Cell Tissue Res DOI 10.1007/s00441-014-1862-4

REVIEW

Histone variants: key players of chromatin Burcu Biterge & Robert Schneider

Received: 16 January 2014 / Accepted: 27 February 2014 # Springer-Verlag Berlin Heidelberg 2014

Abstract Histones are fundamental structural components of chromatin. Eukaryotic DNA is wound around an octamer of the core histones H2A, H2B, H3, and H4. Binding of linker histone H1 promotes higher order chromatin organization. In addition to their structural role, histones impact chromatin function and dynamics by, e.g., post-translational histone modifications or the presence of specific histone variants. Histone variants exhibit differential expression timings (DNA replication-independent) and mRNA characteristics compared to canonical histones. Replacement of canonical histones with histone variants can affect nucleosome stability and help to create functionally distinct chromatin domains. In line with this, several histone variants have been implicated in the regulation of cellular processes such as DNA repair and transcriptional activity. In this review, we focus on recent progress in the study of core histone variants H2A.X, H2A.Z, macroH2A, H3.3, and CENP-A, as well as linker histone H1 variants, their functions and their links to development and disease.

Keywords Chromatin . Histone variant . Function . Epigenetic regulation . Transcription B. Biterge : R. Schneider (*) Institut de Génétique et de Biologie Moléculaire et Cellulaire, CNRS UMR 7104, INSERM U 964, Université de Strasbourg, 67404 Illkirch, France e-mail: [email protected] B. Biterge International Max-Planck Research School, Max Planck Institute of Immunobiology and Epigenetics, 79108 Freiburg im Breisgau, Germany B. Biterge University of Freiburg, Fahnenbergplatz, 79085 Freiburg im Breisgau, Germany

Introduction Eukaryotic DNA is packaged into a complex macromolecular structure called chromatin to facilitate compaction of the genetic material. The DNA is folded and condensed into chromatin in a dynamic manner since it still needs to be accessible to carry out key functions such as replication, transcription, and DNA repair. The first step of compaction is achieved by packaging the naked DNA into the nucleosome, which is the repeating unit of chromatin. One hundred and forty-seven base pairs of DNA are wrapped around the histone octamer containing two copies of each core histone H2A, H2B, H3, and H4 (Luger et al. 1997). The linker histone H1 binds to the nucleosome at the DNA entry–exit point and protects the DNA linking adjacent nucleosomes. Additionally, H1 participates in the formation of higher-order chromatin by further compacting the “beads on a string” chromatin into 30 nm fibers (Robinson and Rhodes 2006). Histones are small basic proteins that can be covalently modified at their flexible N- or C-terminal tails as well as globular domains (Kouzarides 2007). Histone posttranslational modifications (PTMs) affect interactions of histones with DNA and effector proteins, leading to alterations in chromatin structure and function (Choi and Howe 2009; Strahl and Allis 2000). For instance, certain histone PTMs are associated with a more open and accessible chromatin conformation, while others may be found in highly compacted and transcriptionally inactive loci, hinting towards the “histone code” (Turner 1993; Strahl and Allis 2000). Histones are encoded by multiple allelic and non-allelic copies of genes, giving rise to different subtypes with different sequences (Marzluff et al. 2002) (Figs. 1 and 2). Canonical histones are almost exclusively expressed during the S-phase of the cell cycle and incorporated into chromatin in a DNA replicationdependent fashion, whereas replication-independent histone “variants” (replacement histones) are expressed throughout the

