Alternative titles; symbols
HGNC Approved Gene Symbol: H4C1
Cytogenetic location: 6p22.2 Genomic coordinates (GRCh38) : 6:26,021,649-26,022,050 (from NCBI)
The nucleosome is the basic repeat unit of eukaryotic chromatin. The nucleosome core particle consists of an octamer formed by 2 each of the core histones H2A (see 613499), H2B (see 609904), H3 (see 602810), and H4, around which DNA is wrapped. A fifth histone, histone H1 (see 142709), is bound to the linker DNA between nucleosomes and is important for the higher order structure of chromatin. HIST1H4A is a core histone H4 (summary by Marzluff et al. (2002) and Foster and Downs (2005)).
All core histones, including H4 histones, contain a histone fold domain, which is central to the nucleosome core structure, and a flexible N-terminal domain that protrudes from the nucleosome core particle. Like other histones, H4 histones can be subgrouped according to their temporal expression. Replication-dependent histones, such as HIST1H4A through HIST1H4L (602831) and HIST2H4A (142750) are mainly expressed during S phase. In contrast, replication-independent histones, or replacement variant histones, can be expressed throughout the cell cycle. Most replication-dependent H4 histone genes, as well as other core histone genes, are located within histone gene cluster-1 (HIST1) on chromosome 6p22-p21. Two other histone gene clusters, HIST2 and HIST3, are located on chromosomes 1q21 and 1q42, respectively. HIST2 contains 1 replication-dependent H4 gene, HIST2H4A, and there are no H4 genes in HIST3. An additional H4 gene, HIST4H4 (615069), is located on chromosome 12p13.1. In mouse, the Hist1, Hist2, and Hist3 gene clusters are located on chromosomes 13A2-A3, 3F1-F2, and 11B2, respectively. All replication-dependent histone genes are intronless, and they encode mRNAs that lack a poly(A) tail, ending instead in a conserved stem-loop sequence. Unlike replication-dependent histone genes, replication-independent histone genes are solitary genes that are located on chromosomes apart from any other H1 or core histone genes. Some replication-independent histone genes contain introns and encode mRNAs with poly(A) tails. All human and mouse H4 histone genes encode the same protein (summary by Marzluff et al. (2002) and Foster and Downs (2005)).
By genomic sequence analysis, Marzluff et al. (2002) identified the mouse and human HIST1H4A genes. All mouse and human H4 genes, including HIST1H4A, encode the same protein.
By analysis of a YAC contig from chromosome 6p21.3, Albig et al. (1997) characterized a cluster of 35 histone genes, including H4/a.
By genomic sequence analysis, Marzluff et al. (2002) determined that the HIST1 cluster on chromosome 6p22-p21 contains 55 histone genes, including 12 H4 genes. The HIST1H4A gene is the most telomeric H4 gene within the HIST1 cluster. The HIST1 cluster spans over 2 Mb and includes 2 large gaps (over 250 kb each) where there are no histone genes, but many other genes. The organization of histone genes in the mouse Hist1 cluster on chromosome 13A2-A3 is essentially identical to that in human HIST1. The HIST2 cluster on chromosome 1q21 contains 6 histone genes, including 1 H4 gene (HIST2H4A; 142750), and the HIST3 cluster on chromosome 1q42 contains 3 histone genes, but no H4 genes. Hist2 and Hist3 are located on mouse chromosomes 3F1-F2 and 11B2, respectively. An additional H4 gene, HIST4H4 (615069), is located on human chromosome 12p13.1 and mouse chromosome 6G1.
H4 Histone Family
As reviewed by Felsenfeld (1992), detailed biochemical definition of the protein complexes that regulate gene transcription led to reemergence of questions concerning the role of histones. He reviewed evidence suggesting that transcriptional activation requires that transcription factors successfully compete with histones for binding to promoters.
CpG island hypermethylation and global genomic hypomethylation are common epigenetic features of cancer cells. Fraga et al. (2005) characterized posttranslational modifications to histone H4 in a comprehensive panel of normal tissues, cancer cell lines, and primary tumors. They found that cancer cells had a loss of monoacetylated and trimethylated forms of histone H4. These changes appeared early and accumulated during the tumorigenic process, as they showed in a mouse model of multistage skin carcinogenesis. The losses occurred predominantly at the acetylated lys16 and trimethylated lys20 residues of histone H4 and were associated with the hypomethylation of DNA repetitive sequences, a well-known characteristic of cancer cells. Fraga et al. (2005) suggested that the global loss of monoacetylation and trimethylation of histone H4 is a common hallmark of human tumor cells.
