The bacterial nucleoid

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Subhash C. Verma1, Zhong Qian2 , Sankar Adhya3

Public peer review comments will be posted here


Bacterial chromosome or nucleoid is composed of the genomic DNA, RNA, and protein. Nucleoid forms by condensation and functional arrangement of the chromosomal DNA with the help of DNA architectural proteins and specific RNA molecules. Although a high-resolution structure of a bacterial nucleoid is yet to come the five decades of research has established the following salient features: 1) The chromosomal DNA is a negatively supercoiled molecule that folds into ~400 independent plectonemic loops or topological domains of ~ 10 kilobase size; 2) The loops spatially organize into megabase size regions called macrodomains, defined by more frequent physical interactions among DNA sites of the same macrodomain than among those of different macrodomains; 3) The condensed and spatially organized DNA takes form of a helical ellipsoid radially confined in the cell; 4) The DNA in the chromosome appears to have a 3-D structure that dictates gene expression, and 5) The nucleoid is also a dynamic entity, its structure is condition dependent. We hope that current advents of high-resolution microscopy and single molecule analysis with the help of structure determination of the components will lead to a better understanding of the molecular structure and function of the bacterial nucleoid.


In most bacteria, the chromosomal DNA is a covalently closed (circular) double-stranded DNA that encodes the genetic information in a haploid form. The size of the DNA varies from 500,000 to several million base-pairs (bp) encoding from 500 to several thousand genes depending on the organism. The chromosomal DNA is present in the cells in a highly condensed, organized form called nucleoid (nucleus-like), which is not encased by a nuclear membrane as in eukaryotic cells. The isolated nucleoid contains 80% DNA, 10% protein, and 10% RNA by weight [1, 2]. In this exposition, we review our current knowledge about (i) how the chromosomal DNA becomes nucleoid, (ii) the factors involved therein, (iii) what is known about its structure, and (iv) how some of the DNA structural aspects influence gene expression, using the gram-negative bacterium Escherichia coli as a model system. We also enlist some related issues that need to be resolved. This exposition is an extension of recent excellent reviews [3, 4].

There are two essential aspects of nucleoid formation; condensation of a large DNA into a small cellular space and functional organization of DNA in a three-dimensional form. The haploid circular chromosome in E. coli consists of ~ 4.6 x 106 bp. If DNA is relaxed in the B form, it would have a circumference of ~1.5 millimeters (0.332 nm x 4.6 x 106) (Fig. 1A). DNA has an inherent property called persistence length, which describes its stiffness and is the maximum length up to which DNA segments remain straight by resisting the bending enforced by Brownian motion. The standard persistence length is ~50 nm or 150 bp. However, the Brownian motion will generate bends in a DNA molecule longer than the persistence length such as the E. coli chromosomal DNA. Due to the bending such long polymers become substantially condensed without any additional factors; at the thermal equilibrium, they assume a random coil form [5, 6]. The random coil of the E. coli chromosomal DNA (Fig. 1B) would occupy a volume (4 ÷ 3 π r3) of ~ 523 µm3, calculated from the radius of gyration (Rg = (√N a) ÷ √6) where a is the Kuhn length (2 x persistence length), and N is the number of Kuhn length segments in the DNA (total length of the DNA divided by N). Although DNA is already condensed in the random coil form, it still cannot assume the volume of nucleoid which is less than a micron (Fig. 1C). Thus, the inherent property of DNA is not enough: additional factors must help condense DNA further on the order of ~103 (volume of the random coil divided by the nucleoid volume). The second essential aspect of the nucleoid formation is the functional arrangement of DNA. The chromosomal DNA is not only condensed but also functionally organized in a way that is compatible with DNA transaction processes such as replication, recombination, segregation, and transcription.

Almost five decades of research beginning in 1971 [1], has shown that the final form of the nucleoid arises from a hierarchical organization of DNA. At the lowest level (1 kb or less), nucleoid architectural proteins condense and organize DNA by bending, looping, bridging or wrapping DNA. At a higher scale (10-kb), DNA forms plectonemic loops that are topologically independent because of supercoiling diffusion barriers. At the megabase scale, nucleoid contains six spatially organized domains (macrodomains) - four structured and two unstructured - as well as mega loops formed by connections of distal DNA segments that also contribute to condensation. Finally, nucleoid is a helical ellipsoid with regions of highly condensed DNA at the longitudinal axis. We discuss these organizational features of the nucleoid and their molecular basis below.

Nucleoid at ≥1 kb scale: DNA condensation and organization by nucleoid-associated proteins and RNAs

A group of small DNA binding proteins referred to as nucleoid-associated proteins (NAPs), help DNA topology and topography by bending, looping, bridging or wrapping DNA (Fig. 2) [7]. NAPs organize DNA throughout the genome because of a genome-wide distribution of their binding sites (Fig. 3). There are at least 12 NAPs identified in E. coli [8]. NAPs also directly participate in a wide variety of DNA transaction processes such as transcription, DNA replication, recombination, and repair. Here we focus on properties of the most extensively studied NAPs relevant to DNA condensation and organization although some of the organization events may help the transaction reactions.


Histone-like protein from E. coli U93 (HU) is an evolutionarily conserved protein in bacteria [9, 10]. It exists in E. coli as homo- and heterodimers of two subunits HUα and HUβ sharing 69% amino acid identity [11]. ΔhupAB strains show the nucleoid decondensed suggesting a role of HU in nucleoid condensation [12]. HU exhibits low-affinity (10-6 M) non-sequence specific binding to DNA [13]. Because of non-specific binding, HU shows a weak uniform binding signal across the genome (Fig. 3). It appears that this low-affinity binding is involved in nucleoid organization. HU also shows specific high-affinity binding to structurally altered DNA such as cruciform DNA, dsDNA containing a single-stranded break such as nicks, gaps, or forks [13-18]. The high-affinity binding is mostly involved in DNA transaction reactions in site-specific recombination, repair, replication initiation, and gene regulation [19-21].

Although how HU condenses and organizes DNA in vivo is not understood, the following in vitro studies suggest possible mechanisms. According to in vitro studies with linear 0.5 – 2 kb size DNA, the HU heterodimer induces flexible bends in DNA at lower protein concentrations but forms thick rigid nucleoprotein filaments at higher concentrations; homodimers were not studied [22]. Furthermore, a pattern of HU regularly bound on DNA was observed that given the 9 bp DNA as the minimum binding length led to a model in which both DNA and HU dimer wrap around each other in a spiral fashion (Fig. 2A top) [18, 22]. Note that in the magnetic tweezers experiments, the flexible bending results in mild DNA condensation whereas formation of rigid filaments at higher concentrations results in no condensation. Magnetic tweezers allow studying condensation of a single DNA molecule by a DNA binding protein [23]. In a magnetic tweezers experiment, one end of a single linear DNA molecule attaches to a glass surface and the other end to a fluorescently labeled magnetic bead. Under magnetic force, an increase or a decrease in the DNA tether length in the presence of a protein is defined as DNA de-condensation or condensation respectively.

