Search tips
Search criteria 


Logo of narLink to Publisher's site
Nucleic Acids Res. 2013 November; 41(21): 9593–9609.
Published online 2013 August 14. doi:  10.1093/nar/gkt700
PMCID: PMC3834806

H1 histones: current perspectives and challenges


H1 and related linker histones are important both for maintenance of higher-order chromatin structure and for the regulation of gene expression. The biology of the linker histones is complex, as they are evolutionarily variable, exist in multiple isoforms and undergo a large variety of posttranslational modifications in their long, unstructured, NH2- and COOH-terminal tails. We review recent progress in understanding the structure, genetics and posttranslational modifications of linker histones, with an emphasis on the dynamic interactions of these proteins with DNA and transcriptional regulators. We also discuss various experimental challenges to the study of H1 and related proteins, including limitations of immunological reagents and practical difficulties in the analysis of posttranslational modifications by mass spectrometry.


Histones are evolutionarily conserved proteins responsible for condensation, organization and regulation of the DNA within the nucleus of all eukaryotes. The basic structural element of DNA compaction, the nucleosome core particle, is made up of superhelical DNA wrapped about a protein octamer composed of two copies of each core histone H2A, H2B, H3 and H4 (1–4). Structurally, each core histone has a long central helix with a helix-strand-helix motif on each end forming what is termed the histone fold (5). Hydrophobic interactions between two core histone monomers form heterodimers in a head-to-tail configuration called the handshake motif (2–7). The heterodimers of histones H3 and H4 further associate to form tetramers (5,6). The histone octamer is assembled from two H2A–H2B dimers binding opposite the H3–H4 tetramer (7). Micrococcal nuclease digestion of chromatin exposed to increasing salt concentrations shows symmetrical association of ~146 base pairs of left-handed superhelical DNA wrapped ~1.65 turns around the histone octamer forming the nucleosome core particle (5,8–12). Crystallography orients the histone octamer with the H3–H4 tetramer centered between and in direct contact with the DNA entry and exit points and the H2A–H2B tetramer centered opposite. Higher-order chromatin structures are produced through the binding of a linker histone, histone H1, to the nucleosome core particle to form the chromatosome (13–16).

Nucleosomal stabilization facilitated by the chromatosome is provided through the binding of histone H1 to the nucleosomal dyad and the linker DNA entering and exiting the core particle (16–26). Recent •OH radical footprinting experiments show that the positioning of histone H1 at the nucleosomal dyad axis protects an additional 20 base pairs of DNA, 10 base pairs from both the entering and exiting linker DNA, from micrococcal nuclease digestion (8,10,17,25,26). Additional experimental evidence illustrates the influence of histone H1 on chromatin arrangement and compaction (14,19,27–33). However, the specific folding of the 30-nm filament remains controversial and potentially variable in nature (32). In any case, recent studies suggest histone H1 binding provides stabilization and protection through the formation of a dynamic and polymorphic linker histone/linker DNA stem structure (25,26,30,32). Stem-to-stem interactions of neighboring nucleosomes are hypothesized to stabilize folding into higher-order chromatin fibers (26). No matter how the 30-nm chromatin fiber ultimately folds, the influence of histone H1 is dependent on its unique structural characteristics.


Histone H1 has a tripartite structure containing an evolutionarily conserved central globular domain with flanking variable domains. X-ray crystallography of the globular domain of the avian erythrocyte linker histone H5 (considered a member of the H1 family) shows a winged-helix motif consisting of three alpha helices with a C-terminal beta hairpin (34). An antiparallel beta sheet is formed between the C-terminal beta hairpin and a short beta strand connecting the first and second alpha helices (34). Conformational studies on the globular domain of the erythrocyte linker histone show that H5 binds asymmetrically to two DNA duplexes through two clusters of highly conserved, positively charged residues on opposite sides of the globular H5 molecule (18,34). Initial positional studies of linker histone H5 on chicken nucleosomes illustrate the globular domain is located between chromatosomal terminal DNA and DNA near the dyad axis of the nucleosome (20). However, more recent experiments using the globular domain of histone H1.5 show binding at the DNA minor groove of the nucleosomal dyad axis (25). As a result, the globular domain has been shown to mediate the protection of 20 additional base pairs of linker DNA by the chromatosome (17,25,26). Although binding of the globular domain of histone H1 can protect almost two full turns of superhelical DNA from micrococcal nuclease digestion, it is the flanking terminal regions of the linker histone that allow for the formation of higher-order chromatin structures (17).

The amino terminus of histone H1 is considered nominally unstructured, as solution and X-ray crystallographic stuctures have yet to be determined (15). Based on sequence, the N-terminus can be divided into two sub-regions (35). The extreme N-terminal sequence is enriched in hydrophobic residues, whereas a highly basic portion resides close to the globular domain (35). The basic cluster has been linked to globular domain positioning and takes on an alpha helical structure in the presence of DNA, whereas the hydrophobic region remains uncharacterized (36,37). Although the N-terminus of bovine thymus histone H1 has been shown to be non-essential for the formation of higher-order chromatin structures, deletion of the N-terminal domain (NTD) of histone H1 isoforms reduces the binding affinity for chromatin in vitro (36,38,39). Additionally, histone H1 NTD swapping experiments between mouse H1o and H1c show exchange of their chromatin binding affinities via fluorescence recovery after photobleaching analysis (40). These studies suggest the NTD of histone H1 plays a role in proper binding to the nucleosome. However, additional studies are needed to characterize the functionality of the NTD of histone H1.

Similar to the amino terminus of histone H1, the carboxy terminus lacks X-ray crystallographic resolution and is assumed to nominally be a random-coil, or intrinsically disordered, in solution (41–45). In vitro data suggest that on DNA binding at physiological salt concentrations, the C-terminal domain (CTD) of histone H1 takes on a folded conformation dominated by common secondary structural components such as alpha helices, beta sheets, loops and turns (42,45–49). Recent data presented by Fang et al. support the formation of secondary structure, as the carboxy terminus of histone H1 settles into DNA helices, allowing for the formation of the nucleosome stem structure (49). Additionally, the interaction of the CTD with linker DNA has been shown to extend beyond the initial 20 base pairs entering and exiting the nucleosome (45).

The CTD accounts for more than half the linker histone sequence, with ~40% composed of lysine, 20–35% alanine and 15% proline residues (43). Mutational studies on the CTD of histone H1 suggest two distinct functional regions for DNA binding, two 24-amino-acid lengths, facilitate chromatin condensation (44,50). It is hypothesized the remaining CTD length (~50 amino acids) is involved in protein–protein interactions (44). Support for this concept was recently shown through the binding of DNA methyltransferases (DNMT1 and DNMT3B) to the CTD of mouse histone H1 by Yang et al. (51).

The net positive charge imparted on the CTD from the high lysine content allows for regulation of higher-order chromatin structures through DNA backbone charge neutralization (52,53). This allows for low-affinity H1 binding to give rise to the formation of secondary structure in the CTD that permits high-affinity binding (17,36,38,49,50,52,54,55,). In addition to the globular domain, the secondary structure in the CTD enables the formation and stabilization of linker DNA into higher-order chromatin structures (17,25,26,36,38,54,55). The length, charge and number of posttranslational modification (PTM) sites of the C-terminal tails vary between histone H1 isoforms, suggesting that individual H1 variants may play distinct roles in the regulation of higher-order chromatin structure.


The histone H1 gene family in lower organisms is less evolutionarily conserved than that of the core histones. For example, in Saccharomyces cerevisiae, the sequence homology between Hho1, the S. cerevisiae histone H1 homolog, and Homo sapien H1 is 31% identical and 44% similar, whereas histone H4 between the species is 92% identical and 96% similar. Conversely, in higher-order organisms such as the Gallus gallus (chicken), the erythrocyte linker histone, H5, shows high sequence homology (66%) to the human histone H1.0, with the greatest sequence divergence found in the CTD (56). In addition to sequence variation, histone H1 proteins also display a range of structures. For instance, S. cerevisiae Hho1p contains two globular domains, whereas Tetrahymena completely lacks a globular domain (57,58). Eukaryotes also differ in the number of histone H1 variants present. H. sapiens and Mus musculus both have 11 distinct variants, whereas Caenorhabditis elegans has eight and Xenopus laevis has five (59). The H. sapien family of histone H1 proteins contains five somatic variants (H1.1, H1.2, H1.3, H1.4 and H1.5), which are expressed in nearly all cells (60–62). Six additional H1 variants have been identified in specific tissues, such as H1t and H1T2 in the testis, or cell types, such as H1.0 in terminally differentiated cells (56,63–71). Two types of histone H1 genes exist in human cells: replication-independent and replication-dependent genes. The replication-independent H1 genes, such as histone H1(0), exhibit a replacement phenotype (72). These replacement histone H1s are genomically isolated from other histone genes with transcription based on cellular status (72). Whereas replication-independent H1s have been observed throughout the cell cycle, the majority of the histone H1 protein is produced during S phase of the cell cycle (73–77).

