Genomic looping: a key principle of chromatin organization.

Review Article J Mol Microbiol Biotechnol 2014;24:344–359 DOI: 10.1159/000368851

Published online: February 17, 2015

Genomic Looping: A Key Principle of Chromatin Organization Ramon A. van der Valk a Jocelyne Vreede b Frédéric Crémazy a Remus T. Dame a a Leiden Institute of Chemistry and Cell Observatory, Leiden University, Leiden, and b Computational Chemistry, van ‘t Hoff Institute for Molecular Sciences, University of Amsterdam, Amsterdam, The Netherlands

Key Words Bacterial chromatin · Archaeal chromatin · Nucleoid · DNA looping · H-NS protein

tion of these diverse, yet difficult-to-study, structures. DNA looping is universal and a conserved mechanism of genome organization throughout all domains of life. © 2015 S. Karger AG, Basel

© 2015 S. Karger AG, Basel 1464–1801/15/0246–0344$39.50/0 E-Mail [email protected] www.karger.com/mmb

Introduction

The effective volume occupied by the genomes of all forms of life far exceeds that of the cells in which they are contained. Therefore, all organisms have developed mechanisms for compactly folding and functionally organizing their genetic material. Eukaryotic Chromatin Eukaryotic cells contain distinct cellular domains called organelles with specific functions in the cell. Perhaps the most complex organelle is the nucleus, containing the genetic material. The genomic DNA in the nucleus of human somatic cells – with a length of more than 6 billion (6 × 109) bp – is compacted three orders of magnitude. To functionally organize DNA, cells express dedicated architectural proteins. The structural organization imposed by these proteins also impacts the expression of the genes, providing an additional mechanism of gene R.T. Dame Leiden Institute of Chemistry and Cell Observatory NL–2333 CC Leiden (The Netherlands) E-Mail rtdame @ chem.leidenuniv.nl

Downloaded by: Siriraj Medical Library, Mahidol University 202.28.191.34 - 3/20/2015 4:24:39 AM

Abstract The effective volume occupied by the genomes of all forms of life far exceeds that of the cells in which they are contained. Therefore, all organisms have developed mechanisms for compactly folding and functionally organizing their genetic material. Through recent advances in fluorescent microscopy and 3C-based technologies, we finally have a first glimpse into the complex mechanisms governing the 3-D folding of genomes. A key feature of genome organization in all domains of life is the formation of DNA loops. Here, we describe the main players in DNA organization with a focus on DNA-bridging proteins. Specifically, we discuss the properties of the bacterial DNA-bridging protein H-NS. Via two different modes of binding to DNA, this protein is a key driver of bacterial genome organization and provides a link between 3-D organization and transcription regulation. Importantly, H-NS function is modulated in response to environmental cues, which are translated into adapted gene expression patterns. We delve into the mechanisms underlying DNA looping and explore the complex and subtle modula-

proven very insightful. Using this approach, the previously inferred role of CTCF in genomic loop formation could be proven directly [Handoko et al., 2011]. Initial observations of large-scale chromosomal organization were made by fluorescence microscopy [Cremer and Cremer, 2010]. With the advent of 3C-based techniques [de Laat and Dekker, 2012], it is now possible to directly probe the in vivo interaction frequencies between segments along the genome (the formation of loops). These techniques have revealed the importance of genomic loop formation on both local scales and global scales in relation to gene expression and genomic organization. On the global scale, eukaryotic genomes are organized into topological domains (topologically associating domains, TADs) [Dixon et al., 2012; Hou et al., 2012; Nora et al., 2012; Sexton et al., 2012]. These domains are characterized by frequent interactions between loci within a domain and the absence of interactions amongst one another [Dixon et al., 2012]. A key role in delineating the boundaries of these domains is attributed to CTCF, which is enriched at the boundaries and essential in the segregation of TADs [Seitan et al., 2013; Sofueva et al., 2013; Zuin et al., 2014]. In addition to dedicated architectural proteins, other factors, such as DNA supercoiling and macromolecular crowding, also play roles in genome organization. The formation of plectonemic supercoils in DNA (wrapping of DNA duplexes around each other) alters its effective volume. Macromolecular crowding due to the large concentration of macromolecules in the cytoplasm has similar effects. Moreover, there is synergy between the architectural proteins, DNA topology and crowding. Many architectural proteins are capable of constraining DNA supercoils as their affinity for DNA is sensitive to both DNA topology and crowding. These factors are beyond the scope of this article; they are however extensively covered in other review articles [Cunha et al., 2001; Dame, 2005; de Vries, 2010; Dorman, 2006; Marenduzzo et al., 2006; Zimmerman and Murphy, 1996].

Genomic Looping

J Mol Microbiol Biotechnol 2014;24:344–359 DOI: 10.1159/000368851

Bacterial Chromatin Despite the absence of a dedicated organelle in prokaryotes, bacteria and archaea also compact and organize their genetic material within a confined part of the cell, the nucleoid. Conventional light and electron microscopy reveal that the nucleoid is distinct from the surrounding cytoplasm because of differences in their refractive index and electron density, respectively. Deconvolution of epifluorescence microscopy images of fluorescently labeled nucleoids in Escherichia coli reveals the existence of ultrastructure: in G1 (with ori and ter regions – where 345


regulation on top of that provided by conventional transcription factors. An illustrious example of a protein involved in shaping the eukaryotic genome is the histone. In the textbook view, hetero-octameric histones interact with 147 bp of DNA along its circumference to form a nucleosome [Luger et al., 1997]. DNA with arrays of nucleosomes (often referred to as ‘beads-on-a-string’ or the 10-nm fiber) is capable of forming a helical structure, likely mediated by nucleosome-nucleosome interactions: the 30-nm fiber. Although this structure readily forms in vitro under idealized conditions on well-defined substrates, its occurrence in vivo is the subject of debate [Fussner et al., 2012]. These nucleosomal fibers are further organized in loops, spanning large distances along the genome. Such loops are either transient due to thermal motion of the nucleosomal fiber or are mediated/stabilized by specific proteins [Hagstrom and Meyer, 2003; Merkenschlager and Odom, 2013; Phillips and Corces, 2009; Thadani et al., 2012]. The formation of loops has a moderate contribution to genome conformation, yet its primarily role is to impose functional organization. The occurrence of transient random contacts is related to the intrinsic flexibility of the fiber: a flexible fiber exhibits higher internal contact frequencies than a stiff fiber. In recent years, evidence has been accumulating for the existence of specific proteins involved in loop formation. In eukaryotic interphase chromosomes, the best-characterized looping proteins are the structural maintenance of chromosomes (SMC) protein cohesin and the CCCTCbinding factor (CTCF). Cohesin mediates intersegmental DNA-DNA contacts (bridges) in cis or in trans by encircling two chromatin fibers. These ring-like proteins play specific roles in regulating genes by connecting regulatory elements (which can be located thousands of base pairs away) with their target genes [Hadjur et al., 2009]. The CTCF protein contains several zinc finger DNAbinding motifs, allowing it to bind several DNA molecules in tandem [Ohlsson et al., 2001], yet no direct evidence has been found for the formation of CTCF-mediated bridges independent of interacting factors. Instead, genome-wide studies reveal binding of cohesin at CTCFbinding sites suggesting that CTCF recruits cohesin (among other proteins) to form DNA loops [Hadjur et al., 2009]. These sites have been implied in the formation of specific genomic loops based on correlation analysis of genome-wide binding patterns and interaction frequencies along the genome [Hadjur et al., 2009]. Although the conventional 3C approaches are unbiased, enriching the cross-linked material using antibodies – biasing the detection of interactions mediated by specific proteins – has

346


cel and Burgi, 1972]. To date, it is unclear which proteins are involved in the formation of such loops in vivo. However, there are several candidate proteins capable of forming intersegmental bridges in vitro: H-NS, FIS and bacterial SMC proteins [Cui et al., 2008; Dame et al., 2006, 2000; Petrushenko et al., 2010; Skoko et al., 2006]. The H-NS and FIS proteins were shown to be involved in domain formation in vivo [Hardy and Cozzarelli, 2005]. Although all these proteins are capable of fulfilling a role in topological domain formation via looping, their involvement has yet to be proven. However, in the case of H-NS the genome-wide distribution of H-NS-bound regions and their spacing quantitatively agrees with estimates of topological domain size [Noom et al., 2007]. The simplest model based on these data would be that the two replichores are each organized in a series of adjacently stacked loops stabilized at their base by binding of H-NS [Noom et al., 2007]. The role of FIS and the SMC proteins in chromosome organization is covered extensively in other recent review articles [Dorman, 2013; Wang et al., 2013a]. At small length scales, architectural proteins shape the local conformation of the bacterial genome. Many of these proteins (HU, IHF, FIS) operate by bending of the DNA duplex [Pan et al., 1996; Swinger and Rice, 2004; van Noort et al., 2004], resulting in an effective reduction of the volume occupied. HU binds nonspecifically, whereas its homologue IHF and the FIS protein recognize specific target sites. The expression levels and DNA-binding affinity of the latter two are sufficiently high [Ali Azam et al., 1999; Ishihama et al., 2014] that binding to other nonspecific sites along the genome is frequent. Although the genome-wide architectural effects of these proteins are not fully understood, the global architectural role of HU in C. crescentus has been elegantly demonstrated in vivo by systematic measurement of physical distance between pairs of fluorescent markers along the genome [Dame et al., 2011b; Hong and McAdams, 2011]. Moreover, in the same organism it was shown that the internal contact frequencies inside CIDs are higher in the presence of HU than in its absence, indicating that HU effectively compacts these domains [Le et al., 2013]. Archaeal Chromatin Archaea phenotypically closely resemble bacteria. Archaea were originally identified in extreme environments such as high temperatures and extreme pH [Pikuta et al., 2007]. However, evidence is accumulating that archaea are more abundant and widespread than confined to these extreme environments. For instance, archaeal organisms have been recently identified in the van der Valk /Vreede /Crémazy /Dame


