Histone marks mechanism

Histone marks mechanism

We are searching data for your request:

Forums and discussions:
Manuals and reference books:
Data from registers:
Wait the end of the search in all databases.
Upon completion, a link will appear to access the found materials.

I am slightly confused about the mechanisms that makes histone modifications associate with gene expression.

That is, H3K36me3 is believed to be present in actively transcribed genes, H3K27me3 in repressed regions, etc. What is the mechanism that makes the presence of H3K36me3 to be in active regions, etc.? Is it causal or just an association?

First we must remember what a nucleosome is: a segment of DNA packed with proteins called histones. This is the initial step to turn the DNA a more compact structure. Citing the advantages of an extremely compact but flexible structure like chromosomes would make this answer a little longer (if you want, I can add them later), and to answer your question faster, we'd better keep moving on the discussion.
Epigenetics and gene expression are so beautiful that they have many mechanisms to regulate or change gene expression, and you mention one, which is done by reversible covalent modifications on histones. Such modifications are executed by especific proteins, that can do methylation, acetylation, phosphorilation in some aminoacids of histones.
When an aminoacid of a histone is linked with charged groups (e.g: methyl, acetyl) its charge is altered, and depending of the resulting charge (negative or positive), it can reduce or increase the afinity between the histone tail and the adjacent nucleosomes, and therefore, make the nucleosome more compact and less accessible or less compact and more accessible, respectively.
But the main effect of these covalent modifications on histones is that the groups added to the aminoacids act like markers indicating for proteins involved in expression (like transcriptional factors and RNA polymerase) that some genes are switched on or switched off. The patterns of this modifications (which one repress or estimulate gene expression) is not completely understood yet.
An example of how a certain region is chosen to be marked for expression or repression:
Consider an organism that has received a repressed gene related to obesity from its parents and this organism is lazy, eats a lot, has a sedentary life. As a result of all this stimulation, the proteins responsible for altering the histones of the gene related to obesity can be activated or recruited by others proteins, and make covalent modifications in a way that the histones of that gene now are marked for it to be expressed. This organism can pass the new mark to the its children and they will have propensity to obesity due to the activated gene. (This final example was an experiment of epigenetics using rats, I saw a long time ago, so I can't remember the reference).
And about if it's casual or not, the marks of regulation in a sequence of DNA can't be casual. For each one or two modifications of histones there is an enzyme that will only do its function if some event is already triggered or if it's triggered during the cell life depending on the environment the cell is exposed to. So millions of cells of a same tissue that have the same function would not have the same pattern of expression for the main genes related with that function by chance. Therefore, regulation of gene expression is very refined and histone marks are one of the many mechanisms that together dictate if a gene is repressed or expressed.

Epigenetic Inheritance: Histone Bookmarks Across Generations

Multiple circuitries ensure that cells respond correctly to the environmental cues within defined cellular programs. There is increasing evidence suggesting that cellular memory for these adaptive processes can be passed on through cell divisions and generations. However, the mechanisms by which this epigenetic information is transferred remain elusive largely because it requires that such memory survive through gross chromatin remodeling events during DNA replication, mitosis, meiosis and developmental reprogramming. Elucidating the processes by which epigenetic information survives and is transmitted is a central challenge in biology. Here we consider recent advances in understanding mechanisms of epigenetic inheritance with a focus on histone segregation at the replication fork and how an epigenetic memory may get passed through the paternal lineage.


Eukaryote DNA is organized in a highly compact structure, chromatin, that consists of deoxyribonucleic acids and proteins. The DNA double helix is wound up around nucleosomes consisting of histone octamers, including two subunits each of histones H2A, H2B, H3 and H4. A plethora of proteins are involved in maintaining and regulating chromatin structure during DNA replication, transcription, repair, etc. DNA methylation, nucleosome positioning and reversible post-translational modifications of histone proteins govern the spatial organization and accessibility of DNA in chromatin in eukaryote cells. The post-translational modifications of histones, also known as histone marks, include methylation, acetylation, phosphorylation and other covalent chemical moieties that are (reversibly) conjugated to distinct amino acid residues in the histone proteins. These site-specific and co-existing modifications of multiple amino acid residues generate complex combinatorial patterns that may have functional roles in modulating chromatin structure and in the recruitment of specific protein co-factors to distinct domains in chromatin, thereby constituting a highly dynamic regulatory network [1]. Heterochromatin denotes the highly condensed inactive state of chromatin, where genes are repressed due to the inaccessibility of DNA for the transcription machinery. Abnormal function of the heterochromatic state has been linked to several diseases [2]–[4].

In the present work we address several fundamental questions in chromatin biology and histone structure/function relationships: (a) Are histone modifications organized in domains along the chromatin? (b) What is the minimal model able to simulate the formation of heterochromatin domains that is in accordance with experimental results? (c) What are the different mechanisms leading to changes of the histone modification landscape and which are able to switch genes on/off as response to external stimuli?

Several computational and/or mathematical approaches simulate a bistable state of histone modifications, for example switching between a state dominated by H3K9 methylation and the state dominated by H3K9 acetylations [5]–[8]. These studies concentrated on a general stability analysis and memory of such a system, thereby revealing ultrasensitive switching behavior. However, there was no direct comparison of those results to experimentally measured chromatin configurations. In another approach, the formation of multiply modified histones was described by stochastic nonlinear equations [9]. The analysis did not consider specific modifications as the model only counted the number of modifications on a histone without specifying their type. An epigenetic switch was modeled in ref. [10], where the authors studied switching and memory effects of the floral repressor of Arabidopsis with a simple mathematical model implementing nucleation and spreading of the silencing H3K27me3 mark. The data was successfully compared to ChIP data. Furthermore, simulations of the heterochromatin domain around the Oct4 locus in mouse ES cells and fibroblasts showed that this domain and most euchromatic H3K9me3 domains were well-described by a model based on propagation of the marks without taking into account specific boundary or insulator elements [11].

We go further and simulate the formation of heterochromatin over whole human chromosomes. The computer model implements the basic processes of nucleation, propagation and competition of histone marks through stochastic rates. We test whether such a simple model is able to generate stable domains of competing histone modifications. We then compare the results to experimental measurements and study the model's overall behavior.

In the following, we present biological evidence for the rules implemented in our computational model.


Non-protein-coding DNA sequences seem to play a crucial role to nucleate histone modification mediated domain formation. The RNA interference machinery shows activity at dh-dg repeats in yeast DNA [12], [13] leading to heterochromatin formation through a self-amplifying feed-forward regulatory mechanism [14], [15]. In higher eukaryotes, details about the initialization of heterochromatin remain unclear but strong correlations between heterochromatin and diverse satellite-repeats and transposable elements were observed [16], [17], as for instance with SINE-Alu elements in humans [18]. We will refer to these initiating sequences from now on as heterochromatin nucleation sites [19]. Within this scenario, these sequences contain regulatory information over gene transcription that can be fine-tuned to allow the development of different cell types [20], [21]. We will show how this information can be used to generate different heterochromatic states.

