Báo cáo khoa học: Characterization of sequence variations in human histone H1.2 and H1.4 subtypes pptx

Characterization of sequence variations in human histone
H1.2 and H1.4 subtypes
´
Bettina Sarg1, Anna Green2, Peter Soderkvist2, Wilfried Helliger1, Ingemar Rundquist2
ă
and Herbert H. Lindner1
1 Division of Clinical Biochemistry, Biocenter, Innsbruck Medical University, Austria
2 Division of Cell Biology, Linkopings Universitet, Sweden
ă

Keywords
HILIC; linker histones; sequence variants;
SNP; tumor cell lines
Correspondence
H. H. Lindner, Division of Clinical
Biochemistry, Biocenter, Innsbruck Medical
University, Fritz-Pregl-Strasse 3, A-6020
Innsbruck, Austria
Fax: +43 512 507 2876
Tel: +43 512 507 3521
E-mail:
(Received 10 March 2005, revised 10 May
2005, accepted 26 May 2005)
doi:10.1111/j.1742-4658.2005.04793.x

In humans, eight types of histone H1 exist (H1.1–H1.5, H1°, H1t and
H1oo), all consisting of a highly conserved globular domain and less conserved N- and C-terminal tails. Although the precise functions of these isoforms are not yet understood, and H1 subtypes have been found to be
dispensable for mammalian development, it is now clear that specific functions may be assigned to certain individual H1 subtypes. Moreover, microsequence variations within the isoforms, such as polymorphisms or
mutations, may have biological significance because of the high degree of
sequence conservation of these proteins. This study used a hydrophilic
interaction liquid chromatographic method to detect sequence variants

within the subtypes. Two deviations from wild-type H1 sequences were
found. In K562 erythroleukemic cells, alanine at position 17 in H1.2 was
replaced by valine, and, in Raji B lymphoblastoid cells, lysine at position
173 in H1.4 was replaced by arginine. We confirmed these findings by
DNA sequencing of the corresponding gene segments. In K562 cells, a
homozygous GCCfiGTC shift was found at codon 18, giving rise to H1.2
Ala17Val because the initial methionine is removed in H1 histones. Raji
cells showed a heterozygous AAAfiAGA codon change at position 174 in
H1.4, corresponding to the Lys173Arg substitution. The allele frequency of
these sequence variants in a normal Swedish population was found to be
6.8% for the H1.2 GCCfiGTC shift, indicating that this is a relatively frequent polymorphism. The AAAfiAGA codon change in H1.4 was detected
only in Raji cells and was not present in a normal population or in six
other cell lines derived from individuals suffering from Burkitt’s lymphoma. The significance of these sequence variants is unclear, but increasing evidence indicates that minor sequence variations in linker histones
may change their binding characteristics, influence chromatin remodeling,
and specifically affect important cellular functions.

The H1 histones are small basic proteins occurring in
all higher eukaryotes in multiple subtypes that differ
only slightly in their primary sequences. H1 histones
consist of a central, highly conserved globular domain,
while the hydrophilic N- and C-terminal tails exhibit

less sequence conservation. In addition to the heterogeneity of their primary structures, the H1 tails are
also extensively post-translationally modified (e.g.
phosphorylated or ADP-ribosylated) under various
biological conditions. Moreover, the proportion of H1

Abbreviations
CE, capillary electrophoresis; HILIC, hydrophilic interaction liquid chromatography; HPCE, high performance capillary electrophoresis; RFLP,
restriction fragment length polymorphism; SNP, single nucleotide polymorphism; TEAP, triethylammonium phosphate.

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H1 microsequence variants in human cell lines

subtypes varies in a tissue- and species-specific manner,
and the expression of each subtype varies throughout
development and differentiation [1–3].
In the human genome, genes encoding eight different
subtypes of histone H1 have been identified. The genes
encoding H1.1, H1.2, H1.3, H1.4, H1.5 and H1.t are
located on the short arm of chromosome 6, while H1°
is located on chromosome 22 [4] and H1oo on chromosome 3 [5]. Only one copy of each gene is present
[6]. Within these H1 genes several single nucleotide
polymorphisms (SNPs) have been reported (NCBI
Single Nucleotide Polymorphism Database). To our
knowledge, no acquired mutations have been reported
to date in any human H1 gene.
Studies of the structure of H1 histones, their interaction with the nucleosome and their roles in controlling gene activity, indicate that these proteins have
both an essential architectural function and an
important task in regulating transcription [7,8]. The
precise functions of these multiple H1 subtypes and
their modifications are not yet fully understood, but
it has been reported that distinct H1 histone variants
are preferentially localized to particular chromosomal
domains [9–11]. Although individual H1 subtypes are
dispensable for mammalian development [12], it now

