Edited by Sharon R. Long, Stanford University, Stanford, CA, and approved July 2, 2014 (received for review April 23, 2014)
In bacteria, faithful and efficient DNA segregation is intimately linked to the spatial organization of the chromosome. Two distinct organization patterns have been described for bacterial chromosomes (ori-ter and left-ori-right) that appear to arise from distinct segregation mechanisms. Here, we show that the Bacillus subtilis chromosome oscillates between them during a replication–segregation cycle. Our data further suggest that the highly conserved condensin complex and the parABS partitioning system function as the core components that underlie these alternating patterns. We propose that this oscillatory cycle enhances the efficiency of DNA replication and sister chromosome segregation and provides a unifying model for the diverse patterns of chromosome organization observed in bacteria.
Bacterial chromosomes have been found to possess one of two distinct patterns of spatial organization. In the first, called “ori-ter” and exemplified by Caulobacter crescentus, the chromosome arms lie side-by-side, with the replication origin and terminus at opposite cell poles. In the second, observed in slow-growing Escherichia coli (“left-ori-right”), the two chromosome arms reside in separate cell halves, on either side of a centrally located origin. These two patterns, rotated 90° relative to each other, appear to result from different segregation mechanisms. Here, we show that the Bacillus subtilis chromosome alternates between them. For most of the cell cycle, newly replicated origins are maintained at opposite poles with chromosome arms adjacent to each other, in an ori-ter configuration. Shortly after replication initiation, the duplicated origins move as a unit to midcell and the two unreplicated arms resolve into opposite cell halves, generating a left-ori-right pattern. The origins are then actively segregated toward opposite poles, resetting the cycle. Our data suggest that the condensin complex and the parABS partitioning system are the principal driving forces underlying this oscillatory cycle. We propose that the distinct organization patterns observed for bacterial chromosomes reflect a common organization–segregation mechanism, and that simple modifications to it underlie the unique patterns observed in different species.
Central to reproduction is the faithful segregation of replicated chromosomes to daughter cells. In eukaryotes, DNA replication, chromosome condensation, and sister chromatid segregation are separated into distinct steps in the cell cycle that are safeguarded by checkpoint pathways. In bacteria, these processes occur concurrently, posing unique challenges to genome integrity and inheritance (1, 2). In the absence of temporal control, bacteria take advantage of spatial organization to promote faithful and efficient chromosome segregation. The organization of the chromosome dictates where the chromosome is replicated, and the factors that organize and compact the newly replicated DNA play a central role in its segregation (1, 2).
Studies in different bacteria have revealed strikingly distinct patterns of chromosome organization that appear to arise from different segregation mechanisms. In Caulobacter crescentus and Vibrio cholerae chromosome I, the origin and terminus are located at opposite cell poles, with the two replication arms between them, in a pattern referred to as “ori-ter” (3⇓–5). After replication initiation, one of the sister origins is held in place and the other is actively translocated to the opposite cell pole, regenerating the ori-ter organization in both daughter cells (5⇓⇓⇓⇓⇓–11). By contrast, in slow-growing Escherichia coli, the origin is located in the middle of the nucleoid and the two replication arms reside in opposite cell halves, in a “left-ori-right” pattern (12, 13). Replication initiates at midcell, and the duplicated origins segregate to the quarter positions followed by the left and right arms on either side, regenerating the left-ori-right pattern in the two daughter cells.
Although chromosome organization was first analyzed in the Gram-positive bacterium Bacillus subtilis, our understanding of the replication–segregation cycle in this bacterium has remained elusive. In pioneering studies, it was shown that during spore formation, the replicated origins reside at opposite cell poles and the termini at midcell in an ori-ter ter-ori organization (14⇓⇓⇓–18). A similar ori-ter pattern was observed during vegetative growth (15, 19). However, in separate studies, DNA replication was found to initiate at a midcell-localized origin (20, 21). How these disparate patterns fit into a coherent replication–segregation cycle has never been addressed and motivated this study. Our analysis has revealed that the B. subtilis chromosome follows an unexpected and previously unidentified choreography during vegetative growth in which the organization alternates between ori-ter and left-ori-right patterns. Our data further suggest that the highly conserved partitioning system (parABS) and the structural maintenance of chromosomes (SMC) condensin complex, in conjunction with replication initiation, function as the core components for this oscillating cycle. We propose that this cycle enhances the efficiency of DNA replication and sister chromosome segregation and provides a unifying model for the diverse patterns of chromosome organization observed in bacteria.