Wang et al. (2001) reported the purification, molecular identification, and functional characterization of a histone H4-specific methyltransferase, PRMT1 (602950), a protein arginine methyltransferase. PRMT1 specifically methylates arginine-3 of histone H4 in vitro and in vivo. Methylation of arg3 by PRMT1 facilitates subsequent acetylation of H4 tails by p300 (602700). However, acetylation of H4 inhibits its methylation by PRMT1. Most important, a mutation in the S-adenosyl-L-methionine-binding site of PRMT1 substantially crippled its nuclear receptor coactivator activity. Wang et al. (2001) concluded that their findings reveal arg3 of H4 as a novel methylation site by PRMT1 and indicate that arg3 methylation plays an important role in transcriptional regulation.
Agalioti et al. (2002) found that only a small subset of lysines in histones H3 (see 602810) and H4 are acetylated in vivo by the GCN5 acetyltransferase (see 602301) during activation of the interferon-beta gene (IFNB; 147640). Reconstitution of recombinant nucleosomes bearing mutations in these lysine residues revealed the cascade of gene activation via a point-by-point interpretation of the histone code through the ordered recruitment of bromodomain-containing transcription complexes. Acetylation of histone H4 lys8 mediates recruitment of the SWI/SNF complex (see 603111), whereas acetylation of lys9 and lys14 in histone H3 is critical for the recruitment of TFIID (see 313650). Thus, the information contained in the DNA address of the enhancer is transferred to the histone N termini by generating novel adhesive surfaces required for the recruitment of transcription complexes.
Using deuterium exchange/mass spectrometry coupled with hydrodynamic measures, Black et al. (2004) demonstrated that CENPA (117139) and histone H4 form subnucleosomal tetramers that are more compact and conformationally more rigid than the corresponding tetramers of histones H3 and H4. Substitution into histone H3 of the domain of CENPA responsible for compaction was sufficient to direct it to centromeres. Thus, Black et al. (2004) concluded that the centromere-targeting domain of CENPA confers a unique structural rigidity to the nucleosomes into which it assembles, and is likely to have a role in maintaining centromere identity.
Acetylation of histone H4 on lysine-16 (H4-K16Ac) is a prevalent and reversible posttranslational chromatin modification in eukaryotes. To characterize the structural and functional role of this mark, Shogren-Knaak et al. (2006) used a native chemical ligation strategy to generate histone H4 that was homogeneously acetylated at K16. The incorporation of this modified histone into nucleosomal arrays inhibited the formation of compact 30-nanometer-like fibers and impeded the ability of chromatin to form cross-fiber interactions. H4-K16Ac also inhibited the ability of the adenosine triphosphate-utilizing chromatin assembly and remodeling enzyme ACF to mobilize a mononucleosome, indicating that this single histone modification modulates both higher order chromatin structure and functional interactions between a nonhistone protein and the chromatin fiber.
In a screen for endogenous tumor-associated T-cell responses in a primary mouse model of prostatic adenocarcinoma, Savage et al. (2008) identified a naturally arising CD8+ T cell response that is reactive to a peptide derived from histone H4. Despite the ubiquitous nature of histones, T cell recognition of histone H4 peptide was specifically associated with the presence of prostate cancer in these mice. Thus, Savage et al. (2008) concluded that the repertoire of antigens recognized by tumor-infiltrating T cells is broader than previously thought and includes peptides derived from ubiquitous self antigens that are normally sequestered from immune detection.
Dang et al. (2009) reported an age-associated decrease in yeast Sir2 (see SIRT1, 604479) protein abundance accompanied by an increase in histone H4 lysine-16 acetylation and loss of histones at specific subtelomeric regions in replicatively old yeast cells, which results in compromised transcriptional silencing at these loci. Antagonizing activities of Sir2 and Sas2, a histone acetyltransferase, regulate the replicative life span through histone H4 lys16 at subtelomeric regions. Dang et al. (2009) concluded that this pathway, distinct from existing aging models for yeast, may represent an evolutionarily conserved function of sirtuins in regulation of replicative aging by maintenance of intact telomeric chromatin.
Xu et al. (2010) reported that significant amounts of histone H3.3 (see 601128)-H4 tetramers split in vivo, whereas most H3.1 (see 602810)-H4 tetramers remain intact during mitotic division. Inhibiting DNA replication-dependent deposition greatly reduced the level of splitting events, which suggested that (i) the replication-independent H3.3 deposition pathway proceeds largely by cooperatively incorporating 2 new H3.3-H4 dimers, and (ii) the majority of splitting events occurred during replication-dependent deposition. Xu et al. (2010) concluded that 'silent' histone modifications within large heterochromatic regions are maintained by copying modifications from neighboring preexisting histones without the need for H3-H4 splitting events.
