Live-cell superresolution microscopy reveals theorganization of RNA polymerase in thebacterial nucleoidMathew Stracya, Christian Lesterlinb, Federico Garza de Leona, Stephan Uphoffa,c, Pawel Zawadzkia,c,and Achillefs N. Kapanidisa,1

aBiological Physics Research Group, Clarendon Laboratory, Department of Physics, University of Oxford, Oxford OX1 3PU, United Kingdom;bBases Moléculaires et Structurales des Systèmes Infectieux, UMR 5086, Centre National de la Recherche Scientifique, University of Lyon, 69 367 Lyon,France; and cDepartment of Biochemistry, University of Oxford, Oxford OX1 3QU, United Kingdom

Edited by Taekjip Ha, University of Illinois at Urbana–Champaign, Urbana, IL, and accepted by the Editorial Board July 2, 2015 (received for review April22, 2015)

Despite the fundamental importance of transcription, a compre-hensive analysis of RNA polymerase (RNAP) behavior and its rolein the nucleoid organization in vivo is lacking. Here, we used super-resolution microscopy to study the localization and dynamics of thetranscription machinery and DNA in live bacterial cells, at both thesingle-molecule and the population level. We used photoactivatedsingle-molecule tracking to discriminate between mobile RNAPs andRNAPs specifically bound to DNA, either on promoters or transcribedgenes. Mobile RNAPs can explore the whole nucleoid while searchingfor promoters, and spend 85% of their search time in nonspecific in-teractions with DNA. On the other hand, the distribution of specifi-cally bound RNAPs shows that low levels of transcription can occurthroughout the nucleoid. Further, clustering analysis and 3D struc-tured illumination microscopy (SIM) show that dense clusters oftranscribing RNAPs form almost exclusively at the nucleoid periphery.Treatmentwith rifampicin shows that active transcription is necessaryfor maintaining this spatial organization. In faster growth conditions,the fraction of transcribing RNAPs increases, as well as their clustering.Under these conditions, we observed dramatic phase separation be-tween the densest clusters of RNAPs and the densest regions of thenucleoid. These findings show that transcription can cause spatialreorganization of the nucleoid, with movement of gene loci out ofthe bulk of DNA as levels of transcription increase. This work providesa global view of the organization of RNA polymerase and transcrip-tion in living cells.

RNA polymerase | transcription | superresolution | single-moleculetracking | protein-DNA interactions

Cellular functions in bacteria are not compartmentalized asthey are in eukaryotes, yet they are still highly organized,

with specific subcellular regions associated with specific ma-chineries. In Escherichia coli, many of these machineries operateon the bacterial nucleoid, a highly structured and dynamic objectmade primarily by the 4.6-Mbp circular chromosome that iscompacted between 103- and 104-fold compared with the equivalentlinear DNA and segregated from the cytoplasm (1). Transcrip-tion is one of the most abundant processes occurring on thenucleoid DNA; transcription is driven by 1,500–5,000 moleculesof RNA polymerase (RNAP) per cell (2–4) and plays a crucialrole in maintaining both global and local DNA organization. Forexample, blocking cellular transcription with the antibiotic rifam-picin causes loss of global organization reflected by nucleoid ex-pansion (5, 6). Transcription has also been shown to modify thelocal DNA topology by unwinding the DNA double helix (7).Despite this, little is known about the spatial distribution of ac-tive RNAPs in living bacterial cells and how they affect the or-ganization of DNA and the nucleoid.The nucleoid structure depends on growth conditions. Under

slow growth conditions, where each cell contains between one andtwo chromosomes, the nucleoid lacks observable structures and

appears diffuse, occupying most of the cellular volume. By con-trast, under fast growth conditions, where the cells contain betweenfour and eight chromosome equivalents, the nucleoid is highlystructured and displays dramatic variation in local DNA density(8). Growth conditions also influence transcriptional activityon different genes; for example, the overall rate of synthesis ofribosomal RNA increases ∼40-fold in fast growth compared withslow growth conditions (9).This change in the level of expression isreflected in large changes in the spatial distribution of RNAP:Under slow growth conditions, RNAP appears to be fairly ho-mogeneously distributed over the diffuse nucleoid (10), whereasunder fast growth conditions, dense clusters of RNAPs emerge(4, 10). These dense clusters have been likened to transcriptionfactories in eukaryotic cells, where a single site contains multipleRNAPs active on different genes (11, 12); however, they have onlybeen reported in chemically fixed cells, and little is known abouttheir presence, dynamics, or behavior in live E. coli.Transcription and nucleoid structure are also interconnected

through the coupling of transcription and translation. In bacte-ria, ribosomes can bind to nascent mRNA as soon as the ribo-some binding site emerges from the transcribing RNAP (13).This raises an intriguing puzzle, because ribosomes and DNA arespatially separated in E. coli (3, 14, 15), whereas RNAP isstrongly associated with DNA (3, 10). To reconcile this withcotranscriptional translation, it has been hypothesized that

Significance

Transcription is one of the most fundamental processes for life.In eukaryotic cells, transcriptional activity is regulated to alarge degree by chromosome packaging. In bacteria, despitethe absence of a nuclear envelope and many of the DNA-packaging proteins of eukaryotes, the chromosome is stillhighly condensed into a structured object, the nucleoid. Thespatial organization of transcription within the nucleoid andthe effect of transcription on DNA organization remain poorlyunderstood. In this work, we characterize how RNA polymer-ase accesses transcription sites on DNA, and show that activetranscription can cause spatial reorganization of the nucleoid,with movement of gene loci out of the bulk of DNA as levels oftranscription increase.

Author contributions: M.S. and A.N.K. designed research; M.S. and C.L. performed re-search; M.S., F.G.d.L., S.U., and P.Z. contributed new reagents/analytic tools; M.S. analyzeddata; and M.S., C.L., and A.N.K. wrote the paper.

The authors declare no conflict of interest.

This article is a PNAS Direct Submission. T.H. is a guest editor invited by the EditorialBoard.1To whom correspondence should be addressed. Email: a.kapanidis1@physics.ox.ac.uk.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1507592112/-/DCSupplemental.

E4390–E4399 | PNAS | Published online July 29, 2015 www.pnas.org/cgi/doi/10.1073/pnas.1507592112

Dow

nloa

ded

by g

uest

on

Aug

ust 1

8, 2

020

much of cellular transcription must happen at the surface of thenucleoid, and close to the pool of ribosomes (16). However, evi-dence for this remains weak, because studies under moderategrowth conditions did not reveal any RNAP enrichment at thenucleoid periphery (3).Here, we use a combination of live-cell superresolution mi-

croscopy techniques to provide a comprehensive analysis of thebehavior of RNAP at both single-molecule and population levelsin individual cells. By using photoactivation localization micros-copy (PALM) (17) combined with single-particle tracking (18,19) of individual RNAP molecules in live cells, we sort them intoDNA-bound or mobile subpopulations (20). We show that mo-bile RNAPs are uniformly distributed across the nucleoid, sug-gesting that all DNA is probed through random nonspecificinteractions during the promoter search. The distribution ofspecifically bound RNAPs shows that low levels of transcriptioncan occur throughout the nucleoid. However, clustering analysisand 3D structured illumination microscopy (3D SIM) show thatspecifically bound RNAPs are more clustered than the mobilepopulation, and the denser clusters form preferentially at thenucleoid surface, indicating that heavily transcribed genes tendto move out of the bulk of nucleoid DNA. Imaging cellsgrowing in both rich and minimal media showed that clusteringincreases in rich media conditions and that segregation be-tween RNAP and DNA also increased.