In contrast to flexible bends and the formation of rigid filaments, structural studies of both HUαα and HUαβ bound to a 20 bp random DNA sequence revealed that HUαα dimers but not the heterodimers multimerize laterally on DNA in the co-crystal packing causing straightening of DNA axis rather than bending [24]. The multimerization results in DNA network (DNA bunching) expandable both laterally and medially (Fig. 2A bottom). This observation led to a proposal that the formation of DNA networks through lateral HUαα multimerization occurs in vivo and contribute to DNA condensation.

Nucleoid-associated RNAs

The early studies examining the effect of RNase A treatment on isolated nucleoids indicated the participation of RNA in the stabilization of the nucleoid in the condensed state [25]. Moreover, the treatment of RNase A disrupts the DNA fibers, observed by the atomic force microscopy of the nucleoid using the “on-substrate lysis procedure,” into thinner fibers [26]. These findings demonstrated the participation of RNA in the nucleoid structure, but the identity of the RNA molecule(s) remained unknown until recently.

Most of the studies on HU have focused on its DNA binding. However, HU also binds to RNA and prefers DNA-RNA hybrids [27]. Immunoprecipitation of HU-bound RNA coupled to reverse transcription and microarray (RIP-Chip) study as well as analysis of RNA from purified intact nucleoid identified nucleoid-associated RNA molecules that interact with HU [12]. Several of them are non-coding RNAs, and one such RNA named naRNA4 (nucleoid-associated RNA), encoded in a repetitive extragenic palindrome (REP324), connects DNA segments both in vivo and in vitro in the presence of HU [28]. The RNA forms an RNaseH sensitive complex with DNA molecules containing cruciform structures, and the complex formation is critical for DNA-DNA connections. The same complex binds to an antibody against RNA-DNA hybrids. Surprisingly, although HU is essential for the complex formation, it is not present in the final complex indicating its potential role as a catalyst (chaperone) in the naRNA4-mediated DNA condensation process [29]. Like in ΔhupAB strain the nucleoid is decondensed in Δrep324 strain [12].


Integration host factor (IHF) is closely related to HU at the amino acid sequence level and has the same protein fold as in HU [30, 31]. Despite structural similarity, IHF is more of a specific DNA binding protein. A crystal structure of IHF heterodimer in a complex with 35 bp DNA containing a high-affinity IHF binding site shows DNA wrapped around the heterodimer and bent by >160° [30]. It is noteworthy that the bends induced by IHF are nearly coplanar thus do not generate a writhe in the DNA. There are close to 1000 specific binding sites in the genome identified by a chromatin immunoprecipitation followed by deep sequencing (ChIP-Seq) study [32]. IHF is likely to organize DNA by introducing sharp bends at those sites (Fig. 2B). In vitro, IHF also shows non-sequence specific DNA binding that can cause DNA condensation in a magnetic tweezers experiment depending on concentrations of potassium chloride and magnesium chloride [33]. However, whether this activity has any relevance to in vivo DNA condensation is not clear.


Histone-like or heat-stable nucleoid structuring protein (H-NS) was identified as a significant component of isolated nucleoids [34-37]. It exists predominantly as a homodimer in solution at a relatively low concentration (<1 x 10-5 M) and forms oligomers at higher levels [38, 39]. H-NS binds selectively to 458 binding regions in the genome [40]. Although H-NS has been demonstrated to prefer curved DNA formed by repeated A-tracks in DNA sequence [41, 42] the basis of the selectivity is the presence of a conserved sequence motif found in AT-rich regions [43].

Because of oligomerization property, H-NS spreads on DNA and forms two types of nucleoprotein filaments depending on the magnesium concentration in the reaction (Fig. 2C). At less than 5 mM magnesium, H-NS forms rigid nucleoprotein filaments whereas it forms inter- and intra-molecular bridges at greater than five mM magnesium [44-48]. In the magnetic tweezers experiments, the formation of rigid filaments results in straightening of DNA whereas the bridging causes an abrupt reduction in the DNA extension, indicative of substantial DNA folding [47]. Less than five mM physiological concentration of magnesium inside the cells [49] argues that the primary mode of DNA organization by H-NS in vivo is the formation of rigid nucleoprotein filaments.

Interestingly, super-resolution imaging of fluorescently tagged H-NS revealed that 26% of the total protein reside in 2-3 DNA bound clusters and the rest is scattered throughout the genome [50, 51]. Given the distribution of the binding regions across the genome (Fig. 3), the clustering of the protein led to a model in which H-NS brings the distant sites together to form long loops in DNA and thus helps condensation. However, there are contradicting reports regarding the connections between H-NS regulated loci (see below). Moreover, the absence of H-NS does not change the nucleoid volume [51]. A paralog of H-NS, StpA, shows DNA binding properties like H-NS such as binding to curved DNA and formation of rigid filaments [52-55] and is expected to organize DNA in vivo in a similar fashion.


Factor for Inversion Stimulation (Fis) protein is exclusively present in the growth phase [56, 57]. Fis is a specific DNA binding protein that binds with <1 x 10-9 M affinity to DNA sequences containing a 15-bp symmetric motif composed of an A/T-rich central core and a highly conserved G/C pair at the boundaries (GNTYAAAWTTTRANC; Y = pyrimidine, R = purine, W = A or T, and N = any nucleotide) [58-60]. The distance between the DNA recognition helices in helix-turn-helix (HTH) motifs of the Fis homodimer is 25  Å, ~ 8  Å shorter than the pitch of a canonical B-DNA, indicating that the protein must bend and/or twist DNA to bind stably [61, 62]. Consistently, the crystal structure of Fis-DNA complex shows that the distance between the recognition helices remains unchanged whereas DNA curves in the range of 60-75° [59]. There are 1464 Fis binding regions distributed across the genome and the binding motif, identified computationally, matches with that determined from in vitro experiments [40, 63]. Many Fis binding regions occur in tandem such as those in the stable RNA promoters, e.g., p1 promoter of rRNA operon rrnB. According to a model, if tandem sites are helically phased as in the stable RNA promoters, coherent bending by Fis creates a topologically distinct DNA micro-loop (Fig. 2D) [64].

