A model for chromosome organization during the cell cycle in ...as a model system via developed molecular manipulation methods of genes and proteins in live cells. Scientists are attempting

1Scientific RepoRts | 5:17133 | DOI: 10.1038/srep17133

www.nature.com/scientificreports

A model for chromosome organization during the cell cycle in live E. coliYuru Liu, Ping Xie, Pengye Wang, Ming Li, Hui Li, Wei Li & Shuoxing Dou

Bacterial chromosomal DNA is a highly compact nucleoid. The organization of this nucleoid is poorly understood due to limitations in the methods used to monitor the complexities of DNA organization in live bacteria. Here, we report that circular plasmid DNA is auto-packaged into a uniform dual-toroidal-spool conformation in response to mechanical stress stemming from sharp bending and un-winding by atomic force microscopic analysis. The mechanism underlying this phenomenon was deduced with basic physical principles to explain the auto-packaging behaviour of circular DNA. Based on our observations and previous studies, we propose a dynamic model of how chromosomal DNA in E. coli may be organized during a cell division cycle. Next, we test the model by monitoring the development of HNS clusters in live E. coli during a cell cycle. The results were in close agreement with the model. Furthermore, the model accommodates a majority of the thus-far-discovered remarkable features of nucleoids in vivo.

The bacterium is considered a simple life form. Among bacteria, E. coli is the most studied organism as a model system via developed molecular manipulation methods of genes and proteins in live cells. Scientists are attempting to learn more about the mechanisms of maintaining a living organism from E. coli with the goal of understanding how gene transcription is regulated and how a cell reacts to external and internal stimuli1,2. Despite being a simple life form, unexpected patterns of chromosomal DNA compaction and organization within these tiny organisms have been observed2,3. In live E. coli cells supplied with required nutrients, chromosomal DNA is simultaneously amplified and compacted to fit into the confined cytoplasmic volume. E. coli precisely passage their chromosomal DNA and pro-teins through generations in a smooth and elegant manner, underlying an efficient and reproducible large-scale organization1,4.

In bacteria, abundant nucleoid-associated proteins (NAPs) are associated with chromosomes to facilitate the compaction of chromosomal DNA5,6. In E. coli, major NAPs include HNS, HU, Fis, IHF, and StpA7. Each of these NAPs non-specifically binds to DNA and is involved in gene regulation and chromosome compaction8–11. In addition to NAPs, entropy stemming from molecular crowding in a confined intracellular environment contributes to the process of DNA compaction12,13. Theoretical anal-ysis has provided evidence that DNA compaction occurs under the influence of a subtle equilibrium between several competing factors14. With the aid of chromosomal locus labelling and live cell imaging, the amplification and segregation dynamics of chromosomes in live E. coli have been revealed, providing insight into chromosome organization in vivo5. Four macrodomains (MDs) and two less-structured MDs inside the nucleoid have been discovered using homologous recombination rate analysis and fluorescence in situ hybridization (FISH) staining, revealing higher order structural domains in the E. coli chro-mosome15,16. Remarkable features of nucleoids were recently discovered via high spatial and temporal resolution microscopy of live E. coli cells. Two clusters of HNS per copy of E. coli chromosomal DNA were identified by Wang et al., indicating that higher order organization of chromosomal DNA is likely7. Helical ellipsoid nucleoids with two bundles confined by the radial cell periphery were visualized by

Key Laboratory of Soft Matter Physics, Beijing National Laboratory for Condensed Matter Physics, Institute of Physics, Chinese Academy of Sciences, Beijing 100190, China. Correspondence and requests for materials should be addressed to Y.L. (email: [email protected]) or P.W. (email: [email protected])

received: 24 March 2015

accepted: 22 September 2015

Published: 24 November 2015

OPEN

mailto:[email protected]

mailto:[email protected])

www.nature.com/scientificreports/


4D microscopic monitoring of fluorescence-tagged HU proteins, revealing striking nucleoid dynamics during the cell division cycle17. Overall, the organization of chromosomal DNA in live bacteria is quite different from that of eukaryotes (the ‘beads on a string’ conformation) and viral DNA (confined in a tightly compacted capsid). The nucleoid of bacteria is highly compacted but with soft nature to be dynamic accessible to motor proteins18,19.

ResultsAuto-packaging behaviour of plasmid DNA. In our previous study of the effect of platinum drugs on single DNA molecules, the unexpected phenomenon of circular plasmid DNA auto-wrapping in response to transplatin (T) and cisplatin (C) co-ordination was observed20. We next investigated the dynamics and process of plasmid DNA auto-packaging (Fig. 1). Surprisingly, pBR322 DNA molecules froze into a symmetric dual-toroidal-spool conformation after incubation with C and T for several hours, as observed with atomic force microscopic (AFM) analysis. C and T co-ordination (C+T) introduced sharp bends of up to 180° and resulted in the untwisting of local DNA fragments20. However, pBR322 DNA incubated with the same molar concentration of C or T alone did not exhibit this auto-packag-ing behaviour; instead, it mainly formed local micro-loops or bends during the incubation. Given the comparable amount of divalent Pt atoms detected in C and C+T samples by inductively coupled plasma mass spectrometry (ICP-MS), insufficient mechanical stress injected by C alone compared with C+T might underlie the marked difference in DNA conformation between the two groups20. When negative supercoiled plasmid DNA was injected with ~1.3 sharp bends by ~7.6 ± 3.8 (mean ± SD, n = 3) divalent crosslinked Pt atoms per DNA molecule, plasmid DNA began to alter its conformation, and the average writhe number increased to ~3.0 ± 0.1 (mean ± s.e.m., N = 141; 1 h, Fig. 1a,b; intact DNA mean writhe number ~1.9 ± 0.1, mean ± s.e.m., N = 112). Meanwhile, the writhe joints moved from the centre region (0 h) to the periphery (1 h) of the DNA molecules, indicating increased tension inside the DNA mole-cules21 (Supplementary Fig. S1). With the sharp bends increased to ~1.8–2.4 bearing ~11.3–14.7 divalent Pt atoms per DNA molecule, the DNA molecules exhibited writhe numbers as high as ~4.2–5.1 (2 ~ 3 h, Fig. 1a,b). Gradually, the DNA molecules settled into an eight-shaped dual-toroidal-spool conformation and became more tightly compacted, with an average writhe number ~6.1±0.2 (mean±s.e.m., N = 145), during which time divalently linked Pt atoms accumulated up to ~37.6 per DNA molecule (4 h, Fig. 1a,b). The number of sharp bends was calculated as 16 percent of the total Pt atoms divalently crosslinked to DNA, as determined by a combination of ICP-MS and denaturing gel electrophoresis20. The calculated number of sharp bends, and the average increased writhe number counted by AFM (Supplementary Fig. S2) performed on the same DNA sample during the 1–4 h incubation, were coincidently close to each other (Fig. 1b). The highest DNA density was observed in the centre core of the spools by AFM (Supplementary Fig. S1). The rearrangement of intact plasmid DNA into a mature dual-toroidal-spool conformation occurred in five stages, as shown in Fig. 1c and Supplementary Fig. S1. DNA molecules in the dual-toroidal-spool conformation exhibited a symmetric distribution of two parts of DNA content. In each of the spools, DNA was wrapped onto itself, with the ends crossing with each other and looping into the remaining spool.

Mechanism of DNA auto-packaging. By the end of our experiments, over 90% of the DNA mole-cules had uniformly frozen into the dual-toroidal-spool conformation upon sharp bending and unwind-ing stress by C+T. The remaining ~10% of DNA molecules were nicked and retained an open circular conformation. These results demonstrate that the dual-toroidal-spool conformation represents the ener-getically lowest mechanical state and predominates among all the other possible conformations for cir-cular DNA to compensate for sharp bending and untwisting (C+T) mechanical stress in bulk. We next deduced the basic physical principles underlying the predominant conformation. We first analysed the local mechanical effect of external bending factors on a normal B-form DNA fragment (Supplementary Fig. S3). Bending factors exert action F, which is generally perpendicular to the DNA strand and points to bending factors. Thus, DNA will generate a reaction F′ that is opposite to F, according to Newton’s third law. Bended DNA strands generate F′ via their intrinsic tendency to return back to the B-form confor-mation due to the elastic property of helical DNA. Local deformation of DNA strands introduces uneven distributed tension along the DNA backbone. The extra torque mechanical stress continues to propagate along the circular DNA backbone, which cannot be released from the ends as in the case of linear DNA. Macromolecules such as DNA and proteins fluctuate into a lower energetically mechanical state. When the DNA backbone is nearly spherical or ellipsoidal, the deformation of DNA is most evenly distributed across the whole DNA molecule. DNA might then freeze into this configuration (Supplementary Fig. S4a). In the dual-spool conformation, tension along the DNA backbone that is generated inside one spool by the effects of DNA coiling and bending propagates to the entering and exiting ends, where it turns to serve as the supporting force that acts on the resting spool, thereby holding the coiled DNA strands together to prevent them from elastically spreading into B-form DNA (Supplementary Fig. S4b). Meanwhile, the symmetric conformation of the dual-toroidal-spool achieves a self-balanced structure.

The DNA strands are loosely coiled during early stages (0–1 h) and then become more tightly coiled together in each dual-toroidal-spool with increasing injection of C+T mechanical stress (2–4 h Figs 1a and 2a). In turn, writhe formation compensates for untwisting mechanical stress, such that the spools are increasingly compacted to accommodate the generated writhes (i.e., new DNA circles). The DNA



Figure 1. Auto-packaging of plasmid DNA in response to mechanical stress of C and T co-ordination. (a) AFM visualization of plasmid pBR322 DNA molecules gradually packaged into an eight-shaped dual-toroidal-spool conformation. The DNA molecules were incubated with a C and T mixture for ~5 h. Scale bar is 200 nm. (b) Statistical analysis of writhe number per pBR322 DNA molecule in the samples from (a) collected at different incubation times. The amount of divalent bound platinum (Pt) and sharp bends (Sb) per DNA molecule are indicated as the mean ± SD. (c) Distribution of DNA molecule compositions with different conformations during incubation corresponding to (b) under our experimental conditions. The number on horizontal axis refers to stage 0–5. Inset, representative structure of DNA molecules at each stage.



bending energy increases from the periphery (~0.91 kBT per DNA circle) to the centre core (~5.44 kBT per DNA circle) due to the aggravated deformation and shortened radius of DNA circles in the centre of the spool relative to the periphery (2–4 h Fig. 1a and Supplementary Fig. S5).

Dual symmetry was the only observed form of symmetry in the auto-packaged plasmid DNA. Neither triplex symmetry nor quadruplex symmetry were observed. While this is an interesting phenomenon, we cannot address it in the present manuscript. We were only able to calculate the relative probability of dual-spool and multi-spool occurrence inside a circular DNA molecule. The results of this calculation indicated the occurrence of a triplex-spool with N−3 probability (N = all possible distributions of tran-sient forces along a DNA fragment during a period of time) or a quadruplex-spool with N−4 probability to attain mechanically balanced states that allow DNA to freeze, compared with a dual-spool of N−2 probability, in the case of several comparable local toroidal spools formed spontaneously in circular DNA (Supplementary Fig. S6). More practically, a multiple-spool comprised of one circular DNA mol-ecule had an energetically higher mechanical state that fluctuated into a lower mechanical state, such as dual-spool or single-spool. For example, one pBR322 DNA molecule at a length of 4,361 bps compacted into a single-spool with one DNA circle has a total bending energy ~0.695 kBT; if it compacts into

Figure 2. Proposed model for chromosome organization in E. coli. (a) Behaviour of circular plasmid DNA auto-packaging upon sharp bending and unwinding mechanical stress in bulk, as illustrated in a 2D diagram deduced from AFM analysis. (b) Eight-shaped scaffold (green) for the dual-toroidal-spool conformation (left panel). Black arrows indicate tension along the DNA backbone (long arrows) and elastic anti-bending reactions (F′) generated in DNA strands (short arrows). In each spool, DNA wrapped onto itself to form numerous DNA circles (right panel). One half of each DNA circle signifies the entry of DNA strands (green solid lines), while the other half signifies the exit of strands (grey dashed lines). (c) Proposed 3D model of chromosome organization in E. coli. A rotational dual-toroidal-spool (black, adapted from reference 26) with one spool upside down relative to the other was linked by a buckled hinge (blue and red). The hinge region is widened in the image for clarity (see details in Supplementary Figs S7, 8). Each spool retained a counter-clockwise rotational tendency along the long axis relative to the hinge. (d) NAP distribution in nucleoids. Left panel, positions for HNS clusters, as demonstrated previously7. Right panel, only a portion of DNA strands and NAPs were drawn for clarity, illustrating the general dispersed distribution of HU and IHF, and a clustered HNS distribution among nucleoids. (e) Proposed model of dynamic chromosome organization in E. coli during a cell division cycle. DNA strands containing oriC chromosomal loci were pulled from one spool (blue, upper-left panel) to proceed through the replication fork, thereby initiating DNA replication. The newly synthesized DNA collapsed into two new docked spools (purple and yellow, left-bottom panel) during the middle stage of DNA replication. During the late stage of DNA replication, one of the newly docked spools across the remaining parental spool (blue) was disposed at the other half of the cell (right-bottom panel). The docked spools were isolated from replication machinery after the completion of DNA replication and were re-arranged into a dual-toroidal-spool conformation in the ready-to-divide cells (right-upper panel).



dual-spool with one DNA circle in each spool, the total bending energy is ~2.778 kBT, and ~6.255 kBT of the total bending energy for it in triplex-spool with one DNA circle in each spool. A single-spool is an attractive conformation for a circular DNA to freeze into, and it resides in a lower energetic state than the dual-spool. However, circular DNA coiled in one spool configuration could not effectively achieve a self-balanced state (i.e., the coiled spherical DNA strands were not effectively held together by supporting tension). Therefore, the coiled DNA strands underwent outward fluctuation and were constrained by the circular DNA itself. The strands finally settled in a dual-spool self-balanced structure (Supplementary Fig. S9b,c).