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a

b Canonical H2A

H3.1

H3.2

99%

H2A.X

95%

H3.3

96%

H2A.Z

64%

CENP-A

50%

macroH2A

65%

macro domain

Fig. 1 Schematic representations of selected histone H3 (a) and H2A (b) variants. Globular domains and flexible N- or C-terminal tails are displayed as ovals and lines respectively. Patterning in white indicates

difference in amino acid composition of the variant compared to the canonical histone. Percentage of similarity is shown on the left

cell cycle (Albig and Doenecke 1997). Genes encoding canonical histones lack introns and their mRNAs are not polyadenylated but instead have a unique 3’ stem-loop structure, which is key for mRNA stability and translation (Pandey et al. 1990; Dominski and Marzluff 1999). On the other hand, histone variant pre-mRNAs contain introns, which can even undergo alternative splicing as in the case of macroH2A, and are polyadenylated (Rasmussen et al. 1999; Marzluff et al. 2002). Histone variants are deposited onto chromatin by specific histone chaperones and also interact with other chromatin

modifiers (Tagami et al. 2004; Heo et al. 2008; Foltz et al. 2009; Luk et al. 2010, Elsaesser and Allis, 2010) (Table 1). Structural differences introduced by a core histone variant typically affect interactions between histone proteins within a nucleosome, hence their stability as well as the open/compact chromatin conformation. For example, histone variants H2A.Z and H3.3 are mainly linked with an open chromatin conformation and transcriptional activity, while macroH2A deposition stabilizes the nucleosome and is often associated with a repressive chromatin state (Chakravarthy and Luger 2006;

Fig. 2 Alignment of human histone H1 variants amino acid sequences with histone H1.2 (using Jalview). Conserved residues are highlighted by shades of purple, darker color representing higher conservation (Waterhouse et al. 2009)

Cell Tissue Res Table 1 Summary of human core histone variants, selected associated factors, and functional implications (reviewed in Skene and Henikoff 2013, Chen et al. 2013) Histone variant

Associated factors

Potential functions

H2A.X H2A.Z macroH2A H3.3

FACT INO80, SWR1 ATRX HIRA, DAXX/ATRX

DNA double-strand break repair, damage signaling Nucleosome positioning, context dependent gene (de)activation Evidence for poising genes for transcription and stabilizing the differentiated state of the cell Enriched on actively transcribed genes, regulatory regions and in some cases in heterochromatin

CENP-A

HJURP

Centromere identity and stability, recruitment of kinetochore complex

Thakar et al. 2009). In addition, histone variants play important roles in cellular processes, e.g., the H3 variant CENP-A in assembly of the kinetochore complex at the centromere and phosphorylation of H2A.X in DNA repair (Rogakou et al. 1998; Yoda et al. 2000). Consequently, replacement of canonical histones with histone variants adds another level of complexity and a distinct way of modulating chromatin function.

H2A.X DNA double-strand break repair is crucial for genome integrity. Upon damage, the H2A variant H2A.X is phosphorylated at serine 139 by DNA-dependent protein kinases (ATR/ATM), forming so-called γH2A.X foci, which can extend for up to several megabases in mammals (Rogakou et al. 1998 and 1999). This H2A.XS139 phosphorylation facilitates the recruitment of the DNA damage repair machinery, as well as chromatin remodelers such as INO80 and SWR1 (van Attikum et al. 2007). γH2A.X signaling is often associated with a relaxed chromatin conformation, which makes the break site more accessible for, e.g., DNA repair proteins (Fink et al. 2007). Likewise, H2A.X incorporation into chromatin had been reported to impair histone H1 binding, although it seems not to have a significant impact on nucleosome structure, and the phosphorylation of H2A.X appears to enhance the H1 binding impairment as well as destabilize the nucleosome (Li et al. 2010). Moreover, it has been reported that the phosphorylation of H2A.X enhances FACT (facilitates chromatin transcription)-mediated H2A.X eviction from the nucleosome (Heo et al. 2008). After the damage is repaired, γH2A.X must be dephosphorylated or removed from chromatin for the efficient recovery from DNA damage-induced cell cycle arrest. There are contradictory data on H2A.X eviction, as some studies suggest that H2A.X at the double-strand break (DSB) site is replaced by another H2A variant, H2A.Z, whereas other reports suggest that both H2A.X and H2A.Z are removed from the vicinity of the DSB site after repair (PapamichosChronakis et al. 2006; van Attikum et al. 2007). There is also growing evidence for γH2A.X phosphatases such as