Qi et al. (2010) provided multiple lines of evidence establishing PHF8 (300560) as the first monomethyl histone H4 lysine-20 (H4K20me1) demethylase, with additional activities towards histone H3K9me1 and me2. PHF8 is located around the transcriptional start sites of approximately 7,000 RefSeq genes and in gene bodies and intergenic regions. PHF8 depletion resulted in upregulation of H4K20me1 and H3K9me1 at the transcriptional start site and H3K9me2 in the nontranscriptional start sites, respectively, demonstrating differential substrate specificities at different target locations. PHF8 positively regulates gene expression, which is dependent on its H3K4me3-binding PHD and catalytic domains. Importantly, patient mutations significantly compromised PHF8 catalytic function. PHF8 regulates cell survival in the zebrafish brain and jaw development, thus providing a potentially relevant biologic context for understanding the clinical symptoms associated with PHF8 patients. Lastly, genetic and molecular evidence supported a model whereby PHF8 regulates zebrafish neuronal cell survival and jaw development in part by directly regulating the expression of the homeodomain transcription factor MSX1/MSXB (605558), which functions downstream of multiple signaling and developmental pathways.
Liu et al. (2010) reported that PHF8, while using multiple substrates, including H3K9me1/2 and H3K27me2, also functions as an H4K20me1 demethylase. PHF8 is recruited to promoters by its PHD domain based on interaction with H3K4me2/3 and controls G1-S transition in conjunction with E2F1, HCF1 (300019), and SET1A (611052), at least in part, by removing the repressive H4K20me1 mark from a subset of E2F1-regulated gene promoters. Phosphorylation-dependent PHF8 dismissal from chromatin in prophase is apparently required for the accumulation of H4K20me1 during early mitosis, which might represent a component of the condensin II loading process. Accordingly, the HEAT repeat clusters in 2 non-structural maintenance of chromosomes (SMC) condensin II subunits, NCAPD3 (609276) and NCAPG2 (608532), are capable of recognizing H4K20me1, and ChIP-Seq analysis demonstrated a significant overlap of condensin II and H4K20me1 sites in mitotic HeLa cells. Thus, Liu et al. (2010) concluded that the identification and characterization of an H4K20me1 demethylase, PHF8, has revealed an intimate link between this enzyme and 2 distinct events in cell cycle progression.
Fullgrabe et al. (2013) reported that induction of autophagy is coupled to reduction of histone H4 lysine-16 acetylation (H4K16ac) through downregulation of the histone acetyltransferase MOF (MYST1; 609912), and demonstrated that this histone modification regulates the outcome of autophagy. At a genomewide level, Fullgrabe et al. (2013) found that H4K16 deacetylation is associated predominantly with the downregulation of autophagy-related genes. Antagonizing H4K16ac downregulation upon autophagy induction results in the promotion of cell death. Fullgrabe et al. (2013) concluded that their findings established that alteration in a specific histone posttranslational modification during autophagy affects the transcriptional regulation of autophagy-related genes and initiates a regulatory feedback loop, which serves as a key determinant of survival versus death responses upon autophagy induction.
Saredi et al. (2016) found that new histones incorporated during DNA replication provided a signature of postreplicative chromatin that was read by the TONSL (604546)-MMS22L (615614) homologous recombination complex. The ankyrin repeat domain (ARD) of TONSL read histone H4 tails that were unmethylated at lys20, a feature specific to new histones incorporated during DNA replication. TONSL-MMS22L bound new histones H3-H4 before and after incorporation into nucleosomes and remained on replicated chromatin until late G2/M. Recognition of unmethylated H4 lys20 was required for TONSL-MMS22L binding to chromatin and accumulation at challenged replication forks and DNA lesions. Mutations in the ARD of TONSL were toxic and compromised genome stability, cell viability, and resistance to replication stress.