seems clear that linker histones, in general, are essential
for proper development [13] and it was recently found
that one subtype, H1.2, had a specific role in DNA
damage-induced apoptosis [14]. It is useful therefore to
examine the properties and expression of these variants
as this furthers a better understanding of the relevance
of this diversity for particular cellular activities.
The sequence similarity between histones H1.1–H1.5
requires highly efficient analytical methods for their
resolution. To date, the most widely utilized procedures for the study of human H1 proteins have been
PAGE and low-pressure ion-exchange chromatography. Two-dimensional gel electrophoresis allows the
separation of several H1 variants [15,16], and four H1
subtypes were obtained by using BioRex 70 column
chromatography [17–19]. Both PAGE and lowpressure ion-exchange chromatography, however, are
laborious, time-consuming and their resolution is
unsatisfactory.
Recently, we described rapid and simple methods
for the separation of rat and mouse H1 histones by
using RP-HPLC [20,21] and high performance capillary electrophoresis (HPCE) [22–24]. Furthermore, by
applying hydrophilic interaction liquid chromatography (HILIC) excellent fractionations of various posttranslationally modified core [25,26] and linker [27,28]
histones were obtained in both the analytical and the
semipreparative scale.
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B. Sarg et al.

This article evaluates the potential utility of HILIC
as a means of investigating the occurrence of sequence
variations within linker histone subtypes from various
human tumor cell lines. By using HILIC we detected

amino acid substitutions in H1.2 and H1.4 at the protein level. In addition, sequencing of the corresponding
gene segments confirmed these findings at the genome
level. Furthermore, we also screened DNA from 103
healthy individuals to obtain the allele frequency for
these variants.

Results
HILIC is an excellent technique for using to separate
histone proteins and their modified forms [25–29]. This
study aimed to apply and optimize the HILIC technique in order to detect the occurrence of microsequence variations of H1 subtypes isolated from
various human tumor cell lines. As the level of histone
H1 phosphorylation is lower in nondividing than in
proliferating cells, we isolated H1 histones from cell
cultures in the stationary phase, thus making it possible to reduce the occurrence of additional peaks
caused by phosphorylated forms of the parent proteins. A typical separation pattern using a PolyCAT A
column and a two-step sodium perchlorate gradient
(0–0.68 m) in the presence of 70% (v ⁄ v) acetonitrile
and 0.015 m triethylammonium phosphate (TEAP)
(pH 3.0) is shown in Fig. 1A. The CCRF-CEM H1
sample was separated into four peaks. Analysis of several other cell lines showed the same pattern, but the
relative concentrations of the subtypes varied (data not
shown). H1 samples from Raji (Fig. 1B) and K562
(Fig. 1C) cells, however, showed different patterns,
and additional peaks were detected, namely peak 3a in
Fig. 1B and peak 1a in Fig. 1C.
To identify the individual peaks of the chromatograms we digested the various subfractions with chymotrypsin, a protease that specifically hydrolyzes peptide
bonds at the C terminus of Tyr, Phe and Trp. As
human H1 histones contain only one Phe residue, it
was expected that cleavage with chymotrypsin should
produce two peptide fragments. In fact, when the

digested proteins were separated by RP-HPLC, two
main fractions were obtained. An example is shown in
Fig. 2, where a digest of fraction 3 isolated by HILIC
separation of H1 from CCRF-CEM cells (Fig. 1A)
was analyzed by RP-HPLC. The purity and homogeneity of the two fractions was assessed by capillary electrophoresis (CE) (data not shown). To identify the
fractions, amino acid sequencing was performed.
Fraction 1 contained the C-terminal region starting
after Phe at amino acid 105. No sequence data were
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B. Sarg et al.

H1 microsequence variants in human cell lines

A

B

C

Fig. 1. Hydrophilic interaction liquid chromatography (HILIC) separation of H1 histones isolated from human tumor cell lines. H1 histone samples from (A) CCRF-CEM cells, (B) Raji cells, and (C)
K562 cells were analyzed on a PolyCAT A column (4.6
mm · 250 mm) at 23 °C, and at a constant flow of 1.0 mLỈmin)1,
by using a two-step gradient starting at solvent A ⁄ solvent B
(100 : 0) [solvent A: 70% (v ⁄ v) acetonitrile, 0.015 M triethylammoniumphosphate (TEAP, pH 3.0); solvent B: 70% (v ⁄ v) acetonitrile,
0.015 M TEAP (pH 3.0) and 0.68 M NaClO4]. The concentration of
solvent B was increased from 0 to 80% (v ⁄ v) during a time-period
of 5 min and from 80 to 100% (v ⁄ v) during a time-period of 60 min.
The isolated protein fractions (designated 1–4) were desalted by

using RP-HPLC.

