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MINIREVIEW
Structure and function of active chromatin
and DNase I hypersensitive sites
Peter N. Cockerill
Experimental Haematology, Leeds Institute of Molecular Medicine, University of Leeds, UK
Introduction
Our current understanding of chromatin structure
really began in the 1970s when it was demonstrated
that chromatin was built up from nucleosomes [1,2]
and it was found that histones could be acetylated [3].
In the late 1970s and early 1980s it was then recog-
nized that chromatin structure was likely to play a sig-
nificant role in gene regulation. It was discovered that
(a) histone acetylation is enriched in active genes [4],
(b) active genes adopt a more accessible chromatin
conformation [5–7] and (c) gene regulatory elements
are associated with nucleosome-free regions that came
to be known as DNase I hypersensitive sites (DHSs)
[7–10]. This remained a relatively obscure field of
research until the mid-1990s when the current intense
interest in chromatin modifications was prompted by

the discovery that transcription factors recruit histone
modifying enzymes [11] and chromatin remodelling
complexes [12,13]. Since then there has been an explo-
sion of papers on the multitude of chromatin modifica-
tions and the factors that can either create or
recognize them. We now have a very detailed picture
of the chromatin modifications normally associated
with transcription units. Hence, we know that promot-
ers, gene bodies, termination regions and even intro-
n ⁄ exon boundaries have very characteristic signatures
of histone modifications, histone replacements and
Keywords
chromatin; DNase I hypersensitive; gene
regulation; nucleosome; transcription
Correspondence
P. N. Cockerill, Experimental Haematology,
Leeds Institute of Molecular Medicine,
University of Leeds, Wellcome Trust
Brenner Building, St James’s University
Hospital, Leeds LS9 7TF, UK
Fax: +44 113 343 8502
Tel: +44 113 343 8639
E-mail: p.n.cockerill@leeds.ac.uk
(Received 18 December 2010, revised 10
February 2011, accepted 5 April 2011)
doi:10.1111/j.1742-4658.2011.08128.x
Chromatin is by its very nature a repressive environment which restricts the
recruitment of transcription factors and acts as a barrier to polymerases.
Therefore the complex process of gene activation must operate at two levels.
In the first instance, localized chromatin decondensation and nucleosome
displacement is required to make DNA accessible. Second, sequence-specific
transcription factors need to recruit chromatin modifiers and remodellers to
create a chromatin environment that permits the passage of polymerases. In
this review I will discuss the chromatin structural changes that occur at
active gene loci and at regulatory elements that exist as DNase I hypersensi-
tive sites.
Abbreviations
BE, boundary element; ChIP, chromatin immunoprecipitation; CTD, C-terminal domain; DHS, DNase I hypersensitive site; DNMT, DNA
methyltransferase; EM, electron microscopy; GM-CSF, granulocyte macrophage colony-stimulating factor; HAT, histone acetyltransferase;
HDAC, histone deacetylase; Hsp70, heat shock protein 70; IL-4, interleukin-4; LCR, locus control region; MAR, matrix attachment region;
MBD, methyl binding domain; MMTV, mouse mammary tumour virus; MNase, micrococcal nuclease; ncRNA, non-coding RNA; NF1,
nuclear factor 1; NFAT, nuclear factor of activated T cells; PARP, poly(ADP-ribose) polymerase; PEV, position effect variegation;
TCR-a, T cell receptor a; TFIIH, transcription factor II H.
2182 FEBS Journal 278 (2011) 2182–2210 ª 2011 The Author Journal compilation ª 2011 FEBS
nucleosome positions [14–16]. However, these advances
have been accompanied by a relative decrease in the
number of studies aimed at gaining an understanding
of the structural conformation of chromatin, and the
changes in chromatin structure that accompany gene
activation. Furthermore, it has now become common-
place for chromatin immunoprecipitation (ChIP)
assays to be used as a surrogate for true structural
studies. However, these studies cannot by themselves
give a detailed understanding of the relationships
between specific chromatin modifications and chroma-
tin architecture. It is also important to recognize that
the principal function of many modifications is to
embed a specific recognizable code within chromatin
[14,17] as opposed to directly altering chromatin con-
formation per se.
To understand the basis of the fundamental mecha-
nisms that lead to gene activation it is necessary to
appreciate that chromatin is by its very nature
repressed by nucleosomes and highly inaccessible. The
normal process of gene activation involves the ordered
recruitment of factors that assemble on DNA in a
highly cooperative manner. The key point of control
in this process is the restriction of accessibility to the
DNA sequence. One obvious consequence of this is
the fact that the genome encompasses many cryptic
binding sites for transcription factors that are not uti-
lized because they do not exist in the correct context.
In this review I will therefore focus primarily on the
actual chromatin structure of active genes, with regard
to nucleosomal organization and higher order struc-
ture, and the chromatin structure changes that occur
during locus activation. I will discuss the nature of
transcription factor interactions with chromatin, which
can lead to localized nucleosome displacement at
DHSs within regulatory elements, as well as long
range changes in the organization and accessibility of
nucleosomes within chromatin. During the course of
these discussions I will draw upon our own experi-
ences using the highly inducible human granulocyte-
macrophage colony-stimulating factor (GM-CSF) gene
as a model system that undergoes extensive remodel-
ling. It is beyond the scope of this review to enter
into an extensive discussion of the role of all the vari-
ous specific histone modifications and the activities of
the different ATP-dependent chromatin remodelling
complexes. There are many other reviews on these
subjects by the experts in these fields [18–27]. I will
discuss in detail, however, the structural implications
of the cycle of histone acetylation and deacetylation
that accompanies cycles of transcription, and highlight
the special significance of histone H4 lysine 16 acety-
lation.
Basic features of chromatin structure
and the influence of transcription
Nucleosomes are the basic building blocks of
chromatin
Chromatin is built up from nucleosomes which com-
prise  146 bp segments of DNA wrapped around a
symmetrical histone octamer core particle containing
two molecules of each of the histones H2A, H2B, H3
and H4 [28–31]. The approximate positions of the hi-
stones within a nucleosome are depicted in Fig. 1,
H3
H3
H4
H4
H2B
H2A
H2B
H2A
Tetramer
Upper
H2A/H2B
dimer
Octamer
+ 146 bp DNA
H2B
H2A
H3
H4
Top half Bottom half
H2B
H2A
H3
H4
Split view
Lower
H2A/H2B
dimer
+
Fig. 1. Composition of nucleosomes. The assembly of the histone
octamer on DNA is represented by this model which depicts the
incorporation of two H3 ⁄ H4 dimers with an inner core of  60 bp
of DNA, followed by the loading of two H2A ⁄ H2B dimers onto the
flanking DNA segments above and below the H3 ⁄ H4 tetramer.
Throughout the nucleosome, each DNA strand of the helix is con-
tacted by histones at  10 bp intervals. The lighter colour shades
depict the bottom half of the nucleosome, and the exploded view
below the octamer depicts the arrangement of the histones con-
tacting 73 bp of DNA within each half. Note that each H4 molecule
actually bridges two turns of the DNA helix, by contacting the inner
core DNA within one half of the nucleosome plus the DNA at the
exit point of the opposite half.
P. N. Cockerill Active chromatin and DNase I hypersensitive sites
FEBS Journal 278 (2011) 2182–2210 ª 2011 The Author Journal compilation ª 2011 FEBS 2183
which is a greatly simplified version of the X-ray crys-
tal structure obtained at 2.8 A
˚
resolution [32] but is
used here to convey the concept that an H3 ⁄ H4 tetra-
mer making up the inner core is first loaded onto the
central 60 bp of DNA, followed by two H2A ⁄ H2B
dimers which are loaded above and below the H3 ⁄ H4
tetramer onto the flanking DNA segments. Most of
the genome exists in the form of regularly spaced
nucleosomes with a DNA repeat length of  180–
200 bp. Most nucleosomes also recruit either histone
H1 or high mobility group (HMG) proteins (some-
times both) which bind to the outside of the nucleo-
some to form a particle known as the chromatosome,
which occupies  166 bp of DNA [29,33–35]. Within
native chromatin, nucleosomes assemble into higher
order structures, and both the core histone tails and
linker histone H1 (or H5) play major roles in main-
taining higher order chromatin condensation [36–38].
However, even in the absence of histone H1, chains of
nucleosomes spontaneously assemble into a higher
order fibre 30 nm in diameter if physiological levels of
monovalent or divalent cations are present. It requires
just 0.5 mm MgCl
2
,or60mm NaCl, to promote coil-
ing of 10-nm diameter fibres into 30-nm diameter
fibres [37,39]. The 30-nm fibre represents the predomi-
nant type of chromatin structure observed in electron
microscopy (EM) studies of either ruptured interphase
nuclei [40] or metaphase chromosomes that have been
partially dissociated in 1 mm MgCl
2
[39]. The exact
nature of the structure of this fibre is still a subject of
intense debate [41], but it can potentially be repre-
sented either by a double helix with crossed linkers,
where the linkers zigzag across the centre of the fibre
[42,43], or alternatively as a simple solenoid made up
of six nucleosomes per coil [44], where the nucleosomes
interdigitate between adjacent coils [45].
Chromatin fibres are naturally highly condensed
in vivo
Under salt-free conditions, and in the absence of his-
tone H1, chains of nucleosomes can be visualized as
unfolded chains of regularly spaced 10-nm diameter
particles, giving rise to the popular ‘beads on a
string’ images. Unfortunately, this textbook image
has led to the popular misconception that active
gene loci decondense completely into these unfolded
10-nm diameter fibres. In reality, the eukaryotic gen-
ome is assembled in a much more condensed state
under physiological conditions, and exists in confor-
mations at least as complex as 30-nm diameter
fibres, within all but the most actively transcribed
genes [46,47].
Micrographic studies of interphase and prophase
nuclei reveal that most of the genome is actually
assembled at degrees of condensation much higher
than even the 30-nm fibre [47–49]. By EM, chromatin
fibres are typically seen to be 110–170 nm in diameter
during interphase [48] and 200–250 nm in diameter
during prophase [49]. These high levels of chromatin
condensation were also observed within active genes
via a different approach whereby megabase segments
of chromatin were fluorescently labelled inside living
cells [47]. By this means it is possible to visualize genes
aligned in a linear array both before and after induc-
tion of transcription. However, after transcription acti-
vation, the level of compaction detected was still 10- to
30-fold higher than the level of the 30-nm fibre [47].
Similar results were obtained using fluorescence
microscopy of arrays of steroid-inducible mouse mam-
mary tumour virus (MMTV) DNA, where a DNA
compaction ratio of 50- to 1300-fold remained after
induction of transcription [50]. Hence, transcribed
genes can in some cases remain compacted to an
extent far greater than the DNA packing ratio of
30–40 predicted for a 30-nm fibre and 5–10 predicted
for a 10-nm fibre. The exceptions to this are the highly
transcribed genes such as the ribosomal RNA genes
which are so heavily loaded with polymerases that
most of the nucleosomes are evicted and no conven-
tional chromatin fibre remains.