Although Fis recognizes specific DNA sequences, in vitro studies show that Fis can bind to a random DNA sequence with 1 x 10-9 M affinity, and the non-specific binding can contribute to DNA condensation and organization as shown by magnetic tweezers experiment and electrophoretic mobility gel shift assays (EMSA) [65, 66]. Fis causes mild condensation of a single DNA molecule at less than one micromolar concentrations but induces substantial folding through the formation of DNA loops of an average size of ~800-bp at greater than or equal to one micromolar concentration. The loops in magnetic tweezers experiments are distinct from the micro-loops, as they require the formation of high-density DNA-protein complexes achieved by sequence-independent binding. Given that Fis binds to specific sites in the genome whether such loops occur in vivo remains to be demonstrated. However, it was proposed that the in-tandem occurrence of specific sites might drive the formation of localized high-density Fis arrays through a nucleation reaction similar to that of H-NS, and the bridging between the localized regions can create large DNA loops [66]. Since Fis is absent in stationary phase, any condensing chromosomal structure generated by Fis is specific to growing cells.

Nucleoid at 10-kb scale: Topological organization of nucleoid


Because of helical structure, a double-stranded DNA molecule (Fig. 4A) becomes topologically constrained in the covalently closed circular form (Fig. 4B), which eliminates the rotation of the free ends [67]. The number of times the two strands cross each other in a topologically constrained DNA is called linking number (Lk), which is equivalent to the number of helical turns or twists in a circular molecule. Lk of a topological DNA remains invariant, no matter how the DNA molecule is deformed, as long as neither strand is broken. The Lk of DNA in the relaxed form is defined as Lk0. For any DNA, Lk0 can be calculated by dividing the bp length of the DNA by the number of bp per helical turn, which is equal to 10.5 bp for the relaxed B-form DNA. Any deviation from the Lk0 causes supercoiling in DNA. A decrease in Lk (Lk<Lk0) creates negative supercoiling (Fig. 4C-D) whereas an increase in Lk (Lk>Lk0) creates positive supercoiling (Fig. 4E-F) (more detail on supercoiling discussed in [68, 69].

Supercoiled state (Lk ≠ Lk0) results in a transition in DNA structure that can manifest as a change in the number of twists (Fig. 4D and E) (negative <10.5 bp/turn, positive >10.5 bp per turn) and/or in the formation of writhes, also called supercoils (Fig.4C and F). Thus, Lk is mathematically defined as sign dependent sum of the two geometric parameters, twist and writhe. A quantitative measure of supercoiling independent of the size of DNA molecules is the supercoiling density, σ, where σ =∆Lk/Lk0.

Supercoils can adopt two structures; plectoneme and solenoid or toroidal. A plectonemic structure arises from interwinding of the helical axis (Fig. 4C and F). Toroidal supercoils originate when DNA forms several spirals, around an imaginary axis, which can be best illustrated by a garden hose wrapped on a reel or a coiled telephone cord. Supercoils in the plectonemic form are right- and left-handed for a negatively and positively supercoiled DNA respectively, whereas the handedness of the toroidal supercoils is opposite to those of plectonemes. Both plectonemes and toroidal supercoils can be either in a free form or restrained in a bound form with proteins. The best example of the bound toroidal supercoiling in biology is the eukaryotic nucleosome in which DNA wraps around histones [70].

Topological domains structure of the nucleoid

In most bacteria, the DNA is present in the supercoiled form. The circular nature of the E. coli chromosome makes it topologically constrained molecule which is mostly negatively supercoiled with estimated average supercoiling density (σ) of -0.05 [71]. Unlike eukaryotic chromosomes in which there is almost no free form of supercoiling because nucleosomes restrain almost all negative supercoiling through tight bindings of DNA to histones, half of the supercoiling density in the nucleoid is present in the free form whereas nucleoprotein complexes restrain the other half [71]. One of the striking features of the nucleoid is that the free supercoiling is not confined within a single topological domain (Fig. 5A) but instead in independent plectonemic loops or topological domains separated by the supercoiling-diffusion barrier(s) (see below) [72] (Fig. 5B). In other words, a single cut in one supercoiled domain will only relax that loop and not the others. It was estimated that nucleoid consists of ~ 460 topological domains with an average size of ~10 kb per domain [73].

What are the biological implications of the topological organization of the nucleoid? Plectonemic looping should aid in DNA condensation. It is noteworthy that because of branching of the plectonemic structure, it provides less DNA condensation than the toroidal structure. A same size DNA molecule with equal supercoiling densities is more compact in a toroidal form than in a plectoneme form. In addition to condensing DNA, supercoiling promotes disentanglement of DNA by reducing the probability of catenation thus helps in DNA organization [74]. Supercoiling also helps bring two distant sites of DNA in proximity thereby promoting a potential functional interaction between different segments of a DNA and thus helps DNA condensation by DNA looping.

Supercoiling diffusion barriers

Wide variability in the size of domains [73] indicated that the domains form dynamically and the positions of the topological barriers are random. The possible mechanisms responsible for the formation of the barriers could be different: (i) A barrier could form if a protein complex traps the two distinct segments of DNA in a loop. It has been experimentally demonstrated that protein-mediated looping in a supercoiled DNA can create a topological domain [75, 76]. H-NS and Fis appear as potential candidates based on the DNA looping abilities and the distribution of their binding sites. Of them, H-NS appears to be a good candidate because there are 458 H-NS binding regions in the genome with average separating distance of 11 kb that matches with the average topological domain size of ~ 10 kb. (ii) A genetic screen identified, in addition to H-NS and Fis, DksA, and the two carbohydrate metabolic enzymes phosphoglucomutase (Pgm) and transketolase (TktA) as potential candidates that meet the requirement of forming a topological barrier. (iii) Barriers could exist because of attachment of DNA to the cell membrane through a protein which binds to both DNA and cell membrane or through the transertion mechanism (see below). (iv) Restrained supercoils can also act as barriers motion.

Specific catalytic targets of DNA gyrase such as par sequence from pSC101 plasmid and a bacterial interspersed mosaic element (BIME) located between nrdA and nrdB genes in E. coli can impede diffusion of transcription-induced supercoiling in a synthetically designed topological cassette inserted in the chromosome [77]. There are ~ 600 BIMEs distributed across the genome, and several of them contain an IHF binding site and can associate with DNA gyrase providing a hypothesis that BIMEs are a critical determinant for the formation of barriers [78, 79].