mammalian PP2A and HTP-C phosphatase complex in yeast (Chowdhury et al. 2005; Keogh et al. 2006). More recently, the H2A.X tyrosine 142 residue has been identified to be phosphorylated by WSTF, a WICH ATPdependent chromatin remodeling complex subunit (Xiao et al. 2009). Unlike serine 139, tyrosine 142 is ubiquitously phosphorylated and its de-phosphorylation signals DNA damage repair; failure to do so leads to apoptosis (Xiao et al. 2009; Cook et al. 2009). How effector proteins read and interpret these two phosphorylation states of H2A.X remains an interesting question. A recent study shows that MCPH1, an early DNA damage response protein, directly interacts with di-phosphorylated H2A.X (Singh et al. 2012). Furthermore, many anti-cancer drugs rely on the deficiency of tumor cells to repair DNA damage, and they induce formation of γ-H2A.X foci, suggesting γ-H2A.X is a “biodosimeter” that can be used in screening new therapeutic agents for their DNA-damaging capabilities (Ivashkevich et al. 2012).

H2A.Z H2A.Z is one of the most conserved histone variants. It shares higher sequence similarity among different species than to the canonical H2A within the same organism, suggesting a functionally distinct role for H2A.Z (Henikoff et al. 2001). This notion has been supported by many studies indicating that the deletion of the H2A.Z gene is lethal, e.g., in Drosophila, Tetrahymena, and mouse (van Daal and Elgin 1992; Liu et al. 1996; Faast et al. 2001). H2A.Z-containing nucleosomes display an extended acidic patch on their surface, causing subtle destabilization of the interaction between the H2A.Z– H2B dimer and with the H3–H4 tetramer, and also altered linker histone H1 binding (Suto et al. 2000; Thakar et al. 2009). In line with this, H2A.Z is enriched at the nucleosome-depleted region (NDR) of active transcriptional start sites (TSS) in mouse, which is believed to be necessary for the binding of the transcriptional machinery (Nekrasov et al. 2012). Together with DNA-binding proteins such as Foxa2, H2A.Z regulates nucleosome depletion and promotes gene activation in differentiating cells (Li et al. 2012). Additionally, H2A.Z is depleted from DNA-methylated

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genomic regions (Zilberman et al. 2008). The open chromatin conformation caused by the incorporation of H2A.Z can also aid in the recruitment of the repair machinery at the DNA double-strand breaks (Xu et al. 2012). A recent study showed that incorporation of H2A.Z into chromatin increases the stability of mononucleosomes and facilitates higher-order chromatin formation (Chen et al. 2013). This finding is in accordance with the previous studies implicating H2A.Z in gene silencing, as it is localized to the heterochromatin–euchromatin boundary to prevent spreading of heterochromatin, as well as to keep the pericentromeric heterochromatin intact, e.g., for correct centromere formation (Meneghini et al. 2003; Greaves et al. 2007). In ES cells, H2A.Z is mainly found at the promoters of key developmental regulators that are repressed by the polycomb complex, and shows a pattern of distribution similar to Suz12 (Creyghton et al. 2008). Hence, the function of H2A.Z in transcription seems to be context-dependent and could be affected by other chromatin marks present. Unlike the canonical H2A, H2A.Z is constitutively expressed and incorporated into chromatin by the ATPdependent chromatin remodeling complexes SWR1 and INO80 at specific loci by mechanisms uncoupled to DNA replication (Luk et al. 2010; Papamichos-Chronakis et al. 2011). H2A.Z can also be specifically modified, e.g., monomethylated by SETD6 at lysine 7. Levels of H2A.ZK7me1 increase remarkably upon cellular differentiation of mouse embryonic stem cells, suggesting that H2A.ZK7me1 is a marker of cellular differentiation (Binda et al. 2013).