As a first step toward our analysis of chromosome organization in B. subtilis, we modified the cytological tools to visualize chromosomal loci in live cells. Previous fluorescently tagged repressor proteins bound to operator arrays caused partial or complete blocks to replisome progression in B. subtilis (15, 22, 23). We reduced the number of binding sites in the arrays and lowered expression of the repressor protein fusions using a weak constitutive promoter (Materials and Methods). Under these conditions, virtually all cells contained fluorescent foci with no detectable impact on growth rate, nucleoid morphology, or nucleoid size distribution (Fig. S1).
To validate these tools, we analyzed the organization of the replicated chromosomes during spore formation. At early stages in this developmental process, the replicated chromosomes adopt a structure called the axial filament, in which the origins are anchored at the cell poles and the termini are present near midcell (16, 24⇓–26). We visualized six chromosomal loci [−7° (ori) and ±87°, ±120°, and +174° (ter)] during sporulation and determined their population-average, normalized position relative to cell length using automated image analysis software (27) (Fig. 1 A–E). The origins were present close to the cell poles at 0.17 and 0.83, and the termini were located at midcell (0.5). The ±87° and ±120° loci localized linearly between them. These results confirm and extend previous observations (24, 25, 28) that the sporulating chromosomes are organized linearly along their length from the poles to midcell in a C. crescentus-like ori-ter ter-ori organization (Fig. 1B).
Chromosome organization during sporulation. (A–E) Exponentially growing cells were induced to sporulate and imaged after 2 h. All strains contained a mutation in spoIIIE (spoIIIE36) (55) to prevent DNA transport. (A) Chromosome map showing positions of operator arrays. (B) Schematic summary of cellular localization of loci. Representative micrographs of cells labeled with pairs of loci ori-ter (C, Left), ±87° (D, Left), and ±120° (E, Left) are shown. (C–E, Right) Loci position relative to cell length analyzed (Materials and Methods). The centroid of the Gaussian distribution is shown on the top of each histogram. (F and G) Origin number and localization in rich and poor growth media. Origins (green, tetO48/TetR-CFP), nucleoid (blue, HBsu-mYpet), and membrane [red, N-(3-triethylammoniumpropyl)-4-(6-(4-(diethylamino) phenyl) hexatrienyl), pyridinium dibromide (FM4-64)] are shown. (Scale bar: 4 μm.)
The analysis of chromosome organization in C. crescentus and slow-growing E. coli was facilitated by their simple cell cycles with no overlapping rounds of replication (3, 12, 13). When grown in rich media, B. subtilis, like E. coli, undergoes multifork replication and the cells are born with partially replicated chromosomes. In defined rich medium [doubling time (τ) = 35 min], the cells were born with two origins and, on average, the number of origins per cell was 3.1 (n = 1,825) (Fig. 1F). To obtain a simpler DNA replication cycle, we grew B. subtilis in minimal medium supplemented with different carbon sources, as was done previously in E. coli, to reduce growth rate and DNA content. Although we could slow growth in glucose (τ = 48 min), succinate (τ = 77 min), and sorbitol (τ = 98 min), under all conditions tested, the cells were born with two origins, and the average number of origins per cell was 2.8, 2.3, and 2.1 (n > 1,700), respectively (Fig. 1G and Fig. S2A). Even under the slowest growth condition, cells were born with greater than 50% of their chromosome replicated (Fig. S2B). Thus, we were unable to identify conditions in which cells are born with a single unreplicated chromosome (1-N content). These results suggest that there is a link between DNA replication and cell division that ensures cells are born with two almost completely replicated chromosomes under normal growth conditions, a requirement for successful sporulation. We therefore initiated our analysis using conditions in which cells were artificially restricted to a single copy of the chromosome by inhibiting replication initiation.
