Tài liệu Báo cáo khoa học: Post-translational modifications of the linker histone variants and their association with cell mechanisms docx

REVIEW ARTICLE
Post-translational modifications of the linker histone
variants and their association with cell mechanisms
Christopher Wood
1
, Ambrosius Snijders
2
, James Williamson
2
, Colin Reynolds
1
, John Baldwin
3
and Mark Dickman
2
1 School of Pharmacy and Biomolecular Sciences, Liverpool John Moores University, UK
2 Department of Chemical and Process Engineering, University of Sheffield, UK
3 STFC Daresbury Laboratory, Warrington, UK
Introduction – epigenetic mechanisms
and involvement with disease
Epigenetics is the study of heritable changes in gene
expression that occur without changes in DNA
sequence and, as well as being of fundamental impor-
tance in embryonic development, transcription, chro-
matin structure, X-chromosome inactivation, and
genomic imprinting, it is also now recognized as having
a fundamental role in disease [1]. RNA silencing, DNA
methylation and post-translational modifications
(PTMs) of the core and linker histones are the mechan-
isms that collectively define epigenetics, the latter of
which involve the addition of small chemical groups.

The PTMs that are created by this mechanism include,
but are not limited to, acetylation (lysine), phosphoryla-
tion (serine, threonine), methylation (lysine, arginine),
sumoylation (lysine), and ubiquitination (lysine). Other
epigenetic mechanisms may emerge in the future.
Small RNAs
MicroRNAs (miRNAs) are RNA molecules that are
about 22 nucleotides long and encoded into the
Homo sapiens (hereafter ‘human’) genome [2]. They
Keywords
abundance; acid extraction; cancer; cell
cycle; disease; linker histone; MS;
post-translational modification; PTM;
PTM function
Correspondence
C. M. Wood, School of Pharmacy and
Biomolecular Sciences, Liverpool John
Moores University, Liverpool, UK
Fax: +44 0 51 298 2624
Tel: +44 0 51 231 2565
E-mail:
(Received 10 December 2008, revised 23
March 2009, accepted 30 April 2009)
doi:10.1111/j.1742-4658.2009.07079.x
In recent years, a considerable amount of research has been focused on estab-
lishing the epigenetic mechanisms associated with DNA and the core
histones. This effort is driven by the fact that epigenetics is intimately
involved with genomics in a whole range of molecular processes. However,
there is now a consensus that the epigenetics of the linker histones are just as
important. The result of that consensus is that the post-translational modifi-

cations (PTMs) for most of the linker histone variants in human and mouse
have now been established by a number of experimental techniques, foremost
of which is mass spectrometry (MS). MS was also used by our group to
establish the PTMs of the linker histone variants in chicken erythrocytes.
Although it is now known which types of PTM occur at particular locations
on the linker histone variants, there is still a large gap in the knowledge of
how this data relates to function. The focus of this review is an analysis of
the PTM data for the linker histones from several species, but with an empha-
sis on human, mouse, and chicken. Our analysis reveals that certain PTMs
can be clearly correlated with specific functions of the linker histones in par-
ticular cell types, and that unique PTM patterns exist for different cell types.
Abbreviations
CDK, cyclin-dependent kinase; DNMT, DNA methyltransferase; HDAC, histone deacetylase; miRNA, microRNA; MS, mass spectrometry;
PTM, post-translational modification.
FEBS Journal 276 (2009) 3685–3697 ª 2009 The Authors Journal compilation ª 2009 FEBS 3685
are transcribed by RNA polymerase II into primary
miRNAs and afterwards processed by RNase III Dro-
sha and DGCR8 in the nucleus into precursor miR-
NAs. These precursor miRNAs are then exported by
Exportin-5 to the cytoplasm, where they are further
processed by RNase III Dicer into the mature miR-
NAs [2]. Each miRNA is thought to have many targets
and can bind its target mRNA completely or partially.
If there is complete binding, the mRNA is silenced
and degraded; partial binding leads to downregulation
of a gene. It is known that miRNAs are related to
small interfering RNAs and have similar functions. As
small interfering RNAs have been shown to be
involved with DNA methylation and histone modifica-
tions, it is likely miRNAs operate in the same manner

[2]. The fact that miRNAs are located within the
introns of protein-coding genes has led to the belief
that they are activated with their host genes. A poten-
tial way that this may be achieved is via an active
chromatin hub [3].
DNA methylation
In humans, DNA methylation occurs at a cytosine that
precedes a guanine in a CpG dinucleotide sequence,
most often occurring in short stretches of CpG-rich
regions known as CpG islands. Such regions are about
0.5–2 kb long and can be found in the 5¢-region of
approximately 60% of genes, near to their promoters
[4,5]. The cytosine base is modified at the 5-carbon
position of the pyrimidine ring by the covalent addi-
tion of a methyl group (CH
3
) [5]. This modification is
mediated by DNA methyltransferases (DNMTs) acting
in concert with S-adenosylmethionine, which acts as a
methyl donor in the enzymatic reaction. It is believed
that the pattern of DNA methylation is established in
germline cells through the action of de novo DNMT 3a
and DNMT 3b. This pattern of DNA methylation is
maintained subsequent to DNA replication through
the action of DNMT1. The linker DNA can be prefer-
entially methylated in the absence of H1, but the pres-
ence of the latter will inhibit methylation [6]. CpG
dinucleotides are uniformly dispersed in humans, prob-
ably because 5-methylcytosine can be spontaneously
deaminated to form the DNA base thymidine. CpG

dinucleotides outside islands are essentially continu-
ously methylated, leading to the genes where they
reside being unexpressed. This is a necessary feature,
as there is a large amount of noncoding DNA in the
human genome. However, within CpG islands, the di-
nucleotides can be either unmethylated, if the gene is
expressed, or methylated, if it is not expressed. There
are two exceptions to this rule: imprinted genes, and
genes associated with X-chromosome inactivation will
always have their CpG islands methylated.
Histone modifications
The N-terminal tails of the core histones extend
beyond the nuclesomes and can have their characteris-
tics significantly altered by PTMs. H3 has the greatest
number of modifications currently identified, followed
by H4, H2B, and H2A. The C-terminal tails also con-
tain PTMs, but they are few in number, as are those
for the non-tail regions. Lysine acetylation weakens
electrostatic DNA–histone interations, allowing the
recruitment of factors containing bromodomains such
as SWI ⁄ SNF and TFIID [5]. Methylation of H3
Lys10, H3 Lys28 and H4 Lys21 has been associated
with gene silencing, whereas H3 Lys5, H3 Lys37 and
H3 Lys80 (genomic position numbering) correlate with
actively transcribed genes. It is not only the core
histones that are subject to PTMs; the linker histone
H1 can also be modified (see later).
Epigenetic mechanisms in disease
As specific pathologies (syndromes) can be associated
with problems in the epigenetic machinery, and epi-

genetics is fundamental to chromatin structure, those
diseases have become generically known as diseases of
chromatin. For example, abnormal DNA methylation
can cause errors in genomic imprinting, with an
increased risk of Angelmann’s syndrome [7]. However,
epigenetic problems are also implicated in many more
frequently-occurring diseases, such as cancer.
Many cancer types have been shown to have gains in
methylation at CpG islands in the promoters of some
key genes. Such modifications are associated with tran-
scriptional inactivation [8]. The gains in DNA methyla-
tion, or hypermethylation, are responsible for the
underexpression of tumour suppressors such as
p16INK4a and BRCA1 [5]. Early methylation of DNA
may be a sign of tumorigenesis, as happens to the Wnt
pathway in colon cancer [9], and DNMTs are often
overexpressed in solid and wet cancer types [10]. Muta-
tions and amplification of the androgen receptor gene,
without loss of gene expression, play a key role in the
development of advanced, androgen-independent pros-
tate cancer [4]. Methylation of the androgen receptor
promoter is prevalent in androgen-independent prostate
cancer, but less so in androgen-dependent prostate can-
cer [4]. As well as hormonal genes, cell cycle genes are
also affected in prostate cancer; an example is the
methylation-mediated inactivation of the CDKN2A
gene [4]. Methylation also goes awry in haematopoietic
PTMs of linker histones C. Wood et al.
3686 FEBS Journal 276 (2009) 3685–3697 ª 2009 The Authors Journal compilation ª 2009 FEBS
malignancies, and hypermethylation of p16INK4a has