Single-Molecule Tracking Discriminates DNA-Bound andMobile RNAPsRNAP can be either specifically bound to DNA (while inter-acting with promoter regions or with transcribed genes duringtranscription elongation) or can diffuse through the nucleoidsearching for promoters to initiate transcription. Because themovement of DNA loci is extremely slow compared with thediffusion on cytoplasmic proteins (20, 21), and the time to open apromoter and transcribe a gene [>20–100 s (9)] is at least 500-fold longer than nonspecific DNA interactions [30 ms (22)], wereasoned that individual RNAPs could be sorted into specificallybound molecules or mobile (diffusing and binding only tran-siently) based on their intracellular mobility (20, 23). To trackRNAP molecules, we used an endogenous fusion of photoacti-vatable fluorescent protein PAmCherry (24) with the beta′ subunitof RNAP (4). We imaged molecules in live cells by photoactivatingand localizing fluorophores (17), and joining localizations overmultiple frames to obtain trajectories of individual molecules(Fig. 1A) (18). By photoactivating and tracking all available mole-cules in cells grown in minimal media, we estimated the meanRNAP copy number to be 2,710 ± 700 per cell, which increasedwith cell length as expected (Fig. S1). We note that this may be anunderestimate of the true copy number of RNAP due to immatureor nonphotoactivatable PAmCherry molecules (25).To measure RNAP mobility, we calculated an apparent dif-

fusion coefficient (D*) from the mean squared displacement(MSD) of trajectories of individual molecules (20). As expected,the D* distribution for the entire population of RNAP molecules(Fig. 1B) does not fit to an analytical expression for a single dif-fusing species (Fig. S2A). To establish the apparent motion ofmolecules specifically bound to DNA, we used E. coli DNA poly-merase I (Pol1) as a control that shows distinct D* populations forDNA-bound and mobile molecules (SI Materials and Methods andFig. S2B) (20). We then fitted the RNAP D* distribution with atwo-species model with a DNA-bound population (constrainedusing the DNA-bound population in the Pol1 control) and a sec-ond, unconstrained D* species that corresponds to the populationof mobile RNAP molecules. This analysis showed that ∼48% ofRNAPs were bound and ∼52% were mobile (Fig. 1B, Inset); thisresult is in agreement with previous estimates from both fluo-rescence recovery after photobleaching (26) and single-moleculetracking (23). We then determined a D* threshold (0.16 μm2/s)

which preserves the bound-to-mobile ratio, and allows sorting ofindividual trajectories as corresponding to bound or mobileRNAP molecules (SI Materials and Methods). Plotting the spatialdistribution of the sorted molecules results in a 2D cellular mapthat shows the intracellular location of bound and mobileRNAPs in live cells (Fig. 1C).To check whether most DNA-bound RNAPs are indeed ac-

tively transcribing, we treated the cells with the antibiotic ri-fampicin, which binds to RNAP and blocks transcription beyonda 3-nt RNA but does not affect promoter binding, open complexformation, or transcription by RNAPs already in transcriptionelongation (27). Incubation with rifampicin for 30 min is thusexpected to cause most RNAPs to become mobile, with onlypromoter-bound RNAPs remaining bound to DNA, because anyelongating RNAP will complete transcription and dissociatefrom the DNA. Consistent with these expectations, the rifam-picin treatment led to a clear increase (from 52% to 84%) in thefraction of mobile RNAPs (Fig. 1 D and E). In contrast, the ri-fampicin treatment did not change the fraction of mobile DNApolymerase I (Pol1), confirming that the effect seen for RNAP isspecific to blocking active transcription (Fig. S2 E and F). Givenour estimates for the average RNAP copy number per cell, thebound fraction in the rifampicin-treated cells corresponds to∼430 bound RNAPs per cell. This estimate is in good agreementwith results from ChIP experiments, which showed that RNAPremains associated with ∼530 promoters per chromosome afterrifampicin incubation (28).The rifampicin treatment also increased the apparent diffu-

sion of mobile RNAPs (D* increases from 0.357 μm2/s (95%confidence interval: 0.338–0.378 μm2/s) to 0.636 μm2/s (0.609−0.658 μm2/s) (Fig. S2 C and D), likely the result of global chro-mosome decompaction triggered by rifampicin treatment (Fig.S3) (6). Chromosome decompaction leads to less densely packedstrands of DNA, and we therefore expect mobile RNAPs movingbetween more-distal DNA strands to show faster overall apparentdiffusion. In agreement with this interpretation, our control pro-tein Pol1 also showed increased mobility of its mobile populationupon rifampicin treatment [D* increases from 1.05 (1.02–1.07) μm2/sto 1.37 (1.40–1.35) μm2/s; Fig. S2F].

Bound and Mobile RNAPs Show Different SpatialDistributionsThe locations of mobile and bound RNAPs were compared withthe spatial distribution of DNA by staining the nucleoid DNAwith the intercalating fluorescent dye, SYTOX green (Fig. 2A)(29). The spatial distribution of mobile RNAPs in the cell closelyfollowed that of DNA, with only a few excursions of RNAP tracksinto the surrounding cytoplasm, indicating that most promoter-searching RNAP molecules remain within the volume of the nu-cleoid (Fig. 2B). In contrast, bound RNAPs were much moreheterogeneously distributed, and their distribution did not clearlymatch to that of DNA (Fig. 2C).To quantify the spatial clustering of mobile and bound RNAPs,

we calculated the pair correlation function, which gives the rela-tive probability of finding a protein at a given distance away fromanother protein compared with a random distribution (30). Thepair correlation function for molecules in >200 cells shows a clearclustering of bound RNAPs at distances shorter than 150 nm (Fig.2D), as expected for multiple RNAPs simultaneously bound athighly transcribed genes. This clustering was absent in the case ofmobile molecules, pointing to their independent, nonspecific RNAPinteractions throughout the nucleoid.As an additional verification that mobile RNAPs remain

strongly associated with the nucleoid, we compared the spatialdistribution of mobile RNAPs to that of nucleoid-associated heatunstable protein (HU), which reports on nucleoid DNA density(31). Plotting the probability density of HU molecules across thecell short axis (defined here as the x axis, Fig. 2E) through the

Stracy et al. PNAS | Published online July 29, 2015 | E4391

BIOPH

YSICSAND

COMPU

TATIONALBIOLO

GY

PNASPL

US

Dow

nloa

ded

by g

uest

on

Aug

ust 1

8, 2

020

center of the nucleoid (>100 cells) showed that the DNA densityis lowest at the cell periphery. Defining the periphery as theexterior 50% of the cell width (yellow highlighted areas, Fig. 2F)gave only 29% of HU molecules in this region compared with39% expected from a uniform distribution in a cylindrical cellvolume (32). Performing the same analysis on mobile RNAPmolecules showed that they matched extremely well to the dis-tribution of HU, and also have 29% located in the cell peripheryrather than uniformly filling the cell volume. These results con-firm our SYTOX-based results and strongly suggest that mobileRNAPs explore the entire nucleoid, having access to even thevery dense regions of the nucleoid DNA.