replication initiates and terminates – localized at midcell) the nucleoid has the shape of a helical ellipsoid [Fisher et al., 2013; Hadizadeh Yazdi et al., 2012]. A similar model was proposed for the Caulobacter crescentus nucleoid based on the genome-wide interaction frequencies obtained from a 3C-based study [Umbarger et al., 2011]. In this organism, the left and right chromosomal arms are extended between the ori and ter regions, which are polarly localized (rather than at mid-cell as in E. coli) and helically folded around one another. The interpretation in terms of overall structure of similar data for the E. coli genome [Cagliero et al., 2013] is far from trivial. In fact, a superficial interpretation might suggest that only the ori and ter regions are structured. However, the E. coli genome is generally present in multiple copies due to the presence of multiple replication forks. Because of this, the structure of the E. coli chromosome is not uniquely defined for a given state of the cell cycle. Moreover, it lacks the longitudinal symmetry of the C. crescentus chromosome in which the ori is fixed at the distant pole. The E. coli chromosome is organized in distinct structural domains (macrodomains) of roughly 1 Mb in size. Two of these domains are localized around the ori and ter regions [Boccard et al., 2005; Espéli and Boccard, 2006; Niki et al., 2000; Valens et al., 2004]. In addition to these two domains, an additional ‘structured’ macrodomain was identified on each chromosomal arm (the left and right macrodomains). Finally, two ‘less-structured’ macrodomains (again one on each arm) were identified [Valens et al., 2004]. It is unclear how the boundaries of macrodomains are established in E. coli. However, it is evident that several (architectural) proteins are specifically restricted to binding in certain macrodomains [Dame et al., 2011a]. A second genome-wide 3C-based study of C. crescentus revealed the existence of discrete domains along the genome (chromosomal interaction domains or CIDs) [Le et al., 2013]. Twenty-three of such domains were identified ranging in length from several tens to several hundreds of kilobase pairs. These domains are conceptually similar to the TADs described in eukaryotes, although the nature of the boundaries appears different [Dixon et al., 2012; Le et al., 2013]. In fact, based on experiments using transcription inhibitors, a key role in maintaining CID boundaries was attributed to transcription of highly active genes [Le et al., 2013]. In terms of size, these domains fall in between macrodomains and the smaller microdomains described in E. coli. Microdomains, in the order of 10 kb in size, have been described as topologically closed domains [Deng et al., 2005; Hardy and Cozzarelli, 2005] established via loop formation [Postow et al., 2004; Wor-

Genomic Looping

Impact of Architectural Proteins

Modulation of Genome Structure Because of their complex nature, it comes as no surprise that prokaryotic genomes are organized dynamically and change as a function of cell cycle or growth conditions [Ali Azam et al., 1999; Ishihama et al., 2014]. In eukaryotes, histone proteins are posttranslationally modified, directly affecting their architectural properties or recruiting machinery promoting chromatin remodeling [Jenuwein and Allis, 2001]. Contrary to common belief, posttranslational modifications are ubiquitous in bacteria [Cain et al., 2014], yet there is no evidence for posttranslational modifications of the proteins involved in chromatin organization. Rather, in E. coli the expression levels of many nucleoid-associated proteins (NAPs; or subunits of heteromeric NAPs) vary according to growth phase and temperature [Ali Azam et al., 1999; Ishihama et al., 2014]. As different NAPs exhibit different architectural properties, their relative and absolute concentrations will have direct effects on genome folding. Moreover, NAPs often exhibit synergistic or antagonistic activities in relation to their roles in transcription regulation [Browning et al., 2000; van Ulsen et al., 1997], which might be extrapolated to scales relevant to global genome organization [Dame, 2005]. In that case, the structural output of the joint action of NAPs is altered if their relative expression levels are changed. Although these types of mechanisms have not been described in archaea, expression levels of different NAPs are certainly dependent on cell cycle and growth conditions [Wurtzel et al., 2010]. Differences in expression ratios contribute to the functionality of the Alba proteins: the Alba1 and Alba 2 proteins are ∼65% homologous, yet differ in a crucial phenylalanine residue at the dimer-dimer interaction interface [Jelinska et al., 2010]. The absence of this residue in Alba2 reduces the cooperativity in binding of Alba1/Alba2 heterodimers to DNA [Jelinska et al., 2005, 2010; Laurens et al., 2012] compared to Alba1 homodimers. In contrast with bacteria, posttranslational modification of NAPs is common in archaea. Indeed – with the exception of histones – the proteins involved in chromatin organization are extensively modified [Eichler and Adams, 2005]. A notable example is the Alba protein in which a lysine group positioned at the DNA-binding interface can be acetylated, leading to reduced DNA-binding affinity [Bell et al., 2002; Wardleworth et al., 2002]. However, in most cases it is not directly obvious how the posttranslational modifications affect protein functionality, as their DNA-binding affinity itself is often not disturbed. J Mol Microbiol Biotechnol 2014;24:344–359 DOI: 10.1159/000368851

347


human intestinal microbiome [Eckburg et al., 2005] and have been implied in various human diseases and disorders [Conway de Macario and Macario, 2009; Eckburg et al., 2003; Wang et al., 2013b]. The potential relevance to health has increased the interest in these relatively simple yet fascinating organisms, exhibiting a curious blend of bacterial and eukaryotic features. Much of the machinery (RNA polymerase, DNA polymerase, etc.) is more closely related to their counterparts in eukaryotes rather than those in bacteria [Reeve, 2003; Werner, 2007], whereas proteins implied in genome organization and transcription regulation are mostly reminiscent of those of bacteria [Driessen and Dame, 2011]. Information on the global organization of archaeal chromatin in vivo is lacking, although many of the proteins likely involved have long been identified [Driessen and Dame, 2011]. In discussing these proteins it is important to distinguish the two main archaeal branches: Euryarchaea and Crenarchaea. Euryarchaea express homologues of the eukaryotic histone H3 and H4 proteins [Sandman et al., 1990]. These proteins assemble on DNA into tetrameric nucleosomes in vitro on SELEX-optimized binding sites [Marc et al., 2002]. In vivo information on the multimeric state of the protein on DNA derived from classical MNase digestion confirms the existence of tetramers [Ammar et al., 2012; Maruyama et al., 2013; Nalabothula et al., 2013], and in addition suggests that the unit building block is a dimer [Maruyama et al., 2013]. This implies that also hexamers, octamers and other multimeric forms are present, setting them apart from eukaryotic histones that are known to form either tetrameric or octameric nucleosomes [Talbert and Henikoff, 2012]. Crenarchaea do not express histones. Instead, they express proteins that are functionally analogous to the HU, IHF and FIS proteins found in bacteria. Proteins, such as Sul7 and Cren7 bind and deform DNA [Baumann et al., 1994; Guo et al., 2008], resulting in compaction [Driessen et al., 2013]. To date, it is unclear whether the archaeal genome is organized into DNA loops in vivo. It is tempting to speculate that it is, as archaeal and bacterial organisms exhibit similar mechanisms of DNA compaction. In addition to SMC proteins, which are conserved throughout all domains of life, most archaea express Alba (acetylation lowers DNA-binding affinity) proteins. This protein was suggested already many years ago to mediate DNA-DNA bridges [Lurz et al., 1986], but only recently was this property appreciated as a mechanism to regulate genome folding [Driessen and Dame, 2011; Driessen et al., submitted; Laurens et al., 2012; Luijsterburg et al., 2008].

DNA Looping by H-NS in Bacteria

H-NS Functions Revealed through Genome-Wide Analysis Several genome-wide H-NS binding studies have been reported for E. coli and Salmonella Typhimurium using ChIP-chip or ChIP-seq analysis [Grainger et al., 2006; Kahramanoglou et al., 2011; Lucchini et al., 2006; Myers et al., 2013; Navarre et al., 2006; Oshima et al., 2006]. The obtained genomic binding profiles for H-NS are remarkably similar considering the wide variety of experimental conditions tested [Kahramanoglou et al., 2011]. Binding of H-NS occurs in about 250–450 discrete regions spread across the genome, covering 15–17% of its length [Grainger et al., 2006; Kahramanoglou et al., 2011; Noom et al., 2007; Oshima et al., 2006]. On average, H-NS-binding regions are 1.6–2 kb in length [Kahramanoglou et al., 2011; Noom et al., 2007] with few extending up to or beyond 6 kb. Considering that H-NS binds DNA as a dimer that occupies a length of ∼25–30 bp, a typical H-NSbound region of 2 kb should contain roughly 60–80 adjacent H-NS dimers. The H-NS-binding motif (discussed later in this review) is highly enriched in H-NS-binding regions, which are characterized by higher than average AT content [Kahramanoglou et al., 2011]. It has been generally accepted that the H-NS expression level does not change significantly during the exponential phase of growth, but decreases significantly at late stationary phase [Azam and Ishihama, 1999]. However, a 348


follow-up study on NAP expression levels in E. coli showed that the number of H-NS molecules per genome copy decreases at the onset of the early-exponential growth phase [Ishihama et al., 2014]. Considering that the expression level of H-NS is in the order of several tens of thousands molecules per cell, most of the H-NS dimers are DNA bound rather than free in the cytoplasm. The number and length of the detected binding regions increase as cells progress from rapid growth to stationary phase [Kahramanoglou et al., 2011; Noom et al., 2007]. This increase is most evident in the ter domain. The number of genome copies decreases when cells enter slow growth. This will increase the pool of free H-NS molecules available for DNA binding, resulting in lateral extension of existing H-NS-bound regions [Vora et al., 2009]. Interestingly, the length of H-NS-binding regions can differ notably between different data sets. This suggests that – in addition to stochastic effects – the spreading of H-NS is modulated by other factors such as environmental conditions or interplay with other NAPs [Bouffartigues et al., 2007; Browning et al., 2000; van Ulsen et al., 1997]. It is important to keep in mind that the length of the H-NS-binding regions can also differ between individual cells, due to extrinsic noise; thus, it is difficult to accurately capture their size and distribution when averaging over a population. Although H-NS-bound patches are fairly equally distributed along the genome, H-NS binding is enriched at macrodomain boundaries. These regions also strongly overlap with transcriptionally silent extended protein occupancy domains that were identified by analyzing the generic protein coverage along the E. coli genome [Vora et al., 2009; Zarei et al., 2013]. These observations suggest a link between the function of H-NS as a transcriptional regulator and the formation of boundaries insulating topological domains as it was proposed for the CIDs described in C. crescentus [Le et al., 2013]. Considering the ability of H-NS to bridge DNA duplexes, the distribution of binding regions across the genome suggests that H-NS might be key to the formation of DNA loops along the genome and for the maintenance of boundaries between topological domains that shape the nucleoid (fig. 1). However, direct evidence of such a genome-wide activity is still missing. In the genome-wide binding profiles, two classes of HNS-bound regions can be distinguished based on their location in relation to coding regions of the potential target genes [Grainger et al., 2006; Kahramanoglou et al., 2011; Oshima et al., 2006; Ueda et al., 2013]. The first class corresponds to H-NS-binding regions localized in the upvan der Valk /Vreede /Crémazy /Dame


Genome Structure Modulates Transcription Many NAPs have been implied in global gene regulation following environmental cues. This results in coordinated up- or downregulation of large numbers of (often functionally related) genes. One well-studied NAP known to regulate several genes based on environmental cues is H-NS. H-NS is an abundant and versatile nucleoid-structuring protein that is conserved among Gram-negative bacteria. Apart from its preeminent role in genome folding, H-NS also functions as a pleiotropic transcription factor that regulates the transcription of about 8% of the E. coli genes, mainly by repressing their expression [Dorman, 2004]. H-NS is a global determinant of bacterial fitness and adaptation to environmental stress in response to stimuli such as pH, osmolarity, temperature or growth phase. Recent advances in genomics, biophysics and structural biology are starting to shed light on the function of H-NS as a sculptor of the 3-D structure of the genome.