Model for E. coli chromosome organization. Plasmid DNA and bacterial chromosomal DNA are similar in that they are negative supercoiled circular DNA of differing sizes. The mechanical effect of C+T co-ordination to IHF and HU proteins in the local DNA backbone is analogous (i.e., both introduce sharp bends and un-winding of DNA)20,22,23. These analogies intrigue us to speculate on the organiza-tion pattern of chromosomal DNA in E. coli. It is notable that the phenomenon from plasmid DNA are not always feasible for chromosomal DNA. Prokaryotic chromosomal DNA has a molecular weight thousands of times that of plasmid DNA, and thus exhibits large topological domains as well as more complicated compaction pathways. Nevertheless, the underlying physical principles might be universal and applicable to different molecules under similar conditions. Thus, we propose a hypothesis that the chromosomal DNA of E. coli is organized into a dual-toroidal-spool conformation (upper right panel in Fig. 2a). The nucleoid had an eight-shaped scaffold with DNA coiled onto itself in each spool to form a buckled balanced dual-toroidal-spool conformation (Fig. 2b). In this model, the centre core of each spool corresponds to the HNS cluster observed by photoactivated localization microscopy (PALM) in vivo7. The dual-toroidal-spool conformation well accommodates the observation that one copy of chromo-somal DNA bearing two HNS clusters symmetrically positioned at cell quarters. Given that the highest DNA density is detected in the centre core, adjacent DNA strands fall within close proximity of each other, so HNS is spatially accessible and can bridge them because force and proximity of the DNA duplex are important for HNS bridging as revealed by single molecular investigations19,24,25. In this model, HNS serves as spacer chaperone to protect DNA from collision (Fig. 2d). Other NAPs (e.g., HU, IHF and Fis) function as bending and unwinding chaperones that inject mechanical stress and accelerate the compac-tion of chromosomal DNA. Mechanical stress injection enhances tension inside DNA molecules. When certain amounts of HU, IHF and Fis are bound to DNA, compaction plateaus and the tension in DNA strands becomes so high that further binding of NAPs is unfavourable. Thus, compaction is equilibrated and stabilized.

Based on our AFM analysis and other theoretical studies, we propose a 3D model for nucleoid organ-ization as a rotational dual-toroidal-spool (Fig. 2c)26. Each spool shares the same essential configuration because they are developed under similar mechanical stress by the same NAPs. One spool faces to one side, while the other spool is approximately upside down relative to the first (Supplementary Fig. S7). The entering and exiting ends of each spool buckle together and serve as a hinge for the nucleoid. As eight-shaped negative DNA supercoils, each spool exhibits a counter-clockwise 3D rotational tendency along the long axis of the nucleoid relative to the hinge (Supplementary Fig. S8a–c). The two spools rotate relative to the hinge in the middle, and are buckled together to obtain a balanced 3D structure. The orbit of the DNA strands in each spool projected on the main plane is similar to the 2D model, as shown in Fig. 2a and Supplementary Fig. S8d. To compensate for the bending tension, the DNA strands in a spool rotate around the orbit to form a rotational toroidal spool ellipsoid in 3 dimensions, as elucidated in a previous theoretical study26.

Based on our observations and other related work, such as the findings that the replication factory is located at the centre of the E. coli cell, that the DNA strands were pulled through the replication machin-ery during DNA replication27,28 and that the oriC and dif loci have distinct cytoplasmic locations16,29,30, we further propose a dynamic model for chromosome organization during a cell division cycle in live E. coli (Fig. 2e). Firstly, the replication factory located at the centre of E. coli cells pulls the DNA strands from one spool containing the oriC chromosomal loci, thereby initiating replication. The newly synthesised DNA strands are dispersed randomly in the cytoplasm, whereas the chromosomal loci in the Ori region are docked on cellular counterparts near cell quarters by MukB proteins and the migS cis element31,32. Upon the binding of NAPs to newly replicated DNA, the actual torsional constrained newborn DNA33,34, collapses into a single-spool docked on replication machinery during DNA replication (Supplementary Fig. S9). Each sister DNA forms one spool. DNA is continually pulled from the un-replicated parental DNA spool. Newborn DNA strands undergo free diffusion and stress from NAPs binding, and they are wrapped onto the docked sister spools (Fig. 2e). Finally, the docked spool is isolated from the replication machinery when DNA replication is completed, and DNA is released from DNA polymerases. Once released, the single-spool, which contains one copy of sister chromosomal DNA, rearranges into the dual-toroidal-spool conformation to obtain a balanced configuration (Supplementary Fig. S9 b,c).

HNS cluster development in live E. coli. To test our hypotheses, we directly monitored HNS cluster development during a cell cycle in E. coli using PALM. Moreover, we checked the model with published data related to nucleoid behaviour and structural features in vivo to analyse the accommodation ability of the model with the experimental data. Details are available in the Supplementary Discussion.



In our model, HNS clusters correspond to the centre core of DNA spools. A packaged DNA spool bore a clear HNS cluster. We tagged HNS protein with a monomeric photoactivatable fluorescent pro-tein, mEos2, as previously described7. The fusion proteins were expressed from their native promot-ers at the chromosomal loci to obtain an expression approximately at the wild-type level. Cells were imaged in M9 minimal medium at room temperature during the early log phase. Time-lapse snapshots at 10–20 min intervals were taken during the cell division cycle, and the images were analysed sequentially. Representative conformations of HNS clusters at different stages of the replication cycle are shown in Fig. 3. We observed two, three and four typical HNS clusters in newborn, growing and ready-to-divide cells, respectively; these findings were consistent with previous observations7. Furthermore, the dynamic developments of two to three clusters and three to four clusters were clearly visualized by monitoring the same cell during a cell cycle, which strongly supports our model of chromosome organization in live E. coli. In brief, one of the two HNS clusters with an average distance of ~0.88 ± 0.1 μ m (mean ± SD, N = 211; Fig. 3b, upper panel) in newborn cells shrank significantly, whereas the other cluster remained essentially constant (ii, Fig. 3a). The shrinking cluster disappeared around 20–30 min after DNA repli-cation and two new additional comparably smaller HNS clusters showed up at the same half of the cells around ~30–40 min during DNA replication (iii Fig. 3a). The two new clusters grew bigger, whereas the other parental cluster shrank (iii, Fig. 3a). At the late stage of DNA replication, the new HNS cluster in the middle switched locations with the shrinking parental cluster (iii, Fig. 3a). Finally, the parental cluster

Figure 3. HNS cluster development visualized by PALM during a cell division cycle in live E. coli. (a) Typical HNS cluster phases during one cell cycle. Two HNS clusters at a defined distance in newborn cells are shown in (i). After DNA replication, one of the HNS clusters shrank and finally disappeared (ii). Two new clusters, which were smaller compared with the parental HNS clusters, appeared during the middle stage of DNA replication (left, iii). The size of the parental cluster decreased with increasing size of the new clusters (middle, iii). Two large HNS clusters were deposited at the two halves of the cell with the shrinking parental HNS cluster in the middle (right, iii). The middle parental HNS cluster disappeared, leaving two comparable HNS clusters in each half of the cell (left, iv). The HNS clusters subsequently became smeared (middle and right, iv). The HNS clusters remained less visible for a period of time (v). Finally, four new HNS clusters appeared in the pre-division cells (vi). The snapshots-taken time points (min) were labelled on each image, corresponding to the normalized cell cycle of ~120 min. (b) The distribution of distance between HNS cluster pairs (N = 211) in newborn and early growing cells with a cell length ~1.7-2.5 μ m (upper panel) and that (N = 310) in middle-aged and old cells with a cell length ~2.6-3.7 μ m (lower panel). (c) Duration of HNS clusters in phases (i–vi) within the cell cycle. The average time of a cell cycle is ~127 ± 25 (mean ± SD, N = 706) min, during which ~15% cells stop growing, reflecting the diversity of cellular repair ability from 405 nm phototoxicity. The different lengths of individual cell cycles were normalized to 120 min, and the corresponding time points were aligned. Error bars indicate s.e.m. Scale bars are 1 μ m.



disappeared and two big clusters remained in a symmetric distribution in the growing cells. The average distance between was ~1.50 ± 0.15 μ m (mean ± SD, N = 310), with two peaks centered around 1.41 μ m and 1.65 μ m (Fig. 3b, lower panel). The two big clusters smeared and disappeared around 60–80 min in a cell cycle (iv–v, Fig. 3a), followed by the appearance of four new clusters around the end of a cells cycle (vi, Fig. 3a). These observations are in agreement with our proposed model. Once the docked spool was released from the replication fork, it re-arranged itself into a dual-toroidal-spool conformation to obtain a stabilized and balanced structure for circular DNA to be frozen into.

The duration times of each phase were recorded, as shown in Fig. 3c, for ~700 E. coli cell division cycles. The average time of a cell cycle is ~127 ± 25 min (mean ± SD, N = 706). The overlapping duration of adjacent phases was observed due to the heterogeneity of cell recovery ability from 405 nm irradiation. There are two phases in which two HNS clusters appeared in one E. coli cell. One is the newborn and early stage of DNA replication (0–20 min during a cell cycle, Fig. 3c); the other is during the late stage of DNA replication and after DNA replication (60–100 min during a cell cycle, Fig. 3c). Statistical meas-urements of the distance between centres of HNS cluster pairs showed a significant difference between the two phases (Fig. 3b). The distance between HNS cluster pairs in newborn and early growing cells was ~0.88 ± 0.1 μ m (mean ± SD, N = 211) with a narrow distribution profile that was constant regardless of cell length ranged from 1.7 μ m to 2.5 μ m, reflecting a relatively solid structure of HNS cluster pairs in these cells. The average distance of ~1.50 ± 0.15 μ m (mean ± SD, N = 310) between HNS cluster pairs has been observed in middle-aged and old cells and shows a broad distribution. Meanwhile, the distance val-ues varied significantly, with the cell body length ranging from 2.6 μ m to 3.7 μ m, probably reflecting the isolated state of sister nucleoids at this time span. Two peaks centred around 1.41 μ m and 1.65 μ m were observed in middle-aged and old cells, respectively. The following scenarios might explain this observa-tion. Before DNA replication is completed, when two sister DNAs are linked via replication forks, the 1.41 μ m peak value might reflect the average distribution of HNS cluster pairs at this stage. After DNA replication is completed, sister DNA is released from the replication fork and thus the distances between might contribute to the 1.65 μ m peak distribution, which is consistent with previous observations that sister DNAs move freely and fit into each cell half after dissociation from replication machinery17.

