H2A.Z.2.2 is an alternatively spliced histone H2A.Z variant that causes severe nucleosome destabilization.
ABSTRACT The histone variant H2A.Z has been implicated in many biological processes, such as gene regulation and genome stability. Here, we present the identification of H2A.Z.2.2 (Z.2.2), a novel alternatively spliced variant of histone H2A.Z and provide a comprehensive characterization of its expression and chromatin incorporation properties. Z.2.2 mRNA is found in all human cell lines and tissues with highest levels in brain. We show the proper splicing and in vivo existence of this variant protein in humans. Furthermore, we demonstrate the binding of Z.2.2 to H2A.Z-specific TIP60 and SRCAP chaperone complexes and its active replication-independent deposition into chromatin. Strikingly, various independent in vivo and in vitro analyses, such as biochemical fractionation, comparative FRAP studies of GFP-tagged H2A variants, size exclusion chromatography and single molecule FRET, in combination with in silico molecular dynamics simulations, consistently demonstrate that Z.2.2 causes major structural changes and significantly destabilizes nucleosomes. Analyses of deletion mutants and chimeric proteins pinpoint this property to its unique C-terminus. Our findings enrich the list of known human variants by an unusual protein belonging to the H2A.Z family that leads to the least stable nucleosome known to date.
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ABSTRACT: Fluctuations in the ambient environment can trigger chromatin disruptions, involving replacement of nucleosomes or exchange of their histone subunits. Unlike canonical histones, which are available only during S-phase, replication-independent histone variants are present throughout the cell cycle and are adapted for chromatin repair. The H2A.Z variant mediates responses to environmental perturbations including fluctuations in temperature and seasonal variation. Phosphorylation of histone H2A.X rapidly marks double-strand DNA breaks for chromatin repair, which is mediated by both H2A and H3 histone variants. Other histones are used as weapons in conflicts between parasites and their hosts, which suggests broad involvement of histone variants in environmental responses beyond chromatin repair.Trends in cell biology. 01/2014;
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ABSTRACT: One of the most remarkable chromatin remodelling processes occurs during spermiogenesis, the post-meiotic phase of sperm development during which histones are replaced with sperm-specific protamines to repackage the genome into the highly compact chromatin structure of mature sperm. Here we identify Chromodomain helicase DNA binding protein 5 (Chd5) as a master regulator of the histone-to-protamine chromatin remodelling process. Chd5 deficiency leads to defective sperm chromatin compaction and male infertility in mice, mirroring the observation of low CHD5 expression in testes of infertile men. Chd5 orchestrates a cascade of molecular events required for histone removal and replacement, including histone 4 (H4) hyperacetylation, histone variant expression, nucleosome eviction and DNA damage repair. Chd5 deficiency also perturbs expression of transition proteins (Tnp1/Tnp2) and protamines (Prm1/2). These findings define Chd5 as a multi-faceted mediator of histone-to-protamine replacement and depict the cascade of molecular events underlying this process of extensive chromatin remodelling.Nature Communications 01/2014; 5:3812. · 10.74 Impact Factor
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ABSTRACT: In Förster resonance energy transfer (FRET) experiments, extracting accurate structural information about macromolecules depends on knowing the positions and orientations of donor and acceptor fluorophores. Several approaches have been employed to reduce uncertainties in quantitative FRET distance measurements. Fluorophore-position distributions can be estimated by surface accessibility (SA) calculations, which compute the region of space explored by the fluorophore within a static macromolecular structure. However, SA models generally do not take fluorophore shape, dye transition-moment orientation, or dye-specific chemical interactions into account. We present a detailed molecular-dynamics (MD) treatment of fluorophore dynamics for an ATTO donor/acceptor dye pair and specifically consider as case studies dye-labeled protein-DNA intermediates in Cre site-specific recombination. We carried out MD simulations in both an aqueous solution and glycerol/water mixtures to assess the effects of experimental solvent systems on dye dynamics. Our results unequivocally show that MD simulations capture solvent effects and dye-dye interactions that can dramatically affect energy transfer efficiency. We also show that results from SA models and MD simulations strongly diverge in cases where donor and acceptor fluorophores are in close proximity. Although atomistic simulations are computationally more expensive than SA models, explicit MD studies are likely to give more realistic results in both homogeneous and mixed solvents. Our study underscores the model-dependent nature of FRET analyses, but also provides a starting point to develop more realistic in silico approaches for obtaining experimental ensemble and single-molecule FRET data.Biophysical Journal 08/2014; · 3.67 Impact Factor
H2A.Z.2.2 is an alternatively spliced histone H2A.Z
variant that causes severe nucleosome
Clemens Bo ¨nisch1, Katrin Schneider2, Sebastian Pu ¨nzeler1, Sonja M. Wiedemann1,
Christina Bielmeier2, Marco Bocola3, H. Christian Eberl4, Wolfgang Kuegel5,
Ju ¨rgen Neumann2, Elisabeth Kremmer6, Heinrich Leonhardt2,7, Matthias Mann4,7,
Jens Michaelis4,7,8, Lothar Schermelleh2,* and Sandra B. Hake1,7,*
1Department of Molecular Biology, Adolf-Butenandt-Institute, Ludwig-Maximilians-University Munich, 80336
Munich,2Department of Biology, Biozentrum, Ludwig-Maximilians-University Munich, 82152
Planegg-Martinsried,3Department of Biochemistry II, University Regensburg, 93053 Regensburg,4Department
of Proteomics and Signal Transduction, Max-Planck-Institute of Biochemistry, 82152 Martinsried,5Department
of Chemistry, Ludwig-Maximilians-University Munich,6Institute of Molecular Immunology, Helmholtz Center
Munich, German Research Center for Environmental Health,7Center for Integrated Protein Science Munich
(CIPSM), 81377 Munich and8Department of Physics, Ulm University, 89081 Ulm, Germany
Received November 8, 2011; Revised and Accepted March 9, 2012
The histone variant H2A.Z has been implicated in
many biological processes, such as gene regulation
and genome stability. Here, we present the identifi-
cation of H2A.Z.2.2 (Z.2.2), a novel alternatively
spliced variant of histone H2A.Z and provide a com-
prehensive characterization of its expression and
chromatin incorporation properties. Z.2.2 mRNA is
found in all human cell lines and tissues with highest
levels in brain. We show the proper splicing and
in vivo existence of this variant protein in humans.
Furthermore, we demonstrate the binding of Z.2.2 to
complexes and its active replication-independent
deposition into chromatin. Strikingly, various inde-
pendent in vivo and in vitro analyses, such as bio-
chemical fractionation, comparative FRAP studies
of GFP-tagged H2A variants, size exclusion chroma-
tography and single molecule FRET, in combination
with in silico molecular dynamics simulations,
consistently demonstrate that Z.2.2 causes major
structural changes and significantly destabilizes
nucleosomes. Analyses of deletion mutants and
chimeric proteins pinpoint this property to its
unique C-terminus. Our findings enrich the list of
known human variants by an unusual protein
belonging to the H2A.Z family that leads to the
least stable nucleosome known to date.