In contrast to replication-independent H1 expression, the replication-dependent variants are found in a large cluster alongside many of the core histone genes located on the short area of chromosome 6 (6p21-p22) (62,78,79). These histone H1 genes, located in gene cluster HIST1, have paired expression with DNA replication and core histone mRNA expression levels, although specific H1 variants have been shown to have fluctuating expression across S phase of the cell cycle (77,80–84). The mRNA of replication-dependent H1 genes lack a poly(A) tail and introns commonly observed in other protein-coding genes (85,86). Alternatively, somatic H1 genes contain a 3′ stem-loop sequence allowing for rapid translation during DNA replication, while permitting tight regulation of gene expression after the conclusion of S phase (83,86,87). The expression patterns of individual H1 variants are essential to the functional properties of H1 in the chromatin regulatory system.

In addition to the expression of the normal somatic histone variants, several histone H1 sequence variations have been described. Initially, two sequence variants were described in K562 and Raji cells (88). In the K562 cell line, an alanine to valine substitution is observed at position 17 of histone H1.2 (H1.2A17V) (88). A histone H1.4 sequence variant was found in the Raji cell line corresponding to a lysine to arginine substitution at position 173 (H1.4K173R) (88). Finally, an alanine to threonine substitution at position 142 on histone H1.2 was described by mass spectrometry (MS) in HeLa S3 cells (89). Although identified, the function of the sequence variations remains unknown.

Overexpression of histone H1 variants shows functional differences between the isoforms. Experiments overexpressing histone H1c and H1(0) in the mouse 3T3 cell lines led to distinct phenotypes in these cells. Overexpression of H1(0) results in an increase in nucleosomal repeat length and a decline in cell cycle progression (90,91). Conversely, overexpression of murine histone H1c gives rise to an increase in or no change in transcription levels, while conferring no effect on cell cycle progression (91). These overexpression experiments show functional differences between the two variants, although additional experiments with other isoforms are still needed to further elucidate H1 function.


Histone H1 binding to chromatin has been shown to be dynamic in nature, with specific H1 variants divergent in their binding affinity for chromatin (54,55,92–94). It is thought that a high percentage of the total nuclear H1 is bound to nucleosomes at any given time; however, these interactions are individually transient (54,55). Data presented by Lever et al. demonstrate in vivo dynamics of histone H1.1 occur through soluble intermediates, giving rise to a rapid “stop-and-go” movement of H1.1 in the nucleus between random binding sites (54). Others have further demonstrated that the transient binding of H1 variants with nucleosomes is affected by the structure of the H1 variant, PTMs present on H1 and competition for chromatin binding by other nuclear factors.

Histone H1, as described earlier, has a tripartite structure. Of these, the CTD is the primary determinant of the binding dynamics of each specific variant. For example, fluorescence recovery after photobleaching experiments using NTD green fluorescent protein-tagged H1 variants (H1.0-H1.5) show that the variants with the shortest CTDs have the shortest residence times on nucleosomes (93). Additionally, Th’ng et al. show through CTD swapping between H1.1 and H1.4 or H1.5 and truncation experiments with H1.5 that the CTD determines the in vitro binding affinity for the nucleosome (93). Similarly, others have shown truncation of histone H1.1’s CTD reduces the residence time of the variant on the nucleosome ~10-fold in vivo (38,54). While CTD length clearly affects variant nucleosomal residence times, the number of phosphorylations and phosphorylation sites also play a role.

Phosphorylation of histone H1 has many distinct functions, leading to both chromatin condensation and decondensation dependent on the site of phosphorylation and cell cycle context. Histone H1 phosphorylation has been shown to progressively increase as a cell progresses from G1 to mitosis during the cell cycle (95–101). The overall importance of histone H1 phosphorylation was highlighted by several studies showing that changes in histone H1 phosphorylation can prevent entry into mitosis, thus linking histone H1 phosphorylation with the cell cycle (102,103). The phosphorylation of histone H1 during the cell cycle has been theorized to be a two-fold process (92). First, an interphase (G0–S phase) partial phosphorylation that allows for chromatin relaxation and facilitates transcriptional activation (104–107). Second, a maximal phosphorylation during mitosis (M phase) allows for chromatin condensation and separation of chromosomes into daughter cells (95–101). The partial phosphorylation observed in interphase has been shown to induce structural changes in the CTD of H1, which in turn leads to a decreased affinity of histone H1 for DNA (108). Mutational studies mimicking histone H1 phosphorylation have been shown to change the chromatin histone H1 dynamics (109). Additionally, decondensation of chromatin at DNA replication forks has been shown to be a result of histone H1 phosphorylation by cyclin-dependent kinase (CDK) 2 (110). To this end, work by Talasz et al. and Sarg et al. with histone H1.5 suggests interphase phosphorylation only occurs on serine residues at SPK(A)K sequences (H1.5S17p, H1.5S172p and H1.5S188p) (111,112). Zheng et al. have supported this argument by demonstrating H1.2 and H1.4 have serine-only phosphorylation during interphase by MS (H1.2S173p, H1.4S172p and H1.4S187p) (89). The identified interphase phosphorylation sites remained in the mitotic fraction, suggesting preferential hierarchy of phosphorylation on these H1 variants (89). Collectively these studies support the model that interphase phosphorylation on specific histone H1 variants can disrupt DNA–histone interactions, allowing for chromatin relaxation through histone H1 mobilization, and allow for competition and regulation of binding sites on DNA by other nuclear proteins (110,113).

The dynamic nature of histone H1 during interphase allows for regulation of DNA access through several mechanisms (114). First, through condensation of chromatin, histone H1 can limit access of other proteins to chromatin. Lee et al. have shown that phosphorylation of histone H1, mimicking H1 removal from chromatin and decondensation, allows for glucocorticoid-induced transcription of the mouse mammary tumor virus promoter (115). Additionally, phosphorylation of histone H1 has been shown to disrupt the interaction between itself and heterochromatin protein 1α, leading to chromatin decondensation (116). Second, histone H1-bound nucleosomes can limit access of chromatin remodeling complexes. For example, the activity of ATP-dependent SWI/SNF chromatin remodeling complexes are reduced and altered when nucleosomes are bound to H1 (117,118). Additionally, this reduction in SWI/SNF activity can be rescued by phosphorylation of histone H1, suggesting a role for histone H1 phosphorylation in chromatin remodeling (119). However, data by Maier et al. and Clausell et al. have shown that chromatin remodeling complexes can remain active even in the presence of linker histone (24,120). These data suggest specific remodeling complexes can access key nucleosomal elements without the removal of the linker histones. Next, stabilization of the nucleosomal positioning by histone H1 limits the rotational access of specific DNA sequences to transcription factors and other nuclear proteins. This principle was demonstrated by Cheung et al., who showed that estrogen receptor α-mediated transcriptional activity is repressed by H1 via decreased promoter accessibility (121). However, others have demonstrated transcriptional activation of the mouse mammary tumor virus promoter after histone H1 phosphorylation, suggesting a rescue of transcription can be achieved by histone H1 phosphorylation (115,122,123). Finally, histone H1 binding sterically inhibits access of other factors to the chromatin. Herrera et al. have demonstrated histone H1 sterically occludes histone acetyl transferase complexes from acetylating the N-terminal tail of histone H3 (124). Whereas interphase phosphorylation of histone H1 is largely involved in transcriptional regulation, mitotic phosphorylation yields a condensed chromatin state allowing for cell division.

The second phase of histone H1 phosphorylation occurs during mitosis. Similar to interphase phosphorylation, mitotic phosphorylation has been shown to be primarily a result of CDK activity at sites of S/TPXK consensus sequences, although non-CDK mitotic phosphorylations have also been identified (Table 1). First described in the 1970s, mitotic phosphorylation of histone H1 is a maximal phosphorylation resulting in the condensation of chromatin (95–101). Several studies by Deterding et al. using MS have shown reduction in variant-specific histone H1 phosphorylation in response to therapeutics (CDK inhibitors) and hormones (dexamethasone) (145–147). Furthermore, Th’ng et al. have shown through the use of the kinase inhibitor staurosporine that the hyperphosphorylation of histone H1 observed on mitotic chromatin is required to retain condensed chromatin structures (102). Additionally, they established that the inhibition of the H1 kinase by staurosporine arrests cells at the G2/M transition, preventing progression into mitosis (102). This study and others, such as those seen with the topoisomerase inhibitor VM-26, emphasize the importance of histone H1 phosphorylation in cell cycle progression (103,148,149). Collectively, these studies suggest the potential for histone H1 kinase inhibitors as cancer therapeutics.

Table 1.
Histone H1 posttranslational modifications identified by mass spectrometry

An important aspect of histone H1 dynamics that remains unresolved is the degree to which histone chaperones control the dynamics and assembly of the linker histones. Due to their exceptionally high degree of positive charge, the histone proteins can form indiscriminate and deleterious complexes with negatively charged species in the cell such as nucleic acids. A key function of the class of proteins known as histone chaperones is thought to prevent these inappropriate interactions. Although a large number of chaperones have been demonstrated to play a role in core histone transit and assembly, it is not clear whether the movements of histone H1 in the cell and its association with chromatin are mediated by other protein factors or whether they occur spontaneously (150). One potential histone H1 chaperone is the human protein nuclear autoantigenic sperm protein (NASP). NASP has been shown to be associated with both linker and core histones in the cell (151,152). In vitro, NASP is capable of binding to histone H1 with nM affinity and to transfer H1 molecules to DNA (153–155). However, a role for NASP in the cellular dynamics of histone H1 has not been directly demonstrated.