B

Fig. 1. Schematic illustration of a bacterial cell. The folded bacterial genome consists of a plethora of DNA loops of various sizes. These can be subdivided into large DNA loops (A) and small DNA loops (B). Loops are stabilized/formed by DNA bridging proteins. Large DNA loops are primarily implied in global DNA organization. Small DNA loops might be specifically involved in the regulation of specific genes. In B, some DNA-bending proteins are also shown to illustrate the interplay between DNA-organizing proteins. DNA-bending proteins help form and stabilize DNA loops by increasing the flexibility of the DNA and facilitating the formation of DNA bridges.

stream regulatory region of genes, whereas the second class covers the coding sequence (sometimes extending across a whole operon). Several mechanisms for H-NSmediated gene regulation have been reported based on in vitro and in vivo experiments on defined target genes, and these in vivo genome-wide binding patterns may be able to corroborate some of these models. Promoter-Specific Binding A quarter of the total number of H-NS-binding regions is located in operon-upstream intergenic regions, which represent only 8% of the genome [Grainger et al., 2006; Kahramanoglou et al., 2011; Oshima et al., 2006]. Kahramanoglou et al. [2011] identified 597 operons, where H-NS associates to their promoter regions. Indeed, 65% of these operons display a significant change in mRNA levels when wild-type and Δhns strains are compared. Generally, these regions are less than 1 kb in size and correlate positively with RNAP-binding signals [Grainger et al., 2006; Oshima et al., 2006]. However, intergenic H-NS-binding regions often extend into a part of the coding sequence. A close inspection of the H-NSGenomic Looping

binding profiles on the promoters of the known H-NS targets proU and bgl reveals the presence of two majors peaks: one in the intergenic region upstream of the promoter and a second extending into the 5′ coding region (fig. 2a, b). These signals correlate to some extent with the position of the regulatory elements that were previously determined for the proU and bgl promoters in in vitro experiments. In both cases, effective repression relies on the interaction between two AT-rich high-affinity binding elements: an upstream repressive element (URE) located in the promoter region and a downstream repressive element (DRE) in the 5′ coding region [Bouffartigues et al., 2007; Dole et al., 2004; Rimsky et al., 2001]. These elements contain high-affinity binding motifs that likely serve as nucleation sites to attract additional H-NS molecules on the adjacent lower-affinity sites through cooperative binding yielding an extended nucleoprotein complex [Bouffartigues et al., 2007; Lang et al., 2007; Liu et al., 2010; Rimsky et al., 2001]. In the case of the proU operon, binding of H-NS on the high-affinity sites and lateral extension depend on temperature and DNA superhelicity [Bouffartigues et al., 2007]. Surprisingly, whereas the first peak detected using ChIP-seq aligns with the position of the URE identified using DNA footprint assays, the second appears to be shifted inward the coding region of proV. The situation is opposite for bgl, with the DRE coinciding almost perfectly with the intragenic peak, whereas the URE falls in a region devoid of H-NS. Both ChIPseq-binding regions appear to be wider than the regions determined in vitro. A possible explanation for this discrepancy is that ChIP-seq yields snapshots of specific protein-DNA interactions as they occur in living cells, at physiological concentrations of H-NS, levels of supercoiling and macromolecular crowding. Moreover, there might be effects of other DNA-binding proteins. Both URE and DRE display a high level of cooperativity, since elimination of the URE strongly impairs repression induced by the DRE [Nagarajavel et al., 2007]. Synergistic repression via these two elements can be envisioned as a looping mechanism involving a cis-interaction between these two elements via H-NS-mediated bridging (fig. 1, box B) [Bouffartigues et al., 2007]. The mechanistic aspects of these binding modes are explained in detail in a later section. The most straightforward consequence of the formation of such a loop is to impede transcription initiation, either by occluding RNAP from the promoter as described for proU and bgl [Nagarajavel et al., 2007], or to trap RNAP at the promoter as shown for the hdeAB and rrnB genes [Dame et al., 2002; Schroder and Wagner, 2000; Shin et al., 2005]. Moreover, these regulatory eleJ Mol Microbiol Biotechnol 2014;24:344–359 DOI: 10.1159/000368851

349


Color version available online

A


a

b

c

d

Fig. 2. H-NS binding signal intensity for different genes regulated by H-NS in E. coli. a, b Genes bound by H-NS in their promoter and 5′ coding sequence illustrated by the proU and bgl locus. c Intragenic H-NS-binding regions extending across multiple genes illustrated by hsdS. d, e Genes regulated by interplay between H-NS, FIS and IHF illustrated by dps and csg. ChIP-seq H-NSbinding peaks taken from Kahramanoglou et al. [2011] are represented, the thick arrows indicate gene positions along the linear chromosome and flags are transcription start sites. Highlighted

rectangles represent H-NS-binding regions identified in vitro [Bouffartigues et al., 2007; Ogasawara et al., 2010]; high-affinity binding sites determined by DNA footprint [Bouffartigues et al., 2007] assays are indicated by a line superimposed on the ChIP-seq binding peaks. Binding sites determined by DNA footprint assays for FIS and IHF are indicated by boxes along the linear chromosome, whereas binding regions detected by ChIP-seq [Kahramanoglou et al., 2011; Prieto et al., 2012] are indicated by bars parallel to the chromosome.

ments might also be (transiently) associated with other distant genomic regions in line with the ability of H-NS to bridge DNA segments in vitro (fig. 1, box A) [Dame et al., 2000]. Such bridging might be the structural basis for the clustering of H-NS repressed genes as observed in vivo [Wang et al., 2011]. Long regions of DNA not bound by H-NS might emanate from such clusters in the form of large genomic loops.

mediate repression during the elongation phase of transcription by acting as a roadblock. However, when the power of a transcribing RNA polymerase is compared with the stability of a region bound and bridged by H-NS in vitro, one would expect that the H-NS roadblock is not sufficiently stable to bring transcription to a halt [Dame et al., 2006]. As lateral filaments are expected to be at most as stable as bridged complexes, they are also not likely to halt transcription. A possible consequence of H-NS binding is that a pause in transcription is induced and that such a pause has a regulatory role [Larson et al., 2014]. A completely new perspective on intragenic binding of H-NS was the finding that these loci often correspond to

Intragenic-Specific Binding A second class of H-NS-binding regions extends across the coding sequence of genes (fig. 2c) [Kahramanoglou et al., 2011; Ueda et al., 2013]. At these sites, H-NS might 350


van der Valk /Vreede /Crémazy /Dame


e

Structural Determinants of H-NS Function The remarkably versatile roles and complex functionality of H-NS find their basis in the tertiary and quaternary structure of the protein. Although a structure of the full-length protein is still lacking, different structural elements of H-NS have been identified and structurally characterized. The H-NS protein consists of three functional domains, an N-terminal dimerization domain (site 1), a second dimerization domain (site 2) and a C-terminal DNA-binding domain (DBD; fig. 3d). The site 1 dimerization domain contains three helices, with helix 3 forming a coiled coil motif. In this motif, hydrophobic side chains, predominantly leucine residues, form the interaction interface. Hydrophobic residues in the two N-terminal helices also contribute to the stability of the dimerization interface. Two conformations have been found for site 1 [Bloch et al., 2003], which differ in the orientation of the helices in the coiled coil, which is parallel [Esposito et al., 2002] or anti-parallel [Bloch et al., 2003]. Both configurations might be functionally relevant in the light of the reported H-NS-mediated response to changes in osmolarity in vivo [Vreede and Dame, 2012]. A study using molecular dynamics simulations showed that the antiparallel configuration is insensitive to changes in ion concentration, whereas the parallel configuration is stable only in the Genomic Looping

presence of high levels of monovalent salt [Vreede and Dame, 2012]. Four out of five structures of the N-terminal part of H-NS exhibit the anti-parallel conformation, while the parallel conformation is observed only once [Ali et al., 2013; Arold et al., 2010; Bloch et al., 2003; Cerdan et al., 2003; Esposito et al., 2002]. The models shown in this review are based on the anti-parallel structure. The second dimerization domain (site 2) contains two helices, which interact in an anti-parallel orientation with the other dimer. The dimerization interface contains both hydrophobic and hydrophilic interactions, as highlighted in figure 3a. For instance, Arg54 of one monomer forms a salt bridge with Glu74 of its dimerization partner. Tyr61 forms hydrophobic contacts with Leu65 and a hydrogen bond with the backbone carbonyl group of Met64. Site 2 has fewer contacts compared to site 1, and therefore it is considered as a secondary dimerization interface, mediating dimer-dimer contacts. For the C-terminal DBD, several structures have been resolved, all in absence of DNA [Gordon et al., 2011]. These structures reveal a globular α/βfold with the conserved QGR motif, responsible for DNA binding, in a loop [Gordon et al., 2011]. In vitro experiments on specific H-NS-regulated regions revealed that H-NS preferentially binds a short 10-bp AT-rich motif: tCGATAAATT [Lang et al., 2007]. This motif might serve as a nucleation site for H-NS binding that facilitates recruitment of additional H-NS molecules to adjacent sites [Bouffartigues et al., 2007]. A shorter version of the motif is indeed observed throughout all H-NS-binding regions detected by ChIP-seq and ChIP-chip experiments [Grainger et al., 2006; Kahramanoglou et al., 2011]. H-NS as a minor groove-binding protein [Gordon et al., 2011] might recognize AT-rich DNA through its narrower minor groove compared to that of GC-rich DNA. The H-NS linker region, connecting site 2 and the DBD (residues 83–96), is unstructured. This region influences the DNA sequence specificity of H-NS [Fernandez-de-Alba et al., 2013]. Theoretical modeling and simulations have shown that the rigidity of the linker region strongly affects the ability of H-NS to form DNA bridges [Joyeux and Vreede, 2013; Wiggins et al., 2009]. Using the structural information currently available, we constructed a model of an H-NS multimer, with dimerization occurring at both site 1 and site 2 (fig. 3a). H-NS exhibits different modes of binding to DNA: lateral filament formation coinciding with DNA stiffening [Amit et al., 2003] and bridge formation between separate DNA segments by binding in trans [Dame et al., 2006]. H-NS has been proposed to bridge DNA segments through the two DBD found in H-NS dimers. A similar argument would hold for larger multimers J Mol Microbiol Biotechnol 2014;24:344–359 DOI: 10.1159/000368851