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Fig. 2. RP-HPLC analysis of peptide fractions of chymotrypsindigested H1 from CCRF-CEM cells. Peak 3 from H1 histones isolated with hydrophilic interaction liquid chromatography (HILIC)
(Fig. 1A) was digested with chymotrypsin, as described in the
Experimental procedures. The digest (containing  100 lg of
protein) was injected onto a Nucleosil 300-5 C18 column
(250 mm · 3 mm). Analysis was performed at a constant flow of
0.35 mLỈmin)1 with a multistep acetonitrile gradient starting at solvent A ⁄ solvent B (85 : 15) (solvent A: water containing 0.1% (v ⁄ v)
trifluoroacetic acid; solvent B: 85% (v ⁄ v) acetonitrile and 0.1%
(v ⁄ v) trifluoroacetic acid). The concentration of solvent B was
increased linearly from 15 to 23% during a time-period of 25 min,
from 23 to 70% during a time-period of 45 min and from 70 to
100% during a time-period of 5 min.

obtained from fraction 2 because the first amino acid
was blocked, indicating that fraction 2 consisted of the
N-terminal region of this H1 subtype. By using protein
sequence data, the four H1 peaks from CCRF-CEM
cells (Fig. 1A) were identified as follows: peak 1, histone H1.2 (SWISS-PROT P16403); peak 2, histones
H1.3 (SWISS-PROT P16402) + H1.5 (SWISS-PROT
P16401); peak 3, histone H1.4 (SWISS-PROT P10412);
and peak 4, histone H1.5.
Identification of the Raji H1 fractions (Fig. 1B)
yielded the same result for peaks 1, 2 and 4, while
peaks 3a and 3b contained histone H1.4. In order to
exclude the presence of phosphorylated H1.4, which
would be a reasonable explanation for this diversity,
the Raji H1 proteins were incubated with alkaline

phosphatase and subjected to HILIC. Peak 4 showed a
dramatic decrease in size, whereas the two H1.4 peaks
were not affected by the phosphatase (data not
shown). Therefore, peak 4 was identified as a phosphorylated form of H1.5. Surprisingly, the two H1.4
subfractions (Fig. 1B, peaks 3a and 3b) were not separated either by HPCE or RP-HPLC or by different
gel-electrophoretic methods (data not shown). To
examine the structural difference between these two
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H1 microsequence variants in human cell lines

proteins, we analyzed the two main fractions 1 and 2
obtained from chymotrypsin digestion and RP-HPLC
separation (as shown for CCRF-CEM cells in Fig. 2)
of peaks 3a and 3b under HILIC conditions. We
found that fraction 2 derived from peaks 3a and 3b
had the same elution time, while fraction 1 clearly differed. Therefore, the C-terminal domain should be
responsible for the microheterogeneity of histone H1.4.
In order to elucidate the nature of this alteration,
the C-terminal peptides were further cleaved with
endoproteinase Glu-C. Digests were analyzed by
RP-HPLC by using a Nucleosil 300-5 C18 column
(250 · 3 mm; Fig. 3). Fragmentation yielded three
main peptides (I–III), which were identified by Edman
degradation: fraction I (eluting at 27 min) consisted of
a peptide starting at residue 105, fraction II (43 min)
consisted of a peptide starting at residue 115, and fraction III (60 min) consisted of a peptide starting at residue 150. The purity of the fractions obtained was
confirmed by CE (data not shown). Again, all peptides
were analyzed under HILIC conditions, and the result

showed that only the two fractions III contained
peptides with different elution times. Further peptide

Fig. 3. RP-HPLC analysis of peptide fractions of endoproteinase
Glu-C-digested fraction 1 from Raji H1.4. Peak 3a from Raji H1
histones isolated with hydrophilic interaction liquid chromatography
(HILIC) (Fig. 1B) was digested with chymotrypsin and isolated by
RP-HPLC (Fig. 2). The fraction 1 obtained was further digested with
endoproteinase Glu-C, as described in the Experimental procedures, and the digest (containing  30 lg of protein) was injected
onto a Nucleosil 300-5 C18 column (250 mm · 3 mm). Analysis was
performed at a constant flow of 0.35 mLỈmin)1 with a multistep
acetonitrile gradient starting at solvent A ⁄ solvent B (95 : 5) [solvent
A: water containing 0.1% (v ⁄ v) trifluoroacetic acid; solvent B: 85%
(v ⁄ v) acetonitrile and 0.1% (v ⁄ v) trifluoroacetic acid]. The concentration of solvent B was increased linearly from 5 to 20% during a
time-period of 65 min and from 20 to 100% during a time-period of
25 min.

