Structure and function of active chromatin and DNase I hypersensitive sites

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1 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 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: Tel: p.n.cockerill@leeds.ac.uk 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 hypersensitive sites. (Received 18 December 2010, revised 10 February 2011, accepted 5 April 2011) doi: /j x 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 recognized that chromatin structure was likely to play a significant 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 explosion of papers on the multitude of chromatin modifications 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 promoters, gene bodies, termination regions and even intron exon boundaries have very characteristic signatures of histone modifications, histone replacements and 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 FEBS Journal 278 (2011) ª 2011 The Author Journal compilation ª 2011 FEBS

2 P. N. Cockerill Active chromatin and DNase I hypersensitive sites 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 commonplace 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 chromatin 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 conformation per se. To understand the basis of the fundamental mechanisms 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 utilized 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 structure, 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 experiences using the highly inducible human granulocytemacrophage colony-stimulating factor (GM-CSF) gene as a model system that undergoes extensive remodelling. It is beyond the scope of this review to enter into an extensive discussion of the role of all the various 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 acetylation. 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 comprise 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 histones within a nucleosome are depicted in Fig. 1, Upper H2A/H2B dimer H2A H2B Split view H2A H2B Top half H4 H3 Tetramer H3 H3 H4 Octamer bp DNA H4 + H3 H4 H2B H2A Bottom half Lower H2A/H2B dimer H2A H2B 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 contacted 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 contacting 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. FEBS Journal 278 (2011) ª 2011 The Author Journal compilation ª 2011 FEBS 2183

3 Active chromatin and DNase I hypersensitive sites P. N. Cockerill which is a greatly simplified version of the X-ray crystal structure obtained at 2.8 A resolution [32] but is used here to convey the concept that an H3 H4 tetramer 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 bp. Most nucleosomes also recruit either histone H1 or high mobility group (HMG) proteins (sometimes both) which bind to the outside of the nucleosome 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 maintaining 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 coiling of 10-nm diameter fibres into 30-nm diameter fibres [37,39]. The 30-nm fibre represents the predominant 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 represented 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 histone 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 genome is assembled in a much more condensed state under physiological conditions, and exists in conformations 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 nm in diameter during interphase [48] and 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 induction of transcription. However, after transcription activation, 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 mammary 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 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 conventional chromatin fibre remains. The concept of the 30-nm fibre as the universal building block of chromatin in vivo has also been challenged by an independent cryo-em analysis of metaphase 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 nucleosomes within chromosomes exist primarily in a disordered interdigitated state, rather than conforming to the well organized helical structures observed for in vitro reconstituted chromatin fibres. The Balbiani rings observed in polytene chromosomes in Chironomus tentans provide another representation 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 downstream 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 intervening DNA can be seen typically as either nucleosomefree or as a 10-nm fibre. However, even in these highly 2184 FEBS Journal 278 (2011) ª 2011 The Author Journal compilation ª 2011 FEBS

4 P. N. Cockerill Active chromatin and DNase I hypersensitive sites 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 nucleosome-free, but that coding regions return to a conventional 30-nm diameter chromatin fibre once a polymerase has passed. The challenge presented here, then, is to gain a better 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 condensed 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 chromatin 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 polymerases are bound and any genes which are so loaded with polymerases that this does not permit the reassembly 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 heterochromatin 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 decondensed 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 acetylation 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 chromatin domains encompassing active genes are at least twice as sensitive to DNase I digestion as non-transcribed 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 restriction 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 encompassed. 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-globin 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 protected 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 hypersensitivity 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 regulatory 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 initially 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- FEBS Journal 278 (2011) ª 2011 The Author Journal compilation ª 2011 FEBS 2185

5 Active chromatin and DNase I hypersensitive sites P. N. Cockerill expressing cells only [64]. In chicken embryo erythrocytes, 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 closely 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 enhancer 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 enhancerblocking 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 polytene 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 complexes that create islands of active chromatin which counteract the repressive complexes that mediate heterochromatin 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 repressive 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 functional DNA domains within which enhancers and promoters can cooperate, as opposed to demarcating active chromatin domains. Active loci undergo extensive nucleosome mobilization Classical models of chromatin depict chains of regularly spaced nucleosomes that fold up into a helix as highly ordered chromatin fibres. However, this image is really only representative of inactive loci that constitute the bulk of chromatin in the nucleus. The highly regular ordering of nucleosomes is more closely associated 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 relatively few studies which have assessed the organization as opposed to the modification status of active chromatin. 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 conforming to any of the current models of the higher order chromatin fibre. It is even possible that this highly disorganized form of chromatin includes some nucleosomes 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 electrophoretic 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 2186 FEBS Journal 278 (2011) ª 2011 The Author Journal compilation ª 2011 FEBS