In the eukaryotic nucleus, DNA is packaged into chroma-
tin. The fundamental unit of this structure is the nucleo-
some consisting of a histone octamer (two of each H2A,
H2B, H3 and H4) that organizes ?147bp of DNA (1). In
order to allow or prevent nuclear regulatory proteins
access to the DNA, the chromatin structure has to be
flexible anddynamic. Several
controlled chromatin changes, one being the incorpor-
ation of specialized histone variants (2,3).
Variants of the histone H2A family are the most diverse
in sequence and exhibit distinct functions (4,5), com-
prising DNA damage repair, transcriptional regulation,
cell cycle control and chromatin condensation, though
the exact mechanisms of action are not fully understood
yet. Interestingly, the highest sequence variation among
H2A variants is found in the C-terminus, suggesting that
differences in structure and biological function might be
*To whom correspondence should be addressed. Tel: +49 89 2180 75435; Fax: +49 89 2180 75425; Email: email@example.com
Correspondence may also be addressed to Lothar Schermelleh. Tel: +44 1865 613264; Fax: +49 89 2180 74236;
Lothar Schermelleh, Department of Biochemistry, University of Oxford, South Park Road, Oxford OX1 3QU, UK.
Published online 29 March 2012Nucleic Acids Research, 2012, Vol. 40, No. 135951–5964
? The Author(s) 2012. Published by Oxford University Press.
This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/
by-nc/3.0), which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.
primarily attributed to this domain (6–9). One of the best
investigated and highly conserved but also functionally
enigmatic histone variant is H2A.Z. This variant is essen-
tial in most eukaryotes and possesses unique functions
(10,11). H2A.Z is involved in transcriptional regulation,
chromosome segregation and mitosis, acting in an
organism- and differentiation-dependent manner (12,13).
Furthermore, H2A.Z has been implicated in regulating
epigenetic memory (14) and in inhibiting read-through
H2A.Z might play a role in heterochromatin organization
(16), genome stability and chromosome segregation (17).
Despite many efforts to elucidate the exact biological
functions of H2A.Z, its roles have been and remain con-
troversial (18). Furthermore, deregulation of H2A.Z ex-
pression or localization seems to be connected to the
development of several neoplasias (19–23). Interestingly,
in vertebrates two non-allelic genes coding for two highly
similar H2A.Z proteins, H2A.Z.1 and H2A.Z.2, exist (24)
(previously named H2A.Z-1 and H2A.Z-2, prefixes were
changed due to a new histone variant nomenclature;
Talbert P.B., manuscript in preparation). They have a
common origin in early chordate evolution, are both
acetylated on the same N-terminal lysines (25–27) and
might be ubiquitinated on either one of the two
C-terminal lysines (28).
Here, we report the identification and structural char-
acterization of H2A.Z.2.2 (Z.2.2), an unusual alternative
splice form of H2A.Z. We show that Z.2.2 mRNA is
expressed to different degrees in all human cell lines and
tissues examined, with highest levels found in brain. Cell
biological and biochemical analyses consistently reveal the
presence of two distinct Z.2.2 populations within the cell.
The majority of Z.2.2 is freely dispersed in the nucleus,
whereas only a minority is stably incorporated into chro-
matin, most likely through the H2A.Z-specific p400/
NuA4/TIP60 (TIP60) and SRCAP chaperone complexes.
In vivo and in vitro analyses, in agreement with molecular
dynamic (MD) simulations, demonstrate that due to its
unique docking domain Z.2.2 chromatin incorporation
provide compelling evidence that a novel H2A.Z variant
exists in humans that plays a distinct and novel role in
chromatin structure regulation.
MATERIALS AND METHODS
See Supplementary Materials and Methods section for
Cell culture, transfection, FACS and cloning
Cell lines were grown in DMEM medium (PAA) supple-
mented with 10% FCS (Sigma) and 1% penicillin/strepto-
mycin at 37?C and 5% CO2. Cells were transfected using
FuGene HD (Roche Applied Science) according to the
manufacturer’s instructions. For details on cell selection,
FACS and cloning of expression plasmids see Sup-
plementary Materials and Methods section.
RNA expression analysis
RNA isolation and cDNA generation were performed as
previously described (29). Data were analyzed with the
advanced relative quantification tool of the Lightcycler
480 (Roche) software including normalization to HPRT1
and HMBS levels. Statistical evaluation was done using
t-test (two-tailed distribution, heteroscedastic). Total
RNA from different human tissues was commercially
acquired from: Applied Biosystems: normal lung, breast
and tumor breast, lung and ovary; Biochain: tumor lung,
breast, thyroid and bone, normal testis, cerebellum,
cerebral cortex, hippocampus, thalamus and total fetal
brain; amsbio: frontal lobe.
Histone extraction, RP–HPLC purification, sucrose
gradient, cellular fractionation and salt stability
Acid extraction of histones was done as previously
described (30). Histones were separated by RP–HPLC as
previously described (29). Fractions were dried under
vacuum and stored at ?20?C.
Details on MNase digest and sucrose gradient fraction-
ation can be found in Supplementary Materials and
Fractionation and salt stability experiments were
carried out as described previously (31–33) with minor
changes. For details on these methods see Supplementary
Materials and Methods section.
For the generation of a Z.2.2-specific antibody (aZ.2.2), a
GGEKRRCS of Z.2.2 was synthesized (Peptide Specialty
Laboratories GmbH) and coupled to BSA and OVA,
respectively. Development of Z.2.2-specific monoclonal
antibodies in rats was done as previously described (29).
The aZ.2.2 clone 1H11-11 of rat IgG1 subclass was
applied in this study. Rabbit aZ.2.2 antibody (rabbit 2,
bleed 3) was generated by the Pineda-Antiko ¨ rper-Service
company using the identical peptide epitope followed by
affinity purification. Following other primary antibodies
were used: aGAPDH (sc-25778, Santa Cruz), aGFP
(Roche Applied Science), aH2A (ab 13923, abcam), aH3
(ab1791, abcam) and aH2A.Z (C-terminus: ab4174,
abcam; N-terminus: ab18263, abcam). Following second-
ary antibodies and detection kits were used in imm-
unoblots: GFP-Z.2.2 and GFP-Bbd histones (aGFP)
and endogenous Z.2.2 (aZ.2.2) were detected using
HRP-conjugated secondary antibodies (Amersham) with
ECL advance (Amersham), all other proteins were
detected using ECL (Amersham). Detection of recombin-
ant proteins to evaluate histone stoichiometry of in vitro
IRDye-labeled secondary antibodies (LI-COR).
Fluorescence microscopy of cells and chromosomes
Preparation of cells and chromosome spreads for fluores-
cence microscopy was done as previously reported (34).
Wide-fieldfluorescence imagingwasperformed on
5952 Nucleic Acids Research, 2012,Vol.40, No. 13
a PersonalDV microscope system (Applied Precision)
(Olympus), CoolSNAP ES2 interline CCD camera (Pho-
tometrics), Xenon illumination and appropriate filtersets.
Iterative 3D deconvolution of image z-stacks was per-
formed withthe SoftWoRx
package (Applied Precision).