Creation and maintenance of DNA supercoiling in E. coli

Three factors contribute to generating and maintaining chromosomal DNA supercoiling in E. coli: (i) activities of topoisomerases, (ii) the act of transcription, and (iii) supercoiling restraining proteins.


Topoisomerases are a particular category of DNA metabolic enzymes that create or remove supercoiling by breaking and then re-ligating DNA strands [80]. E. coli possess four topoisomerases (Table 1). DNA gyrase introduces negative supercoiling in an ATP-dependent reaction and removes positive supercoiling in the absence of ATP [81]. DNA gyrase is the only topoisomerase in all forms of life that can create negative supercoiling. Topo I acts opposite to DNA gyrase by relaxing the negatively supercoiled DNA [82, 83]. There is genetic evidence to suggest that a balance between the opposing activity of DNA gyrase and Topo I is responsible for maintenance of a steady-state level of negative superhelicity in E. coli. Both enzymes are essential for the survival of E. coli. A null strain of topA, the gene encoding Topo I, survives only because of the presence of suppressor mutations in the genes encoding DNA gyrase [82, 87]. The mutations result in reduced gyrase activities suggesting that excess negative supercoiling caused by the absence of Topo I is compensated by reduced activity of DNA gyrase to introduce negative supercoiling. Although the primary function of Topo IV is to resolve sister chromosomes, it has been shown to also contribute in the steady-state level of negative supercoiling by relaxing negative supercoiling along with Topo I [84, 85]. Topo III is dispensable in E. coli and has no known role in supercoiling [86]. Gyrase inhibition alone does not cause an increase in nucleoid volume. Decondensation upon gyrase inhibition only occurs if the nucleoid is over-condensed due to exposure of cells to chloramphenicol suggesting that other factors can compensate condensation force of gyrase-induced supercoiling [88].


Liu and Wang put forth a twin supercoiling domain model to show that the unwinding of DNA double helix during transcription induces supercoiling in DNA (Fig. 6) [89]. According to the model, If the rotation of transcribing RNA polymerase (RNAP) relative to the helical axis of DNA is hindered and instead RNAP moves along the helical groove of DNA, it would force DNA to rotate on its helical axis. A hindrance in the free rotation of DNA due to a topological constraint would cause DNA in front of RNAP to become over-twisted or positively supercoiled and DNA behind it to become under-twisted or negatively supercoiled. If the template DNA is already negatively supercoiled, this action removes existing negative supercoiling ahead and enhance it behind RNAP. It is noteworthy that any transcription-induced supercoiling would confine to DNA segments of the genes being transcribed in a given topological domain and not contribute to DNA segments of other domains. Supercoiling of the other domains must originate locally by transcription or other means.

DNA gyrase and Topo I can remove the transcription-induced supercoiling by relaxing positive and negative supercoils respectively. However, if the elongation rate exceeds the turnover of the two enzymes, the process of transcription contributes to the steady-state level of supercoiling. Exposure of cells to rifampicin, a transcription blocking drug, causes de-condensation of the nucleoid [90, 91]. The effect is observed only after a few minutes of the exposure suggesting that the inhibition of transcriptional activities reduces supercoiling, hence the chromosome de-condensation.

Supercoiling restraining proteins

Alternatively, or in combination, supercoiling can be manifest in the form of free plectonemes or can be stabilized through binding of restraining proteins in a toroidal form. HU, Fis, H-NS, and MukB (see below) can constrain negatively supercoiled DNA [92-96]. Besides, Fis can modulate supercoiling by repressing gyrA and gyrB transcription through direct binding. There is genetic evidence to suggest that HU maintains supercoiling levels by stimulating DNA gyrase and reducing the activity of Topo I [97, 98]. In support of the genetic studies, HU stimulates DNA gyrase-catalyzed decatenation of DNA in vitro [99]. However, it is unclear how HU modulates the activities of the two enzymes.

In summary, the negative supercoiling of the nucleoid is a result of a combination of the following: (i) a dynamic balance between Topo I and DNA gyrase occurring at both expression level and enzymatic activity of the two proteins. (ii) transcription elongation rate and the turnover rate of Topo I and DNA gyrase in the actively transcribed region, and (iii) restraining of supercoiling by the formation of nucleoprotein complexes.

Nucleoid at megabase scale: Spatial organization of nucleoid

Spatial domains of the nucleoid

The precise location of the replication origin (oriC) and terminus (ter) of the E. coli chromosome inside the cell provided the first evidence that nucleoid is spatially organized [100, 101]. In a newly born E. coli cell, oriC and ter are at the mid-cell and the left and right replication arms (replichores) are present in the opposite cell halves generating the left-ori-right arrangement [102]. The oriC-ter axis lies perpendicular to the cell’s long axis creating a transverse configuration (Fig. 7A). It is noteworthy that other bacteria such as Bacillus subtilis and Caulobacter crescentus have a longitudinal arrangement. The ori and ter are present at the opposite cell poles with the right and left arms to lie side by side.

In E. coli, ~ 1-Mb regions containing DNA segments that colocalized with oriC and ter in fluorescent in situ hybridization were referred to as Ori and Ter macrodomains (MDs) respectively [101]. Later, measurement of site-specific recombination frequencies between pairs of lambda att sites inserted at various distant locations in the chromosome confirmed the Ori and Ter MDs and identified two additional MDs [103, 104]. An MD was defined by a large genomic region whose DNA sites primarily showed recombination with each other but not with those outside of that MD. The two additional MDs were formed by the regions flanking the Ter and were referred to as Left and Right. The four MDs comprised most of the genome except the two genomic regions flanking the Ori which were more flexible and showed recombination with the DNA sites in MDs on both sides (Fig. 7A). The two regions were referred to as less-structured regions (NS). The genetic position of oriC appears to dictate the formation of MDs because the repositioning of the oriC by genetic manipulation results in reorganized nucleoids such that regardless of DNA sequence the genomic regions closest to the oriC always behave as an NS and the regions farther behave as an MD [105].

In a recent study, a high-resolution spatial organization of the nucleoid was revealed using chromosome conformation capture (3C) coupled with deep sequencing [106]. 3C is a molecular method that determines physical proximity, if any, between any two genomic loci in 3D space [107]. The contact map revealed partitioning of the chromosome into two distinct domains. The region surrounding ter formed an insulted domain that overlapped with previously identified the Ter MD (Fig. 7B). The DNA contacts in the Ter occurred in the range of only up to ~280 kb. The rest of the chromosome formed a single domain allowing two modes of DNA communications: long-range contacts in the >280-kb range and mid-range contacts <280-kb. The study revealed two loose regions corresponding to previously identified NS regions whose genomic loci formed contacts at even more considerable distances (Fig. 7B). The boundaries of Ter and the two loose regions segmented the entire chromosome into six regions that overlap with the four MDs and two NS regions defined by the recombination-based assays (Fig. 7B).