The concept of the 30-nm fibre as the universal
building block of chromatin in vivo has also been chal-
lenged by an independent cryo-EM analysis of meta-
phase chromosomes which depicted homogeneous
grainy images of chromatin sections with no evidence
for any discrete higher order fibre formation [51]. The
interpretation of these images was that chains of nucle-
osomes within chromosomes exist primarily in a disor-
dered interdigitated state, rather than conforming
to the well organized helical structures observed for
in vitro reconstituted chromatin fibres.
The Balbiani rings observed in polytene chromo-
somes in Chironomus tentans provide another represen-
tation of very actively transcribed genes. These are
looped out domains of highly decondensed chromatin
containing genes heavily loaded with polymerases. The
elegant EM studies of Balbiani rings by Daneholt and
co-workers [52,53] gave us one of our first glimpses of
the true nature of transcribed chromatin. In this model
system, sequences immediately upstream and down-
stream of genes can be seen in most cases to remain
coiled as 30-nm fibres. In the cases where the RNA
polymerases are the most densely packed, the interven-
ing DNA can be seen typically as either nucleosome-
free or as a 10-nm fibre. However, even in these highly
Active chromatin and DNase I hypersensitive sites P. N. Cockerill
2184 FEBS Journal 278 (2011) 2182–2210 ª 2011 The Author Journal compilation ª 2011 FEBS
transcribed structures there sometimes remain stretches
of condensed 30-nm fibres formed in between more
distantly spaced polymerases [52,53]. This suggests that
chromatin can transiently exist as a decondensed
10-nm fibre during transcription, perhaps even nucleo-
some-free, but that coding regions return to a conven-
tional 30-nm diameter chromatin fibre once a
polymerase has passed.
The challenge presented here, then, is to gain a bet-
ter understanding of the significance of the different
degrees of chromatin condensation and chromatin
modification that prevail in the nucleus, that enable
the appropriate activation of specific gene loci. Clearly,
it is not sufficient to merely think in terms of con-
densed 30-nm chromatin fibres versus open 10-nm
chromatin fibres. We also need to be able to define the
specific mechanisms that create a more dynamic chro-
matin structure in which nucleosomes and chromatin
proteins are more mobile [15,54]. For example, it is
accepted that active gene loci are less condensed and
more accessible than inactive loci, and that a passing
polymerase must at least transiently create openings in
the chromatin fibre. However, in normal interphase
nuclei, it is likely that most sections of most active
genes will remain condensed to at least the level of 30-
nm fibres. The exceptions to this rule will be the actual
sites of ongoing transcription where individual polyme-
rases are bound and any genes which are so loaded
with polymerases that this does not permit the reas-
sembly of nucleosomes.
Active chromatin domains
Evidence from a wide range of sources confirms that
active gene loci are associated with fundamental
changes in chromatin architecture across broad
domains spanning genes. Electron micrographs of
interphase nuclei reveal areas of condensed heterochro-
matin and decondensed euchromatin that are generally
assumed to represent inactive and active chromatin –
although this is now known to be somewhat of an
over-simplification, as some active genes reside within
heterochromatin. Drosophila polytene chromosomes
offer one of the clearest examples of active chromatin
domains whereby active genes appear as highly decon-
densed ‘puffs’.
Active chromatin domains are permissive for
transcription
It is generally accepted that active genes lie within
broad active chromatin domains that carry a variety of
modifications associated with active chromatin [18–23].
The significance of this was highlighted by a study that
found that chromatin domains marked by H3 acetyla-
tion and H3-K4 methylation were permissive for the
stable expression of integrated transgenes, whereas
transgenes integrated at other sites were prone to
silencing [55].
Active genes reside within extensive
nuclease-sensitive domains
It was recognized in the 1970s and 1980s that chroma-
tin domains encompassing active genes are at least
twice as sensitive to DNase I digestion as non-tran-
scribed genes [5–7,56–62]. These studies used either C
o
t
analysis of DNA hybridization kinetics, slot-blot filter
hybridization, or the disappearance of discrete restric-
tion enzyme DNA fragments as a measure of the rate
of DNase I digestion. In many cases it was found that
these accessible domains exhibiting general DNase I
sensitivity extended many kilobases upstream and
downstream of the transcription units they encom-
passed. For example, the chicken lysozyme active
domain extends for about 14 kb upstream and 6 kb
downstream of the gene, and is preferentially sensitive
in the oviduct which expresses lysozyme, but not in
liver or erythrocytes which do not [59]. In the chicken
b-globin locus the DNase I sensitive domain extends
from 6 kb upstream to 8 kb downstream of the gene,
although in this instance the coding sequences are even
more sensitive than the immediate flanking sequences
[7]. In the mouse b-globin locus, the active adult b-glo-
bin genes are in a more nuclease-sensitive domain than
the inactive embryonic globin gene [58]. However,
increased nuclease accessibility does not mean that the
chromatin fibre is completely decondensed. Recent
studies suggest that active genes remain, for the most
part, in a condensed state, with the linker regions pro-
tected within the fibre and no more accessible to
DNase I than the nucleosomes [63]. This study also
suggested that some of the reports of general nuclease
sensitivity might in fact be attributable to the hyper-
sensitivity at the DHSs within these active chromatin
domains.
It was once thought that one DNase I sensitive
domain would correspond to one gene plus its regula-
tory elements. However, this concept is now outdated,
because regulatory elements can reside far from the
genes they control, sometimes existing within inactive
loci. In the case of the lysozyme locus, which was ini-
tially used to help establish the active domain model,
it was later found that its domain encompasses the
ubiquitously expressed Gas41 gene, even though this
domain was thought to be sensitive in lysozyme-
P. N. Cockerill Active chromatin and DNase I hypersensitive sites
FEBS Journal 278 (2011) 2182–2210 ª 2011 The Author Journal compilation ª 2011 FEBS 2185
expressing cells only [64]. In chicken embryo erythro-
cytes, the inactive lysozyme gene has almost the same
DNase I sensitivity as the active Gas41 gene [63]. This
issue was also addressed by a genome-wide analysis
which found that open chromatin domains more clo-
sely correlated with gene density than gene activity,
because inactive genes can also be found within active
gene domains [65].
Demarcation of active chromatin domains
It was once proposed that active chromatin domains
would be demarcated by the rigid attachment of
nuclear matrix attachment regions (MARs or SARs)
to the nuclear skeleton [66–68]. However, there is little
evidence for this [69], and MARs are often found
inside active domains or associated with enhancers
[66,70]. In contrast, there are numerous examples
where the borders of active domains are defined by a
class of DNA elements termed boundary elements
(BEs) (or barrier elements) which block the spread of
repressive chromatin [71,72]. In this regard, BEs may
function in a way that is not quite the same as another
class of elements termed insulators that block enhan-
cer–promoter communication but do not necessarily
demarcate active chromatin domains. The terminology
here can be very confusing, however, because the two
terms are often used interchangeably, and some DNA
elements have both BE and insulator activity [71,72].
BEs were first identified in Drosophila, where they were
found to block position effect variegation (PEV) of
expression of mobile integrated transgenes containing
transposons. One of the best studied such examples
exists in the Drosophila 87A7 heat shock protein 70
(Hsp70) locus where two BEs termed SCS and SCS’
directly flank an inducible active chromatin domain
spanning 12 kb. These BEs function both as enhancer-
blocking insulators [69,73] and as active chromatin
domain boundaries [74,75] that block PEV [76]. The
SCS and SCS’ elements are the prototypes of one of
the major classes of BE in Drosophila, which bind a
protein complex termed BEAF [77]. This complex is
associated with about half of the interbands in poly-
tene chromosomes, and in many cases is present at the
borders of active genes within polytene chromosome
puffs [78].
One of the proposed mechanisms of BE function
involves the recruitment of chromatin modifying com-
plexes that create islands of active chromatin which
counteract the repressive complexes that mediate het-
erochromatin spreading [71,72]. Many BEs are known
to have promoter activity and to recruit chromatin
activators, and in yeast some BEs are in fact tRNA
genes [71,72]. This model of BE function is further
supported by the fact that many components of repres-
sive chromatin complexes, such as the histone H3-K9
methyltransferase SUV39H1 [Su(var)3-9 in Drosophila],
were themselves initially identified via mutations that
blocked PEV [79,80]. These proteins are typically
involved in heterochromatin spreading mediated by
HP1 [71,80]. Conversely, enhancer-blocking insulators
can function by an alternative mechanism. Vertebrate
insulators invariably recruit CTCF which in turn
recruits the cohesin chromosomal cohesion complex
[71,81]. This leads to a model whereby CTCF controls
chromatin looping [82] and defines independent func-
tional DNA domains within which enhancers and pro-
moters can cooperate, as opposed to demarcating
active chromatin domains.
Active loci undergo extensive nucleosome
mobilization
Classical models of chromatin depict chains of regu-
larly spaced nucleosomes that fold up into a helix as
highly ordered chromatin fibres. However, this image
is really only representative of inactive loci that consti-
tute the bulk of chromatin in the nucleus. The highly
regular ordering of nucleosomes is more closely associ-
ated with gene silencing, and with decreased sensitivity
to DNase I [83].
Although it is well known that gene activation
induces alterations in chromatin, there are still rela-
tively few studies which have assessed the organization
as opposed to the modification status of active chro-
matin. Significantly, those studies which have
addressed this issue have typically found that gene
activation is associated with extensive nucleosome
mobilization which results in the formation of a highly
disorganized nucleosome array incapable of conform-
ing to any of the current models of the higher order
chromatin fibre. It is even possible that this highly dis-
organized form of chromatin includes some nucleo-
somes fused together, as there is evidence that adjacent
nucleosomes can in some cases merge to form a single
fused particle [84]. This type of information is difficult
to gather from genome-wide studies that have defined
the average nucleosome positions, because this
approach does not necessarily provide a meaningful
picture of how individual nucleosomes are packaged
within chromatin relative to each other in any one cell.
Nucleosome mobilization is best visualized by elec-
trophoretic size fractionation and southern blot
hybridization of chromatin digested with micrococcal
nuclease (MNase), which cuts primarily in linker
regions. This type of analysis typically reveals ladders
Active chromatin and DNase I hypersensitive sites P. N. Cockerill
2186 FEBS Journal 278 (2011) 2182–2210 ª 2011 The Author Journal compilation ª 2011 FEBS
of regularly spaced discrete oligo-nucleosome bands
for bulk chromatin, but a smeared pattern for active
chromatin. An example of the phenomenon is pre-
sented in Fig. 2A, which shows MNase digestion data
for the human GM-CSF locus in T cells [85,86]. In
unstimulated T cells, where the gene is completely
silent, MNase generates very uniform ladders of evenly
spaced nucleosomes with an average repeat length of
about 190 bp throughout the GM-CSF locus [85,86].
Parallel mapping of nucleosome positions by indirect
end-labelling [85] shows that nucleosomes are posi-
tioned at  200 bp intervals at highly specific locations
throughout at least 6 kb of the locus (Fig. 3A). How-
ever, after gene activation by stimulation of calcium
and kinase signalling pathways, nucleosomes through-
out this 6 kb region adopt a highly disorganized struc-
ture with nucleosomes redistributed to random
positions in both T cells and mast cells. Interestingly,
the degree of nucleosome position randomization is far
more extreme within the first few kilobases of the non-
transcribed upstream region than within the gene itself
(Fig. 2A). This could mean that each cycle of tran-
scription resets the normal spacing of nucleosomes.