been observed in non-Hodgkin’s lymphoma, multiple
myeloma, and acute lymphocytic leukaemia [8].
It is widely accepted that DNA methylation should,
in the right circumstances, be a target for clinical treat-
ment. Accordingly, nucleoside inhibitors that inhibit
DNA methylation, such as azacitidine, decitabine, and
zebularine, have been developed. All three are cytidine
derivatives that irreversibly inhibit DNMTs [11]. As
decitabine contains a deoxyribose group, it is incorpo-
rated into DNA [12]. However, because azacitidine
contains a ribose group, it is initially incorporated into
RNA [12]. Incorporation into DNA occurs when aza-
citidine is converted into 5-aza-2¢-deoxycytidine
diphosphate by ribonucleotide reductase, which is then
phosphorylated, the triphosphate form being incorpo-
rated into DNA in place of the natural base cytosine
[12]. The use of such analogues results in the global
depletion of DNMTs and a subsequent reduction in
DNA methylation.
Although DNA methylation is the most studied epi-
genetic modification in terms of clinical diagnostics,
the mechanism is also important for histone modifica-
tions. DNMTs can interact with histones in two ways.
First, DNA methylated by DNMT can attract proteins
such as MeCP2 that are able to recruit histone deacet-
ylases (HDACs); and, second, DNMTs can themselves
directly recruit HDACs to help silence gene expression
[4]. Most of the literature on interactions with methy-
lated DNA has centred on the core histones H2A,
H2B, H3, and H4, but a complete picture of epigenetic

modifications cannot be obtained until linker histone
PTMs have been factored in. This review analyses the
research effort expended thus far on linker histone
PTMs. For consistency, amino acid positions in
a sequence are referred to by their actual genomic
position, as given in Swiss-Prot and similar databases.
Structure and function of the linker
histone variants
Location and structure of the linker histones
Historically, the location of the globular domain of the
linker histone has been a matter of contention [13].
Currently, although there is a good degree of agree-
ment about the overall parameters of the fibre formed
by folding the zig-zagging chain of nucleosomes in
inactive chromatin, the location of the linker histone
in relation to the nucleosome core particle and linker
DNA is still not known to high resolution (Fig. 1).
However, recent studies [14–16] suggest that the linker
histone is close to the dyad axis of the core particle at
the entry and exit of the DNA. Similarly, the geometry
of one nucleosome in the fibre relative to a DNA-con-
nected nucleosome is also unknown [17–20].
The structure of the linker histone H1 in humans is
characterized by a relatively short N-terminal tail, a
longer C-terminal tail, and a conserved globular
domain [21]. This model extends to most other organ-
isms, two exceptions being Tetrahymena thermophila
and Saccharomyces cerevisiae [22]. The linker histone
H1 variants show the greatest diversity when compared
to the core histones. There is also a great diversifica-

tion in the H1 variants within a single species such as
H. sapiens, predominantly in the N-terminal and C-ter-
minal tails, with, as stated, a conserved globular
domain. However, when similar H1 variants are com-
pared between species, there is a remarkable similarity.
The nearer the species, the less is the divergence, such
that H1.2 in Pan troglodytes has just one amino acid
difference from its human counterpart, and the H1.4
variant in the human and Mus musculus (hereafter
‘mouse’) genomes has 93.6% sequence identity
(Fig. 2). The reason for this is that the H1-variant
genes within a species are paralogues, originating from
gene duplication events, whereas the same H1 gene
between species is an orthologue, originating from an
ancestral gene [23]. In humans, the variants consist of
the following: the somatic subtypes, H1.1–H1.5; a
spermatogenesis subtype, H1t; an oocyte-specific
subtype, H1oo; and a replacement subtype, H1
o
. H1.1–
H1.5, along with H1t, are known as the replication-
independent group, and are mainly expressed in
S-phase. The remaining two, H1oo and H1
o
, are known
as the replication-dependent variants. H1.1–H1.5 and
Fig. 1. The possible locations of the linker histone in relation to the
nucleosome core particle. The globular domain of the linker histone
will be located either symmetrically (left image), or asymmetrically
(right image) [14,15]. Colour assignments are as follows: magenta,

nucleosome core particle; blue, 146 bp of DNA; red, globular
domain of linker histone H1; green, 22 bp of linker DNA; orange,
11 bp of linker DNA.
C. Wood et al. PTMs of linker histones
FEBS Journal 276 (2009) 3685–3697 ª 2009 The Authors Journal compilation ª 2009 FEBS 3687
H1t reside on the short arm of chromosome 6; H1oo
is located on chromosome 3 and H1
o
on chromosome
22. H1.2 and H1.4 predominate in most cell types. The
affinity of the various types seems to depend on their
C-terminal tails. H1.1 and H1.2, with the shortest
C-terminal tails, a low density of positively charged
residues, and the lowest number of cyclin-dependent
kinase (CDK) sites, have the lowest affinity for chro-
matin. A CDK site is identified by the consensus
sequence (S ⁄ T)PXZ, where X is any amino acid and Z
is a basic amino acid. H1.4 and H1.5, with longer
C-terminal tails and more than two (S ⁄ T)PXZ sites,
have the highest affinity for chromatin. With the high-
est content of positively charged residues, H1.3 has an
intermediate affinity for chromatin [23]. The precise
functions of the H1 variants within a cell are only just
starting to be elucidated.
Functions of the linker histones
It was evident from work with unicellular organisms
that the linker histones were not critical for growth
and cell division [23–25]. Following these experiments,
it was speculated that the H1 variants were not global
repressors of transcription, and this has now been

shown to be the case [25]. Depletion of H1 in mam-
mals causes significant changes to chromatin structure.
When chromatin is depleted of H1, there is a reduction
in the nucleosome-repeat length globally and a reduc-
tion in local chromatin compaction [25]. The reduction
in repeat length arises from having fewer than one
linker histone per nucleosome [25]. Depletion of H1 in
mammals also causes a reduction in H3 Lys28 acetyla-
tion, with a smaller reduction in H3 Lys28 trimethyla-
tion, and also leads to a reduction of methylation at
CpG islands in some of the H1-regulated genes [25].
The H1 variants tend to associate with specific tran-
scriptional regulators [23]. For example, H1.1 specifi-
cally associates with BAF, which regulates chromatin
structure [23], and H1.2 has been shown to associate
with p53 [26]. It is thought that the specificity of indi-
vidual variants stems partially from their sequence
diversity, but mostly from PTMs [23]. Thus, the evi-
dence emerging is that the H1 variants have specific
functions. First, individual H1-variant knockout mice
gave rise to specific phenotypes, with distinct effects on
gene expression and chromatin structure [27]. Second,
in the knockout mice referred to, there was no equal
upregulation of the remaining variants, with only par-
ticular variants being able to compensate. Third, there
are differences in the localization of the H1 variants
within the nucleus, and there are variations in their
relative amounts between different cell types [28].
Fourth, the H1 variants have different affinities for
chromatin, can be recruited to specific transcription

factors, and, as we shall see, have particular PTM
patterns.
Significantly, H1.2 is now associated with an extra-
nuclear function. This variant will, upon a DNA dou-
ble-strand break, translocate to the cytoplasm and then
permeabilize the mitochondrial membrane, causing the
release of apoptotic compounds [29–31]. It has been
shown that H1.2 in a cell infected with a virus displays
an increase in mobility [32], and this property may play
an important role in the treatment of cancer [33].
A specific mutation in H1.4 was detected in Raji
cells, but was not detected in 103 healthy individuals
or other Burkitt’s lymphoma cell lines [34]. This could
ARMCX4_HUMAN
H1FX_HUMAN
H1F0_HUMAN
HIST1H1T_HUMAN
HIST1H1A_HUMAN
HIST1H1B_HUMAN
HIST1H1C_HUMAN
HIST1H1E_HUMAN
HIST1H1D_HUMAN
H1fx_MOUSE
Hist1h1t_MOUSE
Hist1h1a_MOUSE
Hist1h1b_MOUSE
Hist1h1c_MOUSE
Hist1h1e_MOUSE
Hist1h1d_MOUSE
XP_899364_MOUSE