Spatial domain-specific proteins


Macrodomain Ter protein (MatP) binds to a 13-bp motif called the macrodomain ter sequence (matS) [108]. There are 23 matS sites present in the Ter domain, one every 35-kb on average. Consistently, the global analysis of MatP binding by ChIP-Seq shows strong enrichment of MatP binding signal in the Ter [109] (Fig. 8A). Consistent with the selective binding of MatP in the Ter domain, fluorescence imaging of MatP reveals the formation of discrete MatP foci colocalizing with the DNA markers of the Ter domain throughout the cell cycle [108].

MatP appeared as a critical player in insulating the ter region from the rest of the chromosome [106]. It promotes DNA-DNA contacts within the Ter but prevents DNA contacts between the Ter and the flanking regions. How does MatP function in the formation of a spatial domain? The lack of MatP caused an increase in the distance between two fluorescent DNA markers located 100-kb apart in the Ter domain indicating that MatP condenses the ter region [110]. MatP can form a loop between two matS sites in vitro and its DNA looping activity depends on tetramerization through coiled-coil interactions between two MatP molecules bound to DNA (Fig. 8D) [110]. The DNA looping activity suggested MatP bridges the matS sites in vivo to organize the Ter domain. (Fig. 8E). Although MatP promoted the DNA-DNA contacts in the ter region, MatP does not specifically connect matS sites, and the MatP mutant unable to form tetramers does not modify the contacts either [106]. These results refute the matS bridging model for the Ter organization and leave the mechanism of MatP action elusive.


MukB shares structural similarity to a family of ATPases called structural maintenance of chromosome proteins (SMC), which participate in higher order chromosome organization in eukaryotes [111]. Two MukB monomers associate via continuous antiparallel coiled-coil interaction forming a 100-nm long rigid rod containing a flexible hinge region in the middle [112, 113]. Due to the flexibility of the hinge region, MukB adopts the characteristic V-shape of the SMC family (Fig. 8B). The SMC proteins form a heterotrimeric complex by associating with non-SMC proteins. The association closes the V formation resulting in large ring-like structures (Fig. 8B). Two of the most studied SMC complexes are condensins and cohesins, required for mitotic chromosome condensation and sister chromatid cohesion respectively, during cell division in eukaryotes [111]. The non-SMC subunits associating with MukB are MukF and MukE, encoded together with MukB in the smtA-mukF-mukE-mukB operon of E. coli [114].

An E. coli mutant defective in any of the subunits of the MukBEF complex exhibits temperature-sensitive lethality and produces anucleate cells, indicative of chromosome mis-segregation [115]. The MukBEF complex colocalizes with oriC and remains so as sister oriCs move to 1/4 and 3/4 quarter positions during replication [116]. Moreover, sister oriCs are aberrantly positioned at the opposite old poles in the mukB null mutant [117]. These findings along with other studies that found a physical and functional link between MukB and Topo IV led to a model that the MukBEF together with Topo IV is required for decatenation as well as repositioning of newly replicated oriCs [109, 118-121].

Single molecule fluorescence microscopy of MukBEF in cells suggests that the minimum functional unit in vivo is a dimer of dimers formed by joining of the two ATP-bound MukBEF complexes through MukF-mediated dimerization [131]. The minimum functional unit forms 1-3 clusters elongated parallel to cell’s long axis each containing on an average ~ 8-10 dimer of the dimers. In B. subtilis, the structural homolog of the MukBEF complex is recruited to centromeric sites (oriC) through a parS/ParB system, and it zips the two replichores through a DNA loop extrusion mechanism while it travels toward the ter [132]. In contrast to the chromosome of B. subtilis, the E. coli chromosome shows no interactions between its right and left arms, perhaps due to the transverse organization, and the parS/ParB system is absent in E. coli [106]. Moreover, despite co-localization of MukBEF complex to oriC in vivo, there is no enrichment of binding in the oriC region [109]. Therefore, MukBEF appears to function via a mechanism different from that of the other SMC-complexes.

The following observations suggest a model that MukBEF organizes newly replicated DNA into a condensed form through supercoiling [74]. First observation is that the topA mutation suppresses mukB - phenotypes suggesting that the increased levels of negative supercoiling resulting from the reduced activity of Topo I due to the topA mutation can compensate for the impairment of the MukBEF function [122]. Second observation is that a gyrase inhibitor can reverse the topA-mediated suppression, and the mukB null mutant itself is hypersensitive to gyrase inhibitors [123]. The DNA transaction activities of the complex reside in the SMC subunit MukB and the ability of MukB to constrain negative supercoiling in vitro supports a direct link between negative supercoiling and in vivo MukBEF function and strengthens the model [124].

The role of MukBEF in organizing and condensing DNA is not restricted during DNA replication but is critical even in the non-replicating cells [125]. It is possible that the chromosome segregation defects observed in the absence of MukB are indirect consequences of the disruption of the nucleoid structure and DNA condensation. In fact, the nucleoid is decondensed in the absence of MukB and becomes over condensed when MukB is over produced [126]. In vitro studies further demonstrated that MukB causes condensation of a single DNA molecule and stabilizes DNA bridges between two separate DNA molecules [127, 128]. Recent biochemical experiments suggest a step-wise mechanism for DNA condensation by MukB [129] (Fig. 8C). First, MukB binds to DNA through the head domain. Subsequently, the oligomerization of MukB dimers via hinge-hinge interactions results in the formation of large topologically isolated domains that are supercoiled into plectonemic loops by DNA gyrase. Alternatively, or in combination, because of its ring-like structure, MukB can also entrap topological DNA loops for which it prefers the circular single-stranded DNA (cssDNA) to a double-stranded circular DNA (Fig. 8B) [130]. However, there are contradicting reports about topological entrapment of DNA by MukB [129].

The recent 3D contact map of E. coli genome in the MukB-depleted cells show that MukB participates in the formation of DNA-DNA interactions in the chromosome except in the Ter domain [106]. How is MukB prevented from acting in the Ter domain? It can be explained by the observations that MatP physically interacts with MukB and thus prevents MukB to localize in the Ter domain [109]. Consistently, DNA binding signal of MatP and MukB in the Ter domain shows a negative correlation (Fig. 8A); the binding signal of MatP is enriched whereas that of MukB is reduced compared to the rest of the genome. In agreement with the displacement of MukB by MatP, MukB is not restricted to function in the Ter domain in a strain already lacking MatP [106].