Furthermore, for genes undergoing moderate levels of
transcription, it is thought that RNA polymerase II
(Pol II) proceeds via a mechanism that actually pre-
vents nucleosome translocation [87]. However, the situ-
ation may be very different at highly transcribed
genes, where closely spaced Pol II molecules can dis-
place the entire histone octamer [88].
As will be discussed in more detail below, there is
widespread evidence for both nucleosome repositioning
and increased chromatin accessibility in the neighbour-
hood of regulatory elements. For example, in mast
cells, GATA factors are able to bind to an accessible
nucleosome-free linker region within the GM-CSF
enhancer, leading to lineage-specific repositioning of
the flanking nucleosomes (Fig. 3B). This involves the
relocation of the upstream nucleosome N0 to a new
position  100 bp further upstream and the down-
stream nucleosomes about 20–30 bp further down-
stream. A similar finding was obtained in studies of
the MMTV long terminal repeat where Oct1 and
nuclear factor 1 (NF1) were sufficient to direct nucleo-
some repositioning [89]. A further consequence of
GATA factor recruitment at the GM-CSF enhancer is
increased accessibility of the linker regions flanking the
two nucleosomes located immediately downstream of
the GATA sites (Fig. 3A) [85]. This appears to repre-
sent a primed active state that precedes the disruption
of these same two nucleosomes upon subsequent
inducible binding of nuclear factor of activated T cells
(NFAT) and AP-1 (to be discussed in more detail
below). A similar situation may exist in the human
interleukin-4 (IL-4) locus, where a total of six nucleo-
some linker regions at the 5¢ end of the gene are
more accessible specifically in type 2 T helper cells that
express IL-4 [90].
Nucleosome mobilization in the 3 kb region between
the GM-CSF enhancer and promoter is dependent
upon this upstream enhancer [85]. In the absence of
the enhancer, inducible nucleosome mobilization in the
upstream region is completely abolished (Fig. 2B).
These findings suggest that one important aspect of
enhancer function is to direct localized nucleosome
mobilization within an active chromatin domain. This
implies that enhancers can function both by recruiting
GM-CSFEnhancer
–2 to –0.6 kb
+1.2 to 2.6 kb
Gene probe
5′ probe
A
B
Non-stimulated
Stimulated
GM-CSF locus
with the
enhancer
deleted
–3 kb
676
418
175
130
bp
Non- Stim.
stim.
MNase
5′ probe
5′ probe
Non- Stim.
stim.
MNase
1847
bp
Gene probe
–185 bp
+ 150 bp
– 180 bp
+ 160 bp
Non-stimulated
Stimulated
RE
RE
Fig. 2. Nucleosome mobilization within the activated GM-CSF
locus. Southern blot analysis of oligo-nucleosome fragments pro-
duced by increasing amounts of MNase digestion and probed
directly with specific GM-CSF locus probes. In this analysis chroma-
tin fragments were prepared from T cells before or after stimulation
of TCR signalling pathways that induce NFAT and AP-1 [85,86].
Nucleosome mobilization is characterized by a smear of random
products at early digestion points, and by the small proportion of
very close packed nucleosomes that are more resistant to MNase
and remain after increased digestion. In this analysis, nucleosomes
have an average repeat length of  190 bp before mobilization,
whereas the closed packed nucleosomes have a repeat length of
 150 bp after mobilization. The densitometric traces of the middle
lanes are shown below each panel and reveal that the predominant
pattern is essentially random after mobilization. (A) Analysis of the
intact GM-CSF locus. (B) Analysis of the GM-CSF locus with a spe-
cific deletion of the 0.7 kb enhancer.
P. N. Cockerill Active chromatin and DNase I hypersensitive sites
FEBS Journal 278 (2011) 2182–2210 ª 2011 The Author Journal compilation ª 2011 FEBS 2187
remodellers that can act within a few kilobases and by
looping to function over larger distances. GM-CSF
enhancer activation is mediated by the inducible tran-
scription factors NFAT and AP-1 which direct the for-
mation of a DHS (Fig. 4, discussed in more detail
below). NFAT ⁄ AP-1 complexes are thought to recruit
CBP ⁄ P300 family histone acetyltransferases (HATs) as
well as SWI ⁄ SNF family chromatin remodelling com-
plexes which may well account for the observed nucle-
osome mobilization [91].
Within the region of nucleosome mobilization
upstream of the GM-CSF gene, it can also be seen
that a fraction of the nucleosomes end up as fragments
of close packed nucleosomes with a repeat length of
just 150 bp which resist digestion (Fig. 2A). It is
inconceivable that such a close packed arrangement
could either accommodate histone H1 or assemble into
a 30-nm chromatin fibre. I have attempted to depict
this chromatin structure transition in Fig. 4A, whereby
a well organized inactive chromatin fibre compacted
by histone H1 is converted to a disorganized active
chromatin fibre that is probably depleted of histone
H1. Because it is so disorganized, active chromatin
may have an intrinsic resistance to folding into a rigid
compacted structure.
Similar nucleosome mobilization within active loci
has been observed in many model systems (which I
have summarized previously [85]) and is not just
restricted to transcribed regions. For example, in the
chicken oviduct, a 2.5 kb region of chromatin just
upstream of the ovalbumin gene undergoes extensive
nucleosome randomization, whereby some chromatin
fragments contract to a nucleosome repeat length of
about 150 bp [92]. This is also observed in mouse B
Nucleosome positions and functional binding sites in the human GM-CSF enhancer
N0 N1 N2 N3
N1 N2 N3
Mast cells
T cells
NFAT
AP-1
Runx1
NFAT
AP
-1
Sp1
GATA
GATA
AP
-1
GATA
-2
GATA-2
1
Bgl II
717
Bgl II
1 800 bp
200
600
400–200
–3289 –2578
Enhancer
GM-CSF
Mast
cells
stim.
T cells
stim.
Mast
cells
non-stim.
T cells
non-stim.
GATA + AP-1
NFAT + AP-1
NFAT + AP-1
GATA
Promoter
Inducible nucleosome reorganisation across the human GM-CSF locus
Enhancer
Strongly positioned
Nucleosome
Runx1
A
B
Fig. 3. Positions of regulatory elements and
nucleosomes within the GM-CSF enhancer.
(A) Relative MNase cleavage at linker
regions that define nucleosome positions in
T cells (blue) and mast cells (red) before and
after stimulation with 4b-phorbol 12-myri-
state 13-acetate and calcium ionophore. The
graphs of MNase cleavage represent the
ratio of the level of MNase digestion in
chromatin divided by the level of cleavage
for purified genomic DNA [85]. The scale
represents position relative to the transcrip-
tion start site. (B) A map showing the posi-
tions of regulatory elements required for
function in either T cells or mast cells.
Shown below are the positions that nucleo-
somes and GATA-2 occupy in unstimulated
T cells and mast cells [85].
Active chromatin and DNase I hypersensitive sites P. N. Cockerill
2188 FEBS Journal 278 (2011) 2182–2210 ª 2011 The Author Journal compilation ª 2011 FEBS
cells expressing Igj within the coding sequences of the
Igj gene, where nucleosome mobilization extends to
just beyond the end of the transcription unit [93,94].
The role of histone H1 in chromatin accessibility
The molecular basis of the general DNase I sensitivity
observed both within and around genes is likely to be
highly complex. At the simplest level, loss of histone
H1 is sufficient to reduce the level of compaction of
the chromatin fibre, and at active genes the amount of
histone H1 is reduced compared with inactive genes
[95–97]. Conversely, addition of histone H1 to active
chromatin results in gene repression [98]. Although sig-
nificant levels of histone H1 do remain at active loci,
the ratio of histone H1:nucleosomes is less than the
1 : 1 predicted for inactive loci, and this may be suffi-
cient to trigger a breakdown of chromatin compaction
[95]. Furthermore, chromatin within nuclei stripped of
histone H1 is about two- to three-fold more sensitive
to DNase I [99], consistent with the increased level of
DNase I sensitivity typically observed at active gene
loci. Histone H1 is also implicated as a factor that
maintains the differential DNase I sensitivity of the
mouse adult and embryonic b-globin genes [58]. How-
ever, it is probably safe to assume that general DNase
I sensitivity arises from the concerted effects of many
of the chromatin modifications associated with active
genes, plus the act of transcription itself. For example,
a recent study found that both acetylation of H4-K16
and eviction of histone H1 were required for the
decompaction of the 30-nm fibre in vitro [100].
Genetic analyses have found that histone H1 is not
as essential for correct gene regulation as previously
thought [97]. Histone H1 can be eliminated from uni-
cellular organisms without much impact, and reduction
of histone H1 levels in mouse stem cells to 50% of
normal levels results in a global reduction in average
nucleosome linker length but not much effect on gene
expression [97,101]. Although this reduction in H1
A
Anatomy of the inducible DNaseI hypersensitive site in the GM-CSF enhancer in T cells
DNase I and MNase
Active chromatin with DHS
and mobilised nucleosomes
with less histone H1
Promoter
Condensed chromatin + histone H1
NFAT +
AP-1
Nucleosome N1
Nucleosome N2
400 bp
140 540
GATA + AP-1
Mast
T cells
RE
NFAT + AP-1
NFAT + AP-1
B
-
1 Bgl II
-
717 Bgl II
-
265 Apa I
-
514 Pst I
NFAT
AP-1
Sp1
Runx
SWI/SNF
NFAT
AP
-1
CBP
SWI/SNF
CBP
Runx
Fig. 4. DHS formation and nucleosome
mobilization at the human GM-CSF locus.
(A) Model of the DHS within the human
GM-CSF enhancer induced by activation of
TCR signalling pathways that induce NFAT
and AP-1 [85,86]. Prior to activation, the
locus exists as an array of regularly spaced
nucleosomes assembled as condensed
chromatin. The induction of the DHS is
accompanied by the eviction of two posi-
tioned nucleosomes that otherwise occupy
two discrete sets of factor binding sites and
block binding of the constitutively expressed
factors Sp1 and Runx1. Upon activation,
NFAT and AP-1 bind cooperatively to com-
posite NFAT ⁄ AP-1 elements within each
nucleosome, and are predicted to support
the formation of enhanceosome-like com-
plexes including co-factors such as CBP and
SWI ⁄ SNF [85]. In vivo footprinting con-
firmed inducible binding of NFAT, AP-1, Sp1
and Runx1 [86,233]. Nuclease digestion
studies have determined that the nucleo-
somes normally occupy  150 bp of DNA
before stimulation, and are replaced by
complexes that protect  50 bp of DNA.
(B) High resolution DHS mapping of the
GM-CSF enhancer in activated T cells and
mast cells by indirect end-labelling [85]. The
protected regions between zones of DNase
I hypersensitivity (arrowed) indicate the
potential presence of enhanceosomes.
P. N. Cockerill Active chromatin and DNase I hypersensitive sites
FEBS Journal 278 (2011) 2182–2210 ª 2011 The Author Journal compilation ª 2011 FEBS 2189
levels is tolerated by stem cells in vitro, the defect is
embryonic-lethal in mice and reveals a need for H1 in
embryonic development.