Fig. 2. Phylogeny tree of human and mouse linker histones. Speciation events are indicated by blue dots and gene duplication by red dots.
HIST1H1C and HIST1H1E are the genes that code for human linker histones H1.2 and H1.4, respectively. Note how the human HIST1H1C
and HIST1H1E genes have a last common ancestor that is a duplication node, which makes these two genes paralogues. However,
HIST1H1C in human and mouse originate from a speciation node, and are therefore orthologues. The phylogeny tree was generated by
TREEFAM [69].
PTMs of linker histones C. Wood et al.
3688 FEBS Journal 276 (2009) 3685–3697 ª 2009 The Authors Journal compilation ª 2009 FEBS
be an example of H1 sequence variation acting as a
marker for a particular phenotype. However, the main
aim of Sarg et al. [34] was to demonstrate the use of a
particular chromatography technique, rather than to
firmly establish H1 sequence variation with disease.
Thus far, then, no in-depth analysis has been per-
formed that attempts to correlate linker histone
sequence or PTM variation with disease. This is not
surprising, especially in the case of the latter, as there
is a potential for many permutations. Nevertheless,
this must be the next phase in the work on H1 PTMs.
H1
o
is a general differentiation-dependent linker his-
tone, and has a similar sequence to the avian H5 vari-
ant. H1
o
will accumulate in a cell, reaching a peak at
terminal differentiation, being initially synthesized in
oocytes and early embryos [35].
Epigenetic control of linker histones was discovered
relatively early, with studies of synchronously dividing
nuclei in the plasmodia of Physarum polycephalum

[36], when it was shown that phosphorylation of linker
histones is strongly implicated in cell cycle control and
that phosphorylation is a precursor of mitosis. It is
now widely accepted that patterns of PTMs on the
core histones can influence transcriptional activity. For
example, acetylation of H3 Lys10 has an inverse rela-
tionship with the amount of DNA methylation [37,38].
As it is also known that the linker histones can affect
DNA methylation [25,39], it is not unreasonable to
conclude that there must be key PTM patterns that
govern – or at least significantly contribute to – the
function of H1, as they do in the core histones.
Although much work has been done to identify specific
PTMs in the linker histones, there is still a gap in our
knowledge of how these affect function.
Phosphorylation of CDK and non-CDK
consensus sites
A PTM pattern for mitosis?
Early work on identifying the sites of phosphorylation
in core and linker histones indicated that there was no
correlation between cell cycle status and the number or
location of phosphorylation sites [40]. Later work, how-
ever, has shown that this is not the case [41], and that
the H1 linker histone of human lymphoblastic T-cells
has phosphorylation states that correlate with the inter-
phase and mitosis stages of the cell cycle. During inter-
phase, it was found that the H1.5 variant was
phosphorylated at Ser18, Ser173 and Ser189, which all
reside in a CDK consensus motif of the form (S ⁄ T)PXZ,
as previously defined. It was found that, during mitosis,

the same three serine phosphorylations were present,
but were also accompanied by phosphorylations at
Thr11, Thr138, and Thr155. The first of these three
threonines is not located within a TPXZ consensus
sequence, but the latter two are. The same pattern of cell
cycle dependency of phosphorylations was found in the
linker histone variants H1.2, H1.3, and H1.4. So, for all
tested linker histone variants, it was established that
only serines were phosphorylated during interphase, but
in mitosis, threonine residues were additionally phos-
phorylated. It was found that, during interphase, the
human lymphoblastic T-cells had a proportion of H1.5
molecules monophosphoryated at a particular residue
and a smaller proportion that was monophosphorylated
on another residue. It was also found that the ratio of
these two subgroups of H1.5 occurred in other cell
types. During mitosis, it was found that H1.5 existed as
two species with five phosphorylations either on Thr11,
Ser18, Thr138, Ser173, and Ser189, or on Thr11, Ser18,
Thr155, Ser173, and Ser189. Therefore, it was concluded
that Thr138 and Thr155 of H1.5 can never be phosphor-
ylated at the same time. There is support for this
hypothesis [42], where the only phosphorylations to be
found on H1.5 were at Thr138 and Thr155. If these two
modifications occurred at the same time, it would be
reasonable to expect that they would be found in equal
abundance. However, whereas the H1.5 peptide with a
phosphorylation on Thr155 was readily detected, that
with a phosphorylation at Thr138 could only be
detected after methanolic HCl was used to convert car-

boxylic groups to their corresponding methyl esters.
Thus, the suggestion is that H1.5 exists as two separate
species, with either Thr138 or Thr155 phosphorylated,
but not both. Wisniewski et al. [43] (Table 1) identified
phosphorylation at Thr138 on H1.5, but not at Thr155,
even though, like Sarg et al. [41] and Garcia et al. [42],
they used HeLa cells. Wisniewski et al. [43] agree with
Sarg et al. [41] that Ser18 of H1.5 is phosphorylated, but
could find no such modifications at Thr11, Ser173 or
Ser189 in human or mouse tissue.
Correlation of N-terminal tail PTMs with function
Sarg et al. [41] could not detect any N-terminal tail
phosphorylations for H1.2, H1.3 and H1.4 for cells that
were in the interphase part of the cell cycle. The reason
given is that there are no (S ⁄ T)PXZ motifs in the N-ter-
minal tails of those variants. This is in conflict with the
work of Garcia et al. [42], who found phosphorylations
on H1.2 at Thr31 and Ser36, on H1.3 at Ser37, and on
H1.4 at Ser2, Thr4, Thr18, Ser27, and Ser36. In com-
paring the work of Wisniewski et al. [43] (Table 1) and
Garcia et al. [42], it can be seen that: H1.4 is phosphory-
lated at Thr18, but not on Ser2 or Thr4; H1.3 is
C. Wood et al. PTMs of linker histones
FEBS Journal 276 (2009) 3685–3697 ª 2009 The Authors Journal compilation ª 2009 FEBS 3689
similarly phosphorylated at Ser37; H1.2 is not phos-
phorylated at Thr31. Thus, it is possible to conclude
that phosphorylation of the N-terminal tails of the H1
variants does occur, but why do some researchers detect
them but others do not? Before addressing this question,
there is also the issue as to what variety of PTMs occur