Although both interphase and mitotic phosphorylation-specific sites have been observed by MS, only a small number of sites have been functionally examined. For example, phosphorylation of H1.4 Ser27 (H1.4S27p) by Aurora B kinase blocks the binding of heterochromatin protein 1α to methylated Lys26 (H1.4K26me), suggesting a cross-talk between these modifications (156,157). Zheng et al. showed that interphase phosphorylation at Ser173 on H1.2 (H1.2S173p) and Ser187 on H1.4 (H1.4S187p) is localized to the nucleoli of HeLa S3 cells (89). Phosphorylated Ser187 (H1.4S187p) was further shown to localize to active rDNA promoters, and phosphorylation at this site can be induced by dexamethasone treatment (89). Ser35 phosphorylation on histone H1.4 (H1.4S35p) by protein kinase A mediates H1.4 removal from the mitotic chromatin, suggesting a mechanism of histone H1 mitotic dynamics (158). However, these few examples are not the only sites characterized, and many sites of histone H1 phosphorylations have yet to be functionally described in a site-specific manner.

Although histone H1 phosphorylation is the most researched PTM, other PTMs such as acetylation, methylation and ubiquitination have also been identified (Table 1). The functional relevance of non-phosphorylation PTMs on histone H1 is just coming to light. For example, lysine acetylation at position 34 on histone H1.4 (H1.4K34ac) by the histone acetyltransferase GCN5 has been linked to transcriptional activation and increased dynamic mobility in vitro (159). Additionally, further evidence for histone modification cross-talk was shown by the ARTD1-mediated PARylation of histone H3, which induces a shift in specificity of the methyltransferase SET7/9 from H3 to histone H1 (160). Kassner et al. further identified new sites of histone H1.4 methylation at Lys-Ala-Lys motifs not previously described (160). The role of other non-phosphorylation PTMs on histone H1 function and dynamic mobility is yet to be explored.


Beyond the function of histone H1 on chromatin condensation, histone H1 (specifically H1.2) has been found to have an extrachromatin function. Konishi et al. found translocation of histone H1.2 to the cytoplasm in response to X-ray-induced DNA double-strand breaks (161). Furthermore, Giné et al. show cytosolic movement of H1.2 in chronic lymphocytic leukemia cells after therapeutic intervention (162). The cytosolic histone H1.2 was shown to induce apoptosis through a Bak-mediated mitochondrial release of cytochrome C, allowing for caspase activation (161,162). However, cytosolic histone H1.2 has been observed in non-apoptotic cells as well (14,63) (unpublished data). Collectively, these results suggest there is a mechanism of regulation for histone H1.2 apoptotic induction beyond localization of H1.2 in the cytoplasm. Data presented by Gréen et al. and our own unpublished data suggest H1 isoforms are phosphorylated in the cytoplasm of non-apoptotic cells, giving an underlying potential for apoptosis regulation or chaperone binding (163).


Although the work described previously illustrates the successes of research focused on histone H1, progress has been limited for several reasons. Availability and specificity of immunological reagents for histone H1 are drastically lacking. As methods using antibodies are the primary means of molecular and biochemical investigation, limitations in quality antibodies have caused severe difficulty in study. As a result, a lag in understanding of the biological function of histone H1 and its PTMs persists. The production of immunological reagents is hindered by several factors. The demand for histone H1 antibodies remains generally low. A recent search of PubMed for primary research articles with histone H1 in the title or abstract across the past decade revealed a steady-state number (<100/year) of publications (Figure 1). A similar search for the core histone H3 showed an increase in publications over the past decade (Figure 1). These data suggest histone H1 research has not seen the same explosive growth evident for the chromatin field.

Figure 1.
A search of PubMed for primary research articles containing histone H1 or histone H3 in the title or abstract. Data show a steady-state low number (<100) of publications for histone H1, whereas histone H3 displays an increasing trend over time. ...

We suspect the lull in histone H1 publications can be attributed to two interdependent factors. First, as discussed earlier, histone H1 displays much lower level of evolutionary conservation than the core histones, suggesting that linker histones are not as fundamentally important to chromatin biology. Interest in histone H1 and histone H1 reagents was also dampened by initial studies that suggested that the linker histones were not essential for cell viability. For example, loss of histone H1 genes in M. musculus, T. thermophyla and S. cerevisiae did not affect viability (164–166). However, the essential role of histone H1 in mammals was subsequently affirmed in a landmark publication from the Skoultchi laboratory. Whereas initial single knockout experiments of H1c, H1d and H1e showed no apparent phenotypical changes, a triple knockout of H1c, H1d and H1e (H1.2, H1.3 and H1.4 mouse orthologs) resulted in a 50% reduction in the H1/nucleosome ratio and nucleosome repeat length, ultimately leading to embryonic lethality by E11.5 (167,168). These data suggest a key structural role of histone H1 is to keep the nucleosomes adequately spaced (168,169). Additional in vitro work by Fan et al. showed triple-H1 knockout in embryonic stem cells leads to substantial changes in chromatin structure with variations in gene expression localized to sites of genes regulated by DNA methylation (27). Findings by Yang et al. have confirmed this work by showing histone H1 can interact with DNA-modifying enzymes such as DNMT1 and DNMT3B to alter gene transcription (51). Furthermore, Lee et al. have shown a complex of histone H1b (H1.5 mouse ortholog) and the MSX1 transcription factor represses mesoderm differentiation through the regulation of the MyoD gene, suggesting a role for specific H1 isoforms in the development of muscle (170).

Second, the high sequence homology between variants of histone H1 hinders the ability to produce high-specificity antibodies for individual variants. For example, CLUSTAL 2.1 alignment of the four most common somatic histone H1 variant sequences shows high amino acid sequence conservation (Figure 2A) (171). Pairwise scoring of the sequence alignments between variants shows 74–87% sequence homology (Figure 2B). Domain analysis and sequence alignments show the divergence in the sequences of the H1 variants is primarily located at the amino and carboxy termini of the H1 molecule (Figure 2A). As a result, distinction between variants of histone H1 would require partial identification of epitopes from one of these two domains. However, the high number of PTMs on the terminal tails of histone H1 adds additional complexity that could alter the affinity of antibodies for a significant fraction of the molecules in a cell. Despite these complications, there have been a number of recent successes (89,112,143,157,172–174). Importantly, the use of peptides based on the divergent sequences in the NH2-terminal tails of the H1 variants has led to the production of variant-specific antibodies for both chicken and mammalian H1 (173,174). In addition, the generation of phosphorylation-specific H1 antibodies has begun to shed light on the signal transduction pathways that are involved in the modification of the linker histones. For example, Hergeth et al. generated a polyclonal antibody against phospho Ser27 of H1.4 (H1.4S27p) and demonstrated this phosphorylation was a result of Aurora B kinase activity (157). Additionally, the Lindner laboratory used the commercially available anti phospho-Thr146 H1 (H1.4T146p) antibody to identify this modification on condensed mitotic chromatin by immunofluorescence (111). Furthermore, Chu et al. generated a H1.4 phospho Ser35 (H1.4S35) antibody to show protein kinase A-induced phosphorylation at this site causes removal of H1.4 from the chromatin (158).

Figure 2.
Sequence alignment of histone H1 variants. (A) Amino acid sequence alignment for the histone H1 variants H1.2, H1.3, H1.4 and H1.5. (B) Pairwise scores of sequence homology. The alignment shows a high homology between the human H1 variants.

The amino and carboxy terminal tails of histone H1 variants are among the most abundantly posttranslationally modified sequences in the cell. For example, Figure 3 depicts the known MS-identified PTMs for the histone H1.4 variant. In line with the data in Figure 3, current literature shows multiple numbers of simultaneous PTMs on histone H1 are regularly identified (125,175). These results suggest antibodies generated toward the tail domains of H1 could result in low specificity based on the PTM combinations present on the H1 molecule and the immunogen used for antibody generation. This effect is similar to that seen with histone H4 and H3 modification-specific antibodies, where antibodies must be generated with distinct combinations of localized PTMs to retain specificity for the epitopes of interest. Consequently, MS has become widely used to analyze histone H1 variants through the ability to bypass the limitations of immunological reagents. However, even MS has limitations when analyzing histone H1.

Figure 3.
An illustration of the MS-identified posttranslational modifications on histone H1.4. Asterisk denotes N-α-acetylation of the N-terminal residue after methionine removal.