Histone replacement at active gene loci
It is established that the act of transcription involves
at least partial transient displacement of histones from
nucleosomes, as well as the substitution of some of the
canonical histones with histone variants. It has been
known since 1983 that active genes are enriched in nu-
cleosomes lacking one molecule of each of histones
H2A and H2B, and that these partially disassembled
nucleosomes are preferentially bound by Pol II in vitro
[102]. As depicted in Fig. 5, RNA polymerase can
recruit facilitator of active transcription (FACT) which
displaces one H2A ⁄ H2B dimer as each nucleosome is
transcribed [103]. Once the polymerase has passed, the
H2A ⁄ H2B dimer is replaced. There is also evidence for
more substantial histone core displacement during
transcription because histone H3.3 is highly enriched
within transcribed or recently transcribed genes [104–
106]. H3.3 is synthesized during interphase whereas
H3.1 and H3.2 are synthesized during S phase. This
may be one reason why H3.3 is found enriched at
active genes. It was once assumed that the presence of
H3.3 in active genes was of little structural signifi-
cance, because H3 variants are structurally very similar
to each other. However, it is now believed that H3.3-
containing nucleosomes are much less stable than
H3.1-containing nucleosomes [107]. Furthermore, H3.3
may suppress histone H1 mediated chromatin compac-
tion, because H3.3-containing nucleosomes appear to
be unable to recruit histone H1 [108].
Regulation of chromatin structure by
poly(ADP-ribose) polymerase (PARP)
Studies in Drosophila and mammals have revealed that
PARP-1, the enzyme that directs modification of
histones by poly ADP ribosylation, can direct either
gene activation or repression [75,109,110]. These
opposing actions appear to work by distinct mecha-
nisms. At repressed loci, PARP-1 can function as a
structural protein whereby it binds to nucleosomes at a
1 : 1 molar ratio in place of histone H1 and, like H1,
it promotes chromatin condensation [110]. In this con-
text, PARP-1 does not PARylate chromatin, and acti-
vation of its enzymatic activity actually relieves
silencing [110]. PARP-1 binds to chromatin by engag-
ing each of the two strands of DNA at the point at
which they exit from the nucleosome, thereby opposing
the actions of transcriptional activators that mobilize
or disassemble nucleosomes [110].
If the enzymatic functions of PARP-1 are activated
in the presence of NAD+ it mediates the PARylation
of both histones and PARP-1 itself, and thereby pro-
motes decondensation of higher order chromatin struc-
ture [75,109]. However, in studies of condensed
chromatin assembled in vitro in the presence of PARP-
1, it was found that chromatin decondensation can be
induced by activation of PARP-1 without PARylation
of the underlying core histones and without disruption
of nucleosomes [110]. In this model system, chromatin
decondensation occurred primarily via auto-PARyla-
tion and loss of binding of PARylated PARP-1 to
chromatin.
PARP-1 was also found to contribute to extensive
remodelling of nucleosomes across the Drosophila
Hsp70 in response to heat shock [75]. This study made
the surprising observation that nucleosomes through-
out the Hsp70A locus were rendered MNase sensitive
after just 1 or 2 min of heat shock. This extensive dis-
ruption or modification of nucleosomes spanned the
entire region defined by the SCS and SCS’ boundary
elements, was independent of transcription, and was
suppressed by RNAi-depletion of PARP-1 [75].
Active genes partition differentially during
chromatin fractionation
Active chromatin has very different physical properties
from inactive chromatin. For example, minichromo-
somes assembled in Xenopus oocytes partition into
inactive soluble chromatin and insoluble active chro-
matin [111]. Early attempts to fractionate native chro-
matin into functionally distinct fractions were
performed by digestion of nuclei with MNase followed
Ac
Ac
HDACs
Histone
hexamer
H2A/H2B
dimer
Pol II
Direction of transcription
HATs
FACT
H3K36
Me2
Ac
RNA
Fig. 5. Model of the chromatin structure in the vicinity of an elon-
gating Pol II complex. Histone acetylation in advance of polymeras-
es is likely to create an open chromatin structure. The advancing
polymerase recruits FACT which partially disassembles the nucleo-
some, allowing Pol II to pass this barrier. Once Pol II has passed,
HDACs such as Rpd3S can be recruited via dimethylated or trime-
thylated H3-K36 and act to return chromatin to the condensed
state.
Active chromatin and DNase I hypersensitive sites P. N. Cockerill
2190 FEBS Journal 278 (2011) 2182–2210 ª 2011 The Author Journal compilation ª 2011 FEBS
by separation based on solubility under different ionic
conditions [93,112]. This involved progressively
fractionating chromatin into (i) highly soluble small
chromatin fragments readily released from nuclei dur-
ing digestion (fraction S1), (ii) the bulk of the remain-
ing chromatin which could be subsequently solubilized
from digested nuclei by extraction in 2 mm EDTA
(fraction S2), and (iii) the residue comprising highly
insoluble chromatin (fraction P). These studies found
that fraction S1 was predominantly mono-nucleosomal
and was highly enriched in transcribed genes, but was
depleted of inactive genes and histone H1; fraction S2
contained classically organized oligo-nucleosomes
depleted of transcribed genes but retaining most of the
nuclear histone H1; fraction P was composed of disor-
ganized chromatin fragments that were also enriched
in active genes [93]. Hence, the S1 fraction represented
the highly accessible and extensively modified active
gene fraction containing highly acetylated nucleo-
somes, which were more soluble and could be released
from highly remodelled chromatin segments that were
tightly associated with the transcription apparatus
[113].
At first it appears paradoxical that the more accessi-
ble active genes should be split between the most and
the least soluble chromatin fractions. However, the
explanation for this observation lies in the fact that
active genes are tightly associated with multi-compo-
nent transcription factor and polymerase complexes at
sites that have been termed transcription factories
[114–116]. The residual insoluble fraction is in essence
equivalent to the ‘nuclear matrix’ fraction that was
shown to be enriched in active genes [117–119]. While
the ‘nuclear matrix’ was originally proposed to be a
true nuclear skeleton organizing the functions of the
nucleus, it may in reality represent an aggregate of all
the active sites in the nucleus, such as transcription
factories, that remain when the inactive chromatin
fraction is removed. These may be the sites bound by
MARs and may explain why MARs often exist along-
side enhancers.
Chromatin structure regulation by
histone acetylation
The role of histone acetylation
Histone modifications help to create a more accessible
and dynamic chromatin environment and thereby play
a major role in making chromatin permissive for tran-
scription [54]. Acetylation of lysines leads to neutraliza-
tion of the positively charged nitrogen atoms that
mediate contacts between histone tails and DNA, ren-
dering individual nucleosomes more unstable and
mobile. These histone tail contacts occur primarily with
the linker DNA rather than the nucleosomal DNA [21].
In contrast, other non-neutralizing modifications such
as methylation may have a less direct impact on struc-
ture, but serve as docking sites for regulatory molecules
such as chromatin remodelling factors.
Acetylation of histone H4-K16 suppresses
chromatin condensation within active genes
In a study of chromatin fibre dynamics, it was revealed
that acetylation of lysine 16 on histone H4 (H4-K16)
was the only modification that was able to destabilize
higher order chromatin structure [120]. In sedimenta-
tion velocity analyses, acetylation of this one amino
acid led to a degree of chromatin fibre decompaction
equivalent to loss of the entire histone H4 tail [120].
The reason for this may be because H4-K16 mediates
interactions with adjacent stacks of nucleosomes within
the 30-nm fibre and its acetylation disrupts H4 tail
secondary structure and salt bridging [121,122]. Subse-
quent EM studies confirmed that acetylation of H4-
K16 led to a breakdown of 30-nm compacted fibres
[100]. A more recent chromatin sedimentation study
also found that H4-K16 acetylation is sufficient to
greatly reduce chromatin folding, whereas combined
acetylation of H4-K5, K8 and K12 had a much more
modest effect [123].
Acetylation of H4-K16 does appear to have special
significance in vivo [21]. Unlike AcH3-K9, which is
mainly confined to promoters, AcH4-K16 is also pres-
ent at elevated levels throughout the transcribed
regions of active genes in human T cells [124]. In a
study in yeast, mutations were introduced alone or in
combination in lysines 5, 8, 12 and 16 in the gene for
histone H4 [125]. Of these, the only mutation that had
a specific effect on patterns of yeast gene expression
was the mutation in H4-K16. In Drosophila, specific
acetylation of H4-K16 is an integral feature of dosage
compensation that results in a global two-fold increase
in gene activity [126]. Interestingly, AcH4-K16 plays
an additional role in countering the repressive effects
of chromatin because it reduces the ability of the ISWI
remodelling complex to reset active chromatin as com-
pacted chromatin [127].
The HAT primarily responsible for the bulk of
AcH4-K16 in vivo is likely to be MOF in mammals
and Drosophila, and its homologue Sas2 in yeast.
MOF is H4-K16 specific and was originally identified
in Drosophila as a component of the dosage compensa-
tion complex [126] in association with MSL1, MSL2
and MSL3 [128,129]. MSL3 specifically binds to
P. N. Cockerill Active chromatin and DNase I hypersensitive sites
FEBS Journal 278 (2011) 2182–2210 ª 2011 The Author Journal compilation ª 2011 FEBS 2191
methylated H3-K36, which promotes recruitment of
MOF to recently transcribed regions, especially at the
3¢ ends of genes where H3-K36me3 is enriched [128].
In mammalian cells MOF also exists as part of a sepa-
rate MOF–MSLv1 complex that co-purifies with
MLL1 and WDR5 which binds to dimethylated and
trimethylated H3-K4, and this is found preferentially
bound at active promoters [128,129]. In genome-wide
analyses in human cells, the enrichment of AcH4-K16
within transcribed genes is closely correlated with
binding of both MOF and Tip60 (equivalent to Esa1
in yeast NuA4) [124,130]. However, in human cells
depletion of MOF, but not Tip60, results in reduced
global levels of AcH4-K16 and defective DNA damage
response [131]. In mouse embryos, MOF is essential
for AcH4-K16, and loss of MOF results in embryonic-
lethal chromatin condensation [132–134]. In yeast,
acetylation of H4-K16 is also required to suppress the
spread of heterochromatin, and Sas2 mutations lead to
spreading of heterochromatin mediated by repressive
Sir protein complexes [135]. Conversely, the hetero-
chromatin protein Sir2 directs deacetylation of
H4-K16 and promotes heterochromatin spreading by
allowing Sir3 to bind to non-acetylated H4-K16 [21].
Although not required for acetylation of H4-K16,
the NuA4 group of HATs are essential for H4-K5, K8
and K12 acetylation [136–138]. In yeast, this group is
made up of NuA4 and Piccolo NuA4 which both uti-
lize Esa1 as the HAT, and Esa1 was found to be essen-
tial for H4-K5, K8 and K12 acetylation [137]. In
mammalian cells, this group is composed of two dis-
tinct complexes which employ different HATs: Tip60
which forms a NuA4-like complex, and HBO1 which
more closely resembles yeast Piccolo NuA4 [138]. In
mammals, it is HBO1 and not Tip60 which is responsi-
ble for the bulk of the global H4-K5, K8 and K12
acetylation [136]. Each of these HATs exists in com-
plexes that include PHD domains that interact with
methylated histone H3-K4 and ⁄ or K36 [138–140].
