-C), and a “E-R” motif (Esa1-Rpd3), both needed for the enzymatic activity and for the substrate recognition [175]. Recently, additional members of this family were identified including Esa1 in yeast, MOF in Drosophila, and HBO1 and MORF in mammals. Despite their structural similarities, the members of this superfamily have various functions in various organisms. They resemble those of the GNAT family as both have an AT domain [176], but differ by the fact that they have different C- and N-termin, leading to different substrates. In addition, MYST family members possess either a chromodomain or an additional zinc finger domain termed the PHD domain [177,178].

p300 and CBP are often referred to as a single entity, since the two proteins are considered as structural and functional homologs and both proteins were subsequently shown be functionally interchangeable. But the two proteins diverge in several functional and structural properties. Indeed, some studies identified phosphorylation residues which are specific for each of the two proteins [179]. Another difference is that, in response to ionizing radiation (IR), p300, but not CBP, is important for apoptosis induction (probably through the activation of p53) [180]. In addition, whereas both proteins are necessary for apoptosis and G1 arrest of F9 embryo carcinoma cells, differentiation and induction of the cell cycle

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inhibitor p21/Cip1 critically depends on p300, while induction of p27/Kip1 requires CBP [181]. The most striking divergence comes from loss of function studies showing that individual knockouts of each of the two proteins resulted in two different phenotypes [182,183]. The specificity of the acetyltransferase activity of both proteins may explain, at least in part, the different functions between the two proteins. For instance, CBP was recently shown to have a preference for acetylating K12 on histone H4, while p300 preferentially acetylates K8 on histone H4 in vivo [184]. However, both proteins were shown to be able to acetylate H3K56 in collaboration with ASF1A histone chaperone. Another histone chaperone, CAF1, is needed for the incorporation of H3K56Ac into the chromatin, notably in response to DNA damage [185]. Other studies showed that H3K56 can be acetylated in the cytoplasm by Gcn5-containing HAT complex called HatB3.1 prior to its transport to the nucleus [186,187], which makes Gcn5 both nuclear and cytoplasmic. Another HAT-like complex in yeast, called Rtt109p, was also defined as responsible for H3K56 acetylation in the cytoplasm [188]. P300/CBP are large proteins (~300?kDa) containing more than 2400 residues. Four interaction domains have been characterized throughout their sequence. These include a bromodomain motif [189,190], which is also found in several other HATs such as Gcn5 and PCAF. P300/CBP have homologs in most metazoas but not in inferior eukaryotes including yeast. They were first identified as transcriptional adaptors for many different transcription factors that directly contact DNA-bound activators. In vitro studies seem to indicate that p300/CBP preferentially acetylate K12 and K15 of H2B, K14 K18 and K56 of H3K5, and K8 of H4 [191]. HAT proteins have also been directly implicated in transcriptional activation brought about by hormone signals. The HAT activity of human coactivators ACTR, SRC-1, and TIF2, which interact with nuclear hormone receptors, confirms the involvement of acetylation in yet another system of transcriptional regulation and defines a unique family of HATs. The members of this family share several similarities including HAT domain in the C-terminus, and an N-terminal, basic helix-loop-helix/PAS region [192], as well as receptor and coactivator interaction domains.


HAT complexes
Most HAT enzymes, alone, are not able to acetylate histones in the context of nucleosomes. However, when present in multisubunit complexes, these enzymes become more stable and more histone-type specific. Furthermore, the substrate of the HAT enzyme may change according to the HAT complex to which it belongs. This modification in specificity is further confirmed by the fact that distinct HAT complexes having distinct substrate specificity may share common subunits. For instance, TRRAP is shared between several HAT complexes, and STAGA and TFTC complexes share all their subunits except some high molecular weight TAFs which are not part of STAGA. HAT complexes were purified in both humans and yeast, were functionally equivalent in the two organisms, and were divided into several families. Between all HAT complexes, GNAT or SAGA-like HAT complexes (SAGA, SLIK, PCAF, STAGA, TFTC) are unique by the fact that they contain TAFs. The enzymatic subunit of these complexes can be represented by Gcn5 or PCAF. To date, two complexes belonging to this group have been discovered in yeast (SAGA and SALSA/SLIK) and three in humans (PCAF, STAGA, TFTC). It is important to note that GNAT or SAGA-like complexes may exist in flies and in mice [193,194]. Subunits of these complexes include Ada proteins, Spt proteins, TAFs, SAP130, and TRRAP. NuA3 (nucleosomal acetyltransferase of histone H3) is one of the yeast HAT complexes identified in the study carried out by Grant et al. [195]. It is a 500-kDa complex and it exclusively acetylates histone H3 in nucleosomes. Peptide sequencing of proteins from the purified NuA3 complex identified Sas3, a MYST protein involved in silencing, as the catalytic HAT subunit of the complex. NuA3 also contains the TBP-associated factor, yTAF(II)30. In addition, Yng1 was identified as a subunit of NuA3: it belongs to PHD finger-containing proteins and was recently found to interact with H3K4me3. The interaction between Yng1 and H3K4me3 seems to promote NuA3 HAT