Limitations in the use of MS for the analysis of histone H1 result from the inability to use common shotgun proteomic methods for analysis. For example, the most commonly used endoproteinase for shotgun proteomic studies is trypsin. In most commonly expressed soluble proteins, trypsin regularly yields peptides of 6–10 amino acids in length due to the relative abundance of lysine and arginine. Additionally, because trypsin cleaves at the C-terminal side of the basic amino acids, each peptide carries at least two sites for protonation, one at the N-terminal and one at the C-terminal lysine or arginine side chain. Thus, when such a peptide is fragmented via tandem MS, two singly charged ions are typically produced. These properties make trypsin ideal for yielding peptides of a mass and charge suitable for liquid chromatography–electrospray ionization–tandem MS. Although ideal for most soluble proteins, trypsin does not work well for histone H1. Figure 4A is a graphical representation of the peptide lengths generated by an in silico digestion of histone H1.4, histone H4 and bovine serum albumin (BSA) with trypsin. The tryptic peptides of H1.4 are very short, with most peptides 5 amino acids or less in length. A similar observation is seen with in silico tryptic digest of histone H4. However, BSA yields peptides of variable length more conducive to MS analysis and increased protein sequence coverage (Table 2). Additionally, the peptides that are generated by trypsin for histone H1.4 have low relative hydrophobicities (Figure 4B, Table 2). This results in low retention of peptides on reversed-phase C18 HPLC columns. Furthermore, of those peptides with hydrophobic properties, appropriate masses and peptide lengths generally correspond to the globular domain of the protein. Collectively, these factors lead to poor sequence coverage when compared with more standard proteins such as BSA. As a result, the use of common bottom-up MS strategies with trypsin is limited.

Figure 4.
A graphical representation of the tryptic peptide length (A) and a histogram of relative hydrophobicities (B) for histone H1.4, histone H4 and bovine serum albumin.
Table 2.
In silico tryptic digestion and relative peptide hydrophobicity

The use of alternative endoproteinases for middle-down proteomics also yields poor digestion results. For example, endoproteinase Glu-C cleaves proteins at the C-terminal side of glutamic acid in ammonium bicarbonate buffer. In silico digests of histone H1.4, histone H4 and BSA with Glu-C, shown in Table 3, yield less-than-optimal peptide lengths and charges. Glu-C digests of histone H1.4 result in long, highly charged peptides not readily suited for liquid chromatography-MS/MS analysis. Additionally, Glu-C cleavage of histone H1.4 gives a peptide encompassing nearly the entire CTD (aa 116–219). Similar results are obtained from in silico digests of histone H4. Conversely, Glu-C digests of BSA yield many suitable peptides in length and charge for MS analysis. Collectively, these results suggest the amino acid sequence of histone H1 does not lend to commonly used bottom-up and middle-down MS strategies.

Table 3.
V8 protease (Glu-C) in silico digestion and charge states

The inability to use bottom-up and middle-down approaches has drastically limited the ability to study histone H1 via MS. However, top-down MS techniques, although limited, have been successfully applied to study histone H1 PTMs. For example in Drosophila melanogaster, Bonet-Costa et al. used top-down MS/MS to map both single and multiple co-existing histone H1 PTMs after collision induced dissociation or electron-capture dissociation (176). Although effectively applied to the single H1 variant in Drosophila, top-down MS/MS on human H1 is severely limited by the necessity for high protein purity, high concentration and separation of the multiple variants. As a result, others have used top-down MS to assess the relative abundance of histone H1 PTMs without fragmentation (89,177,178). For instance, Wang et al. monitored changes in histone H1.5 phosphorylation patterns after drug treatment in acute myeloid leukemia cell lines using intact mass MS (178). While giving the number of modifications and abundances, these approaches do not yield the specific location of the PTM as top-down MS/MS can. Despite these limitations in proteomic methods for the analysis of histone H1, adaptations of these methods in conjunction with state-of-the-art equipment has led to progress in the study of histone H1.


The immunological limitations for studying the function of histone H1 and its PTMs make it a challenging field of research. Although progress has been made, overcoming these difficulties will require combinatorial mass spectral methods. The use of a top-to-bottom proteomics approach will facilitate targeted characterization of specific histone H1 variants and PTMs of interest where a single MS method may fail. Site-directed mutagenesis and the application of single and multiple reaction monitoring experiments to histone H1 variants will allow for further functional descriptions without the necessity for immunological reagents. Collectively, the use of such methods will unlock the specific cellular functions of each histone H1 variant and their respective PTMs.


Funding of this review was provided by grants from the National Institutes of Health [R21 DK082634, R01 CA107106 and R01 GM62970]; and support from The Ohio State University. Funding for open access charge: [R01 CA107106 and R01 GM62970].

Conflict of interest statement. None declared.