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B. Sarg et al.

sequencing of fraction III revealed that the two H1.4
HILIC peaks differed from one another by a single
amino acid substitution – lysine at position 173 (peak
3b) was replaced by arginine (peak 3a). This result was
further confirmed by subjecting the two H1.4 subfractions obtained by HILIC (Fig. 1B) to electrospray ionization mass spectrometry analysis. For peak 3a we
found a mass of 21802.4 Da and for peak 3b a mass
of 21774.8 Da, the latter being in close agreement with
the wild-type H1.4 mass, which was calculated to be
21776.1 Da. The mass difference observed between the

two peaks was 27.6 Da, and this corresponds to the
mass difference between lysine and arginine. This
microsequence variant H1.4 Lys173Arg, found in Raji
cells in the same concentrations as the wild-type H1.4,
was detected neither in CCRF-CEM and K562 cells
nor in several other human cell lines (e.g. U937,
HL60) or in human tissue (placenta, testis).
A further microheterogeneity was found when analyzing the K562 H1 sample by HILIC (Fig. 1C). Peak
1, which was a single fraction in CCRF-CEM and Raji
cells, and identified as histone H1.2, was separated into
two subfractions (peaks 1a and 1b) in K562 cells.
RP-HPLC separation after chymotrypsin digestion of
the two fractions 1a and 1b yielded two main fragments each (similar to Fig. 2). HILIC analysis revealed
that fraction 2 from 1a had a shorter elution time than
did fraction 2 from 1b, whereas no differences were
observed between the fraction 1 samples. Further cleavage of the fraction 2 samples with endoproteinase
Glu-C followed by peptide sequencing showed that the
proteins differed by one amino acid out of a total of
212: peak 1a contained valine in position 17, while
peak 1b contained wild-type alanine. Histone H1.2
from all cell lines and tissues investigated contained
only alanine at this location.
To confirm the H1.2 Ala17Val sequence variation, a
183 bp PCR product was amplified in the 5¢-UTR, and
the start of the coding sequence of the H1.2 gene corresponding to the N-terminal tail of the protein. PCR
products from K562, Raji and wild-type blood donors
were sequenced. K562 DNA contained a homozygous
CfiT substitution at nucleotide position 578 in the
H1.2 gene, resulting in a change, in codon 18, from
GCC to GTC (Fig. 4), encoding alanine and valine,

respectively. This change in codon 18 corresponds to
the substitution at amino acid position 17 in the H1.2
protein, as the initiating methionine is removed after
translation in all H1 histones. Traces of the wild-type
C in position 578 were also detected (Fig. 4).
To obtain the population frequency of the
H1.2 g578 CfiT substitution, 103 healthy individuals
were screened by using a restriction fragment length
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B. Sarg et al.

H1 microsequence variants in human cell lines

Fig. 4. DNA sequencing of H1.2 PCR products.

Fig. 6. DNA sequencing of H1.4 PCR products.

A

B

C

Fig. 5. Restriction fragment length polymorphism (RFLP) analysis
of H1.2 PCR products. Lanes 1 and 2, blood donors, heterozygous
for g578 CfiT. Lane 3, wild-type blood donor. Lane 4, K562, homozygous for g578 CfiT. Lane 5, uncleaved control. Lane 6, 100 bp
ladder.

polymorphism (RFLP) assay. The wild-type H1.2 PCR
product was cleaved into three fragments by BsuRI,
while a PCR product containing the g578 CfiT substitution was cleaved into two fragments (Fig. 5). We
found 10 individuals to be heterozygous and two
homozygous for the g578 CfiT substitution, resulting
in an allele frequency of 6.8% in this population. The
presence of the g578 CfiT substitution in samples displaying the 130 bp fragment in the RFLP analysis was
confirmed by DNA sequencing.
To detect the H1.4 Lys173Arg substitution, a 217 bp
fragment of the H1.4 gene, corresponding to the C terminus of the H1.4 protein, was amplified by PCR. Raji,
K562 and wild-type blood donor PCR products were
subjected to DNA sequencing. Raji cells were heterozygous for an AAA to AGA codon change at position
174 (Fig. 6), resulting in a Lys173Arg substitution in
histone H1.4. To determine the allele frequency of this
g1250 AfiG substitution in the H1.4 gene, a denaturating HPLC method was developed (Fig. 7). Heteroduplex and mutant homoduplex formation was
FEBS Journal 272 (2005) 3673–3683 ª 2005 FEBS

Fig. 7. Denaturing HPLC of PCR-amplified H1.4 fragments. (A)
Wild-type blood donor. (B) Raji, heterozygous for g1250 AfiG. (C)
Raji and wild-type PCR products in a 1 : 1 ratio.

resolved after mixing all PCR products with wild-type
PCR products of H1.4. No G alleles were detected in
the normal population of 206 alleles studied. As the Raji
cell line was derived from an individual suffering from
Burkitt’s lymphoma, six other cell lines established from
Burkitt’s lymphoma were screened for the g1250 AfiG
substitution. However, none of these cell lines contained
this sequence variation.