FRAP and exponential fitting
For details see Supplementary Materials and Methods
Stable isotope labeling with amino acids in cell culture
(SILAC) and mass spectrometric identification of
H2A.Z-specific chaperone complexes
HeLa cells expressing GFP-Z.2.1 or GFP-Z.2.2 were
SILAC labeled and nuclear extracts were prepared as
described before (35,36). High-resolution LC MS/MS
analysis was performed on an Orbitrap platform: details
on the experimental procedure are found in Supple-
mentary Materials and Methods section. Mass spectro-
metric (MS) operation and raw data analysis (37) are
described in Supplementary Materials and Methods
section. A complete list of all proteins identified is found
in Supplementary Table S1.
Immunoflurescence microscopy of cell cycle-dependent
GFP-Z.2.1 and GFP-Z.2.2 chromatin incorporation
Details on the experimental labeling (38) and microscopy
procedures are found in Supplementary Materials and
Expression of recombinant human histone proteins in
Escherichia coli, in vitro octamer and nucleosome
Histones were expressed, purified and assembled into
octamers as described (39) and mononucleosomes were
assembled on DNA containing the 601-positioning
sequence (40) according to (39,41). For details on
in vitro octamer and nucleosome reconstitution, see Sup-
plementary Materials and Methods section.
Single molecule Fo ¨ rster resonance energy transfer
(smFRET) single molecule burst analysis followed by the
removal of multi-molecular events (42–45) are described in
moleculeFo ¨ rster resonanceenergytransfer
Molecular modeling and MD simulations
The molecular modeling suite YASARA-structure version
9.10.29 was employed, utilizing the AMBER03 force field
(46) for the protein and the general amber force field
(GAFF) (47) throughout this study. The partial charges
were computed using the AM1/BCC procedure (48) as
implemented in YASARA structure (49). The starting
point for molecular modeling was the crystal structure of
a nucleosome core particle containing the histone variant
H2A.Z (PDB 1F66) (50). Missing side chain atoms were
added (Glu E 634). The missing N-terminal and C-terminal
residues were not modeled, although they might interact
with the neighboring DNA, e.g. in the case of missing
GKKGQQKTV). All structures were solvated in a water
box with 0.9% NaCl and neutralized (51). The structures
were initially minimized using steepest descent and
simulating annealing procedures. All deletions and muta-
tions were introduced sequentially using YASARA struc-
ture. MD simulations were carried out at 300K over 2.5ns
in an NPT ensemble using PME. All simulations were per-
formed four times using various starting geometries. The
2.5ns MD trajectories were sampled every 25ps, resulting
in 100 simulation frames per run, which were evaluated
after an equilibration phase of 500ps to derive statistical
averages and properties of the corresponding variant.
Finally, the interaction energy of H2A and H3 was
calculated from a simulation of the solvated octamer and
the isolated (H3–H4)2tetramer or the isolated respective
H2A.Z–H2B dimer. The interaction energy is calculated as
energy difference of the solvated octamer minus the
solvated (H3–H4)2tetramer and H2A.Z–H2B dimer.
Alternative splicing of H2A.Z.2 occurs in vivo
Two non-allelic intron-containing genes with divergent
promoter sequences that code for H2A.Z variants exist in
vertebrates (24,27). In humans, the H2A.Z.2 (H2AFV)
primary transcript is predicted to be alternatively spliced
(Supplementary Figure S1A). Using PCR and confirmed
by sequencing we detected not only H2A.Z.2.1 (Z.2.1) but
also H2A.Z.2.2 (Z.2.2) mRNA, though none of the other
splice variants in human cells (Supplementary Figure S1B)
showing that the H2A.Z.2 primary transcript is indeed al-
ternatively spliced in vivo. Interestingly, database searches
found Z.2.2 mRNA to be predicted in chimpanzee (Pan
(Nomascus leucogenys) as well. In addition, the coding
sequence of the unique exon 6 was present downstream
of the H2AFV locus of several other primate genomes,
such as gorilla (Gorilla gorilla gorilla), macaque (Macaca
mulatta), orangutan (Pongo abelii) and white-tufted-ear
marmoset (Callithrix jacchus) (data not shown). In all of
these primates, with the exception of marmoset, the result-
ing protein sequence, if translated, is 100% identical to the
unique human Z.2.2 peptide. Further searches revealed
that the genomes of horse, and to a certain extent also
rabbit and panda bear, contain sequences downstream of
their H2AFV loci that could, if translated, lead to proteins
with some similarities to human Z.2.2, although they are
much more divergent and even longer (rabbit, panda bear).
Due to these differences, it is highly likely that those species
do not express a Z.2.2 protein homolog. Surprisingly, we
could not detect Z.2.2-specific sequences in mouse, rat or
other eukaryotic genomes, suggesting that Z.2.2 might be
Next, we wanted to determine to what degree all three
H2A.Z mRNAs are expressed in different human cell lines
and tissues and performed quantitative PCR (qPCR).
Nucleic Acids Research, 2012,Vol.40, No. 13 5953
Z.2.2 mRNA was present to different degrees in all human
cell lines and tissues tested, though less abundant than Z.1
and Z.2.1 mRNAs that are expressed in similar amounts
(Supplementary Figure S1C and D). Z.2.2 constituted
between 5% and 15% of total Z.2 transcripts in all cell
lines and tissues, with the exception of brain, where it was
statistically significant upregulated (p=1.7?10?4; Figure
1A). In some regions of this particular organ Z.2.2 ac-
counted for up to 50% of all Z.2 transcripts pointing
toward an exciting brain-specific function of this novel
Encouraged by our findings we next investigated
whether the endogenous protein is present in vivo. The
distinctive feature of Z.2.2 is its C-terminus that is 14
amino acids shorter and contains six amino acids differ-
ences compared to Z.2.1 (Figure 1B). Due to this
shortened C-terminal sequence, ubiquitination sites at
positions K120 and K121 (28) and part of the H3/H4
docking domain (50) are lost in Z.2.2. We generated
antibodies against Z.2.2’s unique C-terminal amino acids
(aZ.2.2) in rats and rabbits and confirmed their specificity
in immunoblots (IB) with recombinant Z.2.1 and Z.2.2
proteins (Supplementary Figure S1E and data not
shown). We extracted histones from several human and
mouse cell lines, purified them by reversed phase–high
performance liquid chromatography (RP–HPLC) and
analyzed obtained fractions by IB (Figure 1C). Using
aZ.2.2 (polyclonal rabbit), we observed a signal of the
calculated weight of Z.2.2 that elutes shortly before Z.1-
and Z.2.1-containing fractions in all human samples.
Similar results were obtained with a monoclonal aZ.2.2
rat antibody (data not shown). In agreement with the
finding that Z.2.2-specific exon 6 sequences are mainly
restricted to primate genomes, we could detect Z.2.2
protein in human but not in mouse cells (Figure 1C). In
summary, our data show that Z.2.2 protein indeed exists
in vivo, albeit at a low expression level.
