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

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

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

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

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

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

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

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-

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-

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

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

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

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].

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

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

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

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-

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

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,

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-

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

(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

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

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

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
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 2195
defining nucleosome locations. This is partly because
H2AZ nucleosomes only protect  120 bp of DNA

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

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
Active chromatin and DNase I hypersensitive sites P. N. Cockerill
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

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

?
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

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

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

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

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
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 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

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-

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-

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

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