RNAP Spends 85% of Its Promoter Search Time Bound toNonspecific DNAWe then focused on the process of promoter search, whereinRNAP must identify specific substrate DNA sequences in themidst of huge amounts of nonspecific chromosomal DNA (22).The high level of association between mobile RNAPs and DNAand the absence of excursions into the cell cytoplasm furthersuggest a high level of transient nonspecific RNAP interactionswith DNA during the promoter search. Consistent with this, theapparent diffusion coefficient for mobile RNAPs in our experi-ments, D*mobile, is an order of magnitude smaller than the ex-pected free diffusion of RNAP, given its size (see SI Materials andMethods), indicating an interconversion between 3D diffusion andtransient binding.To determine the fraction of time that RNAP spends non-

specifically bound to DNA, two apparent diffusion coefficientsneed to be obtained: the D* of DNA-bound RNAP molecules(D*bound) and the D* for intracellular RNAP diffusion in the ab-sence of DNA (D*free). Although obtaining D*bound is straight-forward (extracted using the control experiments with Pol1; Fig. 1Band Fig. S2), obtainingD*free is more challenging, because virtuallyall RNAPs in wild-type E. coli are located within the nucleoid,complicating the study of freely diffusing RNAP molecules.To characterize the intracellular diffusion of RNAP in the

absence of the nucleoid, we used a temperature-sensitive DnaCmutation to generate a minimal-DNA E. coli strain with longDNA-free endcaps (Fig. 3A, Inset). At nonpermissive tempera-tures, these cells were unable to initiate DNA replication butkept elongating, yielding long cells containing a single chromo-some. We also treated cells with norfloxacin (a DNA gyrase in-hibitor) to enhance DNA compaction and increase the relativevolume of DNA-free endcaps. In these minimal-DNA cells,there is a dramatic increase in the mobility of RNAP comparedwith wild-type cells (Fig. 3A). Tracking RNAPs molecules lo-cated only in the DNA-free cell ends allowed us to determine theD*free distribution, which was centered at ∼1.1 μm2/s (Fig. 3B).The x axis distribution of RNAP molecules located in the DNA-free endcaps agreed well with the expected distribution formolecules evenly distributed through across the cell (Fig. 3D).To correct for the effects of confinement due to the small cell

volume, motion blurring, and localization error on the experi-mentally observed D*, and obtain accurate unbiased D values, weused Monte Carlo simulations of Brownian motion within an av-erage-size cell endcap (Fig. 3C) (19, 20, 33). By scanning throughD values and analyzing simulated tracks as we did for experi-mental tracks, we found that a diffusion coefficient Dfree of2.6 μm2/s best matched our data (dashed black line, Fig. 3B). Thisdiffusion is within the theoretically expected range (2.5–3.6 μm2/s),given the size of RNAP (see SI Materials and Methods and Fig.S4), and noticeably faster than previous estimates [0.7 μm2/s (23)].

Fig. 1. Single-molecule tracking allows mobility-based categorizing of in-dividual RNAPs as DNA-bound or mobile. (A) Example trajectories of indi-vidual RNAPs with specifically bound molecules colored red and mobilemolecules colored blue. (Scale bar, 1 μm.) (B) The distribution of apparentdiffusion coefficients, D*, for 69,900 RNAP molecules in live cells can befitted with two diffusing species (Inset), giving a ratio of 48% bound to 52%diffusing. Using these values allows a D* threshold to be determined tocategorize bound (red) and mobile (blue) molecules. (C) Representative ex-ample cells show the spatial distribution of categorized RNAP trajectoriescolored according to their D* value. (D) The distribution of D* values for39800 RNAP molecules in cells after incubation with rifampicin. (E) Example

cells after rifampicin treatment show fewer bound molecules. Mobile RNAPsexplore most of the cell volume due to the expansion of the nucleoid.

E4392 | www.pnas.org/cgi/doi/10.1073/pnas.1507592112 Stracy et al.

Dow

nloa

ded

by g

uest

on

Aug

ust 1

8, 2

020

Performing similar simulations on nucleoid-confined molecules,we found that the D*mobile of 0.36 μm2/s measured in wild-type cellsmatched a Dmobile of 0.4 μm2/s. We can then determine the fractionof time, x, that the interconverting species, Dmobile, spends bindingnonspecifically to DNA using Dmobile = xDbound + (1-x)Dfree (34).Using our experimentally determined D values to solve this equa-tion shows that RNAP spends 85% of its promoter search timenonspecifically bound to DNA. This high amount of nonspecificbinding could explain the very high spatial correlation of mobileRNAP with DNA observed in Fig. 2.

Transcribing RNAPs Tend to Cluster at the Edge of theNucleoidSimple visual inspection of the spatial distribution of boundRNAPs showed that they were much more frequently located atthe periphery of the nucleoid compared with the mobile population(Fig. 4A, i). To quantify this, we plotted the probability distri-bution of mobile and bound RNAPs across the cell short axis (xaxis) through the center of the nucleoid, from >200 cells. BoundRNAPs showed a broader distribution than the nucleoid (shownby both the HU distribution and mobile RNAP distribution),with 41% of them located in the cell periphery (yellow high-lighted areas, Fig. 4A, ii), compared with 29% for the nucleoid.This broader distribution is evident along the y axis as well as thex axis, with bound RNAPs also located at the periphery of thenucleoid toward the cell endcaps (Fig. S5).The relative localization of RNAP and nucleoid DNA was

further characterized using 3D SIM coimaging of RNAP−GFPand DAPI-stained DNA in live cells (Fig. 4A, iii and Movie S1).