GFP-Z.2.2 is partially incorporated into chromatin
Having demonstrated the existence of this novel variant
in vivo, we next sought to clarify whether Z.2.2 constitutes
a bona fide histone by being part of the chromatin struc-
ture. Due to high background of all our aZ.2.2 antibodies
in IB with cell extracts (data not shown), we generated
A G G K A G K D S G K A K T K A V S R S Q R A G L Q F P V G R I H R H L K S R T
A G G K A G K D S G K A K A K A V S R S Q R A G L Q F P V G R I H R H L K T R T
A G G K A G K D S G K A K A K A V S R S Q R A G L Q F P V G R I H R H L K T R T
T S H G R V G A T A
T S H G R V G A T A
T S H G R V G A T A
I T P R H L Q L A I R G D E E L D S L I
I T P R H L Q L A I R G D E E L D S L I
I T P R H L Q L A I R G D E E L D S L I
A V Y S A A I L E Y L T A E V L E L A G N A S K D L K V K R
A V Y S A A I L E Y L T A E V L E L A G N A S K D L K V K R
A V Y S A A I L E Y L T A E V L E L A G N A S K D L K V K R
K A T I A G G G V I P H I H K S L I G K K G
K A T I A G G G V I P H I H K S L I G K K G
K A T I A G G E K R R C S
Q Q K T V
Q Q K T A
H3/H4 docking domain
Z.2.2 (% Z.2 transcripts)
lung tumor 1lung tumor 2
breast tumor 1
breast tumor 2
total fetal brain
seuss i tn i a r bseuss i t n i a r bnonsen i l l l ec
reversed phase-HPLC fractions
Figure 1. Identification of Z.2.2. (A) qPCR with cDNA from different human cell lines and tissues using primers specific for Z.2.1 and Z.2.2. Data
were normalized to HPRT1 and HMBS expression levels. Controls generated without reverse transcriptase (no RT) were used to assess amplification
threshold. Shown are the levels of Z.2.2 mRNA as percentages of total Z.2 transcripts (Z.2.1+Z.2.2). For an evaluation of absolute expression levels
see Supplementary Figure S1C and D. (B) Amino acid alignment of human Z.1, Z.2.1 and Z.2.2 proteins using ClustalW Alignment (MacVector
10.0.2). Identical amino acids are highlighted in dark gray, similar amino acids in light gray and changes are set apart on white background. Known
acetylation sites are depicted with stars and ubiquitination sites with circles. A schematic representation of the secondary structure of Z.1 and Z.2.1 is
shown below the alignment, including depiction of the H3/H4 docking domain (50). M6 and M7 boxes indicate regions important for H2A.Z-specific
biological functions in D. melanogaster (60). (C) IB analyses of RP–HPLC purified fractions from different human (HEK293, HeLa, HeLa Kyoto
and U2OS) and mouse (NIH3T3) cell lines using a polyclonal rabbit aZ.2.2 and aH2A.Z (aZ, C-terminal) antibodies. Recombinant Z.2.2 protein
(rZ.2.2) was loaded in the first lane as positive control for aZ.2.2 antibody. Similar results were obtained when using a monoclonal rat aZ.2.2
antibody (data not shown).
5954Nucleic Acids Research, 2012,Vol.40, No. 13
HeLa Kyoto cell lines stably expressing GFP-tagged H2A
Expression levels of GFP-tagged histone variants were
determined by FACS (Supplementary Figure S2A) and
by comparing expression levels of GFP-tagged variants
with endogenous H2A.Z proteins in IB (Supplementary
Figure S2B). GFP-Z.1 and -Z.2.1 were expressed in
similar amounts as the endogenous H2A.Z protein, and
GFP-Z.2.2 expression levels were considerably lower than
those of other GFP-tagged H2A variants, with the excep-
tion of GFP-H2A.Bbd (Barr body deficient; Bbd). These
data show that all GFP-H2A variants were not expressed
in abnormal amounts in cell clones used for further
In fluorescence microscopy, GFP-Z.2.2 exhibited a sole
but rather diffuse nuclear distribution similar to GFP-
Bbd, suggesting that both variants might have similar
properties (Figure 2A). Additionally, GFP-Z.2.2 was
detected in condensed mitotic chromosomes, with a faint
residual staining in the surrounding area (Figure 2B), sug-
gesting that it is incorporated into chromatin, although to
a lesser extent than other GFP-H2A variants. To discrim-
inate between a potential non-specific DNA binding and
nucleosomal incorporation of Z.2.2 we purified mono-
GFP-Z.2.2 was detected by IB in fractions containing
mononucleosomes (Figure 2C), suggesting that Z.2.2 is
indeed a nucleosomal constituent.
To analyze the extent of Z.2.2 chromatin incorporation
in more detail, we isolated soluble (sol) and chromatin
(chr) fractions from HK-GFP cells. IB analyses revealed,
as expected, that similar to GFP-Bbd, GFP-Z.2.2 is pre-
dominantly nuclear soluble, with only minor amounts
present in chromatin (Figure 3A). Based on fractionation
and fluorescence imaging results, we hypothesized that
this novel variant behaves in a different manner as
compared to other H2A variants with regard to chromatin
exchange mobility in vivo. To test this prediction, we per-
(FRAP) experiments with HK-GFP cells. Using spinning
disk confocal microscopy we monitored the kinetic
behavior of H2A variants with variable intervals over
2min (short-term) up to several hours (long-term) after
bleaching a 5mm?5mm square nuclear region (Figure
3B and Supplementary Figure S3). As expected, GFP
GFP-H2A, -Z.1 and -Z.2.1 showed a slow recovery,
which is in agreement with a previous report (52).
GFP-Bbd has been described to exhibit low nucleosomal
stability and a fast FRAP kinetic (53), which we also
observed in our experiment. Interestingly, GFP-Z.2.2
showed an even faster recovery than GFP-Bbd, with
?80% of initial fluorescence reached after 1min. Careful
assessment and bi-exponential fitting of FRAP data
allowed us to also calculate ratios of fractions with fast,
intermediate and slow recovery and their respective
half-time of recovery (t1/2) as an indication of exchange
rate thereby revealing quantitative differences between
Z.2.2 and other H2A variants (Figure 3D, Supplementary
Figure S3C and E). For Z.2.2 as well as for Bbd, we
identified a fast fraction of unbound or very transiently
interacting molecules (78%, t1/2?1.1s and 52%, t1/2
?2.5s, respectively; for comparison GFP t1/2 ?0.4s)
and a substantially slower fraction with a t1/2 in
the range of 7–9min. In contrast, GFP-H2A, -Z.1
and -Z.2.1 showed no fast mobile fraction but intermedi-
ate slow fractions with t1/2in the range of 8–17min and a
second even slower class exchanging with a t1/2of a few
hours. For comparison, we measured the linker histone
H1.0 (54–57) and the histone binding protein HP1a
(58,59), both DNA-associated proteins, and found that
HP1a shows an overall much faster recovery than all
H2A variants. In contrast to Z.2.2 and Bbd, no
unbound fraction of H1.0 was detected. More import-
antly, with regards to the bound Z.2.2 and Bbd fractions
overall H1.0 showed a faster recovery, arguing against
an unspecific DNA-association of Z.2.2 and Bbd. In
agreement with cell biological and biochemical analyses,
Z.2.2 BbdGFPH2A Z.1 Z.2.1
Figure 2. Z.2.2 localizes to the nucleus and is partially incorporated
into chromatin. (A) Fluorescence imaging of stably transfected HeLa
Kyoto cells shows nuclear localization of all GFP-H2A variants
(middle). DNA was counterstained with DAPI (top). Overlay of both
channels in color is shown at the bottom (Merge; GFP: green, DAPI:
blue). Scale bar=5mm. (B) Deconvolved images of metaphase spreads
of HeLa Kyoto cells stably expressing GFP-H2A variants (middle).