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activity at K14 of H3 and transcription at a subset of targeted ORFs [196]. In vitro studies on NuA3, like those on Ada, indicated that both complexes failed to interact with activation domains or to activate transcription in a specific way [197,198]. NuA4 (nucleosomal acetyltransferase of histone H4)/TIP60 complex is another yeast HAT complex identified by Grant et al. [195] (complex 2), at the same time as SAGA, NuA3, and ADA. Its human homolog is called TIP60 [199]. As with Gcn5, NuA4 and TIP60 enzymes are able to acetylate histones H4, H3, and H2A when in free form but they are not capable of acetylating histones folded into nucleosomes and their activities seem to depend on the presence of other proteins in the context of multisubunit complexes. These complexes also seem to restrict their activities to histones H4 and H2A [199]. A homolog of these complexes was also recently identified in Drosophila (dmTIP60) [193,200]. Further studies identified three new complexes that share several subunits of the TIP60/ NuA4 complex. The first two complexes have been identified in humans in vivo and they are very similar to TIP60: these are the p400 complex [201] and another complex that contains TRRAP-BAF53-TIP48-TIP49 [202]. P400 is deprived of any HAT activity but can hydrolyze ATP [193]. The second complex has a HAT activity; however, the enzyme responsible for this activity is still not defined. The third complex was identified in yeast, and it represents a sort of “mini” NuA4 complex containing only three subunits (Tip60p/NuA4-Ing3-Epc1). This complex is termed “Piccolo NuA4” and its homolog also exists in humans and seems to represent the catalytic core of TIP60 [203].

Histone deacetylases
There are three distinct families of histone deacetylases: the class I and class II histone deacetylases, and the class III NAD-dependent enzymes of the Sir family. They are involved in multiple signaling pathways and are present in numerous repressive chromatin complexes. Similarly to HATs, these enzymes do not appear to show much specificity for a particular acetyl group. However, yeast enzyme Hda1 seems to have higher specificity for H3 and H2B whereas Hos2 is specific for H3 and H4. The fission yeast class III deacetylase Sir2 and its human homolog SirT2 preferentially deacetylate H4K16ac [204]. Recent reports indicated that Sir2/SirT2 is also able to deacetylate H3K56 [185].


Role of histone acetylation in transcription regulation
The “traditional” role of histone acetylation is transcription regulation. The first evidence of the involvement of HATs in transcription dates back to 1964, when it was observed that chromatin regions of actively transcribed genes tend to have hyperacetylated histones [6]. The addition of acetyl groups to histone tails was proposed to neutralize the histone charge, which weakens histone-DNA interaction, relaxing the chromatin structure and facilitating the access of transcription machinery [205]. For example, work from Craig Peterson’s laboratory demonstrated that the incorporation of H4K16Ac into nucleosomal arrays impedes the formation of compacted chromatin fibers and prevents the ATP-mediated chromatin remodeling factors from mediating nucleosome sliding [206,207]. In addition, two other mechanisms by which histone acetylation facilitates transcription have been proposed. First, there is evidence that histone acetylation may serve as a specific docking site for the recruitment of transcription regulators [208–211]. Second, histone acetylation may also act in combination with other histone modifications (methylation, phosphorylation, and ubiquitination) to form the “histone code” that dictates biological outcomes including gene transcription [9,212]. HAT complexes from both GNAT and MYST families were shown to be recruited to activator-bound nucleosomes resulting in transcriptional activation [89,213,214]. The recruitment of SAGA leads to acetylation of promoter-proximal H3, whereas recruitment of NuA4 results in a broader domain of H4 acetylation (?3?kbp) [214]. This hyperacetylation of histones was linked to transcription activation [215], and NuA4-dependent acetylation of histone H4 was shown to affect transcription of specific genes such as His4, Lys2 [216],