SIM produces a superresolved visualization of the nucleoid vol-ume and edges, enabling better assessment of the relative positionof the RNAP clusters in three dimensions. We observed that theoverall RNAP signal overlaps with the nucleoid; however, thedensest RNAP clusters are nearly always located at the nucleoidedge (Fig. 4B, iii and Movie S2). Projections onto the z–x axishighlight that dense RNAP clusters that appear to be located inthe center of the cell in x−y projections are in fact located at thesurface of the nucleoid, where the DNA density is lower.We confirmed this observation by clustering analysis of DNA-

bound RNAPs from PALM data sets. We used a density-basedclustering algorithm (4, 35) to distinguish nonclustered RNAPs(single and pairs of RNAPs) from those in dense clusters (>6molecules). The resulting plot shows that dense clusters of boundRNAPs are clearly located at the nucleoid periphery (71% in theperiphery), whereas nonclustered RNAPs were located through-out the nucleoid (33% in the periphery) (Fig. 4B, ii). This showsthat, although low levels of transcription can occur throughout thenucleoid, dense RNAP clusters (i.e., more highly transcribed genes)were located at the nucleoid periphery, exactly at the interfacebetween the DNA and the ribosome-enriched cytoplasm. When weperformed the same analysis on another nucleoid associated pro-tein, the histone-like nucleoid-structuring (H-NS) protein, we sawthe opposite trend, with dense clusters located in the nucleoid center(only 12% in the periphery), and small clusters essentially matchingthe nucleoid density (30% in the periphery; Fig. S6). This resultestablishes that the clustering of NAPs does not universally lead toexclusion from the nucleoid, and adds weight to the hypothesis thatactive transcription drives RNAP segregation from the nucleoid.

Fig. 2. Comparison of categorized RNAPs with the distribution of DNA. (A) An example cell shown in a brightfield image (Left, used for cell segmentation),and in a SYTOX fluorescence image (Right, showing the nucleoid DNA distribution); x axis, short cell axis; y axis, long cell axis. (Scale bar, 1 μm.) (B) Thedistribution of mobile molecule trajectories (blue lines/bars) closely matched the distribution of DNA (green line) as shown in histogram projections along thex and y cell axes. (C) The distribution of bound RNAPs in the same example cell is more heterogeneous, and does not closely follow the DNA distribution.(D) Pair correlation of bound molecules (from 256 cells) shows a more clustered distribution than mobile molecules. (E) The average x axis distribution ofmolecules is measured from many cells by taking the relative distance from the cell midline through the center of the nucleoid, with −1 and 1 representingthe cell membrane. Short cells between 1.6 μm and 2.5 μm long were chosen as they have a single nucleoid, centrally located along the y axis. (F) Plot of the xaxis distribution of HU molecules from 256 cells shows the average DNA density is highest at the center and lowest at the cell periphery, with 29% ofmolecules found in the exterior 50% of the cell width (yellow highlighted area), compared with the expected distribution from molecules evenly occupyingthe full cylindrical cell volume (39% expected in cell periphery; dashed gray line). The distribution of mobile RNAP molecules (from 256 cells) matched ex-tremely well to the distribution of HU.

Stracy et al. PNAS | Published online July 29, 2015 | E4393

BIOPH

YSICSAND

COMPU

TATIONALBIOLO

GY

PNASPL

US

Dow

nloa

ded

by g

uest

on

Aug

ust 1

8, 2

020

Blocking transcription with rifampicin abolished this spatialpattern, as the distribution of bound RNAPs (on promoters, butnot transcribing) and mobile RNAPs became almost identical, with31% of both distributions located in the cell periphery (Fig. 4C, ii).Consistent with this, SIM images show homogenous, overlappingRNAP and DNA distributions (Fig. 4C, ii and Movie S3). Thisdemonstrates that active transcription is necessary for maintainingthis characteristic spatial distribution of RNAP with respect to thenucleoid DNA.We then examined the effect of active translation on the RNAP

spatial profile, because cotranscriptional translation has also beenproposed to play an important role in the nucleoid structure.According to the transertion model (36), transmembrane proteinsproduced by cotranscriptional translation indirectly attach tran-scribed genes to the cell membrane, promoting nucleoid decom-paction and potentially helping chromosome segregation. To blocktranslation, we used chloramphenicol (Fig. 4D), an antibiotic thatblocks protein chain elongation by inhibiting the peptidyl trans-ferase activity of the 50S ribosomal subunit. Consistent with pre-vious reports (16), SIM imaging showed that chloramphenicoltriggers a dramatic compaction of the nucleoid into a ring-likestructure that locates at the center of the cell (Fig. 4D, iii); how-ever, RNAP molecules still cluster at the nucleoid surface (MovieS4). These results are supported by PALM results, which show thatthe periphery bias of bound RNAPs is maintained; indeed, manycells showed distinct rings of bound RNAPs surrounding the nu-cleoid (Fig. 4D, i). On average, 46% and 33% of bound and mobileRNAPs, respectively, were located in the periphery (Fig. 3D, ii).These results suggest that transertion indeed promotes chromo-some expansion but is not the main driving force for the globalspatial localization of large RNAP clusters to the nucleoid surface.

Transcription in Rich Media Occurs in Dense DynamicClusters at the Nucleoid PeripheryAs growth rate increases, transcription on most genes is reduced;however, a small number of genes, particularly stable RNAs(such as ribosomal RNAs), become very heavily transcribed (9,10). When we performed tracking and sorting of RNAPs in cells

grown in rich media, we found that a significantly higher fractionof RNAPs (63%) were bound, compared with minimal mediaconditions (48%; Fig. 5A and Fig. S7 A and B). This differenceindicates a higher fraction of RNAPs engaged in transcription,despite the fact that cells in rich media have roughly half thenumber of active genes relative to minimal media (37).In cells grown in rich media, and subsequently chemically fixed,

RNAP has been shown to form dense clusters, which were notobserved in minimal media (4, 10); however, little is known abouttheir presence, dynamics, or behavior in live E. coli. The obser-vation of these large RNAP clusters, reminiscent of “transcriptionfactories” in eukaryotic cells (11), raised the question of whetherthese are static structures that stably anchor in cells and pulltranscribed DNA through them, or whether RNAPs assemble inlarge numbers on active promoters and simply follow the motionof the segregating chromosome during transcription. To addressthis question, we measured the mobility of RNAP clusters usingGFP-labeled RNAP in live cells growing in rich media (10). Time-lapse images over a 2-min timescale showed that these structureswere dynamic (Movie S5). To compare their mobility with that ofDNA loci, we labeled the DNA close to the origin of replication(Ori) on the chromosome by using an array of TetO operator sitesand chromosomally expressed TetR-YFP. Computing the MSDfor tracked RNAP clusters and DNA loci over the same timescale,we found that clusters have similar mobility to DNA (Fig. 5B).This indicates that these foci were moving along with the tran-scribed DNA region and are unlikely to be rigidly tethered to staticcellular structures.To characterize the size of these RNAP clusters in live cells, we

performed rapid localization microscopy with high photoactivation(38). Typical PALM imaging takes several minutes per super-resolved image, which would blur these RNAP clusters due to theirmovement over this timescale. Chemical fixation, on the otherhand, distorts biological structures (39). To capture sharp super-resolved PALM images in live cells, we localized RNAPs photo-activated at high density and analyzed using a crowded-field lo-calization algorithm (38), achieving a 50-nm resolution at 15 s perimage. These snapshots of RNAP localizations confirm that