macroH2A The largest histone variant macroH2A (∼40 kDa, almost three times the size of canonical H2A) is named after its macrodomain-containing C-terminal globular domain. Macrodomain-containing proteins are found in all kingdoms of life and show affinity for NAD+ metabolites such as ADP-ribose and poly(ADP-ribose) (PAR) (Till and Ladurner 2009). This is very interesting, since it could suggest that macroH2A has a role in sensing the metabolic state of the cell and linking it with chromatin. Mammals have two genes encoding for macroH2A and, unlike most other histone genes, alternative splicing takes place, resulting in 3 isoforms: macroH2A1.1, macroH2A1.2, and macroH2A2, which will collectively be referred to as macroH2A in this review for simplicity (Chadwick and Willard 2001; Rasmussen et al. 1999). MacroH2A-containing nucleosomes show high structural similarity to canonical nucleosomes and wrap the same amount of DNA (Chakravarthy et al. 2005; Changolkar and Pehrson 2002). However, a four amino acid difference within the L1 loop, which participates in interactions between H2A–

H2B dimers, gives rise to more stable nucleosomes that have increased resistance to nuclease digestion and more compact chromatin (Chakravarthy and Luger 2006; Changolkar and Pehrson 2002; Muthurajan et al. 2011). The role initially attributed to macroH2A in mammals was transcriptional repression, taking into account its enrichment on the transcriptionally inactive X chromosome (Xi), its association with the inactive allele of imprinted genes, and the impeding effect it exerts on transcriptional elongation (Costanzi and Pehrson 1998; Choo et al. 2006; Doyen et al. 2006). Nevertheless, subsequent studies have indicated macroH2A enrichment on inducible genes, as well as bivalent genes (harboring both H3K27me3 and H3K4me marks) in stem cells (Azuara et al. 2006; Buschbeck et al. 2009; Creppe et al. 2012). It remains bound to these genes before and after transcription, implying a role for macroH2A in establishing the necessary chromatin environment for “poised” genes to be transcribed (Gamble et al. 2010). Additionally, macroH2A is expressed at low levels in stem cells and is upregulated during differentiation, preventing reprogramming of induced pluripotent stem cells and after nuclear transfer (Pasque et al. 2011; Gaspar-Maia et al. 2013; Barrero et al. 2013). The molecular mechanism underlying macroH2Amediated gene repression remains elusive to date. One possible explanation is that macroH2A may directly prevent transcription factor binding and chromatin remodeling (Angelov et al. 2003). On the other hand, macroH2A could act indirectly, as it inhibits p300-dependent histone acetylation in vitro (Doyen et al. 2006). Moreover, ATRX (α-Thalassemia/ Mental Retardation syndrome X-linked) negatively regulates macroH2A incorporation into chromatin by an unknown mechanism (Ratnakumar et al. 2012). MacroH2A is overexpressed in Huntington’s disease patients, as well as in steatosis-associated hepatocellular carcinoma, although it is downregulated in breast and lung tumors (Hu et al. 2011; Rappa et al. 2013; Sporn et al. 2009).

H3.3 Histone H3 exists in three distinct non-centromeric subtypes in metazoans. H3.1 and H3.2 are canonical H3 proteins, which are highly expressed in S-phase and incorporated into chromatin during DNA replication. H3.2 differs from H3.1 by only one amino acid (Hake and Allis 2006). In contrast to this, the variant H3.3, which differs from H3.1 by 5 amino acids, can be incorporated into chromatin both coupled to and independent of DNA replication, and its expression is maintained throughout the cell cycle (Ahmad and Henikoff 2002). Although the changes in amino acid sequence may seem subtle, H3.1 and H3.3 exhibit differential characteristics and H3.3 can replace H3.1 at transcriptionally active genes, as initially thought to be triggered by transcriptional activity and histone turnover (Mito