The presence of genomic CpG islands is strongly correlated to transcriptional activity, and makes CpG islands candidates for nucleation sites for transcriptionally activating histone marks. CpG islands exhibit a high abundance of demethylated DNA, enrichment of H3K4me2/3, H3K9ac and H3K14ac marks [22]–[24]. The underlying mechanisms involve the protein Cfp1 that associates with unmethylated CpG islands in vivo and recruits H3K4 methyltransferases to nearby histones [25]. SINEAlu elements and CpG appear to be experimentally well characterized nucleation sites. Other types of nucleation sites were neglected in the model.

According to these relations between DNA sequence and chromatin state, both repressive and activating histone marks are occurring in specific gene regions and are initiated by their respective nucleation sites. However, different cell types exhibit distinct transcriptome profiles reflected in different histone modifications [26]. Within chromatin, an effective regulatory mechanism is required for switching transcriptional activity of large genomic regions and for the formation of distinct transcription patterns. One might argue that the presence of predefined fixed nucleation sites for initiation of histone marks within the genome is incompatible with the presence of different transcriptional states. We demonstrate that predefined nucleation sites and histone modifications will indeed provide features that allow for a dynamic switching behavior of genes and genomic regions.


Chromosomes exhibit regions mediated by histone modifications that expand over considerable ranges along the DNA strand. Heterochromatin domains represent one type of these regions. Reinforcing mechanisms lead to the formation of heterochromatin domains enriched in H3K9me2 and H3K9me3 marks [27]. Di- and trimethylation of lysine 9 on histone H3 by the methyltransferases G9a and Suv39h1 reflect the repressed state of heterochromatin [28]–[30] that is maintained by several proteins through a positive feedback loop [31], [32]. Heterochromatin protein-1 (HP1) recognizes H3K9 methylation [33]–[35] and it interacts with Suv39h1 [36], [37], that is recruited by neighboring H3K9me2 sites [35] and thereby stabilizes the heterochromatin state. HP1 also recruits the DNA methyltransferases DNMT1 that itself associates with G9a. G9a sets the H3K9me2 mark [38], [39]. HP1 establishes the spatially dense chromatin structure and recruits histone deacetylases and DNA methyltransferases to strengthen this state [40], [41]. This loop is further stimulated as HP1 associates to itself [42] leading to the propagation of H3K9me2 and H3K9me3.

Transcriptionally active regions contain not only H3K4me3 but also the other methylation states H3K4me2 and H3K4me1. It was found that the USF protein binds to specific DNA sequences and mediates H3K4 methylation and histone acetylation [43], which are both related to gene activation. There is strong correlation between H3K4 methylations and e.g. the acetylation marks H3K14ac, H3K18ac [44], [45]. Generally, local recruitment of histone acetyltransferases activities seems to counteract the spreading of heterochromatin [46]–[48]. This whole machinery suggests a propagation mechanism for euchromatin formation.


How are the efficient molecular processes for the propagation of heterochromatin domains prevented from reaching a state where heterochromatin completely occupies the chromosomes? A total occupation would lead to a complete shut-down of gene transcription. Boundary elements, also called insulators, are defined as genetic regions where the propagation of histone marks are stalled. In a DNA strand with only nearest-neighbor interactions, the boundary must permanently flank both ends of the to-be-protected domain to shield it from silencing [49].

Passive insulators that prevent the setting of H3K9 methylation or HP1 association without actively recruiting histone-modifying enzymes are not able to stop heterochromatin propagation [50]. The condensed three-dimensional conformation of the DNA strand allows propagation of heterochromatin marks between co-located non-neighboring nucleosomes leading to the propagation into the to-be-insulated region. Furthermore, passive insulators would be required to form a static and stable barrier. Otherwise, heterochromatin domains would be able to temporally spread into the region. Hence, effective insulation can only work when the insulator actively maintains a region of histone marks that antagonize the setting of the heterochromatin forming constituents on the entire region.

Wang et al. [51] present a model that explains both fixed boundaries, with a specific actively recruiting boundary element, and broader ones where euchromatin related histone modifications gradually change in to the ones related to the repressive state. We will show that both scenarios are possible in our simulations, mainly depending on the distribution of nucleation sites.

Histone marks that are related to gene activation seem also to be main players in controlling heterochromatin formation. Domains decorated with H3 and H4 acetylation marks and the H3K4 methylation mark prevent heterochromatin from spreading over the entire chromosomes [46], [52]. H3K4me3 competes directly with the heterochromatin state as it inhibits the methylation of H3K9 by Suv39h1 [53], [54]. Similarly, H3K9ac inhibits histone deacetylases and interrupts the interaction between HP1 and chromatin [55].

After identifying the processes propagation, nucleation and competition as main actors in histone domain formation, it is possible to construct a theoretical model for the distribution of active and inactive chromatin domains by assuming that the competing histone domains become initiated from their respective nucleation sites. We will test this assumption with a computational model that implements the basic underlying rules.

The Discovery of Histone Lys Crotonylation

When histone Kcr was first identified, it was found to be evolutionary conserved from yeast to human, occurring broadly in all core histones (H2A, H2B, H3, and H4), as well as linker histone H1 and marked active promoters and potential enhancers (Tan et al., 2011). Similar to Kac, Kcr also occurs on the ε-amino group of the lysine side chain, where it neutralizes the positive charge of this residue. The loss in positive charge on histone Lys residues weakens DNA interaction, thus making chromatin less compact and accessible to DNA-binding factors. In support of a potential cis-function of Kcr on chromatin structure, H3K122cr-H4 containing tetrasomes that were subjected to thermal stability assays, were found to be less stable compared to unmodified H3-H4 tetrasomes (Suzuki et al., 2016). Consistent with this, the ability of Kcr to destabilise nucleosome structure has been proposed to be part of a compensatory mechanism during chromatin-to-nucleoprotamine transition, an essential process during spermatogenesis as discussed in the spermatogenesis section below (Montellier et al., 2013). Montellier et al. (2013) showed that incorporation of a histone H2B variant, TH2B, is essential for the final transformation of dissociating nucleosomes into protamine packed structures. In the absence of TH2B, cells compensate by upregulating H2B and programming nucleosome instability to reach that of wild type cells through targeted histone modifications, including crotonylation of H3K122 and H4K77. This in turn allows the histone replacement to take place. Furthermore, modified histone lysine residues can mediate trans-effects through recruitment of effector proteins containing specific reader modules. This is particularly important for histone Kcr, where the crotonyl group is a four-carbon chain containing a C𠄼 π bond that results in a rigid planar conformation, which is unique among histone acylations. The extended hydrocarbon chain of the crotonyl group increases the hydrophobicity and bulk of the Lys residue compared to acetylation (Sabari et al., 2017). These differences in the biophysical properties of the crotonyl group provide an important mechanism of specificity for reader interaction, as described in detail below.