HNS cluster development was also investigated using the multifork chromosome of E. coli cells cul-tured under an LB-agarose pad (Fig. 4). This experiment revealed symmetrically outward binary growth and division of the parental chromosome into two sister nucleoids, which was consistent with previous investigations35,36. The number of HNS clusters in fast growing cells was typically one, three or two during a cell cycle (Fig. 4c). The HNS clusters were found to be essentially aligned under slow growing condi-tions, and the angles between them were centred around ~180 degree (Fig. 4a,b). However, the three HNS clusters under fast growing conditions had a triangular distribution pattern, with the angles between them ranging from ~40 to 180 degrees (Fig. 4a,b). This observation indicates that the two sister DNA masses are not linear but rather bilobed, which is consistent with previous morphological observations of nucleoids under LB-pads35. The distance between adjacent HNS clusters is ~0.43 ± 0.11 μ m (mean ± SD, N = 255) in new-born cells (Fig. 4d), which is significantly shorter than that observed in slow grow-ing cells (~0.88 μ m). The HNS cluster distance in middle aged and old cells averaged ~0.89 ± 0.30 μ m (mean ± SD, N = 239), with one peak centred at approximately 0.7 μ m for adjacent clusters and another peak centred at approximately 1.37 μ m for distal clusters (Fig. 4e). These observations agree with our model that chromosomal DNA is organized into a single-spool docking on replication forks prior to the completion of DNA replication.

DiscussionThe fundamental mechanism underlying the precise and regular auto-organization of very long chro-mosomal DNA in a living cell remains unknown. Extensive theoretical and experimental studies have reported that NAPs in bacteria, or histones in mammal cells, play an important role in initiating the wrapping of DNA through electrostatic interactions with its phosphate backbone23,37–39, and further result in DNA compaction via protein–protein interactions9,14,40,41. However, in the present study, we found that circular plasmid DNA mechanically stressed by local torque rearranged its conformation into a balanced dual-toroidal-spool conformation (Fig. 1). The accumulation of ~11.7 to 37.6 local torques per 4,361 bps plasmid DNA introduced a global transition of DNA conformation from a random super-coiled structure to an eight-shaped dual-spool configuration over a time scale of 3–5 hours. Theoretically, canonical B-form DNA retained the lowest energetic state, in which minimal mechanical stress exists. When DNA was bent or twisted, it responded as a stiff, springy molecule in conformational flux until a new low energetic state was achieved. The auto-packaging behaviour of plasmid DNA in response to torque mechanical stress from small molecules, such as the platinum complex, was reported and inves-tigated for the first time in the present study. These findings could provide new clues for understanding the mystery of DNA auto-compaction in bacterial cells.

Bacterial chromosomal DNA is highly compacted into a nucleoid. Nucleoid DNA is considered a highly mechanically stressed object whose intrinsic mechanical features play governing roles17,42. The findings of our PALM studies strongly support this opinion (Figs 3 and 4). Time sequential evolutions of HNS clusters both in slow and fast growing E. coli cells demonstrate their unique regular patterns during cell cycles, indicating the existence of unknown factors which govern these evolutions. In particu-lar, characteristic HNS cluster pairs at a relative constant distance (~0.88 μ m) were readily observed in



new born cells and early growing cells (0–30 min) under slow growing conditions. Over this time span, the cells are known to contain approximately one copy of chromosomal DNA. Subsequently, these HNS pairs developed into three clusters and then became two new clusters again during the late stage of DNA replication and after DNA replication (60–100 min). The new HNS pairs have a significantly different distance value (i.e., 1.41 to 1.65 μ m). The cells over this time span are known to contain approximately two copies of chromosomal DNA. The specific “two-three-two-four” mode of HNS cluster evolution during a cell cycle under slow growing conditions, and the “one-three-two” mode of HNS cluster devel-opment within a cell division cycle in cells under fast growing conditions, are observed regularly in growing E. coli cells. These experimental data agree well with our model of chromosome organization in

Figure 4. HNS cluster development pattern in a multifork E. coli chromosome. (a) The distribution pattern of three HNS clusters in cells under slow growing conditions (M9 minimal media) as shown in upper panel, is different from that of cells under fast growing conditions (LB-agarose pad, lower panel). (b) The distribution of angles θ as indicated in (a) under fast (N = 238) or slow (N = 133) growing conditions. (c) HNS clusters in a multifork chromosome developed through a ‘one-three-two’ mode over a cell cycle. The representative phase of HNS clusters in E. coli cells growing under an LB-agarose pad is shown as i to iv for an amplification cycle. i, pre-division cells; ii, newborn cells; iii, growing cells; iv, pre-division cells. (d) Distribution of distance between adjacent HNS clusters (N = 255) in newborn and early growing cells under an LB-agarose pad. (e) Distribution of distance between adjacent HNS clusters (N = 239) in late growth and pre-division cells growing under LB-agarose pad. (f) Proposed model for the organization pattern of multifork chromosomal DNA corresponding to (c). Scale bars are 500 nm.



E. coli. Based on AFM single molecular observations and PALM evaluation, we have explained a novel organization pattern for circular chromosomal DNA in living E. coli cells (Fig. 2). However, due to the limitations of experimental techniques, we cannot prove our model by direct observation of circular DNA compaction by NAPs in vivo and/or in vitro. We leave this problem open for future investigations to discover more about the mechanisms underlying chromosomal DNA compaction. From the findings in the current study and previous reports, we propose that NAP binding introduces mechanical stress to chromosomal DNA such that stressed chromosomal DNA rearranges its conformation into one docked spool conformation (Fig. 4) or a dual-toroidal-spool conformation (Fig. 3, i and vi) depending on the isolate state of DNA. Our findings indicate that the unique intrinsic physical properties of DNA might be the determining factor underlying auto-packaging behaviours of these giant biomolecules in cells.

We tested the ability of our model to accommodate previously published experimental data. It has been difficult to directly observe the compaction pattern of DNA in a nucleoid due to molecular crowd-ing in live bacteria and the loss of the original morphology of the nucleoid after cell lysis17,43. However, in recent years, a number of well-designed investigations have provided insight into the dynamic organ-ization of the nucleoid in living E. coli2. Surprisingly, a majority of these experimental results support our model for dynamic chromosomal organization in E. coli during a division cycle. For example, six macrodomains representing a higher order of nucleoid organization coincided with the temporal and spatial distribution of DNA strands at significantly different fluctuating states, as described by our model (Supplementary Discussion)15,44. The distribution pattern of pairs of chromosomal loci was easily explained by the dual-toroidal-spool configuration45,46. Our model accommodates the possibility that one copy of chromosomal DNA bears two HNS clusters exactly located at cell quarters7, as well as the asymmetry of the newly split sister nucleoid42,47. The 3D model of the rotational dual-toroidal-spool with an upside down relationship between the two spools explains the helicity of the nucleoid without apparent handedness (Supplementary Discussion and Supplementary Fig. S7)17,35. Our findings prove that sister nucleoids are basically non-intermingling objects that are separated from each other in a single coordinated transition with global chromosomal movements17,42,47. Overall, our observations provide the direct experimental proof of the auto-packaging behavior of circular plasmid DNA upon unwinding and bending mechanical stress by the platinum complex. The proposed model for dynamic compaction of the nucleoid may expand our understanding of the mechanisms and pathway of chromosome organization in E. coli. The present study provides new perspectives for investigating fundamental mechanisms of chromosome organization and gene regulation in bacteria and eukaryotes in the future.

Finally, let us forget the impression that DNA is a loosely relaxed string with local bends by proteins.Let us appreciate DNA as an elastic, rigid (in nanometer-scale), fluctuant biomolecule which tends to

auto-package onto itself with the aid of chaperone NAPs.Though no special physical or chemical laws in cells, we cannot help wondering, adoring and admir-

ing how well organized a cell can be.

Methods summary. Atomic force microscopy analysis was performed essentially the same as described previously20. Bacterial strain construction of the chromosomal fusion HNS-mEos2 MG1655 strain was performed according to previous methods7,48. Detailed procedures for imaging and data pro-cessing are available in the Supplementary Methods.

References1. Lovett, S. T. & Segall, A. M. New views of the bacterial chromosome. EMBO Rep 5, 860–864 (2004).2. Ptacin, J. L. & Shapiro, L. Chromosome architecture is a key element of bacterial cellular organization. Cell Microbiol 15, 45–52

(2013).3. Toro, E. & Shapiro, L. Bacterial chromosome organization and segregation. Cold Spring Harb Perspect Biol 2, a000349 (2010).4. Wiggins, P. A., Cheveralls, K. C., Martin, J. S., Lintner, R. & Kondev, J. Strong intranucleoid interactions organize the Escherichia

coli chromosome into a nucleoid filament. Proc Natl Acad Sci USA 107, 4991–4995 (2010).5. Reyes-Lamothe, R., Wang, X. & Sherratt, D. Escherichia coli and its chromosome. Trends Microbiol 16, 238–245 (2008).6. Luijsterburg, M. S., Noom, M. C., Wuite, G. J. & Dame, R. T. The architectural role of nucleoid-associated proteins in the

organization of bacterial chromatin: a molecular perspective. J Struct Biol 156, 262–272 (2006).7. Wang, W., Li, G. W., Chen, C., Xie, X. S. & Zhuang, X. Chromosome organization by a nucleoid-associated protein in live

bacteria. Science 333, 1445–1449 (2011).8. Dorman, C. J. H-NS: a universal regulator for a dynamic genome. Nat Rev Microbiol 2, 391–400 (2004).9. Browning, D. F., Grainger, D. C. & Busby, S. J. Effects of nucleoid-associated proteins on bacterial chromosome structure and

gene expression. Curr Opin Microbiol 13, 773–780 (2010).10. Johansson, J. et al. Nucleoid proteins stimulate stringently controlled bacterial promoters: a link between the cAMP-CRP and

the (p)ppGpp regulons in Escherichia coli. Cell 102, 475–485 (2000).11. Dorman, C. J. Co-operative roles for DNA supercoiling and nucleoid-associated proteins in the regulation of bacterial

transcription. Biochem Soc Trans 41, 542–547 (2013).12. Marenduzzo, D., Micheletti, C. & Cook, P. R. Entropy-driven genome organization. Biophys J 90, 3712–3721 (2006).13. Jun, S. & Mulder, B. Entropy-driven spatial organization of highly confined polymers: lessons for the bacterial chromosome. Proc

Natl Acad Sci USA 103, 12388–12393 (2006).14. Joyeux, M. & Vreede, J. A model of H-NS mediated compaction of bacterial DNA. Biophys J 104, 1615–1622 (2013).15. Valens, M., Penaud, S., Rossignol, M., Cornet, F. & Boccard, F. Macrodomain organization of the Escherichia coli chromosome.

EMBO J 23, 4330–4341 (2004).16. Niki, H., Yamaichi, Y. & Hiraga, S. Dynamic organization of chromosomal DNA in Escherichia coli. Genes Dev 14, 212–223

(2000).


1 0Scientific RepoRts | 5:17133 | DOI: 10.1038/srep17133

17. Fisher, J. K. et al. Four-dimensional imaging of E. coli nucleoid organization and dynamics in living cells. Cell 153, 882–895 (2013).

18. Pelletier, J. et al. Physical manipulation of the Escherichia coli chromosome reveals its soft nature. Proc Natl Acad Sci USA 109, E2649–2656 (2012).

19. Dame, R. T., Noom, M. C. & Wuite, G. J. Bacterial chromatin organization by H-NS protein unravelled using dual DNA manipulation. Nature 444, 387–390 (2006).

20. Liu, Y. R. et al. Transplatin enhances effect of cisplatin on both single DNA molecules and live tumor cells. Archives of Biochemistry and Biophysics 536, 12–24 (2013).

21. Sankararaman, S. & Marko, J. F. Formation of loops in DNA under tension. Phys Rev E Stat Nonlin Soft Matter Phys 71, 021911 (2005).

22. Swinger, K. K. & Rice, P. A. IHF and HU: flexible architects of bent DNA. Curr Opin Struct Biol 14, 28–35 (2004).23. Rice, P. A. & Making DNA do a U-turn: IHF and related proteins. Curr Opin Struc Biol 7, 86–93 (1997).24. Dame, R. T., Wyman, C. & Goosen, N. H-NS mediated compaction of DNA visualized by atomic force microscopy. Nucleic Acids

Res 28, 3504–3510 (2000).25. Liu, Y., Chen, H., Kenney, L. J. & Yan, J. A divalent switch drives H-NS/DNA-binding conformations between stiffening and

bridging modes. Genes Dev 24, 339–344 (2010).26. Kulic, I. M., Andrienko, D. & Deserno, M. Twist-bend instability for toroidal DNA condensates. Europhys Lett 67, 418–424

(2004).27. Uphoff, S., Reyes-Lamothe, R., Garza de Leon, F., Sherratt, D. J. & Kapanidis, A. N. Single-molecule DNA repair in live bacteria.