Crystal Structure
Sekulic et al. (2010) reported the crystal structure of a subnucleosomal heterotetramer, (CENP-A-H4)2 (CENP-A, 117139, in complex with histone H4), that reveals 3 distinguishing properties encoded by the residues that comprise the CENP-A targeting domain (CATD): (1) a CENP-A-CENP-A interface that is substantially rotated relative to the H3-H3 interface; (2) a protruding loop L1 of the opposite charge as that on H3; and (3) strong hydrophobic contacts that rigidify the CENP-A-H4 interface. Residues involved in the CENP-A-CENP-A rotation are required for efficient incorporation into centromeric chromatin, indicating specificity for an unconventional nucleosome shape. DNA topologic analysis indicated that CENP-A-containing nucleosomes are octameric with conventional left-handed DNA wrapping. Sekulic et al. (2010) concluded that CENP-A marks centromere location by restructuring the nucleosome from within its folded histone core.
Elsasser et al. (2012) reported the crystal structures of the DAXX (603186) histone-binding domain with a histone H3.3-H4 dimer, including mutants within DAXX and H3.3, together with in vitro and in vivo functional studies that elucidated the principles underlying H3.3 recognition specificity. Occupying 40% of the histone surface-accessible area, DAXX wraps around the H3.3-H4 dimer, with complex formation accompanied by structural transitions in the H3.3-H4 histone fold. DAXX uses an extended alpha-helical conformation to compete with major interhistone, DNA, and ASF1 interaction sites. Elsasser et al. (2012) concluded that their structural studies identified recognition elements that read out H3.3-specific residues, and functional studies addressed the contribution of gly90 in H3.3 and glu225 in DAXX to chaperone-mediated H3.3 variant recognition specificity.
Histone IV genes are highly conserved across evolution. Delange and Smith (1971) noted that, in their 110 amino acids, histone IV genes of cattle and garden peas differ by only 2 residues.
Heintz et al. (1981) concluded that the human histone genes are clustered in the genome but are not arranged into recognizable repeating units. The lack of organization of the human histone genes (as contrasted with those of invertebrates or of Xenopus laevis) may reflect the diminished requirement for rapid synthesis of large quantities of histone proteins during early mammalian development.
Kedes and Maxson (1981) found that the histone genes in man, mouse, chicken, and toad show a dispersed topology; they are scattered and separated by long stretches of nonhistone DNA. In an article subtitled 'Paradigm Lost,' the authors referred to 'this newly discovered diaspora.'
Marzluff et al. (2002) provided a nomenclature for replication-dependent histone genes located within the HIST1, HIST2, and HIST3 clusters. The symbols for these genes all begin with HIST1, HIST2, or HIST3 according to which cluster they are located in. The H2A, H2B, H3, and H4 genes were named systematically according to their location within the HIST1, HIST2, and HIST3 clusters. For example, HIST1H4A is the most telomeric H4 gene within HIST1, and HIST1H4L (602831) is the most centromeric. In contrast, the H1 genes, all of which are located within HIST1, were named according to their mouse homologs. Thus, HIST1H1A (142709) is homologous to mouse H1a, HIST1H1B (142711) is homologous to mouse H1b, and so on.
Szabo et al. (1978) presented nucleic acid hybridization data indicating that chromosome 7 carries gene(s) coding for histone H4 protein. Steffensen (1979) presented evidence that all 5 histone genes in man are clustered at 7q2. Yunis and Chandler (1979) located the histone genes to bands 7q32-36 and the homologous chromosome segments in chimpanzee, gorilla, and orangutan.
A clone containing a human histone gene cluster in the order H3-H4-H1-H2A-H2B was isolated by Clark et al. (1981), as cited by Hentschel and Birnstiel (1981). Sierra et al. (1982) likewise found an arrangement of the histone genes different from that in the sea urchin and Drosophila.
Carozzi et al. (1984) isolated an H1 histone gene from a 15-kb human DNA genomic sequence. The presence of H2A, H2B, H3 and H4 genes in this same 15-kb fragment demonstrated that these genes are clustered.
By study of mouse-human cell hybrids and by in situ hybridization, Green et al. (1984) showed that H3 and H4 histone genes are on 1q, probably 1q21. From in situ hybridization, Tripputi et al. (1986) concluded that histone genes map to at least 3 different chromosomes: 1, 6, and 12. Some may be nonexpressed pseudogenes. They commented that the number of histone genes is between 100 and 200. The histones have the distinction of being the only proteins coded by repetitive DNA. Tanguay et al. (1987) reported in situ hybridization data corroborating those of Tripputi et al. (1986), using a heterologous probe containing the 5 histone genes of Drosophila. They found that the main concentrations of grains were at 6p12-q21, 12q11-q22, and 1cen-q25. Allen et al. (1989) reported the conflicting assignment of histones 3 and 4 to human chromosome 6.
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