Merged images in color are shown below (GFP: green; DAPI: blue).
Scale bar=10mm. (C) Chromatin from HeLa Kyoto cells stably ex-
pressing GFP-Z.2.2 was digested with MNase followed by a purifica-
tion of mononucleosomes using sucrose gradient centrifugation.
Isolated DNAfrom subsequent
analyzed by agarose gel electrophoresis (left). Fractions containing
pure mononucleosomes (marked with asterisk) were combined and
analyzed by IB (right) using aGFP antibody for the presence of
GFP-Z.2.2 (top), and aH3 (bottom).
Nucleic Acids Research, 2012,Vol.40, No. 135955
these data clearly demonstrate that a large fraction of the
splice variant Z.2.2 is very rapidly exchanged or chromatin
unbound, and a minor population is incorporated into
Z.2.2’s unique docking domain, but not its shortened
length, weakens chromatin association
The functional importance of specific C-terminal domains
of H2A.Z has previously been demonstrated by nucleo-
somal structure analyses (7,50) and in rescue experiments
in flies (60). Since the C-terminus of Z.2.2 is shorter and
has a distinct sequence when compared to Z.1 and Z.2.1, it
is not clear which of these features determines Z.2.2’s
Therefore, we generated deletion and domain-swap
constructs (Supplementary Figure S3D) for FRAP experi-
Supplementary Figure S3B). Surprisingly, C-terminal
(Z.2.1113) to mimic the shortened length of Z.2.2 did not
affect their original mobility in short-term and only
modestly in long-term FRAP. Hence, the mere shortening
of the C-terminus is not sufficient to weaken stable
To investigate whether the unique six C-terminal amino
acids of Z.2.2 are sufficient to generate highly mobile
proteins, we created a further C-terminally truncated
GFP-H2A construct (H2A105) and added the Z.2.2
specific C-terminal six amino acids (H2A105+CZ.2.2).
Although both mutant constructs are slightly more
mobile than H2A111, their indistinguishable recovery
kinetics demonstrate that the unique six C-terminal
amino acids of Z.2.2 alone are not sufficient to cause its
extreme mobility in vivo.
To explore whether the complete Z.2.2 docking domain
is able to induce high-protein mobility, we transferred the
respective domain of either Z.2.1 (amino acids 91–127) or
Z.2.2 (amino acids 91–113) onto a C-terminally truncated
H2A (H2A88+CZ.2.1 and H2A88+CZ.2.2, respectively).
Interestingly, only the docking domain of Z.2.2, but not
the one of Z.2.1, confers high mobility. In conclusion,
the six unique C-terminal amino acids of Z.2.2 prevent
chromatin-association of a large proportion of this
protein, but only when present in the context of
the preceding H2A.Z-specific docking domain sequence.
Z.2.2 interacts with H2A.Z-specific TIP60 and SRCAP
chaperone complexes and is deposited into chromatin
outside of S-phase
Our so far obtained data strongly imply that at least a
minor amount of the cellular Z.2.2 protein is incorporated
into nucleosomes. Since previous studies have shown that
chromatin remodelers specifically exchange canonical
H2A–H2B with H2A.Z–H2B dimers within nucleosomes
(10,61), we wondered if such complexes are also able to
actively deposit Z.2.2 into chromatin. HK cells and HK
cells stably expressing GFP-Z.2.1 or -Z.2.2 were SILAC
labeled, soluble nuclear proteins isolated, GFP-tagged
Z.2.1 and Z.2.2-associated proteins precipitated using
GFP nanotrap beads and identified by quantitative mass
spectrometry (Figure 4 and Supplementary Table S1 for a
complete list of all identified proteins). Whereas the
majority of proteins are background binders clustering
Figure 3. The majority of Z.2.2 protein is nuclear soluble and highly
mobile in a sequence-dependent manner. (A) HK-GFP cells were sub-
jected to biochemical fractionation. Fractions sol and chr of identical
cell equivalents were probed in IB with aGFP (top), aH2A (middle)
and aGAPDH (bottom). (B) FRAP quantification curves of average
GFP signal relative to fluorescence intensity prior to bleaching are
depicted for GFP, GFP-tagged wild-type H2A variants, linker histone
H1.0 and heterochromatin protein 1a (HP1a). Mean curves of 10–29
cells are shown for each construct. Error bars are omitted for clarity.
(C) FRAP quantification curves similar to (B) are depicted for GFP,
GFP-tagged wild-type H2A, Z.2.1, Z.2.2 and mutant constructs.
(D) Quantitative evaluation of FRAP curves. Plot shows calculated
mobility fraction sizes of different wild-type and mutant H2A variant
constructs, as well as H1.0 and HP1a, based on bi-exponential fitting of
FRAP data. Error bars indicate SD (see Supplementary Figure S3 for
long-term FRAP and for numerical values).
5956Nucleic Acids Research, 2012,Vol.40, No. 13
around 0, specific interactors can be found on the right
side having a high ratio H/L or ratio L/H for Z.2.1 and
Z.2.2, respectively. In accordance with previous studies
(62–65), we found GFP-Z.2.1 to be part of two major
complexes, the SRCAP and the p400/NuA4/TIP60
(TIP60) complexes (Figure 4A), as we were able to
detect all of their thus far identified members, with the
exception of actin, as significant outliers. Interestingly,
GFP-Z.2.2 also associated with both SRCAP and TIP60
complexes (Figure 4B), showing an almost identical
binding composition as GFP-Z.2.1 (Figure 4C). These
results strongly imply that Z.2.2 is, similar to other
through specific chaperone complexes.
Based on these results, we predicted that Z.2.1 and Z.2.2
should be incorporated into chromatin in a highly similar
spatial manner. Since both SRCAP and TIP60 chaperone
complexes are evolutionary conserved between different
species, we tested mouse C127 cells that do not express
endogenous Z.2.2 for their ability to deposit GFP-Z.2.2.