The Discovery of Histone Lys Crotonylation

When histone Kcr was first identified, it was found to be evolutionary conserved from yeast to human, occurring broadly in all core histones (H2A, H2B, H3, and H4), as well as linker histone H1 and marked active promoters and potential enhancers (Tan et al., 2011). Similar to Kac, Kcr also occurs on the ε-amino group of the lysine side chain, where it neutralizes the positive charge of this residue. The loss in positive charge on histone Lys residues weakens DNA interaction, thus making chromatin less compact and accessible to DNA-binding factors. In support of a potential cis-function of Kcr on chromatin structure, H3K122cr-H4 containing tetrasomes that were subjected to thermal stability assays, were found to be less stable compared to unmodified H3-H4 tetrasomes (Suzuki et al., 2016). Consistent with this, the ability of Kcr to destabilise nucleosome structure has been proposed to be part of a compensatory mechanism during chromatin-to-nucleoprotamine transition, an essential process during spermatogenesis as discussed in the spermatogenesis section below (Montellier et al., 2013). Montellier et al. (2013) showed that incorporation of a histone H2B variant, TH2B, is essential for the final transformation of dissociating nucleosomes into protamine packed structures. In the absence of TH2B, cells compensate by upregulating H2B and programming nucleosome instability to reach that of wild type cells through targeted histone modifications, including crotonylation of H3K122 and H4K77. This in turn allows the histone replacement to take place. Furthermore, modified histone lysine residues can mediate trans-effects through recruitment of effector proteins containing specific reader modules. This is particularly important for histone Kcr, where the crotonyl group is a four-carbon chain containing a C𠄼 π bond that results in a rigid planar conformation, which is unique among histone acylations. The extended hydrocarbon chain of the crotonyl group increases the hydrophobicity and bulk of the Lys residue compared to acetylation (Sabari et al., 2017). These differences in the biophysical properties of the crotonyl group provide an important mechanism of specificity for reader interaction, as described in detail below.


Five major families of histones exist: H1/H5, H2A, H2B, H3, and H4. [2] [4] [5] [6] Histones H2A, H2B, H3 and H4 are known as the core histones, while histones H1/H5 are known as the linker histones.

The core histones all exist as dimers, which are similar in that they all possess the histone fold domain: three alpha helices linked by two loops. It is this helical structure that allows for interaction between distinct dimers, particularly in a head-tail fashion (also called the handshake motif). [7] The resulting four distinct dimers then come together to form one octameric nucleosome core, approximately 63 Angstroms in diameter (a solenoid (DNA)-like particle). Around 146 base pairs (bp) of DNA wrap around this core particle 1.65 times in a left-handed super-helical turn to give a particle of around 100 Angstroms across. [8] The linker histone H1 binds the nucleosome at the entry and exit sites of the DNA, thus locking the DNA into place [9] and allowing the formation of higher order structure. The most basic such formation is the 10 nm fiber or beads on a string conformation. This involves the wrapping of DNA around nucleosomes with approximately 50 base pairs of DNA separating each pair of nucleosomes (also referred to as linker DNA). Higher-order structures include the 30 nm fiber (forming an irregular zigzag) and 100 nm fiber, these being the structures found in normal cells. During mitosis and meiosis, the condensed chromosomes are assembled through interactions between nucleosomes and other regulatory proteins.

Histones are subdivided into canonical replication-dependent histones that are expressed during the S-phase of the cell cycle and replication-independent histone variants, expressed during the whole cell cycle. In animals, genes encoding canonical histones are typically clustered along the chromosome, lack introns and use a stem loop structure at the 3' end instead of a polyA tail. Genes encoding histone variants are usually not clustered, have introns and their mRNAs are regulated with polyA tails. Complex multicellular organisms typically have a higher number of histone variants providing a variety of different functions. Recent data are accumulating about the roles of diverse histone variants highlighting the functional links between variants and the delicate regulation of organism development. [10] Histone variants from different organisms, their classification and variant specific features can be found in "HistoneDB 2.0 - Variants" database.

The following is a list of human histone proteins:

Super family Family Subfamily Members
Linker H1 H1F H1F0, H1FNT, H1FOO, H1FX

The nucleosome core is formed of two H2A-H2B dimers and a H3-H4 tetramer, forming two nearly symmetrical halves by tertiary structure (C2 symmetry one macromolecule is the mirror image of the other). [8] The H2A-H2B dimers and H3-H4 tetramer also show pseudodyad symmetry. The 4 'core' histones (H2A, H2B, H3 and H4) are relatively similar in structure and are highly conserved through evolution, all featuring a 'helix turn helix turn helix' motif (DNA-binding protein motif that recognize specific DNA sequence). They also share the feature of long 'tails' on one end of the amino acid structure - this being the location of post-translational modification (see below). [11]

Archaeal histone only contains a H3-H4 like dimeric structure made out of the same protein. Such dimeric structures can stack into a tall superhelix ("hypernucleosome") onto which DNA coils in a manner similar to nucleosome spools. [12] Only some archaeal histones have tails. [13]

The distance between the spools around which eukaryotic cells wind their DNA has been determined to range from 59 to 70 Å. [14]

In all, histones make five types of interactions with DNA:

    and hydrogen bonds between side chains of basic amino acids (especially lysine and arginine) and phosphate oxygens on DNA
  • Helix-dipoles form alpha-helixes in H2B, H3, and H4 cause a net positive charge to accumulate at the point of interaction with negatively charged phosphate groups on DNA between the DNA backbone and the amide group on the main chain of histone proteins
  • Nonpolar interactions between the histone and deoxyribose sugars on DNA
  • Non-specific minor groove insertions of the H3 and H2B N-terminal tails into two minor grooves each on the DNA molecule

The highly basic nature of histones, aside from facilitating DNA-histone interactions, contributes to their water solubility.

Histones are subject to post translational modification by enzymes primarily on their N-terminal tails, but also in their globular domains. [15] [16] Such modifications include methylation, citrullination, acetylation, phosphorylation, SUMOylation, ubiquitination, and ADP-ribosylation. This affects their function of gene regulation.

In general, genes that are active have less bound histone, while inactive genes are highly associated with histones during interphase. [17] It also appears that the structure of histones has been evolutionarily conserved, as any deleterious mutations would be severely maladaptive. All histones have a highly positively charged N-terminus with many lysine and arginine residues.