in the N-terminal tails. Sarg et al. [41] found that only
H1.5 was modified in the N-terminal tail in human cells;
more recent work by Wisniewski et al. [43] and Snijders
et al. [44] (Table 1) has, excluding the N-terminus acety-
lations, identified eight N-terminal tail PTMs in cultured
human cells and seven in Gallus gallus (hereafter
‘chicken’) erythrocytes, respectively. Although the over-
all number of these modifications is low, the density of
modifications is much the same as in the rest of the lin-
ker histones. It is the shortness of the N-terminal tails
Table 1. Alignment of chicken, mouse and human PTMs. Each PTM-containing sequence in humans has been aligned with the similar
sequence in the other two species, which may or may not contain a PTM. Symbols: a, acetylation; d, deamidation; f, formylation; m, methyl-
ation; p, phosphorylation; u, ubiquitination; 2m, dimethylation; 2m ⁄ f, dimethylation and ⁄ or formylation; a ⁄ m, acetylation and ⁄ or monomethy-
lation; a ⁄ f, acetylation and ⁄ or formylation; 2m ⁄ f, dimethylation and ⁄ or formylation. The ‘a-’ in the second column (first PTM location) refers
to N-terminal acetylation. The data for human and mouse were taken from [43], and the data for chicken were taken from [44].
Chicken
H101 a-STAAPP AmKA K A K A T K K K 2m ⁄ fKK dNK
H110 a-STAAPA AK A K A K AT K KK 2m ⁄ fKK dNK
H102 a-STAAPS AK A K P K ATK KK 2m ⁄ fKK dNK
H103 a-A pTAAPA AK A K A K ATK KK 2m ⁄ fK 2mK dNK
H11L a-STAPAA AK A K A K AT K KK 2m ⁄ fKK dNK
H11R a-A
pTA – A A A aKA K A K AT K KK 2m ⁄ fKK dNK
H5 a-pT pSA pS– P A – A amKR pS pST A K Q 2m ⁄ fKN D R
Mouse
H1.0 a-TNpSA aK– –––K KS DSA KQK E DK
H1.1 a-pS pTAASaKPaK mKA K K A pSQ K ufKK a ⁄ mKN aK
H1.2 a-pSAAAAaKAKKmKR a ⁄ mK pS pSK aK ufKK a ⁄ mKN afK
H1.3 a-STAAP2mK pTKKT Ra ⁄ mK pS pS ua ⁄ mK a
K ufKK a ⁄ mKN afK

H1.4 a-pS pTAAPK pTKKK RmK pS pS a ⁄ mK aK ufKK a ⁄ mKN afK
H1.5 a-pS pTA E P K pSKKK mR mK pTS K K K K mKNK
Human
H1.0 a-TNSA–– –––K KS DSA KQK E DK
H1.1 a-pST V A S K P aKK P K K A S Q KfK aKK NK
H1.2 a-pSTAAAaKAKKA RaK pSS auK aK fK aKK NaK
H1.3 a-STAAPaK pTKKK RaK pSS auK aK fK a
KK NaK
H1.4 a-STAAPaK pTKKA RaK pSS auK aK fK aKK NaK
H1.5 a-pST A E P aK pSKKK RK TSaK K K K K N afK
Chicken
H101 aKKR RTAaKK pSKKTKK aKS A K S K
H110 aKKR RPAaKK pSKKTKK aKS T K S K
H102 aKKK KPAaKK pSKKTKK aKS T K S K
H103 aKKR KPAaKK pSKKTKK aKS T mKSK
H11L aKKR KSAaKK pSKKTaKK K S T a ⁄ mKS K
H11R K K R K S A aKK pSKKTaKK K S – a ⁄ mKS K
H5 A KR G– STKAKKTKA KSaKK T A
Mouse
H1.0 T K R K A V A – A K A K K K A T K V K –
H1.1 uKKK uK aKAA–TKK–KK KSKK SK
H1.2 aKKmfKQ G A T pTT K aKA aKK K S aKS – –
H1.3 aKKmfKKGATTT KKAaKK KSKK SK
H1.4 aKKmfKK G A A T T aK aKA aKK KSKK pSK
H1.5 aKKK KGTA–TKaKA aKK KSKK SK
Human
H1.0 T K R F K A A – T A K – K K A T K A K K
H1.1 a ⁄ fK aKK – KST–TKK–RK KNKK SK
H1.2 a ⁄ fK aKK K G A A pTT K K pT aKK K pSK K S aK
H1.3 a ⁄ fK aKK K G A A pTT KKAaK a ⁄ mKK S K K S K

H1.4 a ⁄ fK aKK K G A T pTTKKAaK a ⁄ mKK pSK K
pSK
H1.5 K K K K G pTKGT KK–aK a ⁄ mKK S K K S K
PTMs of linker histones C. Wood et al.
3690 FEBS Journal 276 (2009) 3685–3697 ª 2009 The Authors Journal compilation ª 2009 FEBS
that accounts for the low numbers. Therefore, it is now
possible to say that the N-terminal tails do, in fact,
contain a range of different types of PTM.
Returning to the issue of abundancy, Garcia et al.
[42] had to use two techniques to increase the number
of peptides with certain PTMs. First, protein digests
were treated with propionylation reagent to convert
monomethylated and endogenously unmodified amino
groups on the side chains of lysine residues and N-ter-
mini to propionyl amides. Second, it was found that
certain phosphorylated peptides (predominantly origi-
nating from the N-terminal tail) were of such low abun-
dance that, in order to obtain stronger spectra, they
were subjected to enrichment by immobilized metal
affinity chromatography [45]. Prior to using this tech-
nique, Garcia et al. [42] converted the carboxylic
groups to methyl esters with the use of methanolic
HCl. This modification decreases the strength of bind-
ing of nonphosphorylated linker histones to the immo-
bilized metal affinity chromatography column. They
can then be washed off before eluting the phospho-
linker histones. It should be noted that the practice of
methyl esterification is currently not widely used, owing
to problems with side reactions [46]. Deterding et al.
[47], who only analysed the H1.4 linker histone variant

in human and mouse tissue, found that, in both species,
only Thr18 in the N-terminal tail region was phosphor-
ylated. However, the signal for this in the human tissue
was so noisy in comparison with signals for phosphory-
lated residues in the C-terminal tail that confirmation
could only be obtained by reference to the mouse signal,
which was less noisy. With high-mass accuracy mass
spectrometers now readily available, this should be less
of a problem in the future. We can perhaps, then,
hypothesize that although N-terminal tail modifications
of linker histones do occur, they may be less abundant
than those of the globular domain and C-terminal tail.
The degree to which they are less abundant remains to
be established, as does the biological significance of that
fact.
Correlation of C-terminal tail PTMs with function
Sarg et al. [41] found that H1.2, H1.3 and H1.4 had
fewer phosphorylations in the C-terminal tail region
than H1.5. In H1.2, Ser173 is phosphorylated, as are
Ser189 in H1.3 and Ser172 and Ser187 in H1.4. Garcia
et al. [42] found that H1.2 was phosphorylated at
Ser173, Thr146, and Thr154. It is worth noting that
these two latter modifications occur in TPXZ motifs
and, if Sarg et al. [41] is correct, may reflect the fact that
the cells are in mitosis. The H1.3 variant in Garcia et al.
[42] was found to be phosphorylated at Ser189, Thr147,
Thr155, and Thr180. The latter threonine is in a non-
CDK consensus site, and seems to be an anomaly. The
occurrence of phosphorylated Thr147 and Thr155 in
the C-terminal tail of H1.3 probably has the same

explanation as their occurrence in H1.2. Deterding et al.
[47] identified the same H1.4 C-terminal tail phosphory-
lations in human and mouse tissue as Sarg et al. [41].
As can be seen from Table 2, Wisniewski et al. [43]
detected no phosphorylation on Ser172 of H1.4. For
H1.2 and H1.3, the work of Wisniewski et al. [43] agrees
with that of Garcia et al. [42], noting that in Table I of
the former paper the phosphorylation on Thr173 of
H1.2 is a typographical error (should be Ser174).
Table 2 lists the phosphorylations that have been
detected more than once in the research described
above. It therefore represents those sites that are most
likely to be modified at reasonable levels of abundance.
Most mass spectrometry (MS) analysis of PTMs has
been performed on cultured cell lines. It has been
shown that methylation can be readily detected in tis-
sue, but is extremely rare in cultured cells [43]. Other
potential problems with cultured cells are discussed
later.
Analysis of other PTMs
It is now accepted that acetylation and methylation of
the core histones are key regulators of transcription.
Although phosphorylation of the linker histones has
attracted the most attention, recent results from vari-
ous MS analyses have shown that acetylation and
methylation are also key modifications of the linker
histones.
Lysine and N-terminus acetylations
Acetylation of the Na-terminus involves the cotransla-
tional cleavage of a methionine, followed by acetyla-