chapter 3 Mechanisms of Histone Modifications
ribosomal proteins, and heat-shock proteins [217]. Arabi and colleagues have shown that TRRAP (a subunit of many HAT complexes) is recruited by c-Myc to the promoters of Pol I transcribed genes. The recruitment of TRRAP leads to increased histone acetylation, followed by recruitment of RNA polymerase I and activation of rRNA transcription [218]. Interaction between several activators and Tra1 (yeast homologs of TRRAP) cofactor was demonstrated in yeast and this interaction is essential for efficient transcriptional activation [219]. For example, c-Myc binding correlates with regions of acetylated histones [220]. The effect of Myc oncoproteins on chromatin structure was studied in more details by Knoepfler and coworkers, who found that c-Myc and N-Myc are involved in the widespread maintenance of active chromatin, probably through upregulation of GCN5 [221]. In mammals, TRRAP has also been implicated in the regulation of transcription. For instance, TRRAP activates the transcription of target genes through the recruitment of Tip60 and Gcn5/PCAF to their promoters, thus acetylating histones H4 and H3, respectively [222,223]. H3K56Ac was also implicated in transcriptional activation. H3K56 residue is facing the major groove of the DNA within the nucleosome, so it is in a particularly good position to affect histone/DNA interactions when acetylated [224–226].

Role of histone acetylation in DNA repair
While the role of HAT enzymes in transcriptional regulation is well established [227–229], a plethora of recent reports has also implicated HATs and histone acetylation in DNA damage detection and DNA repair. TATA box-binding protein-free TAFII (TFTC), a complex containing Gcn5 HAT, appears to preferentially acetylate histone H3 in nucleosomes containing UV-damaged DNA in mammalian cells [230], whereas STAGA (SPT3-TAFII31GCN5L acetyltransferase), another Gcn5 containing HAT complex, associates with UV-damage-binding factor [231]. Yeast strains with mutations in the N-terminal tail of histone H4, a subject for acetylation, were shown to be deficient in both DNA DSB repair and replication-coupled repair, and Esa1 (catalytic component of the yeast NuA4) was found to be responsible for this acetylation. Tip60 (mammalian homolog of Esa1) was also shown to be important in DNA DSB repair following genotoxic stress [199]. In addition, mutations in Yng2, a component of the yeast NuA4 HAT complex, results in hypersensitivity to and inefficient repair of DNA damage caused by genotoxic agents that induce replication fork stall [232]. Finally, mutations in either specific lysine residues in histone H3 or the yeast acetyltransferase HAT1 result in hypersensitivity to DNA DSB-inducing agents [233]. Mechanistic data for the role of acetylation in DNA repair has arisen from several recent reports. Binding of the NuA4 HAT complex at sites of DNA damage and site-specific histone H4 acetylation were found to occur concomitantly with histone H2A phosphorylation after induction of DSBs [80,90]. Additionally, histone H3 acetylation is an abundant modification of newly synthesized histones and defects in this acetylation result in sensitivity to DNA damaging agents that cause DNA breaks during replication [224]. Furthermore, localized histone H3 and H4 acetylation and deacetylation is triggered by homology directed repair of DSBs. Consistent with this finding, Gcn5 and Esa1 HATs are recruited to chromatin around a DSB induced by HO endonuclease in yeast [234]. Alongside histone modification on the amino-terminal tails of histones, histone core modifications also play a role in DNA repair. This is exemplified by the role of H3K56 acetylation in response to DNA damage. In budding yeast acetylation of H3K56 is deposited on newly synthesized histones during S phase and disappears in G2. However, in the presence of DNA damage the deacetylases for H3K56, Hst3, and Hst4 (two paralogs of Sir2) are downregulated and the modification persists [235,236]. The Rtt109 enzyme, which acetylates H3K56, has recently been implicated in genome stability and DNA replication [225,237,238]. Furthermore, recent evidence has revealed that histone acetylation by TRRAP/TIP60 HAT is important for recruitment/loading of repair proteins to sites of DNA DSBs and homology-directed DNA repair [239]. These


section I Molecular Mechanisms of Epigenetics
findings lead to a model in which induction of DSBs leads to the recruitment of the TIP60/ NuA4 complex to DSBs and concomitant acetylation of H4 N-terminal tails [4,240].

Summary and perspectives
Research on chromatin modifications is a newly emerging field that holds the promise of further advancing our understanding of tumorigenesis and facilitating the development of novel strategies to prevent, diagnose, and treat cancer. Chromatin modifications act in a coordinated and orderly fashion to regulate cellular processes such as transcription, DNA replication, and DNA repair. These processes may be regulated by TRRAP/HAT and there is an intimate and self-reinforcing cross-talk and interdependence between histone-modifying complexes and other histone-modifying activities such as acetylation, phosphorylation, and methylation. Consistent with the critical function of histone modifications in key cellular processes, a large body of evidence has suggested that these complexes are intimately linked to human pathologies. Most notably, recent genetic and molecular studies have directly implicated histone modifications and histone-modifying complexes in human cancer. The fact that epigenetic alterations are, in contrast to genetic changes, reversible, has important implications for human cancer treatment as aberrant histone modifications are potential molecular targets for therapeutic intervention in human malignancies.

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