6 P. N. Cockerill Active chromatin and DNase I hypersensitive sites A Enhancer bp kb RE GM-CSF 2 to 0.6 kb +1.2 to 2.6 kb 5 probe Gene probe Nonstim. 185 bp bp Stim. MNase bp 1847 RE Nonstim. Stim. MNase B GM-CSF locus with the enhancer deleted Non-stimulated Stimulated 5 probe Gene probe 5 probe 180 bp bp Non-stimulated Stimulated Fig. 2. Nucleosome mobilization within the activated GM-CSF locus. Southern blot analysis of oligo-nucleosome fragments produced by increasing amounts of MNase digestion and probed directly with specific GM-CSF locus probes. In this analysis chromatin 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 specific deletion of the 0.7 kb enhancer. of regularly spaced discrete oligo-nucleosome bands for bulk chromatin, but a smeared pattern for active chromatin. An example of the phenomenon is presented 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 positioned at 200 bp intervals at highly specific locations throughout at least 6 kb of the locus (Fig. 3A). However, after gene activation by stimulation of calcium and kinase signalling pathways, nucleosomes throughout this 6 kb region adopt a highly disorganized structure 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 nontranscribed upstream region than within the gene itself (Fig. 2A). This could mean that each cycle of transcription 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 prevents nucleosome translocation [87]. However, the situation may be very different at highly transcribed genes, where closely spaced Pol II molecules can displace 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 neighbourhood 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 downstream nucleosomes about bp further downstream. A similar finding was obtained in studies of the MMTV long terminal repeat where Oct1 and nuclear factor 1 (NF1) were sufficient to direct nucleosome 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 represent 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 nucleosome 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 FEBS Journal 278 (2011) ª 2011 The Author Journal compilation ª 2011 FEBS 2187

7 Active chromatin and DNase I hypersensitive sites P. N. Cockerill A Inducible nucleosome reorganisation across the human GM-CSF locus Strongly positioned Nucleosome GATA + AP-1 NFAT + AP-1 NFAT + AP-1 Enhancer Mast cells stim. GATA Promoter T cells stim. Mast cells non-stim. T cells non-stim. B Nucleosome positions and functional binding sites in the human GM-CSF enhancer Enhancer Mast cells T cells Bgl II GATA GATA AP-1 GATA-2 GATA-2 Sp1 NFAT AP-1 AP-1 NFAT Runx1 GM-CSF N1 N2 N3 N0 N1 N2 N3 717 Bgl II bp Runx1 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-myristate 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 transcription start site. (B) A map showing the positions of regulatory elements required for function in either T cells or mast cells. Shown below are the positions that nucleosomes and GATA-2 occupy in unstimulated T cells and mast cells [85]. 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 transcription factors NFAT and AP-1 which direct the formation 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 complexes which may well account for the observed nucleosome 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 2188 FEBS Journal 278 (2011) ª 2011 The Author Journal compilation ª 2011 FEBS

8 P. N. Cockerill Active chromatin and DNase I hypersensitive sites A Anatomy of the inducible DNaseI hypersensitive site in the GM-CSF enhancer in T cells bp 540 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 positioned 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 composite NFAT AP-1 elements within each nucleosome, and are predicted to support the formation of enhanceosome-like complexes including co-factors such as CBP and SWI SNF [85]. In vivo footprinting confirmed inducible binding of NFAT, AP-1, Sp1 and Runx1 [86,233]. Nuclease digestion studies have determined that the nucleosomes 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. B NFAT + AP-1 DNase I and MNase Nucleosome N1 Condensed chromatin + histone H1 GATA + AP-1 NFAT + AP-1 NFAT + AP-1 Mast T cells RE Sp1-1 Bgl II NFAT Apa I Pst I Bgl II CBP AP-1 SWI/SNF Nucleosome N2 AP-1 CBP NFAT SWI/SNF Runx Runx Promoter Active chromatin with DHS and mobilised nucleosomes with less histone H1 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 significant 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 sufficient 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]. However, 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 unicellular 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 FEBS Journal 278 (2011) ª 2011 The Author Journal compilation ª 2011 FEBS 2189

