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Briefings in Functional Genomics Advance Access originally published online on May 10, 2006
Briefings in Functional Genomics 2006 5(3):228-243; doi:10.1093/bfgp/ell020
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© The Author 2006. Published by Oxford University Press. All rights reserved. For permissions, please email: journals.permissions@oxfordjournals.org

Special Issue Papers

Histone variants—the structure behind the function

Juan Ausió

Juan Ausió, Department of Biochemistry and Microbiology, University of Victoria, Victoria, BC, Canada V8W 3P6. E-mail: jausio{at}uvic.ca


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 DYNAMIC CHROMATIN
 HISTONE H2A VARIANTS SPECIALIZE...
 HISTONE H3 VARIANTS
 OTHER CORE HISTONE VARIANTS
 CORE HISTONE VARIANTS AND...
 HISTONE H1--MICROHETEROGENEITY...
 CONCLUSIONS
 Acknowledgements
 References
 
In recent years, the chromatin field has witnessed a renewed interest in histone variants as pertaining to their structural role, but mainly because of the functional specificity they impart to chromatin. In this review, I am going to discuss several of the most recent structural studies on core histone (H2A.Bbd, H2A.Z, H2A.X, macroH2A, H3.3, CENP-A) and linker histone variants (histone H1 microheterogeneity) focusing on their role in nucleosome stability and chromatin fibre dynamics with special emphasis on their possible functional implications. The data accumulated to date indicates that histone variability plays an important role in the histone-mediated regulation of chromatin metabolism. Understanding and deciphering the underlying structural amino acid code behind such variability remains one of the most exciting future challenges in chromatin research.

Keywords: histones, chromatin, nucleosome, stability, dynamics


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
DYNAMIC CHROMATIN
 HISTONE H2A VARIANTS SPECIALIZE...
 HISTONE H3 VARIANTS
 OTHER CORE HISTONE VARIANTS
 CORE HISTONE VARIANTS AND...
 HISTONE H1--MICROHETEROGENEITY...
 CONCLUSIONS
 Acknowledgements
 References
 
In 1947, at the peak over the controversy of the histone/DNA nature of the genetic material Edgar and Ellen Stedman [1] wrote:

‘... the old view of the composition of the cell nucleus, that the basic protein of cell nuclei can provide the material of which genes are composed is one which, to say the least, has never seem satisfying. While histone is usually regarded as, and probably is, a fairly simple protein, its degree of complexity is not, in fact, known and it may therefore been argued that it is sufficiently complex to fulfill this function’.

This inconclusive, yet hesitant statement, appears in retrospective to be quite timely as there is now plenty of evidence in support of the important structural and epigenetic role of the ‘fairly simple’ histones in mediating gene expression. Furthermore, histones also play important additional roles in other metabolic chromatin functions, such as maintenance of the chromosome integrity (DNA repair), DNA recombination (meiosis) and the process of DNA replication itself. In fact, it has been the proposal of a histone language and deciphering of a potential histone code [2, 3] resulting from histone post-translational modifications (PTMs) [4, 5] and histone variants [6] that have been in part responsible for bringing much recent attention and renewed interest to the chromatin field.

At the time when the conventional nomenclature of histones was finally established at the CIBA Foundation Symposium on Structure and Function of Chromatin in 1974 [7], there was already evidence to suggest that in addition to each of the five major histone types (H1, H2A, H2B, H3 and H4) [8] each of them included some minor variant forms that could in turn be subject to PTMs (i.e. acetylation, methylation, phosphorylation, ubiquitination and poly-ADP-ribosylation) (see [9–11] for early reviews). From a structural point of view, histone variants (which are often replication-independent) [12] can be classified into homomorphous and heteromorphous families [13, 14] depending on the extent of their amino acid sequence departure from the main canonical isoforms (whose genes are usually expressed during the S phase of the cell cycle). Homomorphous variants involve only a few amino acid changes (i.e. H2A.1 and H2A.2; H3.1, H3.2 and H3.3) whilst heteromorphous variants involve changes that affect larger portions of the histone molecule [i.e. H2A.X, H2A.Z, macroH2A (mH2A), H2A Barr body-deficient (H2A.Bbd) and centromeric protein A (CENP-A)].

Histones and their variants are responsible for organizing the chromatin complex [15] and within this context they can be classified into core histones (H2A/H2B, H3/H4) and linker histones (histones of the H1 family). Core histones consist of a dimerizing central histone-fold domain [16] which is flanked by N- and C-terminal unstructured domains commonly referred to as ‘tails’, and as their name indicate they are responsible for creating a protein ‘core’ around which 146 bp of DNA are wrapped in approximately two left-handed superhelical turns giving rise to a particle that is known with the name of nucleosome core particle (NCP). Linker histones consist of a winged helix motif [17] flanked by N- and C-terminal tails, bind to the linker DNA connecting adjacent nucleosomes and are responsible for the higher order of folding into a 30 nm chromatin fibre.

In this review, I am going to focus on several recently unveiled dynamic structural aspects of histone variants, both at the nucleosome and at the chromatin fibre levels, with emphasis on their role and possible implications for their defined chromatin function.


   DYNAMIC CHROMATIN
TOP
 ABSTRACT
 INTRODUCTION
 DYNAMIC CHROMATIN
HISTONE H2A VARIANTS SPECIALIZE...
 HISTONE H3 VARIANTS
 OTHER CORE HISTONE VARIANTS
 CORE HISTONE VARIANTS AND...
 HISTONE H1--MICROHETEROGENEITY...
 CONCLUSIONS
 Acknowledgements
 References
 
The two major functions of the chromatin nucleoprotein complex are: folding of DNA to fit the genome within the limited space available in the nucleus of the eukaryotic cell and modulation of the chromatin metabolism [15]. Implicit to these functions is the notion that this complex cannot be structurally rigid but rather a highly dynamic system [4]. Thus, although a lot of the highly valuable structural information about the nuclesome has been obtained from the crystallographic analysis of NCP in recent years [18, 19], this information is quite limited when trying to understand the dynamics of chromatin. Several highly complementary structural methods of analysis, have been, or are currently being used to help overcome the intrinsic limitations of the crystallographic data. In what follows next, I am going to briefly refer to some of the technology used for this purpose as it pertains to chromatin.

Analytical ultracentrifuge
Sedimentation velocity analysis in the analytical ultracentrifuge is extremely sensitive to changes in the tertiary and quaternary structure of macromolecular complexes and, hence is very suitable for the study of chromatin [20]. In addition to monitoring conformational changes of NCPs and nucleosome array (chromatin fibre) folding, the method is also useful for determining chromatin stability (DNA/histone dissociation fluctuations) [20].

Fluorescence recovery after photobleaching (FRAP)
This method utilizes cells that have been either transiently or stably transfected with green fluorescence protein (GFP)-tagged histones. They are then analysed using confocal microscopy, after bleaching certain areas of the nucleus, and the fluorescence recovery (molecular diffusion) from the unbleached to the bleached areas is monitored [21, 22]. The method allows for the determination of the rates at which histones associate and dissociate from the chromatin template in situ [23] within the living cell and, hence provides a useful tool for the study of chromatin dynamics in vivo.

Fluorescence resonance energy transfer (FRET)
As with the analytical ultracentrifuge, this represents a rather classical method of analysis which still retains an enormous potential for structural analysis [24]. With this technique, two regions of the chromatin complex are labelled with different fluorescent labels. When one of these two fluorophores (donor) is excited by an incident light beam (at a suitable wavelength), and a second fluorophore (acceptor) is at a close distance (20–100 Å), an excited state energy from the former can be transferred to the acceptor. Monitoring this process allows one to determine the relative position (movement) of the two fluorophores. The technique has been recently applied to the study of NCP dynamics [25–27].

Restriction enzyme accessibility assays
This method monitors the accessibility of restriction enzyme cutting sites in chromatin complexes reconstituted with sequence-defined DNA templates. It has been applied to the characterization of DNA unwrapping from NCP [28, 29] and nucleosome arrays [30].

One potential problem with this technique is the lack of information on the role that the high concentration of divalent ions required for the restriction enzyme digestions has on the intrinsic dynamic properties of the chromatin complexes.

Single molecule analysis
In this type of analysis, individual molecules or complexes are analysed using a variety of techniques: atomic force microscope (AFM), optical tweezers (OT) and magnetic tweezers (MT) [31]. This is perhaps one of the most powerful, recently developed set of techniques for the study of chromatin stability and folding [32]. Recently, it has been used in combination with FRET to study the fluctuations of the DNA wrapping in the nucleosome [33]. However, its use in the analysis of chromatin complexes consisting of histone variants has not been forthcoming yet and it may turn out to be very informative.


   HISTONE H2A VARIANTS SPECIALIZE THE NUCLEOSOME FOR DEDICATED FUNCTIONS
TOP
 ABSTRACT
 INTRODUCTION
 DYNAMIC CHROMATIN
 HISTONE H2A VARIANTS SPECIALIZE...
HISTONE H3 VARIANTS
 OTHER CORE HISTONE VARIANTS
 CORE HISTONE VARIANTS AND...
 HISTONE H1--MICROHETEROGENEITY...
 CONCLUSIONS
 Acknowledgements
 References
 
The heteromorphic members of this histone H2A type (Figure 1A) exhibit significantly major sequence changes at both the N- and C-terminal ends of their molecules with substantially larger predominance of those affecting the carboxy end [4, 14]. While the implications of the N-terminal heterogeneity still remain unclear, most of the recent work has centred its attention on the variability that affects the C-terminal domain. Indeed, early studies by Eickbush and coworkers [34] took advantage of an endogenous protease that selectively cleaves H2A.1 between V(114) and L (115) releasing the last 15 amino acids, to show that this region is critical for the stability of the histone core octamer [35]. An observation which might have consequences for nucleosome stability. Also, this is the region that exits the NCP close to its dyad axis [18, 36] at a position near the binding site of the winged helix domain of the linker histones in the nucleosome [37] and its disruption of this binding could affect the folding of the chromatin fibre.