Discussion
A major question concerning the expression of individual H1 subtypes is whether they have coevolved
with functional differences. Although the precise function of H1 isoforms has yet to be determined, several
observations suggest distinct and nonoverlapping roles
for individual H1 variants. Estimation of the rates of
nucleotide substitution for mammalian H1 subtypes
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H1 microsequence variants in human cell lines

H1a–H1e during evolution showed evidence of their
functional differentiation [30]. The study revealed that
the rates of nucleotide substitution differed not only
among subtypes, but also among domains. For all
subtypes, the synonymous substitution rate greatly
exceeded the nonsynonymous rate, and the terminal
domains were more variable than the central globular
domain. We detected two nonsynonymous substitutions in human H1 histones. One caused alanine to
be substituted by valine in the N-terminal region
(position 17) of histone H1.2 in K562 cells, leading to
two subfractions using HILIC. The H1.2 Ala17Val
variant constituted the major fraction of the H1.2
protein extracted, while a minor fraction of wild-type
H1.2 was present. In agreement with this, K562
DNA was found to carry a homozygous GCC to
GCT change at codon 18 in the H1.2 gene. There
were remains of the wild-type C at g578 (Fig. 4),
explaining the minor wild-type H1.2 peak in the
HILIC chromatogram. This result is probably

explained by the nondiploid karyotype of the K562
cells. The H1.2 g578 CfiT sequence variant was also
found to be present in a normal population, with the
allele frequency 6.8%, and is therefore concluded
to be a polymorphism. The H1.2 Ala17Val was
expressed in K562 cells and therefore probably also
in normal individuals carrying this gene variant.
However, H1 histones show a high redundancy in
knockout organisms, and the deletion of one or more
subtypes causes increased expression of those remaining [12,31]. Therefore, it cannot be completely ruled
out that normal individuals carrying the H1.2 g578
CfiT are devoid of Ala17Val H1.2 expression. This
polymorphic site in the H1.2 protein has previously
been recognized in human spleen [18]. The corresponding SNP was recently reported (NCBI SNP database refSNP ID rs 2230653). In addition, three other
SNPs have been reported in H1.2, all leading to
synonymous changes (NCBI SNP). The role of the
N-terminal tail of H1 histones is unclear, but is
believed to be involved in positioning of the globular domain on the nucleosome [32]. The N- and
C-terminal tails of histone H1 adopt a random coil in
solution [33]. On binding of histone H1° to DNA,
significant parts of the N terminus are likely to take
on an a-helical structure [34], and this is probably
also the case for other H1 subtypes. As the tails of
H1 histones do not adopt their native conformation
until they bind to chromatin, it is hard to predict the
structural changes that a single amino acid substitution
may trigger. Substitution of valine for alanine may
affect the predicted a-helical structure of the tail as
valine has a less stabilizing effect on an a-helical struc3678

B. Sarg et al.

ture. If the structure is affected by the polymorphism,
the positioning of H1.2 on the nucleosome, or the
binding of H1.2 to chromatin, may be affected.
A further nonsynonymous substitution prompted
the replacement of lysine with arginine in the C-terminal tail (position 173) of histone H1.4 in Raji cells.
By using HILIC, histone H1.4 was separated into two
peaks: one wild-type H1.4; and one Lys173Arg H1.4.
This microsequence variant was found for the first
time and was present in stationary-phase cells in similar amounts as wild-type H1.4. Genetic analysis of
Raji cells showed a heterozygous H1.4 g1250 AfiG
substitution, prompting alteration of codon 174 from
AAA to AGA, in agreement with the Lys173Arg substitution. This sequence variant was not present in the
103 normal individuals that were screened, or in six
other Burkitt’s lymphoma cell lines, implying that the
Lys173Arg substitution is probably a mutation or a
rare polymorphism detected, thus far, only in Raji
cells. Denaturing HPLC, used to screen for this genetic
variant, provides a sensitive and highly specific method
for investigating sequence variations [35], and the possibility of false negative results is unlikely. Histone
H1.4 mRNA (GenBank NM_005321) has been reported to contain a different polymorphism (NCBI
SNP refSNP ID rs2298090), c455 AfiG, causing a
Lys152Arg substitution (NP_005312.1).
The C-terminal tail of histone H1 is believed to be
responsible for the condensation of chromatin [32,36],
and the condensing property of rat H1d probably
resides in a C-terminal 34 amino acid stretch [37].
Different H1 subtypes may have different chromatincondensing properties [38,39]. On binding to DNA, the
C-terminal tail probably adopts the structure of a segmented a helix [40]. Replacement of lysine with arginine may affect the secondary structure of the