Hereby we should be able to distinguish whether SRCAP
and TIP60 complexes are sufficient for deposition, or if
GFP-Z.2.1 and -Z.2.2 were transiently expressed in C127
cells, S-phase stages highlighted by EdU-incorporation
and co-localization patterns visualized by fluorescence
microscopy (Figure 5). GFP-Z.2.1 and -Z.2.2 showed an
almost identical chromatin localization and deposition
pattern, suggesting that Z.2.2 is, like Z.2.1, deposited
through SRCAP and TIP60 complexes. In accordance
with a recent study, we observed an enrichment of both
H2A.Z variants in facultative heterochromatin regions in
interphase nuclei (66). Surprisingly, although H2A.Z is
expressed in all cell cycle phases (67), and GFP-Z.2.1
and -Z.2.2 expression is driven by a constitutive active
promoter, chromatin deposition of both proteins is
underrepresented at replication foci. This result underlines
our findings that Z.2.2 interacts with all members of both
TIP60 and SRCAP complexes and is actively and not pas-
sively deposited, as would have been the case during
S-phase when nucleosomes are highly exchanged.
Structural changes in Z.2.2’s C-terminus prevent histone
octamer folding and enhance DNA breathing on
structurally destabilized nucleosomes
Our findings thus far imply that Z.2.2 is incorporated
into nucleosomes and most likely targeted by TIP60 and
Figure 4. Z.2.2 associates with H2A.Z-specific SRCAP and TIP60
chaperone complexes. GFP-pull-downs for H2A.Z-specific chaperone
complexes are shown. HK cells stably expressing GFP-Z.2.1 (A) and
GFP-Z.2.2 (B) were SILAC-labeled and subjected to single-step affinity
purifications of soluble nuclear proteins in a ‘forward’ (GFP-Z.2.1)
or ‘reverse’ (GFP-Z.2.2) pull-down using GFP nanotrap beads. In
Figure 4. Continued
each panel the ratio of the identified proteins after MS is plotted.
Proteins known to interact with H2A.Z are indicated in the following
way: members of the SRCAP complex in red, members of the TIP60
complex in blue and shared subunits in purple. Potential novel
H2A.Z-interacting proteins are shown as green dots (‘other outliers’)
and are distinguished from background binders (gray dots) and con-
taminants (yellow dots). See also Supplementary Table S1 for a list of
all identified proteins. (C) List of the SRCAP and TIP60 complex
members and their normalized binding intensity to Z.2.1 or Z.2.2.
Note that for comparison reasons the obtained H/L ratios of
GFP-Z.2.2 binders (numbers in brackets) were calculated in the corres-
ponding L/H ratios. See also Supplementary Table S1 for a list of all
identified proteins and their normalized H/L ratios.
Nucleic Acids Research, 2012,Vol.40, No. 135957
SRCAP complexes. Then why does a large fraction of the
cellular Z.2.2 protein pool shows a high mobility and is
freely dispersed in the nucleus? One plausible possibility is
that Z.2.2 severely destabilizes nucleosomes due to its di-
vergent C-terminal docking domain and is hence rapidly
exchanged. To test this hypothesis, we used an in vitro
reconstitution system.Recombinant humanH2A
variants together with H3, H2B and H4 (Supplementary
Figure S4A) were refolded by dialysis, and formed
complexes purified by size exclusion chromatography. As
expected, both H2A and Z.2.1 containing samples readily
formed histone octamers (Figure 6A, solid lines). Bbd
served as a negative control, because it has been
demonstrated to not form octamers under these condi-
tions (41), a result we also observed (Figure 6A left,
dotted line). Interestingly, in accordance with our FRAP
data, Z.2.2 behaved like Bbd in that it only formed Z.2.2–
H2B dimers, but did not complex together with (H3–H4)2
tetramers to generate octamers (Figure 6A right, dotted
line), which was further confirmed by SDS–PAGE
analyses of the separate fractions (Figure 6B). Thus, like
for Bbd the incorporation of Z.2.2 destabilizes the inter-
face between Z.2.2–H2B dimers and (H3–H4)2tetramers
in a C-terminal sequence dependent manner (Sup-
plementary Figure S4B and C). In conclusion, the Z.2.2
Although no Z.2.2 containing histone octamers could
be generated in vitro, our results using GFP-Z.2.2 strongly
suggest that Z.2.2 can be part of nucleosomes. To test this
in vitro and to evaluate the effect of Z.2.2 on nucleosome
stability, we reconstituted mononucleosomes by mixing
Z.2.2–H2B dimers, (H3–H4)2tetramers and DNA con-
taining a ‘Widom 601’ DNA positioning sequence in a
2:1:1 ratio. As controls, we reconstituted H2A or Z.2.1
containing nucleosomes by mixing octamers and DNA
in a 1:1 ratio. As expected, analysis of all nucleosomes
by native PAGE showed a single band before and after
heat shift (Figure 7A), indicating a unique position on the
‘Widom 601’ DNA template. Purification of nucleosomes
from a native gel and analysis of the protein content by
GFP EdUDAPIGFP/ EdUmergeGFPEdU DAPIGFP/ EdUmerge
Figure 5. Z.2.1 and Z.2.2 are actively deposited into chromatin and are under-represented at replication foci. C127 cells transiently expressing
GFP-Z.2.1 (left) and -Z.2.2 (right) were pulse labeled with EdU to visualize replication foci and to identify S-phase stages. DNA was counterstained
with DAPI and analyzed by wide-field deconvolution microscopy. To remove the unbound fraction in GFP-Z.2.2 expressing cells, an in situ
extraction was performed prior to fixation. Cells in early, middle and late S-phases were distinguished due to their characteristic differential EdU
replication labeling patterns of eu- and heterochromatic regions. Merged images in color are shown alongside (GFP: green; EdU: red; DAPI: blue).
204060 80100 120
Figure 6. Z.2.2 does not constitute stable histone octamers with H2B,
H3 and H4 in vitro. (A) Size exclusion chromatography of refolding
reactions using recombinant human H3, H4 and H2B proteins to-
gether with either H2A (solid line) or Bbd (dashed line) (left overlay)
or with either Z.2.1 (solid line) or Z.2.2 (dashed line) (right overlay).
Peaks corresponding to aggregates, histone octamers, tetramers or
dimers are labeled respectively. (B) Fractions corresponding to H2A-
containing octamers, Bbd-containing tetramers and dimers (left) or
Z.2.1-containing octamers and Z.2.2-containing tetramers and dimers
(right) were analyzed by 18% SDS–PAGE and stained with Coomassie
5958Nucleic Acids Research, 2012,Vol.40, No. 13
showed that Z.2.2 was indeed incorporated into nucleo-
evaluated for their resistance to MNase cleavage as an
indicator of stably organized nucleosomes and to deter-
mine nucleosomal DNA
Supplementary Figure S5). We observed fragments corres-
ponding to protected nucleosomal DNA with the length of
146bp for all variant nucleosomes tested. The appearance
of smaller, subnucleosomal fragments indicates that DNA
breathing occurred (68). Interestingly, DNA of Z.2.2
nucleosomes is less protected, since subnucleosomal frag-
ments were obtained at lower MNase concentrations than
with H2A or Z.2.1 nucleosomes. Additionally, at higher
MNase concentrations a stable DNA fragment of about
120bp was most abundant for Z.2.2 nucleosomes
(Supplementary Figure S5), indicating that this might be
the preferred DNA length wrapped around this octamer.