Proc Natl Acad Sci USA 110, 8063–8068 (2013).28. Reyes-Lamothe, R., Sherratt, D. J. & Leake, M. C. Stoichiometry and architecture of active DNA replication machinery in

Escherichia coli. Science 328, 498–501 (2010).29. Lau, I. F. et al. Spatial and temporal organization of replicating Escherichia coli chromosomes. Mol Microbiol 49, 731–743 (2003).30. Mercier, R. et al. The MatP/matS site-specific system organizes the terminus region of the E. coli chromosome into a macrodomain.

Cell 135, 475–485 (2008).31. Danilova, O., Reyes-Lamothe, R., Pinskaya, M., Sherratt, D. & Possoz, C. MukB colocalizes with the oriC region and is required

for organization of the two Escherichia coli chromosome arms into separate cell halves. Mol Microbiol 65, 1485–1492 (2007).32. Yamaichi, Y. & Niki, H. migS, a cis-acting site that affects bipolar positioning of oriC on the Escherichia coli chromosome. EMBO

J 23, 221–233 (2004).33. Moulin, L., Rahmouni, A. R. & Boccard, F. Topological insulators inhibit diffusion of transcription-induced positive supercoils

in the chromosome of Escherichia coli. Mol Microbiol 55, 601–610 (2005).34. Postow, L., Hardy, C. D., Arsuaga, J. & Cozzarelli, N. R. Topological domain structure of the Escherichia coli chromosome. Genes

Dev 18, 1766–1779 (2004).35. Hadizadeh Yazdi, N., Guet, C. C., Johnson, R. C. & Marko, J. F. Variation of the folding and dynamics of the Escherichia coli

chromosome with growth conditions. Mol Microbiol 86, 1318–1333 (2012).36. Youngren, B., Nielsen, H. J., Jun, S. & Austin, S. The multifork Escherichia coli chromosome is a self-duplicating and self-

segregating thermodynamic ring polymer. Genes Dev 28, 71–84 (2014).37. Bloch, V. et al. The H-NS dimerization domain defines a new fold contributing to DNA recognition. Nat Struct Biol 10, 212–218

(2003).38. Suto, R. K., Clarkson, M. J., Tremethick, D. J. & Luger, K. Crystal structure of a nucleosome core particle containing the variant

histone H2A.Z. Nat Struct Biol 7, 1121–1124 (2000).39. Bhowmick, T. et al. Targeting Mycobacterium tuberculosis nucleoid-associated protein HU with structure-based inhibitors. Nat

Commun 5, 4124 (2014).40. Arold, S. T., Leonard, P. G., Parkinson, G. N. & Ladbury, J. E. H-NS forms a superhelical protein scaffold for DNA condensation.

Proc Natl Acad Sci USA 107, 15728–15732 (2010).41. Thanbichler, M., Viollier, P. H. & Shapiro, L. The structure and function of the bacterial chromosome. Curr Opin Genet Dev 15,

153–162 (2005).42. Joshi, M. C. et al. Escherichia coli sister chromosome separation includes an abrupt global transition with concomitant release

of late-splitting intersister snaps. Proc Natl Acad Sci USA 108, 2765–2770 (2011).43. Thacker, V. V. et al. Bacterial nucleoid structure probed by active drag and resistive pulse sensing. Integr Biol (Camb) 6, 184–191

(2014).44. Espeli, O., Mercier, R. & Boccard, F. DNA dynamics vary according to macrodomain topography in the E. coli chromosome. Mol

Microbiol 68, 1418–1427 (2008).45. Wang, X., Liu, X., Possoz, C. & Sherratt, D. J. The two Escherichia coli chromosome arms locate to separate cell halves. Genes

Dev 20, 1727–1731 (2006).46. Nielsen, H. J., Ottesen, J. R., Youngren, B., Austin, S. J. & Hansen, F. G. The Escherichia coli chromosome is organized with the

left and right chromosome arms in separate cell halves. Mol Microbiol 62, 331–338 (2006).47. Bates, D. & Kleckner, N. Chromosome and replisome dynamics in E. coli: loss of sister cohesion triggers global chromosome

movement and mediates chromosome segregation. Cell 121, 899–911 (2005).48. Datsenko, K. A. & Wanner, B. L. One-step inactivation of chromosomal genes in Escherichia coli K-12 using PCR products. Proc

Natl Acad Sci USA 97, 6640–6645 (2000).

AcknowledgementsWe are grateful to Prof. H.L. Shi of Institute of Theoretical Physics, Chinese Academy of Science for kindly providing the bacteria strains and plasmids. We thank Dr. Q. Zhang for strains and constructs, and Y.L. Zhai for technical assistance. This work was supported by the National Natural Science Foundation of China (Grants 11204363, 11274374 and 61275192) and the National Basic Research Program of China (973 Program) (Grants 2013CB837200).

Author ContributionsM.L. and P.Y.W. supervised the project. Y.R.L., M.L. and P.Y.W. designed experiments. Y.R.L. performed the AFM, HNS construction and imaging experiments, and analysed AFM and HNS data. H.L. carried out the setup of imaging instruments and analysed the data. P.X., M.L., W.L., S.X.D. and P.Y.W. analysed the data and discussed the manuscript. Y.R.L. wrote the paper.


1 1Scientific RepoRts | 5:17133 | DOI: 10.1038/srep17133

Additional InformationSupplementary information accompanies this paper at http://www.nature.com/srepCompeting financial interests: The authors declare no competing financial interests.How to cite this article: Liu, Y. et al. A model for chromosome organization during the cell cycle in live E. coli. Sci. Rep. 5, 17133; doi: 10.1038/srep17133 (2015).

This work is licensed under a Creative Commons Attribution 4.0 International License. The images or other third party material in this article are included in the article’s Creative Com-

mons license, unless indicated otherwise in the credit line; if the material is not included under the Creative Commons license, users will need to obtain permission from the license holder to reproduce the material. To view a copy of this license, visit http://creativecommons.org/licenses/by/4.0/

http://www.nature.com/srep

http://creativecommons.org/licenses/by/4.0/

Supplementary Information

A model for chromosome organization during the cell cycle in live E. coli

Yuru Liu*, Ping Xie, Pengye Wang*, Ming Li, Hui Li, Wei Li, Shuoxing Dou

Key Laboratory of Soft Matter Physics, Beijing National Laboratory for Condensed Matter

Physics, Institute of Physics, Chinese Academy of Sciences, Beijing 100190, China

Supplementary Figures

Supplementary Fig. S1. Montage of AFM images of DNA molecules in different stages of

the formation of dual-toroidal-spool conformation. The development of intact supercoiled

plasmid DNA to dual-toroidal-spool conformation involves five stages in bulk. stg.0, Intact

plasmid DNA; stg.1, local small toroidal supercoils formed in DNA; stg.2, local irregular

toroidal spools with different sizes; stg.3, loose two toroidal spools with roughly comparable

engulfment of DNA contents; stg.4, compacted dual-toroidal-spool with symmetric

conformation; stg.5, tightly wrapped dual-toroidal-spool with a condensed centre core. The

images are 0.5 μm in width.

Supplementary Fig. S2. Identification of writhe information on DNA molecules. The

writhe is defined as crossing of two DNA fragments in our study. The AFM images of DNA

molecules (i.e., DNA 1 to 4, at different stages of DNA compaction) were enlarged and the

DNA backbone was tracked. The writhe points where two DNA fragments cross each other

were recorded by red circles. Scale bars are 100 nm.

Supplementary Fig. S3. Diagram of DNA response to bending mechanical stress. a, The

canonical B-form DNA has the lowest energetically mechanical state in which minimal

mechanical stress exists (i). When a normal B-form DNA was bent by external bending

factors, bending factors generated action F (brown arrow, iii), which was overall

perpendicular to DNA strands and pointed to the side of bending factors; correspondingly,

deformed DNA generated reactions F ́(black arrow, iv) with an opposite direction to F. b, F

and F ́ increased with increasing DNA bending. The bent DNA demonstrated a tendency to

return back to B-form DNA because of the intrinsic elastic property of helical DNA.

Supplementary Fig. S4. Diagram of the mechanism for circular DNA to compensate

sharp bending and untwisting mechanical stress.

a, Diagram to explain the mechanism for the formation of DNA spools in mechanically

stressed circular DNA. When DNA was bent or untwisted by binding factors, local strands

were deformed. Thus, the deformed part was mechanically stressed to obtain a higher

energetic state. The uneven distributed tension and mechanical stress along the DNA

backbone resulted in fluctuation. Fluctuation continued to propagate along the DNA backbone,

and additional twisting mechanical stress could not be released from the ends of DNA as

linear DNA does. When nearly spherical or ellipsoid curvature of DNA strands appeared,

deformation was most evenly distributed along DNA, so fluctuation decreased and DNA was

likely to be frozen into this conformation. Therefore, mechanically stressed DNA fell into a

new lowest energetic state in which fluctuation along the DNA chain was mostly reduced by

the self-balancing effect and evenly distributed curvature of DNA. For a circular DNA, a

DNA spool was a more stable structure than local toroidal supercoils (inside the grey solid

line box).

b, Diagram illustrating the coiling process of DNA spool in mechanically stressed circular

DNA. The increasing untwisting effect by external factors on circular DNA continued to

disturb DNA. DNA compensated for these effects by increasing its writhe number. Thus,

DNA became more tightly compact to accommodate the increasing writhes, i.e., new DNA

circles. MS injection means mechanical stress injection. F is the tension along DNA, which

increased with the accumulation of torque injections. F serves as the supporting force to act

on the remaining spool.

Supplementary Fig. S5. Diagram of DNA coiling onto itself to accommodate new writhes

upon mechanical stress injection. New writhes accompanied with new DNA circles were

introduced into the DNA spool to compensate for untwisting mechanical stress. In the lower

panel, the new circles of DNA are drawn in different colours for clarity. One half of DNA

circle is entering DNA strand and the other half is exiting DNA strand. The results of AFM

analysis show that the circles became significantly smaller from the periphery to the centre of

the spool. In this case, the bending energy of each circle increased from the periphery to the

centre core of the spool.

To estimate the bending energy of each DNA circle, linear elastic theory of a linear spring

was applied (as described in a teaching program in Massachusetts Institute of Technology,

http://www.mit.edu/~kardar/teaching/projects/dna_packing_website/bending.html).

The bending energy is of the form E = 1/2 k x2,

The energy to bend the DNA chain is of the following form,

E =1

2𝑘𝐿 (

𝑑𝑡

𝑑𝑠)

2

where dt/ds is the derivative of the tangent vector to each simulated DNA Kuhn segment of the

DNA chain. We simplified each DNA circle being bent into a circle with radius R, dt/ds = 1/R. L is

the length of DNA. Thus, the bending energy of each DNA circle is expressed as follows,

E =1

2𝑘𝐿 𝑅2⁄ =

1

2𝑘(2𝜋𝑅) 𝑅2⁄ = 𝑘𝜋

𝑅⁄

where k is ξPkBT and ξp is the persistent length of DNA. The bending energy increases with the

shortening of radius of DNA circles.

Supplementary Fig. S6. Diagram of the mechanism for dual symmetry predomination in

mechanically stressed circular DNA molecules. a, If three or more comparable local DNA

spools (green circles) formed spontaneously inside a circular DNA, whether faced to one side

(i, iii) or both sides (ii, iv), the odds for occurrence of a moment when the tension along all

the hinge DNA strands (blue lines) reaches an exactly balanced state would be very rare. Thus,

significant fluctuation continuously propagated inside the circular DNA molecules until a

balanced structure formed.

b, In a fluctuating DNA molecule, the tension F1 generated from one developing spool acting

on the hinge part dynamically changed its direction and value, similar to F2. In a simplified

estimation, if we assume that the possible distribution pattern for F1 and F2 pair is N, then the

balanced state is unique (where F1 = F2 with exact opposite directions between F1 and F2).

The odds of occurrence for one DNA section at the hinge part in a balanced state is:

CN1 = 1/N = N−1.

Thus, the odds for a balanced dual-spool occurrence is as follows: Rd = CN1 × CN

1 = N−2,

where the two crossing DNA sections are required to reach a balanced state simultaneously.

The odds for a balanced triplex-spool occurrence is as follows: Rt = CN1 × CN

1 × CN1 = N−3;

the odds for a balanced quadruplex-spool occurrence is: Rq = CN1 × CN

1 × CN1 × CN

1 = N−4.