Figure 1
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  		Figure 1: Histone H2A variants and their involvement in nucleosome dynamics and conformation. (A) Amino acid sequence alignment of heteromorphous human H2A histone variants. The degree of similarity among the different sequences is proportional to the dark intensity of the shaded areas. Also shown are the {alpha}-helices and ß-turns corresponding to the histone-fold domain and macro domain of mH2A. (B) Nucleosomes are highly dynamic nucleoprotein complexes [4, 25]. In solution, the DNA associated with the histone octamer rapidly fluctuates between a folded (closed) (1) and an open (2) conformation [25, 33, 138]. Histone variants can influence this process which likely plays a very important role in the process of histone H2A–H2B dimer exchange (3) involved in chromatin metabolism. The different intensities of the grey arrows depict the importance of the H2A variants in shifting the equilibrium towards an open (1) or a tightly bound histone octamer (2). The shorter arrows in the case of H2A.X–H2B dimers indicate the low level of H2A.X exchange in the cell [77]. The question mark indicates where the relationship is still in question. Histone H2A–H2B displacement actively participates in replication [140], transcription [141], and DNA repair [142]. Displacement of the histone H2A–H2B dimers leads to a characteristic enhanced nuclease sensitivity in the region close to the dyad axis of the nucleosomal DNA (black arrowhead) [183]. (1) and (2) have been modelled after the crystallographic structure of the NCP [18]. For clarity, the two H2A–H2B dimers have been removed in (3).

 
Histone H2A.Bbd destabilizes the histone octamer
Histone H2A.Bbd appears to have occupied a primitive position in the lineage of the histone H2A family [38], but the occurrence of the gene and its expression did not occur until late in evolution during the appearance of vertebrates [39]. This histone variant which has only 48% identity to canonical H2A (Figure 1A) was first described in 2001 by Chadwick and Willard [40] and is shown to be excluded from the inactive X chromosome from where its name was derived.

The recombinant version of the protein does not form stable octamers when reconstituted with equimolar amounts of H3-H4-H2B [41, 42], although the mixture is able to form NCPs in the presence of DNA [41, 43]. However, H2A.Bbd-containing NCPs exhibit a relaxed conformation compared with the native NCPs under a broad range of salt concentrations as determined by analytical ultracentrifuge and by FRET [41, 43]. This is most likely the result of the flanking DNA being released from the NCP constraints [41] with the whole particle adopting a more open structure which is highly reminiscent of what is observed in NCPs with the highly acetylated histones [4]. Interestingly, H2A.Bbd was found to be co-localized with acetylated histones during both metaphase and interphase. Thus, it is possible to envision the two structural effects acting synergistically to produce a chromatin domain that is more amenable to transcription [44]. Indeed, FRAP experiments using GFP–H2A.Bbd transfected cells showed that this variant exchanges more rapidly than canonical H2A in chromatin, a fact that would be consistent with the exchange of H2A–H2B dimers associated with transcription (see subsequently) and with the lower stability of the H2A.Bbd histone octamer. The high dynamic flexibility of NCPs consisting of this histone variant when compared with NCPs consisting of other H2A variants is clearly reflected on the ease with which the former can be assembled and disassembled by nucleosome assembly protein (NAP-I) [45].

Histone H2A.Z—a histone variant with controversial roles
If there are any histone variants that have been most extensively studied from a functional perspective, these are without a doubt H2A.Z and H2A.X. Because of the extensive literature available in this and the following section, I am going to refer mainly to the recently published reviews focusing on the more limited amount of structural information available for both of these variants.

To date, histone H2A.Z is one of the few variants that has been shown to be indispensable for survival [46]. At the functional level, the role of this histone variant has been [47] and still remains extremely controversial, as claims for its involvement in gene inactivation [48] and activation (or both) [49, 50] are currently being made. In strong support of the silencing role comes the recent association of H2A.Z with heterochromatin binding protein (HP) (i.e. HP1-{alpha}) [51]. Equally supportive of an active role in transcription is the finding of the association of this variant with chromatin remodelling complexes involved in transcription such as SWR1 [52–55].

The functional controversy has been mirrored by the structural characterization of NCP consisting of H2A.Z. The first crystallographic structure of H2A.Z-containing NCPs suggested the existence of a ‘subtle destabilization of the interaction between the (H2A.Z–H2B) dimer and the (H3–H4) tetramer’ and predicted that the coexistence of canonical H2A–H2B and H2A.Z–H2B dimers within the same nucleosome is unlikely due to changes in the interfaces between the two types of dimers [56]. In agreement with the NCP destabilizing role, it was shown that NCPs reconstituted with recombinant-H2A.Z–H2B dimers exhibited an ionic strength-dependent reduced stability as analysed in the analytical ultracentrifuge [57]. H2A.Z-containing NCPs appear to have an enhanced thermal-dependent mobility [58]. However, a more recent characterization of the salt-dependent stability using FRET by the same group that crystallized the NCP indicated that H2A.Z stabilizes the histone octamer within the NCP [59]. Suggesting a H2A.Z-chromatin stabilizing role are also the early results from Jim Davie's lab using salt-elution of histones from hydroxyapatite (HAP)-adsorbed chromatin which showed that H2A.Z eluted at much higher salt concentrations than the canonical H2A–H2B dimer [60]. Nevertheless, all these data are seemingly hard to reconcile with the intrinsically reduced stability of the H2A.Z–H2B dimer itself [61].

Hence, like the functional studies, those dealing with the structural role of H2A.Z appear to be equally unsettled. A recent article from our group has shown the existence of two H2A.Z variants in chicken erythrocytes which only differ by three amino acids [62]. The presence of different subtypes of H2A.Z and/or the occurrence of differential PTMs (such as acetylation) [49, 63–65] may help to explain some of the structural and functional discrepancies described previously.

Histone H2A.X—maintaining chromosome integrity
Because of its connection with double-strand break (DSB) DNA repair and genome integrity, the functional role of this histone variant has been studied extensively [38, 66–68]. Histone H2A.X has also been involved in apoptosis [69], variable (diversity) joining [V(D)J] recombination [70], meiosis [71, 72] and replication [73]. Given all these important functional implications, it is not surprising that histone H2A.X has coevolved in parallel with the major canonical H2A variant [38]. Despite all this, the structural studies with this variant are lacking and only a limited amount of information is available. It is possible that like H2A.Z, an important structural role of this variant is mediated by its own PTMs and/or those of other histone nucleosomal partners [38] in addition to its interactions with non-nucleosomal partners such as the chromatin remodelling complexes Inositol 80-containing complex (INO80) and Swi2/Snf2-related adenosine triphosphate (SWRI) [74, 75].

In metazoans, H2A.X is evenly distributed throughout the genome with approximately one to two H2A.X molecules every ten nucleosomes [76]. FRAP results have shown that H2A.X has a low diffusional mobility [77]. Upon DSB damage, the H2A.X histones of extensive regions flanking the damaged site become reversibly phosphorylated [78]. Indeed, it was the discovery that the very C-terminal SQEY motif of mammalian H2A.X becomes phosphorylated by DNA-dependent protein kinase (DNA-PK) upon DSB damage [79] that brought much attention to this variant. Whether this phosphorylation has any effect on the structural characteristics of the NCP or the folding of the chromatin fibre remains yet to be elucidated. The only indirect information in this regard comes from a study carried out in yeast [80], where mutants were created in which the serine 129 of the native SQEL motif was replaced by glutamic acid (EQEL). The results [by supercoiling, micrococcal nuclease (MNase) digestions] obtained with these mutants suggested that chromatin adopts a more relaxed structure upon H2A.X phosphorylation [14, 80] which would be consistent with the homologous recombination (HR) repair mechanism in this organism. However, it remains to be determined whether this is also true in metazoans [which preferentially use non-homologous end-joining (NHEJ) for repair and HR in meiosis]. Whether or not this phosphorylation, which takes place in the vicinity of the histone H1 binding to the nucleosome, has any effect on such binding also needs to be established.

MacroH2A—a highly dynamic chromatin silencer
Like H2A.Bbd, mH2A is another recently discovered H2A variant, which, in contrast to the former, was first identified by its occurrence in the X-inactivated (Xi) chromosome of mammalian females [81–86]. Phylogenetically, mH2A has been a recent evolutionary acquisition which appeared during the evolution of vertebrates [38]. Approximately, one of every 30 nucleosomes in the cell contains one such variant [87]. The mH2A appears to be absent from the terminally differentiated cells such as the nucleated erythrocytes of fish, amphibian, reptiles and birds and from mature spermatozoa [88]. However, in addition to the inactive female X-chromosome, mH2A is also found in the tissues of vertebrate males that do not undergo this form of chromosome compensation [89]. Indeed, despite its absence from mature spermatozoa, mH2A has been found to be present during spermatogenesis (meiosis) [72, 90, 91] where it is associated with pericentric chromatin [92].

The mH2A is a heteromorphous H2A variant that has a tri-partite organization consisting of an N-terminal H2A-like histone-fold followed by a non-histone region (NHR) that contains a random coil and a C-terminal highly structured globular domain (Figure 1A). The latter has been called the ‘macro’ domain and shows high similarity to yeast macro domains [93]. Recent experimental evidence has shown that this domain can differentially bind ADP-ribosyl metabolites [88, 93, 94]. The NHR comprises two-thirds of the molecular mass of the whole protein.

This histone variant exists as two isoforms which are the product of alternative gene splicing: mH2A1.1, that accumulates throughout differentiation and development, and mH2A1.2, that has a constant level of expression [95]. The splicing site is located in a portion of the gene corresponding to the ligand binding pocket, suggesting a difference in the ability of the isoforms to bind nucleotide ligands [94]. Though the structural differences are minor, mH2A1.2 cannot bind nucleotides while mH2A1.1 can bind NAD metabolites [94]. Nevertheless, the functional significance of this is still unclear.

At the structural level, the crystallographic structure of the reconstituted NCP consisting of the H2A-like portion [96] as well as the macro domain have been recently published [94, 96]. Sedimentation velocity analysis demonstrated that in solution, the mH2A-NCPs had a very asymmetric conformation as a result of the NHR extending outwards. This is also supported by the ready accessibility of DNase I at the site where the random coil domain of the NHR exits the particle close to the dyad axis [88]. Sedimentation analysis using sucrose gradients revealed that mH2A stabilizes the NCP [89] in agreement with previous results using salt-dependent dissociation of histones from HAP-adsorbed chromatin [87, 88]. In this regard, it is important to notice that this variant has been shown to interfere with transcription factor binding and switch/sucrose non-fermenting (SWI/SNF) nucleosome remodelling [97]. Two-dimensional (2D)-polyacrylamide gel electrophoresis (PAGE), western blot analysis of the native mH2A-containing NCPs from chicken liver revealed the existence of two distinct populations, one of which consisted of an acid-labile PTM which was conditionally ascribed to poly-ADP ribosylation [89]. The low electrophoretic mobility fraction was found to be highly enriched in the MNase-resistant heterochromatin fraction [89].