These data suggest that increased DNA breathing occurs
in Z.2.2 nucleosomes, which as a result might be less
stable. To quantify nucleosome stability in vitro we
measured salt-dependent changes in nucleosome structure
using smFRET (69). In line with the results presented
above, Z.2.2 containing recombinant nucleosomes lost
their compact structure at lower salt concentrations than
Z.2.1 or H2A-containing ones (Figure 7D). To investigate
whether the observed Z.2.2-dependent nucleosome desta-
bilization is true in the context of chromatin, we isolated
chromatin from HK cells expressing GFP-H2A variants
and incubated it with buffer containing increasing
amounts of salt. Histones that remained stable chroma-
tin components were precipitated and detected by IB
Z.2.2-containing nucleosomes disintegrated between 200
and 400mM NaCl, and were therefore even less stable
than Bbd-containing ones. In summary, incorporation
of Z.2.2 leads to a severely reduced nucleosome stability
due to C-terminal sequence dependent changes in its
docking domain and subsequent loss of its interaction
with histone H3.
Our FRAP data suggest that the Z.2.2 C-terminal
amino acids might have a direct influence on the nucleo-
somal structure by affecting interactions with DNA and/
or adjacent histones. Based on the existing structural data
(50), we performed MD simulations of nucleosomes con-
In addition, we also included the deletion mutant
Z.2.1113, which did not show any change in short-term
FRAP (Figure 3C), but some increase in mobility in
long-term FRAP (Supplementary Figure S3B) in our
assay. These in silico models revealed that changes in the
C-terminus of H2A.Z strongly affect its protein structure
(Figure 8A). Strikingly, different statistical descriptors
over the MD-trajectory like distance and mobility
(B-factor) show in contrast to Z.1 and Z.2.1113unique
properties for the Z.2.2 tail. Only Z.2.2 leads to a substan-
tial structural change in the C-terminus resulting in an
increased distance to histone H3, which in turn makes a
hydrogen bond interaction between peptide backbone NH
of Cys112 in Z.2.2 and the oxygen in the Gln55 side chain
in H3 impossible (Figure 8B). Additionally, an increase in
the B-factor for Z.2.2 indicates a substantially enhanced
mobility of Z.2.2’s C-terminus (Figure 8C). We also
calculated the Z.2.2–H3 interaction energy and observed
a switch from negative to positive values in the case of
Z.2.2 suggesting that this histone variant destabilizes the
nucleosome (Figure 8D). In summary, these data suggest
that the C-terminal sequence of Z.2.2 leads to a more
dynamic structure that in turn loses binding to histone
50 200 400 600input
0300 400 500600700
fraction closed nucleosomes
(normalized to 0 mM NaCl)
Figure 7. Z.2.2-containing nucleosomes are less resistant to MNase
digestion and increased ionic strength. (A) H2A, Z.2.1 or Z.2.2 con-
taining nucleosomes were assembled on DNA by salt gradient depos-
ition, incubated at 4?C or 37?C to evaluate DNA positioning and
separated by a native 5% PAGE gel. (B) Agarose-gel-electro-eluted
material from (A) was analyzed by 18% SDS–PAGE and Coomassie
stained to evaluate stoichiometry of histones after nucleosome assembly
(top). Stars indicate H2A variants that were used for assembly. Further
evaluation of histone stoichiometry after nucleosome assembly was
done by IB using a LI-COR instrument (bottom). Assembled nucleo-
somes containing Z.2.1 or Z.2.2 were immunoblotted and the amount
of histones was visualized using an aH3 antibody (top) and an
N-terminal aZ antibody (recognizes all H2A.Z variants, bottom).
(C) Mononucleosomes containing either H2A, Z.2.1 or Z.2.2 were
digested with increasing concentrations of MNase and extracted
DNA was separated using Bioanalyzer. Stars indicate DNA length of
146bp. For detailed electropherogram analyses of fragment lengths in
each sample see Supplementary Figure S5. (D) Mononucleosomes
containing either H2A, Z.2.1 or Z.2.2 histones together with double
dye labeled DNA were incubated with increasing amounts of salt.
smFRETmeasurement values of
normalized to 0mM NaCl. Error bars represent SEM of six measure-
ments. (E) Chromatin from HK-GFP cells was isolated and incubated
with increasing amounts of salt. Chromatin-bound histones were
precipitated and detected by IB using aGFP antibody. Staining with
aH2A was used as loading control.
each saltconcentration were
Nucleic Acids Research, 2012,Vol.40, No. 135959
H3 and destabilizes the nucleosomal structure, providing a
reasonable explanation for the observed in vivo and
in vitro data.
In this work, we have identified a previously unknown
histone H2A.Z variant and provide a comprehensive
characterization of its nucleosomal properties. This alter-
natively spliced variant, Z.2.2, is present to different
degrees in all human cell lines and tissues investigated,
with a significant enrichment in brain. Z.2.2 contains a
shortened and in six amino acids divergent C-terminus
compared to Z.1 and Z.2.1 that is necessary, but not
sufficient, to weaken chromatin association. Only in
the context of the unique Z.2.2 docking domain does
the C-terminal sequence negatively affect nucleosome
stability in vitro and in vivo. To our knowledge,
Z.2.2 has the strongest destabilizing effect on nucleosomal
structure compared to other histone H2A variants
reported to date.
Only one other histone variant, macroH2A, has been
shown thus far to be alternatively spliced (70). Here, like
our observation with H2A.Z, two independent genes
mH2A1 and mH2A2 exist in mammals, with only
mH2A1 being alternatively spliced resulting in functional
different proteins (71). In our study, we demonstrate that
the human H2A.Z.2 (H2AFV) primary transcript is alter-
natively spliced generating Z.2.1 and Z.2.2 mRNAs and
proteins. These observations suggest that Z.2.2 is tightly
regulated in a tissue-specific manner through alternative
splicing and/or RNA stability. Our findings now raise the
intriguing possibility that alternative splicing of histone
variants might not be rare but more common than previ-
ously thought. If true, it will be of interest to reevaluate
other intron-containing histone variant genes with regard
to their possible alternative transcripts and protein
Bioinformatic genome analyses revealed the existence of
Z.2.2-specific sequences only in humans, old and new
world primates and to some extend in other mammals,
with the exclusion of mouse, rat and even lower eukary-
otes. It remains to be seen, whether Z.2.2’s evolution is
indeed limited to primates only. Primate-specific gene
products have been often identified in human brain
and reproductive tissues (72), supporting the notion that
distance H3 αN to
H2A.Z C-terminus (Å)
chain C chain G
Figure 8. Unique Z.2.2 C-terminal amino acids cause significant changes in protein and nucleosome structure. (A) In silico models of Z.1, Z.2.1113
and Z.2.2 C-terminal C-chains (yellow; from amino acids 84 to C-terminus, including the complete docking domain) together with the E-chain of
histone H3 (blue; amino acids 33–60, including aN-helix). Side (left) and frontal views (right) of four MD simulations are shown respectively.