1. Kornberg RD. Chromatin structure: a repeating unit of histones and DNA. Science. 1974;184:868–871. [PubMed]
2. Olins AL, Olins DE. Spheroid chromatin units (v bodies) Science. 1974;183:330–332. [PubMed]
3. Thomas JO, Kornberg RD. An octamer of histones in chromatin and free in solution. Proc. Natl Acad. Sci. USA. 1975;72:2626–2630. [PubMed]
4. Oudet P, Gross-Bellard M, Chambon P. Electron microscopic and biochemical evidence that chromatin structure is a repeating unit. Cell. 1975;4:281–300. [PubMed]
5. Arents G, Burlingame RW, Wang BC, Love WE, Moudrianakis EN. The nucleosomal core histone octamer at 3.1 A resolution: a tripartite protein assembly and a left-handed superhelix. Proc. Natl Acad. Sci. USA. 1991;88:10148–10152. [PubMed]
6. Kornberg RD, Thomas JO. Chromatin structure; oligomers of the histones. Science. 1974;184:865–868. [PubMed]
7. Eickbush TH, Moudrianakis EN. The histone core complex: an octamer assembled by two sets of protein-protein interactions. Biochemistry. 1978;17:4955–4964. [PubMed]
8. Whitlock JP, Simpson RT. Removal of histone H1 exposes a fifty base pair DNA segment between nucleosomes. Biochemistry. 1976;15:3307–3314. [PubMed]
9. Noll M, Kornberg RD. Action of micrococcal nuclease on chromatin and the location of histone H1. J. Mol. Biol. 1977;109:393–404. [PubMed]
10. Simpson RT. Structure of the chromatosome, a chromatin particle containing 160 base pairs of DNA and all the histones. Biochemistry. 1978;17:5524–5531. [PubMed]
11. Richmond TJ, Finch JT, Rushton B, Rhodes D, Klug A. Structure of the nucleosome core particle at 7 A resolution. Nature. 1984;311:532–537. [PubMed]
12. Luger K, Mäder AW, Richmond RK, Sargent DF, Richmond TJ. Crystal structure of the nucleosome core particle at 2.8 A resolution. Nature. 1997;389:251–260. [PubMed]
13. Finch JT, Klug A. Solenoidal model for superstructure in chromatin. Proc. Natl Acad. Sci. USA. 1976;73:1897–1901. [PubMed]
14. Thoma F, Koller T, Klug A. Involvement of histone H1 in the organization of the nucleosome and of the salt-dependent superstructures of chromatin. J. Cell. Biol. 1979;83:403–427. [PMC free article] [PubMed]
15. Van Holde KE. New York: Springer-Verlag; 1989. Chromatin.
16. Carruthers LM, Bednar J, Woodcock CL, Hansen JC. Linker Histones Stabilize the Intrinsic Salt-Dependent Folding of Nucleosomal Arrays: Mechanistic Ramifications for Higher-Order Chromatin Folding. Biochemistry. 1998;37:14776–14787. [PubMed]
17. Allan J, Hartman PG, Crane-Robinson C, Aviles FX. The structure of histone H1 and its location in chromatin. Nature. 1980;288:675–679. [PubMed]
18. Goytisolo FA, Gerchman SE, Yu X, Rees C, Graziano V, Ramakrishnan V, Thomas JO. Identification of two DNA-binding sites on the globular domain of histone H5. EMBO J. 1996;15:3421–3429. [PubMed]
19. Bednar J, Horowitz RA, Grigoryev SA, Carruthers LM, Hansen JC, Koster AJ, Woodcock CL. Nucleosomes, linker DNA, and linker histone form a unique structural motif that directs the higher-order folding and compaction of chromatin. Proc. Natl Acad. Sci. USA. 1998;95:14173–14178. [PubMed]
20. Zhou YB, Gerchman SE, Ramakrishnan V, Travers A, Muyldermans S. Position and orientation of the globular domain of linker histone H5 on the nucleosome. Nature. 1998;395:402–405. [PubMed]
21. Widom J. Chromatin structure: linking structure to function with histone H1. Curr. Biol. 1998;8:R788–91. [PubMed]
22. Thomas JO. Histone H1: location and role. Curr. Opin. Cell Biol. 1999;11:312–317. [PubMed]
23. Sivolob A, Prunell A. Linker histone-dependent organization and dynamics of nucleosome entry/exit DNAs. J. Mol. Biol. 2003;331:1025–1040. [PubMed]
24. Maier VK, Chioda M, Rhodes D, Becker PB. ACF catalyses chromatosome movements in chromatin fibres. EMBO J. 2008;27:817–826. [PubMed]
25. Syed SH, Goutte-Gattat D, Becker N, Meyer S, Shukla MS, Hayes JJ, Everaers R, Angelov D, Bednar J, Dimitrov S. Single-base resolution mapping of H1-nucleosome interactions and 3D organization of the nucleosome. Proc. Natl Acad. Sci. USA. 2010;107:9620–9625. [PubMed]
26. Meyer S, Becker NB, Syed SH, Goutte-Gattat D, Shukla MS, Hayes JJ, Angelov D, Bednar J, Dimitrov S, Everaers R. From crystal and NMR structures, footprints and cryo-electron-micrographs to large and soft structures: nanoscale modeling of the nucleosomal stem. Nucleic Acids Res. 2011;39:9139–9154. [PMC free article] [PubMed]
27. Fan Y, Nikitina T, Zhao J, Fleury TJ, Bhattacharyya R, Bouhassira EE, Stein A, Woodcock CL, Skoultchi AI. Histone H1 depletion in mammals alters global chromatin structure but causes specific changes in gene regulation. Cell. 2005;123:1199–1212. [PubMed]
28. Hizume K, Yoshimura SH, Takeyasu K. Linker histone H1 per se can induce three-dimensional folding of chromatin fiber. Biochemistry. 2005;44:12978–12989. [PubMed]
29. Schalch T, Duda S, Sargent DF, Richmond TJ. X-ray structure of a tetranucleosome and its implications for the chromatin fibre. Nature. 2005;436:138–141. [PubMed]
30. Robinson PJJ, Fairall L, Huynh VAT, Rhodes D. EM measurements define the dimensions of the ‘30-nm’ chromatin fiber: evidence for a compact, interdigitated structure. Proc. Natl Acad. Sci. USA. 2006;103:6506–6511. [PubMed]
31. Robinson PJJ, Rhodes D. Structure of the ‘30 nm’ chromatin fibre: a key role for the linker histone. Curr. Opin. Struct. Biol. 2006;16:336–343. [PubMed]
32. Wong H, Victor J-M, Mozziconacci J. An all-atom model of the chromatin fiber containing linker histones reveals a versatile structure tuned by the nucleosomal repeat length. PLoS One. 2007;2:e877. [PMC free article] [PubMed]
33. Routh A, Sandin S, Rhodes D. Nucleosome repeat length and linker histone stoichiometry determine chromatin fiber structure. Proc. Natl Acad. Sci. USA. 2008;105:8872–8877. [PubMed]
34. Ramakrishnan V, Finch JT, Graziano V, Lee PL, Sweet RM. Crystal structure of globular domain of histone H5 and its implications for nucleosome binding. Nature. 1993;362:219–223. [PubMed]
35. Bohm L, Mitchell TC. Sequence conservation in the N-terminal domain of histone H1. FEBS Lett. 1985;193:1–4. [PubMed]
36. Allan J, Mitchell T, Harborne N, Bohm L, Crane-Robinson C. Roles of H1 domains in determining higher order chromatin structure and H1 location. J. Mol. Biol. 1986;187:591–601. [PubMed]
37. Vila R, Ponte I, Collado M, Arrondo JL, Jiménez MA, Rico M, Suau P. DNA-induced alpha-helical structure in the NH2-terminal domain of histone H1. J. Biol. Chem. 2001;276:46429–46435. [PubMed]
38. Hendzel MJ, Lever MA, Crawford E, Th'ng JPH. The C-terminal domain is the primary determinant of histone H1 binding to chromatin in vivo. J. Biol. Chem. 2004;279:20028–20034. [PubMed]
39. Öberg C, Belikov S. The N-terminal domain determines the affinity and specificity of H1 binding to chromatin. Biochem. Biophys. Res. Commun. 2012;420:321–324. [PubMed]
40. Vyas P, Brown DT. N- and C-terminal domains determine differential nucleosomal binding geometry and affinity of linker histone isotypes H10 and H1c. J. Biol. Chem. 2012;287:11778–11787. [PubMed]
41. Bradbury EM, Danby SE, Rattle HW, Giancotti V. Studies on the role and mode of operation of the very-lysine-rich histone H1 (F1) in eukaryote chromatin. Histone H1 in chromatin and in H1 - DNA complexes. Eur. J. Biochem. 1975;57:97–105. [PubMed]
42. Roque A. DNA-induced Secondary Structure of the Carboxyl-terminal Domain of Histone H1. J. Biol. Chem. 2005;280:32141–32147. [PubMed]
43. Hansen JC, Lu X, Ross ED, Woody RW. Intrinsic protein disorder, amino acid composition, and histone terminal domains. J. Biol. Chem. 2006;281:1853–1856. [PubMed]
44. Lu X, Hamkalo B, Parseghian MH, Hansen JC. Chromatin condensing functions of the linker histone C-terminal domain are mediated by specific amino acid composition and intrinsic protein disorder. Biochemistry. 2009;48:164–172. [PMC free article] [PubMed]
45. Caterino TL, Fang H, Hayes JJ. Nucleosome linker DNA contacts and induces specific folding of the intrinsically disordered H1 carboxyl-terminal domain. Mol. Cell. Biol. 2011;31:2341–2348. [PMC free article] [PubMed]
46. Clark DJ, Hill CS, Martin SR, Thomas JO. Alpha-helix in the carboxy-terminal domains of histones H1 and H5. EMBO J. 1988;7:69–75. [PubMed]
47. Hamiche A, Schultz P, Ramakrishnan V, Oudet P, Prunell A. Linker histone-dependent DNA structure in linear mononucleosomes. J. Mol. Biol. 1996;257:30–42. [PubMed]
48. Roque A, Ponte I, Suau P. Role of charge neutralization in the folding of the carboxy-terminal domain of histone H1. J. Phys. Chem. B. 2009;113:12061–12066. [PubMed]
49. Fang HH, Clark DJD, Hayes JJJ. DNA and nucleosomes direct distinct folding of a linker histone H1 C-terminal domain. Nucleic Acids Res. 2012;40:1475–1484. [PMC free article] [PubMed]