Role of NAPs in the spatial organization of nucleoid

A high-resolution contact map of individual mutants of NAPs, Fis, H-NS, and HU revealed their role in the spatial organization of nucleoid [106]. HU and Fis emerged as key players in promoting long-range DNA-DNA contacts occurring throughout the chromosome including the insulated ter region. There was no correlation between the binding profiles of the two NAPs and the observed contacts. Therefore, whether the DNA organization modes of HU and Fis at the lower scale (discussed before) play a role in higher-order organization remains to be tested. The study did not examine naRNAs, but a study using the original 3C method showed that some of the HU-mediated DNA interactions require the presence of the naRNA4 [28]. The naRNA4 promotes long-range DNA contacts without HU suggesting that the RNA also participates with other NAPs in forming contacts [28]. HU also appears to act together with MukB to promote long-range DNA-DNA interactions because the DNA contact pattern of the MukB-depleted cells outside the Ter domain was remarkably similar to that of HU deficient cells. It is unclear how MukB and HU potentially act together in promoting DNA-DNA interactions. The study did not find H-NS-mediated long-range contacts including the connections between H-NS regulated genes that were reported before [50]. Instead, H-NS was found to prevent short-range contacts within the H-NS binding regions possibly due to the formation of rigid nucleoprotein filaments by spreading [106].

Spatial organization of functionally related genes

A study examining pairwise physical distances between the seven rRNA operons that are genetically separated from each other by as much as two million bp found that all the operon except the rrnC are present in physical proximity [133]. In another example, GalR, a specific transcription regulator of the galactose regulon comprising genes encoding enzymes for transport and metabolism of the sugar D-galactose [134], forms an interaction network of GalR binding sites that are scattered across the chromosome [135]. GalR forms only one to two foci in the cells [135] and can self-assemble into large ordered structures [136] suggesting the multimerization of DNA-bound GalR causes the long-distance interactions. The spatial proximity of the functionally related genes perhaps occurs to increase an output of the biological process; facilitation of the ribosome assembly and the co-regulation of the genes in case of the rrn operons and GalR respectively, but it could also contribute in the folding and spatial organization of the nucleoid. Based on the theoretical modeling, it has been proposed that the gene regulatory network requiring the colocalization of the transcription regulatory genes and the target genes is the basis for the topological organization of the nucleoid [137].

Nucleoid at global scale: Nucleoid shape and structure

Nucleoid is a helical ellipsoid confined radially in the cell

It remains a significant challenge to accurately describe the hierarchical organization of the chromosomal DNA resulting in the formation of a functional nucleoid even at the 10-kb resolution let alone any single nucleotide resolution. However, we have significantly improved our understanding of the global structure and shape of the nucleoid. Conventional transmission electron microscopy (TEM) of chemically fixed E. coli cells portrayed nucleoid as an irregularly-shaped organelle. However, wide-field fluorescence imaging of live nucleoid in 3D revealed a discrete ellipsoid shape (Fig. 9) [4, 138, 139]. A close juxtaposition of the nucleoid along its entire length to the cell periphery only in the radial dimension indicated radial confinement of the nucleoid [138]. Furthermore, 3D fluorescence imaging revealed two striking features of the nucleoid. It is curved without preferential handedness (Fig. 9) [139] and consists of high-density regions or bundles at the central core and low-density regions at the periphery (Fig. 1C) [138, 140].

How is the helical nature of the nucleoid determined? It could be an internal feature of the nucleoid defined solely by how DNA is condensed and organized by supercoiling, NAPs, and MD-specific proteins. Either removal of the cell wall or inhibition of cell wall synthesis induces an increase in the radius of the E. coli cell causing the rod-shaped cells to become round-shaped. The nucleoid of these cells also lose its ellipsoid shape and becomes round-shaped due to an increase of helical radius and a decrease of helical pitch [138]. These observations suggested an alternative model that nucleoid is forced to assume a curved shape because of the radial confinement within a cylindrical cell whose radius is smaller than the bendable length or persistence length of the entire nucleoid. Furthermore, because all the detectable fluorescent signal constituted the ellipsoid shape of the nucleoid visualized by fluorescence microscopy, it was suggested that a nucleoid is a self-adhering object [138]. DNA-membrane contacts were invoked but not ruled out in defining the nucleoid shape and helical organization.

Nucleoid-membrane connections

It is possible that the DNA-membrane connections are not visible in the fluorescence imaging because of less DNA density. Cell-fractionation and electron microscopy studies first indicated a possibility of DNA-membrane connections [141, 142]. The transertion, a mechanism of concurrent transcription, translation, and insertion of the membrane proteins can form transient DNA-membrane contacts [143]. Fluorescence imaging of the chromosomal loci of the two membrane proteins LacY and TetA has provided direct evidence of transertion [144]. The induction of the protein expression caused repositioning of the chromosomal loci toward the membrane. There is another mechanism independent of translation for the positioning of the membrane-protein encoding loci on the membrane. The bglF transcript localizes to the membrane, and the membrane-target signal is in the transcript itself [145]. Since membrane-protein encoding genes are distributed throughout the genome, dynamic DNA-membrane contacts through transertion or alternative mechanisms can act as nucleoid expansion force functioning in opposition to the condensation forces to maintain an optimal condensation level. The formation of highly condensed nucleoids upon the exposure of E. coli cells to chloramphenicol that blocks translation provides support for the expansion force of transient DNA-membrane contacts formed through transertion [146, 147]. The round shape of the over-condensed nucleoids upon chloramphenicol treatment also suggests a role of transertion-mediated DNA-membrane contacts in defining the ellipsoid shape of the nucleoid.

Nucleoid dynamics

Nucleoid structure must be remodeled after or simultaneously with chromosome replication. Sister oriCs go through a positional rearrangement during DNA replication and segregation to recreate the transverse configuration of the nucleoid in the daughter cells demonstrating that nucleoid is spatially organized and yet dynamic [102]. Nucleoid structure is also dependent on the physiological state of the cells. The high-resolution contact map of cells in stationary phase revealed a nucleoid reorganization [106]. The long-range contacts in the ter region were more pronounced in the stationary phase than the growth phase. Furthermore, the boundaries of chromosome interaction domains in the stationary phase were different from those found in the growth phase.