See Supplementary Figure S7 for complete in silico model of H2A.Z-containing nucleosome. (B) Simulated distances between peptide backbone
NH of amino acids 112 in H2A.Z (His or Cys, respectively) variants and the oxygen in the Gln55 sidechain in H3 based on in silico nucleosome
models containing either Z.1 (white), Z.2.1113(light gray) or Z.2.2 (dark gray) proteins. Error bars represent SD of four independent simulations.
(C) Simulated mobility measuring B-factor values between amino acids 108 and 113 in respective H2A.Z variant C-termini. Error bars represent SD
of four independent simulations. (D) Simulated interaction energy between tetramer versus respective H2A.Z variant-containing dimers.
5960 Nucleic Acids Research, 2012,Vol.40, No. 13
their RNAs and proteins might be essential to adaptive
changes leading to human development and further specu-
lates that primate-specific genes might be important in re-
productive function and disease. Since we have found
Z.2.2 transcripts to be strongly enriched in brain samples
of higher brain function in comparison to other tissues and
cell types, it will be of great interest to determine in future
studies, if this novel variant might play an important
functional role in this particular organ. These observations
also raise the interesting question of how alternative
splicing and/or differential stability of H2AFV transcripts
are tissue specifically regulated.
Another intriguing feature of Z.2.2 is its influence on
nucleosome stability. Although Z.2.2 localizes exclusively
to the nucleus, only a minor proportion is stably
incorporated into chromatin. The only other exception
in humans known thus far is Bbd, which has previously
been demonstrated to destabilize the nucleosome structure
(41,53,73). Bbd, similar to Z.2.2, is a shorter H2A variant
with an unusual C-terminus and a considerable different
primary histone fold sequence that might explain its
ability to destabilize nucleosomes. In agreement, a recent
study demonstrated that the incomplete C-terminal
docking domain of Bbd results in structural alterations
in nucleosomes and that those are in turn associated
with an inability of the chromatin remodeler RSC to
both remodel and mobilize nucleosomes (8). Z.2.2, on
the other hand, is identical to Z.2.1, except that its
C-terminus is 14 amino acids shorter and in six amino
acids altered. How can this small change in Z.2.2’s
primary sequence lead to such drastic effects on chromatin
We show that Z.2.2 can be part of a bona fide nucleo-
some and that it interacts with the H2A.Z-specific TIP60
and SRCAP chaperone complexes. These complexes have
been shown to catalyze the exchange of H2A–H2B dimers
with H2A.Z–H2B dimers in nucleosomes and our finding
therefore suggests that both complexes are also involved in
an active chromatin incorporation of Z.2.2. Supporting
this idea is the observation that both Z.2.1 and Z.2.2 are
incorporated into chromatin in a replication-independent
manner, even in mouse cells that do not express endogen-
ous Z.2.2. Both H2A.Z variants are not primarily de-
posited at replication foci, not even in middle S-phase
when facultative heterochromatin is replicated, where the
majority of the H2A.Z protein pool is found in interphase
cells (66). Our findings are in agreement with a model
proposed by Hardy and Robert, in which H2A.Z
variants are randomly deposited into chromatin by
specific chaperone complexes in a replication-independent
manner coupled to a subsequent targeted H2A.Z deple-
tion (74). As a consequence, an enrichment of H2A.Z at
non-transcribed genes and heterochromatin regions over
several cell generations can be observed (74). It might be
possible that INO80 facilitates this eviction function, as it
has been shown to exchange nucleosomal H2A.Z–H2B
dimers with free H2A–H2B dimers (75). It will be of
interest in future studies to determine whether Z.2.2
exchange is subjected to a similar mechanism. Taken
together, our findings strongly imply that Z.2.2 is
actively deposited into chromatin through the interaction
Nevertheless, a large fraction of Z.2.2 protein is not chro-
matin bound and we have mapped the region crucial for
high FRAP mobility to its docking domain. In addition to
Z.2.2’s unique C-terminal amino acids this region spans
the highly conserved acidic patch responsible for depos-
ition (76), the M6 region that is functionally essential in
fly H2A.Z (60) and required for the interaction with
the SWR1 complex in yeast (77). Strikingly, in silico simu-
lation of Z.2.2 predicted dynamic structural changes
that in turn weaken interaction with histone H3 and
destabilize the nucleosome structure. Such a gross struc-
tural alteration explains why Z.2.2 is not able to form
stable octamers in vitro and leads to enhanced DNA
breathing in a nucleosomal context. Hence, Z.2.2 incorp-
oration into chromatin disrupts nucleosomes more easily
and supports a model in which Z.2.2 is more rapidly
exchanged than Z.2.1.
What functional outcome might Z.2.2 cause when
incorporated into chromatin? And how is the variant
composition of Z.2.2 containing nucleosomes? It has
been shown that a special class of nucleosomes contain-
ing both H2A.Z and H3.3 variants exists in humans (78).
These nucleosomes are enriched at promoters, enhancers
and insulator region and promote the accessibility of
transcription factors to these DNA regions (78), most
likely due to their extreme sensitivity to disruption (79).
Since these studies nicely demonstrate that differential
nucleosome stabilities due to the incorporation of
different histone variants influence transcriptional regula-
tion, it is tempting to speculate that Z.2.2 might also
affect chromatin-related processes. Future experiments
will shed light on Z.2.2 function(s), especially with
regard to its increased expression in human brain areas,
and explain why and where nucleosomal destabilization is
needed. This is of particular interest, since Bbd that also
leads to nucleosomal destabilization is almost exclusively
present in testis (80–82) in contrast to the apparently ubi-
quitously expressed Z.2.2, possibly pointing toward
distinct roles of both destabilizing H2A variants in dif-
ferent tissues. Our data suggest that additional interest-
ing, yet unidentified, histone variants may exist and await
evolutionaryconserved chaperone complexes.
Supplementary Data are available at NAR Online:
Supplementary Table S1, Supplementary Figures S1–S7
and Supplementary Materials and Methods.
The authors thank S. Dambacher, M. Kador, H. Klinker,
C. Mehlhorn, M. Holzner and F. Mu ¨ ller-Planitz for
advice and help. The authors also thank U. Rothbauer
for providing various GFP-Trap beads to initially test
GFP pull-down efficiency. The authors are grateful to
D. Rhodes, E. Bernstein, R. Schneider, M. Hendzel, T.
Misteli and S. Matsunaga for reagents. The authors are
indebted to E. Bernstein for critical examination of the
Nucleic Acids Research, 2012,Vol.40, No. 13 5961
manuscript. The authors thank especially the Hake lab
and members of the Adolf-Butenandt Institute for con-
German Research Foundation (DFG), SFB Transregio 5
(to S.B.H., E.K., H.L. and L.S.); the BioImaging Network
Munich and the German Ministry for Education and
Research (BMBF) (to H.L.); the Center for Integrated
Protein Science Munich (CIPSM) (to H.L., J.M., M.M.
and S.B.H.); the European Research Council (to J.M.);
Molecular and CellularLife
(to S.P., S.M.W. and K.S.); the International Doctorate
Program NanoBio Technology (IDK-NBT) (to K.S.).
Funding for open access charge: DFG.
Conflict of interest statement. None declared.
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