Core histones are found in the nuclei of eukaryotic cells and in most Archaeal phyla, but not in bacteria. [13] However the linker histones have homologs in bacteria. [18] The unicellular algae known as dinoflagellates were previously thought to be the only eukaryotes that completely lack histones, [19] however, later studies showed that their DNA still encodes histone genes. [20] Unlike the core histones, lysine-rich linker histone (H1) proteins are found in bacteria, otherwise known as nucleoprotein HC1/HC2. [18]

It has been proposed that histone proteins are evolutionarily related to the helical part of the extended AAA+ ATPase domain, the C-domain, and to the N-terminal substrate recognition domain of Clp/Hsp100 proteins. Despite the differences in their topology, these three folds share a homologous helix-strand-helix (HSH) motif. [11]

Archaeal histones may well resemble the evolutionary precursors to eukaryotic histones. [13] Furthermore, the nucleosome (core) histones may have evolved from ribosomal proteins (RPS6/RPS15) with which they share much in common, both being short and basic proteins. [21] Histone proteins are among the most highly conserved proteins in eukaryotes, emphasizing their important role in the biology of the nucleus. [2] : 939 In contrast mature sperm cells largely use protamines to package their genomic DNA, most likely because this allows them to achieve an even higher packaging ratio. [22]

There are some variant forms in some of the major classes. They share amino acid sequence homology and core structural similarity to a specific class of major histones but also have their own feature that is distinct from the major histones. These minor histones usually carry out specific functions of the chromatin metabolism. For example, histone H3-like CENPA is associated with only the centromere region of the chromosome. Histone H2A variant H2A.Z is associated with the promoters of actively transcribed genes and also involved in the prevention of the spread of silent heterochromatin. [23] Furthermore, H2A.Z has roles in chromatin for genome stability. [24] Another H2A variant H2A.X is phosphorylated at S139 in regions around double-strand breaks and marks the region undergoing DNA repair. [25] Histone H3.3 is associated with the body of actively transcribed genes. [26]

Compacting DNA strands Edit

Histones act as spools around which DNA winds. This enables the compaction necessary to fit the large genomes of eukaryotes inside cell nuclei: the compacted molecule is 40,000 times shorter than an unpacked molecule.

Chromatin regulation Edit

Histones undergo posttranslational modifications that alter their interaction with DNA and nuclear proteins. The H3 and H4 histones have long tails protruding from the nucleosome, which can be covalently modified at several places. Modifications of the tail include methylation, acetylation, phosphorylation, ubiquitination, SUMOylation, citrullination, and ADP-ribosylation. The core of the histones H2A and H2B can also be modified. Combinations of modifications are thought to constitute a code, the so-called "histone code". [27] [28] Histone modifications act in diverse biological processes such as gene regulation, DNA repair, chromosome condensation (mitosis) and spermatogenesis (meiosis). [29]

The common nomenclature of histone modifications is:

  • The name of the histone (e.g., H3)
  • The single-letter amino acid abbreviation (e.g., K for Lysine) and the amino acid position in the protein
  • The type of modification (Me: methyl, P: phosphate, Ac: acetyl, Ub: ubiquitin)
  • The number of modifications (only Me is known to occur in more than one copy per residue. 1, 2 or 3 is mono-, di- or tri-methylation)

So H3K4me1 denotes the monomethylation of the 4th residue (a lysine) from the start (i.e., the N-terminal) of the H3 protein.

Examples of histone modifications in transcriptional regulation
Type of
H3K4 H3K9 H3K14 H3K27 H3K79 H3K36 H4K20 H2BK5 H2BK20
mono-methylation activation [30] activation [31] activation [31] activation [31] [32] activation [31] activation [31]
di-methylation repression [33] repression [33] activation [32]
tri-methylation activation [34] repression [31] repression [31] activation, [32]
repression [31]
activation repression [33]
acetylation activation [35] activation [34] activation [34] activation [36] activation

A huge catalogue of histone modifications have been described, but a functional understanding of most is still lacking. Collectively, it is thought that histone modifications may underlie a histone code, whereby combinations of histone modifications have specific meanings. However, most functional data concerns individual prominent histone modifications that are biochemically amenable to detailed study.

Chemistry Edit

Lysine methylation Edit

The addition of one, two, or many methyl groups to lysine has little effect on the chemistry of the histone methylation leaves the charge of the lysine intact and adds a minimal number of atoms so steric interactions are mostly unaffected. However, proteins containing Tudor, chromo or PHD domains, amongst others, can recognise lysine methylation with exquisite sensitivity and differentiate mono, di and tri-methyl lysine, to the extent that, for some lysines (e.g.: H4K20) mono, di and tri-methylation appear to have different meanings. Because of this, lysine methylation tends to be a very informative mark and dominates the known histone modification functions.

Glutamine serotonylation Edit

Recently it has been shown, that the addition of a serotonin group to the position 5 glutamine of H3, happens in serotonergic cells such as neurons. This is part of the differentiation of the serotonergic cells. This post-translational modification happens in conjunction with the H3K4me3 modification. The serotonylation potentiates the binding of the general transcription factor TFIID to the TATA box. [37]

Arginine methylation Edit

What was said above of the chemistry of lysine methylation also applies to arginine methylation, and some protein domains—e.g., Tudor domains—can be specific for methyl arginine instead of methyl lysine. Arginine is known to be mono- or di-methylated, and methylation can be symmetric or asymmetric, potentially with different meanings.

Arginine citrullination Edit

Enzymes called peptidylarginine deiminases (PADs) hydrolyze the imine group of arginines and attach a keto group, so that there is one less positive charge on the amino acid residue. This process has been involved in the activation of gene expression by making the modified histones less tightly bound to DNA and thus making the chromatin more accessible. [38] PADs can also produce the opposite effect by removing or inhibiting mono-methylation of arginine residues on histones and thus antagonizing the positive effect arginine methylation has on transcriptional activity. [39]

Lysine acetylation Edit

Addition of an acetyl group has a major chemical effect on lysine as it neutralises the positive charge. This reduces electrostatic attraction between the histone and the negatively charged DNA backbone, loosening the chromatin structure highly acetylated histones form more accessible chromatin and tend to be associated with active transcription. Lysine acetylation appears to be less precise in meaning than methylation, in that histone acetyltransferases tend to act on more than one lysine presumably this reflects the need to alter multiple lysines to have a significant effect on chromatin structure. The modification includes H3K27ac.

Serine/threonine/tyrosine phosphorylation Edit

Addition of a negatively charged phosphate group can lead to major changes in protein structure, leading to the well-characterised role of phosphorylation in controlling protein function. It is not clear what structural implications histone phosphorylation has, but histone phosphorylation has clear functions as a post-translational modification, and binding domains such as BRCT have been characterised.