   HISTONE H3 VARIANTS
TOP
 ABSTRACT
 INTRODUCTION
 DYNAMIC CHROMATIN
 HISTONE H2A VARIANTS SPECIALIZE...
 HISTONE H3 VARIANTS
OTHER CORE HISTONE VARIANTS
 CORE HISTONE VARIANTS AND...
 HISTONE H1--MICROHETEROGENEITY...
 CONCLUSIONS
 Acknowledgements
 References
 
Histone H3.3—the functional implications of small compositional changes
Histone H3.3 provides a very good example of homomorphous variation. In humans, only four amino acids are different in the composition of this histone when compared with the major canonical H3.1. In H3.3, S31 replaces A31 within the N-terminal domain and A87, G90, S96 replace S87, M90, C96 within the second helix of the histone-fold domain. During the course of evolution, the split between the two types appears to have occurred several times, and hence they cannot be considered separate lineages [98]. Despite all this, the two variants exhibit important functional differences. In contrast to H3.1, H3.3 has been shown to accumulate in some tissues during development in both humans and mice [99, 100], and it is enriched in actively transcribed regions of the genome [101] where it replaces H3.1 during transcriptional elongation [102]. The variant H3.3 also plays a very important role in spermatogenesis [72, 103].

Part of the involvement of this histone replacement variant in chromatin dynamics is mediated by the chaperone protein histone regulator A (HIRA). The HIRA complex participates in the assembly of H3.3–H4 dimers [104] into NCPs during the processes of transcription [104], replication [104] and in the assembly of the male pronucleus immediately after fertilization of the egg by sperm [105]. Remarkably however, recent evidence suggests that the amino acid differences between this histone variant and the canonical variant H3.1 may be enough to facilitate the NCP assembly/disassembly process alone [45]. The molecular details of how such small sequence compositional variability can account for this observation awaits further study.

CENP-A—defining the nucleosome structure of the centromeres
The hallmark of centromeric chromatin, CENP-A is a histone variant which replaces H3 in the NCP [106] (see [107–110] for recent reviews). As with H2A.Z, this variant is indispensable for survival [111]. This variant has also been referred to as chromatin-associated protein CSE4 (capping enzyme suppressor 4-p; (Cse4p) in yeast or centromere identifier (Cid) in Drosophila. It is one of the most rapidly evolving members of the histone H3 family which cannot be considered an independent lineage, but rather, an ‘orphan’ that has arisen several times during evolution [98] as a result of an adaptive process that affects the whole molecule [112].

From a structural point of view, the protein consists of a highly variable and essential N-terminal sequence (which varies inter-species in both length and composition) [113] followed by a histone-fold domain, which in the case of human CENP-A, displays 62% identity to the canonical H3 counterpart [114]. Mutagenic analysis in yeast has shown that both domains perform different essential roles [115].

At the nucleosome level, the information we have about the conformation imparted by this variant has been sparse but forthcoming in recent years. In vitro reconstitution experiments with CENP-A purified from HeLa cells were able to produce NCPs with DNase I footprints and AFM images almost identical to those of native NCPs [116]. Using deuterium exchange/mass spectrometry and hydrodynamic analysis, it has been recently shown that CENP-A and histone H4 form tetramers that are more compact and rigid than those assembled from canonical counterparts [117]. These features have been attributed to the synergistic action of the first loop (L1) and second {alpha}-helix ({alpha}-2) of the histone-fold of CENP-A and they are suggested to be responsible for targeting the CENP-A–H4 complexes to centromeres [117] in a way that is independent of the underlying DNA sequence. It is important to notice here that the sequence variability of H3.3 also affected the {alpha}-2 region of the corresponding histone-fold.

At the level of the chromatin fibre, the information available is much more limited. It is worth mentioning in this regard, that the long N-terminal region of histone H3 interacts with the linker DNA and is involved, together with histone H1, in the maintenance of the higher order structure of the chromatin fibre [118, 119]. Hence, it is quite possible that this region of CENP-A, which has been postulated to bind to the narrow groove of DNA [120], plays a critical role in further defining the topological characteristics of centromeric chromatin which has been shown to be more compact than pericentric or bulk chromatin [121]. Whether this is further facilitated by the additional binding of other centromeric proteins (CENP-B, CENP-C, CENP-H, CENP-I) [122, 123] and/or requires histone H1 has not yet been established.


   OTHER CORE HISTONE VARIANTS
TOP
 ABSTRACT
 INTRODUCTION
 DYNAMIC CHROMATIN
 HISTONE H2A VARIANTS SPECIALIZE...
 HISTONE H3 VARIANTS
 OTHER CORE HISTONE VARIANTS
CORE HISTONE VARIANTS AND...
 HISTONE H1--MICROHETEROGENEITY...
 CONCLUSIONS
 Acknowledgements
 References
 
The sperm-specific H2B variants
In great contrast to H2A and H3 histones, H2B and particularly H4 exhibit a low level of amino acid sequence variability, a fact that has been ascribed, at least in part, to the preferential role that these histones play in maintaining the interactions that hold together the histone octamer within the NCP [124]. Still, histone H2B has a relatively long N-terminal tail that protrudes from the NCP between the two DNA gyres [at superhelix location (SHL) 4.5 and –2.5/–3] [18], can form internucleosomal histone–DNA interactions [125] and plays an important role in NCP mobility and dynamics [126, 127]. Therefore, histone variation within this domain may have important chromatin conformational implications.

Interestingly, most of the variability observed in H2B seems to occur in the sperm of vertebrate [90] and invertebrate [128] organisms and in male gametes in plants [129, 130]. In humans, three testis-specific variants have been described: TH2B [131], human testis-specific H2B (hTSH2B) [132] and H2B family member W testis-specific (H2BFWT) [133].

The long N-terminal domains of echinoderm sperm histones consist of highly characteristic repetitive motifs, SPR/KR/K [128], that are well-known to bind to the minor groove of DNA [134]. These tails may participate in maintaining a distinctive NCP organization [135] leading to a high extent of chromatin compaction observed in the sperm of these organisms. This compaction probably results from partial DNA charge neutralization and by establishing internucleosomal interactions.

As for vertebrate variants, hTSH2B is the only H2B human sperm histone variant whose role in nucleosome conformation has been recently characterized in detail [136]. As in the case of H2A.Bbd, it was found that that hTSH2B lowered the stability of the histone octamer without affecting its ability to produce NCPs that were structurally and dynamically undistinguishable from particles consisting of canonical H2B [136]. The source of sequence variation of hTSH2B resides mainly in the occurrence of amino acid transitions to S/T residues. It is thus possible that the structural effects of this H2B variant on the NCP and chromatin fibre are phosphorylation mediated. However, despite this rather isolated work, structural studies on other vertebrate sperm H2B variants are for their most part still lacking, and hence their implications for chromatin are unknown.

In an interesting twist to the H2A.Z research, it has very recently been found that in Trypanosoma brucei, this histone variant pairs with a rather specialized H2BV variant [137]. The intriguing possibility of this being the case in higher eukaryotes deserves further exploration. As pointed out by the authors of this article, there are several potential H2B candidates in the human genome for this role.


   CORE HISTONE VARIANTS AND CHROMATIN DYNAMICS
TOP
 ABSTRACT
 INTRODUCTION
 DYNAMIC CHROMATIN
 HISTONE H2A VARIANTS SPECIALIZE...
 HISTONE H3 VARIANTS
 OTHER CORE HISTONE VARIANTS
 CORE HISTONE VARIANTS AND...
HISTONE H1--MICROHETEROGENEITY...
 CONCLUSIONS
 Acknowledgements
 References
 
In solution and within the cell, nucleosomes are highly dynamic structures. In vitro, the conformation of the NCP fluctuates between folded and unfolded states [25, 33] (Figure 1B) with an affinity of the histone octamer for the template DNA that depends on both temperature and octamer concentration [138] as well as the ionic conditions [139]. In vivo, most of the metabolic functions of chromatin involve the dynamic exchange of histone H2A–H2B dimers. There is now evidence of different sorts to support this exchange during DNA replication [140], transcription [141] and repair [142]. Although in many instances the process is facilitated or assisted by ATP-dependent [143, 144] and independent chromatin assembly complexes (such as NAP-1, nucleoplasmin, chromatin assembly factor-1(CAF-1) or SWI/SNF, or facilitates transcription (FACT) just to mention a few) [145–148], the intrinsic NCP plasticity in solution and the incorporation of histone variants also play a critical role.

The sections described above provide a glimpse of how core histone variants themselves can actively participate in nucleosome dynamics. Core histones of the H2A family of variants provide several good examples. The FRET studies on H2A.Bbd [43] indicate that this variant can exchange very readily between different chromatin domains in a way that most likely drives the equilibrium between the closed (1) and open conformation (2) of the NCP shown in Figure 1B towards an open nucleosome conformation (2) as indicated by the thickness of arrow 2 in this figure.

In contrast to H2A.Bbd, the salt resilience toward NCP destabilization displayed by mH2A [88, 89] suggests that in this instance the equilibrium is shifted in the opposite direction (Figure 1B). It is important to note here that the shift in the equilibrium appears to be only temporary. The presence of mH2A during spermiogenesis and its absence from terminally differentiated cells suggests that the inactivation role of this variant is transient. It is likely that this variant dynamically exchanges to temporarily maintain certain repressed states of chromatin, but that other variants are responsible for a more permanent structures. In agreement with this, it has been shown that mH2A exhibits a mutually exclusive relationship with linker histones, suggesting some functional redundancy between these proteins [88].