351


sites of spurious transcription (arising from weak promoters) of noncoding and antisense RNAs or due to inefficient Rho-dependent termination [Peters et al., 2012; Purtov et al., 2014; Saxena and Gowrishankar, 2011; Singh and Grainger, 2013]. In agreement with the proposed effect of pausing on transcript elongation, H-NS contributes to the suppression of antisense transcription by binding close to intragenic Rho-binding sites and facilitating Rho termination [Peters et al., 2012]. An alternative explanation is that H-NS inhibits pervasive transcription in the cell, and that spurious ncRNA synthesis can account for the fitness defect observed in Δhns mutants [Singh et al., 2014]. H-NS binding might preclude low-affinity RNAP recruitment on intragenic regions, enhancing transcription initiation at high-affinity canonical promoters [Singh and Grainger, 2013; Singh et al., 2014]. It is not clear whether intragenic silencing and Rho-specific termination rely on lateral filament formation or also involve DNA bridging. Whereas DNA-DNA bridging is not crucial to many of the mechanisms of repression by H-NS, its intrinsic cooperative nature may enhance the stability of repressive complexes, possibly by engaging in spurious and transient interactions with other genomic segments.


b

a

c

Fig. 3. a Model of E. coli H-NS assembly containing 6 H-NS monomers. To construct a model for a full-length H-NS monomer, we used the structural information available for H-NS, residues 2–83, from S. Typhimurium (PDB code 3NR7 [Arold et al., 2010]) and the NMR structure of the C-terminal domain, residues 91–137 (PDB-code 2L93 [Gordon et al., 2011]). Missing parts of the sequence (residues 1 and 84–90) were modelled as a random coil. The structure of residues 2–83 of S. Typhimurium H-NS also contained information on the dimerization sites. Using these structures as templates, we combined 8 H-NS monomers into a multimer. This structure was relaxed in an energy minimization procedure, employing the conjugate gradient method, with the AMBER03 force field [Duan et al., 2003] using the GROMACS molecular simulation software [Pronk et al., 2013]. Nonbonded interactions were cut off at 1.1 nm, and long-range electrostatics were treated with the particle mesh Ewald method [Darden et al., 1993; Essmann et al., 1995]. Only part of the octamer is displayed, with the dangling monomers cropped from the picture. The struc-

tures of site 1 and site 2 are shown in close-up, with residues involved in dimerization highlighted as stick models. b Models of heterodimers between H-NS and homologous proteins StpA and Hfp. The sequences of H-NS, StpA, Hfp, H-NSB and H-NST were aligned using ClustalW [Larkin et al., 2007]. This sequence alignment, together with the structure of H-NS (residues 2–83, PDB code 3NR7 [Arold et al., 2010]), served as input for a comparative modelling procedure to construct structural models for H-NSStpA and H-NS-Hfp heterodimers using MODELLER version 9.13 [Eswar et al., 2006]. Relevant residues at the dimerization interface are shown as stick models. c The crystal structure of H-NS in complex with Hha (PDB-code 4ICG) [Ali et al., 2013]. d Sequence alignment of H-NS homologues. Highlighted boxes indicate the three functional domains; site 1, site 2, and the DBD. The core residues directly involved in dimerization are highlighted. This alignment was generated with ClustalW2. Residues in italics in the H-NST, Hha, YdgT and Gp5.5 sequences are not aligned.

in which pairs of DBD are present with regular spacing. Based on the notion that long segments of DNA are bridged by large series of adjacent H-NS proteins [Dame et al., 2006] forming bridged regions with a cross-section of a few nanometers [Wiggins et al., 2009], it was thus proposed that such multimeric H-NS assemblies mediate the interactions between two DNA segments [Arold et al.,

2010]. The modes of H-NS binding to DNA might be affected by environmental conditions [Amit et al., 2003; Liu et al., 2010]. Switching between the two binding modes (lateral filament formation and bridging DNA) might be a mechanism to modulate and fine-tune H-NS function at different loci.

352




d

The tertiary and quaternary structure of the protein can provide direct clues as to how such modulation may take place and which functional elements of the protein can be targeted. A direct response to salt conditions can be envisioned in relation to electrostatic interactions. As such, the finding that three positively charged residues at positions 41–43 drive repulsion between the individual subunits within a dimer when these charges are not shielded, could be very relevant [Vreede and Dame, 2012]. Such interactions involve protonatable groups that can also be affected by pH changes, altering the protonation states. It is to date unclear and highly speculative whether such mechanisms operate in vivo. On the contrary, evidence is accumulating that interactions between H-NS and protein partners modulate the functional activity of H-NS [Ali et al., 2011, 2013; Leonard et al., 2009; Muller et al., 2010; Ueda et al., 2013; Williamson and Free, 2005]. Evident targets for H-NS-modulating proteins are the two dimerization sites, where binding of a partner could enhance or reduce the stability of subunit interactions. An additional option is to affect DNA binding by reducing or increasing the effective affinity of the H-NS- protein complex for DNA. Nature indeed provides solutions based on this rationale, with partial or full-length homologues, or distinct proteins targeting either one of two dimerization domains and affecting DNA binding [Ali et al., 2011, 2013; Williamson and Free, 2005]. H-NS Function Is Regulated via Protein Partners The functional properties of H-NS can be modulated via direct interaction with protein partners. These H-NSinteracting proteins can be subdivided into two categories: (1) proteins with full or partial homology to the H-NS amino acid sequence and (2) proteins completely unrelated to H-NS. Most Gram-negative bacteria express at least two variants of the H-NS protein; while pathogenic species can express three or more variants of H-NS [Muller et al., 2010; Williamson and Free, 2005]. These variants can be genomically encoded or carried on a plasmid. In this review, we limit ourselves to discussing the genomically encoded H-NS variants. Full-Length H-NS Homologues StpA, Hfp and H-NSB E. coli expresses at least one H-NS homologue, StpA (suppressor of td mutant phenotype A). This protein has a high degree of sequence homology (58%) with H-NS throughout the three different functional domains [Zhang Genomic Looping

and Belfort, 1992]. This homology allows StpA to form heterodimers with H-NS at both dimerization sites [Johansson et al., 2001]. StpA negatively regulates its own expression, much like H-NS, but StpA and H-NS also repress each other’s expression [Sonden and Uhlin, 1996; Zhang et al., 1996]. H-NS and StpA have similar properties, exhibiting both DNA bridging [Dame et al., 2005] and DNA stiffening [Lim et al., 2012]. Although these properties have not been systematically investigated under identical experimental conditions, there might be subtle differences in DNA-binding behavior: H-NS cannot bridge two DNA duplexes precoated with H-NS [Dame et al., 2006], while StpA is capable of forming bridges even if both duplexes are precoated by StpA [Lim et al., 2012]. Moreover, the structure of StpA-bridged complexes is different from H-NS-bridged complexes, promoting parallel- rather than antiparallel-oriented DNA duplexes in the bridged complex [Dame et al., 2005]. Despite the high level of similarity, StpA expression only partially complements the Δhns phenotype. Comparison of the genome-wide association profiles reveals that the binding regions of StpA and H-NS overlap, and that StpA loses two thirds of its DNA-binding sites in an hns– strain [Uyar et al., 2009]. Moreover, StpA is susceptible to proteolytic cleavage, except when forming heterodimers with H-NS [Johansson et al., 2001]. These observations suggest that StpA is a co-regulator that is functionally dependent on the presence of H-NS. Because of the high sequence homology, yet functional disparities, it is likely that StpA modulates H-NS function by altering the DNA-binding properties of H-NS. Figure 3b shows homology models of H-NS-StpA dimers, modeled on the structures of site 1 and site 2. The main interactions, including the hydrophobic core in site 1 and the hydrophobic and hydrophilic aspects of site 2 are retained in these dimers. These models suggest that StpA and H-NS can form heterodimers (with altered DNA binding properties) via both dimerization sites of H-NS. Such H-NSStpA heterodimers could be next assembled into heteromultimers. The uropathogenic E. coli strains CFT073 and 536 encode a third H-NS-like protein, called H-NSB and Hfp, respectively [Muller et al., 2010; Williamson and Free, 2005]. These proteins have a similar degree of homology with H-NS as StpA (63%), primarily in the N-terminal dimerization domain. In figure 3b, homology models of H-NS-Hfp dimers are shown for both site 1 and site 2, revealing that the nature of the interaction interface is not much different from that of the H-NS homodimer. Indeed, in vitro experiments show that Hfp can form J Mol Microbiol Biotechnol 2014;24:344–359 DOI: 10.1159/000368851

353


Modulation of H-NS Functionality

‘Partial’ Homologues of H-NS: H-NST and Ler Several proteins that share homology with part of H-NS have been identified that modulate H-NS function through interactions with dimerization site 1 or site 2. An example of a site 1 H-NS homologue is the H-NST protein, found in several enteropathogenic E. coli species [Williamson and Free, 2005]. H-NST was found to relieve H-NS-mediated repression of bacterial virulence genes, such as the LEE operon [Levine et al., 2014]. The H-NST protein is believed to interfere with H-NS function by forming H-NS heterodimers via dimerization site 1 (fig. 3). Because H-NST lacks part of dimerization site 2, it could have a dominant negative effect by impairing H-NS multimerization capacities. In addition to modulation of H-NS multimerization, H-NST is also – unexpectedly – itself capable of binding regulatory regions in the LEE operon, and of competing with H-NS to relieve the repression of this operon [Levine et al., 2014]. Another partial H-NS homologue known to modulate H-NS function is the Ler (locus of enterocyte effacementencoded regulator) protein. The largest degree of homology between H-NS and Ler is localized in the C-terminal DBD of the two proteins. Ler, much like H-NS, can form multimers via its N-terminal dimerization domain [Mellies et al., 2011]. Ler is believed to form rigid filaments along the DNA [Bhat et al., 2014; Mellies et al., 2011; Winardhi et al., 2014]. In addition, it may be capable of forming DNA bridges in vitro on specific sequences [Bhat et al., 2014]. Ler cannot form heterodimers with H-NS due to the differences in site 1, where Ler has two polar residues aligned with hydrophobic interfacial residues in H-NS (Thr16, Gln30). Ler also seems to lack part of dimerization site 2. As Ler antagonizes H-NS-mediated repression of genes, it is classified as a transcriptional activator. Both Ler and H-NS bind the LEE operon, and compete to modulate the expression of the LEE genes. Ler has a higher DNA-binding affinity than H-NS, and when ex-