In fact, Rt and Rq may even be lower than the estimation above because the spontaneous

occurrence of comparable triplex-spool or quadruplex-spool is rarer than dual-spool in

solution.

Supplementary Fig. S7. Diagram of 3D spatial relationship between two spools in

dual-toroidal-spool conformation. a, When circular DNA was bent and untwisted by NAPs,

the locally developed spools shared similar structural features (i). This phenomenon was due

to the same mechanical effects introduced by the same types of NAPs into different parts of

circular DNA. Each spool (green, i) bore similar handedness, and both were negative

supercoiled toroidal spools because of the untwisting effects of NAPs. In reality,

conformation (i) does not exist. Given the intrinsic rotational tendency of negative supercoils

of DNA, conformation (i) rotates into conformation (ii) with a crossover hinge part. b,

Therefore, the two spools inside a dual-toroidal-spool retained a substantially upside down

spatial 3D relationship between them. The image of the DNA spool (black) was copied and

adapted from reference [1].

Supplementary Fig. S8. Diagram for structural details of dual-toroidal-spool

conformation. a, Two DNA ellipsoids connected with a hinge was basically the 3D

morphology for the dual-toroidal-spool conformation. Each spool had the same 3D rotational

tendency along the long axis to the hinge (blue arrows), and shared the same structural

features. b, Structural features of the hinge of the dual-toroidal-spool conformation. They

crossed over with each other but were not intertwined. Each spool rotated on the hinge in the

middle between them, buckling the two DNA strands (blue and red) of the hinge together.

Left panel, the projection of the hinge onto the x–y plane. Right panel, the projection of the

hinge onto the x–z plane. c, The dual-toroidal-spool is essentially an eight-shaped negative

supercoil. In negative supercoils of circular DNA, DNA strands retain right-handed rotational

tendency. Therefore, each spool in (a) has a counterclockwise rotational tendency along the

long axis to the hinge. The DNA image in the left panel was adapted from reference [2]. d, 3D

structural feature of the DNA spool. DNA strands in each spool are wrapped with almost

evenly distributed curvature, with the projection of DNA strands onto the x–y plane (left)

serving as an orbit (middle). In 3D solution, the DNA strands in spool rotated along the orbit

to compensate for the bending stress to form an ellipsoid rotational spool (right), as elucidated

by a previous theoretical investigation [1]. The dashed line indicates linking to the hinge (left

and middle).

Supplementary Fig. S9. Diagram of mechanism for docked spool formation during cell

division cycle. a, Newly synthesised DNA (blue) docked on two replication forks was bound

by NAPs, which are abundant in the cytoplasm of E. coli. The bindings of NAPs and other

replication-associated proteins introduced mechanical stress and torsional constraints into

replicating sister DNA, so the mechanically stressed sister DNA auto-wrapped into a

single-spool conformation held by replication factories and achieved the lowest energetic

mechanical state (green).

b and c, The underlying mechanism is that the single-spool is in a lower energetic state than

the dual-spool when both of them developed from the same DNA molecule. Considering that

the overall radius of DNA circles distributed in a single-spool is bigger than that in a

dual-spool, the total bending energy of DNA single-spool is lower than that of DNA

dual-spool. Therefore, a single-spool is more energetically favourable than multiple-spools for

DNA to settle down. In a single-spool, the holding force either from replication factories or

other comparable spools provided an external force to hold itself in shape. Without a holding

force, the isolated single-spool spread outwards because of the intrinsic elastic property of

helical DNA; thus, the spool dissembled. Finally, a dual-spool formed because of the

self-balancing effect for governing DNA to fluctuate into the lowest energetic state. Although

a dual-spool has a higher energetic state than a single-spool, it is the lowest energetic structure

among other possible conformations of mechanically stressed circular DNA (except

single-spool). Dual-spool demonstrates good self-balance, so it is most favourable for an

isolated circular DNA to be frozen into among other possibilities as mentioned in

Supplementary Fig. S6.

Supplementary discussion

The organization of nucleoids in live E. coli remains largely unknown even

though E. coli has been used as a classical model organism under extensive

investigations by various molecular genetic approaches for the past decades. With the

application of new techniques available in recent years, our understanding of the

features of nucleoids has greatly improved [3-5]. Although chromosome DNA in

bacteria is highly compacted to fit into a confined cytoplasmic volume, nucleoids

possess a soft nature that differs from viral DNA tightly compacted in a capsid [6].

Nucleoids do not always fill out all the cytoplasmic space, and they move as a solid

without changing much of their shape and volume inside the cytoplasm [7, 8]. The

nucleoid can even retain part of their compactness over a time scale of tens of minutes

after they are isolated from lysed cells [6, 9]. These observations suggest a soft but

solid nucleoid, which is rather in a highly organized form than in random coils.

E. coli lacks a mitotic system and an equivalent prokaryotic mechanism to

separate sister chromosomes. There is no defined special cytoskeletal apparatus or

docking proteins on the inner membrane to mediate the segregation of sister DNA [10,

11]. Bates et al. have shown that loss of the adhesion of sister nucleoid triggers global

chromosome movements and mediates chromosome segregation, demonstrating a

unique feature for bacterial sister chromosome segregation [12]. In the present study,

we proposed a model to elucidate organization and segregation of chromosome DNA

in live E. coli during a cell cycle. The model accommodates the thus-far-discovered

remarkable features of nucleoids in vivo. We focus the discussion in the following six

sections.

1. Six MDs of nucleoids display their particular spatial and temporal

distribution of DNA contents with varied mobility during cell cycles.

Higher order organization of bacterial chromosome is poorly understood. The

organization of the chromosome into MDs was first discovered by Niki and

colleagues based on FISH observations [13]. The MDs Ori and Ter were defined as

the 1 Mb regions flanking the oriC or dif chromosomal locus. All the chromosomal

loci in Ori MD or Ter MD have their respective unique localisations, which are

distinct from the rest of the chromosome. Four MDs and two less-structured regions

were further observed by analyzing the recombination efficiency using a site-specific

recombination system [14]. Two of the MDs defined by recombination efficiency,

which were similar to the Ter and Ori MDs, were Right and Left MDs, respectively.

Two less-structured regions flanking the Ori MD were named NSR and NSL MDs

(Supplementary Discussion Diagram 1a). The recombination efficiency in NSR and

NSL MDs was much lower than that in the other four MDs.

Supplementary Discussion Diagram 1. Mobility features of DNA strands in six MDs elucidated by

our model. (a) Map of MD distribution on E. coli chromosome according to a previous description [14]

and the time points selected in a cell cycle. The green region containing oriC and migS is Ori MD. The

black regions flanking Ori MD are NSL and NSR MDs. The blue region is Ter MD, and it is flanked by

Right (purple) and Left MDs (orange). (b) In our model, the migS locus was amplified at T1 and then

docked onto cell quarters. The ~1 MB DNA flanking this locus was also docked onto cell quarters [15, 16].

The NS MDs underwent free diffusion for a significant time span, and they were randomly disposed in the

cytoplasm (from their replication to ~T3). At ~T3, thus-far-replicated sister DNA collapsed into a DNA

spool docked on replication factories. From ~T3–T6, newly amplified DNA strands briefly underwent free

diffusion, and wrapped on the developing spools where fluctuation was significant. At T7, the maturation

of dual-toroidal-spool reduced fluctuation by achieving a self-balanced and low energetic mechanical

state. (c) The fluctuation state of DNA from each MD during a cell cycle was scored into four levels. Level

i, constraint by cytoplasmic components; level ii, fluctuated in a mature DNA spool; level iii, fluctuated in a

developing DNA spool; and level iv, free diffusion.

The cytological and molecular mechanisms underlying MD development are

currently unknown. Based on the observations in the present study, the spatial and

temporal distribution and organization of chromosome DNA during a cell cycle were

proposed (Supplementary Discussion Diagram 1b). In brief, each DNA spool

occupied half of an E. coli cell. When replication of chromosome DNA was initiated,

the oriC locus was first pulled to pass through the replication factory located at the

centre of the cell. Newly synthesised oriC and the loci in Ori MD were then tethered

and disposed at cell quarters by MukB protein and migS cis element, as revealed

previously [15, 16]. Thus, Ori MD was diffusion constraint. The replicated DNA

region outside Ori MD in the spool underwent free diffusion for a significant time

until the formation of two additional new spools at around the middle stage of DNA

replication. These DNA regions corresponded to the NSL and NSR MDs flanking Ori

MD. The DNA contents located in the Right and Left MDs were replicated. The

newly amplified DNA fragments from the Right and Left MDs underwent a brief

period of free diffusion, and wrapped onto the already formed docked spools. Thus,

free diffusion was minimal for Ori, Right, Left, and Ter MDs, but significant for NS

MDs.

During a cell division cycle, DNA fragments in nucleoids are mostly in a dynamic

fluctuation state at different levels for a special time span (Supplementary Discussion

Diagram 1c). We scored the fluctuation state based on four levels. The lowest level (i)

is diffusion constraint by the docking system to associate with other cytoplasmic

components. The second level (ii) involves DNA fluctuation on mature

dual-toroidal-spool without local docking systems. The dual-toroidal-spool is a

well-balanced conformation in the lowest energetic state, which allows DNA to freeze

to stop significant fluctuation. In the third level (iii), DNA fluctuates on a newly

formed, dynamically fluctuating, non-balanced spool. The highest level (iv) is free

diffusion. DNA in this level is not packaged yet. In particular, NS MDs spend almost

the entire cell cycle to achieve a higher fluctuation state (iii and iv). They are the only

MDs that have to freely diffuse for a significant time during a cell cycle. This feature

might explain why they are regarded as less-structured regions in previous

observations. By contrast, Ori and Ter MDs are in the lowest fluctuation state (i) for

almost the entire cell cycle by associating with MukB or MatP, respectively, and they

are docked on defined cytoplasmic positions [16, 17]. Right and Left MDs are in a

relative median state of fluctuation (ii and iii) in which DNA is organized into mature

or developing spool. Our model was in agreement with the results of a previous study

presenting the diffusion speeds and travel distance of chromosomal loci in each MD

in E. coli [18] (Supplementary Discussion Published Data 1). The highest mobility

and greatest travel distance were observed for the loci in NS MDs in their study.

The DNA fluctuation state coincident with homologous recombination efficiency

would be an interesting issue to be further addressed for deeper understanding of

mechanism for recombination.

Supplementary Discussion Published Data 1. Mobility features of DNA strands in six MDs [18]. (a)

Fluorescent chromosomal loci distribution during a cell cycle. NSR-1 exhibited the longest travel distance

in the cytoplasm among the four chromosomal loci. (b) The travel distance of chromosomal loci in

different MDs. (c) The diffusion speeds of chromosomal loci in different MDs.

The distribution of MDs on the chromosome map presents several geometric

features. The detailed distribution lengths and positions of MDs on the chromosome

map indicate the subtle relationship between MDs and DNA replication

(Supplementary Discussion Diagram 2). Firstly, two NS MDs had different sizes and

Ori MD was not centred on the oriC locus. Meanwhile, the lengths of DNA fragments

from oriC to the ends of each NS MDs were largely comparable (O-N1 and O-N2

fragments, Supplementary Discussion Diagram 2). This geometric feature might

reflect the sudden collapse of newly synthesised sister DNA into more stable

conformations, thereby preventing the free diffusion of DNA contents replicated after

NS MDs to end a ‘non-structured’ state of those DNA parts. Considering the

replication initiating from oriC and preceding bidirectional with the same speed, the

comparable lengths of O-N1 and O-N2 indicate a subtle relationship with DNA

replication. Secondly, the length of NSR + Ori + NSL was barely equal to that of

Right + Ter + Left on the chromosome map, which coincides with the time of collapse

of newly synthesised DNA into docked spools at the middle stage of DNA replication.

In this paper, we propose new perspectives regarding DNA fluctuation and replication

to explain the origin of MDs of nucleoids, which might provide new clues for further

investigations.

Supplementary Discussion Diagram 2. Geometric features of six MDs in the genetic map of E. coli.

(a) Geometric features of MD distribution in the chromosome map subtly indicate the relationship

between MDs and DNA replication. Firstly, the length of oriC to each end of NS MDs (i.e., O-N1 and O-N2

fragments) was roughly comparable. Secondly, NSL + Ori + NSR fragment was barely equal to

Right + Ter + Left fragment.

2. Left and right chromosome arms are organized in separate cell halves

stochastically.