Despite its even distribution throughout the genome and the high extent of similarity to the canonical form (Figure 1A), H2A.X appears to exhibit a highly restricted mobility [77] (Figure 1B). With H2A.Z the situation appears to be more complex. The most recent structural results imply an intra- and inter-nucleosomal stabilizing role for this variant [27, 149] that in the latter instance has been ascribed to an enhanced interaction between the acidic C-terminal patch of this variant and histone H4 of adjacent nucleosomes [51]. Nevertheless, although supported by early crystallographic data [18], this interpretation contrasts with the polymorphic packing of nucleosomes within the recently published structure of a tetranucleosome complex [150], which is likely to be enhanced within the more structurally heterogeneous chromatin fibre.

A theoretical study using elastic network models [151] based on the plethora of NCP crystal structures currently available [19] has shown that in general, histone variants exhibit higher motilities and weaker correlations between internal motions in NCPs than those displayed by the canonical counterparts [151]. However, the extent with which the amino acid changes involved in histone variation affect the protein–protein interfaces and local alterations of the ionic environment [152] to contribute to this enhanced plasticity, still need to be experimentally determined. This knowledge is imperative for a complete understanding of the detailed molecular events involved in the dynamic equilibria shown in Figure 1B.


   HISTONE H1—MICROHETEROGENEITY MATTERS
TOP
 ABSTRACT
 INTRODUCTION
 DYNAMIC CHROMATIN
 HISTONE H2A VARIANTS SPECIALIZE...
 HISTONE H3 VARIANTS
 OTHER CORE HISTONE VARIANTS
 CORE HISTONE VARIANTS AND...
 HISTONE H1--MICROHETEROGENEITY...
CONCLUSIONS
 Acknowledgements
 References
 
Eleven different linker histone variants have been identified to date in mammals: Seven somatic variants: (H1.1–H1.5, H10 and H1x) [4, 153–155], three spermatogenic variants: H1t [156], H1T2 [157] and HILS1 [158, 159] and an oocyte-specific H1foo [160]. Highly specialized histone H1 variants are also expressed in the sperm of many invertebrates including the long histone H1s from the sperm of echinoderms [128] and the highly specialized protamine-like protein I (PL-I) that are thought to be evolutionary linked to protamines (see Eirin-Lopez et al. [161] for a recent review). Except for H1.1–H1.5, most of the other histone H1 variants are expressed in a replication-independent cell cycle and development-dependent way [162]. In this section, I am going to focus mainly on recent functional and structural aspects of somatic histone H1 microheterogeneity. For a recent review on developmentally regulated histone H1 variants, the reader is referred to Khochbin [162].

Interestingly, the concept of histone variants within this family and of histone sequence variability itself had its origins in the early work on calf thymus H1 histones by Kinkade and Cole [163, 164] who noted that somatic histone H1 exhibited sequence microheterogeneity (Figure 2A). Although Cole [165, 166] himself wrote several reviews about the possible implications of this histone H1 microheterogeneity (see also for a more recent review [167]), evidence for a multifaceted functionality of this family both at the level of somatic microheterogeneity (H1.1–H1.5) (Figure 2A) and at the level of differentiation-specific variants (H10, H1t) has long remained elusive. Indeed, and quite unexpectedly, experimentally costly knock out-experiments carried out in different organisms in recent years seem to suggest that the function of this variability may have a large overlapping redundancy (reviewed in [4, 168]).


Figure 2
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  		Figure 2: Histone H1 microheterogeneity and chromatin dynamics. (A) Amino acid sequence alignment of the human somatic H1 histones (H1.1–H1.5) to highlight the extent of microheterogeneity between these proteins. The degree of similarity among the different sequences is proportional to the dark intensity of the shaded areas. The nomenclature followed for the designation of these variants is that of Albig and his collegues [154] with the nomenclature of Seyedin and Kistler [184] shown in parentheses. The {alpha}-helices and ß-turns of the winged helix globular domain are indicated. (B) Schematic representation of the dynamic model of linker histone binding to chromatin [177]. Histone H1 microheterogeneous variants (black dots) are continuously exchanging at different rates [178] between folded and unfolded regions of chromatin (indicated by the reversible arrows). Local concentration may change in response to different stimuli (initiation of transcription, transcriptional elongation, DNA repair, DNA replication) that may involve selective PTMs of the linker histones being exchanged as well as core histone PTMs and the differential presence of core histone variants themselves. This dynamic process is in addition influenced by a broader network of chromatin interacting proteins [177, 180] such as heterochromatin binding proteins (HP) (i.e. HP1-{alpha}) and methyl binding domain proteins (MBD) such as MeCP2 and high mobility group proteins (HMG).

 
Despite this, the non-random distribution of linker histone variants in the genome has now been well-documented [169] and mammalian linker histone variants have been shown to differentially affect gene expression in vivo [170]. Not only that, but histone H1.2 is able to preferentially bind to a regulatory sequence of the gene for histone H3.2 [171] has been directly involved in DNA DSB-induced apoptosis [172], and interacts with Msx1 (a transcription factor involved in myogenic gene expression) [173]. Furthermore, there is recent evidence showing that the long-term evolution of the replication-dependent and replication-independent somatic linker histone variants (H1.1–H1.5) has taken place though a process of ‘birth-and-death’ evolution with a strong purifying selection [174, 175]. Accordingly, the degree of interspecific conservation of these variants is higher than that observed at the intraspecific level [174]; an observation which also comes in support of specialized structural and functional roles of these variants. Thus, although the functional specificity of linker histone variants does not appear to be as well-defined as with core histone variants, all of the above evidence suggests that their structural variability is functionally relevant.

The less defined functional character of linker histones can perhaps be better understood from the structural details of their association with chromatin. In comparison with core histones, linker histones have long been known to be less tightly associated with chromatin and much more mobile [176], a fact that has been recently corroborated by in situ FRAP experiments [21, 22]. The rapid and dynamic exchange of histone H1 between different segments of chromatin (Figure 2B) is a very important determinant of the extent of fibre folding and interchromatin fibre association. A model for the dynamic association of linker histones with chromatin [177] is shown in Figure 2B. Accordingly, histone H1 is dynamically associated with the chromatin fibre in a transient mode. Recent FRAP experiments have shown the existence of ‘dramatic’ differences in the binding affinities of the different H1 subtypes [178]. Hence, their movement and local distribution is likely to vary from one type to another. It will additionally depend on other factors such as: core histone variant (and PTM) composition of individual nucleosomes [14] along the fibre, transcription activation pathways [177], and/or the extent of the histone H1 PTMs themselves (i.e. phosphorylation) [178, 179]. Furthermore, linker histones may operate in conjunction with a ‘network’ of other chromatin-binding proteins such as high mobility group (HMG) proteins, HP1 or methyl CpG-binding protein (MeCP2) [177,180] so as to define permissive (euchromatin) and repressive (heterochromatin) DNA domains. Therefore, as it was stated at the beginning of this section, if the main role of linker histones is to maintain and stabilize the folding of chromatin, it should not be entirely surprising that there is some degree of redundancy despite their well-supported structural and functional specialization.


   CONCLUSIONS
TOP
 ABSTRACT
 INTRODUCTION
 DYNAMIC CHROMATIN
 HISTONE H2A VARIANTS SPECIALIZE...
 HISTONE H3 VARIANTS
 OTHER CORE HISTONE VARIANTS
 CORE HISTONE VARIANTS AND...
 HISTONE H1--MICROHETEROGENEITY...
 CONCLUSIONS
Acknowledgements
 References
 
Structural evidence gathered so far indicates that the amino acid sequence variability of histones constrained and maintained throughout evolution can, by itself or in conjunction with PTMs, regulate chromatin dynamics in a way that has important functional implications. The epigenetic language of the histone PTMs (histone code) is ultimately determined by this prevalent underlying (amino acid code) of the different histone variants. At the core histone level, a representative example of this can be found in the SQE[Y/L/F] motif at the C-terminal end of H2A.X that provides a sequence motif specificity for the DNA-PK involved in double-stranded DNA repair [38]. At the linker histone level, the yet to be clearly established sequence motif responsible for the apoptotic specificity of H1.2 [172] provides another example.

The histone-fold, the winged helix domain and the ‘disordered’ N- and C-terminal tails [181] are ‘simple’ structures that could be achieved through many different amino acid combinations. The fact that the amino acid locations in the molecule are so conserved throughout evolution within the different variants indicate that most of the amino acid variation observed is functionally relevant. Deciphering the details of the amino acid code responsible for the manifold structural and functional implications of all the histone variants such as, for instance, the amino acid motif involved in the recognition of H2A.Z by SWR1 and/or INO80 may still take some time. This effort, however, may prove very rewarding in understanding the overall epigenetic dimension of these ‘fairly simple proteins’ we call histones.

In the interim, it may also be worthwhile to pursue some additional questions raised by several sections of this review. For instance, do the different docking domains of H2A variants prevent them from forming heterologous NCPs consisting of H2A–H2B dimers with mixed variants? What are the histone H2B variant partners of the different H2A variants? In this regard, the recent finding that H2A.Z interacts with a highly specialized H2BV in T. brucei [137] is very insightful and hopefully similar analyses carried out in metazoans will soon follow. What is the structural and functional relevance of H2A.Z sequence microheterogeneity? Does the compositional microheterogeneity of H3 variants affect the structure of NCPs? Given the instability of the H2A.Z–H2B dimer, how does the replacement of the canonical dimer take place? In the case of H3.3, it appears that this variant interacts with chaperone proteins that may mediate its exchange with canonical H3 during elongation in actively transcribed genes. The recent observation that this variant may operate synergistically with other variants such as H2A.Bbd to facilitate the assembly and disassembly process [45] sheds important light on some of the possible mechanisms involved. However, considering that H3.3 is by far, much more abundant than H2A.Bbd, the full implications of H3.3 involvement in the loss of NCP stability and the molecular details of the exchange process need yet to be established.

Considering the intensity of the research, currently ongoing, on the topic of this review which can be easily attested by the large number of other recent reviews in the same area of chromatin research [38, 39, 124, 152, 182], the answer to several of these questions are likely to be forthcoming soon.