To analyze vegetatively growing cells with 1-N DNA content, we inhibited replication initiation using a temperature-sensitive mutant of the helicase loader (DnaB) (29, 30). Cells were grown in rich medium at the permissive temperature (30 °C, τ = 57 min) to early exponential phase and then shifted to the restrictive temperature (42 °C, τ = 33 min) (Fig. 2A). After 1 h, 84% of the cells (n = 1,224) had a single-lobed nucleoid containing one replication origin, indicating that the chromosome content was reduced to 1-N (Fig. 2A). After 1.5 h, >90% of the cells (n = 2,399) had a single origin. Furthermore, as observed previously by McGinness and Wake (31), the nucleoids adopted a bilobed structure (Fig. 2A). Strikingly, in most cases (92%, n = 2,196), the origin was located at or close to the center of the two lobes (Fig. 2 A, C, D, and F). This localization pattern is dramatically different from sporulating cells, in which origins are at the extreme poles (Fig. 1C), and vegetatively growing cells with unperturbed replication, in which origins are frequently localized at the nucleoid periphery (14, 16) (Fig. 1 F and G and Fig. S2A). To determine whether each nucleoid lobe represents a chromosome arm, we visualized pairs of loci on the left and right arms. Eighty percent of the cells (n = 1,633) with fluorescently tagged loci at ±87° and 88% of the cells (n = 1,634) with tagged loci at ±120° had these markers in separate nucleoid lobes (Fig. 2 B and E). We also monitored loci at −87° and −120° on the same chromosome arm. In 86% of the cells (n = 1,804), the two loci were present in the same nucleoid lobe (Fig. 2 B and E). Furthermore, in the majority of these cells, the −87° locus was closer to the center of the two lobes and the −120° locus was closer to the nucleoid edge (Fig. 2 B and E).
Left-ori-right organization in cells with a single chromosome. Cells with a dnaB(ts) mutation were shifted from 30 °C to 42 °C for 1.5 h to block new rounds of initiation. (A) Origin localization before and after temperature shift. Origins (green), nucleoid (red), and membrane (white) are shown. (B) Representative micrographs of cells labeled at pairs of loci: ±87° (Left), ±120° (Center), and −87° and −120° (Right). (C) Cells labeled at ori (red) and ter (green). (D) Cells labeled at ori (red) and expressing RTP-YFP (green). (E and F) Quantitative analysis of loci positions. Origin localization in dnaB(ts) 1.5 h after temperature shift and degradation of SMC (G) or ScpB (H) is shown. (Scale bars: 4 μm.)
Analysis of a locus close to the replication terminus (+174°) revealed that most cells had a ter locus directly between the two lobes (64%, n = 2,196) or in close proximity to this position (17%) (Fig. 2 C and F). However, in 18% of the cells, the terminus was present near the edge of the nucleoid. To visualize a larger region of the terminus, we used YFP fusion to the replication termination protein (RTP) (32). RTP binds to nine termination sites spanning ∼430 kb of the terminus region (33). RTP-YFP localized as a single focus, clusters of foci, or multiple foci spanning the bilobed nucleoid (Fig. 2D), consistent with the terminus region connecting the ends of the two lobes. Similar results were obtained using different strain backgrounds and different growth media, as well as when initiation was blocked using dnaA(ts) or induction of SirA, an inhibitor of DnaA (34, 35). Collectively, these data indicate that under conditions in which B. subtilis is restricted to 1-N content, the nucleoid is organized in a pattern like the E. coli chromosome during slow growth, with the two arms present on either side of the origin and genetic loci arranged linearly from the middle toward the poles. Finally, the terminus region connects the two ends.