Effects on transcription Edit

Most well-studied histone modifications are involved in control of transcription.

Actively transcribed genes Edit

Two histone modifications are particularly associated with active transcription:

Trimethylation of H3 lysine 4 (H3K4me3) This trimethylation occurs at the promoter of active genes [40] [41] [42] and is performed by the COMPASS complex. [43] [44] [45] Despite the conservation of this complex and histone modification from yeast to mammals, it is not entirely clear what role this modification plays. However, it is an excellent mark of active promoters and the level of this histone modification at a gene's promoter is broadly correlated with transcriptional activity of the gene. The formation of this mark is tied to transcription in a rather convoluted manner: early in transcription of a gene, RNA polymerase II undergoes a switch from initiating' to 'elongating', marked by a change in the phosphorylation states of the RNA polymerase II C terminal domain (CTD). The same enzyme that phosphorylates the CTD also phosphorylates the Rad6 complex, [46] [47] which in turn adds a ubiquitin mark to H2B K123 (K120 in mammals). [48] H2BK123Ub occurs throughout transcribed regions, but this mark is required for COMPASS to trimethylate H3K4 at promoters. [49] [50] Trimethylation of H3 lysine 36 (H3K36me3) This trimethylation occurs in the body of active genes and is deposited by the methyltransferase Set2. [51] This protein associates with elongating RNA polymerase II, and H3K36Me3 is indicative of actively transcribed genes. [52] H3K36Me3 is recognised by the Rpd3 histone deacetylase complex, which removes acetyl modifications from surrounding histones, increasing chromatin compaction and repressing spurious transcription. [53] [54] [55] Increased chromatin compaction prevents transcription factors from accessing DNA, and reduces the likelihood of new transcription events being initiated within the body of the gene. This process therefore helps ensure that transcription is not interrupted.

Repressed genes Edit

Three histone modifications are particularly associated with repressed genes:

Trimethylation of H3 lysine 27 (H3K27me3) This histone modification is deposited by the polycomb complex PRC2. [56] It is a clear marker of gene repression, [57] and is likely bound by other proteins to exert a repressive function. Another polycomb complex, PRC1, can bind H3K27me3 [57] and adds the histone modification H2AK119Ub which aids chromatin compaction. [58] [59] Based on this data it appears that PRC1 is recruited through the action of PRC2, however, recent studies show that PRC1 is recruited to the same sites in the absence of PRC2. [60] [61] Di and tri-methylation of H3 lysine 9 (H3K9me2/3) H3K9me2/3 is a well-characterised marker for heterochromatin, and is therefore strongly associated with gene repression. The formation of heterochromatin has been best studied in the yeast Schizosaccharomyces pombe, where it is initiated by recruitment of the RNA-induced transcriptional silencing (RITS) complex to double stranded RNAs produced from centromeric repeats. [62] RITS recruits the Clr4 histone methyltransferase which deposits H3K9me2/3. [63] This process is called histone methylation. H3K9Me2/3 serves as a binding site for the recruitment of Swi6 (heterochromatin protein 1 or HP1, another classic heterochromatin marker) [64] [65] which in turn recruits further repressive activities including histone modifiers such as histone deacetylases and histone methyltransferases. [66] Trimethylation of H4 lysine 20 (H4K20me3) This modification is tightly associated with heterochromatin, [67] [68] although its functional importance remains unclear. This mark is placed by the Suv4-20h methyltransferase, which is at least in part recruited by heterochromatin protein 1. [67]

Bivalent promoters Edit

Analysis of histone modifications in embryonic stem cells (and other stem cells) revealed many gene promoters carrying both H3K4Me3 and H3K27Me3, in other words these promoters display both activating and repressing marks simultaneously. This peculiar combination of modifications marks genes that are poised for transcription they are not required in stem cells, but are rapidly required after differentiation into some lineages. Once the cell starts to differentiate, these bivalent promoters are resolved to either active or repressive states depending on the chosen lineage. [69]

Other functions Edit

DNA damage Edit

Marking sites of DNA damage is an important function for histone modifications. It also protects DNA from getting destroyed by ultraviolet radiation of sun.

Phosphorylation of H2AX at serine 139 (γH2AX) Phosphorylated H2AX (also known as gamma H2AX) is a marker for DNA double strand breaks, [70] and forms part of the response to DNA damage. [25] [71] H2AX is phosphorylated early after detection of DNA double strand break, and forms a domain extending many kilobases either side of the damage. [70] [72] [73] Gamma H2AX acts as a binding site for the protein MDC1, which in turn recruits key DNA repair proteins [74] (this complex topic is well reviewed in [75] ) and as such, gamma H2AX forms a vital part of the machinery that ensures genome stability. Acetylation of H3 lysine 56 (H3K56Ac) H3K56Acx is required for genome stability. [76] [77] H3K56 is acetylated by the p300/Rtt109 complex, [78] [79] [80] but is rapidly deacetylated around sites of DNA damage. H3K56 acetylation is also required to stabilise stalled replication forks, preventing dangerous replication fork collapses. [81] [82] Although in general mammals make far greater use of histone modifications than microorganisms, a major role of H3K56Ac in DNA replication exists only in fungi, and this has become a target for antibiotic development. [83]

DNA repair Edit

H3K36me3 has the ability to recruit the MSH2-MSH6 (hMutSα) complex of the DNA mismatch repair pathway. [84] Consistently, regions of the human genome with high levels of H3K36me3 accumulate less somatic mutations due to mismatch repair activity. [85]

Chromosome condensation Edit

Addiction Edit

Epigenetic modifications of histone tails in specific regions of the brain are of central importance in addictions. [91] [92] [93] Once particular epigenetic alterations occur, they appear to be long lasting "molecular scars" that may account for the persistence of addictions. [91]

Cigarette smokers (about 15% of the US population) are usually addicted to nicotine. [94] After 7 days of nicotine treatment of mice, acetylation of both histone H3 and histone H4 was increased at the FosB promoter in the nucleus accumbens of the brain, causing 61% increase in FosB expression. [95] This would also increase expression of the splice variant Delta FosB. In the nucleus accumbens of the brain, Delta FosB functions as a "sustained molecular switch" and "master control protein" in the development of an addiction. [96] [97]

About 7% of the US population is addicted to alcohol. In rats exposed to alcohol for up to 5 days, there was an increase in histone 3 lysine 9 acetylation in the pronociceptin promoter in the brain amygdala complex. This acetylation is an activating mark for pronociceptin. The nociceptin/nociceptin opioid receptor system is involved in the reinforcing or conditioning effects of alcohol. [98]

Methamphetamine addiction occurs in about 0.2% of the US population. [99] Chronic methamphetamine use causes methylation of the lysine in position 4 of histone 3 located at the promoters of the c-fos and the C-C chemokine receptor 2 (ccr2) genes, activating those genes in the nucleus accumbens (NAc). [100] c-fos is well known to be important in addiction. [101] The ccr2 gene is also important in addiction, since mutational inactivation of this gene impairs addiction. [100]

The first step of chromatin structure duplication is the synthesis of histone proteins: H1, H2A, H2B, H3, H4. These proteins are synthesized during S phase of the cell cycle. There are different mechanisms which contribute to the increase of histone synthesis.