Transcription directs transient histone
acetylation
The regulated process of transcription is accompanied
by an ordered sequence of transient histone modifica-
tions that directly impact upon chromatin structure
across transcribed genes. There is also evidence that
transcription initiation is a cyclical process [141,142],
involving alternate assembly and disassembly of an
open chromatin structure at promoters [143–145], as is
described in more detail in another review paper in this
issue [146]. This cyclical process is accompanied by
transient sequential histone acetylation and deacetyla-
tion, and transient recruitment of remodellers and
transcription factors.
A cycle of transcription commences with the recruit-
ment of transcription factors and co-factors bound at
the promoter, which modify the local chromatin struc-
ture and enable the assembly of the pre-initiation com-
plex. In yeast, transcription factors typically recruit
HATs such as SAGA and NuA3, which mainly acety-
late histone H3, and NuA4 which acetylates histone H4
on K5, K8 and K12. This cascade of events leads to
recruitment of transcription factor II H (TFIIH) which
phosphorylates Pol II at the serines at position 5 (Ser5)
within the heptapeptide repeats of the C-terminal
domain (CTD) of Pol II [147]. This modification pro-
motes the recruitment of histone H3-K4 histone meth-
yltransferases (HMTs) such as Set1 and MLL1,
typically as part of the COMPASS complex. This class
of HMTs introduces the H3-K4me3 mark, which is
predominantly found at the 5¢ ends of active or recently
transcribed genes [148]. In mammals, Set1 and MLL
exist in stable association with both WDR5, a protein
that specifically interacts with dimethylated and trime-
thylated H3-K4 [22,149,150], and MOF [149]. This pro-
vides a mechanism to both amplify H3-K4 methylation
and decondense chromatin by introducing AcH4-K16.
H3-K4me3 also recruits the Isw1 chromatin remodel-
ling ATPase to the promoter to prevent premature ini-
tiation of transcription elongation [137].
The initiation and elongating phases of the tran-
scription cycle are characterized by distinct sets of his-
tone and Pol II modifications [19,151]. Following
promoter clearance, the elongating phase of transcrip-
tion is associated with phosphorylation of CTD hepta-
peptide repeat Ser2 by P-TEFb ⁄ cdk9. This phase of
transcription can also be regulated by specific tran-
scription factors because P-TEFb can be recruited by
nuclear factor jB (NF-jB), c-Myc, MyoD and
GATA-1 [147]. Furthermore, recruitment of P-TEFb
by c-Myc is instrumental in releasing proximal paused
Pol II in many mammalian genes [152]. During the
elongation phase, the Pol II CTD Ser2 phosphate
modification plays a direct role in recruiting the HMT
Set2 which marks regions downstream of the promoter
with H3-K36me3. Histone phosphorylation can also
act as a trigger driving the onset of transcription elon-
gation in mammalian cells [153]. Phosphorylation of
histone H3S10 by Pim1 kinase enables the recruitment
of both MOF and P-TEFb via interactions involving
the adaptor protein 14-3-3 and the bromodomain pro-
tein BRD4 which is recruited via AcH4-K16 and phos-
pho H3 [153].
In yeast it is apparent that the histone acetylation
associated with transcription elongation is only a very
Active chromatin and DNase I hypersensitive sites P. N. Cockerill
2192 FEBS Journal 278 (2011) 2182–2210 ª 2011 The Author Journal compilation ª 2011 FEBS
transient event, whereby the H3-K36me3 (or me2)
modification plays a vital role in returning the chroma-
tin structure of a gene to the deacetylated state follow-
ing a cycle of transcription [19,21,54,130,154–158].
Hence, it is possible that RNA polymerase travels
within a moving window of decondensed active chro-
matin, with the chromatin structure returning to the
condensed state once the polymerase has passed. This
cycle is summarized in Fig. 5. The K36me3 or K36me2
H3 modifications, which are introduced during tran-
scription, can function as a docking site for Eaf3,
which is a component of the yeast Rpd3S histone
deacetylase (HDAC) complex [156]. This complex
directs histone deacetylation in the wake of the tran-
scribing polymerase and serves the important function
of suppressing spurious transcription from cryptic pro-
moters within genes [19,21,154–157,159]. Because this
process maintains the body of active genes in a con-
densed state most of the time it serves to suppress
cryptic transcription initiation. This is one reason why
active genes are not routinely observed as unfolded 10-
nm diameter chromatin fibres. Curiously, a similar
mechanism also operates immediately downstream of
the promoter in yeast genes, where H3-K4me2 engages
Set3 to recruit an HDAC complex containing the
Rpd3-like protein Hos2 [160]. This is thought to limit
the spread of nucleosome modification emanating from
promoter-associated factors. Hence, the maintenance
of a condensed chromatin structure throughout the
entire body of a gene may rely on the combined
actions of Set3 at the 5¢ end and Set2 at the 3¢ end.
In yeast the cycle of transient co-transcriptional
chromatin opening may be driven by factors carried
by the polymerase itself. The recruitment of SAGA
to transcribed coding regions is also supported by
Pol II CTD Ser5 phosphorylation by TFIIH
[161,162]. SAGA acetylates nucleosomes on histone
H3 and aids their eviction during transcription, and
is required for efficient transcription [161,162]. The
phosphorylation of Pol II CTD Ser5, plus the histone
H3 methylation by Set1 or Set2, also function to
promote co-transcriptional recruitment of NuA4, and
the subsequent acetylation of H4 promotes recruit-
ment of RSC which destabilizes nucleosomes [163].
However, the specific function of CTD Ser5 phos-
phorylation during transcription is unclear, because
TFIIH kinase activity can be suppressed without
major defects in the function of the elongating poly-
merase and its role may be more closely related to
mRNA capping [164].
The acetylation of H4-K16 by other HATs may
represent an additional important aspect of this cycle
that enables transcription elongation, because this
modification is known to be sufficient to trigger
unfolding of the condensed 30-nm diameter fibre.
However, whether AcH4-K16 is the key target for
Rpd3S within transcribed genes in yeast is not
entirely clear because, in contrast to human T cells
[124], AcH4-K16 is not typically enriched within
active gene coding sequences in yeast [165], and
Rpd3S is required for the deacetylation of all sites
except for H4-K16 within heterochromatin [166]. Nev-
ertheless, in Drosophila there is a direct relationship
between H4-K16 acetylation and H3-K36 methyla-
tion, whereby a reduction in H3-K36me3 leads to an
embryonic-lethal accumulation of acetylation specifi-
cally at H4-K16 [159]. In this study it was found that
dimethylated and trimethylated H3-K36 had opposing
effects. Hence, it was suggested that H3-K36me2
might function first to recruit an H4-K16 HAT such
as MOF to enable transcription elongation, followed
by conversion of the dimethyl to a trimethyl state
and the recruitment of an HDAC to reform a
repressed state. The principal role of H3-K36 methyl-
ation in mammalian cells remains far from clear. This
modification can recruit the HATs MOF and HBO1,
and both factors are found enriched within active
genes [124,130,140]. H3-K36me3 is recognized by
both JADE1, which is associated with HBO1 [140],
and MSL3, which is associated with MOF [128].
There is additional evidence from genome-wide
analyses suggesting that an acetylation ⁄ deacetylation
cycle also occurs during transcription in human T
cells [124,130]. The levels of both HATs and HDACs,
especially HDAC6, are higher within active genes,
and the levels present are in each case directly pro-
portional to gene activity [130]. The level of HDAC
recruitment also increases in direct proportion to the
level of histone acetylation present. In contrast, only
a minor proportion of HDACs are found associated
with inactive genes, and when HDACs are present
they are found in genes that are transcriptionally
poised by H3-K4 methylation [130]. A similar situa-
tion exists in yeast where the HDAC Hos2 was found
to be both preferentially associated with active genes
and required for efficient transcription [167]. Overall,
these studies suggest that HDACs do indeed play a
vital role in controlling the cycle of gene transcrip-
tion, and not just gene repression. However, one dif-
ference between yeast and humans is that, in yeast,
HDACs are recruited to active genes via H3-K36me3,
whereas in humans both Tip60 and HDAC6 appear
to be recruited to active genes directly by phosphory-
lated Pol II [130].
Two key issues that remain to be fully resolved in
this model of a cycle of histone acetylation and deacet-
P. N. Cockerill Active chromatin and DNase I hypersensitive sites
FEBS Journal 278 (2011) 2182–2210 ª 2011 The Author Journal compilation ª 2011 FEBS 2193
ylation are (a) which specific histone modifications are
the most important to open up chromatin structure
ahead of Pol II and enable efficient transcription, and
(b) what specific mechanisms might acetylate active
genes in association with the elongating Pol II. In
addition to yeast SAGA and NuA4, the mammalian
HAT HBO1 is also potentially able to be recruited to
and to acetylate H4 on K5, K8 and K12, as well as
H3 throughout active gene coding regions [139,140].
HBO1 may itself promote transcription elongation,
and it can be recruited to transcribed chromatin via its
close interaction with ING4 which binds H3-K4me3
and JADE1 which binds to H3-K36me3 [139,140].
JADE1 also has an additional PHD domain that inter-
acts with non-methylated histone H3, meaning that it
can direct recruitment of HBO1 both in advance of
and in the wake of a transcribing polymerase.
The other most likely candidates driving histone
acetylation during transcription are the Elongator
complex, which has intrinsic HAT activity [168], and
COMPASS, which can promote histone acetylation
indirectly [22]. Although COMPASS is mainly associ-
ated with promoters, both COMPASS and Elongator
can travel together with the elongating polymerase.
COMPASS employs Set1 to introduce methylated
H3-K4 which can then recruit HAT complexes.
H3-K4me3 is recognized by Chd1 within the SAGA
complex [22] and by Yng1 within the NuA3 complex
[169]. The Elp3 component of the Elongator complex
is a Gcn5-like HAT which, like SAGA and NuA3,
preferentially targets histone H3 and so is not an obvi-
ous candidate driving H4-K16 acetylation [170]. This
is more likely to require a histone H4 HAT such as
Sas2, Tip60 or MOF. However, the use of arginine-
substituted histones in yeast indicated that H3-K14
and H4-K8 are both significant targets of the human
Elongator HAT Elp3 [171].
Paradoxically, another potential mechanism for acet-
ylation also involves H3-K36me3. Eaf3, which recog-
nizes H3-K36, is a component of both the HAT NuA4
and the HDAC Rpd3S [21,154,155]. Furthermore,
NuA4-dependent acetylation of H4-K8 is decreased in
the absence of H3-K36 methylation [137]. However,
this mechanism of NuA4 recruitment to sites of tran-
scription seems unlikely because yeast NuA4 does not
bind to nucleosomes containing methylated H3-K36
[172]. Rpd3S recognizes this modification via the com-
bined actions of Eaf3 and the PHD domain protein
Rco1 [172]. If the Nu4 ⁄ Yng2 PHD domain is replaced
by the Rpd3S ⁄ Rco1 PHD domain, then this results in
mis-targeting to H3-K36me3 and also leads to activa-
tion of cryptic promoters within genes [172]. Further-
more, mutation of Eaf3 results in an increase in
histone acetylation, indicating that it cannot be the
principal factor recruiting HATs to sites of transcrip-
tion [154,155].