The nucleoid reorganization in stationary phase could be brought about by changes in expression levels of NAPs [148] and the Muk subunits. The copy number of MukB increases two-fold in stationary phase [125, 149]. Fis is only present in the growth phase [150]. Fis levels rise upon entry into exponential phase and then rapidly decline while cells are still in the exponential phase reaching undetectable levels in the stationary phase [150]. HUαα is the predominant form in the early exponential phase whereas heterodimeric form becomes predominant in the stationary phase with minor amounts of the homodimers [151]. This transition has functional consequences regarding nucleoid dynamics because the two forms appear to organize and condense DNA differently; both homo- and heterodimer form filaments but only the homodimer can bring multiple DNA segments together to form a DNA network (Fig. 2A) [24]. Supercoiling can act in a concerted manner with the protein factors to reorganize the nucleoid. The overall supercoiling level decreases in the stationary phase, but supercoiling exhibits a different pattern at the regional level [152].

During the prolonged stationary phase, nucleoid morphology undergoes massive transformation mainly due to starvation-induced DNA binding protein Dps [153]. The nucleoid exhibits ordered toroidal structures [154]. As the stationary phase continues, Dps forms DNA crystalline assemblies that act as a mean to protect the nucleoid from the DNA damaging agents during starvation [154].

Nucleoid structure-function: Nucleoid structure and gene expression

NAPs and gene expression

A 3D structure of DNA within nucleoid is linked to gene expression at perhaps different scales. How the higher-order nucleoid structure and its dynamics influence gene expression is an entirely open question. At least, we have a better understanding of how local changes in DNA topology such as DNA bending, stiffening, bridging and looping induced by NAPs influence gene expression.

H-NS spreads laterally along AT-rich DNA where high-affinity sites containing the conserved sequence motif function as nucleation centers [42, 155]. The frequent occurrence of the motif within an H-NS binding region and the unusually long length of an H-NS binding region are consistent with spreading of the protein [40]. The spreading of H-NS from the high-affinity sites over an extended region of DNA renders a promoter such as the proU and bgl promoters inaccessible to RNAP [42]. The spreading can not only render the promoter inaccessible to RNAP and other activators for transcription initiation but also affect transcription elongation, termination, and RNAP pausing [156] . This mechanism of transcription inhibition is called gene silencing. Through gene silencing, H-NS acts as a global repressor preferentially inhibiting transcription of horizontally transferred genes because of their AT-rich content [40, 43]. At some promoters such the rrnB p1 promoter, instead of making a promoter inaccessible to RNAP, H-NS forms DNA bridges on both sides of RNAP thereby trapping the RNAP in an open initiation complex [157].

Specific binding of HU at the gal promoter facilitates the formation of a DNA loop that keeps the gal operon repressed in the absence of the inducer [158]. Coherent bending of DNA by Fis at the stable RNA promoters results in the formation of a topologically distinct DNA loop that activates transcription via the torsional transmission [64]. DNA bending by IHF differentially controls transcription from the two tandem promoters of the ilvGMEDA operon in E. coli [159, 160].

DNA supercoiling and gene expression

A two-way interconnectedness exists between DNA supercoiling and gene transcription [161]. Negative supercoiling of the promoter region can stimulate transcription by facilitating the promoter melting and by increasing the DNA binding affinity of a protein regulator. According to the twin supercoiling domain model, supercoiling induced by transcription of a gene can influence transcription of other nearby genes through a supercoiling relay. One such example is the activation of the leu-500 promoter [161]. Because of the segregation of the chromosomal DNA into independent topological domains, a local change in supercoiling of one topological domain will affect transcription of supercoiling-sensitive promoters (SSG) only in that domain [73]. Therefore, the topological organization provides a regulatory mechanism to coordinate expression of SSG in different topological domains independently. A genome-scale map of supercoiling showed that genomic regions have different steady-state supercoiling densities indicating that level of supercoiling differs in the individual topological domains [152]. As a result, a global change in supercoiling can result in differential expression of SSG in the different domains depending on the existing level of supercoiling in each domain.

The effect of supercoiling on gene expression can be mediated by NAPs that directly or indirectly influence supercoiling. The effect of HU on gene expression appears to involve a change in supercoiling and perhaps a higher-order DNA organization. 1266 genes are differentially expressed in a HU deficient strain [32]. A positive correlation between gyrase binding and upregulation of the genes in the HU double mutant suggests that changes in supercoiling are responsible for differential expression. Another transcriptome study also found a connection between supercoiling and gene expression for HU [162]. Furthermore, HU was found to be responsible for a positional effect on gene expression by insulating transcriptional units by constraining transcription-induced supercoiling [163]. E38K and V42L mutations in HUα dramatically change gene expression program of E. coli altering morphology, physiology, and metabolism making the mutant strain more invasive to mammalian cells [164, 165]. The dramatic effect is concomitant with nucleoid compaction and increased positive supercoiling [24, 166]. The mutant protein is an octamer in contrast to the wild-type dimer and wraps DNA on its surface in a right-handed manner restraining positive supercoils [166]. These studies show that mutations in HU could lead to a dramatic impact on nucleoid structure that in turn results in significant phenotypic changes.

Although HU appears to control gene expression by modulating supercoiling density, the exact molecular mechanism remains unknown. Since MukB and HU have emerged as a critical player in long-range DNA interactions, it will be worthwhile to compare the effect of the two proteins on global gene expression. No one has yet analyzed the impact of MukB on gene expression.

Future perspective and challenges

A highly resolved and complete structure of the bacterial nucleoid is still not available, although studies of isolated nucleoid began in the 70s. As is known now, the chromosomal DNA is a supercoiled molecule. NAPs organize the DNA through DNA bending, bridging and looping activities. This organization leads to higher-order organizations of DNA into topologically distinct domains, and the large (megabase) size spatial domains or MDs involving the location of ori and ter. NAPs and MD specific proteins MukBEF and MatP appear to collaborate in that. Supercoiling density of individual topological domains created and maintained by the two topoisomerases with opposing functions and transcriptional activities help the spatial organization of the entire chromosome by promoting interactions among distant genomic loci. Supercoiling and topological domain formation also help in proper regulation of gene transcription. The inherent properties of DNA, plectonemic supercoiling, and DNA organization activities of NAPs and MukBEF provide condensation force to sufficiently reduce the volume of the chromosomal DNA to fit within the cell volume, and imparts a hierarchical organization, and the condensed chromosomal DNA results in a functional nucleoid with a helical ellipsoid shape.