Fig. 3. DNA-free diffusion of RNAP. (A) Example minimal-DNA cell (Inset); temperature-sensitive DnaC mutant cells are grown at a nonpermissive tem-perature to give long cells with a single centrally located chromosome. Diffusion of RNAP in minimal-DNA cells (blue columns, 97,900 molecules) is muchfaster than wild-type cells (gray columns, 69,900 molecules). (B) Tracking RNAPs only in the DNA-free cell endcaps (green bars, 2,400 molecules) allows the free3D diffusion to be determined. (C) Simulated molecular tracks undergoing Brownian diffusion within a confined cell endcap volume. Analyzing the tracksusing the same protocols as the experimental data gives an estimated accurate D value that best matches the experimental data. Scanning through D valuesfrom 1 μm2/s to 5 μm2/s, the best value was D = 2.6 μm2/s (black dashed line in B). (D) Plot of the x axis distribution of RNAP molecules from 72 DNA-free cellendcaps; 40% of these RNAP molecules are found in the exterior 50% of the cell width (yellow highlighted area); 39% of molecules evenly occupying the fullcylindrical cell volume are expected to be found in the periphery (dashed gray line).

E4394 | www.pnas.org/cgi/doi/10.1073/pnas.1507592112 Stracy et al.

Dow

nloa

ded

by g

uest

on

Aug

ust 1

8, 2

020

Fig. 4. Transcribing RNAPs cluster at the nucleoid periphery. (A) (i) An example cell growing in minimal media with trajectories of mobile (blue) and bound(red) RNAPs shows the difference in location of transcribing RNAPs compared with the mobile population. (ii) Probability density distribution of categorizedRNAPs in the nucleoid across the x axis for 256 cells. Mobile RNAPs (blue line) follow the distribution of the nucleoid (HU distribution; gray line), with29 ± 0.7% located in the exterior 50% of the cell width (yellow highlighted area). Transcribing molecules have a significantly broader distribution(P < 0.001), with 41 ± 0.9% of molecules locating in the periphery. (iii ) The 3D SIM images of live cells in minimal media show that the densest regionsof RNAP (red) are located at the edge of the nucleoid (blue). Projections onto the z–x axis highlight that dense regions apparently located in the centerof the cell in X−Y projections are in fact above or below the central bulk of the DNA. (B) (i ) Example cells showing clustering of bound RNAP molecules,with clustered RNAPs (>6 molecules) colored purple and nonclustered RNAPs (single or pairs of RNAPs) colored green. (ii ) x-axis distribution of clusteredand nonclustered bound RNAPs in 120 cells shows that nonclustered RNAPs are distributed throughout the nucleoid, whereas dense clusters form at theperiphery (71 ± 9%). (iii ) SIM images confirm that, although the RNAP distribution frequently overlaps with the DNA distribution, the densest RNAPregions locate at the cell edge (see also Movie S2). (C ) (i) An example cell after rifampicin incubation. (ii) After rifampicin incubation, the width dis-tributions of mobile and bound RNAPs become almost identical (P > 0.05), with 31 ± 0.8% of mobile molecules and 31 ± 1.6% of bound molecules foundin the cell periphery. (iii) SIM images show DNA and RNAP distributions fill most of the cell volume homogeneously. (D) (i) An example cell afterchloramphenicol incubation. (ii) The width distribution shows that 46 ± 1.7% of bound RNAP is found in the periphery compared with 33 ± 0.4% ofmobile molecules. (iii) SIM image showing a cell with a compacted nucleoid with RNAPs located at the edge of the bulk of DNA.

Stracy et al. PNAS | Published online July 29, 2015 | E4395

BIOPH

YSICSAND

COMPU

TATIONALBIOLO

GY

PNASPL

US

Dow

nloa

ded

by g

uest

on

Aug

ust 1

8, 2

020

Fig. 5. Organization of transcription in rich media. (A) The fractions of bound RNAPs determined per cell for different media. In rich media (531 cells), a significantlylarger fraction of RNAP is specifically bound compared with minimal media (883 cells). Controls after chemical fixation (322 cells) and after rifampicin incubation (204cells) show concomitant increases and decreases in bound fraction, respectively. (B) Localizing dense RNAP foci in live cells (Inset) allows them to be tracked in time-lapse experiments (Movie S5). MSDs of RNAP foci (red) and labeled DNA loci (blue). (C) Representative cells imagedwith fast acquisition PALM, showing all localizations(Top), and clustered RNAPs, with number of molecules in clusters indicated (Bottom). (D) Histogram of the cluster sizes (in number of RNAPs) for 218 cells in rich mediaand 298 cells in minimal media. (E) Example fields of view showing the DAPI-stained nucleoids of live cells imaged with 3D SIM. Surface contour plots highlight thenucleoid structure. (F) The 3D surface renderings of RNAP−GFP (red) and DNA (DAPI, blue) distributions in example cells grown inminimal (Left) and rich (Right) media.

E4396 | www.pnas.org/cgi/doi/10.1073/pnas.1507592112 Stracy et al.

Dow

nloa

ded

by g

uest

on

Aug

ust 1

8, 2

020

clustering is much more extensive under fast growth conditionscompared with minimal media conditions (Fig. 5C). To quantifythis difference, we calculated the pair correlation distribution ofthe localizations, which showed an increase in clustering at pairwisedistances less than ∼150 nm for rich media compared with minimalmedia (Fig. S7 C and D).To get an estimate of the number of RNAP molecules per cluster

in cells grown in different media, we clustered our localizationsusing a density-based algorithm (4, 35). We determined the meannumber of localizations per molecule (by particle tracking using thesame acquisition protocol but lower photoactivation) and used thisto convert localizations to numbers of molecules (Fig. 5D). Ourresults showed that the cluster sizes in live cells match broadly toprevious estimates in fixed cells (4): We observed intermediate-sizeclusters in minimal media (∼50 molecules), and large clusters (up to500 molecules) in rich media.SIM imaging in live cells confirmed that the organization of the

nucleoid also changes as growth rate increases. In minimal media,the nucleoid is relatively homogenous (Fig. 5E, Left), whereas inrich media, nucleoids are more heterogeneous, with distinctstructures (Fig. 5E, Right). In rich media, the very dense RNAPclusters clearly form at the nucleoid edge, showing stronger seg-regation than in minimal media, with very little overlap betweenRNAP and DNA (Fig. 5F and Movie S6). Interestingly, the RNAP

clusters form not just at the cell periphery but also in areas of low-density DNA throughout the cell volume (Fig. 5F, Right). Com-paring rich and minimal media shows that increasing RNAPclustering correlates with increased bias of RNAP to the edge ofthe nucleoid. In rich media, this effect is more dramatic becausenot only is transcription on a few genes increased heavily (leadingto increased clustering) but transcription is also reduced on mostof the remaining genes, which could allow this DNA to be moretightly compacted.

DiscussionUsing live-cell superresolution microscopy, single-molecule track-ing, and diffusion simulations, we have shown how RNAP locatespromoters distributed throughout the nucleoid, and how activetranscription causes spatial reorganization of the DNA.