Yeast Edit

Yeast carry one or two copies of each histone gene, which are not clustered but rather scattered throughout chromosomes. Histone gene transcription is controlled by multiple gene regulatory proteins such as transcription factors which bind to histone promoter regions. In budding yeast, the candidate gene for activation of histone gene expression is SBF. SBF is a transcription factor that is activated in late G1 phase, when it dissociates from its repressor Whi5. This occurs when Whi5 is phosphorylated by Cdc8 which is a G1/S Cdk. [102] Suppression of histone gene expression outside of S phases is dependent on Hir proteins which form inactive chromatin structure at the locus of histone genes, causing transcriptional activators to be blocked. [103] [104]

Metazoan Edit

In metazoans the increase in the rate of histone synthesis is due to the increase in processing of pre-mRNA to its mature form as well as decrease in mRNA degradation this results in an increase of active mRNA for translation of histone proteins. The mechanism for mRNA activation has been found to be the removal of a segment of the 3' end of the mRNA strand, and is dependent on association with stem-loop binding protein (SLBP). [105] SLBP also stabilizes histone mRNAs during S phase by blocking degradation by the 3'hExo nuclease. [106] SLBP levels are controlled by cell-cycle proteins, causing SLBP to accumulate as cells enter S phase and degrade as cells leave S phase. SLBP are marked for degradation by phosphorylation at two threonine residues by cyclin dependent kinases, possibly cyclin A/ cdk2, at the end of S phase. [107] Metazoans also have multiple copies of histone genes clustered on chromosomes which are localized in structures called Cajal bodies as determined by genome-wide chromosome conformation capture analysis (4C-Seq). [108]

Link between cell-cycle control and synthesis Edit

Nuclear protein Ataxia-Telangiectasia (NPAT), also known as nuclear protein coactivator of histone transcription, is a transcription factor which activates histone gene transcription on chromosomes 1 and 6 of human cells. NPAT is also a substrate of cyclin E-Cdk2, which is required for the transition between G1 phase and S phase. NPAT activates histone gene expression only after it has been phosphorylated by the G1/S-Cdk cyclin E-Cdk2 in early S phase. [109] This shows an important regulatory link between cell-cycle control and histone synthesis.

Histones were discovered in 1884 by Albrecht Kossel. [110] The word "histone" dates from the late 19th century and is derived from the German word "Histon", a word itself of uncertain origin - perhaps from the Greek histanai or histos.

In the early 1960s, before the types of histones were known and before histones were known to be highly conserved across taxonomically diverse organisms, James F. Bonner and his collaborators began a study of these proteins that were known to be tightly associated with the DNA in the nucleus of higher organisms. [111] Bonner and his postdoctoral fellow Ru Chih C. Huang showed that isolated chromatin would not support RNA transcription in the test tube, but if the histones were extracted from the chromatin, RNA could be transcribed from the remaining DNA. [112] Their paper became a citation classic. [113] Paul T'so and James Bonner had called together a World Congress on Histone Chemistry and Biology in 1964, in which it became clear that there was no consensus on the number of kinds of histone and that no one knew how they would compare when isolated from different organisms. [114] [111] Bonner and his collaborators then developed methods to separate each type of histone, purified individual histones, compared amino acid compositions in the same histone from different organisms, and compared amino acid sequences of the same histone from different organisms in collaboration with Emil Smith from UCLA. [115] For example, they found Histone IV sequence to be highly conserved between peas and calf thymus. [115] However, their work on the biochemical characteristics of individual histones did not reveal how the histones interacted with each other or with DNA to which they were tightly bound. [114]

Also in the 1960s, Vincent Allfrey and Alfred Mirsky had suggested, based on their analyses of histones, that acetylation and methylation of histones could provide a transcriptional control mechanism, but did not have available the kind of detailed analysis that later investigators were able to conduct to show how such regulation could be gene-specific. [116] Until the early 1990s, histones were dismissed by most as inert packing material for eukaryotic nuclear DNA, a view based in part on the models of Mark Ptashne and others, who believed that transcription was activated by protein-DNA and protein-protein interactions on largely naked DNA templates, as is the case in bacteria.

During the 1980s, Yahli Lorch and Roger Kornberg [117] showed that a nucleosome on a core promoter prevents the initiation of transcription in vitro, and Michael Grunstein [118] demonstrated that histones repress transcription in vivo, leading to the idea of the nucleosome as a general gene repressor. Relief from repression is believed to involve both histone modification and the action of chromatin-remodeling complexes. Vincent Allfrey and Alfred Mirsky had earlier proposed a role of histone modification in transcriptional activation, [119] regarded as a molecular manifestation of epigenetics. Michael Grunstein [120] and David Allis [121] found support for this proposal, in the importance of histone acetylation for transcription in yeast and the activity of the transcriptional activator Gcn5 as a histone acetyltransferase.

The discovery of the H5 histone appears to date back to the 1970s, [122] and it is now considered an isoform of Histone H1. [2] [4] [5] [6]

Epigenetics and Development

Histone Methylation

Histone methylation occurs when methyltransferases add a methyl group to arginine or lysine (or possibly histidine) residues (reviewed in Bannister and Kouzarides, 2011 reviewed in Greer and Shi, 2012 ). In addition to methylation, several proposed methods for histone demethylation have been confirmed (reviewed in Bannister and Kouzarides, 2011 ). Unlike acetyl groups, methyl groups do not change the histone's charge (reviewed in Bannister and Kouzarides, 2011 ). Additionally, methyltransferases can add multiple methyl groups to a single arginine or lysine, mono-, di-, or even, in the case of lysine, tri-methylating the residue while histidines have only been found to be monomethylated (reviewed in Bannister and Kouzarides, 2011 reviewed in Greer and Shi, 2012 ). The altered gene expression resulting from histone methylation is not as clear cut as with histone acetylation. Both the location of the methyl group and whether that location is mono-, di-, or tri-methylated can determine whether the result is an increase or decrease in gene expression (reviewed in Greer and Shi, 2012 ). At some locations, a certain histone methylation marker may lead to both gene expression and repression at different times (reviewed in Greer and Shi, 2012 ).