354


pressed Ler effectively competes with H-NS on the LEE operon [Winardhi et al., 2014]. Alternative Modulators of H-NS Function Not all proteins known to functionally interact with H-NS are sequence and/or structural homologues. The Hha protein and its paralogue YdgT were first identified as transcriptional regulators of the virulence factor hemolysin in pathogenic E. coli [Godessart et al., 1988]. Copurification experiments revealed that H-NS and Hha interact [Nieto et al., 2002]. Although initial experiments were unable to resolve the stoichiometry between Hha and H-NS heteromers [Garcia et al., 2005], a recent cocrystal structure of Hha and the N-terminal region of H-NS revealed that Hha and H-NS bind in a 1:1 ratio [Ali et al., 2013]. In this structure two individual Hha molecules bind at the dimerization site 1 in an H-NS dimer [Ali et al., 2013]. Although varying accounts of HhaDNA-binding affinity have been reported [Ali et al., 2013; Nieto et al., 2000], Hha is not considered to be a DNAbinding protein in the absence of H-NS. However, by forming hetero-tetramers Hha might be capable of increasing the DNA-binding affinity of H-NS [Ali et al., 2013]. ChIP-chip analysis revealed that the two proteins share more than 70% of their binding sites on the genome (under conditions of Hha overexpression) [Ueda et al., 2013]. This co-localization primarily occurs in long stretches of H-NS detected within coding regions. Of the 134 genes identified as being affected by either Hha or its paralogue YdgT, 98% are regulated by H-NS [Ueda et al., 2013]. The association of Hha with H-NS enhances its ability to bridge DNA and form loops [van der Valk et al., unpubl. results], suggesting a possible mechanism by which Hha affects gene expression on local as well as global scales. H-NS function can also be modulated via multimerization site 2. No proteins endogenous to E. coli with such functionality have been described, but a protein thought to operate in this manner is the viral T7 gene product 5.5 (Gp5.5) [Ali et al., 2011; Studier, 1981; Zhu et al., 2012]. Interestingly, there is very little homology between H-NS and Gp5.5 for both site 1 and site 2 dimerization domains (fig. 3d). Therefore, it is unlikely that Gp5.5 interacts with H-NS and forms heterodimers. Nevertheless, gp5.5 is capable of interacting with dimerization site 2 using tRNA as a cofactor [Ali et al., 2011; Zhu et al., 2012]. It is not clear to date whether gp5.5 interferes with binding of H-NS along the genome. However, it has become clear that H-NS interferes with T7 DNA replication, and this inhibition is relieved by the interaction with Gp5.5 [Zhu et al., 2012]. van der Valk /Vreede /Crémazy /Dame


heterodimers with H-NS [Muller et al., 2010]. Analogous to H-NS and StpA, Hfp is capable of regulating its own expression and Hfp and H-NS cross-regulate each other’s expression. Hfp expression is growth stage dependent, peaking during the stationary phase [Muller et al., 2010]. Interestingly, StpA and Hfp are expressed under different growth conditions, such that the two proteins are not simultaneously expressed in the cell. This suggests somewhat divergent functionality leading to subtle differences in their H-NS modulatory potential and regulatory role.

Genomic Looping

terplay of NAPs affects genome organization based solely on the current in vivo genomic data. Therefore, it will be important to correlate genome-wide binding profiles of NAPs and gene expression analysis with genome-wide interaction frequencies from 3C technologies. By combining these data with spatial relationships between genomic segments obtained using fluorescence microscopy, this information can be used as constraints for modeling 3-D genome structure and its interplay with gene activity. At the other end of the spectrum, mathematical modeling of polymers incorporating biologically relevant features such as loop formation will provide testable hypotheses that aid in understanding how the architectural properties of NAPs affect the high-order structure of the genome. A crucial issue that needs to be addressed is the fact that information in genomic studies is averaged over the population. In this light, the advent of single- and lowcell sequencing [Shapiro et al., 2013], ChIP [Dahl and Collas, 2009] and chromosome conformation capture [Nagano et al., 2013] paves the way to determining 3-D genome structure in bacteria. The combination of these techniques coupled with structural and mechanistic knowledge acquired for DNA bridging proteins will reveal the sheer scale and complexity of DNA looping found in all domains of life. Acknowledgments Research on the topic of this review is supported by grants from the Netherlands Organization for Scientific Research (864.08.001, Athena 700.58.802), High Tech Systems and Materials Nano NextNL program 8B, the FOM Foundation for Fundamental Research on Matter program ‘Crowd management: The physics of genome processing in complex environments’, and the Human Frontier Science Program (RGP0014/2014). The authors acknowledge Mariliis Tark-Dame for stimulating discussions on genome looping.

References

Ali SS, Beckett E, Bae SJ, Navarre WW: The 5.5 protein of phage T7 inhibits H-NS through interactions with the central oligomerization domain. J Bacteriol 2011;193:4881–4892. Ali SS, Whitney JC, Stevenson J, Robinson H, Howell PL, Navarre WW: Structural insights into the regulation of foreign genes in Salmonella by the Hha/H-NS complex. J Biol Chem 2013;288:13356–13369. Ali Azam T, Iwata A, Nishimura A, Ueda S, Ishihama A: Growth phase-dependent variation in protein composition of the Escherichia coli nucleoid. J Bacteriol 1999;181:6361–6370.


355


Interplay between H-NS and Other NAPs Although the role of NAPs in high-order chromosome folding in vivo still needs to be firmly established, it is likely that the spatial structure of the genome is orchestrated via interplay between these different proteins. Correlating the genome-wide binding profiles available for H-NS and other NAPs can be informative in understanding this interplay. In addition to H-NS, the genomic binding pattern of several NAPs including FIS and IHF has been investigated using ChIP-seq and ChIP-chip [Grainger et al., 2006; Kahramanoglou et al., 2011; Myers et al., 2013; Prieto et al., 2012; Ueda et al., 2013]. Similar to H-NS, these NAPs bind abundantly across the bacterial genome. The binding regions for FIS and IHF are small compared to the binding regions of H-NS (averaging 2,000 bp in size), reflecting the binding of individual functional proteins rather than extended filaments. Strikingly, most of the genes bound by H-NS are also enriched in FIS and IHF [Grainger et al., 2006; Myers et al., 2013]. It is likely that expression of these genes is in part coordinated by competition between different NAPs (antagonistic effects) or by a combination of their effects on the local DNA structure (synergistic effects) as illustrated in figure 1b. In the absence of physical interactions between different types of NAPs (besides interactions between paralogues), both synergistic and antagonistic effects are attributed to structural interplay [Luijsterburg et al., 2008]. Alignment of the genome-wide binding regions of H-NS, FIS and IHF yields correlations not expected based on in vitro analyses. The expression of dps and that of the csg operon are two examples of regulation involving association of NAPs to the promoter region [Altuvia et al., 1994; Grainger et al., 2008; Ogasawara et al., 2010]. Genome-wide binding profiles obtained during early- and mid-exponential growth phase reveal that only H-NS associates to the dps promoter, whereas FIS and IHF bind in the coding sequence and in the 3′ region of the gene, respectively (fig. 2d). This is surprising as in vitro experiments revealed FIS- and IHF-binding sites at the promoter [Altuvia et al., 1994; Grainger et al., 2008]. Similarly, at the csgD gene only H-NS is found on the promoter during exponential growth and in stationary phase in genomewide binding studies (fig. 2e). This is in apparent disagreement with the antagonistic interplay of these proteins observed in previous in vitro experiments [Ogasawara et al., 2010]. In addition to evident differences in conditions between in vitro and in vivo experiments, these apparent discrepancies might be related to stringency parameters in the algorithms used for processing of raw data. It is thus difficult to extrapolate how the in-

356

Cerdan R, Bloch V, Yang Y, Bertin P, Dumas C, Rimsky S, Kochoyan M, Arold ST: Crystal structure of the N-terminal dimerisation domain of VicH, the H-NS-like protein of Vibrio cholerae. J Mol Biol 2003;334:179–185. Conway de Macario E, Macario AJ: Methanogenic archaea in health and disease: a novel paradigm of microbial pathogenesis. Int J Med Microbiol 2009;299:99–108. Cremer T, Cremer M: Chromosome territories. Cold Spring Harbor Perspect Biol 2010; 2:a003889. Cui Y, Petrushenko ZM, Rybenkov VV: MukB acts as a macromolecular clamp in DNA condensation. Nat Struct Mol Biol 2008; 15: 411– 418. Cunha S, Woldringh CL, Odijk T: Polymer-mediated compaction and internal dynamics of isolated Escherichia coli nucleoids. J Struct Biol 2001;136:53–66. Dahl JA, Collas P: Microchip: chromatin immunoprecipitation for small cell numbers. Methods Mol Biol 2009;567:59–74. Dame RT: The role of nucleoid-associated proteins in the organization and compaction of bacterial chromatin. Mol Microbiol 2005; 56: 858–870. Dame RT, Kalmykowa OJ, Grainger DC: Chromosomal macrodomains and associated proteins: implications for DNA organization and replication in Gram negative bacteria. PLoS Genet 2011a;7:e1002123. Dame RT, Luijsterburg MS, Krin E, Bertin PN, Wagner R, Wuite GJ: DNA bridging: a property shared among H-NS-like proteins. J Bacteriol 2005;187:1845–1848. Dame RT, Noom MC, Wuite GJ: Bacterial chromatin organization by H-NS protein unravelled using dual DNA manipulation. Nature 2006;444:387–390. Dame RT, Tark-Dame M, Schiessel H: A physical approach to segregation and folding of the Caulobacter crescentus genome. Mol Microbiol 2011b;82:1311–1315. Dame RT, Wyman C, Goosen N: H-NS mediated compaction of DNA visualised by atomic force microscopy. Nucleic Acids Res 2000;28: 3504–3510. Dame RT, Wyman C, Wurm R, Wagner R, Goosen N: Structural basis for H-NS-mediated trapping of RNA polymerase in the open initiation complex at the rrnB P1. J Biol Chem 2002;277:2146–2150. Darden T, York D, Pedersen L: Particle mesh Ewald – an N.log(N) method for Ewald sums in large systems. J Chem Phys 1993;98:10089– 10092. de Laat W, Dekker J: 3C-based technologies to study the shape of the genome. Methods 2012; 58:189–191. Deng S, Stein RA, Higgins NP: Organization of supercoil domains and their reorganization by transcription. Mol Microbiol 2005; 57: 1511–1521.