The right and left chromosome arms of E. coli are located in different cell halves,

as observed in fluorescent loci pairs on separate arms of the chromosome lying in

separate halves of a cell [19, 20]. Our model explained this phenomenon as the loci

pairs distributed in different spools in a nucleoid. Furthermore, our model also

accommodated the contradictory data in each group that ~10%–20% cells exhibited

reversed distribution patterns to the major pattern in previous studies (Supplementary

Discussion Published Data 2c). We attempt to explain the underlying mechanism as

follows. After completion of DNA replication, one docked spool containing one sister

chromosome DNA re-arranged into dual-toroidal-spool conformation. In this

transition, nearly half of the DNA contents of chromosome coiled into each spool of

the dual-toroidal-spool conformation. Re-arrangement occurs under conditions when

Ori MD and Ter MD are associated with cellular components, because Ori MD is

located near cell quarters and Ter MD is in the centre of the cell [13, 15, 16, 21]. Thus,

the two domains were likely distributed in separated spools because of spatial

constraints (Supplementary Discussion Diagram 3). Other parts of chromosome DNA

isolated in the cytoplasm could be coiled into either spool. Theoretically, numerous

possibilities exist for a circular chromosome DNA to be engulfed into nearly equal

halves (along dashed lines, Supplementary Discussion Diagram 3). In reality, given

the initial coiling of newly replicated Ori + NSL + NSR into a docked spool, followed

by gradual wrapping of Right + Left + Ter MDs, the confinement of cellular space,

crowding of cytoplasmic molecules, association with cytoplasmic components and

influence of the stochastically binding of NAPs, etc., there might be preferred

pathways for binary engulfment of DNA contents into dual-spool in dividing E. coli

cells. The pathway is predicted to be energetically favourable and biochemical

stochastically. In the present study, we could not address and investigate such an issue.

Based on the published experimental data [19, 20] (Supplementary Discussion

Published Data 2), there seems to be a bias of binary engulfment of chromosome

DNA contents along the axis largely across Ori and Ter directions rather than along

the axis across NS MDs or Right and Left MDs directions. This would be an

interesting issue to be investigated in future studies to understand more about the

pathways of chromosome packaging in live E. coli.

Balanced conformation is preferred for a final stable ‘frozen’ nucleoid, so nearly

half of DNA was engulfed into one spool of the dual-toroidal-spool conformation.

The loci pairs near each other in the chromosome map were at higher odds to be

distributed in the same spool than in a separate spool as checked from a cell

population, which was consistent with the experimental observations (Supplementary

Discussion Published Data 2a–c). However, close pairs may be engulfed into different

spools, so small portions of cells retained different patterns of loci dispositions to the

major portions of cells (Supplementary Discussion Published Data 2c). Interestingly,

the loci pairs which were separated about 180° on the circular chromosome, though

farther away in terms of left-arm-to-right-arm distance than other pairs separated

~120° on the genetic map, displayed higher occurrence rate to be engulfed into same

spool than that for 120° pairs, probably due to 180° loci pairs lying near edges of one

half chromosome DNA [19].

Supplementary Discussion Diagram 3. Pathways for stochastic rearrangement of single-spool

containing one sister chromosome DNA into a dual-toroidal-spool conformation. Two immobilised

chromosomal regions (Ori and Ter MDs) exist in E. coli. When a single-spool rearranged into dual-spool

with comparable size, the binary engulfment pathways (orange dashed lines) occurred stochastically

under the influence of cytoplasmic events.

a

b

c

Supplementary Discussion Published Data 2. Distribution pattern of chromosomal loci pairs in

cell halves [19, 20]. (a, b) Occurrence of chromosomal loci pairs located in the same spool or different

spools, which was correlated with their distance in the chromosome map. The loci pairs near each other

exhibited high occurrence in the same spool, whereas the pairs far from each other displayed high

occurrence in different spools. (c) In each group, the majority of cells exhibited the dominant pattern of

disposition of loci pairs, whereas a small portion cells displayed the reverse pattern. The open bar

indicates same cell halves, and the solid bar indicates separate cell halves. Two loci pairs ~180° away

from each other on the genetic map (1,4; green arrows) exhibited a higher occurrence of dispositions into

the same cell halves compared with the pairs ~120° away from each other (2,3).

3. Dynamic distribution of chromosomal loci revealed the global conformation

reorganization of nucleoid during a cell cycle.

The application of high-resolution fluorescence microscopy techniques and

powerful molecular genetics tools provides insight into the dynamic localisation of

individual chromosomal loci during cell division cycles. The oriC and dif loci have

unique targeting locations, whereas other loci are distributed linearly between them

based on their relative positions in the chromosome map [7, 13]. However, we

observed a diversity in gene loci distribution pattern apart from the linear pattern,

possibly reflecting the stochastic process of chromosome compaction. For example, as

shown in Supplementary Discussion Published Data 3a, the gln locus is much closer

to the oriC locus than to the lac locus in the gene map, but the lac locus appears

nearer to oriC than gln in the nucleoid according to FISH analysis [12]. This result

can be easily understood if they are distributed in a DNA toroidal spool. The relative

positions of loci in newly bilobed sister nucleoid altered dramatically compared with

those in newborn cells, indicating a global conformation re-arrangement of the sister

nucleoid. A similar phenomenon was observed previously [18] (Supplementary

Discussion Published Data 3b). These observations could possibly reflect the

transition of nucleoid conformation during cell division to allow the locus to

dramatically change its location.

Supplementary Discussion Published Data 3. Localisation of chromosomal loci in newborn cells

differs from that in middle-aged cells [12, 18]. (a, b) Fluorescent chromosomal loci in newborn cells

(upper panels) were distributed in a pattern different from that in middle-aged cells (lower panels). The

diagram indicating the corresponding conformation of chromosome DNA is shown on the right panels.

4. Asymmetry of newly bilobed sister nucleoid.

An unexpected feature of chromosome organization in E. coli is the asymmetry of

newly split sister nucleoids. When bilobed nucleoids initially appear, they are

asymmetric at around the late stage of DNA replication [10, 12] (Supplementary

Discussion Published Data 4). The bilobed nucleoids are shaped like dumbbells with a

prominent central connection. The two lobes then become equal in size, and they are

disposed symmetrically in the cytoplasm with no central connection when DNA

replication is complete. This finding agrees well with the model we proposed that one

of the docked spools transferred across Ter site to locate in other side of the cell (T3

to T4 transition, Supplementary Discussion Diagram 1b) during the late stage of DNA

replication. Strikingly, the travel of DNA spool acting as a solid explains the

‘dramatic one co-ordination pulsed transition’ as described by the authors [10, 12].

This translocation leaves significant extra room at the original side for the sub-sized

sister nucleoid, which might make it clearly visible by DAPI staining. Meanwhile, the

prominent central connection could be very possibly the newly synthesised DNA

strands from un-replicated parental DNA to be the entering and exiting ends of each

sister DNA spool. Due to molecular crowding and confinement of bacteria, it has been

difficult to observe directly the dynamic organization of nucleoid in vivo. Previous

remarkable findings on nucleoid behaviours in vivo provided precious clues for

understanding and determining a possible underlying mechanism that orchestrates the

organization and replication of bacterial nucleoids.

Supplementary Discussion Published Data 4. Asymmetry of new splitting sister nucleoid in

E. coli [10, 12]. (a, b) Newly split sister nucleoids in a configuration drawn in the bottom right panel in (a)

and bottom left panel in (b). One of the thus-far-replicated sister nucleoids at midcell translocated across

the non-replicated parental DNA to the other side of cells, resulting in the asymmetry of newly split

nucleoids.

5. Development of HNS clusters in nucleoids in vivo.

HNS is a global gene silencer and a main structural protein in the organization of

bacterial chromosome [22]. HNS has two modes to shape DNA, namely, the stiffen

mode and bridging mode [23]. The stiffen mode is related to gene regulation, and the

bridging mode directly modulates DNA condensation. Recently, single molecular

studies have revealed the detailed behavioural characteristics of HNS. Divalent ions

can switch the transition between two modes [23], and a high off-rate of HNS

bridging mode ensures the condensation of DNA but leaves DNA accessible for motor

proteins [24]. With the application of super-resolution PALM, the remarkable features

of HNS clusters in live E. coli have been revealed, which are distinct from other

NAPs dispersed throughout nucleoids [25]. Wang et al. found two HNS clusters in

newborn cells, and three to four HNS clusters in growing and ready-to-divide cells

(Supplementary Discussion Published Data 5a). One copy of chromosome DNA bears

two HNS clusters symmetrically distributed along the long axis of nucleoids (¼ and ¾

positions). This phenomenon is reminiscent of the auto-packaging behaviour of

plasmid DNA, resulting in a dual-toroidal-spool conformation with the highest DNA

density at the centre core of spools (also ¼ and ¾ positions). The HNS bridging mode

depends significantly on the proximity of DNA strands [24]. Thus, HNS is presumed

to reside in the centre core of DNA spools because of the close proximity of DNA

strands in this area, whereas HNS are not accessible to form significant DNA bridges

in the outer area with relatively loosely condensed DNA. HNS serves as a spacer and

also stabilizer chaperone of nucleoid in our proposed model. This role agrees with the

experimental findings, i.e., the deletion of HNS gene has only a small effect on the

growth rate of bacteria strain [26, 27]; nucleoids purified from the HNS-null strain

expand approximately five fold more than that observed in WT bacteria [9].

The dynamic development tendency for HNS clusters was schemed in

Supplementary Discussion Published Data 5b. How can two clusters develop into

three? How can three clusters develop into four to be ready for division? The dynamic

pathways were explained by our proposed model for chromosomal DNA organization,

and verified by in vivo observation of HNS clusters using PALM. We would like to

propose several factors involved in the organization of nucleoids in E. coli based on

our observations and related knowledge. First, the supercoiled circular DNA itself

underwent turbulent fluctuation of conformation upon injection of bending and

unwinding mechanical stress by NAPs, and the stress could not be released from the

ends of DNA as easily as linear DNA. Instead, the stress propagated along the circular

DNA backbone. Stressed nucleoids gradually settled down into a self-balanced

conformation to obtain the lowest energetic state. Second, when DNA replication was

initiated, the DNA strands were pulled to be processed through the replication factory

in E. coli [28, 29]. In this course, locally bound chaperone NAPs are likely to

dissociate by the pulling force generated (~30 pN), and the torque stress on this

portion of DNA is relieved. Concomitantly, topological insulators of nucleoids ensure

that the local pulling and torque relieving effects do not travel far to trigger global

topological conformation alteration [30, 31]. Third, NAPs such as HNS, HU, Fis and

IHF ect., are not the governor to shape and hold tightly a compact nucleoid, instead,

they are dynamically bound and dissociated from nucleoid and the overall torque

effect introduce DNA to transit from free diffusion state into packaged state. However,

the fundamental mechanism for auto-organization of highly ordered chromosome

DNA in cells is waiting to be discovered as the proceedings of single molecular

studies and dynamic in vivo investigations in future.

Supplementary Discussion Published Data 5. HNS cluster distribution in E. coli cells [25]. (a) Two

HNS clusters in newborn cells, three HNS clusters in growing cells and four HNS clusters in ready-to

divide cells were previously observed in E. coli. (b) The dynamic developing tendency of HNS clusters

was schemed in middle panel. Upper panel, DNA replication stage. Bottom panel, chromosome DNA

conformation during a cell cycle. I, early stage of DNA replication; II, middle stage of DNA replication; III,

complete of DNA replication; and IV, pre-division stage.

6. Dynamic observation of morphology of nucleoids in live E. coli.

The dynamic morphology of nucleoids has been extensively studied in recent

years [8, 17, 32-35]. New microscopic techniques and labelling strategies have

revolutionised our opinions of nucleoids, as revealed by super temporal and spatial

resolution microscopy. The striking features of nucleoids are addressed and discussed

in this section.

4D imaging of E. coli nucleoids with mCherry-tagged HU protein, a dynamic

helical ellipsoid was presented [33] (Supplementary Discussion Published Data 6a).

The discrete, dynamic ellipsoids within a nucleoid were typically found in growing E.

coli cells. This observation was consistent with our model; the fluctuant developing

DNA spools are reminiscent of dynamic ellipsoids. DAPI staining of E. coli cells

growing in Lennox rich medium revealed two to four blocks of DNA in cells [17]

(Supplementary Discussion Published Data 6b). In such growth conditions, DNA

replication proceeded without an isolation state from replication forks. Therefore,

single-spool could be the favoured packaging conformation for DNA mass. The DNA

blocks display similar morphology as DNA spools.