Key Points

  • Chromatin is a dynamic nucleoprotein complex whose structural characterization requires the use of appropriate biophysical tools (such as, for instance, analytical ultracentrifuge, FRAP and FRET) in addition to the powerful static data provided by X-ray crystallography.
  • The histone H2A family includes a broad spectrum of variants whose functional implications for chromatin have been extensively characterized. However, the structural aspects behind the function are in many instances still not clear and/or controversial (i.e. H2A.Z and phosphorylated H2A.X).
  • Core histone variants and their PTMs play a critical role in the modulation of chromatin dynamics.
  • Histone H1 microheterogeneity exhibits a non-random distribution in the genome and is an important component of chromatin metabolism. The latter is mediated by the highly dynamic differential association of the distinct H1 variants with chromatin.

 


   Acknowledgements
TOP
 ABSTRACT
 INTRODUCTION
 DYNAMIC CHROMATIN
 HISTONE H2A VARIANTS SPECIALIZE...
 HISTONE H3 VARIANTS
 OTHER CORE HISTONE VARIANTS
 CORE HISTONE VARIANTS AND...
 HISTONE H1--MICROHETEROGENEITY...
 CONCLUSIONS
 Acknowledgements
References
 
I would like to thank Ron Finn and José Maria Eirín Lopez for their skilful computer assistance and help in the preparation of the figures and to Wade Abbott, Lindsay Frehlick and Anita Thambirajah for their critical and careful reading of the manuscript. This work was supported by Canadian Institutes of Health Research (CIHR) grant MOP-57718.


   FOOTNOTES
 
Juan Ausió (PhD, University of Barcelona) has worked in the area of structural and functional characterization of histones and chromatin since 1976. He has made important contributions to the biophysical characterization of the nucleosome core particle in solution [Weizmann Institute of Science, Israel (1981–84)] and acetylated chromatin [Biochemistry and Biophysics Department, Oregon State University (1984–86)]. His current work on histone variants is carried out at the Department of Biochemistry and Microbiology at the University of Victoria where he is a Professor.


   References
TOP
 ABSTRACT
 INTRODUCTION
 DYNAMIC CHROMATIN
 HISTONE H2A VARIANTS SPECIALIZE...
 HISTONE H3 VARIANTS
 OTHER CORE HISTONE VARIANTS
 CORE HISTONE VARIANTS AND...
 HISTONE H1--MICROHETEROGENEITY...
 CONCLUSIONS
 Acknowledgements
 References
 