de Vries R: DNA condensation in bacteria: interplay between macromolecular crowding and nucleoid proteins. Biochimie 2010; 92: 1715– 1721. Dixon JR, Selvaraj S, Yue F, Kim A, Li Y, Shen Y, Hu M, Liu JS, Ren B: Topological domains in mammalian genomes identified by analysis of chromatin interactions. Nature 2012; 485: 376–380. Dole S, Nagarajavel V, Schnetz K: The histonelike nucleoid structuring protein H-NS represses the Escherichia coli bgl operon downstream of the promoter. Mol Microbiol 2004; 52:589–600. Dorman CJ: H-NS: a universal regulator for a dynamic genome. Nat Rev Microbiol 2004; 2: 391–400. Dorman CJ: DNA supercoiling and bacterial gene expression. Sci Prog 2006;89:151–166. Dorman CJ: Genome architecture and global gene regulation in bacteria: making progress towards a unified model? Nat Rev Microbiol 2013;11:349–355. Driessen RP, Dame RT: Nucleoid-associated proteins in Crenarchaea. Biochem Soc Transact 2011;39:116–121. Driessen RP, Meng H, Suresh G, Shahapure R, Lanzani G, Priyakumar UD, White MF, Schiessel H, van Noort J, Dame RT: Crenarchaeal chromatin proteins Cren7 and Sul7 compact DNA by inducing rigid bends. Nucleic Acids Res 2013;41:196–205. Driessen RPC, Peeters E, Werner F, Dame RT: Global gene regulation and genome organization in archaea. Nat Rev Microbiol, submitted. Duan Y, Wu C, Chowdhury S, Lee MC, Xiong GM, Zhang W, Yang R, Cieplak P, Luo R, Lee T, Caldwell J, Wang JM, Kollman P: A pointcharge force field for molecular mechanics simulations of proteins based on condensedphase quantum mechanical calculations. J Comput Chem 2003;24:1999–2012. Eckburg PB, Bik EM, Bernstein CN, Purdom E, Dethlefsen L, Sargent M, Gill SR, Nelson KE, Relman DA: Diversity of the human intestinal microbial flora. Science 2005;308:1635–1638. Eckburg PB, Lepp PW, Relman DA: Archaea and their potential role in human disease. Infect Immun 2003;71:591–596. Eichler J, Adams MW: Posttranslational protein modification in archaea. Microbiol Mol Biol Rev 2005;69:393–425. Espéli O, Boccard F: Organization of the Escherichia coli chromosome into macrodomains and its possible functional implications. J Struct Biol 2006;156:304–310. Esposito D, Petrovic A, Harris R, Ono S, Eccleston JF, Mbabaali A, Haq I, Higgins CF, Hinton JC, Driscoll PC, Ladbury JE: H-NS oligomerization domain structure reveals the mechanism for high order self-association of the intact protein. J Mol Biol 2002;324:841–850. Essmann U, Perera L, Berkowitz ML, Darden T, Lee H, Pedersen LG: A smooth particle mesh Ewald method. J Chem Phys 1995;103:8577– 8593.



Altuvia S, Almiron M, Huisman G, Kolter R, Storz G: The dps promoter is activated by OxyR during growth and by IHF and sigma S in stationary phase. Mol Microbiol 1994; 13: 265– 272. Amit R, Oppenheim AB, Stavans J: Increased bending rigidity of single DNA molecules by H-NS, a temperature and osmolarity sensor. Biophys J 2003;84:2467–2473. Ammar R, Torti D, Tsui K, Gebbia M, Durbic T, Bader GD, Giaever G, Nislow C: Chromatin is an ancient innovation conserved between Archaea and Eukarya. eLife 2012;1:e00078. Arold ST, Leonard PG, Parkinson GN, Ladbury JE: H-NS forms a superhelical protein scaffold for DNA condensation. Proc Natl Acad Sci USA 2010;107:15728–15732. Azam TA, Ishihama A: Twelve species of the nucleoid-associated protein from Escherichia coli. Sequence recognition specificity and DNA binding affinity. J Biol Chem 1999;274: 33105–33113. Baumann H, Knapp S, Lundback T, Ladenstein R, Hard T: Solution structure and DNA-binding properties of a thermostable protein from the archaeon Sulfolobus solfataricus. Nat Struct Biol 1994;1:808–819. Bell SD, Botting CH, Wardleworth BN, Jackson SP, White MF: The interaction of Alba, a conserved archaeal chromatin protein, with Sir2 and its regulation by acetylation. Science 2002;296:148–151. Bhat A, Shin M, Jeong JH, Kim HJ, Lim HJ, Rhee JH, Paik SY, Takeyasu K, Tobe T, Yen H, Lee G, Choy HE: DNA looping-dependent autorepression of LEE1 P1 promoters by Ler in enteropathogenic Escherichia coli (EPEC). Proc Natl Acad Sci USA 2014;111:E2586–E2595. Bloch V, Yang Y, Margeat E, Chavanieu A, Auge MT, Robert B, Arold S, Rimsky S, Kochoyan M: The H-NS dimerization domain defines a new fold contributing to DNA recognition. Nat Struct Biol 2003;10:212–218. Boccard F, Esnault E, Valens M: Spatial arrangement and macrodomain organization of bacterial chromosomes. Mol Microbiol 2005; 57: 9–16. Bouffartigues E, Buckle M, Badaut C, Travers A, Rimsky S: H-NS cooperative binding to highaffinity sites in a regulatory element results in transcriptional silencing. Nat Struct Mol Biol 2007;14:441–448. Browning DF, Cole JA, Busby SJ: Suppression of FNR-dependent transcription activation at the Escherichia coli nir promoter by Fis, IHF and H-NS: modulation of transcription initiation by a complex nucleo-protein assembly. Mol Microbiol 2000;37:1258–1269. Cagliero C, Grand RS, Jones MB, Jin DJ, O’Sullivan JM: Genome conformation capture reveals that the Escherichia coli chromosome is organized by replication and transcription. Nucleic Acids Res 2013;41:6058–6071. Cain JA, Solis N, Cordwell SJ: Beyond gene expression: the impact of protein post-translational modifications in bacteria. J Proteomics 2014;97:265–286.

Genomic Looping

Handoko L, Xu H, Li G, Ngan CY, Chew E, Schnapp M, Lee CW, Ye C, Ping JL, Mulawadi F, Wong E, Sheng J, Zhang Y, Poh T, Chan CS, Kunarso G, Shahab A, Bourque G, Cacheux-Rataboul V, Sung WK, Ruan Y, Wei CL: CTCF-mediated functional chromatin interactome in pluripotent cells. Nat Genet 2011;43:630–638. Hardy CD, Cozzarelli NR: A genetic selection for supercoiling mutants of Escherichia coli reveals proteins implicated in chromosome structure. Mol Microbiol 2005;57:1636–1652. Hong SH, McAdams HH: Compaction and transport properties of newly replicated Caulobacter crescentus DNA. Mol Microbiol 2011; 82:1349–1358. Hou C, Li L, Qin ZS, Corces VG: Gene density, transcription, and insulators contribute to the partition of the Drosophila genome into physical domains. Mol Cell 2012;48:471–484. Ishihama A, Kori A, Koshio E, Yamada K, Maeda H, Shimada T, Makinoshima H, Iwata A, Fujita N: Intracellular concentrations of transcription factors in Escherichia coli: 65 species with known regulatory functions. J Bacteriol 2014;196:2718–2727. Jelinska C, Conroy MJ, Craven CJ, Hounslow AM, Bullough PA, Waltho JP, Taylor GL, White MF: Obligate heterodimerization of the archaeal Alba2 protein with Alba1 provides a mechanism for control of DNA packaging. Structure 2005;13:963–971. Jelinska C, Petrovic-Stojanovska B, Ingledew WJ, White MF: Dimer-dimer stacking interactions are important for nucleic acid binding by the archaeal chromatin protein Alba. Biochem J 2010;427:49–55. Jenuwein T, Allis CD: Translating the histone code. Science 2001;293:1074–1080. Johansson J, Eriksson S, Sonden B, Wai SN, Uhlin BE: Heteromeric interactions among nucleoid-associated bacterial proteins: localization of StpA-stabilizing regions in H-NS of Escherichia coli. J Bacteriol 2001;183:2343–2347. Joyeux M, Vreede J: A model of H-NS mediated compaction of bacterial DNA. Biophys J 2013; 104:1615–1622. Kahramanoglou C, Seshasayee AS, Prieto AI, Ibberson D, Schmidt S, Zimmermann J, Benes V, Fraser GM, Luscombe NM: Direct and indirect effects of H-NS and Fis on global gene expression control in Escherichia coli. Nucleic Acids Res 2011;39:2073–2091. Lang B, Blot N, Bouffartigues E, Buckle M, Geertz M, Gualerzi CO, Mavathur R, Muskhelishvili G, Pon CL, Rimsky S, Stella S, Babu MM, Travers A: High-affinity DNA binding sites for H-NS provide a molecular basis for selective silencing within proteobacterial genomes. Nucleic Acids Res 2007; 35: 6330– 6337. Larkin MA, Blackshields G, Brown NP, Chenna R, McGettigan PA, McWilliam H, Valentin F, Wallace IM, Wilm A, Lopez R, Thompson JD, Gibson TJ, Higgins DG: Clustal W and clustal X version 2.0. Bioinformatics 2007; 23: 2947– 2948.