Mobile RNAPs and Promoter Search. In minimal media conditions,approximately half of RNAPs are mobile and diffusing within thenucleoid, engaging in frequent nonspecific interactions withDNA as they search for promoters. Contrary to a recent study(40), we see that mobile RNAPs are not excluded from any partof the nucleoid. Indeed, mobile RNAPs show striking agreementwith the DNA distribution, with the densest regions of DNAcorrelating with the densest concentrations of mobile RNAPs.This implies that searching RNAPs can find promoters locatedthroughout the nucleoid (Fig. 6A).The fraction of specifically bound molecules, fbound, can tell us

about the approximate timescale for an average RNAP to find apromoter and engage in transcription, tmobile, because fbound =tbound/(tbound + tmobile) (20). In minimal media, fbound ≈ 0.5,hence the search time, tmobile, is roughly the same as the averagetranscription event, on the order of 30–120 s (considering theapproximately timescales for promoter opening, promoter es-cape, and transcriptional elongation, and the length of the av-erage gene). During this search process, RNAP spends about85% of its time nonspecifically bound to DNA, which is a similarpercentage to that seen for the lac repressor (34). In rich media,fbound is nearly 30% higher (0.63); therefore the average searchtime should be only ∼60% the duration of the average tran-scription event.

Bound RNAPs Found in Clusters Are Biased Toward the NucleoidPeriphery. Transcribing RNAPs are more clustered than mobilemolecules. Interestingly, nonclustered transcribing RNAPs arelocated throughout the nucleoid; however, the densest clustersare preferentially located at the edge of the nucleoid where theDNA density is lowest. It appears, therefore, that although lowlevels of transcription can occur on genes distributed throughoutthe nucleoid, the most heavily transcribed genes locate to thenucleoid periphery. Incubation with rifampicin shows that activetranscription is necessary to maintain these genes at the pe-riphery. These results provide compelling evidence that tran-scription itself is the driving force for locating active genes to thenucleoid edge, rather than a mechanism whereby regions of thechromosome are inaccessible to RNAP (40), leaving only genesat the periphery available to be transcribed.

Mechanism of RNAP Spatial Organization. One mechanism thatcould cause highly transcribed genes to locate to the nucleoidperiphery is cotranscriptional translation (12, 32). Ribosomes arestrongly segregated from the E. coli nucleoid (3, 14, 15). Statis-tical models have suggested that this phase separation betweenthe ribosome-containing cytoplasm and the DNA can be ex-plained by simple entropic forces: The DNA polymer avoids thewalls to maximize conformational entropy, and the polysomes(multiple 70S ribosomes on a single mRNA) occupy the emptyspace near the walls to maximize translational entropy (41).Although mRNA-bound 70S ribosomes are excluded from the

Fig. 6. Mechanisms of gene spatial organization. (A) RNAPs and free ribo-some subunits can explore the entire nucleoid search for specific nucleic acidsequences. RNAP starts transcribing a gene within the nucleoid, and the firstribosome binds to the emerging mRNA. As the polyribosome grows andother RNAPs start transcribing, entropic forces favor movement away fromthe bulk of the nucleoid. (B) In the case of rRNA transcription, there is nocoupled translation, but rRNA operons are extremely highly transcribed.Multiple RNAPs on the same gene may also drive movement of the genetoward the periphery of the nucleoid.

Stracy et al. PNAS | Published online July 29, 2015 | E4397

BIOPH

YSICSAND

COMPU

TATIONALBIOLO

GY

PNASPL

US

Dow

nloa

ded

by g

uest

on

Aug

ust 1

8, 2

020

nucleoid, it has recently been shown that free 50S and 30Ssubunits do have access to the nucleoid center (32, 42). Theseresults indicate that both transcription and translation can beginin the center of the nucleoid (Fig. 6A). Transcription initiationand early elongation produce a nascent mRNA that contains aribosome binding site, which nucleates the recruitment and as-sembly of one or several ribosomes when translation is initiated.It seems likely that the large DNA−RNAP−ribosome complexesare also entropically excluded from the bulk of the nucleoid andtend to migrate toward the nucleoid periphery, thus leading tothe spatial separation of genes depending on their level of ex-pression, with highly expressed genes enriched at the surface of thenucleoid. The inner volume of the nucleoid is consequently lessmetabolized, potentially allowing it to become more highly com-pacted. Consistent with this proposal, in minimal media, the nu-cleoid is less structured, showing less variation in local DNAdensity due to the fact that genes are expressed throughout thechromosome. In contrast, in rich media, fewer genes are expressed,thus allowing large parts of the chromosome to be inactive, givingrise to more structured and compacted nucleoids.In rich media, most transcription occurs on the ribosomal RNA

operons. Despite the fact that these transcripts are not translated,there appears to be even stronger segregation between RNAPclusters and nucleoid DNA in rich media compared with minimalmedia. Coupled transcription and translation is therefore not theonly mechanism of locating RNAP clusters to the nucleoid edge.In rich media, the active genes are fewer but are more heavilytranscribed; for example, ribosomal operons can have up to 80transcribing RNAPs on them at any one time (4). The largenumber of RNAPs, which may also be coupled with associatedribosome assembly cofactors (43), are likely to exert similar en-tropic bias out of the nucleoid DNA as coupled RNAP−mRNA−ribosome complexes (Fig. 6B).Our results also indicate that other potential mechanisms for

creating the phase separation between RNAP and the nucleoidhave significant shortcomings. For example, because MreB andRNAP copurify in cell extracts and interact in vitro (4), it hasbeen proposed that RNAP clusters could be anchored to thecytoskeleton. Although our data do not rule out some interactionbetween RNAP and the cytoskeleton, we believe it is unlikely forthis to be a primary mechanism for locating RNAP clusters tothe nucleoid periphery, for two reasons. Firstly, tracking ofRNAP clusters in rich media showed that they have very similarmobility to DNA, suggesting that they are not rigidly tethered tothe cytoskeleton. Secondly, 3D SIM imaging showed that in richmedia, where the nucleoid forms more heterogeneous structures,RNAP clusters form not just at the cell periphery but also in thecentral volume of the cell in low-density regions of the nucleoidthat are distal from the cell membrane.Transertion has also been proposed to play a role in nucleoid

decompaction because blocking translation with chloramphenicolcauses nucleoid compaction. Indeed, individual genes expressing

membrane proteins have been shown to migrate to the membranewhen expressed (44). However, we show that clusters of boundRNAPs still remain at the nucleoid edge after chloramphenicolincubation, indicating that transertion might play a role in thenucleoid conformation but is clearly not the driving force behindlocalization of large RNAP clusters to the nucleoid surface.Taken together, our work shows that transcription can cause

spatial reorganization of the nucleoid, with movement of geneloci out of the bulk of DNA as levels of transcription increase.The peripheral localization of transcription is reminiscent of thatof DNA repair (45), supporting the view that DNA metabolismactivities involving large machineries locate preferentially at thenucleoid periphery, where large protein complex formation isfacilitated. In rich media conditions, the number of RNAPclusters observed per cell (10), and the number of RNAPs foundin each cluster (4), has led to the hypothesis that these clustersrepresent RNAPs active on multiple heavily transcribed genesspatially located to the same site, analogous to transcriptionfactories in eukaryotes (11, 46). Similarly, during DNA breakrepair, distant DNA loci are brought together at the periphery ofthe nucleoid (45). The localization of heavily transcribed genesoutside the bulk of DNA may therefore also facilitate spatialclustering of genes located at distant sites on the chromosome.