50. Lu X. Identification of specific functional subdomains within the linker histone H10 C-terminal domain. J. Biol. Chem. 2004;279:8701–8707. [PubMed]
51. Yang S-M, Kim BJ, Norwood Toro L, Skoultchi AI. H1 linker histone promotes epigenetic silencing by regulating both DNA methylation and histone H3 methylation. Proc. Natl Acad. Sci. USA. 2013;110:1708–1713. [PubMed]
52. Clark DJ, Kimura T. Electrostatic mechanism of chromatin folding. J. Mol. Biol. 1990;211:883–896. [PubMed]
53. Subirana JA. Analysis of the charge distribution in the C-terminal region of histone H1 as related to its interaction with DNA. Biopolymers. 1990;29:1351–1357. [PubMed]
54. Lever MA, Th'ng JP, Sun X, Hendzel MJ. Rapid exchange of histone H1.1 on chromatin in living human cells. Nature. 2000;408:873–876. [PubMed]
55. Misteli T, Gunjan A, Hock R, Bustin M, Brown DT. Dynamic binding of histone H1 to chromatin in living cells. Nature. 2000;408:877–881. [PubMed]
56. Doenecke D, Tönjes R. Differential distribution of lysine and arginine residues in the closely related histones H1° and H5. J. Mol. Biol. 1986;187:461–464. [PubMed]
57. Wu M, Allis CD, Richman R, Cook RG, Gorovsky MA. An intervening sequence in an unusual histone H1 gene of Tetrahymena thermophila. Proc. Natl Acad. Sci. USA. 1986;83:8674–8678. [PubMed]
58. Ali T, Thomas JO. Distinct properties of the two putative ‘globular domains’ of the yeast linker histone, Hho1p. J. Mol. Biol. 2004;337:1123–1135. [PubMed]
59. Izzo A, Kamieniarz K, Schneider R. The histone H1 family: specific members, specific functions? Biol. Chem. 2008;389:333–343. [PubMed]
60. Eick S, Nicolai M, Mumberg D, Doenecke D. Human H1 histones: conserved and varied sequence elements in two H1 subtype genes. Eur. J. Cell Biol. 1989;49:110–115. [PubMed]
61. Albig W, Kardalinou E, Drabent B, Zimmer A, Doenecke D. Isolation and characterization of two human H1 histone genes within clusters of core histone genes. Genomics. 1991;10:940–948. [PubMed]
62. Albig W, Doenecke D. The human histone gene cluster at the D6S105 locus. Hum. Genet. 1997;101:284–294. [PubMed]
63. Carozzi N, Marashi F, Plumb M, Zimmerman S, Zimmerman A, Coles LS, Wells JR, Stein G, Stein J. Clustering of human H1 and core histone genes. Science. 1984;224:1115–1117. [PubMed]
64. Drabent B, Kardalinou E, Doenecke D. Structure and expression of the human gene encoding testicular H1 histone (H1t) Gene. 1991;103:263–268. [PubMed]
65. Drabent B, Franke K, Bode C, Kosciessa U, Bouterfa H, Hameister H, Doenecke D. Isolation of two murine H1 histone genes and chromosomal mapping of the H1 gene complement. Mamm. Genome. 1995;6:505–511. [PubMed]
66. Yamamoto T, Horikoshi M. Cloning of the cDNA encoding a novel subtype of histone H1. Gene. 1996;173:281–285. [PubMed]
67. Tanaka M, Hennebold JD, Macfarlane J, Adashi EY. A mammalian oocyte-specific linker histone gene H1oo: homology with the genes for the oocyte-specific cleavage stage histone (cs-H1) of sea urchin and the B4/H1M histone of the frog. Development. 2001;128:655–664. [PubMed]
68. Yan W. HILS1 is a spermatid-specific linker histone H1-like protein implicated in chromatin remodeling during mammalian spermiogenesis. Proc. Natl Acad. Sci. USA. 2003;100:10546–10551. [PubMed]
69. Martianov I, Brancorsini S, Catena R, Gansmuller A, Kotaja N, Parvinen M, Sassone-Corsi P, Davidson I. Polar nuclear localization of H1T2, a histone H1 variant, required for spermatid elongation and DNA condensation during spermiogenesis. Proc. Natl Acad. Sci. USA. 2005;102:2808–2813. [PubMed]
70. Happel N, Schulze E, Doenecke D. Characterisation of human histone H1x. Biol. Chem. 2005;386:541–551. [PubMed]
71. Tanaka H, Matsuoka Y, Onishi M, Kitamura K, Miyagawa Y, Nishimura H, Tsujimura A, Okuyama A, Nishimune Y. Expression profiles and single-nucleotide polymorphism analysis of human HANP1/H1T2 encoding a histone H1-like protein. Int. J. Androl. 2006;29:353–359. [PubMed]
72. Pehrson JR, Cole RD. Histone H1 subfractions and H10 turnover at different rates in nondividing cells. Biochemistry. 1982;21:456–460. [PubMed]
73. Gurley LR, Walters RA, Tobey RA. The metabolism of histone fractions. IV. Synthesis of histones during the G1-phase of the mammalian life cycle. Arch. Biochem. Biophys. 1972;148:633–641. [PubMed]
74. Appels R, Ringertz NR. Metabolism of F1 histone in G1 and G0 cells. Cell Differ. 1974;3:1–8. [PubMed]
75. Tarnowka MA, Baglioni C, Basilico C. Synthesis of H1 histones by BHK cells in G1. Cell. 1978;15:163–171. [PubMed]
76. Wu RS, Bonner WM. Separation of basal histone synthesis from S-phase histone synthesis in dividing cells. Cell. 1981;27:321–330. [PubMed]
77. Plumb M, Marashi F, Green L, Zimmerman A, Zimmerman S, Stein J, Stein G. Cell cycle regulation of human histone H1 mRNA. Proc. Natl Acad. Sci. USA. 1984;81:434–438. [PubMed]
78. Albig W, Drabent B, Kunz J, Kalff-Suske M, Grzeschik KH, Doenecke D. All known human H1 histone genes except the H1(0) gene are clustered on chromosome 6. Genomics. 1993;16:649–654. [PubMed]
79. Albig W, Kioschis P, Poustka A, Meergans K, Doenecke D. Human histone gene organization: nonregular arrangement within a large cluster. Genomics. 1997;40:314–322. [PubMed]
80. Hohmann P, Cole RD. Hormonal effects on amino acid incorporation into lysine-rich histones in the mouse mammary gland. J. Mol. Biol. 1971;58:533–540. [PubMed]
81. Felsenfeld G. Chromatin. Nature. 1978;271:115–122. [PubMed]
82. Sizemore SR, Cole RD. Asynchronous appearance of newly synthesized histone H1 subfractions in HeLa chromatin. J. Cell. Biol. 1981;90:415–417. [PMC free article] [PubMed]
83. Heintz N, Sive HL, Roeder RG. Regulation of human histone gene expression: kinetics of accumulation and changes in the rate of synthesis and in the half-lives of individual histone mRNAs during the HeLa cell cycle. Mol. Cell. Biol. 1983;3:539–550. [PMC free article] [PubMed]
84. Plumb M, Stein J, Stein G. Coordinate regulation of multiple histone mRNAs during the cell cycle in HeLa cells. Nucleic Acids Res. 1983;11:2391–2410. [PMC free article] [PubMed]
85. Pandey NB, Chodchoy N, Liu TJ, Marzluff WF. Introns in histone genes alter the distribution of 3′ ends. Nucleic Acids Res. 1990;18:3161–3170. [PMC free article] [PubMed]
86. Dominski Z, Marzluff WF. Formation of the 3′ end of histone mRNA. Gene. 1999;239:1–14. [PubMed]
87. Sittman DB, Graves RA, Marzluff WF. Histone mRNA concentrations are regulated at the level of transcription and mRNA degradation. Proc. Natl Acad. Sci. USA. 1983;80:1849–1853. [PubMed]
88. Sarg B, Gréen A, Söderkvist P, Helliger W, Rundquist I, Lindner HH. Characterization of sequence variations in human histone H1.2 and H1.4 subtypes. FEBS J. 2005;272:3673–3683. [PubMed]
89. Zheng Y, John S, Pesavento JJ, Schultz-Norton JR, Schiltz RL, Baek S, Nardulli AM, Hager GL, Kelleher NL, Mizzen CA. Histone H1 phosphorylation is associated with transcription by RNA polymerases I and II. J. Cell. Biol. 2010;189:407–415. [PMC free article] [PubMed]
90. Brown DT, Alexander BT, Sittman DB. Differential effect of H1 variant overexpression on cell cycle progression and gene expression. Nucleic Acids Res. 1996;24:486–493. [PMC free article] [PubMed]
91. Gunjan A, Alexander BT, Sittman DB, Brown DT. Effects of H1 histone variant overexpression on chromatin structure. J. Biol. Chem. 1999;274:37950–37956. [PubMed]
92. Talasz H, Sapojnikova N, Helliger W, Lindner H, Puschendorf B. In vitro binding of H1 histone subtypes to nucleosomal organized mouse mammary tumor virus long terminal repeat promotor. J. Biol. Chem. 1998;273:32236–32243. [PubMed]
93. Th'ng JPH, Sung R, Ye M, Hendzel MJ. H1 family histones in the nucleus. Control of binding and localization by the C-terminal domain. J. Biol. Chem. 2005;280:27809–27814. [PubMed]
94. Orrego M, Ponte I, Roque A, Buschati N, Mora X, Suau P. Differential affinity of mammalian histone H1 somatic subtypes for DNA and chromatin. BMC Biol. 2007;5:22. [PMC free article] [PubMed]
95. Bradbury EM, Inglis RJ, Matthews HR, Sarner N. Phosphorylation of very-lysine-rich histone in Physarum polycephalum. Correlation with chromosome condensation. Eur. J. Biochem. 1973;33:131–139. [PubMed]
96. Gurley LR, Walters RA, Tobey RA. Sequential phosphorylation of histone subfractions in the Chinese hamster cell cycle. J. Biol. Chem. 1975;250:3936–3944. [PubMed]
97. Hohmann P, Tobey RA, Gurley LR. Phosphorylation of distinct regions of f1 histone. Relationship to the cell cycle. J. Biol. Chem. 1976;251:3685–3692. [PubMed]