Supplementary Discussion Published Data 6. Morphology of nucleoids in E. coli cells. (a) 4D

imaging of mCherry-tagged HU revealed dynamic ellipsoid structures of nucleoids [33]. (b) DAPI staining

of nucleoids in MG1655 strain in Lennox rich medium, showing two to four blocks of chromosome DNA in

cells [17]. The enlarged images of the regions enclosed in white-line-boxes are listed at right and bottom

panels.

(Notably, numerous DAPI-stained nucleoids in live E. coli cells under slow growth conditions demonstrate

linear fluorescent morphology, as shown in Supplementary Discussion Published Data 4b. The difference

between linear and blocks appearance possibly derives from growth conditions. Under fast growing

condition, nucleoid is more compacted than in the slow growing media, which may be due to faster

synthesis rate of NAPs than that in cells growing in slow growing media. The more compacted, thus more

isolated nucleoid, may allow more obvious visualisation of transition of DNA global conformations than

loosely compacted nucleoid where fluorescent DNA blocks were not separated enough and blurred with

each other.)

Using an inducible GFP-tagged Fis protein to image the entire chromosome, the

growth and division of nucleoids during multifork replication was observed in cells

growing in grooves under LB-agarose pad [8] (Supplementary Discussion Published

Data 7a). One block of chromosome DNA in pre-division cells became bilobed in

newborn cells, and the two lobes were separated and isolated in the next cell cycle.

The organization and segregation patterns of the multifork E. coli chromosome are

schemed in Supplementary Discussion Published Data 7b and c. In brief, chromosome

DNA is organized into single-spool docking on replication forks before the

completion of DNA replication. Given that the initiation of a new round of DNA

replication occurs earlier than the end of whole chromosome replication, the newly

synthesised sister DNA is stressed by NAPs binding and collapses into two new

single-spools before the disassembly of the parental spool. Sister spools grow and the

parental spool shrinks as multiple fork replication proceeds. The parental spool finally

turns into a central connection and disappears. Thus, the sister nucleoids are isolated

from one another.

Supplementary Discussion Published Data 7. Morphology of multifork E. coli chromosome during

cell division cycles [8]. (a) The GFP-tagged Fis protein allows visualisation of nucleoids during a cell

cycle. (b) Diagram of a cycle of multiple-fork replication of chromosome DNA. (c) Proposed DNA

organizational pattern of nucleoids in different ages during a cell cycle corresponding to (b).

Conformations ii and iii explain the bilobed transition of nucleoids observed experimentally (white line

boxes, in a). i and iv indicate nucleoids with central connection (green arrows, in a). Replication forks

were omitted in ii–iv.

Helical bundles of nucleoids in E. coli have been proposed by both experimental

and theoretical investigations [7, 8, 33]. In direct scanning of nucleoids by monitoring

GFP-tagged Fis protein or mCherry-tagged HU protein, significant helical bundles

were observed using image analysis by deconvolution [8, 33] (Supplementary

Discussion Published Data 8a,b). In these observations, right-handed and left-handed

helicity were found even along the length of a single nucleoid, indicating no particular

handedness in nucleoids. In our 3D model, the DNA strands in dual-toroidal-spool

rotated along their orbits to compensate for the bending tension stress, as proposed by

theoretical analysis [1]. Thus, global helicity was introduced into the nucleoid by this

rotation. The upside down relationship between the two spools sharing similar

configuration (Supplementary Fig. S7) resulted in right-handedness in one spool and

left handedness in the other spool as observing the nucleoid from same perspective,

thereby eliminating the overall handedness of DNA bundles in a nucleoid

(Supplementary Discussion Published Data 8d). Another feature of our model for

nucleoids is that DNA density increased from the periphery to the centre core of the

spool. The bending energy was significantly higher in the centre core than that in the

periphery (Supplementary Fig. S5). HU and Fis proteins were sensitive to tension; the

association rate significantly decreased when the tension increased [36, 37]. Thus, the

centre core of mature DNA spools of nucleoids was dominated by non-bending NAPs,

such as HNS in bridging mode, which was favoured by closely juxtaposing DNA in

the centre core. Other DNA bending proteins had difficulty in conquering the bending

energy in DNA located in the centre core, so they were mainly abundant at the

periphery of spools. The periphery DNA bundle morphology derived from

dual-toroidal-spool conformation of nucleoids is schematically shown in

Supplementary Discussion Published Data 8 e and f, which was in agreement with the

observations documented.

Supplementary Discussion Published Data 8. Helical bundles of nucleoids. (a, b) The nucleoid was

scanned by fluorescent Fis or HU proteins, and helical bundles of the nucleoid were revealed previously

[8, 33]. (c) DNA conformations proposed for newborn cells (arrow, a) with multifork replication of

chromosome DNA (i, ii). Green spot, oriC locus; red square, dif locus; triangle, replication fork. The

binding profile of HU or Fis is shown in the middle panel. The centre core (dark blue) was void of

significant HU or Fis protein binding because of high bending energy, whereas the periphery of the

nucleoid (light blue) was bound by abundant HU and Fis proteins. (d) DNA conformation for newborn

nucleoids in slow growth conditions (arrow, b) is shown in the left panel. The helical bundles of nucleoids

originate from the 3D rotation of DNA strands along their orbits, as elucidated by the 3D model of

dual-toroidal-spool conformation. The upside down relationship eliminated the particular handedness

inside a nucleoid. The protein binding profile is in the right panel. (e) Sections of nucleoids along the main

plane (blue arrows, left panel) are shown on the right. The DNA strands on each section are drawn by

black lines. m.p. indicates main plane as indicated in (d). (f) At lateral view from both sides of nucleoid,

the overall curvature of a dual-spool was simplified shown in the boxes. Each spool contributes a “C”

shape curvature. The “two-tandem-C” shape resembles the curvature of nucleoids in (a) and (b).

Supplementary Discussion Published Data 9. Distribution of fluorescent HU proteins along the

long axis of the nucleoid. (a) Distribution of fluorescent HU protein along the y-slices shows a

two-helical-bundle pattern, as presented previously (iii, left panel) [33]. The dynamic fluctuation of

fluorescent signals was significant on the 5 s scale, as shown in the right panel. Fluctuation was probably

due to the association–dissociation dynamics of HU proteins to DNA, as well as turbulent fluctuation of

DNA strands in nucleoids. (b) The images of nucleoids during the early stage of DNA replication. At this

time point (before T1, as mentioned by the authors), one spool in dual-spool conformation already

replicated and slightly shrank, based on the lack of fluorescence in the lower panels (panels below ii).

This image was selected as a representative for comparison with a mature dual-toroidal-spool because

~62±14% newborn cells underwent dynamic fluctuation of their conformation with a sub-matured

un-balanced distribution of two HNS clusters, as observed in our experiments. DNA replication was often

observed after the appearance of two symmetrically comparable HNS clusters. Thus, the nucleoid in

early DNA replication stage was considered to retain, at least partially, the balanced mature

dual-toroidal-spool conformation with aids of topological insulators in E. coli. (c) Corresponding y-slices

of nucleoid based on our model. The green lines in the left panel indicate the positions of arbitrary slices.

Given that the centre core of each spool (dark blue) prevented significant binding of HU protein, the

periphery of the spool was bound with abundant HU proteins. Therefore, the fluorescence signals from

the top to bottom exhibited a “one-two-one-two-one” pattern based on our model (right panel), which is

substantially same as the pattern shown in (b, white line box). Due to the counterclockwise rotational

tendency of the spool along the long axis of nucleoid to the hinge, the two spools rotated and finally

buckled together to form a small dihedral angle between the two spools (dashed line, right panel), in

comparison with the case without rotational tendency of DNA spools to the hinge (dashed line, middle

panel). Coincidently, experimental observations display a dihedral angle between the two planes (the

planes across two bundles, red-line-box i and ii in b).

As recently reported, nucleoids in live E. coli exhibit striking features in temporal

and spatial organizations [33]. The authors performed 4D scanning of nucleoids using

mCherry-tagged HU protein. They observed ‘dual longitudinal helical bundles’ of

nucleoids. Surprisingly, the observations fit well with our model. As shown in

Supplementary Discussion Published Data 9, nucleoids illuminated by fluorescent HU

protein largely displayed a ‘one-two-one-two-one’ pattern of HU-tagged bundles from

one pole to the other pole. We explain this feature with our proposed model as follows.

The centre core lacked HU proteins, but HU was abundant at the periphery. Thus, two

bundles were observed in the middle part of the spool, but one bundle was detected in

the distal ends of the spool (Supplementary Discussion Published Data 9c). When the

nucleoids were in the process of replication, specifically, in the late phase of DNA

replication (Supplementary Discussion Published Data 10), the newly formed docked

spool or isolated spool underwent turbulent fluctuation. The conformation was not

balanced yet, so we presumed that tension in the centre core of such nucleoids was not

high enough to avoid significant HU binding. At these time points, HU proteins

exhibited one large alternating bundle with a fluorescent density slightly higher than

that of nucleoids in newborn cells.

There is a counterclockwise 3D rotational tendency of the spool along the long

axis of the nucleoid to the hinge, as suggested by our model (Supplementary Fig. S8).

Coincidently, the experimental observation demonstrated a small angle between the

two lines passing through both bundle signals in one slice (between i and ii,

Supplementary Discussion Published Data 9b). The slices of periphery DNA bundle

distribution in dual-toroidal-spool conformation are schematically shown in

Supplementary Discussion Published Data 9c, which were in agreement with the

experimental results.

It is striking that the pulsed nucleoid growth in length at defined time points

during a cell cycle was revealed in their study [33] (Supplementary Discussion

Published Data 11). The first pulse was observed when DNA replication was halfway

finished, which coincided with the collapse of newly synthesised DNA into two

docked spools in our model. At this time point, the authors mentioned the growth

direction towards the old pole of cells, which was in agreement with our observation

that two additional spools were initially distributed at the side of the old pole. There is

the second pulse of growth of nucleoid toward the other cell pole. We attributed this

second pulse to the translocation of one docked spool from the centre to across the

remaining parental spool during the late age of DNA replication. In another study,

sister chromosomes were described to lose cohesion and separate in a single

coordinate transition involving most or all of the thus-far-replicated regions at late

stage of DNA replication [12]; these findings were consistent with our observations

by PALM. The latter pulse growth of nucleoids was explained by our model as

follows. At third pulse, DNA replication was completed and the docked new spools

were isolated from the replication factory to move freely to fit into their respective

cell halves. At fourth pulse, two isolated single-spools rearranged into

dual-toroidal-spool conformation, which slightly elongated the length of nucleoids for

geometric reasons. To our surprise, our model presenting the dynamic organization of

chromosomes in E coli was in good agreement with the direct experimental

observations.

Due to limitation of manuscript, we cannot discuss many other interesting

published data about bacterial chromosome organization. Overall, in order to probe

the complicated question of chromosome DNA organization in E. coli, a new

perspective in combination of physical deduction and biological integration was

regarded in the present study for understanding of the underlying mechanisms. It has

been extremely difficult to directly monitor DNA compaction in a crowded tiny E.

coli cell. However, considering that chemical and physical principles govern every

biochemical reaction and behaviour of single molecules, and also considering that the

complicated and subtle experimental signs from in vivo observations are the

integrated reflections of cytoplasmic events, we checked the model deduced from

single molecular observations with thus-far-discovered in vivo behaviours of

nucleoids from the literature. Our proposed model accommodated majority of the

remarkable features of nucleoids in E. coli previously observed in vivo. The DNA

itself plays a major role in its organization, as shown in the present study. NAPs are

the auxiliary factors to deform local DNA strands, so the accumulation of the local

torque effect induced a global transition of DNA conformation from free diffusion

into a packaging spool, which was the major opinion of this study. The findings

provide new perspectives to understand chromosome organization in E. coli. The

underlying physical principles may expand our understanding of the auto-organization

behaviours of DNA in other bacteria and eukaryotes.

Supplementary Discussion Published Data 10. Fluorescent HU bundles in replicating nucleoids (a)

[33]. We would like to explain the T2 bundles with the diagram in (b). The newly formed DNA spools

located at upper and bottom panels, are abundantly bound with HU at this time span when a spool is still

immature. Whereas in the middle panel, the parental spool retains partially its original mature

conformation during replication, thus the centre core lacking HU binding resulted in two bundles of HU.

We would like to attribute the irregular fluorescent bundles to “newly-developing-spool” as shown in (c),

wherein the turbulent fluctuation of the spools and prosperously HU binding may thus result in

multi-bundles of HU proteins in irregular shape along the nucleoid.