Materials and MethodsComplete details of materials and methods are available in SI Materials andMethods. In brief, cells were grown in either M9 minimal or EZ Rich DefinedMedia (Teknova) supplemented with 0.2% glucose to OD of ∼0.2 andimmobilized for imaging on 1% agarose pads. Where indicated, cells wereincubated with 50 μg/mL rifampicin or 100 μg/mL chloramphenicol for30 min. Live cell single-molecule-tracking PALM, rapid high-density PALM,and RNAP-GFP cluster tracking experiments were performed on a custom-built total internal reflection fluorescence (TIRF) microscope. PALM data forsingle-molecule-tracking analysis was localized using custom-written MATLABsoftware (MathWorks). Fluorophore images were localized to 40-nm precisionby elliptical Gaussian fitting. Localizations within a radius of 0.48 μm in con-secutive frames were linked into tracks. An apparent diffusion coefficient, D*,was calculated from the mean-squared displacement (MSD) for each track witha minimum of 4 steps. 3D-Structured illumination (SIM) imaging was per-formed on a DeltaVision OMX V3 (Applied Precision/GE Healthcare) equippedwith a Blaze SIM module. Complete details of PALM and SIM microscopy aredescribed in SI Materials and Methods. Data analysis and simulations wereperformed in MATLAB; full details are presented in SI Materials and Methods.

ACKNOWLEDGMENTS. We thank Kieran Finan for the gift of bacterial strains,and we thank David Sherratt and Amy Upton for their help constructingstrains. This work was supported by the European Commission SeventhFramework Programme Grant FP7/2007-2013 HEALTH-F4-2008-201418, UKBiotechnology and Biological Sciences Research Council Grant BB/H01795X/1,and European Research Council Grant 261227 (to A.N.K.). M.S. was supportedby the Engineering and Physical Sciences Research. F.G.d.L. was supported bythe Consejo Nacional de Ciencia y Technología/I2T2. S.U. was supported by aSir Henry Wellcome Postdoctoral Fellowship and a Junior Research Fellowshipat St. John’s College Oxford. C.L. and P.Z. were supported by a Wellcome TrustProgramme Grant to David Sherratt (WT083469MA).

1. Sherratt DJ (2003) Bacterial chromosome dynamics. Science 301(5634):780–785.2. Grigorova IL, Phleger NJ, Mutalik VK, Gross CA (2006) Insights into transcriptional

regulation and sigma competition from an equilibrium model of RNA polymerasebinding to DNA. Proc Natl Acad Sci USA 103(14):5332–5337.

3. Bakshi S, Siryaporn A, Goulian M, Weisshaar JC (2012) Superresolution imaging ofribosomes and RNA polymerase in live Escherichia coli cells. Mol Microbiol 85(1):21–38.

4. Endesfelder U, et al. (2013) Multiscale spatial organization of RNA polymerase inEscherichia coli. Biophys J 105(1):172–181.

5. Dworsky P, Schaechter M (1973) Effect of rifampin on the structure and membraneattachment of the nucleoid of Escherichia coli. J Bacteriol 116(3):1364–1374.

6. Cabrera JE, Cagliero C, Quan S, Squires CL, Jin DJ (2009) Active transcription of rRNAoperons condenses the nucleoid in Escherichia coli: Examining the effect of tran-scription on nucleoid structure in the absence of transertion. J Bacteriol 191(13):4180–4185.

7. Moulin L, Rahmouni AR, Boccard F (2005) Topological insulators inhibit diffusion oftranscription-induced positive supercoils in the chromosome of Escherichia coli. MolMicrobiol 55(2):601–610.

8. Hadizadeh Yazdi N, Guet CC, Johnson RC, Marko JF (2012) Variation of the folding

and dynamics of the Escherichia coli chromosome with growth conditions. Mol

Microbiol 86(6):1318–1333.9. Bremer H, Dennis PP (1996) Escherichia coli and Salmonella: Cellular and Molecular

Biology, ed Neidhardt FC (Am Soc Microbiol Press, Washington, DC).10. Cabrera JE, Jin DJ (2003) The distribution of RNA polymerase in Escherichia coli is

dynamic and sensitive to environmental cues. Mol Microbiol 50(5):1493–1505.11. Cook PR (1999) The Organization of replication and transcription. Science 284(5421):

1790–1795.12. Jin DJ, Cabrera JE (2006) Coupling the distribution of RNA polymerase to global gene

regulation and the dynamic structure of the bacterial nucleoid in Escherichia coli.

J Struct Biol 156(2):284–291.13. Proshkin S, Rahmouni AR, Mironov A, Nudler E (2010) Cooperation between trans-

lating ribosomes and RNA polymerase in transcription elongation. Science 328(5977):

504–508.14. Hobot JA, et al. (1985) Shape and fine structure of nucleoids observed on sections of

ultrarapidly frozen and cryosubstituted bacteria. J Bacteriol 162(3):960–971.

E4398 | www.pnas.org/cgi/doi/10.1073/pnas.1507592112 Stracy et al.

Dow

nloa

ded

by g

uest

on

Aug

ust 1

8, 2

020

15. Wang W, Li GW, Chen C, Xie XS, Zhuang X (2011) Chromosome organization by anucleoid-associated protein in live bacteria. Science 333(6048):1445–1449.

16. Jin DJ, Cagliero C, Zhou YN (2013) Role of RNA polymerase and transcription in theorganization of the bacterial nucleoid. Chem Rev 113(11):8662–8682.

17. Betzig E, et al. (2006) Imaging intracellular fluorescent proteins at nanometer reso-lution. Science 313(5793):1642–1645.

18. Manley S, et al. (2008) High-density mapping of single-molecule trajectories withphotoactivated localization microscopy. Nat Methods 5(2):155–157.

19. English BP, et al. (2011) Single-molecule investigations of the stringent responsemachinery in living bacterial cells. Proc Natl Acad Sci USA 108(31):E365–E373.

20. Uphoff S, Reyes-Lamothe R, Garza de Leon F, Sherratt DJ, Kapanidis AN (2013) Single-molecule DNA repair in live bacteria. Proc Natl Acad Sci USA 110(20):8063–8068.