Histone acetylation regulates not just the unfolding
of the 30-nm fibre but also influences the ability to
unwrap DNA from the nucleosome. Histone H3-K56
plays a special role in this context. This lysine is part
of the globular core region of histone H3, and is
located at the point where the DNA exits from the
nucleosome. Acetylation of H3-K56 weakens the
interactions with DNA at this critical position and
thereby contributes to nucleosome disassembly at pro-
moters [173,174]. However, this acetylation event is
not thought to be coupled directly to the transcrip-
tion cycle, but involves the replacement of disassem-
bled nucleosomes at promoters with newly
synthesized acetylated histones [173]. This means that
nucleosomes at promoters are likely to be highly
dynamic once they have undergone a cycle of nucleo-
some replacement. The many additional lysine acety-
lation events directed by transcription factors will
further loosen contacts between histone tails and
DNA to create a more open structure that is more
readily mobilized by remodellers and transcription
complexes.
Localized chromatin modifications
within regulatory elements
Gene expression control by distal and proximal
regulatory elements
It is typical for higher eukaryotic genes to be con-
trolled not just by the proximal promoter but by one
or more distal elements as well. These include elements
that have been defined as either enhancers or locus
control regions (LCRs), depending on the assay used
to identify them. In some cases these elements are
located far upstream or downstream, or inside genes,
and even inside adjacent genes. Hence, it is not always
obvious which elements control which genes, and the
identification of essential distal elements typically has
to be accomplished by experimental means using
genetic manipulation.
The first LCR to be formally identified was the
mammalian b-globin LCR, which exists as a cluster of
DHSs, including a classical enhancer, several kilobases
upstream of the b-globin gene cluster [175]. This LCR
was first found to be required for correctly regulated
expression of transgenes and for the expression of all
globin genes, but it was then found that it is not actu-
ally essential for the maintenance of an open chroma-
tin structure [176]. In essence, an LCR is a region that
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includes an enhancer that is required for correct
expression of a transgene.
The concept of a gene having a single essential LCR
is itself somewhat outdated, as others have established
that transgenes can also be correctly expressed by
inserting a full complement of cis regulatory elements,
and there is not necessarily one dominant control ele-
ment [177]. Furthermore, gene locus activation typi-
cally involves direct interactions within chromatin that
bring multiple upstream and downstream elements
together with the promoter at sites that have been
termed active chromatin hubs [178–181]. This process
involves chromatin looping to bring all these elements
together. Furthermore, active chromatin hubs can
incorporate several co-expressed active genes [182],
plus their enhancers, and may in fact be no different
to the active sites termed transcription factories
[183,184]. The co-localization of many active promot-
ers and enhancers at one site is an efficient way of sus-
taining active transcription by maintaining threshold
levels of polymerases and transcription factors. How-
ever, there is also some evidence indicating that any
one specific enhancer can only activate one target gene
at any one moment in time [185].
Distal regulatory elements such as enhancers and
LCRs recruit many of the same factors as promoters,
and in many cases even support non-coding (nc) RNA
transcription [186,187]. Their main mechanism of
action may therefore be simply to supply factors to the
promoter at the chromatin hub. However, some distal
elements appear to function not by looping but by
directing ncRNA transcription in the direction of the
locus to be activated. For example, in the case of the
pituitary-specific human growth hormone locus,
ncRNA transcription initiates within an LCR located
15 kb upstream of the gene and is required for efficient
expression, even though it does not reach the gene
itself [188]. These ncRNA transcripts proceed towards
the gene, through a non-expressed B cell specific gene,
but terminate several kilobases before the actual
growth hormone gene. In other cases, such as the yeast
fbp1 gene, ncRNAs initiate upstream of the gene, tran-
scribe through the gene, and progressively convert the
chromatin to an open configuration [189]. Within the
b-globin locus, ncRNA transcription plays a role in
the development control of globin gene switching from
fetal e-globin to adult b-globin gene expression [190].
Genetic recombination in T cells and B cells is also
enhanced by ncRNA transcription. In the T cell recep-
tor a (TCR-a) locus, ncRNA transcription from an
upstream element proceeds through the V regions, and
is required for efficient recombination, presumably
because it establishes an open chromatin structure
[191]. The IgH locus intronic transcripts may serve the
same function in B cells [187].
Active promoters appear as nucleosome-free
regions together with variant histones
As summarized in a recent review paper [26], promot-
ers can be divided into (a) constitutively active open
promoters that are intrinsically depleted of nucleo-
somes and (b) highly regulated covered promoters
where specific transcription factors act to disrupt a
positioned nucleosome and create a nucleosome-
depleted region [26,192]. Nucleosomes tend to natu-
rally assemble at intrinsically defined positions,
determined by the underlying sequence, and nucleo-
some positioning evolves in parallel with the specific
mode of regulation of a promoter [193]. Open promot-
ers often contain poly(dA):poly(dT) tracts which are
not easily assembled into nucleosomes. Along the same
lines, promoters can also be roughly divided into (a)
TATA-containing promoters where the TATA is
blocked by a nucleosome which can be disrupted by
regulated factors which activate the gene, and (b)
TATA-free constitutively active promoters that intrin-
sically exist in nucleosome-free regions [26,194]. Cov-
ered promoters are naturally more dependent on
chromatin remodelling complexes [26].
Constitutively open promoters, as well as induced
active promoters, are typically depleted of a single
nucleosome upstream of the transcription start site,
and have positioned nucleosomes directly adjacent
which contain the histone variant H2AZ [195–197].
Regions immediately downstream of active or recently
active promoters are also enriched in the histone vari-
ant H3.3 [104–106]. One reason why histones H2AZ
and H3.3 are enriched at sites of transcription is
because they are the predominant replacement histones
used by the genome during interphase. This is also
consistent with studies showing that there is a high
rate of turnover of nucleosomes at active promoters
and their flanking regions [15,26]. In yeast, H2AZ is
found at most promoters, not just active promoters
[26,196]. However, another major reason for the pro-
moter-specific localization of H2AZ is the genome-
wide INO80-directed removal of non-acetylated H2AZ
at sites other than promoters [198].
Although open promoter regions are often assumed
to be nucleosome-free, a recent study has found that
DHSs throughout the genome are occupied by highly
unstable nucleosomes containing both H2AZ and H3.3
[107,199,200]. This specific combination renders nucle-
osomes so unstable that they typically either disassem-
ble or are digested away during the assay process of
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FEBS Journal 278 (2011) 2182–2210 ª 2011 The Author Journal compilation ª 2011 FEBS 2195
defining nucleosome locations. This is partly because
H2AZ nucleosomes only protect  120 bp of DNA
from MNase, not 146 as is the case for conventional
nucleosomes [201]. Hence, the best way to view DHSs
is as highly dynamic sites that alternate between being
assembled into an unstable H2AZ ⁄ H3.3 nucleosome,
being nucleosome-free, and being bound by factors.
Furthermore, H2AZ and H3.3 are not found just at
promoters but also at other classes of regulatory ele-
ment such as enhancers, insulators and Polycomb
response elements [15,199,200]. H2AZ is enriched at
TATA-less promoters and may therefore play a role of
maintaining this class of promoter in an open confor-
mation [196].
Many promoters contain CG islands that form
unstable nucleosomes
The majority of CG sequences in the genome carry the
5-methylcytosine modification that is introduced and
maintained by DNA methyltransferases (DNMTs) on
both strands [202,203]. Furthermore, most of the
ancestral CG elements in the genome have been lost
during the course of evolution due to the high muta-
tion rate of methylated cytosine to thymidine. How-
ever, in mammals, 70% of promoters exist within CG
islands which are characterized by their very high CG
content and have an intrinsic resistance to DNA meth-
ylation. CG-island promoters tend to be constitutively
and ⁄ or ubiquitously active promoters and it is likely
that their constant state of activity overcomes mecha-
nisms that would lead to repression by DNA methyla-
tion. A recent study has uncovered another intrinsic
property of CG islands that helps to explain their
functions as promoter elements. Their general G ⁄ C-
richness is unfavourable for nucleosome assembly,
meaning that CG islands form inherently unstable
nucleosomes [204]. This allows some factors to readily
engage their targets within chromatin, which under
other circumstances would be dependent upon remod-
elling factors such as SWI ⁄ SNF to render their binding
sites accessible [204].
CG islands maintain active chromatin domains
free of DNA methylation
DNA methylation is one of the most prevalent mecha-
nisms employed within the genome to maintain inac-
tive regions in a repressed state, and it is also one of
the most stable modifications [202,203]. Although
DNA methylation often has little effect on gene
expression when it is dispersed at low level throughout
the body of a gene, it is highly repressive when present
at high density at CG islands [203]. The many poten-
tial mechanisms controlling the balance between acti-
vation and repression of CG islands are represented in
Fig. 6. Methylated CG elements recruit proteins con-
taining methyl binding domains (MBDs) which assem-
ble complexes containing repressive HMT and DNMT
proteins that act in a concerted fashion to maintain
both DNA methylation and H3-K9 methylation [202].
However, CG islands usually exist as constitutively
active regions that exclude DNA methylation, and
thereby evade this mechanism of repression. CG
islands typically carry the H3-K4me3 modification and
are maintained in a constitutively active state by
CXXC domain proteins that bind to non-methylated
CG sequences. These proteins include MLL1 and the
CG-binding protein Cfp1 that itself recruits both Set1
and MLL1 [205–208]. This provides a mechanism to
maintain H3-K4me3 even in the absence of transcrip-
tion [208], and this modification also suppresses
recruitment of the DNMT3a ⁄ DNMT3L complex
which directs de novo DNA methylation [209]. How-
ever, many CG islands are in fact promoters [203,210],
meaning that they can also recruit Set1 which intro-
duces H3-K4me3 in a transcription-dependent manner
(Fig. 6) [148]. CG islands also recruit the CXXC
domain protein KDM2A, which demethylates H3-
K36me2 [211], and may thereby prevent the recruit-
ment of HDAC complexes that recognize this modifi-
cation. In yeast it is known that H3-K36me2 and H3-
K36me3 can suppress transcription by recruiting
HDACs [154–157]. If a similar mechanism operates in
mammalian cells, then KDM2A may suppress this
pathway and maintain histone acetylation at tran-
scribed CG islands.
It is becoming increasingly likely that CG islands
remain free of DNA methylation because they also
recruit factors that lead to elimination of DNA meth-
ylation. Foremost among these is the CXXC domain
protein Tet1 that belongs to a family of proteins that
convert 5-methylcytosine to 5-hydroxymethylcytosine
[212,213]. This may subsequently lead to loss of the
modification either directly by excision or indirectly if
the hydroxy methylated state is not recognized by
maintenance DNMTs following DNA replication.