  1. Stedman E, Stedman E. The chemical nature and functions of the components of the cell nuclei. Cold Spring Harb Symp Quant Biol 1947; 12:224–36.[Abstract/Free Full Text]
  2. Tsanev R, Sendov B. Possible molecular mechanism for cell differentiation in multicellular organisms. J Theor Biol 1971; 30:337–93.[CrossRef][Web of Science][Medline]
  3. Strahl BD, Allis CD. The language of covalent histone modifications. Nature 2000; 403:41–5.[CrossRef][Medline]
  4. Ausió J, Abbott DW. The Role of Histone Variability in Chromatin Stability and Folding. Amsterdam, The Netherlands: Elsevier 2004.
  5. Turner BM. Reading signals on the nucleosome with a new nomenclature for modified histones. Nat Struct Mol Biol 2005; 12:110–2.[CrossRef][Web of Science][Medline]
  6. Sarma K, Reinberg D. Histone variants meet their match. Nat Rev Mol Cell Biol 2005; 6:139–49.[CrossRef][Web of Science][Medline]
  7. Bradbury EM. Histone Nomenclature. Amsterdam: Associated Scientific Publ 1975.
  8. Bradbury EM. Histone nomenclature. Methods Cell Biol 1977; 16:179–81.[Medline]
  9. DeLange RJ, Smith EL. Chromosomal Proteins. NY: Academic Press 1979.
  10. Wu RS, Panusz HT, Hatch CL, et al. Histones and their modifications. CRC Crit Rev Biochem 1986; 20:201–63.[Web of Science][Medline]
  11. Stellwagen RH, Cole RD. Chromosomal proteins. Annu Rev Biochem 1969; 38:951–90.[CrossRef][Web of Science][Medline]
  12. Grove GW, Zweidler A. Regulation of nucleosomal core histone variant levels in differentiating murine erythroleukemia cells. Biochemistry 1984; 23:4436–43.[CrossRef][Medline]
  13. West MH, Bonner WM. Histone 2A, a heteromorphous family of eight protein species. Biochemistry 1980; 19:3238–45.[CrossRef][Medline]
  14. Ausió J, Abbott DW, Wang X, et al. Histone variants and histone modifications: a structural perspective. Biochem Cell Biol 2001; 79:693–708.[CrossRef][Web of Science][Medline]
  15. van Holde KE. Chromatin. NY: Springer-Verlag 1988.
  16. Arents G, Moudrianakis EN. The histone fold: a ubiquitous architectural motif utilized in DNA compaction and protein dimerization. Proc Natl Acad Sci USA 1995; 92:11170–4.[Abstract/Free Full Text]
  17. Graziano V, Gerchman SE, Wonacott AJ, et al. Crystallization of the globular domain of histone H5. J Mol Biol 1990; 212:253–7.[CrossRef][Web of Science][Medline]
  18. Luger K, Mader AW, Richmond RK, et al. Crystal structure of the nucleosome core particle at 2.8 A resolution. Nature 1997; 389:251–60.[CrossRef][Medline]
  19. Chakravarthy S, Park YJ, Chodaparambil J, et al. Structure and dynamic properties of nucleosome core particles. FEBS Lett 2005; 579:895–8.[CrossRef][Web of Science][Medline]
  20. Ausió J. Analytical ultracentrifugation and the characterization of chromatin structure. Biophysical Chemistry 2000; 86:141–53.[CrossRef][Web of Science][Medline]
  21. Misteli T, Gunjan A, Hock R, et al. Dynamic binding of histone H1 to chromatin in living cells. Nature 2000; 408:877–81.[CrossRef][Medline]
  22. Lever MA, Th’ng JP, Sun X, et al. Rapid exchange of histone H1.1 on chromatin in living human cells. Nature 2000; 408:873–6.[CrossRef][Medline]
  23. Kimura H. Histone dynamics in living cells revealed by photobleaching. DNA Repair (Amst) 2005; 4:939–50.[CrossRef][Medline]
  24. Selvin PR. The renaissance of fluorescence resonance energy transfer. Nat Struct Biol 2000; 7:730–4.[CrossRef][Web of Science][Medline]
  25. Li G, Levitus M, Bustamante C, et al. Rapid spontaneous accessibility of nucleosomal DNA. Nat Struct Mol Biol 2005; 12:46–53.[CrossRef][Web of Science][Medline]
  26. Li G, Widom J. Nucleosomes facilitate their own invasion. Nat Struct Mol Biol 2004; 11:763–9.[CrossRef][Web of Science][Medline]
  27. Park YJ, Dyer PN, Tremethick DJ, et al. A new fluorescence resonance energy transfer approach demonstrates that the histone variant H2AZ stabilizes the histone octamer within the nucleosome. J Biol Chem 2004; 279:24274–82.[Abstract/Free Full Text]
  28. Anderson JD, Widom J. Sequence and position-dependence of the equilibrium accessibility of nucleosomal DNA target sites. J Mol Biol 2000; 296:979–87.[CrossRef][Web of Science][Medline]
  29. Widom J. Equilibrium and dynamic nucleosome stability. Methods Mol Biol 1999; 119:61–77.[Medline]
  30. Jaskelioff M, Gavin IM, Peterson CL, et al. SWI-SNF-mediated nucleosome remodeling: role of histone octamer mobility in the persistence of the remodeled state. Mol Cell Biol 2000; 20:3058–68.[Abstract/Free Full Text]
  31. Leuba SH, Bennink ML, Zlatanova J. Single-molecule analysis of chromatin. Methods Enzymol 2004; 376:73–105.[CrossRef][Web of Science][Medline]
  32. Zlatanova J, Leuba SH. Chromatin fibres, one-at-a-time. J Mol Biol 2003; 331:1–19.[CrossRef][Web of Science][Medline]
  33. Tomschik M, Zheng H, van Holde K, et al. Fast, long-range, reversible conformational fluctuations in nucleosomes revealed by single-pair fluorescence resonance energy transfer. Proc Natl Acad Sci USA 2005; 102:3278–83.[Abstract/Free Full Text]
  34. Eickbush TH, Watson DK, Moudrianakis EN. A chromatin-bound proteolytic activity with unique specificity for histone H2A. Cell 1976; 9:785–92.[CrossRef][Web of Science][Medline]
  35. Eickbush TH, Godfrey JE, Elia MC, et al. H2a-specific proteolysis as a unique probe in the analysis of the histone octamer. J Biol Chem 1988; 263:18972–8.[Abstract/Free Full Text]
  36. Usachenko SI, Bavykin SG, Gavin IM, et al. Rearrangement of the histone H2A C-terminal domain in the nucleosome. Proc Natl Acad Sci U S A 1994; 91:6845–9.[Abstract/Free Full Text]
  37. Zhou YB, Gerchman SE, Ramakrishnan V, et al. Position and orientation of the globular domain of linker histone H5 on the nucleosome. Nature 1998; 395:402–5.[CrossRef][Medline]
  38. Li A, Eirin-Lopez JM, Ausio J. H2AX: tailoring histone H2A for chromatin-dependent genomic integrity. Biochem Cell Biol 2005; 83:505–15.[CrossRef][Web of Science][Medline]
  39. Henikoff S, Furuyama T, Ahmad K. Histone variants, nucleosome assembly and epigenetic inheritance. Trends Genet 2004; 20:320–6.[CrossRef][Web of Science][Medline]
  40. Chadwick BP, Willard HF. A novel chromatin protein, distantly related to histone H2A, is largely excluded from the inactive X chromosome. J Cell Biol 2001; 152:375–84.[Abstract/Free Full Text]
  41. Bao Y, Konesky K, Park YJ, et al. Nucleosomes containing the histone variant H2A.Bbd organize only 118 base pairs of DNA. Embo J 2004; 23:3314–24.[CrossRef][Web of Science][Medline]
  42. Finn RM, Abbott WD. Ausió J. H2A quo vadis? The role of histoneH2A variability, Chemtracts 2005. (in press).
  43. Gautier T, Abbott DW, Molla A, et al. Histone variant H2ABbd confers lower stability to the nucleosome. EMBO Rep 2004; 5:715–20.[CrossRef][Web of Science][Medline]
  44. Angelov D, Verdel A, An W, et al. SWI/SNF remodeling and p300-dependent transcription of histone variant H2ABbd nucleosomal arrays. Embo J 2004; 23:3815–24.[CrossRef][Web of Science][Medline]
  45. Okuwaki M, Kato K, Shimahara H, et al. Assembly and disassembly of nucleosome core particles containing histone variants by human nucleosome assembly protein I. Mol Cell Biol 2005; 25:10639–51.[Abstract/Free Full Text]
  46. Clarkson MJ, Wells JR, Gibson F, et al. Regions of variant histone His2AvD required for Drosophila development. Nature 1999; 399:694–7.[CrossRef][Medline]
  47. Dryhurst D, Thambirajah AA, Ausio J. New twists on H2A.Z: a histone variant with a controversial structural and functional past. Biochem Cell Biol 2004; 82:490–7.[CrossRef][Web of Science][Medline]
  48. Guillemette B, Bataille AR, Gevry N, et al. Variant histone H2A.Z is globally localized to the promoters of inactive yeast genes and regulates nucleosome positioning. PLoS Biol 2005; 3:e384.[CrossRef][Medline]
  49. Bruce K, Myers FA, Mantouvalou E, et al. The replacement histone H2A.Z in a hyperacetylated form is a feature of active genes in the chicken. Nucleic Acids Res 2005; 33:5633–9.[Abstract/Free Full Text]
  50. Raisner RM, Hartley PD, Meneghini MD, et al. Histone variant H2A.Z marks the 5' ends of both active and inactive genes in euchromatin. Cell 2005; 123:233–48.[CrossRef][Web of Science][Medline]
  51. Fan JY, Rangasamy D, Luger K, et al. H2A.Z alters the nucleosome surface to promote HP1alpha-mediated chromatin fibre folding. Mol Cell 2004; 16:655–61.[CrossRef][Web of Science][Medline]
  52. Wu WH, Alami S, Luk E, et al. Swc2 is a widely conserved H2AZ-binding module essential for ATP-dependent histone exchange. Nat Struct Mol Biol 2005; 12:1064–71.[CrossRef][Web of Science][Medline]
  53. Krogan NJ, Keogh MC, Datta N, et al. A Snf2 family ATPase complex required for recruitment of the histone H2A variant Htz1. Mol Cell 2003; 12:1565–76.[CrossRef][Web of Science][Medline]
  54. Mizuguchi G, Shen X, Landry J, et al. ATP-driven exchange of histone H2AZ variant catalyzed by SWR1 chromatin remodeling complex. Science 2004; 303:343–8.[Abstract/Free Full Text]
  55. Kobor MS, Venkatasubrahmanyam S, Meneghini MD, et al. A protein complex containing the conserved Swi2/Snf2-related ATPase Swr1p deposits histone variant H2A.Z into euchromatin. PLoS Biol 2004; 2:E131.[CrossRef][Medline]
  56. Suto RK, Clarkson MJ, Tremethick DJ, et al. Crystal structure of a nucleosome core particle containing the variant histone H2A.Z. Nature Struct Biol 2000; 7:1121–4.[CrossRef][Web of Science][Medline]
  57. Abbott DW, Ivanova VS, Wang X, et al. Characterization of the stability and folding of H2A.Z chromatin particles: implications for transcriptional activation. J Biol Chem 2001; 276:41945–9.[Abstract/Free Full Text]
  58. Flaus A, Rencurel C, Ferreira H, et al. Sin mutations alter inherent nucleosome mobility. Embo J 2004; 23:343–53.[CrossRef][Web of Science][Medline]
  59. Park Y-J, Dyer PN, Tremethick DJ, Luger K. A new FRET approach demonstrates that the histone variant H2A.Z stabilizes the histone octamer within the nucleosome. J Biol Chem 2004; 279:24274–82.[Abstract/Free Full Text]
  60. Li W, Nagaraja S, Delcuve GP, et al. Effects of histone acetylation, ubiquitination and variants on nucleosome stability. Biochem J 1993; 296:(Pt 3)737–44.[Web of Science][Medline]
  61. Placek BJ, Harrison LN, Villers BM, et al. The H2A.Z/H2B dimer is unstable compared to the dimer containing the major H2A isoform. Protein Sci 2005; 14:514–22.[CrossRef][Web of Science][Medline]
  62. Coon JJ, Ueberheide B, Syka JE, et al. Protein identification using sequential ion/ion reactions and tandem mass spectrometry. Proc Natl Acad Sci USA 2005; 102:9463–8.[Abstract/Free Full Text]
  63. Babiarz JE, Halley JE, Rine J. Telomeric heterochromatin boundaries require NuA4-dependent acetylation of histone variant H2A.Z in Saccharomyces cerevisiae. Genes Dev 2006; 20:700–10.[Abstract/Free Full Text]
  64. Keogh MC, Mennella TA, Sawa C, et al. The Saccharomyces cerevisiae histone H2A variant Htz1 is acetylated by NuA4. Genes Dev 2006; 20:660–5.[Abstract/Free Full Text]