C-terminal tail and the binding of H1.4 to chromatin,
as arginine offers additional hydrogen-bonding abilities
to DNA as compared to lysine. Lysine 173 in H1.4 is
situated in an SPKK motif, one of the known sites for
H1 phosphorylation. As far as we know, there are no
differences in the phosphorylation behavior of SPKK
and SPRK motifs.
Identification of numerous linker histone variants in
vertebrates suggests that these proteins may play specialized roles. Recent investigations using gene-targeting
techniques, however, suggest that the specific timing of
expression may have a greater functional significance
than the nature of the individual H1 subtypes [2,41].
It is, however, not clear to what extent the function of
H1 variants depends on their primary sequence or on
the specific timing of their expression. The literature
presents arguments in favor of both possibilities [2,7].
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The selective effect of linker histones on transcription of individual genes was demonstrated by using
an in vivo system for inducible overexpression of different H1 subtypes (H1°, H1c) in mouse cells [42,43].
Subtype-specific effects were shown to be related to
differences in the structure of the globular domains
[44]. As H1 subtypes differ not only in primary
sequence but also in turnover rate and extent of
phosphorylation, they have the potential to add a
great deal of flexibility to chromatin structure and
transcriptional activation.

Both genes homologous to H1.2 and H1.4 have been
disrupted in mouse, each resulting in viable and fertile
knockout mice [12], indicating that these individual subtypes are dispensable and that compensatory effects
reside between the subtypes, thus keeping H1 stoichiometry intact. Despite the dispensability of wild-type H1
subtypes, however, microsequence variants may have
biological significance. Recent evidence reveals a highly
specific function for H1.2 in DNA damage-induced
apoptosis [14]. As the somatic H1 subtypes H1.1–H1.5
show a high degree of sequence conservation, such specificity must rely on subtle differences in amino acid
sequence. Therefore, as histone H1 is implicated in chromatin organization, cell differentiation, gene regulation
and apoptosis, these processes may be affected by minor
sequence variations, including SNPs.
In conclusion, we have demonstrated the remarkably
high resolving power of HILIC by using this technique
to separate sequence variants within human linker histone subtypes. We were thus able to detect an Ala17Val substitution in histone H1.2 in K562 cells, as well
as a Raji-specific H1.4 Lys173Arg sequence variation
at the protein level. These observations were confirmed
at the genetic level. The significance of these variations
is unclear, but it seems increasingly clear that minor
sequence variations in linker histones may affect
important cellular functions in vivo.

Experimental procedures
Chemicals
Sodium perchlorate (NaClO4), trifluoroacetic acid and triethylamine were purchased from Fluka (Buchs, Switzerland). All other chemicals were purchased from Merck
(Darmstadt, Germany), unless indicated otherwise.

Cell lines and culture conditions
CCRF-CEM acute lymphoblastic leukemia cells, Raji cells
(originally derived from patients with Burkitt’s lymphoma)

and K562 erythroleukemic cells were cultured in RPMI-

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H1 microsequence variants in human cell lines

1640 medium (Biochrom, Berlin, Germany) supplemented
with 10% (v ⁄ v) fetal bovine serum, penicillin (60 lgỈmL)1)
and streptomycin (100 lgỈmL)1) in the presence of 5%
(v ⁄ v) CO2. The cells were seeded at a density of 8 · 104
cellsỈmL)1 and harvested after 7 days to accumulate cells in
stationary phase.

Preparation of H1 histones
Human cells (6–7 · 109) were collected by centrifugation
(800 g for 10 min). H1 histones were extracted with perchloric acid (5%, w ⁄ v) according to the procedure of Lindner et al. [28].

HPLC
The equipment used for HPLC consisted of two 114M
pumps, a 421A system controller and a Model 165 UV-visible-region detector (Beckman Instruments, Palo Alto, CA,
USA). The effluent was monitored at 210 nm and the peaks
were recorded by using Beckman System Gold software.