tion of the second residue of the N-terminal tail.
Initial, non-MS work showed that only H1.0 in human
cells and H5 in avian cells existed in forms that had
the Na-terminus both acetylated and unacetylated
[48,49]. However, it has now been shown that the
Table 2. Phosphorylations of the human linker histones. Those
modifications that have been identified more than once in the
papers referred to are shown. The numbers in parentheses refer to
the appropriate references.
H1.2 (P16403) H1.3 (P16402) H1.4 (P10412) H1.5 (P16401)
pS36 [42,43],
pT146 [42,43]
pS37 [42,43],
pT147 [42,43]
pT18 [42,43,47],
pS172 [41,42,47]
pS18 [41,43],
pT138 [41–43]
pS173 [41–43] pS189 [41–43] pS187 [41–43,47] pT155 [41,42]
C. Wood et al. PTMs of linker histones
FEBS Journal 276 (2009) 3685–3697 ª 2009 The Authors Journal compilation ª 2009 FEBS 3691
former modification not only occurs on all the linker
histone variants in human and chicken [42–44], but
that is also the most abundant type of acetylation.
Linker histones do exist with their Na-termini
unacetylated, but are significantly fewer in number.
Although the function of Na-terminus acetylation is
unclear, it was noted in earlier work [48,49] that
whereas the ratio of acetylated to unacetylated Na-ter-
mini remained constant in avian H5 erythrocytes from

newly hatched and adult chickens, it increased for
H1.0 in ageing rat tissues. As H1.0 is associated with
differentiation and is most abundant in terminally dif-
ferentiated cells, there may well be a correlation
between Na-terminus acetylation and differentiation.
Not all methionines are cotranslationally cleaved and
can therefore become acetylated. This process is not
widespread, but it has been shown to be present in
recent work [43,44] (Table 1).
Garcia et al. [42] found that H1.2, H1.3 and H1.4 in
human cells all had just one site of lysine acetylation,
and on the same residue, Lys64 (or Lys65, depending
on the variant). Considerably more acetylations – up
to nine – were found in H1.4 [43] (Table 1). The glob-
ular domain was found to contain the largest number
of acetylations: Lys52, Lys64, Lys85 and Lys97; all of
these are thought to be involved with DNA binding
[43]. The abundance of lysine methylation can be
attributed to the fact that certain types of human cell
were rapidly proliferating. In mouse tissue, the spleen
was found to contain the most acetylations, because
lymphopoiesis is associated with rapid cell division. In
mouse tissue containing mostly differentiated cells, e.g.
liver, the number of acetylations was much lower.
Lysine methylation
Methylation of lysine in linker histone proteins has
been reported in human HeLa cells [42,43], although
there is a difference in the number of identified sites.
Garcia et al. [42] found that, in H1.4, Lys26 and Ser27
were simultaneously methylated and phosphorylated,

respectively. The point is made that the aforemen-
tioned residues occur in the sequence KARKSAGA
(residues 23–30), which is similar to one found in the
core histone H3 (VARKSAPA, residues 25–31).
Within H3 there are well-known adjacent methylation
and phosphorylation sites at Lys9-Ser10 and Lys27-
Ser28 that are involved with transcription. Thus, the
same argument is made for H1.4, by virtue of it, too,
having adjacent methylation and phosphorylation sites.
There is support for these assertions [49]; however,
Wisniewski et al. [43] (Table 1) found no methylations
in this region of H1.4, or of the other variants. Puta-
tive sites of methylation were identified at Lys169 in
H1.4 and H1.5, or Lys170 in H1.3. In mouse, the H1.4
variant has no modifications at Lys26, and at posi-
tion 27 there is an alanine, rather than a serine [43].
Ubiquitination and formylation
For the first time, ubiquitinations of the histone lin-
ker protein were identified by MS [43] (Table 1). It
was found that Lys46 was ubiquitinated in H1.2,
H1.3 and H1.4 in human HeLa cells, but not in
MCF7 cells. In mouse tissue, Lys116 of H1.1 and
Lys46 of H1.2 and H1.3 in the spleen were the only
sites of ubiquitination. The fact that both cell lines
were cultured in the same growth medium, and have
the same doubling time, increases the probability that
these ubiquitinations are unique for HeLa cells.
Another putative novel modification found is that of
formylation [43]; H1.1, H1.2, H1.3, H1.4 and H1.5 in
human MCF7 cells were all found to be formylated.

Whereas H1.5 was uniquely formylated on Lys88, the
others were similarly modified on Lys90 (H1.2 num-
ber). In mouse tissue, the most frequently occurring
formylation site was Lys63. Formylation of lysines
has been shown to arise as a result of oxidative dam-
age to DNA [50]. Snijders et al. [44] (Table 1) identi-
fied a single site of lysine dimethylation at Lys71, but
were unable to distinguish between dimethylation and
formylation.
Perturbation of phosphorylations
by external mechanisms
It has been clearly shown in several studies that phos-
phorylation can be imposed by external influences
[41,47,51]. This is an important phenomenon, and
means that those processes will be able to influence the
cell cycle.
Garcia et al. [42] found that growing T. thermophila
cells had site-specific higher levels of phosphorylation
than when they were being starved. Phosphorylated
Thr47 was enriched in growing cells by a factor of
seven as compared with starved cells. Similarly, phos-
phorylated Thr35 was also found to be enriched by a
factor of four in growing cells. It is perhaps impor-
tant that these two residues occur in (S ⁄ T)PXZ motifs
(as defined). It was found that in Drosophila melanog-
aster embryos, phosphorylated Ser11 was associated
with mitosis and that the proportion of this post-
translational modification decreased as those embryos
aged [52]. These experiments clearly show that the
amount of phosphorylated H1 is a function of cell

activity. Villar-Garea and Imhof [52] concluded that,
PTMs of linker histones C. Wood et al.
3692 FEBS Journal 276 (2009) 3685–3697 ª 2009 The Authors Journal compilation ª 2009 FEBS
in mammalian cells, phosphorylation in mitosis only
occurs in the N-terminal tail. However, it has been
shown that, during mitosis, phosphorylations also
occur in the C-terminal tail, and in (S ⁄ T)PXZ motifs
[41].
Deterding et al. [47] analysed human UL3 cells
(derived from the osteosarcoma cell line U2OS) treated
with dexamethasone, CVT313, or CGP74514 (dexa-
methasone is a synthetic glucocorticoid used in the treat-
ment of autoimmune diseases; CGP74514 and CVT313
are CDK1 and CDK2 inhibitors, respectively), and
looked at the phosphorylation state of the linker
histones using MS. It was found that treatment with all
of these compounds reduced the global level of phos-
phorylation of the H1.2 and H1.4 isoforms. Although
the work did not go so far as to establish site-specific
patterns of phosphorylation related to compound and
dosage, it did establish, by the use of antibodies, that the
level of phosphorylated Thr18 in the N-terminal tail of
H1.4 was reduced by treatment with any one of these
three compounds.
In the examples discussed, extensive use was made of
cultured cells. There is, of course, nothing wrong with
this in the substantial majority of cases. However,
culturing cells may have an impact on overall PTM pat-
terns. In particular, differences may arise in the struc-
tural and biochemical properties of a cultured cell (and

hence PTM patterns), particularly when the cells are
grown on a monolayer 2D medium. Normal cells grown
in such a medium can display a nuclear structure that is
different to their in vivo structure [53,54]. Use of a 3D
culture medium better mimics the extracellular matrix
[53], and the cells should therefore have a nuclear struc-
ture that is more representative of the in vivo structure.
If the nuclear structure of cultured cells can be altered,
then there will be a concomitant change in the biochem-
istry of those cells [54].
Existence of global PTM patterns
in different cell types
The strong evidence emerging is that specific PTM pat-
terns occurring on DNA and particular sets of proteins
can be correlated with cell type. The inference from
this is that there will be a change in a cell’s PTM pat-
tern when it progresses from a normal to diseased
state, and that, accordingly, such changes can be
detected and made the target of clinical intervention
[55,56]. However, although changes in the PTM pat-
terns of particular proteins between normal and dis-
eased cells have been detected [55,56], can the concept
be taken to the lower level of chromatin? This has
already been shown to be the case in three sets of
mouse cells [57]. A proportion of murine embryonic
stem cells, embryonic fibroblasts and embryonic carci-
noma cells were grown in standard cell growth med-
ium, with the remainder having trichostatin A, an
HDAC inhibitor, added, the aim being to mimic dis-
ease-induced hyperacetylation of histones. Two