  65. Millar CB, Xu F, Zhang K, et al. Acetylation of H2AZ Lys 14 is associated with genome-wide gene activity in yeast. Genes Dev 2006; 20:711–22.[Abstract/Free Full Text]
  66. Foster ER, Downs JA. Histone H2A phosphorylation in DNA double-strand break repair. Febs J 2005; 272:3231–40.[CrossRef][Medline]
  67. Thiriet C, Hayes JJ. Chromatin in need of a fix: phosphorylation of H2AX connects chromatin to DNA repair. Mol Cell 2005; 18:617–22.[CrossRef][Web of Science][Medline]
  68. Moore JD, Krebs JE. Histone modifications and DNA double-strand break repair. Biochem Cell Biol 2004; 82:446–52.[CrossRef][Web of Science][Medline]
  69. Rogakou EP, Nieves-Neira W, Boon C, et al. Initiation of DNA fragmentation during apoptosis induces phosphorylation of H2AX histone at serine 139. J Biol Chem 2000; 275:9390–5.[Abstract/Free Full Text]
  70. Chen HT, Bhandoola A, Difilippantonio MJ, et al. Response to RAG-mediated VDJ cleavage by NBS1 and gamma-H2AX. Science 2000; 290:1962–5.[Abstract/Free Full Text]
  71. Mahadevaiah SK, Turner JM, Baudat F, et al. Recombinational DNA double-strand breaks in mice precede synapsis. Nature Genetics 2001; 27:271–6.[CrossRef][Web of Science][Medline]
  72. Lewis JD, Abbott DW, Ausio J. A haploid affair: core histone transitions during spermatogenesis. Biochemistry and Cell Biology 2003; 81:131–40.[CrossRef][Web of Science][Medline]
  73. Ward IM, Chen J. Histone H2AX is phosphorylated in an ATR-dependent manner in response to replicational stress. J Biol Chem 2001; 276:47759–62.[Abstract/Free Full Text]
  74. Morrison AJ, Highland J, Krogan NJ, et al. INO80 and gamma-H2AX interaction links ATP-dependent chromatin remodeling to DNA damage repair. Cell 2004; 119:767–5.[CrossRef][Web of Science][Medline]
  75. Downs JA, Allard S, Jobin-Robitaille O, et al. Binding of chromatin-modifying activities to phosphorylated histone H2A at DNA damage sites. Mol Cell 2004; 16:979–90.[CrossRef][Web of Science][Medline]
  76. Pilch DR, Sedelnikova OA, Redon C, et al. Characteristics of gamma-H2AX foci at DNA double-strand breaks sites. Biochem Cell Biol 2003; 81:123–9.[CrossRef][Web of Science][Medline]
  77. Siino JS, Nazarov IB, Svetlova MP, et al. Photobleaching of GFP-labeled H2AX in chromatin: H2AX has low diffusional mobility in the nucleus. Biochem Biophys Res Commun 2002; 297:1318–23.[CrossRef][Web of Science][Medline]
  78. Keogh MC, Kim JA, Downey M, et al. A phosphatase complex that dephosphorylates gammaH2AX regulates DNA damage checkpoint recovery. Nature 2005; 439:497–501.
  79. Rogakou EP, Pilch DR, Orr AH, et al. DNA double-stranded breaks induce histone H2AX phosphorylation on serine 139. J Biol Chem 1998; 273:5858–68.[Abstract/Free Full Text]
  80. Downs JA, Lowndes NF, Jackson SP. A role for Saccharomyces cerevisiae histone H2A in DNA repair. Nature 2000; 408:1001–4.[CrossRef][Medline]
  81. Chadwick BP, Willard HF. Cell cycle-dependent localization of macroH2A in chromatin of the inactive X chromosome. J Cell Biol 2002; 157:1113–23.[Abstract/Free Full Text]
  82. Costanzi C, Pehrson JR. Histone macroH2A1 is concentrated in the inactive X chromosome of female mammals. Nature 1998; 393:599–601.[CrossRef][Medline]
  83. Pehrson JR, Fuji RN. Evolutionary conservation of histone macroH2A subtypes and domains. Nucleic Acids Research (Online) 1998; 26:2837–42.
  84. Chadwick BP, Valley CM, Willard HF. Histone variant macroH2A contains two distinct macrochromatin domains capable of directing macroH2A to the inactive X chromosome. Nucleic Acids Res 2001; 29:2699–705.[Abstract/Free Full Text]
  85. Chadwick BP, Willard HF. Barring gene expression after XIST: maintaining facultative heterochromatin on the inactive X. Semin Cell Dev Biol 2003; 14:359–67.[CrossRef][Web of Science][Medline]
  86. Chadwick BP, Willard HF. Chromatin of the Barr body: histone and non-histone proteins associated with or excluded from the inactive X chromosome. Hum Mol Genet 2003; 12:2167–78.[Abstract/Free Full Text]
  87. Pehrson JR, Fried VA. MacroH2A, a core histone containing a large nonhistone region. Science 1992; 257:1398–400.[Abstract/Free Full Text]
  88. Abbott DW, Laszczak M, Lewis JD, et al. Structural characterization of macroH2A containing chromatin. Biochemistry 2004; 43:1352–9.[CrossRef][Medline]
  89. Abbott DW, Chadwick BP, Thambirajah AA, et al. Beyond the Xi: macroH2A chromatin distribution and post-translational modification in an avian system. J Biol Chem 2005; 280:16437–45.[Abstract/Free Full Text]
  90. Churikov D, Zalenskaya IA, Zalensky AO. Male germline-specific histones in mouse and man. Cytogenet Genome Res 2004; 105:203–14.[CrossRef][Web of Science][Medline]
  91. Govin J, Caron C, Lestrat C, et al. The role of histones in chromatin remodelling during mammalian spermiogenesis. Eur J Biochem 2004; 271:3459–69.[Web of Science][Medline]
  92. Turner JM, Burgoyne PS, Singh PB. M31 and macroH2A1.2 colocalise at the pseudoautosomal region during mouse meiosis. J Cell Sci 2001; 114:3367–75.[Medline]
  93. Karras GI, Kustatscher G, Buhecha HR, et al. The macro domain is an ADP-ribose binding module. EMBO J 2005; 24:1911–20.[CrossRef][Web of Science][Medline]
  94. Kustatscher G, Hothorn M, Pugieux C, et al. Splicing regulates NAD metabolite binding to histone macroH2A. Nat Struct Mol Biol 2005; 12:624–5.[CrossRef][Web of Science][Medline]
  95. Pehrson JR, Costanzi C, Dharia C. Developmental and tissue expression patterns of histone macroH2A1 subtypes. J Cell Biochem 1997; 65:107–13.[CrossRef][Web of Science][Medline]
  96. Chakravarthy S, Gundimella SK, Caron C, et al. Structural characterization of the histone variant macroH2A. Mol Cell Biol 2005; 25:7616–24.[Abstract/Free Full Text]
  97. Angelov D, Molla A, Perche PY, et al. The histone variant macroH2A interferes with transcription factor binding and SWI/SNF nucleosome remodeling. Molecular Cell 2003; 11:1033–41.[CrossRef][Web of Science][Medline]
  98. Malik HS, Henikoff S. Phylogenomics of the nucleosome. Nature Structural Biology 2003; 10:882–91.[CrossRef][Web of Science][Medline]
  99. Wu RS, Tsai S, Bonner WM. Changes in histone H3 composition and synthesis pattern during lymphocyte activation. Biochem 1983; 22:3868–73.[CrossRef][Medline]
  100. Zweidler A. Histone Genes: Structure, Organization and Regulation. New York: Wiley and Sons 1984.
  101. Mito Y, Henikoff JG, Henikoff S. Genome-scale profiling of histone H3.3 replacement patterns. Nat Genet 2005; 37:1090–7.[CrossRef][Web of Science][Medline]
  102. Schwartz BE, Ahmad K. Transcriptional activation triggers deposition and removal of the histone variant H3.3. Genes Dev 2005; 19:804–14.[Abstract/Free Full Text]
  103. Hennig W. Chromosomal proteins in the spermatogenesis of Drosophila. Chromosoma 2003; 111:489–94.[Web of Science][Medline]
  104. Tagami H, Ray-Gallet D, Almouzni G, et al. Histone H3.1 and H3.3 complexes mediate nucleosome assembly pathways dependent or independent of DNA synthesis. Cell 2004; 116:51–61.[CrossRef][Web of Science][Medline]
  105. Loppin B, Bonnefoy E, Anselme C, et al. The histone H3.3 chaperone HIRA is essential for chromatin assembly in the male pronucleus. Nature 2005; 437:1386–90.[CrossRef][Medline]
  106. Palmer DK, O'Day K, Wener MH, et al. A 17-kD centromere protein (CENP-A) copurifies with nucleosome core particles and with histones. J Cell Biol 1987; 104:805–15.[Abstract/Free Full Text]
  107. Smith MM. Centromeres and variant histones: what, where, when and why? Curr Opin Cell Biol 2002; 14:279–85.[CrossRef][Web of Science][Medline]
  108. Henikoff S, Dalal Y. Centromeric chromatin: what makes it unique? Curr Opin Genet Dev 2005; 15:177–84.[CrossRef][Web of Science][Medline]
  109. Fukagawa T. Centromere DNA, proteins and kinetochore assembly in vertebrate cells. Chromosome Res 2004; 12:557–67.[CrossRef][Web of Science][Medline]
  110. Sullivan KF. A solid foundation: functional specialization of centromeric chromatin. Curr Opin Genet Dev 2001; 11:182–8.[CrossRef][Web of Science][Medline]
  111. Howman EV, Fowler KJ, Newson AJ, et al. Early disruption of centromeric chromatin organization in centromere protein A (Cenpa) null mice. Proc Natl Acad Sci USA 2000; 97:1148–53.[Abstract/Free Full Text]
  112. Cooper JL, Henikoff S. Adaptive evolution of the histone fold domain in centromeric histones. Mol Biol Evol 2004; 21:1712–8.[Abstract/Free Full Text]
  113. Tyler-Smith C, Floridia G. Many paths to the top of the mountain: diverse evolutionary solutions to centromere structure. Cell 2000; 102:5–8.[CrossRef][Web of Science][Medline]
  114. Sullivan KF, Hechenberger M, Masri K. Human CENP-A contains a histone H3 related histone fold domain that is required for targeting to the centromere. J Cell Biol 1994; 127:581–92.[Abstract/Free Full Text]
  115. Chen Y, Baker RE, Keith KC, et al. The N terminus of the centromere H3-like protein Cse4p performs an essential function distinct from that of the histone fold domain. Mol Cell Biol 2000; 20:7037–48.[Abstract/Free Full Text]
  116. Yoda K, Ando S, Morishita S, et al. Human centromere protein A (CENP-A) can replace histone H3 in nucleosome reconstitution in vitro. Proc Natl Acad Sci USA 2000; 97:7266–271.[Abstract/Free Full Text]
  117. Black BE, Foltz DR, Chakravarthy S, et al. Structural determinants for generating centromeric chromatin. Nature 2004; 430:578–82.[CrossRef][Medline]
  118. Leuba SH, Bustamante C, van Holde K, et al. Linker histone tails and N-tails of histone H3 are redundant: scanning force microscopy studies of reconstituted fibres. Biophys J 1998; 74:2830–9.[Web of Science][Medline]
  119. Marion C, Roux B, Coulet PR. Role of histones H1 and H3 in the maintenance of chromatin in a compact conformation. Study with an immobilized enzyme. FEBS Lett 1983; 157:317–21.[CrossRef][Web of Science][Medline]
  120. Malik HS, Vermaak D, Henikoff S. Recurrent evolution of DNA-binding motifs in the Drosophila centromeric histone. Proc Natl Acad Sci USA 2002; 99:1449–54.[Abstract/Free Full Text]
  121. Gilbert N, Allan J. Distinctive higher-order chromatin structure at mammalian centromeres. Proc Natl Acad Sci USA 2001; 98:11949–54.[Abstract/Free Full Text]
  122. Ando S, Yang H, Nozaki N, et al. CENP-A, -B, and -C chromatin complex that contains the I-type alpha-satellite array constitutes the prekinetochore in HeLa cells. Mol Cell Biol 2002; 22:2229–41.[Abstract/Free Full Text]
  123. Amor DJ, Kalitsis P, Sumer H, et al. Building the centromere: from foundation proteins to 3D organization. Trends Cell Biol 2004; 14:359–68.[CrossRef][Web of Science][Medline]
  124. Pusarla RH, Bhargava P. Histones in functional diversification. FEBS J 2005; 272:5149–68.[CrossRef][Medline]
  125. Zheng C, Hayes JJ. Intra- and inter-nucleosomal protein-DNA interactions of the core histone tail domains in a model system. J Biol Chem 2003; 278:24217–24.[Abstract/Free Full Text]