Larson MH, Mooney RA, Peters JM, Windgassen T, Nayak D, Gross CA, Block SM, Greenleaf WJ, Landick R, Weissman JS: A pause sequence enriched at translation start sites drives transcription dynamics in vivo. Science 2014;344:1042–1047. Laurens N, Driessen RP, Heller I, Vorselen D, Noom MC, Hol FJ, White MF, Dame RT, Wuite GJ: Alba shapes the archaeal genome using a delicate balance of bridging and stiffening the DNA. Nat Commun 2012;3:1328. Le TB, Imakaev MV, Mirny LA, Laub MT: Highresolution mapping of the spatial organization of a bacterial chromosome. Science 2013; 342:731–734. Leonard PG, Ono S, Gor J, Perkins SJ, Ladbury JE: Investigation of the self-association and hetero-association interactions of H-NS and StpA from Enterobacteria. Mol Microbiol 2009;73:165–179. Levine JA, Hansen AM, Michalski JM, Hazen TH, Rasko DA, Kaper JB: H-NST induces LEE expression and the formation of attaching and effacing lesions in enterohemorrhagic Escherichia coli. PloS One 2014;9:e86618. Lim CJ, Whang YR, Kenney LJ, Yan J: Gene silencing H-NS paralogue StpA forms a rigid protein filament along DNA that blocks DNA accessibility. Nucleic Acids Res 2012; 40: 3316–3328. Liu Y, Chen H, Kenney LJ, Yan J: A divalent switch drives H-NS/DNA-binding conformations between stiffening and bridging modes. Genes Dev 2010;24:339–344. Lucchini S, Rowley G, Goldberg MD, Hurd D, Harrison M, Hinton JC: H-NS mediates the silencing of laterally acquired genes in bacteria. PLoS Pathogens 2006;2:e81. Luger K, Mader AW, Richmond RK, Sargent DF, Richmond TJ: Crystal structure of the nucleosome core particle at 2.8 Å resolution. Nature 1997;389:251–260. Luijsterburg MS, White MF, van Driel R, Dame RT: The major architects of chromatin: architectural proteins in bacteria, archaea and eukaryotes. Crit Rev Biochem Mol Biol 2008;43: 393–418. Lurz R, Grote M, Dijk J, Reinhardt R, Dobrinski B: Electron microscopic study of DNA complexes with proteins from the Archaebacterium Sulfolobus acidocaldarius. EMBO J 1986; 5:3715–3721. Marc F, Sandman K, Lurz R, Reeve JN: Archaeal histone tetramerization determines DNA affinity and the direction of DNA supercoiling. J Biol Chem 2002;277:30879–30886. Marenduzzo D, Finan K, Cook PR: The depletion attraction: an underappreciated force driving cellular organization. J Cell Biol 2006; 175: 681–686. Maruyama H, Harwood JC, Moore KM, Paszkiewicz K, Durley SC, Fukushima H, Atomi H, Takeyasu K, Kent NA: An alternative beads-ona-string chromatin architecture in Thermococcus kodakarensis. EMBO Rep 2013;14:711–717.


357


Eswar N, Webb B, Marti-Renom MA, Madhusudhan MS, Eramian D, Shen MY, Pieper U, Sali A: Comparative protein structure modeling using MODELLER. Curr Protoc Protein Sci 2007;chapter 2:unit 2.9. Fernandez-de-Alba C, Berrow NS, Garcia-Castellanos R, Garcia J, Pons M: On the origin of the selectivity of plasmidic H-NS towards horizontally acquired DNA: linking H-NS oligomerization and cooperative DNA binding. J Mol Biol 2013;425:2347–2358. Fisher JK, Bourniquel A, Witz G, Weiner B, Prentiss M, Kleckner N: Four-dimensional imaging of E. coli nucleoid organization and dynamics in living cells. Cell 2013;153:882–895. Fussner E, Strauss M, Djuric U, Li R, Ahmed K, Hart M, Ellis J, Bazett-Jones DP: Open and closed domains in the mouse genome are configured as 10-nm chromatin fibres. EMBO Rep 2012;13:992–996. Garcia J, Cordeiro TN, Nieto JM, Pons I, Juarez A, Pons M: Interaction between the bacterial nucleoid associated proteins HHA and H-NS involves a conformational change of HHA. Biochem J 2005;388:755–762. Godessart N, Munoa FJ, Regue M, Juarez A: Chromosomal mutations that increase the production of a plasmid-encoded haemolysin in Escherichia coli. J Gen Microbiol 1988;134: 2779–2787. Gordon BR, Li Y, Cote A, Weirauch MT, Ding P, Hughes TR, Navarre WW, Xia B, Liu J: Structural basis for recognition of AT-rich DNA by unrelated xenogeneic silencing proteins. Proc Natl Acad Sci USA 2011;108:10690–10695. Grainger DC, Goldberg MD, Lee DJ, Busby SJ: Selective repression by Fis and H-NS at the Escherichia coli dps promoter. Mol Microbiol 2008;68:1366–1377. Grainger DC, Hurd D, Goldberg MD, Busby SJ: Association of nucleoid proteins with coding and non-coding segments of the Escherichia coli genome. Nucleic Acids Res 2006; 34: 4642–4652. Guo L, Feng Y, Zhang Z, Yao H, Luo Y, Wang J, Huang L: Biochemical and structural characterization of Cren7, a novel chromatin protein conserved among Crenarchaea. Nucleic Acids Res 2008;36:1129–1137. Hadizadeh Yazdi N, Guet CC, Johnson RC, Marko JF: Variation of the folding and dynamics of the Escherichia coli chromosome with growth conditions. Mol Microbiol 2012; 86: 1318–1333. Hadjur S, Williams LM, Ryan NK, Cobb BS, Sexton T, Fraser P, Fisher AG, Merkenschlager M: Cohesins form chromosomal cis-interactions at the developmentally regulated IFNG locus. Nature 2009;460:410–413. Hagstrom KA, Meyer BJ: Condensin and cohesin: more than chromosome compactor and glue. Nat Rev Genet 2003;4:520–534.

358

Ohlsson R, Renkawitz R, Lobanenkov V: CTCF is a uniquely versatile transcription regulator linked to epigenetics and disease. Trends Genet 2001;17:520–527. Oshima T, Ishikawa S, Kurokawa K, Aiba H, Ogasawara N: Escherichia coli histone-like protein H-NS preferentially binds to horizontally acquired DNA in association with RNA polymerase. DNA Res 2006;13:141–153. Pan CQ, Finkel SE, Cramton SE, Feng JA, Sigman DS, Johnson RC: Variable structures of FisDNA complexes determined by flanking DNA-protein contacts. J Mol Biol 1996; 264: 675–695. Peters JM, Mooney RA, Grass JA, Jessen ED, Tran F, Landick R: Rho and Nusg suppress pervasive antisense transcription in Escherichia coli. Genes Dev 2012;26:2621–2633. Petrushenko ZM, Cui Y, She W, Rybenkov VV: Mechanics of DNA bridging by bacterial condensin MukBEF in vitro and in singulo. EMBO J 2010;29:1126–1135. Phillips JE, Corces VG: CTCF: master weaver of the genome. Cell 2009;137:1194–1211. Pikuta EV, Hoover RB, Tang J: Microbial extremophiles at the limits of life. Crit Rev Microbiol 2007;33:183–209. Postow L, Hardy CD, Arsuaga J, Cozzarelli NR: Topological domain structure of the Escherichia coli chromosome. Genes Dev 2004; 18: 1766–1779. Prieto AI, Kahramanoglou C, Ali RM, Fraser GM, Seshasayee AS, Luscombe NM: Genomic analysis of DNA binding and gene regulation by homologous nucleoid-associated proteins IHF and HU in Escherichia coli K12. Nucleic Acids Res 2012;40:3524–3537. Pronk S, Pall S, Schulz R, Larsson P, Bjelkmar P, Apostolov R, Shirts MR, Smith JC, Kasson PM, van der Spoel D, Hess B, Lindahl E: Gromacs 4.5: a high-throughput and highly parallel open source molecular simulation toolkit. Bioinformatics 2013;29:845–854. Purtov YA, Glazunova OA, Antipov SS, Pokusaeva VO, Fesenko EE, Preobrazhenskaya EV, Shavkunov KS, Tutukina MN, Lukyanov VI, Ozoline ON: Promoter islands as a platform for interaction with nucleoid proteins and transcription factors. J Bioinform Comput Biol 2014;12:1441006. Reeve JN: Archaeal chromatin and transcription. Mol Microbiol 2003;48:587–598. Rimsky S, Zuber F, Buckle M, Buc H: A molecular mechanism for the repression of transcription by the H-NS protein. Mol Microbiol 2001;42:1311–1323. Sandman K, Krzycki JA, Dobrinski B, Lurz R, Reeve JN: HMF, a DNA-binding protein isolated from the hyperthermophilic archaeon methanothermus fervidus, is most closely related to histones. Proc Natl Acad Sci USA 1990;87:5788–5791. Saxena S, Gowrishankar J: Modulation of Rhodependent transcription termination in Escherichia coli by the H-NS family of proteins. J Bacteriol 2011;193:3832–3841.


Schroder O, Wagner R: The bacterial DNA-binding protein H-NS represses ribosomal RNA transcription by trapping RNA polymerase in the initiation complex. J Mol Biol 2000; 298: 737–748. Seitan VC, Faure AJ, Zhan Y, McCord RP, Lajoie BR, Ing-Simmons E, Lenhard B, Giorgetti L, Heard E, Fisher AG, Flicek P, Dekker J, Merkenschlager M: Cohesin-based chromatin interactions enable regulated gene expression within preexisting architectural compartments. Genome Res 2013;23:2066–2077. Sexton T, Yaffe E, Kenigsberg E, Bantignies F, Leblanc B, Hoichman M, Parrinello H, Tanay A, Cavalli G: Three-dimensional folding and functional organization principles of the Drosophila genome. Cell 2012;148:458–472. Shapiro E, Biezuner T, Linnarsson S: Single-cell sequencing-based technologies will revolutionize whole-organism science. Nat Rev Genet 2013;14:618–630. Shin M, Song M, Rhee JH, Hong Y, Kim YJ, Seok YJ, Ha KS, Jung SH, Choy HE: DNA loopingmediated repression by histone-like protein H-NS: specific requirement of Eσ70 as a cofactor for looping. Genes Dev 2005; 19: 2388– 2398. Singh SS, Grainger DC: H-NS can facilitate specific DNA-binding by RNA polymerase in AT-rich gene regulatory regions. PLoS Genet 2013;9:e1003589. Singh SS, Singh N, Bonocora RP, Fitzgerald DM, Wade JT, Grainger DC: Widespread suppression of intragenic transcription initiation by H-NS. Genes Dev 2014;28:214–219. Skoko D, Yoo D, Bai H, Schnurr B, Yan J, McLeod SM, Marko JF, Johnson RC: Mechanism of chromosome compaction and looping by the Escherichia coli nucleoid protein Fis. J Mol Biol 2006;364:777–798. Sofueva S, Yaffe E, Chan WC, Georgopoulou D, Vietri Rudan M, Mira-Bontenbal H, Pollard SM, Schroth GP, Tanay A, Hadjur S: Cohesinmediated interactions organize chromosomal domain architecture. EMBO J 2013;32:3119– 3129. Sonden B, Uhlin BE: Coordinated and differential expression of histone-like proteins in Escherichia coli: regulation and function of the H-NS analog StpA. EMBO J 1996; 15: 4970– 4980. Studier FW: Identification and mapping of five new genes in bacteriophage T7. J Mol Biol 1981;153:493–502. Swinger KK, Rice PA: IHF and HU: flexible architects of bent DNA. Curr Opin Struct Biol 2004;14:28–35. Talbert PB, Henikoff S: Chromatin: packaging without nucleosomes. Curr Biol 2012; 22:R1040–R1043. Thadani R, Uhlmann F, Heeger S: Condensin, chromatin crossbarring and chromosome condensation. Curr Biol 2012; 22:R1012– R1021.