changes were detected: (a) PTM patterns alter for ‘dis-
eased’ cell lines; and (b) those cell lines have unique
and specific PTM patterns. The PTMs that were being
monitored resided on the H3 and H4 core histones.
However, there is nothing to suggest that the linker
histones should not display the same global property,
and such changes involving just phosphorylation were
discussed in an earlier section. The process of disease-
induced alteration of global PTM patterns has also
been observed in human colon adenocarcinoma cell
lines [58]. MS is of fundamental importance when it
comes to detecting combinations of PTMs on a single
protein. This has been demonstrated on human embry-
onic stem cells [59].
Table 1 shows the PTMs detected in human,
mouse and chick cells [43,44]. From the data for
chick cells, it can clearly be observed that six of the
linker histones have identical PTMs at amino acids
71, 84, 147, and 189. Unlike human linker histones,
the chick variants have very similar sequences, and
it would be easy to dismiss this observation with the
argument that near-identical sequences will inevitably
have the same PTM pattern. However, this line of
argument would ignore two important facts. First,
the chick linker histones can – like their human
counterparts – be associated with specific and differ-
ent functions; and, second, the cells, being erythro-
cytes, are terminally differentiated. It is therefore
possible to say that the PTM patterns in Table 1 for
chick cells can be identified as being unique for

terminally differentiated chick erythrocyte cells. How-
ever, as previously mentioned, although MS can
detect many modifications, it does have restrictions,
such as difficulty in distinguishing PTMs that have
near-identical masses [44].
It was mentioned earlier that mouse tissue with the
higher replication rate has higher levels of linker his-
tone acetylation. This can be taken as evidence that
unique linker histone PTM patterns also exist in live
tissue, and not just in cultured cells [43]. It is possible
to come up with a list of PTMs that are either absent
in MCF7 cells and present in HeLa cells, or present in
MCF7 cells but missing in HeLa cells (Table 3). As
mentioned earlier, the two cell lines were grown in the
same media, so it is clearly possible to distinguish the
two human cell lines by comparison of the PTMs on
their linker histones.
C. Wood et al. PTMs of linker histones
FEBS Journal 276 (2009) 3685–3697 ª 2009 The Authors Journal compilation ª 2009 FEBS 3693
Conclusions
The evidence accumulating from MS and other bio-
physical experiments considerably strengthens the
hypothesis that not only can the linker histone vari-
ants be associated with specific functions, but PTMs
thereon can also uniquely identify particular cell
types. Indeed, this is now becoming the accepted par-
adigm [60]. Those functional capabilities even, as in
the case of H1.2, have an extranuclear reach. PTMs
modulate the range of functions covered by the linker
histone variants and, by analogy with the core hi-

stones, each of those functions will have a distinct
PTM signature.
The extraction of the H1 variants from cells or tis-
sue has the potential to alter PTM states. It is there-
fore necessary that gentle procedures should be used.
Acid extraction of H1, although efficient, can instigate
the reversal of labile PTMs, such as histidine phos-
phorylation [61–63]. A range of different acids have
been used to extract linker histones, including per-
chloric acid [43,52], sulfuric acid [42,64], and hydro-
chloric acid [65]. Extraction by salt is gentler and just
as efficient at isolating H1 histones [44,63]. Although it
is the extraction of the linker histones from tissue and
cell cultures that has a high potential to alter PTMs,
purification of the extracts can be considered to be a
benign step in the process of isolation. Purification of
histones in general can involve a myriad of processes,
and these have been discussed in detail elsewhere [66].
It was found in one case that phosphorylation of
threonines in H1.5, namely Thr138 and Thr155, was
associated with cells in mitosis [41]. Some support was
provided for this principle [42], where both of these
residues were found to be phosphorylated, although
Thr138 only with some difficulty. In another case, it
was found that only Thr138 was phosphorylated [43].
There is a clear conflict here, so is it possible to distin-
guish the respective cases? In the first case, the cell
lines were specifically treated to put them into mitosis;
this was not so in the second and third cases. How-
ever, in two cases [42,43], it seems unlikely that there

would have been a significant number of cells in
mitosis. In addition, in MS experiments, absence of a
condition is not proof of its nonexistence.
Whereas, initially, it was found that PTMs in the
N-terminal tail of most of the H1 variants did not
occur – an exception being H1.5 [41] – it is now clear
that there are, in fact, numerous modifications,
although they seem to be less abundant [43,44]. Acety-
lation of the N-terminus of H1 is the most abundant
modification, although it has been shown that the un-
acetylated form does exist [43,44]. There seems to be no
consensus on the significance of N-terminus acetylation.
However, as the amount of H1.0 with an acetylated
N-terminus has been observed to increase in ageing rat
tissue [48,67], and given that H1.0 is most abundant in
terminally differentiated cells, there may be a link
between N-terminus acetylation and differentiation.
From work on human cells and mouse tissue [43], it
can be clearly seen that the amount of acetylated H1 is
a function of the replication rate, with most acetyla-
tions occurring in rapidly replicating tissue, and the
least in the most slowly replicating tissue. Confirma-
tion of this comes from work on chicken erythrocytes
[44], where it was found that there are relatively few
acetylations in the chicken H1 variants. This is because
the erythrocyte sample material comprises cells that
are largely terminally differentiated.
Phosphorylation is correlated with growth rates [64]
and can be significantly increased. The addition of
compounds that influence the cell cycle will cause

changes in the levels of phosphorylation of the linker
histone isoforms [47].
Particular cell lines can be identified with particular
patterns of PTMs on the core and linker histones [43],
and variations in those patterns – having been associ-
ated with oncogenic progression [68] – are primary
candidates for pharmacological intervention.
Taken as a whole, the data from the experiments
discussed herein clearly show that it is possible to asso-
ciate specific PTM patterns in the linker histones with
particular functions, and that unique patterns of PTMs
exist for diseased cells when compared with normal
cells, and between cells of different types.
Table 3. PTMs that uniquely identify human MCF7 from HeLa
cells. ‘+’ indicates a modification that is present on MCF7 linker
histones but is missing from HeLa linker histones. ‘)’ indicates a
modification that is missing from MCF7 linker histones but is pres-
ent on HeLa linker histones. The symbols in italics are as defined
for Table 1.
MCF7 PTMs compared with HeLa PTMs
H1.5: +aK17
H1.1: )aK22
H1.2, H1.3, H1.4: +aK34
H1.2, H1.3, H1.4: )uK46
H1.2, H1.3, H1.4: )aK52
H1.1, H1.2, H1.3, H1.4: )aK64
H1.1, H1.2, H1.3, H1.4: )aK85
H1.5: +a ⁄ fK88
H1.1, H1.2, H1.3, H1.4: )fK90
H1.1, H1.2, H1.3, H1.4: +aK97