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et al. 2005; Schwartz and Ahmad 2005). In line with this, H3.3 is enriched in marks linked to active transcription such as H3K4me3, whereas H3.1 is typically associated with more repressive histone marks (Hake et al. 2006). Recent findings point out that H3.3 incorporation results in an open chromatin conformation and promotes transcription by impairing higherorder chromatin formation despite little effect on stability of mononucleosomes (Chen et al. 2013). Moreover, H3.3 colocalizes with another histone variant, H2A.Z, at the promoters of active genes and marks the nucleosome-depleted region (Jin et al. 2009). H3.3 incorporation has also been linked with epigenetic memory of active gene state in the absence of transcription (Ng and Gurdon 2008). The functional diversity of H3.1 and H3.3 may stem from their specific incorporation by histone chaperones. H3.1 is deposited by CAF1 (Chromatin Assembly Factor-1) while HIRA deposits H3.3 at transcriptionally active sites (Tagami et al. 2004; Elsässer and Allis 2010). Surprisingly, Daxx (death domain-associated protein) has been identified as another H3.3-specific histone chaperone that, together with chromatin remodeler ATRX (α-thalassemia/mental retardation syndrome protein), deposits H3.3 at telomeres and pericentric heterochromatin (Drané et al. 2010; Goldberg et al. 2010; Lewis et al. 2010). A recent study showed H3.3 localization at the PML-NBs (promyelocytic leukemia nuclear bodies), which have previously been implicated in oncogeneinduced senescence, and H3.3 was lost from pericentromeric heterochromatin upon Daxx or PML depletion (Ferbeyre et al. 2000; Corpet et al. 2013). Amino acid differences between H3.1/3.2 and H3.3 may also play a direct role in chromatin dynamics and the posttranslational modification patterns of H3, e.g., alanine at position 31 in H3.1 is a serine in H3.3 that can be phosphorylated during mitosis (Hake et al. 2005). In addition, glycine 90 of H3.3 is essential for Daxx binding (Elsässer et al. 2012). H3.3 (and its chaperones) have also been implicated in cancers. Mutations of H3.3 at glycine 34 to valine or arginine and lysine 27 (which can be methylated or acetylated) to methionine have been found in 31 % of pediatric glioma tumors (Schwartzentruber et al. 2012). The substitution of H3.3 K27 inhibits Polycomb repressive complex 2 (PRC2) activity, affecting overall H3K27me3 levels as well as downstream gene expression patterns, and is associated with reduced survival in pediatric glioma patients (Lewis et al. 2013; Chan et al. 2013).

CENP-A Centromeres are special chromatin domains where the kinetochore complex assembles to create a platform for spindle microtubule attachment during cell division to facilitate sister chromatid segregation (Amor et al. 2004). Although human centromeric DNA is commonly composed of arrays of α-

satellite repeats, the DNA sequence alone is not sufficient for the formation of functional centromeres (Yang et al. 2000; Voullaire et al. 1993). This suggests an epigenetic mechanism for establishing centromere identity. A particular H3 variant called cenH3, CENP-A (centromere protein A) in mammals, replaces canonical H3 at most centromeric regions and is essential for propagation and maintenance of the centromere (Yoda et al. 2000). Human CENP-A shares 50 % sequence identity with histone H3.1. Again, a specific chaperone, in this case HJURP (Holliday junction recognition protein), deposits CENP-A at the centromere in early G1 phase in a replication-independent manner (Jansen et al. 2007; Foltz et al. 2009; Dunleavy et al. 2009). The composition of CENP-A-containing nucleosomes has been under discussion for the last decade. Initial atomic force microscopy analysis indicated that CENP-A-containing nucleosomes had reduced height compared to canonical nucleosomes, which suggested that they could be composed of a histone tetramer, rather than an octamer (Dalal et al. 2007; Dimitriadis et al. 2010). However, multiple groups have recently challenged this concept of CENP-A containing halfnucleosomes by showing that CENP-A nucleosomes are octameric throughout the cell cycle (Miell et al. 2013; Hasson et al. 2013; Padeganeh et al. 2013), and they do not confer reduction in nucleosome height (Codomo et al. 2014; Walkiewicz et al. 2014). Many CENP-A post-translational modification sites have been identified, including trimethylation of Gly1 and phosphorylation of Ser16 and Ser18, which are suggested to influence centromeric chromatin conformation (Bailey et al. 2013). Furthermore, phosphorylation of the CENP-A N-terminal tail is required for proper mitotic progression (Goutte-Gattat et al. 2013). CENP-A expression is elevated in epithelial ovarian cancer cells compared to the noncancerous tissues and is correlated with poor patient survival, suggesting CENP-A could be a prognostic marker and a potential target for epigenetic therapy (Qiu et al. 2013).