  126. Hamiche A, Kang JG, Dennis C, et al. Histone tails modulate nucleosome mobility and regulate ATP-dependent nucleosome sliding by NURF. Proc Natl Acad Sci USA 2001; 98:14316–21.[Abstract/Free Full Text]
  127. Sivolob A, Lavelle C, Prunell A. Sequence-dependent nucleosome structural and dynamic polymorphism. Potential involvement of histone H2B N-terminal tail proximal domain. J Mol Biol 2003; 326:49–63.[CrossRef][Web of Science][Medline]
  128. Poccia D. Male Germ Line Specific Histones of Sea Urchins and Sea stars. In Jamieson BGM, Ausió J, Justine JL (Eds.). Advances in Spermatozoal Phylogeny and Taxonomy. Mémoires du Muséum National d'Histoire Naturelle, Paris, France 1995 vol; 166: pp. 475–89.
  129. Ueda K, Tanaka I. The appearance of male gamete-specific histones gH2B and gH3 during pollen development in Lilium longiflorum. Dev Biol 1995; 169:210–7.[CrossRef][Web of Science][Medline]
  130. Ueda K, Kinoshita Y, Xu ZJ, et al. Unusual core histones specifically expressed in male gametic cells of Lilium longiflorum. Chromosoma 2000; 108:491–500.[CrossRef][Web of Science][Medline]
  131. Shires A, Carpenter MP, Chalkley R. A cysteine-containing H2B-like histone found in mature mammalian testis. J Biol Chem 1976; 251:4155–8.[Free Full Text]
  132. Zalensky AO, Siino JS, Gineitis AA, et al. Human testis/sperm-specific histone H2B (hTSH2B). Molecular cloning and characterization. J Biol Chem 2002; 277:43474–80.[Abstract/Free Full Text]
  133. Churikov D, Siino J, Svetlova M, et al. Novel human testis-specific histone H2B encoded by the interrupted gene on the X chromosome. Genomics 2004; 84:745–56.[CrossRef][Web of Science][Medline]
  134. Suzuki M. SPKK, a new nucleic acid-binding unit of protein found in histone. Embo J 1989; 8:797–804.[Web of Science][Medline]
  135. Bavykin SG, Usachenko SI, Zalensky AO, et al. Structure of nucleosomes and organization of internucleosomal DNA in chromatin. J Mol Biol 1990; 212:495–11.[CrossRef][Web of Science][Medline]
  136. Li A, Maffey AH, Abbott WD, et al. Characterization of nucleosomes consisting of the human testis/sperm-specific histone H2B variant (hTSH2B). Biochem 2005; 44:2529–35.[CrossRef][Medline]
  137. Lowell JE, Kaiser F, Janzen JC, et al. Histone H2A.Z dimerizes with a novel variant H2B and is enriched at repetitive DNA in Trypanosome brucei. J Cell Sci 2005; 118:5721–30.[Abstract/Free Full Text]
  138. Wu C, Travers A. Relative affinities of DNA sequences for the histone octamer depend strongly upon both the temperature and octamer concentration. Biochem 2005; 44:(3)14329–4.[CrossRef][Medline]
  139. Ausio J, Seger D, Eisenberg H. Nucleosome core particle stability and conformational change. Effect of temperature, particle and NaCl concentrations, and crosslinking of histone H3 sulfhydryl groups. J Mol Biol 1984; 176:77–104.[CrossRef][Web of Science][Medline]
  140. Annunziato AT. Split decision: what happens to nucleosomes during DNA replication? J Biol Chem 2005; 280:12065–8.[Free Full Text]
  141. van Holde KE, Lohr DE, Robert C. What happens to nucleosomes during transcription? J Biol Chem 1992; 267:2837–40.[Free Full Text]
  142. Shen X, Mizuguchi G, Hamiche A, et al. A chromatin remodelling complex involved in transcription and DNA processing. Nature 2000; 406:541–4.[CrossRef][Medline]
  143. Fan HY, Narlikar GJ, Kingston RE. Noncovalent modification of chromatin: different remodeled products with different ATPase domains. Cold Spring Harb Symp Quant Biol 2004; 69:183–92.[CrossRef][Web of Science][Medline]
  144. Lusser A, Kadonaga JT. Chromatin remodeling by ATP-dependent molecular machines. Bioessays 2003; 25:1192–200.[CrossRef][Web of Science][Medline]
  145. Loyola A, Almouzni G. Histone chaperones, a supporting role in the limelight. Biochim Biophys Acta 2004; 1677:3–11.[Medline]
  146. Prado A, Ramos I, Frehlick LJ, et al. Nucleoplasmin: a nuclear chaperone. Biochem Cell Biol 2004; 82:437–45.[CrossRef][Web of Science][Medline]
  147. Ehrenhofer-Murray AE. Chromatin dynamics at DNA replication, transcription and repair. Eur J Biochem 2004; 271:2335–49.[Web of Science][Medline]
  148. Sims RJ 3rd, Belotserkovskaya R, Reinberg D. Elongation by RNA polymerase II: the short and long of it. Genes Dev 2004; 18:2437–68.[Abstract/Free Full Text]
  149. Fan JY, Gordon F, Luger K, et al. The essential histone variant H2A.Z regulates the equilibrium between different chromatin conformational states. Nat Struct Biol 2002; 9:172–6.[Web of Science][Medline]
  150. Schalch T, Duda S, Sargent DF, et al. X-ray structure of a tetranucleosome and its implications for the chromatin fibre. Nature 2005; 436:138–41.[CrossRef][Medline]
  151. Ramaswamy A, Bahar I, Ioshikhes I. Structural dynamics of nucleosome core particle: comparison with nucleosomes containing histone variants. Proteins 2005; 58:683–96.[CrossRef][Web of Science][Medline]
  152. Marino-Ramirez L, Kann MG, Shoemaker BA, et al. Histone structure and nucleosome stability. Expert Rev Proteomics 2005; 2:719–29.[CrossRef][Medline]
  153. Happel N, Schulze E, Doenecke D. Characterisation of human histone H1x. Biol Chem 2005; 386:541–51.[CrossRef][Web of Science][Medline]
  154. Albig W, Meergans T, Doenecke D. Characterization of the H1.5 gene completes the set of human H1 subtype genes. Gene 1997; 184:141–8.[CrossRef][Web of Science][Medline]
  155. Parseghian MH, Henschen AH, Krieglstein KG, et al. A proposal for a coherent mammalian histone H1 nomenclature correlated with amino acid sequences. Protein Sci 1994; 3:575–87.[Web of Science][Medline]
  156. Seyedin SM, Kistler WS. Isolation and characterization of rat testis H1t. An H1 histone variant associated with spermatogenesis. J Biol Chem 1980; 255:5949–54.[Abstract/Free Full Text]
  157. Martianov I, Brancorsini S, Catena R, et al. Polar nuclear localization of H1T2, a histone H1 variant, required for spermatid elongation and DNA condensation during spermiogenesis. Proc Natl Acad Sci USA 2005; 102:2808–13.[Abstract/Free Full Text]
  158. Yan W, Ma L, Burns KH, et al. HILS1 is a spermatid-specific linker histone H1-like protein implicated in chromatin remodeling during mammalian spermiogenesis. Proc Natl Acad Sci USA 2003; 100:10546–51.[Abstract/Free Full Text]
  159. Iguchi N, Tanaka H, Yomogida K, et al. Isolation and characterization of a novel cDNA encoding a DNA-binding protein (Hils1) specifically expressed in testicular haploid germ cells. Int J Androl 2003; 26:354–65.[CrossRef][Web of Science][Medline]
  160. Tanaka M, Kihara M, Meczekalski B, et al. H1oo: a pre-embryonic H1 linker histone in search of a function. Mol Cell Endocrinol 2003; 202:5–9.[Web of Science][Medline]
  161. Eirin-Lopez JM, Frehlick LJ, Ausio J. Protamines, in the footsteps of linker histone evolution. J Biol Chem 2005; 281:1–4.[Medline]
  162. Khochbin S. Histone H1 diversity: bridging regulatory signals to linker histone function. Gene 2001; 271:1–12.[CrossRef][Web of Science][Medline]
  163. Kinkade JM Jr, Cole RD. The resolution of four lysine-rich histones derived from calf thymus. J Biol Chem 1966; 241:5790–7.[Abstract/Free Full Text]
  164. Kinkade JM Jr, Cole RD. A structural comparison of different lysine-rich histones of calf thymus. J Biol Chem 1966; 241:5798–805.[Abstract/Free Full Text]
  165. Cole RD. Microheterogeneity in H1 histones and its consequences. Int J Pept and Protein Res 1987; 30:433–49.
  166. Cole RD. A minireview of microheterogeneity in H1 histone and its possible significance. Anal Biochem 1984; 136:24–30.[CrossRef][Web of Science][Medline]
  167. Parseghian MH, Hamkalo BA. A compendium of the histone H1 family of somatic subtypes: an elusive cast of characters and their characteristics. Biochem Cell Biol 2001; 79:289–304.[CrossRef][Web of Science][Medline]
  168. Brown DT. Histone H1 and the dynamic regulation of chromatin function. Biochem Cell Biol 2003; 81:221–7.[CrossRef][Web of Science][Medline]
  169. Parseghian MH, Newcomb RL, Hamkalo BA. Distribution of somatic H1 subtypes is non-random on active vs. inactive chromatin II: distribution in human adult fibroblasts. J Cell Biochem 2001; 83:643–59.[CrossRef][Web of Science][Medline]
  170. Alami R, Fan Y, Pack S, et al. Mammalian linker-histone subtypes differentially affect gene expression in vivo. Proc Natl Acad Sci USA 2003; 100:5920–5.[Abstract/Free Full Text]
  171. Kaludov NK, Pabon-Pena L, Seavy M, et al. A mouse histone H1 variant, H1b, binds preferentially to a regulatory sequence within a mouse H3.2 replication-dependent histone gene. J Biol Chem 1997; 272:15120–7.[Abstract/Free Full Text]
  172. Konishi A, Shimizu S, Hirota J, et al. Involvement of histone H1.2 in apoptosis induced by DNA double-strand breaks. Cell 2003; 114:673–88.[CrossRef][Web of Science][Medline]
  173. Lee H, Habas R, Abate-Shen C. MSX1 cooperates with histone H1b for inhibition of transcription and myogenesis. Science 2004; 304:1675–8.[Abstract/Free Full Text]
  174. Eirin-Lopez JM, Gonzalez-Tizon AM, Martinez A, et al. Birth-and-death evolution with strong purifying selection in the histone H1 multigene family and the origin of orphon H1 genes. Mol Biol Evol 2004; 21:1992–2003.[Abstract/Free Full Text]
  175. Eirin-Lopez JM, Ruiz MF, Gonzalez-Tizon AM, et al. Common evolutionary origin and birth-and-death process in the replication-independent histone H1 isoforms from vertebrate and invertebrate genomes. J Mol Evol 2005; 61:398–407.[CrossRef][Web of Science][Medline]
  176. Caron F, Thomas JO. Exchange of histone H1 between segments of chromatin. J Mol Biol 1981; 146:513–37.[CrossRef][Web of Science][Medline]
  177. Bustin M, Catez F, Lim JH. The dynamics of histone H1 function in chromatin. Mol Cell 2005; 17:617–20.[CrossRef][Web of Science][Medline]
  178. Th’ng JP, Sung R, Ye M, et al. H1 family histones in the nucleus. Control of binding and localization by the C-terminal domain. J Biol Chem 2005; 280:27809–14.[Abstract/Free Full Text]
  179. Hendzel MJ, Lever MA, Crawford E, et al. The C-terminal domain is the primary determinant of histone H1 binding to chromatin in vivo. J Biol Chem 2004; 279:20028–34.[Abstract/Free Full Text]
  180. Luger K, Hansen JC. Nucleosome and chromatin fiber dynamics. Curr Opin Struct Biol 2005; 15:188–96.[CrossRef][Web of Science][Medline]
  181. Hansen JC, Lu X, Ross ED, et al. Intrinsic protein disorder, amino acid composition, and the histone terminal domains. J Biol Chem 2006; 281:1853–6.
  182. Chakravarthy S, Bao Y, Roberts VA, et al. Structural characterization of histone H2A variants. Cold Spring Harb Symp Quant Biol 2004; 69:227–34.[CrossRef][Web of Science][Medline]
  183. Dong F, van_Holde KE. Nucleosome positioning is determined by the (H3-H4)2 tetramer. Proc Natl Acad Sci USA 1991; 88:10596–600.[Abstract/Free Full Text]
  184. Seyedin SM, Kistler WS. H1 histone subfractions of mammalian testes. 1. Organ specificity in the rat. Biochem 1979; 18:1371–5.[CrossRef][Medline]

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