HILIC
Whole human H1 samples were analyzed on a PolyCAT A
column (4.6 mm · 250 mm; 5 lm particle pore size; 30 nm
pore size; ICT, Vienna, Austria) at 23 °C, and at a constant
flow of 1.0 mLỈmin)1, by using a two-step gradient starting at
solvent A ⁄ solvent B (100 : 0) [solvent A: 70% (v ⁄ v) acetonitrile, 0.015 m TEAP, pH 3.0; solvent B: 70% (v ⁄ v) acetonitrile, 0.015 m TEAP (pH 3.0) and 0.68 m NaClO4]. The
concentration of solvent B was increased from 0 to 80%

(v ⁄ v) during a time-period of 5 min and from 80 to 100%
(v ⁄ v) during a time-period of 60 min. The isolated protein
fractions were desalted by using RP-HPLC. Histone fractions
obtained in this manner were collected and, after adding
0.01 m 2-mercaptoethanol, freeze-dried and stored at )20 °C.

RP-HPLC
The peptides obtained by limited chymotrypsin digestion of
human H1 histones were separated by using a Nucleosil
300-5 C18 column (250 mm · 3 mm internal diameter; 5 lm
particle pore size; end-capped; Macherey-Nagel, Duren,
ă
Germany). Samples of  100 lg were injected onto the column. Chromatography was performed within 70 min at a
constant flow of 0.35 mLỈmin)1 with a multistep acetonitrile
gradient starting at solvent A ⁄ solvent B (85 : 15) [solvent
A: water containing 0.1% (v ⁄ v) trifluoroacetic acid; solvent
B: 85% (v ⁄ v) acetonitrile and 0.1% (v ⁄ v) trifluoroacetic
acid]. The concentration of solvent B was increased linearly
from 15 to 23% during a time-period of 25 min, from 23 to
70% during a time-period of 45 min and from 70 to 100%
during a time-period of 5 min.

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H1 microsequence variants in human cell lines

Human H1 peptide fractions obtained by digestion
with endoproteinase Glu-C were separated by using the
same column and solvents as described above. The concentration of solvent B was increased linearly from 5 to

20% during a time-period of 65 min and from 20 to
100% during a time-period of 25 min. Fractions obtained
in this manner were collected and, after adding 20 lL of
2-mercaptoethanol (0.2 m), were lyophilized and stored at
)20 °C.

B. Sarg et al.

tion, DNA samples collected randomly from 103 normal
individuals in south-east Sweden were screened. The individuals were selected from a population register and were
22–77 years of age (mean age, 52 years; SD, 17 years; 47%
men and 53% women). The design was approved by Linkoă
ping University Hospital Ethical Committee. DNA was
extracted from the blood samples by using the QIAampÒ
DNA Blood Maxi kit (Qiagen).

PCR amplification and RFLP analysis of H1.2
Chymotrypsin digestion
Histone H1 subfractions ( 100 lg), obtained from human
cell lines by HILIC fractionation, were digested with
a-chymotrypsin (EC 3.4.21.1) (Sigma type I-S, 1 ⁄ 150, w ⁄ w)
in 100 lL of 100 mm sodium acetate buffer (pH 5.0) for
30 min at room temperature. The digest was subjected to
RP-HPLC.

Endoproteinase Glu-C digestion
Histone H1.4 fractions ( 50 lg) were digested with Staphylococcus aureus V8 Protease (Boehringer Mannheim,
Mannheim, Germany; 1 : 20, w ⁄ w) in 50 lL of 50 mm
NH4HCO3 buffer (pH 7.8) for 5 h at 37 °C. Histone H1.2
fractions ( 50 lg) were digested in 50 lL of 25 mm phosphate buffer (pH 7.8) for 6 h at room temperature. The

digests were subjected to RP-HPLC.

Peptide sequencing
Peptide sequencing was performed on an Applied Biosystems Inc. (ABI, Foster City, CA, USA) Model 492 Procise
protein sequenator. Typically, 5–100 pm of a peptide sample was run for 3–40 cycles, as required for an unambiguous identification.

Mass-spectrometric analysis
Determination of the molecular masses of the two histone
H1.4 subfractions obtained by the HILIC run was carried
out by electrospray ionization mass spectrometry using an
LCQ ion trap instrument (ThermoFinnigan, San Jose, CA,
USA). Samples (15 lg) were dissolved in 50% (v ⁄ v) aqueous methanol, containing 0.1% (v ⁄ v) formic acid, and
injected into the ion source.