21. Elmore S, Müller M, Vischer N, Odijk T, Woldringh CL (2005) Single-particle tracking oforiC-GFP fluorescent spots during chromosome segregation in Escherichia coli. J StructBiol 151(3):275–287.

22. Wang F, et al. (2013) The promoter-search mechanism of Escherichia coli RNA poly-merase is dominated by three-dimensional diffusion. Nat Struct Mol Biol 20(2):174–181.

23. Bakshi S, Dalrymple RM, Li W, Choi H, Weisshaar JC (2013) Partitioning of RNA po-lymerase activity in live Escherichia coli from analysis of single-molecule diffusivetrajectories. Biophys J 105(12):2676–2686.

24. Subach FV, et al. (2009) Photoactivatable mCherry for high-resolution two-colorfluorescence microscopy. Nat Methods 6(2):153–159.

25. Durisic N, Laparra-Cuervo L, Sandoval-Álvarez A, Borbely JS, Lakadamyali M (2014)Single-molecule evaluation of fluorescent protein photoactivation efficiency using anin vivo nanotemplate. Nat Methods 11(2):156–162.

26. Bratton BP, Mooney RA, Weisshaar JC (2011) Spatial distribution and diffusive motionof RNA polymerase in live Escherichia coli. J Bacteriol 193(19):5138–5146.

27. Campbell EA, et al. (2001) Structural mechanism for rifampicin inhibition of bacterialRNA polymerase. Cell 104(6):901–912.

28. Grainger DC, Hurd D, Harrison M, Holdstock J, Busby SJW (2005) Studies of the dis-tribution of Escherichia coli cAMP-receptor protein and RNA polymerase along theE. coli chromosome. Proc Natl Acad Sci USA 102(49):17693–17698.

29. Bakshi S, et al. (2014) Nonperturbative imaging of nucleoid morphology in live bac-terial cells during an antimicrobial peptide attack. Appl Environ Microbiol 80(16):4977–4986.

30. Sengupta P, et al. (2011) Probing protein heterogeneity in the plasma membraneusing PALM and pair correlation analysis. Nat Methods 8(11):969–975.

31. Fisher JK, et al. (2013) Four-dimensional imaging of E. coli nucleoid organization anddynamics in living cells. Cell 153(4):882–895.

32. Sanamrad A, et al. (2014) Single-particle tracking reveals that free ribosomal subunitsare not excluded from the Escherichia coli nucleoid. Proc Natl Acad Sci USA 111(31):11413–11418.

33. Stracy M, Uphoff S, Garza de Leon F, Kapanidis AN (2014) In vivo single-moleculeimaging of bacterial DNA replication, transcription, and repair. FEBS Lett 588(19):3585–3594.

34. Elf J, Li G-W, Xie XS (2007) Probing transcription factor dynamics at the single-mol-ecule level in a living cell. Science 316(5828):1191–1194.

35. Ester M, Kriegel H, Jörg SXX (1996) A density-based algorithm for discovering clustersin large spatial databases with noise. Second International Conference on KnowledgeDiscovery and Data Mining (KDD-96), eds Simoudis E, Han J, Fayyad UM (Assoc Ad-vancement Artif Intell, Palo Alto, CA), pp 226–231.

36. Woldringh CL (2002) The role of co-transcriptional translation and protein translocation(transertion) in bacterial chromosome segregation. Mol Microbiol 45(1):17–29.

37. Tao H, Bausch C, Richmond C, Blattner FR, Conway T (1999) Functional genomics: ex-pression analysis of Escherichia coli growing on minimal and rich media. J Bacteriol181(20):6425–6440.

38. Holden SJ, Uphoff S, Kapanidis AN (2011) DAOSTORM: An algorithm for high- densitysuper-resolution microscopy. Nat Methods 8(4):279–280.

39. Schnell U, Dijk F, Sjollema KA, Giepmans BNG (2012) Immunolabeling artifacts andthe need for live-cell imaging. Nat Methods 9(2):152–158.

40. Spahn C, Cella-Zannacchi F, Endesfelder U, Heilemann M (2015) Correlative super-reso-lution imaging of RNA polymerase distribution and dynamics, bacterial membrane andchromosomal structure in Escherichia coli. Methods Appl Fluoresc 3(1):014005.

41. Mondal J, Bratton BP, Li Y, Yethiraj A, Weisshaar JC (2011) Entropy-based mechanismof ribosome-nucleoid segregation in E. coli cells. Biophys J 100(11):2605–2613.

42. Bakshi S, Choi H, Mondal J, Weisshaar JC (2014) Time-dependent effects of tran-scription- and translation-halting drugs on the spatial distributions of the Escherichiacoli chromosome and ribosomes. Mol Microbiol 94(4):871–887.

43. Shajani Z, Sykes MT, Williamson JR (2011) Assembly of bacterial ribosomes. Annu RevBiochem 80:501–526.

44. Libby EA, Roggiani M, Goulian M (2012) Membrane protein expression triggers chro-mosomal locus repositioning in bacteria. Proc Natl Acad Sci USA 109(19):7445–7450.

45. Lesterlin C, Ball G, Schermelleh L, Sherratt DJ (2014) RecA bundles mediate homologypairing between distant sisters during DNA break repair. Nature 506(7487):249–253.

46. Papantonis A, Cook PR (2013) Transcription factories: Genome organization and generegulation. Chem Rev 113(11):8683–8705.

47. Datsenko KA, Wanner BL (2000) One-step inactivation of chromosomal genes in Es-cherichia coli K-12 using PCR products. Proc Natl Acad Sci USA 97(12):6640–6645.

48. Tokunaga M, Imamoto N, Sakata-Sogawa K (2008) Highly inclined thin illuminationenables clear single-molecule imaging in cells. Nat Methods 5(2):159–161.

49. Michalet X, Berglund AJ (2012) Optimal diffusion coefficient estimation in single-particle tracking. Phys Rev E Stat Nonlin Soft Matter Phys 85(6 Pt 1):061916.

50. Sliusarenko O, Heinritz J, Emonet T, Jacobs-Wagner C (2011) High-throughput, sub-pixel precision analysis of bacterial morphogenesis and intracellular spatio-temporaldynamics. Mol Microbiol 80(3):612–627.

51. Kumar M, Mommer MS, Sourjik V (2010) Mobility of cytoplasmic, membrane, andDNA-binding proteins in Escherichia coli. Biophys J 98(4):552–559.

52. Elowitz MB, Surette MG, Wolf P-E, Stock JB, Leibler S (1999) Protein mobility in thecytoplasm of Escherichia coli. J Bacteriol 181(1):197–203.

53. Ando T, Skolnick J (2010) Crowding and hydrodynamic interactions likely dominate invivo macromolecular motion. Proc Natl Acad Sci USA 107(43):18457–18462.

Stracy et al. PNAS | Published online July 29, 2015 | E4399

BIOPH

YSICSAND

COMPU

TATIONALBIOLO

GY

PNASPL

US

Dow

nloa

ded

by g

uest

on

Aug

ust 1

8, 2

020