Mellies JL, Benison G, McNitt W, Mavor D, Boniface C, Larabee FJ: Ler of pathogenic Escherichia coli forms toroidal protein-DNA complexes. Microbiology 2011;157:1123–1133. Merkenschlager M, Odom DT: CTCF and cohesin: linking gene regulatory elements with their targets. Cell 2013;152:1285–1297. Muller CM, Schneider G, Dobrindt U, Emody L, Hacker J, Uhlin BE: Differential effects and interactions of endogenous and horizontally acquired H-NS-like proteins in pathogenic Escherichia coli. Mol Microbiol 2010;75:280–293. Myers KS, Yan H, Ong IM, Chung D, Liang K, Tran F, Keles S, Landick R, Kiley PJ: Genomescale analysis of Escherichia coli FNR reveals complex features of transcription factor binding. PLoS Genet 2013;9:e1003565. Nagano T, Lubling Y, Stevens TJ, Schoenfelder S, Yaffe E, Dean W, Laue ED, Tanay A, Fraser P: Single-cell Hi-C reveals cell-to-cell variability in chromosome structure. Nature 2013;502: 59–64. Nagarajavel V, Madhusudan S, Dole S, Rahmouni AR, Schnetz K: Repression by binding of H-NS within the transcription unit. J Biol Chem 2007;282:23622–23630. Nalabothula N, Xi L, Bhattacharyya S, Widom J, Wang JP, Reeve JN, Santangelo TJ, FondufeMittendorf YN: Archaeal nucleosome positioning in vivo and in vitro is directed by primary sequence motifs. BMC Genomics 2013; 14:391. Navarre WW, Porwollik S, Wang Y, McClelland M, Rosen H, Libby SJ, Fang FC: Selective silencing of foreign DNA with low GC content by the H-NS protein in Salmonella. Science 2006;313:236–238. Nieto JM, Madrid C, Miquelay E, Parra JL, Rodriguez S, Juarez A: Evidence for direct proteinprotein interaction between members of the enterobacterial Hha/YmoA and H-NS families of proteins. J Bacteriol 2002;184:629–635. Nieto JM, Madrid C, Prenafeta A, Miquelay E, Balsalobre C, Carrascal M, Juarez A: Expression of the hemolysin operon in Escherichia coli is modulated by a nucleoid-protein complex that includes the proteins Hha and H-NS. Mol Gen Genet 2000;263:349–358. Niki H, Yamaichi Y, Hiraga S: Dynamic organization of chromosomal DNA in Escherichia coli. Genes Dev 2000;14:212–223. Noom MC, Navarre WW, Oshima T, Wuite GJ, Dame RT: H-NS promotes looped domain formation in the bacterial chromosome. Curr Biol 2007;17:R913–R914. Nora EP, Lajoie BR, Schulz EG, Giorgetti L, Okamoto I, Servant N, Piolot T, van Berkum NL, Meisig J, Sedat J, Gribnau J, Barillot E, Bluthgen N, Dekker J, Heard E: Spatial partitioning of the regulatory landscape of the X-inactivation centre. Nature 2012;485:381–385. Ogasawara H, Yamada K, Kori A, Yamamoto K, Ishihama A: Regulation of the Escherichia coli csgD promoter: interplay between five transcription factors. Microbiology 2010; 156: 2470–2483.

Genomic Looping

Vreede J, Dame RT: Predicting the effect of ions on the conformation of the H-NS dimerization domain. Biophys J 2012;103:89–98. Wang W, Li GW, Chen C, Xie XS, Zhuang X: Chromosome organization by a nucleoid-associated protein in live bacteria. Science 2011; 333:1445–1449. Wang X, Montero Llopis P, Rudner DZ: Organization and segregation of bacterial chromosomes. Nat Rev Genet 2013a;14:191–203. Wang J, Qi J, Zhao H, He S, Zhang Y, Wei S, Zhao F: Metagenomic sequencing reveals microbiota and its functional potential associated with periodontal disease. Sci Rep 2013b;3: 1843. Wardleworth BN, Russell RJ, Bell SD, Taylor GL, White MF: Structure of Alba: an archaeal chromatin protein modulated by acetylation. EMBO J 2002;21:4654–4662. Werner F: Structure and function of archaeal RNA polymerases. Mol Microbiol 2007; 65: 1395–1404. Wiggins PA, Dame RT, Noom MC, Wuite GJ: Protein-mediated molecular bridging: a key mechanism in biopolymer organization. Biophys J 2009;97:1997–2003. Williamson HS, Free A: A truncated H-NS-like protein from enteropathogenic Escherichia coli acts as an H-NS antagonist. Mol Microbiol 2005;55:808–827. Winardhi RS, Gulvady R, Mellies JL, Yan J: Locus of enterocyte effacement-encoded regulator (Ler) of pathogenic Escherichia coli competes off nucleoid-structuring protein (H-NS) through noncooperative DNA binding. J Biol Chem 2014;289:13739–13750. Worcel A, Burgi E: On the structure of the folded chromosome of Escherichia coli. J Mol Biol 1972;71:127–147.

Wurtzel O, Sapra R, Chen F, Zhu Y, Simmons BA, Sorek R: A single-base resolution map of an archaeal transcriptome. Genome Res 2010;20: 133–141. Zarei M, Sclavi B, Cosentino Lagomarsino M: Gene silencing and large-scale domain structure of the E. coli genome. Mol Biosyst 2013; 9:758–767. Zhang A, Belfort M: Nucleotide sequence of a newly-identified Escherichia coli gene, StpA, encoding an H-NS-like protein. Nucleic Acids Res 1992;20:6735. Zhang A, Rimsky S, Reaban ME, Buc H, Belfort M: Escherichia coli protein analogs StpA and H-NS: regulatory loops, similar and disparate effects on nucleic acid dynamics. EMBO J 1996;15:1340–1349. Zhu B, Lee SJ, Tan M, Wang ED, Richardson CC: Gene 5.5 protein of bacteriophage T7 in complex with Escherichia coli nucleoid protein H-NS and transfer RNA masks transfer RNA priming in T7 DNA replication. Proc Natl Acad Sci USA 2012;109:8050–8055. Zimmerman SB, Murphy LD: Macromolecular crowding and the mandatory condensation of DNA in bacteria. FEBS Lett 1996; 390: 245– 248. Zuin J, Dixon JR, van der Reijden MI, Ye Z, Kolovos P, Brouwer RW, van de Corput MP, van de Werken HJ, Knoch TA, van Ijcken WF, Grosveld FG, Ren B, Wendt KS: Cohesin and CTCF differentially affect chromatin architecture and gene expression in human cells. Proc Natl Acad Sci USA 2014;111:996–1001.


359


Ueda T, Takahashi H, Uyar E, Ishikawa S, Ogasawara N, Oshima T: Functions of the Hha and Ydgt proteins in transcriptional silencing by the nucleoid proteins, H-NS and StpA, in Escherichia coli. DNA Res 2013;20:263–271. Umbarger MA, Toro E, Wright MA, Porreca GJ, Bau D, Hong SH, Fero MJ, Zhu LJ, Marti-Renom MA, McAdams HH, Shapiro L, Dekker J, Church GM: The three-dimensional architecture of a bacterial genome and its alteration by genetic perturbation. Mol Cell 2011; 44: 252–264. Uyar E, Kurokawa K, Yoshimura M, Ishikawa S, Ogasawara N, Oshima T: Differential binding profiles of StpA in wild-type and H-NS mutant cells: a comparative analysis of cooperative partners by chromatin immunoprecipitation-microarray analysis. J Bacteriol 2009; 191:2388–2391. Valens M, Penaud S, Rossignol M, Cornet F, Boccard F: Macrodomain organization of the Escherichia coli chromosome. EMBO J 2004; 23:4330–4341. van Noort J, Verbrugge S, Goosen N, Dekker C, Dame RT: Dual architectural roles of HU: formation of flexible hinges and rigid filaments. Proc Natl Acad Sci USA 2004;101:6969–6974. van Ulsen P, Hillebrand M, Zulianello L, van de Putte P, Goosen N: The integration host factor-DNA complex upstream of the early promoter of bacteriophage Mu is functionally symmetric. J Bacteriol 1997;179:3073–3075. Vora T, Hottes AK, Tavazoie S: Protein occupancy landscape of a bacterial genome. Mol Cell 2009;35:247–253.

Chromatin Organization by Repetitive Elements (CORE): A Genomic Principle for the Higher-Order Structure of Chromosomes.

Characterization of genomic vitamin D receptor binding sites through chromatin looping and opening.

How chromatin looping and nuclear envelope attachment affect genome organization in eukaryotic cell nuclei.

Enhancer-promoter interactions are encoded by complex genomic signatures on looping chromatin.

Reactivation of developmentally silenced globin genes by forced chromatin looping.

Genome organization: A vision of 3D chromatin organization.

Systematic identification of protein combinations mediating chromatin looping.

Capture of associated targets on chromatin links long-distance chromatin looping to transcriptional coordination.

Correction: Depletion of the Chromatin Looping Proteins CTCF and Cohesin Causes Chromatin Compaction: Insight into Chromatin Folding by Polymer Modelling.

Organization of DNA in chromatin.

Depletion of the chromatin looping proteins CTCF and cohesin causes chromatin compaction: insight into chromatin folding by polymer modelling.

Chromatin regulators of genomic imprinting.

A 3D map of the human genome at kilobase resolution reveals principles of chromatin looping.

Chromatin organization in Entamoeba histolytica.

Levels of granular organization of chromatin fibres.

Organization of spacer DNA in chromatin.

Organization of transcribed regions of chromatin.

Diversity in the organization of centromeric chromatin.

Chromatin Domains: The Unit of Chromosome Organization.

Comparative analysis of metazoan chromatin organization.

Chromatin-like organization of the adenovirus chromosome.

Histone variants: key players of chromatin.

Towards a predictive model of chromatin 3D organization.

Nucleosome organization and chromatin dynamics in telomeres.