9 Active chromatin and DNase I hypersensitive sites P. N. Cockerill RNA Ac Ac H3K36 Me2 HDACs Histone hexamer Pol II FACT 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 H2A/H2B dimer Direction of transcription 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 nucleosomes 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 [ ]. 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 significance, 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 compaction, because H3.3-containing nucleosomes appear to be unable to recruit histone H1 [108]. Ac HATs Fig. 5. Model of the chromatin structure in the vicinity of an elongating Pol II complex. Histone acetylation in advance of polymerases is likely to create an open chromatin structure. The advancing polymerase recruits FACT which partially disassembles the nucleosome, allowing Pol II to pass this barrier. Once Pol II has passed, HDACs such as Rpd3S can be recruited via dimethylated or trimethylated H3-K36 and act to return chromatin to the condensed state. 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 mechanisms. 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 context, PARP-1 does not PARylate chromatin, and activation of its enzymatic activity actually relieves silencing [110]. PARP-1 binds to chromatin by engaging 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 promotes decondensation of higher order chromatin structure [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-parylation 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 throughout the Hsp70A locus were rendered MNase sensitive after just 1 or 2 min of heat shock. This extensive disruption 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, minichromosomes assembled in Xenopus oocytes partition into inactive soluble chromatin and insoluble active chromatin [111]. Early attempts to fractionate native chromatin into functionally distinct fractions were performed by digestion of nuclei with MNase followed 2190 FEBS Journal 278 (2011) ª 2011 The Author Journal compilation ª 2011 FEBS

10 P. N. Cockerill Active chromatin and DNase I hypersensitive sites 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 during digestion (fraction S1), (ii) the bulk of the remaining 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 disorganized 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 nucleosomes, 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 accessible 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-component transcription factor and polymerase complexes at sites that have been termed transcription factories [ ]. The residual insoluble fraction is in essence equivalent to the nuclear matrix fraction that was shown to be enriched in active genes [ ]. 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 alongside 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 transcription [54]. Acetylation of lysines leads to neutralization of the positively charged nitrogen atoms that mediate contacts between histone tails and DNA, rendering 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 structure, 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 sedimentation 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]. Subsequent 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 present 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 compacted 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 compensation complex [126] in association with MSL1, MSL2 and MSL3 [128,129]. MSL3 specifically binds to FEBS Journal 278 (2011) ª 2011 The Author Journal compilation ª 2011 FEBS 2191

11 Active chromatin and DNase I hypersensitive sites P. N. Cockerill 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 separate 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 embryoniclethal chromatin condensation [ ]. 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 heterochromatin 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 [ ]. In yeast, this group is made up of NuA4 and Piccolo NuA4 which both utilize Esa1 as the HAT, and Esa1 was found to be essential for H4-K5, K8 and K12 acetylation [137]. In mammalian cells, this group is composed of two distinct 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 responsible for the bulk of the global H4-K5, K8 and K12 acetylation [136]. Each of these HATs exists in complexes that include PHD domains that interact with methylated histone H3-K4 and or K36 [ ]. Transcription directs transient histone acetylation The regulated process of transcription is accompanied by an ordered sequence of transient histone modifications 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 [ ], 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 deacetylation, and transient recruitment of remodellers and transcription factors. A cycle of transcription commences with the recruitment of transcription factors and co-factors bound at the promoter, which modify the local chromatin structure and enable the assembly of the pre-initiation complex. In yeast, transcription factors typically recruit HATs such as SAGA and NuA3, which mainly acetylate 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 promotes the recruitment of histone H3-K4 histone methyltransferases (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 trimethylated H3-K4 [22,149,150], and MOF [149]. This provides a mechanism to both amplify H3-K4 methylation and decondense chromatin by introducing AcH4-K16. H3-K4me3 also recruits the Isw1 chromatin remodelling ATPase to the promoter to prevent premature initiation of transcription elongation [137]. The initiation and elongating phases of the transcription cycle are characterized by distinct sets of histone and Pol II modifications [19,151]. Following promoter clearance, the elongating phase of transcription is associated with phosphorylation of CTD heptapeptide repeat Ser2 by P-TEFb cdk9. This phase of transcription can also be regulated by specific transcription 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 elongation 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 and the bromodomain protein BRD4 which is recruited via AcH4-K16 and phospho H3 [153]. In yeast it is apparent that the histone acetylation associated with transcription elongation is only a very 2192 FEBS Journal 278 (2011) ª 2011 The Author Journal compilation ª 2011 FEBS