H1.2: )pT146
H1.2: +pT165
H1.2: +aK169
H1.4: +aK169
PTMs of linker histones C. Wood et al.
3694 FEBS Journal 276 (2009) 3685–3697 ª 2009 The Authors Journal compilation ª 2009 FEBS
Acknowledgements
We would like to thank A. Evans for his advice on
cell growth rates and S. Lambert for his advice and
assistance in the preparation of the chicken linker hi-
stones. Both of the aforementioned are based in the
School of Pharmacy and Biomolecular Sciences, Liver-
pool John Moores University.
References
1 Bowman RV, Yang IA, Semmler AB & Fong KM (2006)
Epigenetics of lung cancer. Respirology 11, 355–365.
2 Chuang JC & Jones PA (2007) Epigenetics and micro-
RNAs. Pediatr Res 61, 24R–29R.
3 Wood CM (2008) Molecular kinetics and targeting
within the nucleus. Curr Chem Biol 2, 229–236.
4 Li LC, Carroll PR & Dahiya R (2005) Epigenetic
changes in prostate cancer: implication for diagnosis
and treatment. J Natl Cancer Inst 97, 103–115.
5 Lohrum M, Stunnenberg HG & Logie C (2007) The
new frontier in cancer research: deciphering cancer
epigenetics. Int J Biochem Cell Biol 39, 1450–1461.
6 Takeshima H, Suetake I & Tajima S (2008) Mouse
Dnmt3a preferentially methylates linker DNA and is
inhibited by histone H1. J Mol Biol 283 , 810–821.
7 Li T, Vu TH, Ulaner GA, Littman E, Ling JQ, Chen

HL, Hu JF, Behr B, Giudice L & Hoffman AR (2005)
IVF results in de novo DNA methylation and histone
methylation at an Igf2-H19 imprinting epigenetic
switch. Mol Hum Reprod 11, 631–640.
8 Galm O, Herman JG & Baylin SB (2006) The funda-
mental role of epigenetics in hematopoietic malignan-
cies. Blood Rev 20, 1–13.
9 Suzuki H, Watkins DN, Jair KW, Schuebel KE,
Markowitz SD, Chen WD, Pretlow TP, Yang B,
Akiyama Y, Van Engeland M et al. (2004) Epigenetic
inactivation of SFRP genes allows constitutive WNT
signaling in colorectal cancer. Nat Genet 36, 417–422.
10 Esteller M, Fraga MF, Paz MF, Campo E, Colomer D,
Novo FJ, Calasanz MJ, Galm O, Guo M, Benitez J
et al. (2002) Cancer epigenetics and methylation.
Science 297, 1807–1808.
11 Lu Q, Qiu X, Hu N, Wen H, Su Y & Richardson BC
(2006) Epigenetics, disease, and therapeutic interven-
tions. Ageing Res Rev 5, 449–467.
12 Baylin SB (2005) DNA methylation and gene silencing
in cancer. Nat Clin Pract Oncol 2, S4–S11.
13 Allan J, Hartman PG, Crane-Robinson C & Aviles FX
(1980) The structure of histone H1 and its location in
chromatin. Nature 288, 675–679.
14 Fan L & Roberts VA (2006) Complex of linker histone
H5 with the nucleosome and its implications for chro-
matin packing. Proc Natl Acad Sci USA 103, 8384–
8389.
15 Brown DT, Izard T & Misteli T (2006) Mapping the
interaction surface of linker histone H1 with the nucleo-

some of native chromatin in vivo. Nat Struct Mol Biol
13, 250–255.
16 Routh A, Sandin S & Rhodes D (2008) Nucleosome
repeat length and linker histone stoichiometry determine
chromatin fiber structure. Proc Natl Acad Sci USA 105,
8872–8877.
17 Robinson PJ, Fairall L, Huynh VA & Rhodes D (2006)
EM measurements define the dimensions of the ‘30-nm’
chromatin fiber: evidence for a compact, interdigitated
structure. Proc Natl Acad Sci USA 103, 6506–6511.
18 Bordas J, Perez-Grau L, Koch MH, Vega MC & Nave
C (1986) The superstructure of chromatin and its con-
densation mechanism. II. Theoretical analysis of the
X-ray scattering patterns and model calculations. Eur
Biophys J 13, 175–185.
19 Dorigo B, Schalch T, Kulangara A, Duda S, Schroeder
RR & Richmond TJ (2004) Nucleosome arrays reveal
the two-start organization of the chromatin fiber. Sci-
ence 306, 1571–1573.
20 Thoma F, Koller T & Klug A (1979) Involvement of
histone H1 in the organization of the nucleosome and
of the salt-dependent superstructures of chromatin.
J Cell Biol 83, 403–427.
21 Ramakrishnan V, Finch JT, Graziano V, Lee PL &
Sweet RM (1993) Crystal structure of globular domain
of histone H5 and its implications for nucleosome bind-
ing. Nature 362, 219–223.
22 Khochbin S (2001) Histone H1 diversity: bridging
regulatory signals to linker histone function. Gene 271,
1–12.

23 Izzo A, Kamieniarz K & Schneider R (2008) The his-
tone H1 family: specific members, specific functions?
Biol Chem 389, 333–343.
24 Shen X & Gorovsky MA (1996) Linker histone H1
regulates specific gene expression but not global tran-
scription in vivo. Cell 86, 475–483.
25 Fan Y, Nikitina T, Zhao J, Fleury TJ, Bhattacharyya
R, Bouhassira EE, Stein A, Woodcock CL & Skoultchi
AI (2005) Histone H1 depletion in mammals alters glo-
bal chromatin structure but causes specific changes in
gene regulation. Cell 123, 1199–1212.
26 Kim K, Choi J, Heo K, Kim H, Levens D, Kohno K,
Johnson EM, Brock HW & An W (2008) Isolation and
characterization of a novel H1.2 complex that acts as a
repressor of p53-mediated transcription. J Biol Chem
283, 9113–9126.
27 Gabrilovich DI, Cheng P, Fan Y, Yu B, Nikitina E,
Sirotkin A, Shurin M, Oyama T, Adachi Y, Nadaf S
et al. (2002) H1(0) histone and differentiation of
dendritic cells. A molecular target for tumor-derived
factors. J Leukoc Biol 72, 285–296.
28 Th’ng JP, Sung R, Ye M & Hendzel MJ (2005) H1
family histones in the nucleus. Control of binding and
C. Wood et al. PTMs of linker histones
FEBS Journal 276 (2009) 3685–3697 ª 2009 The Authors Journal compilation ª 2009 FEBS 3695
localization by the C-terminal domain. J Biol Chem
280, 27809–27814.
29 Konishi A, Shimizu S, Hirota J, Takao T, Fan Y,
Matsuoka Y, Zhang L, Yoneda Y, Fujii Y, Skoultchi
AI et al. (2003) Involvement of histone H1.2 in apopto-

sis induced by DNA double-strand breaks. Cell 114 ,
673–688.
30 Gillespie DA & Vousden KH (2003) The secret life of
histones. Cell 114, 655–656.
31 Yan N & Shi Y (2003) Histone H1.2 as a trigger for
apoptosis. Nat Struct Biol 10, 983–985.
32 Conn KL, Hendzel MJ & Schang LM (2008) Linker
histones are mobilized during infection with herpes
simplex virus type 1. J Virol 82, 8629–8646.
33 Okamura H, Yoshida K, Amorim BR & Haneji T
(2008) Histone H1.2 is translocated to mitochondria
and associates with Bak in bleomycin-induced apoptotic
cells. J Cell Biochem 103, 1488–1496.
34 Sarg B, Gre
´
en A, So
¨
derkvist P, Helliger W, Rundquist
I & Lindner HH (2005) Characterization of sequence
variations in human histone H1.2 and H1.4 subtypes.
FEBS J 272, 3673–3683.
35 Clarke HJ, McLay DW & Mohamed OA (1998) Linker
histone transitions during mammalian oogenesis and
embryogenesis. Dev Genet 22, 17–30.
36 Bradbury EM, Inglis RJ & Matthews HR (1974) Con-
trol of cell division by very lysine rich histone (F1)
phosphorylation. Nature 247, 257–261.
37 He J, Yang Q & Chang LJ (2005) Dynamic DNA meth-
ylation and histone modifications contribute to lenti-
viral transgene silencing in murine embryonic