Supplementary Discussion Published Data 11. Discontinuous sequential elongation pulses of

nucleoid at defined times in the cell cycle [33]. (a-b) Four discontinuous pulses in the growing of the

length of nucleoid, at T1 to T4 in the cell cycle (b). Meanwhile, a short period of nucleoid shortening was

detected following the period of length increase (a). (c) The corresponding increase in length of nucleoid

at T1 to T4 was highlighted in ii to v, as suggested by our model. The events are briefly described as

follows: at T1, freely defusing DNA collapsed into two additional spools; at T2, one of the new spools

translocated to the other side of cells; at T3, the docked new spools isolated from the replication factory

and moved into their cell halves; at T4, each spool containing one sister chromosome rearranged into the

final balanced dual-toroidal-spool conformation. We speculate that the short period of nucleoid length

shortening as shown in (a) may possibly due to the continuous compaction of the spools thus make it

more tightly and shrink a bit transiently, or the central connection pull the spools back a bit elastically in

response to the sudden movements arising from collapse into spools (T1) or translocation (T2).

Supplementary methods

Atomic force microscopy

The plasmid DNA pBR322 was platinated and subsequently imaged by AFM as

described previously [38]. pBR322 DNA at 5 ng/μl was incubated with C (ri = 0.4, ri

is the molar ratio of platinum complex to nucleotide), or with a C–T mixture (at 1:1

molar ratio) in 10 mM NaClO4 at 37 °C for 5 h in the dark. At 1, 2, 3, 4, and 5 h of

incubation, one portion (~2 µl) of the sample was subjected to AFM imaging. The

reaction was stopped in the remaining portions of samples, which were then purified

and subjected to inductively coupled plasma mass spectrometry (ICP–MS) test to

measure the bound platinum amount per DNA molecule. Samples of >5 h incubation

were excluded from analysis because of the intense condensation of plasmid DNA

molecules by platination, which is beyond the scope of this study. The sharp bends per

DNA molecule were calculated according to the relationship that the amount of sharp

bends equals 16% of the total divalent bound platinum amount per DNA molecule, as

determined by ICP–MS and denaturing gel electrophoresis analyses [38]. The writhe

number was checkd manually using the acompanying software of AFM. The results of

writhe number were confirmed by measuring the height value of the points where

DNA strands crossed over with each other. In such joints, the height value was about

twice or more of that of a dsDNA strand.

Construction mEos2-tagged HNS endogenous strain

The parent strain was MG1655 [39]. The chromosomal fusion of mEos2 and HNS

gene was constructed using λ Red recombination [25, 40]. MG1655 bacterial

strain-carrying pKD46 plasmids was electronically competent, and transformed with

100–500 ng of linear functional cassette DNA flanked with homologous sequences of

HNS by an electroporation machine (Bio-Rad, USA). The mEos2 expression plasmids

were purchased from Addgene (USA) [41]. The FRT flanked chloramphenicol

resistance gene (CAM) selection cassette was purchased (Novagene, Germany). The

mEos2 and CAM selection cassette was ligated by NotI restriction enzyme site. The

functional cassette for HNS-mEos2 fusion construction was prepared by PCR,

amplifying the mEos2-FRT-Chlora-FRT fragments with primers containing the ~56 nt

homologous sequences flanking the HNS gene. Successful recombination was first

selected by CAM antibiotic resistance marker, followed by colony PCR screening and

gene loci sequencing. The primers used are listed below.

H1-P1-HNS-mEos2,

CAAAAAAGCAATGGATGAGCAAGGTAAATCCCTCGACGATTTCCTGATCAA

GCAAATGAGTGCGATTAAGCCAG (55nt homology extensions)

H2-P2-HNS-mEos2,

GGACAATAAAAAATCCCGCCGCTGGCGGGATTTTAAGCAAGTGCAATCTAC

AAAAGATAATACGACTCACTATAGGG (57nt homology extensions)

Sample preparation for imaging

For imaging, the colony containing successful fusion constructs was selected and

incubated in 3 ml of LB media for 5 h. The cells were diluted at 1:10,000 into M9

minimal media supplied with vitamin and amino acids (Invitrogen, USA) or LB

media, and cultured at 250 rpm and 37 °C overnight. The cells were dilluted at

1:1000–1:2000 into fresh M9 media (or LB media) and cultured at the same

conditions for 2–4 h to reach an OD600 of ~0.1 the next morning. Approximately

200 µl of the culture was disposed onto a Petri dish precoated with 0.3%

poly-L-lysine as described previously [42]. The cells were allowed to immobilise onto

the Petri dish for 30–45 min before imaging, and maintained at room temperature.

Subsequently, 3 ml of fresh M9 complete media (or LB-agarose pad) was added into

the Petri dish, and the cells were subjected to imaging.

Imaging and data analysis

A fluorescent microscope and EMCCD were used as described previously [43].

The associated items (e.g., oil immersion objective, laser, mechanical shutter) were

purchased and assembled as described prevously [25]. The excitation and emission

paths were established as previously performed [25]. In our study, one to multiple cell

cycles were monitored by fluorescent HNS-mEos2 clusters. Extensive emmission at

405 nm would harm cellular DNA and other cytoplamic components. Cells need 10–

15 min to recover from single PALM imaging to continue to grow [28]. To reduce

phototoxicity by illumination at 405 nm, time lapse snapshots were obtained at time

intervals of 15, 20, and 25 min. 2-4 decades of mEos2 fluorophore per cell at each

time was photoactivated by the 405 nm excitation (30-60 W·cm-2) over the duration

of 100-500 ms. Two to three snapshots were taken under 20 W·cm-2 561 nm

excitation at 150 ms per frame. Subsequently, the fluorophore was photobleached.

Bright field images of the cell contours were acquired before PALM imaging. The

original images from ~700 cells was processed by image J software and sequentially

analysed based on the time during a cell cycle. The data were integrated to analyse the

reprensentive phase information of HNS cluster developments during the cell division

cycle.

Supplementary Reference

[1] I.M. Kulic, D. Andrienko, M. Deserno, Europhys Lett 67 (2004) 418-424.

[2] A. Rich, S. Zhang, Nat Rev Genet 4 (2003) 566-572.

[3] I.F. Lau, S.R. Filipe, B. Soballe, O.A. Okstad, F.X. Barre, D.J. Sherratt, Mol Microbiol 49 (2003)

731-743.

[4] M. Thanbichler, P.H. Viollier, L. Shapiro, Curr Opin Genet Dev 15 (2005) 153-162.

[5] R. Reyes-Lamothe, X. Wang, D. Sherratt, Trends Microbiol 16 (2008) 238-245.

[6] J. Pelletier, K. Halvorsen, B.Y. Ha, R. Paparcone, S.J. Sandler, C.L. Woldringh, W.P. Wong, S. Jun,

Proc Natl Acad Sci U S A 109 (2012) E2649-2656.

[7] P.A. Wiggins, K.C. Cheveralls, J.S. Martin, R. Lintner, J. Kondev, Proc Natl Acad Sci U S A 107

(2010) 4991-4995.

[8] N. Hadizadeh Yazdi, C.C. Guet, R.C. Johnson, J.F. Marko, Mol Microbiol 86 (2012) 1318-1333.

[9] V.V. Thacker, K. Bromek, B. Meijer, J. Kotar, B. Sclavi, M.C. Lagomarsino, U.F. Keyser, P.

Cicuta, Integr Biol (Camb) 6 (2014) 184-191.

[10] M.C. Joshi, A. Bourniquel, J. Fisher, B.T. Ho, D. Magnan, N. Kleckner, D. Bates, Proc Natl Acad

Sci U S A 108 (2011) 2765-2770.

[11] E. Toro, L. Shapiro, Cold Spring Harb Perspect Biol 2 (2010) a000349.

[12] D. Bates, N. Kleckner, Cell 121 (2005) 899-911.

[13] H. Niki, Y. Yamaichi, S. Hiraga, Genes Dev 14 (2000) 212-223.

[14] M. Valens, S. Penaud, M. Rossignol, F. Cornet, F. Boccard, EMBO J 23 (2004) 4330-4341.

[15] Y. Yamaichi, H. Niki, EMBO J 23 (2004) 221-233.

[16] O. Danilova, R. Reyes-Lamothe, M. Pinskaya, D. Sherratt, C. Possoz, Mol Microbiol 65 (2007)

1485-1492.

[17] R. Mercier, M.A. Petit, S. Schbath, S. Robin, M. El Karoui, F. Boccard, O. Espeli, Cell 135 (2008)

475-485.

[18] O. Espeli, R. Mercier, F. Boccard, Mol Microbiol 68 (2008) 1418-1427.

[19] H.J. Nielsen, J.R. Ottesen, B. Youngren, S.J. Austin, F.G. Hansen, Mol Microbiol 62 (2006)

331-338.

[20] X. Wang, X. Liu, C. Possoz, D.J. Sherratt, Genes Dev 20 (2006) 1727-1731.

[21] A. Thiel, M. Valens, I. Vallet-Gely, O. Espeli, F. Boccard, PLoS Genet 8 (2012) e1002672.

[22] C.J. Dorman, Nat Rev Microbiol 2 (2004) 391-400.

[23] Y. Liu, H. Chen, L.J. Kenney, J. Yan, Genes Dev 24 (2010) 339-344.

[24] R.T. Dame, M.C. Noom, G.J. Wuite, Nature 444 (2006) 387-390.

[25] W. Wang, G.W. Li, C. Chen, X.S. Xie, X. Zhuang, Science 333 (2011) 1445-1449.

[26] T. Atlung, F.G. Hansen, J Bacteriol 184 (2002) 1843-1850.

[27] R. Srinivasan, D. Chandraprakash, R. Krishnamurthi, P. Singh, V.F. Scolari, S. Krishna, A.S.

Seshasayee, Mol Biosyst 9 (2013) 2021-2033.

[28] S. Uphoff, R. Reyes-Lamothe, F. Garza de Leon, D.J. Sherratt, A.N. Kapanidis, Proc Natl Acad

Sci U S A 110 (2013) 8063-8068.

[29] R. Reyes-Lamothe, D.J. Sherratt, M.C. Leake, Science 328 (2010) 498-501.

[30] L. Moulin, A.R. Rahmouni, F. Boccard, Mol Microbiol 55 (2005) 601-610.

[31] L. Postow, C.D. Hardy, J. Arsuaga, N.R. Cozzarelli, Genes Dev 18 (2004) 1766-1779.

[32] S. Adachi, T. Fukushima, S. Hiraga, Genes Cells 13 (2008) 181-197.

[33] J.K. Fisher, A. Bourniquel, G. Witz, B. Weiner, M. Prentiss, N. Kleckner, Cell 153 (2013)

882-895.

[34] B. Youngren, H.J. Nielsen, S. Jun, S. Austin, Genes Dev 28 (2014) 71-84.

[35] G.S. Gordon, D. Sitnikov, C.D. Webb, A. Teleman, A. Straight, R. Losick, A.W. Murray, A.

Wright, Cell 90 (1997) 1113-1121.

[36] B. Xiao, H. Zhang, R.C. Johnson, J.F. Marko, Nucleic Acids Res 39 (2011) 5568-5577.

[37] B. Xiao, R.C. Johnson, J.F. Marko, Nucleic Acids Res 38 (2010) 6176-6185.

[38] Y.R. Liu, C. Ji, H.Y. Zhang, S.X. Dou, P. Xie, W.C. Wang, P.Y. Wang, Archives of Biochemistry

and Biophysics 536 (2013) 12-24.

[39] Y. Hao, Z.J. Zhang, D.W. Erickson, M. Huang, Y. Huang, J. Li, T. Hwa, H. Shi, Proc Natl Acad

Sci U S A 108 (2011) 12473-12478.

[40] K.A. Datsenko, B.L. Wanner, Proc Natl Acad Sci U S A 97 (2000) 6640-6645.

[41] S.A. McKinney, C.S. Murphy, K.L. Hazelwood, M.W. Davidson, L.L. Looger, Nat Methods 6

(2009) 131-133.

[42] J. Elf, G.W. Li, X.S. Xie, Science 316 (2007) 1191-1194.

[43] H. Li, Z.W. Duan, P. Xie, Y.R. Liu, W.C. Wang, S.X. Dou, P.Y. Wang, PLoS One 7 (2012) e45465.

Documents

A model for chromosome organization during the cell cycle in ...as a model system via developed molecular manipulation methods of genes and proteins in live cells. Scientists are attempting