DNA samples
Genomic DNA from various cell lines was extracted by
using the DneasyTM tissue kit (Qiagen, Hilden, Germany)
and examined for sequence variations in codon 18 of H1.2
and in codon 174 of H1.4. To obtain the frequency of the
two polymorphisms in H1.2 and H1.4 in a normal popula-

3680

A 183 bp fragment of the H1.2 gene (HIST1H1C, GenBank
X57129) was amplified by using the PCR primers
5¢-CCCAGGCGCTGCTTC-3¢ (nucleotides 469fi483 of
the H1.2 gene) and 5¢-CTCTGACACCGGGGGAC-3¢
(nucleotides 651fi635 of the H1.2 gene). The PCR was performed with 50 ng of DNA in a 20 lL reaction, containing
1 mm MgCl2, 0.025 mL)1 Taq DNA polymerase, 20 mm
Tris ⁄ HCl, pH 8.4, 50 mm KCl, 1 lm of each primer (all

Life Technologies, Gaithersburg, MD, USA) and 200 lm of
each dATP, dCTP, dGTP and dTTP (Amersham Pharmacia Biotech, Piscataway, NJ, USA). After an initial denaturation at 94 °C for 2 min, amplification was performed for
35 cycles with denaturation at 94 °C for 1 min, annealing
at 57 °C for 1 min and extension at 72 °C for 1 min, in a
thermal cycler (PTC-200; MJ Research, Watertown, MA,
USA). The reaction was completed with an extension step
at 72 °C for 7 min. RFLP analysis was carried out by
digesting the 183 bp PCR product with 10 U BsuRI
(HaeIII) (MBI Fermentas, St Leon-Rot, Germany), at
37 °C overnight. BsuRI recognizes the sequence 5¢-GGCC3¢, and the PCR product from wild-type DNA was digested
into three fragments of 21, 53 and 109 bp. Digestion of the
PCR product from DNA containing the g578 CfiT substitution in the recognition sequence produced two fragments,
one of 53 bp and one of 130 bp. The digested PCR
products were analyzed on Tris ⁄ borate ⁄ EDTA-agarose gels
containing 1% (w ⁄ v) agarose (BioRad, Hercules, CA,
USA), 3% (w ⁄ v) NuSieve GTG Agarose (FMC BioProducts, Rockland, ME, USA) and ethidium bromide
(0.5 lgỈmL)1). The fragments were visualized under UV
transillumination and photographed by using a Polaroid
camera.

PCR amplification of H1.4 and polymorphism
detection by using denaturing HPLC
A 217 bp fragment of the H1.4 gene (HIST1H1E, GenBank
M60748) was PCR amplified by using the primers 5¢-GA
AGAGCGCCAAGAAGACC-3¢ (nucleotides 1173fi1191
of the H1.4 gene) and 5¢-CTACTTTTTCTTGGCTGCCG
(nucleotides 1389fi1370 of the H1.4 gene), using the same
conditions as described above. For mutation analysis, a
denaturating HPLC system (WAVEÒ Nucleic Acid Frag-

FEBS Journal 272 (2005) 3673–3683 ª 2005 FEBS


B. Sarg et al.

ment Analysis System; Transgenomic, Crewe, UK) was
used. All samples were mixed with wild-type PCR product,
which had previously been subjected to DNA sequence analysis, in a 1 : 1 ratio to ensure detection of g1250 AfiG
homozygous mutants. Before analysis, the samples were
denaturated at 95 °C for 4 min and then gradually
cooled, by 1 °CỈmin)1, until 25 °C was reached, to allow
heteroduplex formation.
The optimal melting temperature for the fragment was
calculated by using the wave software, and a temperature
of 63.3 °C was used for analysis. The flow rate was
0.9 mLỈmin)1 and the total run time 7.2 min. Samples
(20 lL) were injected onto the DNA Sep Column at 54%
buffer A (0.1 m triethylammonium acetate, pH 7.0) and
46% buffer B [0.1 m triethylammonium acetate, pH 7.0,
and 25% acetonitrile (v ⁄ v)] and heteroduplexes were separated by using a gradient starting at 49% buffer A and 51%
buffer B, and gradually increasing to 40% buffer A and
60% buffer B.

DNA sequencing
Direct cycle sequencing of H1.2 and H1.4 PCR products
was performed with the corresponding forward PCR primer, using Thermo Sequenase radiolabeled terminator cycle
sequencing kit (USB Corporation, Cleveland, OH, USA),
and labeling with 33P dideoxy nucleotides (Amersham Pharmacia Biotech), according to the manufacturers’ recommendations. Prior to sequencing, the PCR products were
purified and concentrated by using GFX PCR DNA and
the Gel band purification kit (Amersham Pharmacia Biotech). The labeled products from the sequencing reaction

were separated on 6% (w ⁄ v) polyacrylamide gels, containing 6 m urea, in a gel apparatus (OWL) at 70 W constant
power. After electrophoresis, the gel was dried and exposed
to X-ray film.

Acknowledgements
We thank A. Devich, A. Molbaek and S. Gstrein for
their excellent technical assistance. This work, as part
of the European Science Foundation EUROCORES
Programme EuroDYNA, was supported by funds from
the Austrian Science Foundation (project I23-B03) and
the EC Sixth Framework Programme under Contract
no. ERAS-CT-2003-980409 and in part by the Swedish
Cancer Society.

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