Advancements in imaging technologies can pave the way for direct visualization of higher-order nucleoid structure in vivo at high resolution. Advanced electron microscopy methods combining high-pressure freezing and cryo-sectioning to preserve native ultrastructure [167-169], a recently developed approach to enhance DNA signal [170] and 3D reconstruction [171] hold promises in the direction of knowing the 3D arrangement of the entire chromosomal DNA. Although many issues about nucleoid fine structure and functions remain to be resolved, the most curious and exciting are the followings:

  1. Is there a condition dependent 3-D structure of the DNA within the chromosome?
  2. What is the nature of the supercoiling diffusion barriers segregating the nucleoid into independent topological domains?
  3. What are the molecular mechanisms by which distant DNA-DNA contacts are made mainly using the RNA molecule?
  4. Is there a stable attachment of the membrane to DNA?
  5. How are environmental cues transmitted to change the chromosomal structure?


Table 1. E. coli DNA topoisomerases
Topoisomerase Type Function Single- or double-stranded cleavage
Topoisomerase I IA Removes (-) supercoiling SS
Topoisomerase III IA Removes (-) supercoiling SS
Topoisomerase IV IIA Removes (-) supercoiling DS
DNA gyrase IIA Creates (-) supercoiling and removes (+) supercoiling DS

Figure legends

Fig. 1: Formation of a bacterial nucleoid 

A.  A circle with circumference the length of the circular chromosome in E. coli.

B.  A random coil form adopted by the circular chromosome of E. coli at thermal equilibrium.

C.  A 3D volume of nucleoid visualized by HU-mCherry. Blue to red represents increasing DNA density.  Taken from Gall Le et al. 2016.

Fig. 2: Nucleoid at ≥1 kb scale.  Possible DNA structures formed by nucleoid-associated proteins (NAPs).

A.  Flexible bending in DNA induced by binding of one HU heterodimer per 9 bp DNA. Straightening of DNA caused by non-specific binding of HUαβ. DNA networking by lateral multimerization of HUαα on DNA.

B.   Sharp coplanar DNA bending induced by IHF at its specific site.

C.   H-NS spreads on DNA and forms a stiff filament at less than five mM magnesium and induces Inter- and intra-molecular DNA bridging at greater than five mM magnesium.

D.  Coherent DNA bending introduced by binding of Fis at the helically phased binding sites.

Fig. 3: Distribution of nucleoid-associated proteins on the E. coli genome.

Genome-wide DNA binding signal obtained by ChIP-Seq of Fis (red), H-NS (green for growth phase and red for stationary phase), HU (blue), and IHF (orange) in growth and stationary phases in E. coli.  No data are shown for Fis in the stationary phase because it is absent. The figure was prepared in Circos using the data in Kahramanoglou, C. et al. 2011 and Prieto, A., et al. 2012.    

Fig. 4: DNA supercoiling. See the text for details

Note: the figure F will be modified to change the right-handed writhes to the left-handed writhes for the final submission.    

Fig. 5: The chromosomal DNA within the nucleoid is segregated into independent plectonemic loops.

A.   A single topological domain consisting of plectonemes. A single cut anywhere would be sufficient to relax the supercoiling tension of the entire domain.

B.    The nucleoid consists of independent plectonemic loops that are segregated by topological barriers (green spheres represent hypothetical barriers).  A single cut in one loop will only relax that loop and not the others. Estimated number of the loops is ~460, but only four loops with a different number of supercoils are shown for simplicity.  The double-stranded DNA is represented as a line.  

Fig. 6: Twin supercoiling domain model for transcription-induced supercoiling

Helical unwinding during transcription generates two supercoiling domains.  An elongating RNA polymerase complex cannot rotate around the helical axis of DNA and instead forces DNA to rotate.  If DNA is topologically constrained (represented here by shaded bars), it becomes overwound (positively supercoiled) ahead of the complex and underwound (negatively supercoiled) behind it.  Supercoiling is represented here only as a change in the number of twists, but it can also manifest as writhes or both.

Fig. 7: Nucleoid is spatially organized into megabase domains.

A.   On the left is a circular genetic map of E. coli chromosome showing the genomic regions comprising the spatial domains (macrodomains) identified by fluorescence imaging and recombination-based assays.  On the right is a cartoon of an E. coli cell with the condensed nucleoid spatially organized into the domains shown in the circular map on the right. The black dots in the Ori and Ter domains represent oriC and ter respectively.

B.    A scalogram representation of the distribution of 3C contacts made by the chromosomal regions.  The chromosome is divided into 5 kb bins.  The colors represent the fraction of the total cumulated contacts made by a bin with the flanking regions of increasing size (step size of one bin or 5 kb).  The abrupt changes in the signal along the chromosomal regions identify three regions, two loose regions L1 and L2 defined by large blue areas and small red areas and the constrained region encompassing ter defined by small blue and large red areas. The dotted line is the boundary of the three regions in comparison with the macrodomains. Figure B is taken from Lioy VS et al. 2018.  NSR and NSL are NS-right and NS-left respectively.

Fig. 8: Spatial domain specific proteins MatP and MukB and possible mechanism of DNA organization by them.  

A.   Distribution of MatP (green) and MukB (red) on the E. coli chromosome.  The figure was generated in Circos using the processed ChIP-Seq data containing the binding peaks from Nolivos S et al. 2016. 

B.    On the left is the architecture of the E. coli MukBEF complex in the absence of ATP. Two MukE molecules and one MukF molecule form a heterotrimeric complex which then forms a characteristically elongated hexameric structure (MukE2F)2 through MukF dimerization. The complex associates with MukB via interaction between MukF and the cap region of the MukB head.  The ATP binding induces dimerization of the MukB head domains and forces the detachment of one MukF molecule and two MukE molecules associated with it. As a result, a ring is formed that can topologically entrap DNA (on the right).

C.    A stepwise mechanism for DNA condensation by MukB.

D.   A structure of MatP-matS complex (PDB: 4D8J). MatP is in the tetramer form resulting from the oligomerization of the two DNA bound MatP dimers.

E.    A matS-bridging model for DNA condensation in the Ter macrodomain by MatP. The tetramerization of the matS-bound matP dimers bridges the matS sites forming large DNA loops thus condense DNA.

Fig. 9 Nucleoid is a helical ellipsoid

A.   Isosurface model of the nucleoid visualized by HU-mCherry. The image is taken from Fisher JK et al 2012.  Red and green dots are the centroids of individual slices made by cross-sectioning the nucleoid perpendicular to its long axis. 

B.   Z-stack images of the nucleoid visualized by Fis-GFP highlighting the nucleoid curvature. On the right is the calculated writhe of the nucleoid which is equivalent to the centroid path in A. Image is taken from Hadizadeh YN et al. 2012.


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