Linker histone variants Histone H1 is important for higher-order chromatin formation and compaction (Robinson and Rhodes 2006). It binds the nucleosomal core particle close to the DNA entry–exit point and protects the linker DNA between adjacent nucleosomes. H1 stabilizes the interaction of DNA with the nucleosome and prevents ATP-dependent remodeling of the chromatin (Hill 2001). The classical model is that compaction introduced by H1 could lead to inaccessibility to transcription factors and to RNA polymerase; therefore, H1 was originally associated with gene repression. However, a more dynamic role for H1 has also been proposed. FRAP (fluorescence recovery after photobleaching) experiments showed that H1 continuously

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associates with and dissociates from chromatin (Misteli et al. 2000). Similar to the core histones, H1 is a tripartite protein: it consists of a short N-terminal tail, a globular domain containing the histone fold motif and a C-terminal tail that makes up almost half of the protein. This C-terminal tail largely affects the affinity to chromatin (Hendzel et al. 2004), and H1 variants with shorter C-terminal tails have shorter residence times on chromatin (Th’ng et al. 2005). Furthermore, distinct H1 post-translational modifications are involved in altering the accessibility of chromatin to the transcriptional machinery in a cell cycle- or gene-specific manner and/or H1 mobility (Harshman et al. 2013). For instance, H1.4 K34 acetylation is enriched at TSS of active genes, and increases H1 mobility and recruits general transcription factors, suggesting H1 is a fine tuner of transcription rather than merely a general repressor, as well as a mechanism of regulation of H1 function via post-translational modifications (Kamieniarz et al. 2012). Whether and how histone chaperones control the dynamics of H1 is still an open question. Linker histone H1 is expressed in various forms with divergent sequences. In human, there are 11 H1 subtypes: H1.0–1.5 and H1.X are expressed in many different somatic cell types, whereas H1t, H1T2, HILS1, and H1oo are germline-specific (Izzo et al. 2008). H1 genes originated from gene duplication events and acquired sequence variety and differential expression patterns (spatial and temporal) during the course of evolution and are therefore likely to have functional divergence (Th’ng et al. 2005). Nonetheless, this is hard to prove since single knockouts of H1 variants in mouse do not have strong phenotypes, but result in the upregulation of the expression of other subtypes, suggesting a compensatory mechanism amongst different H1 subtypes (Fan et al. 2001). Interestingly, a more recent study indicated that depleting single H1 variants by RNAi (rather than constitutive knockout that results in compensation) could lead to alterations in expression levels of different gene subsets as well as distinct phenotypic effects (Sancho et al. 2008). Furthermore, knockout of three H1 subtypes, which results in a 50 % reduction in total protein levels of H1, is embryonic-lethal (Fan et al. 2005). In addition, it is extremely difficult to produce antibodies specific to only one type of histone H1, which is currently a main limitation in the field. A recent study employed a DamID approach to circumvent this lack of H1-specific antibodies, and performed a first mapping analysis for the somatic H1 variants H1.1–H1.5 in human cells. This study demonstrated distinct localization patterns for individual somatic H1 subtypes and unique subtype combinations in functionally different chromatin domains (Izzo et al. 2013). The expression of H1.1–1.5 and H1t is tightly restricted to S-phase and is DNA-replication-dependent; therefore, they should be considered as subtypes rather than variants (Malik and Henikoff 2003). Histones H1.2–1.5 are expressed in almost all tissues, while H1.1 is expressed in a subset of