carcinoma cells. J Virol 79, 13497–13508.
38 Wu J, Wang SH, Potter D, Liu JC, Smith LT, Wu YZ,
Huang TH & Plass C (2007) Diverse histone modifica-
tions on histone 3 lysine 9 and their relation to DNA
methylation in specifying gene silencing. BMC Genomics
8, 131, doi:10.1186/1471-2164-8-131.
39 Gilbert N, Thomson I, Boyle S, Allan J, Ramsahoye B
& Bickmore WA (2007) DNA methylation affects
nuclear organization, histone modifications, and linker
histone binding but not chromatin compaction. J Cell
Biol 177, 401–411.
40 Gurley LR, Valdez JG & Buchanan JS (1995) Charac-
terization of the mitotic specific phosphorylation site of
histone H1. Absence of a consensus sequence for the
p34cdc2 ⁄ cyclin B kinase. J Biol Chem 270, 27653–
27660.
41 Sarg B, Helliger W, Talasz H, Fo
¨
rg B & Lindner HH
(2006) Histone H1 phosphorylation occurs site-specifi-
cally during interphase and mitosis: identification of a
novel phosphorylation site on histone H1. J Biol Chem
281, 6573–6580.
42 Garcia BA, Busby SA, Barber CM, Shabanowitz J,
Allis CD & Hunt DF (2004) Characterization of
phosphorylation sites on histone H1 isoforms by
tandem mass spectrometry. J Proteome Res 3, 1219–
1227.
43 Wisniewski JR, Zougman A, Kru
¨

ger S & Mann M
(2007) 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
6, 72–87.
44 Snijders AP, Pongdam S, Lambert SJ, Wood CM, Bald-
win JP & Dickman MJ (2008) Characterization of post-
translational modifications of the linker histones H1
and H5 from chicken erythrocytes using mass spectrom-
etry. J Proteome Res 7, 4326–4335.
45 Ficarro SB, McCleland ML, Stukenberg PT, Burke DJ,
Ross MM, Shabanowitz J, Hunt DF & White FM
(2002) Phosphoproteome analysis by mass spectrometry
and its application to Saccharomyces cerevisiae. Nat
Biotechnol 20, 301–305.
46 Ma M, Kutz-Naber KK & Li L (2007) Methyl
esterification assisted MALDI FTMS characterization
of the orcokinin neuropeptide family. Anal Chem 79,
673–681.
47 Deterding LJ, Bunger MK, Banks GC, Tomer KB &
Archer TK (2008) 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 7, 2368–2379.
48 Alami R, Fan Y, Pack S, Sonbuchner TM, Besse A,
Lin Q, Greally JM, Skoultchi AI & Bouhassira EE
(2003) Mammalian linker-histone subtypes differentially
affect gene expression in vivo. Proc Natl Acad Sci USA
100, 5920–5925.

49 Kuzmichev A, Jenuwein T, Tempst P & Reinberg D
(2004) Different EZH2-containing complexes target
methylation of histone H1 or nucleosomal histone H3.
Mol Cell 14, 183–193.
50 Jiang T, Zhou X, Taghizadeh K, Dong M & Dedon PC
(2007) N-formylation of lysine in histone proteins as a
secondary modification arising from oxidative DNA
damage. Proc Natl Acad Sci USA 104, 60–65.
51 Banks GC, Deterding LJ, Tomer KB & Archer TK
(2001) Hormone-mediated dephosphorylation of specific
histone H1 isoforms. J Biol Chem 276, 36467–36473.
52 Villar-Garea A & Imhof A (2008) Fine mapping of
posttranslational modifications of the linker histone
H1 from Drosophila melanogaster. PLoS ONE 3,
e1553, doi:10.1371/journal.pone.0001553.
53 Lelie
`
vre SA, Weaver VM, Nickerson JA, Larabell CA,
Bhaumik A, Petersen OW & Bissell MJ (1998) Tissue
phenotype depends on reciprocal interactions between
the extracellular matrix and the structural organization
of the nucleus. Proc Natl Acad Sci USA 95, 14711–
14716.
54 Zink D, Fischer AH & Nickerson JA (2004) Nuclear
structure in cancer cells. Nat Rev Cancer 4, 677–
687.
PTMs of linker histones C. Wood et al.
3696 FEBS Journal 276 (2009) 3685–3697 ª 2009 The Authors Journal compilation ª 2009 FEBS
55 Gong CX, Liu F, Grundke-Iqbal I & Iqbal K (2005)
Post-translational modifications of tau protein in

Alzheimer’s disease. J Neural Transm 112, 813–838.
56 Huq M, Gupta P & Wei LN (2008) Post-translational
modifications of nuclear co-repressor RIP140: a thera-
peutic target for metabolic diseases. Curr Med Chem 15,
386–392.
57 Dai B & Rasmussen TP (2007) Global epiproteomic
signatures distinguish embryonic stem cells from
differentiated cells. Stem Cells 25, 2567–2574.
58 Godman CA, Joshi R, Tierney BR, Greenspan E,
Rasmussen TP, Wang HW, Shin DG, Rosenberg DW
& Giardina C (2008) HDAC3 impacts multiple onco-
genic pathways in colon cancer cells with effects on
Wnt and vitamin D signalling. Cancer Biol Ther 7,
1570–1580.
59 Phanstiel D, Brumbaugh J, Berggren WT, Conard K,
Feng X, Levenstein ME, McAlister GC, Thomson JA
& Coon JJ (2008) Mass spectrometry identifies and
quantifies 74 unique histone H4 isoforms in differentiat-
ing human embryonic stem cells. Proc Natl Acad Sci
USA 105, 4093–4098.
60 Sancho M, Diani E, Beato M & Jordan A (2008)
Depletion of human histone H1 variants uncovers
specific roles in gene expression and cell growth.
PLoS Genet 4, e1000227, doi:10.1371/journal.pgen.
1000227.
61 Klumpp S & Krieglstein J (2002) Phosphorylation and
dephosphorylation of histidine residues in proteins. Eur
J Biochem 269, 1067–1071.
62 Auze
´

ly-Velty R & Rinaudo M (2003) Synthesis of
starch derivatives with labile cationic groups. Int J Biol
Macromol 31, 123–129.
63 Shechter D, Dormann HL, Allis CD & Hake SB (2007)
Extraction, purification and analysis of histones. Nat
Protoc 2, 1445–1457.
64 Garcia BA, Joshi S, Thomas CE, Chitta RK, Diaz RL,
Busby SA, Andrews PC, Ogorzalek Loo RR, Shabano-
witz J, Kelleher NL et al. (2006) Comprehensive
phosphoprotein analysis of linker histone H1 from
Tetrahymena thermophila. Mol Cell Proteomics 5, 1593–
1609.
65 Bonaldi T, Imhof A & Regula JT (2004) A combination
of different mass spectroscopic techniques for the
analysis of dynamic changes of histone modifications.
Proteomics 4, 1382–1396.
66 Lindner HH (2008) Analysis of histones, histone vari-
ants, and their post-translationally modified forms.
Electrophoresis 29, 2516–2532.
67 Mizzen CA, Dou Y, Liu Y, Cook RG, Gorovsky MA
& Allis CD (1999) Identification and mutation of phos-
phorylation sites in a linker histone. Phosphorylation of
macronuclear H1 is not essential for viability in tetra-
hymena. J Biol Chem 274, 14533–14536.
68 Krueger KE & Srivastava S (2006) Posttranslational
protein modifications: current implications for cancer
detection, prevention and therapeutics. Mol Cell Proteo-
mics 5, 1799–1810.
69 Li H, Coghlan A, Ruan J, Coin LJ, He
´

riche
´
JK,
Osmotherly L, Li R, Liu T, Zhang Z, Bolund L et al.
(2006) TreeFam: a curated database of phylogenetic
trees of animal gene families. Nucleic Acids Res 34,
D572–D580.
C. Wood et al. PTMs of linker histones
FEBS Journal 276 (2009) 3685–3697 ª 2009 The Authors Journal compilation ª 2009 FEBS 3697