Human cells are also known to produce DNA glycosy-
lases capable of excising both 5-hydroxymethylcytosine
[214] and 5-methylcytosine [215]. The best candidates
for such glycosylases are the thymine glycosylases
TDG and MBD4 [215–217] which can remove the
methyl cytosine base [218] and most likely remove
hydroxymethylcytosine as well. Methylated cytosines
resemble thymine, which allows for some cross-reactiv-
ity. TDG may perform this function more widely in
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2196 FEBS Journal 278 (2011) 2182–2210 ª 2011 The Author Journal compilation ª 2011 FEBS
the genome as it can be directly recruited to promoters
by transcription factors, presumably to regulate DNA
methylation [215], while MBD4 also contains an MBD
that can target MBD4 to methylated CG [216]. The
role of Tet1 in the control of DNA methylation levels
within specific target genes was confirmed by studies
which found that downregulation of Tet1 expression
led to decreased expression and increased DNA meth-
ylation at the Nanog locus [213]. However, a different
conclusion was drawn from a study of myeloid malig-
nancies which found that TET2 mutations were associ-
ated with a slight decrease, not an increase, in the
average level of DNA methylation [219]. Paradoxically,
in the same study it was found that overexpression of
TET2 in cell lines led to the disappearance of DNA
methylation detectable by methylcytosine antibodies.
Hence, it is still too early to draw firm conclusions as
to the true in vivo role of TET family proteins in the
control of DNA methylation and genome function.
CG islands can also be targets for dysregulation in
diseased states whereby they can become hypermethy-
lated and maintained in a repressed state by MBD
proteins [202]. As summarized in Fig. 6, repressed CG
islands are likely to be maintained in an inactive state
by a variety of cooperating repressive complexes. In
particular, these are likely to involve HP1, which binds
to H3-K9me3, and methylated DNA binding proteins
that recruit the transferases that introduce this modifi-
cation. The high density of CG elements found within
CG islands may also help explain why they are typi-
cally maintained predominantly in either the highly
methylated or the highly demethylated state. This is
because any repressive modification is likely to be rap-
idly removed if it occurs at an active CG island,
whereas any active chromatin modifications or DNA
demethylation events are likely to be rapidly reversed
if they occur in a repressed CG island. In contrast,
similar events occurring at isolated CG elements are
not under the same pressure and are therefore likely to
behave in a more independent fashion. Hence, it is the
concerted action of many cooperating factors that
drives the balance towards one state or the other.
Active cis regulatory elements exist as DHSs
The binding of transcription factor complexes to the
different types of cis regulatory elements described
above typically generates stretches of  150–400 bp of
Active CG island
Repressed CG island
HDAC
MBD
Me
CG
Me
CG
Me
CG
HMT
Me3K9
H3
Ac
DNMT
Me3K9
H3
HMT
Me
CG
TF
MLL1
CXXC
OH
Me
CG
CG
CG
Tet1
CXXC
MLL1
CXXC
Me3K4
H3
Ac
CG
CG
TF
?
DNMT3L
Cfp1
CXXC
Set1
DNMT3
Tet2
HP1
HP1
Me
CG
KDM2A
CXXC
CG
KDM2A
CXXC
Me2K36
H3
Me
H3
K36
HDAC
MBD4
TDG
?
OH
meC
+
meC
Active CG island
promoter
OH
Me
CG
CG CG
Tet1
CXXC
MLL1
CXXC
Me
3
K4
H3
Ac
CG CG
TF?
DNMT3L
Cfp1
CXXC
Set1
DNMT3
Tet2
CG
KDM2A
CXXC
Me
H3
K36
CCCCGCCCC
Sp1
Pol II
Set1
TAF
TFIIB
OH
Me
CG
Tet2
Fig. 6. Models depicting the factors that act
in concert to maintain either an active or a
repressed state at CG islands. Activators
are shown in green and repressors are
shown in red.
P. N. Cockerill Active chromatin and DNase I hypersensitive sites
FEBS Journal 278 (2011) 2182–2210 ª 2011 The Author Journal compilation ª 2011 FEBS 2197
highly accessible DNA that exist as DHSs. If the same
elements are examined in tissues where they do not
function, then they are unlikely to exist as DHSs. This
general description covers most classes of regulatory
elements, including enhancers, promoters, LCRs,
silencers, insulators and BEs. An excellent example of
this general concept is the human GM-CSF enhancer
which forms an inducible DHS in all cell types where
inducible enhancer activity can be detected, and only
in those cell types [220]. In cases such as this, DHSs
are highly dynamic and can form and disappear in
parallel with the induction and depletion of specific
factors. As also discussed in detail elsewhere in an
accompanying review in this issue [146], steroid-induc-
ible elements similarly form highly dynamic DHSs.
For example, at the rat TAT gene enhancer, induction
of a steroid receptor leads to induction of a DHS 2 kb
upstream of the gene [221]. This is an extremely rapid
and dynamic event, because the DHS forms within just
10–20 min of induction by steroid, and it disappears as
fast as it forms once steroid is withdrawn [222]. Fine
mapping of this DHS indicates that nucleosomes are
destabilized within the DHS, as opposed to being
repositioned sideways, and linker histones are depleted
[223]. Once DHSs at enhancers and promoters have
been created, they invariably incorporate the variant
histones H3.3 and H2A.Z either within or directly
adjacent to the DHS [107,200].
One mechanism likely to be employed to disrupt
chromatin at many DHSs is the recruitment of the
SWI ⁄ SNF family of chromatin remodelling enzymes
[24–26,224]. These factors are noted for their ability to
remodel or eject nucleosomes in a highly dynamic pro-
cess by disturbing DNA–histone interactions. Unlike
ISWI, SWI2 ⁄ SNF2 is able to create open loops of
DNA within nucleosomes, as well as structures resem-
bling di-nucleosomes [24–26,224]. This SWI⁄ SNF
remodelling activity is opposed by other remodellers
such as ISWI which tend to recreate regular repressed
chromatin structure [24–26]. SWI2⁄ SNF2 recruitment
by transcription factors is a widely used mechanism of
gene activation. For example, several zinc finger tran-
scription factors, including EKLF, can directly recruit
SWI2 ⁄ SNF2 to chromatin to create a DHS [225]. In
parallel, incorporation of H2AZ into flanking nucleo-
somes is likely to be mediated by the related remodel-
ler SWR-C [198].
DHSs can also be erased by specific mechanisms. In
the chicken lysozyme locus, a DHS at a CTCF-depen-
dent silencer is erased by the process of inducible
ncRNA transcription which leads to eviction of CTCF
and the repositioning of a nucleosome over the CTCF
site [226]. Different classes of remodelling enzymes can
have opposing actions on chromatin. In one such
example, TFE3 recruits ACF to create a DHS within
the IgH intronic enhancer, whereas PU.1 can subse-
quently recruit Mi2b to erase this DHS [227]. Interest-
ingly, in this example, PU.1 remains bound to the
enhancer even after the DHS is erased.
Historically, it has been very difficult to say defini-
tively whether or not a specific DHS does or does not
still contain a nucleosome. DHSs are themselves
dynamic, and a certain percentage of cells in a popula-
tion will still retain nucleosomes within a DHS even if
the nucleosomes have been evicted from the DHS in
the majority of cells. DHSs can also contain destabi-
lized nucleosomes that do not protect a conventional
146 bp from MNase, and can contain histone variants.
Hence, we should avoid trying to rigidly define DHSs
as always representing nucleosome-free regions as
opposed to being regions containing modified nucleo-
somes. These are dynamic structures, and both of these
situations are likely to be encountered at different
moments in time or at different regulatory elements.
However, due to the dynamic nature of DHSs, it is
usually possible to design alternative experiments to
show that within a population of cells DHSs can exist
as either nucleosomes or conversely as nucleosome-free
regions [85,86,200,223]. ChIP assays do not always
adequately address this issue because they typically
show that one of several chromatin states can exist,
not that it is the only state at a specific DHS. Never-
theless, for the most part it has been widely assumed
that DHSs do represent nucleosome-free regions, and
in many cases this appears to be true at least part of
the time [15]. This is best typified by (a) the EM
images of the SV40 viral mini-chromosome which
reveal a region of  400 bp spanning the enhancer,
promoter and origin of replication which is devoid of
nucleosomes [10], and (b) the yeast PHO5 promoter
which loses contact with histones upon activation
[228,229].
Fine structure of DHSs
DHSs typically occupy a region that would otherwise
be occupied by one or two, and sometimes three,
nucleosomes. There are now many well defined exam-
ples whereby nucleosomes occupying promoters or
enhancers are replaced by multi-protein complexes of
regulatory factors termed enhanceosomes [230]. These
complexes are created via the cooperative assembly of
multiple transcription factors, plus co-factors such as
CBP, to several closely linked binding sites within a
discrete region. A corollary of this concept is that
binding sites for transcription factors are not normally
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widely distributed across a broad region but are tightly
clustered to enable these cooperative interactions. Sig-
nificantly, CBP and p300 have multiple specific interac-
tion surfaces that can interact with several different
transcription factors within the same complex, and it
has been proposed that these enzymes serve an addi-
tional structural role as scaffolds for the assembly or
stabilization of enhanceosomes [231]. These highly
complex structures are unlikely to permit coexistence
of a nucleosome. DHSs can show considerable varia-
tion and complexity in the number of nucleosomes dis-
rupted, the histone composition of the remaining
nucleosomes, and the number and nature of protein
complexes they interact with.
Much has been learnt from well studied model sys-
tems of inducible DHSs such as the yeast PHO5 pro-
moter [228,229], the interferon-b promoter [230] and
the human GM-CSF enhancer [85,86]. Activation of
the GM-CSF enhancer in T cells is accompanied by
replacement of two positioned nucleosomes (N1 and
N2) by two multi-protein complexes that each interact
with and protect  50–100 bp of DNA (Fig. 4). This
seems to be a dynamic and stochastic process because
a low level of nucleosome occupancy remains after
activation [85,86], and not all cells in a population
express the GM-CSF gene upon stimulation [232].
Nevertheless, the enhanceosome-like complexes are
stable enough to generate strong footprints in MNase
and DNase I digestion assays [85,86]. Figure 4 depicts
a hypothetical model of the organization of nucleo-
somes and enhanceosomes within the enhancer after
activation. In this model, I have taken care to depict
all the components as close as possible to their actual
size. Note that the length of DNA exposed by loss of
just two nucleosomes is very extensive compared with
linker regions and that the enhanceosome complexes
are much bigger than the nucleosomes they replace.
The key driver in this model is likely to be NFAT,
which is essential for the creation of many DHSs
observed in mammalian cytokine genes [91]. NFAT,
in turn, plays a major role in assisting the binding of
AP-1 via cooperative binding, and creates an accessi-
ble environment that also enables Sp1 and Runx1
binding [85,86,233]. Both NFAT and AP-1 have the
potential to recruit the HATs CBP and p300 which
may allow the formation of stable enhanceosomes.
AP-1, and potentially NFAT, are also able to recruit
SWI ⁄ SNF family complexes which most probably
contribute to both the disruption and ejection of
nucleosomes within the enhancer and the mobilization
of nucleosomes in the flanking sequences. AP-1 func-
tion is dependent upon the SWI⁄ SNF family protein
Brg1 [234]. NFAT is also known to be required for
the recruitment of the SWI ⁄ SNF family protein Brg1
to the DHS at the IL-5 ⁄ IL-4⁄ IL-13 LCR in T cells,
where Brg1 is required for the creation of this DHS
[235].