98. D'Anna JA, Gurley LR, Deaven LL. Dephosphorylation of histones H1 and H3 during the isolation of metaphase chromosomes. Nucleic Acids Res. 1978;5:3195–3208. [PMC free article] [PubMed]
99. Gurley LR, D'Anna JA, Barham SS, Deaven LL, Tobey RA. Histone phosphorylation and chromatin structure during mitosis in Chinese hamster cells. Eur. J. Biochem. 1978;84:1–15. [PubMed]
100. Matsumoto Y, Yasuda H, Mita S, Marunouchi T, Yamada M. Evidence for the involvement of H1 histone phosphorylation in chromosome condensation. Nature. 1980;284:181–183. [PubMed]
101. Ajiro K, Borun TW, Cohen LH. Phosphorylation states of different histone 1 subtypes and their relationship to chromatin functions during the HeLa S-3 cell cycle. Biochemistry. 1981;20:1445–1454. [PubMed]
102. Th'ng JP, Guo XW, Swank RA, Crissman HA, Bradbury EM. Inhibition of histone phosphorylation by staurosporine leads to chromosome decondensation. J. Biol. Chem. 1994;269:9568–9573. [PubMed]
103. Gurley LR, Valdez JG, Buchanan JS. Characterization of the mitotic specific phosphorylation site of histone H1. Absence of a consensus sequence for the p34cdc2/cyclin B kinase. J. Biol. Chem. 1995;270:27653–27660. [PubMed]
104. Taylor WR, Chadee DN, Allis CD, Wright JA, Davie JR. Fibroblasts transformed by combinations of ras, myc and mutant p53 exhibit increased phosphorylation of histone H1 that is independent of metastatic potential. FEBS Lett. 1995;377:51–53. [PubMed]
105. Chadee DN, Taylor WR, Hurta RA, Allis CD, Wright JA, Davie JR. Increased phosphorylation of histone H1 in mouse fibroblasts transformed with oncogenes or constitutively active mitogen-activated protein kinase. J. Biol. Chem. 1995;270:20098–20105. [PubMed]
106. Herrera RE, Chen F, Weinberg RA. Increased histone H1 phosphorylation and relaxed chromatin structure in Rb-deficient fibroblasts. Proc. Natl Acad. Sci. USA. 1996;93:11510–11515. [PubMed]
107. Chadee DN, Allis CD, Wright JA, Davie JR. Histone H1b phosphorylation is dependent upon ongoing transcription and replication in normal and ras-transformed mouse fibroblasts. J. Biol. Chem. 1997;272:8113–8116. [PubMed]
108. Roque A, Ponte I, Arrondo JLR, Suau P. Phosphorylation of the carboxy-terminal domain of histone H1: effects on secondary structure and DNA condensation. Nucleic Acids Res. 2008;36:4719–4726. [PMC free article] [PubMed]
109. Dou Y, Bowen J, Liu Y, Gorovsky MA. Phosphorylation and an ATP-dependent process increase the dynamic exchange of H1 in chromatin. J. Cell. Biol. 2002;158:1161–1170. [PMC free article] [PubMed]
110. Alexandrow MG, Hamlin JL. Chromatin decondensation in S-phase involves recruitment of Cdk2 by Cdc45 and histone H1 phosphorylation. J. Cell. Biol. 2005;168:875–886. [PMC free article] [PubMed]
111. Sarg B, Helliger W, Talasz H, Förg B, Lindner HH. Histone H1 phosphorylation occurs site-specifically during interphase and mitosis: identification of a novel phosphorylation site on histone H1. J. Biol. Chem. 2006;281:6573–6580. [PubMed]
112. Talasz H, Sarg B, Lindner HH. Site-specifically phosphorylated forms of H1.5 and H1.2 localized at distinct regions of the nucleus are related to different processes during the cell cycle. Chromosoma. 2009;118:693–709. [PubMed]
113. Contreras A, Hale TK, Stenoien DL, Rosen JM, Mancini MA, Herrera RE. The dynamic mobility of histone H1 is regulated by cyclin/CDK phosphorylation. Mol. Cell. Biol. 2003;23:8626–8636. [PMC free article] [PubMed]
114. Bustin M, Catez F, Lim JH. The dynamics of histone H1 function in chromatin. Mol. Cell. 2005;17:617–620. [PubMed]
115. Lee HL, Archer TK. Prolonged glucocorticoid exposure dephosphorylates histone H1 and inactivates the MMTV promoter. EMBO J. 1998;17:1454–1466. [PubMed]
116. Hale TK, Contreras A, Morrison AJ, Herrera RE. Phosphorylation of the linker histone H1 by CDK regulates its binding to HP1alpha. Mol. Cell. 2006;22:693–699. [PubMed]
117. Hill DA, Imbalzano AN. Human SWI/SNF nucleosome remodeling activity is partially inhibited by linker histone H1. Biochemistry. 2000;39:11649–11656. [PubMed]
118. Ramachandran A. Linker histone H1 modulates nucleosome remodeling by human SWI/SNF. J. Biol. Chem. 2003;278:48590–48601. [PubMed]
119. Horn PJ, Carruthers LM, Logie C, Hill DA, Solomon MJ, Wade PA, Imbalzano AN, Hansen JC, Peterson CL. Phosphorylation of linker histones regulates ATP-dependent chromatin remodeling enzymes. Nat. Struct. Biol. 2002;9:263–267. [PubMed]
120. Clausell J, Happel N, Hale TK, Doenecke D, Beato M. Histone H1 subtypes differentially modulate chromatin condensation without preventing ATP-dependent remodeling by SWI/SNF or NURF. PLoS One. 2009;4:e0007243. [PMC free article] [PubMed]
121. Cheung E, Zarifyan AS, Kraus WL. Histone H1 represses estrogen receptor α transcriptional activity by selectively inhibiting receptor-mediated transcription initiation. Mol. Cell. Biol. 2002;22:2463–2471. [PMC free article] [PubMed]
122. Bhattacharjee RN, Banks GC, Trotter KW, Lee HL, Archer TK. Histone H1 phosphorylation by Cdk2 selectively modulates mouse mammary tumor virus transcription through chromatin remodeling. Mol. Cell. Biol. 2001;21:5417–5425. [PMC free article] [PubMed]
123. Koop R, Di Croce L, Beato M. Histone H1 enhances synergistic activation of the MMTV promoter in chromatin. EMBO J. 2003;22:588–599. [PubMed]
124. Herrera JE, West KL, Schiltz RL, Nakatani Y, Bustin M. Histone H1 is a specific repressor of core histone acetylation in chromatin. Mol. Cell. Biol. 2000;20:523–529. [PMC free article] [PubMed]
125. Wiśniewski JR, Zougman A, Krüger S, Mann M. Mass spectrometric mapping of linker histone H1 variants reveals multiple acetylations, methylations, and phosphorylation as well as differences between cell culture and tissue. Mol. Cell. Proteomics. 2007;6:72–87. [PubMed]
126. Molina H, Horn DM, Tang N, Mathivanan S, Pandey A. Global proteomic profiling of phosphopeptides using electron transfer dissociation tandem mass spectrometry. Proc. Natl Acad. Sci. USA. 2007;104:2199–2204. [PubMed]
127. Meierhofer D, Wang X, Huang L, Kaiser P. Quantitative analysis of global ubiquitination in HeLa cells by mass spectrometry. J. Proteome. Res. 2008;7:4566–4576. [PMC free article] [PubMed]
128. Wang B, Malik R, Nigg EA, Körner R. Evaluation of the low-specificity protease elastase for large-scale phosphoproteome analysis. Anal. Chem. 2008;80:9526–9533. [PubMed]
129. Olsen JV, Blagoev B, Gnad F, Macek B, Kumar C, Mortensen P, Mann M. Global, in vivo, and site-specific phosphorylation dynamics in signaling networks. Cell. 2006;127:635–648. [PubMed]
130. Yu L-R, Zhu Z, Chan KC, Issaq HJ, Dimitrov DS, Veenstra TD. Improved titanium dioxide enrichment of phosphopeptides from HeLa cells and high confident phosphopeptide identification by cross-validation of MS/MS and MS/MS/MS spectra. J. Proteome. Res. 2007;6:4150–4162. [PubMed]
131. Carrascal M, Ovelleiro D, Casas V, Gay M, Abian J. Phosphorylation analysis of primary human T lymphocytes using sequential IMAC and titanium oxide enrichment. J. Proteome. Res. 2008;7:5167–5176. [PubMed]
132. Dephoure N, Zhou C, Villén J, Beausoleil SA, Bakalarski CE, Elledge SJ, Gygi SP. A quantitative atlas of mitotic phosphorylation. Proc. Natl Acad. Sci. USA. 2008;105:10762–10767. [PubMed]
133. Mayya V, Lundgren DH, Hwang S-I, Rezaul K, Wu L, Eng JK, Rodionov V, Han DK. Quantitative phosphoproteomic analysis of T cell receptor signaling reveals system-wide modulation of protein-protein interactions. Sci. Signal. 2009;2 ra46. [PubMed]
134. Weiss T, Hergeth S, Zeissler U, Izzo A, Tropberger P, Zee BM, Dundr M, Garcia BA, Daujat S, Schneider R. Histone H1 variant-specific lysine methylation by G9a/KMT1C and Glp1/KMT1D. Epigenetic Chromatin. 2010;3:7. [PMC free article] [PubMed]
135. Lu A, Zougman A, Pudełko M, Bebenek M, Ziółkowski P, Mann M, Wiśniewski JR. Mapping of lysine monomethylation of linker histones in human breast and its cancer. J. Proteome. Res. 2009;8:4207–4215. [PubMed]
136. Wagner SA, Beli P, Weinert BT, Nielsen ML, Cox J, Mann M, Choudhary C. A proteome-wide, quantitative survey of in vivo ubiquitylation sites reveals widespread regulatory roles. Mol. Cell. Proteomics. 2011;10 M111.013284. [PMC free article] [PubMed]
137. Wiśniewski JR, Zougman A, Mann M. Nepsilon-formylation of lysine is a widespread post-translational modification of nuclear proteins occurring at residues involved in regulation of chromatin function. Nucleic Acids Res. 2008;36:570–577. [PMC free article] [PubMed]
138. Ohe Y, Hayashi H, Iwai K. Human spleen histone H1. Isolation and amino acid sequences of three minor variants, H1a, H1c, and H1d. J. Biochem. 1989;106:844–857. [PubMed]