Histone lysine methyltransferases in biology and disease

The precise temporal and spatial coordination of histone lysine methylation dynamics across the epigenome regulates virtually all DNA-templated processes. A large number of histone lysine methyltransferase (KMT) enzymes catalyze the various lysine methylation events decorating the core histone proteins. Mutations, genetic translocations and altered gene expression involving these KMTs are frequently observed in cancer, developmental disorders and other pathologies. Therapeutic compounds targeting specific KMTs are currently being tested in the clinic, although overall drug discovery in the field is relatively underdeveloped. Here we review the biochemical and biological activities of histone KMTs and their connections to human diseases, focusing on cancer. We also discuss the scientific and clinical challenges and opportunities in studying KMTs.

Lysine methylation was first described in 1959 on a bacterial flagellar protein 5 and soon thereafter identified on histone proteins 6 . Indeed, the core histones contain numerous evolutionarily conserved lysine residues that are methylated in vivo. In humans, the canonical lysine methylation sites are found on histone H3 at lysine 4 (H3K4), lysine 9 (H3K9), lysine 27 (H3K27), lysine 36 (H3K36) and lysine 79 (H3K79), and on histone H4 at lysine 20 (H4K20). These modifications regulate an array of chromatin functions (Fig. 1b) 1 . In addition to these canonical sites, there are several less well characterized sites of lysine methylation on the core histones (for example, H3K23me, H3K63me3, H45me1 and H4K12me1) (Fig. 1c) 4,7 . Together, the substantial numbers of methylation sites and differentially methylated states present in histones illustrate the potential complexity that this signaling system can provide in the regulation of chromatin biology and how its deregulation can lead to disease.

Change history

Almenara J, Rosato R, Grant S (2002) Synergistic induction of mitochondrial damage and apoptosis in human leukemia cells by flavopiridol and the histone deacetylase inhibitor suberoylanilide hydroxamic acid (SAHA). Leukemia 16: 1331–1343

Bali P, Pranpat M, Bradner J, Balasis M, Fiskus W, Guo F, Rocha K, Kumaraswamy S, Boyapalle S, Atadja P, Seto E, Bhalla K (2005) Inhibition of histone deacetylase 6 acetylates and disrupts the chaperone function of heat shock protein 90: a novel basis of antileukemia activity of histone deacetylase inhibitors. J Biol Chem 280: 26729–26734

Bereshchenko OR, Gu W, Dalla-Favera R (2002) Acetylation inactivates the transcriptional repressor BCL6. Nat Genet 32: 606–613

Butler LM, Agus DB, Scher HI, Higgins B, Rose A, Cordon-Cardo C, Thaler HT, Rifkind RA, Marks PA, Richon VM (2000) Suberoylanilide hydroxamic acid, an inhibitor of histone deacetylase, suppresses the growth of prostate cancer cells in vitro and in vivo. Cancer Res 60: 5165–5170

Chinnaiyan P, Vallabhaneni G, Armstrong E, Huang SM, Harari PM (2005) Modulation of radiation response by histone deacetylase inhibition. Int J Radiat Oncol Biol Phys 62: 223–229

Cohen LA, Amin S, Marks PA, Rifkind RA, Desai D, Richon VM (1999) Chemoprevention of carcinogen-induced mammary tumorigenesis by the hybrid polar cytodifferentiation agent, suberanilohydroxamic acid (SAHA). Anticancer Res 19: 4999–5005

Cohen LA, Marks PA, Rifkind RA, Amin S, Desai D, Pittman B, Richon VM (2002) Suberoylanilide hydroxamic acid (SAHA), a histone deacetylase inhibitor, suppresses the growth of carcinogen-induced mammary tumors. Anticancer Res 22: 1497–1504

Desai D, Das A, Cohen L, el Bayoumy K, Amin S (2003) Chemopreventive efficacy of suberoylanilide hydroxamic acid (SAHA) against 4-(methylnitrosamino)-1-(3-pyridyl)-1-butanone (NNK)-induced lung tumorigenesis in female A/J mice. Anticancer Res 23: 499–503

Duvic M, Talpur R, Zhang C, Goy A, Richon VM, Frankel SR (2005) Phase II trial of oral suberoylanilide hydroxamic acid (SAHA) for cutaneous T-cell lymphoma (CTCL) unresponsive to conventional therapy. J Clin Oncol 23 (suppl): 577s (abstract 6571)

Eyupoglu IY, Hahnen E, Buslei R, Siebzehnrubl FA, Savaskan NE, Luders M, Trankle C, Wick W, Weller M, Fahlbusch R, Blumcke I (2005) Suberoylanilide hydroxamic acid (SAHA) has potent anti-glioma properties in vitro, ex vivo and in vivo. J Neurochem 93: 992–999

Fenrick R, Hiebert SW (1998) Role of histone deacetylases in acute leukemia. J Cell Biochem Suppl 30–31: 194–202

Finnin MS, Donigian JR, Cohen A, Richon VM, Rifkind RA, Marks PA, Breslow R, Pavletich NP (1999) Structures of a histone deacetylase homologue bound to the TSA and SAHA inhibitors. Nature 401: 188–193

Glaser KB, Staver MJ, Waring JF, Stender J, Ulrich RG, Davidsen SK (2003) Gene expression profiling of multiple histone deacetylase (HDAC) inhibitors: defining a common gene set produced by HDAC inhibition in T24 and MDA carcinoma cell lines. Mol Cancer Ther 2: 151–163

Gregory PD, Wagner K, Horz W (2001) Histone acetylation and chromatin remodeling. Exp Cell Res 265: 195–202

Grignani F, De Matteis S, Nervi C, Tomassoni L, Gelmetti V, Cioce M, Fanelli M, Ruthardt M, Ferrara FF, Zamir I, Seiser C, Grignani F, Lazar MA, Minucci S, Pelicci PG (1998) Fusion proteins of the retinoic acid receptor-alpha recruit histone deacetylase in promyelocytic leukaemia. Nature 391: 815–818

Grunstein M (1997) Histone acetylation in chromatin structure and transcription. Nature 389: 349–352

Gui CY, Ngo L, Xu WS, Richon VM, Marks PA (2004) Histone deacetylase (HDAC) inhibitor activation of p21WAF1 involves changes in promoter-associated proteins, including HDAC1. Proc Natl Acad Sci USA 101: 1241–1246

Halkidou K, Gaughan L, Cook S, Leung HY, Neal DE, Robson CN (2004) Upregulation and nuclear recruitment of HDAC1 in hormone refractory prostate cancer. Prostate 59: 177–189

He LZ, Guidez F, Tribioli C, Peruzzi D, Ruthardt M, Zelent A, Pandolfi PP (1998) Distinct interactions of PML-RARalpha and PLZF-RARalpha with co-repressors determine differential responses to RA in APL. Nat Genet 18: 126–135