In activated mast cells the fine structure of the DHS
within the GM-CSF enhancer is even more complex
than in T cells, with several peaks of nuclease hyper-
sensitivity [85]. Mast cells express GATA-2 as well as
NFAT and AP-1, and GATA-2 initiates the formation
of an additional discrete GATA-2 ⁄ AP-1 enhanceo-
some-like complex existing upstream of the two NFA-
T ⁄ AP-1 complexes. Evidence for the existence of these
distinct enhanceosomes is provided in the form of the
discrete protected regions that are visible when the
GM-CSF DHS is analysed at high resolution in acti-
vated T cells and mast cells [85] (Fig. 4).
Interactions between transcription
factors and chromatin
It is generally accepted that most transcription factors
do not have an intrinsic ability to bind efficiently to
recognition sequences within chromatin, and it is also
evident that most consensus sequences within the gen-
ome are not utilized. This is for a variety of reasons.
Most importantly, chromatin normally exists in a
highly folded condensed state that renders most sites
inaccessible. Even if a nucleosome is accessible, most
transcription factors are likely to find it difficult to
bind to a target sequence if it is assembled into a
nucleosome, especially if the binding site is oriented
towards the surface of the nucleosome. Hence, the key
to efficient interactions between transcription factors
and chromatin lies in the concentrated effect of multi-
ple factors that cooperate in the destabilization of
chromatin at specific sites located close together. Part
of this cooperativity comes from the fact that some
transcription factors have specialized functions in
either opening up chromatin or recruiting nucleosome
destabilizing complexes. Furthermore, a key aspect of
gene control is likely to be the maintaining of posi-
tioned nucleosomes over regulatory elements until the
appropriate signals are delivered that are able to desta-
bilize these nucleosomes.
Pioneer factors gain entry into compacted
chromatin fibres
There is evidence that the initiation of chromatin de-
condensation and the assembly of transcription factor
complexes is a two-stage process. In the first phase,
specialized transcription factors termed pioneer factors
perform the role of creating accessible sites within
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FEBS Journal 278 (2011) 2182–2210 ª 2011 The Author Journal compilation ª 2011 FEBS 2199
histone H1 compacted chromatin fibres, without dis-
rupting the underlying nucleosomes [236,237]. Once an
opening has been created, then binding sites for other
transcription factors become more accessible. Fork-
head family transcription factors such as FoxA1 and
FoxO1 are the best known example of pioneer factors
[236,237]. Forkhead proteins contain a winged helix
domain that has structural similarities with histone
H1. Hence, these proteins may compete with histone
H1 in condensed chromatin fibres to create localized
disruptions within repressed chromatin domains.
FoxA1 and FoxO1 interact with histones H3 and H4
and can thereby disrupt inter-nucleosomal interactions
[236,237]. Because of the unique role that they fulfil,
pioneer factors should not be confused with other fac-
tors that act more generally to destabilize individual
nucleosomes but may not be able to bind to repressed
chromatin.
Transcription factors cooperate to destabilize
nucleosomes
Several studies have shown that there is an intrinsic
cooperativity in the binding of transcription factors to
nucleosomes [238–240]. Even in the absence of pro-
tein–protein interactions, factors can assist each other’s
binding to DNA within nucleosomes. This principle is
illustrated in the model shown in Fig. 7A. In this sce-
nario, factor B is only able to bind to an internal bind-
ing site within the nucleosome once factor A has
disrupted the histone–DNA contacts at a site closer to
the nucleosome boundary. This is essentially a simple
case of factors working together to progressively unzip
the nucleosome. An excellent example of this principle
comes from the work of the Workman laboratory
which found that binding of purified transcription fac-
tors to in vitro assembled mono-nucleosomes is inher-
ently cooperative and is very context dependent [238].
This study found that GAL4, USF or NF-jB alone
were inefficient at binding to nucleosomes if the bind-
ing site was centred 46 bp from the nucleosome
boundary, but they could bind to a site centred 20 bp
inside the boundary. However, once any one of these
factors was bound at the periphery, a second different
factor was then able to bind efficiently to a more inter-
nal site. As depicted in Fig. 7B, four different exam-
ples of pairs of binding sites were employed in the
study to demonstrate this principle [238]. The effect
was clearly mediated at the level of chromatin destabi-
lization, because no such dependence was observed on
free DNA. One reason why factors can bind to the
first 20 bp of nucleosomal DNA near the nucleosome
boundaries is because the DNA–histone interactions
are relatively weak in these regions [241]. Note that
this progressive unwrapping of DNA from the nucleo-
some is also likely to be supported by H3-K56 acetyla-
tion, which underlies the DNA exit points, but
suppressed by histone H1 or PARP-1 which cover the
DNA exit points.
Nucleosome positions in the genome are likely to
evolve in the context of transcription factor binding
sites. For example, in the osteocalcin promoter,
RUNX1 binds to a site located at the point where
DNA exits the nucleosome, which makes it easier for
RUNX1 to disrupt DNA–histone contacts and engage
its binding site [242]. RUNX1 cannot bind if the same
site is assembled further inside a nucleosome.
There are many other instances where it has been
shown that transcription factors cooperate to bind to
DNA within nucleosomes. For example, several mole-
cules of GATA-1 can act together to disrupt a nucleo-
some assembled onto an array of six GATA sites
[243]. NF-E2 is able to promote GATA-1 binding to
chromatin [244]. In some cases there is a clear hierar-
chy in binding. Steroid receptors can bind to nucleo-
somes and promote the binding of other factors such
as NF1 which do not bind alone [245,246]. NF1 is
probably typical of a class of factor whose binding is
dependent upon prior disruption of the underlying
chromatin [247,248]. Sp1 is a ubiquitous factor that is
also thought to be unable to bind to chromatin unless
other factors or intrinsic nucleosome positioning
A B
Factor B cannot
bind alone
Factor A alone
can bind
Factor A promotes
binding of factor B
155 bp DNA templates for
nucleosome assembly
A
20
44
A B
B
Gal4 USF
18
43
Gal4
USF
20
44
NF- kBUSF
20
46
Gal4 NF-kB
Nucleosome length DNA
Fig. 7. Transcription factors cooperate in binding to and disrupting
nucleosomes. (A) Many transcription factors have difficulty binding
to internal sites in nucleosomes. This model depicts a nucleosomal
length fragment of DNA assembled into nucleosomes. In this sce-
nario, a factor can readily bind to a site near the edge of a nucleo-
some but not to a site closer to the nucleosome dyad. However, if
one factor binds first to destabilize histone contacts near the DNA
exit point, then this creates enough destabilization and accessibility
to allow a second factor to bind to an internal adjacent site. (B) Pre-
viously defined examples of DNA templates and combinations of
transcription factors that conform to the rules of engagement
depicted in (A) [238].
Active chromatin and DNase I hypersensitive sites P. N. Cockerill
2200 FEBS Journal 278 (2011) 2182–2210 ª 2011 The Author Journal compilation ª 2011 FEBS
maintain its binding sites as nucleosome-free regions
[86,248].
Some transcription factors clearly have the ability to
bind to internal sites within nucleosomes while others
do not. For example, Fos⁄ Jun heterodimers are able
to disrupt nucleosomal structure and can promote the
binding of other factors such as SRY that do not bind
in the absence of Fos ⁄ Jun [239]. In contrast, Sp1 can
only bind very weakly to in vitro reconstituted nucleo-
somes [249]. Relative to its binding to free DNA, Sp1
binding to nucleosomes in vitro is 20-fold weaker at
peripheral sites in nucleosomes and 100-fold weaker at
central sites [249]. Furthermore, there is in fact little
evidence that Sp1 does bind to nucleosomes in vivo in
the absence of other factors [86,240]. Interestingly, the
relatively weak affinity of Sp1 for nucleosomes is
reflected by the observations that Sp1 sites typically
reside in constitutively active CG island promoters
which form unstable nucleosomes [204], or exist in
TATA-less promoters which tend to exclude nucleo-
somes [250].
There is considerable additional evidence from
in vivo footprinting studies for cooperative binding of
transcription factors to nucleosomes. In the GM-CSF
enhancer, Sp1 and RUNX1 are unable to bind to their
sites, which are located within positioned nucleosomes,
until these nucleosomes are disrupted by other induc-
ible factors [86,233]. In response to induction of
NFAT and AP-1, these two nucleosomes are evicted
and replaced by NFAT ⁄ AP-1 ⁄ Sp1 and NFAT⁄ AP-1 ⁄
RUNX1 enhanceosome-like complexes (Fig. 2) [85,86].
Others have also found that Sp1 is unable to bind to
inducible promoters prior to induction [248]. Similarly,
in a TCR gene enhancer, CREB and RUNX1 are
unable to bind to their sites in vivo if adjacent TCF or
ETS sites are mutated [240]. In many cases, it appears
that in vivo binding of factors does indeed correlate
with nucleosome eviction [85].
Some transcription factors are invariably associated
with DHSs. A genome-wide analysis of glucocorticoid
receptor binding to chromatin found that it always
bound at DHSs, whereby it either bound to a pre-
existing DHS or it induced a DHS upon binding [251].
The Ca
2+
-inducible transcription factor NFAT is vir-
tually always associated with Ca
2+
-inducible DHSs
that are suppressed by the drug cyclosporin A which
targets NFAT [91]. NF-jB was also first identified by
virtue of its ability to induce a DHS at the Igj enhan-
cer, and is closely linked with DHS induction.
The challenge in this field will be to distinguish
between (a) factors that intrinsically disrupt nucleo-
somes, analogous to the role of pioneer factors in open-
ing up condensed chromatin fibres, (b) groups of
factors that bind en masse to cooperatively disrupt nu-
cleosomes, and (c) factors that rely on the recruitment
of remodelling activities. In reality, we should expect to
find that most chromatin remodelling events and fac-
tors that contribute to DHS formation are likely to be
dependent upon ATP-dependent chromatin remodel-
lers. For example, NF-E2 is thought to be instrumental
in the creation of a DHS at HS2 of the human b-globin
LCR, via a process which is ATP-dependent, and this
enables the subsequent binding of GATA-1 [244].
Conclusions
It is clear that active chromatin has a structure that is
very different from inactive chromatin and that struc-
tural studies are indeed required to gain a detailed
understanding of the true nature of active chromatin.
There are many levels at which gene activation is facil-
itated by increasing chromatin accessibility, whether it
be by unfolding 30-nm chromatin fibres, modifying the
histones, or mobilizing and evicting nucleosomes. It is
equally important that chromatin is able to return to
an inactive state once a cycle of transcription has com-
pleted. This whole process is supported by highly
cooperative processes involving multiple transcription
factors, enzymes and remodellers to ensure a correct
pattern of regulation of the genome.
Acknowledgements
I thank Constanze Bonifer for her helpful advice and
comments on the manuscript, and Karolin Luger and
Jacques Cote for informative discussions. I thank Gra-
ham Goodwin for giving me a good initiation into the
study of active chromatin. I would like to acknowledge
the role of the late Dontcho Staynov in initiating the
project that resulted in this collection of review papers.
The work of Peter Cockerill is supported by Leukae-
mia and Lymphoma Research, the Biotechnology and
Biological Sciences Research Council, and Yorkshire
Cancer Research.
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