tissues and H1t expression is specific to testis (Doenecke et al. 1997). In contrast to this, H1.0, H1.X, H1T2, HILS1, and H1oo are expressed throughout the cell cycle and incorporated into chromatin in a replication-independent manner, and are thus categorized as H1 variants. Replicationdependent and -independent H1 genes are found at different genomic regions. Histone H1.1–1.5 and H1t-encoding genes are located in the major histone cluster together with other core histone genes on chromosome 6, whereas the others are scattered around the genome (Terme et al. 2011). What is currently known about the “replacement” H1 variants is limited. Histones HILS1 (histone H1-like protein in spermatids 1) and H1T2 are testis-specific H1 variants. HILS1 is expressed in the late spermatid stage of spermatogenesis and helps condense the chromatin (Yan et al. 2003). H1T2 specifically localizes to a chromatin domain at the apical pole in round and elongating spermatids, which is crucial for spermatid elongation and exchange of histones with protamines, thus chromatin condensation (Martianov et al. 2005). H1T2 knockout males are sterile due to reduced sperm motility and competency for fertilization (Martianov et al. 2005; Tanaka et al. 2005). H1oo is the oocyte-specific variant of H1 and is important for meiotic maturation of the oocyte (Furuya et al. 2007). It is incorporated into and condenses the sperm chromatin after fertilization (Mizusawa et al. 2010). Recent studies have pointed out that H1oo prevents differentiation by maintaining the expression of pluripotency genes (Hayakawa et al. 2012). H1.0 represents up to 80 % of the H1 transcripts in highly differentiated cells. It is suggested to have a role in the regulation of pluripotency genes during differentiation (Terme et al. 2011). Histone H1.X is the least-studied H1 variant. It is mainly enriched in less accessible chromatin domains and adopts a cell cycle-dependent nuclear distribution as it accumulates in the nucleolus during G1 phase, while during S and G2 phases, it is distributed evenly in the nucleus (Happel et al. 2005; Stoldt et al. 2007). A recent study identified the first Drosophila H1 histone variant, an embryonic stage-specific H1, called dBigH1 that regulates zygotic genome activation and is replaced by somatic dH1 after cellularization (Pérez-Montero et al. 2013). Histone H1 has also been implicated in diseases such as cancer. H1 is lost from chromatin in senescent cells, which is an important mechanism of the cell to prevent immortalization and tumorigenesis (Funayama et al. 2006). Likewise, depletion of H1 in breast cancer cell lines can induce senescence (Sancho et al. 2008). Moreover, H1 expression in prostate cancer positively correlates with increased cancer cell proliferation, suggesting that H1 expression may contribute to overcome growth arrest (Sato et al. 2012). Genes encoding for linker histone H1 variants have also been found to be frequently mutated in follicular lymphoma patients, which could account for the epigenetic basis of follicular lymphoma (Li et al. 2014).

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Concluding remarks The growing evidence for the importance of histone variants in regulation of chromatin structure, dynamics, and function has attracted more attention to the field in recent years. However, the mechanisms of histone variant action, incorporation into and removal from chromatin, are not yet completely understood. Therefore, future studies will shed light on the roles of histone variants beyond being structural components of chromatin and contribute to our current understanding of epigenetic regulation. Acknowledgements We thank Poonam Bheda for critical reading of the manuscript and Adam Fiseha Kebede for helpful discussions. Work in the RS laboratory is supported by the Fondation pour la Recherche Médicale, the Agence Nationale de Recherche (ANR), INSERM, La Ligue National Contre La Cancer (Equipe Labellise) and an ERC starting grant.

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