139. Gauci S, Helbig AO, Slijper M, Krijgsveld J, Heck AJR, Mohammed S. Lys-N and trypsin cover complementary parts of the phosphoproteome in a refined SCX-based approach. Anal. Chem. 2009;81:4493–4501. [PubMed]
140. Vaquero A, Scher M, Lee D, Erdjument-Bromage H, Tempst P, Reinberg D. Human SirT1 interacts with histone H1 and promotes formation of facultative heterochromatin. Mol. Cell. 2004;16:93–105. [PubMed]
141. Choudhary C, Kumar C, Gnad F, Nielsen ML, Rehman M, Walther TC, Olsen JV, Mann M. Lysine Acetylation Targets Protein Complexes and Co-Regulates Major Cellular Functions. Sci. Signal. 2009;325:834. [PubMed]
142. Daub H, Olsen JV, Bairlein M, Gnad F, Oppermann FS, Körner R, Greff Z, Kéri G, Stemmann O, Mann M. Kinase-selective enrichment enables quantitative phosphoproteomics of the kinome across the cell cycle. Mol. Cell. 2008;31:438–448. [PubMed]
143. Happel N, Stoldt S, Schmidt B, Doenecke D. M phase-specific phosphorylation of histone H1.5 at threonine 10 by GSK-3. J. Mol. Biol. 2009;386:339–350. [PubMed]
144. Kim JE, Tannenbaum SR, White FM. Global phosphoproteome of HT-29 human colon adenocarcinoma cells. J. Proteome. Res. 2005;4:1339–1346. [PubMed]
145. Banks GC, Deterding LJ, Tomer KB, Archer TK. Hormone-mediated dephosphorylation of specific histone H1 isoforms. J. Biol. Chem. 2001;276:36467–36473. [PubMed]
146. Deterding LJ, Banks GC, Tomer KB, Archer TK. Understanding global changes in histone H1 phosphorylation using mass spectrometry. Methods. 2004;33:53–58. [PubMed]
147. Deterding LJ, Bunger MK, Banks GC, Tomer KB, Archer TK. Global changes in and characterization of specific sites of phosphorylation in mouse and human histone H1 isoforms upon CDK inhibitor treatment using mass spectrometry. J. Proteome Res. 2008;7:2368–2379. [PMC free article] [PubMed]
148. Roberge M, Th'ng J, Hamaguchi J, Bradbury EM. The topoisomerase II inhibitor VM-26 induces marked changes in histone H1 kinase activity, histones H1 and H3 phosphorylation, and chromosome condensation in G2 phase and mitotic BHK cells. J. Cell. Biol. 1990;111:1753–1762. [PMC free article] [PubMed]
149. Thiriet C, Hayes JJ. Linker histone phosphorylation regulates global timing of replication origin firing. J. Biol. Chem. 2009;284:2823–2829. [PubMed]
150. Ransom M, Dennehey BK, Tyler JK. Chaperoning histones during DNA replication and repair. Cell. 2010;140:183–195. [PMC free article] [PubMed]
151. Richardson RT, Batova IN, Widgren EE, Zheng LX, Whitfield M, Marzluff WF, O'Rand MG. Characterization of the histone H1-binding protein, NASP, as a cell cycle-regulated somatic protein. J. Biol. Chem. 2000;275:30378–30386. [PubMed]
152. Wang H, Ge Z, Walsh STR, Parthun MR. The human histone chaperone sNASP interacts with linker and core histones through distinct mechanisms. Nucleic Acids Res. 2012;40:660–669. [PMC free article] [PubMed]
153. Richardson RT, Alekseev OM, Grossman G, Widgren EE, Thresher R, Wagner EJ, Sullivan KD, Marzluff WF, O'Rand MG. Nuclear autoantigenic sperm protein (NASP), a linker histone chaperone that is required for cell proliferation. J. Biol. Chem. 2006;281:21526–21534. [PubMed]
154. Finn RM, Browne K, Hodgson KC, Ausió J. sNASP, a histone H1-specific eukaryotic chaperone dimer that facilitates chromatin assembly. Biophys. J. 2008;95:1314–1325. [PubMed]
155. Wang H, Walsh STR, Parthun MR. Expanded binding specificity of the human histone chaperone NASP. Nucleic Acids Res. 2008;36:5763–5772. [PMC free article] [PubMed]
156. Daujat S, Zeissler U, Waldmann T, Happel N, Schneider R. HP1 binds specifically to Lys26-methylated histone H1.4, whereas simultaneous Ser27 phosphorylation blocks HP1 binding. J. Biol. Chem. 2005;280:38090–38095. [PubMed]
157. Hergeth SP, Dundr M, Tropberger P, Zee BM, Garcia BA, Daujat S, Schneider R. Isoform-specific phosphorylation of human linker histone H1.4 in mitosis by the kinase Aurora B. J. Cell Sci. 2011;124:1623–1628. [PubMed]
158. Chu CS, Hsu PH, Lo PW, Scheer E, Tora L, Tsai HJ, Tsai MD, Juan LJ. Protein kinase A-mediated serine 35 phosphorylation dissociates histone H1.4 from mitotic chromosome. J. Biol. Chem. 2011;286:35843–35851. [PMC free article] [PubMed]
159. Kamieniarz KK, Izzo AA, Dundr MM, Tropberger PP, Ozretic LL, Kirfel JJ, Scheer EE, Tropel PP, Wisniewski JRJ, Tora LL, et al. A dual role of linker histone H1.4 Lys 34 acetylation in transcriptional activation. Genes Dev. 2012;26:797–802. [PubMed]
160. Kassner II, Barandun MM, Fey MM, Rosenthal FF, Hottiger MOM. Crosstalk between SET7/9-dependent methylation and ARTD1-mediated ADP-ribosylation of histone H1.4. Epigenetic Chromatin. 2013;6:1–1. [PMC free article] [PubMed]
161. Konishi A, Shimizu S, Hirota J, Takao T, Fan Y, Matsuoka Y, Zhang L, Yoneda Y, Fujii Y, Skoultchi AI, et al. Involvement of histone H1.2 in apoptosis induced by DNA double-strand breaks. Cell. 2003;114:673–688. [PubMed]
162. Giné E, Crespo M, Muntañola A, Calpe E, Baptista MJ, Villamor N, Montserrat E, Bosch F. Induction of histone H1.2 cytosolic release in chronic lymphocytic leukemia cells after genotoxic and non-genotoxic treatment. Haematologica. 2008;93:75–82. [PubMed]
163. Gréen A, Lönn A, Peterson KH, Ollinger K, Rundquist I. Translocation of histone H1 subtypes between chromatin and cytoplasm during mitosis in normal human fibroblasts. Cytometry A. 2010;77:478–484. [PubMed]
164. Shen X, Yu L, Weir JW, Gorovsky MA. Linker histones are not essential and affect chromatin condensation in vivo. Cell. 1995;82:47–56. [PubMed]
165. Sirotkin AM, Edelmann W, Cheng G, Klein-Szanto A, Kucherlapati R, Skoultchi AI. Mice develop normally without the H1(0) linker histone. Proc. Natl Acad. Sci. USA. 1995;92:6434–6438. [PubMed]
166. Patterton HG, Landel CC, Landsman D, Peterson CL, Simpson RT. The biochemical and phenotypic characterization of Hho1p, the putative linker histone H1 of Saccharomyces cerevisiae. J. Biol. Chem. 1998;273:7268–7276. [PubMed]
167. Fan Y, Sirotkin A, Russell RG, Ayala J, Skoultchi AI. Individual somatic H1 subtypes are dispensable for mouse development even in mice lacking the H1(0) replacement subtype. Mol. Cell. Biol. 2001;21:7933–7943. [PMC free article] [PubMed]
168. Fan Y, Nikitina T, Morin-Kensicki EM, Zhao J, Magnuson TR, Woodcock CL, Skoultchi AI. H1 linker histones are essential for mouse development and affect nucleosome spacing in vivo. Mol. Cell. Biol. 2003;23:4559–4572. [PMC free article] [PubMed]
169. Woodcock CL, Skoultchi AI, Fan Y. Role of linker histone in chromatin structure and function: H1 stoichiometry and nucleosome repeat length. Chromosome Res. 2006;14:17–25. [PubMed]
170. Lee H, Habas R, Abate-Shen C. MSX1 cooperates with histone H1b for inhibition of transcription and myogenesis. Science. 2004;304:1675–1678. [PubMed]
171. Meergans T, Albig W, Doenecke D. Varied expression patterns of human H1 histone genes in different cell lines. DNA Cell Biol. 1997;16:1041–1049. [PubMed]
172. Cole F, Fasy TM, Rao SS, de Peralta MA, Kohtz DS. Growth factors that repress myoblast differentiation sustain phosphorylation of a specific site on histone H1. J. Biol. Chem. 1993;268:1580–1585. [PubMed]
173. Sancho M, Diani E, Beato M, Jordan A. Depletion of human histone H1 variants uncovers specific roles in gene expression and cell growth. PLoS Genet. 2008;4:e1000227. [PMC free article] [PubMed]
174. Trollope AF, Sapojnikova N, Thorne AW, Crane-Robinson C, Myers FA. Linker histone subtypes are not generalized gene repressors. Biochim. Biophys. Acta. 2010;1799:642–652. [PubMed]
175. Garcia BA, Busby SA, Barber CM, Shabanowitz J, Allis CD, Hunt DF. Characterization of phosphorylation sites on histone H1 isoforms by tandem mass spectrometry. J. Proteome. Res. 2004;3:1219–1227. [PubMed]
176. Bonet-Costa C, Vilaseca M, Diema C, Vujatovic O, Vaquero A, Omeñaca N, Castejón L, Bernués J, Giralt E, Azorín F. Combined bottom-up and top-down mass spectrometry analyses of the pattern of post-translational modifications of Drosophila melanogaster linker histone H1. J. Proteomics. 2012;75:4124–4138. [PubMed]
177. Su X, Jacob NK, Amunugama R, Lucas DM, Knapp AR, Ren C, Davis ME, Marcucci G, Parthun MR, Byrd JC, et al. Liquid chromatography mass spectrometry profiling of histones. J. Chromatogr. B Analyt. Technol. Biomed. Life Sci. 2007;850:440–454. [PMC free article] [PubMed]
178. Wang L, Harshman SW, Liu S, Ren C, Xu H, Sallans L, Grever M, Byrd JC, Marcucci G, Freitas MA. Assaying pharmacodynamic endpoints with targeted therapy: flavopiridol and 17AAG induced dephosphorylation of histone H1.5 in acute myeloid leukemia. Proteomics. 2010;10:4281–4292. [PMC free article] [PubMed]

Articles from Nucleic Acids Research are provided here courtesy of Oxford University Press