He LZ, Tolentino T, Grayson P, Zhong S, Warrell Jr RP, Rifkind RA, Marks PA, Richon VM, Pandolfi PP (2001) Histone deacetylase inhibitors induce remission in transgenic models of therapy-resistant acute promyelocytic leukemia. J Clin Invest 108: 1321–1330

Johnstone RW, Licht JD (2003) Histone deacetylase inhibitors in cancer therapy: is transcription the primary target? Cancer Cell 4: 13–18

Kelly WK, O'Connor OA, Krug LM, Chiao JH, Heaney M, Curley T, Macgregore-Cortelli B, Tong W, Secrist JP, Schwartz L, Richardson S, Chu E, Olgac S, Marks PA, Scher H, Richon VM (2005) Phase I study of an oral histone deacetylase inhibitor, suberoylanilide hydroxamic Acid, in patients with advanced cancer. J Clin Oncol 23: 3923–3931

Kelly WK, Richon VM, O'Connor O, Curley T, MacGregor-Curtelli B, Tong W, Klang M, Schwartz L, Richardson S, Rosa E, Drobnjak M, Cordon-Cordo C, Chiao JH, Rifkind R, Marks PA, Scher H (2003) Phase I clinical trial of histone deacetylase inhibitor: suberoylanilide hydroxamic acid administered intravenously. Clin Cancer Res 9: 3578–3588

Kim MS, Blake M, Baek JH, Kohlhagen G, Pommier Y, Carrier F (2003) Inhibition of histone deacetylase increases cytotoxicity to anticancer drugs targeting DNA. Cancer Res 63: 7291–7300

Lemercier C, Brocard MP, Puvion-Dutilleul F, Kao HY, Albagli O, Khochbin S (2002) Class II histone deacetylases are directly recruited by BCL6 transcriptional repressor. J Biol Chem 277: 22045–22052

Marchion DC, Bicaku E, Daud AI, Richon V, Sullivan DM, Munster PN (2004) Sequence-specific potentiation of topoisomerase II inhibitors by the histone deacetylase inhibitor suberoylanilide hydroxamic acid. J Cell Biochem 92: 223–237

Marks P, Rifkind RA, Richon VM, Breslow R, Miller T, Kelly WK (2001) Histone deacetylases and cancer: causes and therapies. Nat Rev Cancer 1: 194–202

Marks PA, Dokmanovic M (2005) Histone deacetylase inhibitors: discovery and development as anticancer agents. Expert Opin Investig Drugs 14: 1497–1511

Marks PA, Richon VM, Miller T, Kelly WK (2004) Histone deacetylase inhibitors. Adv Cancer Res 91: 137–168

Nimmanapalli R, Fuino L, Stobaugh C, Richon V, Bhalla K (2003) Cotreatment with the histone deacetylase inhibitor suberoylanilide hydroxamic acid (SAHA) enhances imatinib-induced apoptosis of Bcr-Abl-positive human acute leukemia cells. Blood 101: 3236–3239

Ocker M, Alajati A, Ganslmayer M, Zopf S, Luders M, Neureiter D, Hahn EG, Schuppan D, Herold C (2005) The histone-deacetylase inhibitor SAHA potentiates proapoptotic effects of 5-fluorouracil and irinotecan in hepatoma cells. J Cancer Res Clin Oncol 131: 385–394

Rahmani M, Reese E, Dai Y, Bauer C, Payne SG, Dent P, Spiegel S, Grant S (2005) Coadministration of histone deacetylase inhibitors and perifosine synergistically induces apoptosis in human leukemia cells through Akt and ERK1/2 inactivation and the generation of ceramide and reactive oxygen species. Cancer Res 65: 2422–2432

Rahmani M, Yu C, Dai Y, Reese E, Ahmed W, Dent P, Grant S (2003) Coadministration of the heat shock protein 90 antagonist 17-allylamino- 17-demethoxygeldanamycin with suberoylanilide hydroxamic acid or sodium butyrate synergistically induces apoptosis in human leukemia cells. Cancer Res 63: 8420–8427

Richon VM, Sandhoff TW, Rifkind RA, Marks PA (2000) Histone deacetylase inhibitor selectively induces p21WAF1 expression and gene-associated histone acetylation. Proc Natl Acad Sci USA 97: 10014–10019

Rundall BK, Denlinger CE, Jones DR (2004) Combined histone deacetylase and NF-kappaB inhibition sensitizes non-small cell lung cancer to cell death. Surgery 136: 416–425

Sakajiri S, Kumagai T, Kawamata N, Saitoh T, Said JW, Koeffler HP (2005) Histone deacetylase inhibitors profoundly decrease proliferation of human lymphoid cancer cell lines. Exp Hematol 33: 53–61

Sandor V, Robbins AR, Robey R, Myers T, Sausville E, Bates SE, Sackett DL (2000) FR901228 causes mitotic arrest but does not alter microtubule polymerization. Anticancer Drugs 11: 445–454

Secrist JP, Zhou X, Richon VM (2003) HDAC inhibitors for the treatment of cancer. Curr Opin Investig Drugs 4: 1422–1427

Song J, Noh JH, Lee JH, Eun JW, Ahn YM, Kim SY, Lee SH, Park WS, Yoo NJ, Lee JY, Nam SW (2005) Increased expression of histone deacetylase 2 is found in human gastric cancer. APMIS 113: 264–268

Ungerstedt JS, Sowa Y, Xu WS, Shao Y, Dokmanovic M, Perez G, Ngo L, Holmgren A, Jiang X, Marks PA (2005) Role of thioredoxin in the response of normal and transformed cells to histone deacetylase inhibitors. Proc Natl Acad Sci USA 102: 673–678

Verdin E, Dequiedt F, Kasler HG (2003) Class II histone deacetylases: versatile regulators. Trends Genet 19: 286–293

Warrener R, Beamish H, Burgess A, Waterhouse NJ, Giles N, Fairlie D, Gabrielli B (2003) Tumor cell-selective cytotoxicity by targeting cell cycle checkpoints. FASEB J 17: 1550–1552

Zhu P, Martin E, Mengwasser J, Schlag P, Janssen KP, Gottlicher M (2004) Induction of HDAC2 expression upon loss of APC in colorectal tumorigenesis. Cancer Cell 5: 455–463


We thank U. Nguyen and Y. David for help with tissue culture G. Laevsky for advice on microscopy P. Lewis for providing biotinylated H3 peptides C.D. Allis, G. Debelouchina, Z. Brown, B. Wang and C. Jenness for helpful discussions and K. Jani for careful proofreading of this manuscript. NIH-3T3 cells were a gift from J. Schwarzbauer (Princeton University). Funding provided by the Swiss National Science Foundation (postdoctoral fellowships to M.M.M. and B.F.) and the US National Institutes of Health (grant R01-GM107047 to T.W.M.).