In E. coli, the left-ori-right organization is shifted to an ori-ter pattern in cells lacking the condensin complex MukBEF (36, 37). We used degradable variants of B. subtilis SMC or its partner protein ScpB to investigate whether the left-ori-right organization similarly depends on the condensin complex. Degradation of SMC (and ScpB separately) was induced when dnaB(ts) cells were shifted to the restrictive temperature to block replication initiation. After 1.5 h, 91% of SMC-degraded cells (n = 1,005) had a more extended chromosome structure, with the replication origin located at the extreme edge of the nucleoid (Fig. 2G and Fig. S3). Similar results were obtained after ScpB degradation (Fig. 2H). The Spo0J (ParB) protein in B. subtilis has been shown to recruit SMC complexes to the origin region (28, 38). Accordingly, we investigated the organization of the nucleoid in its absence (Fig. S3). In the majority of the Spo0J mutant cells (70%, n = 999), the origin of replication localized to the edge of the nucleoid. This phenotype was principally due to the absence of Spo0J-mediated recruitment of SMC rather than the loss of the parABS partitioning activity, because cells lacking Soj (ParA) that are not impaired in SMC recruitment (28) retained the left-ori-right pattern (Fig. S3). We conclude that origin-localized condensin complexes play a critical role in generating the E. coli-like pattern.
The two distinct patterns of chromosome organization described above were observed under extreme conditions. In one, the origins were physically tethered to the cell poles during bacterial differentiation (24, 26); in the other, replication initiation was blocked and the cells were restricted to 1-N content. To investigate whether exponentially growing cells with unperturbed DNA replication possess either pattern, we used time-lapse fluorescence microscopy to visualize chromosomal loci during replication–segregation cycles. To ensure the best time resolution, we used minimal medium supplemented with sorbitol (τ = 98 min). First, we visualized the origin (Fig. 3A). The cells were born with two origins at opposite edges of the nucleoid and remained at the nucleoid periphery for the rest of the replication cycle. Previous studies suggested that replication initiates in the middle of the nucleoid (15, 20). However, in our movies, the origin foci became brighter at or close to the nucleoid periphery, indicative of their replication at this site (Fig. 3A and Fig. S4A, red carets). Similarly, in snapshot images, replisomes colocalized with the brighter origin foci at the nucleoid edge (Fig. S4B). Shortly after their replication, the two new origins migrated as a unit to the middle of the nucleoid (Fig. 3A and Fig. S4A, yellow carets). This event was followed by their bidirectional segregation to opposite edges of the nucleoid. This cycle is consistent with snapshot images in which most cells have two origins at the outer edges of the nucleoid (16, 19) (Fig. 1 F and G and Fig. S2A).
Alternating patterns of chromosome organization during the cell cycle. (A) Time-lapse progression (5-min intervals) of cells labeled at the origin (green) and nucleoid (red) grown in minimal medium supplemented with sorbitol. Red carets highlight replication initiation at the edge of the nucleoid. Yellow carets show duplicated origins at the center of the nucleoid before their segregation. (B) Time-lapse progression (10-min intervals) of cells labeled at −87° (green) and +87° (red). Black arrows highlight resolution of the left and right loci. (C) Representative micrograph of cells labeled at three chromosomal loci: origin (blue, mCherry-Spo0J), −87° (green), and +87° (red). Stages in the replication–segregation cycle are indicated (I–III) and are interpreted in the schematic model (Right). Origin, −87°, and +87° loci are labeled as blue, green, and red balls, respectively. Chromosome arms are shown as gray lines. (Scale bar: 4 μm.)
Next, we tracked markers on the left and right chromosome arms. For most of the cell cycle, the ±87° loci localized adjacent to each other. However, for a brief period (20–30 min), the two loci separated, localizing on either side of the nucleoid (Fig. 3B, black arrow, and Fig. S5). After this period of resolution, the two loci were replicated, segregated, and colocalized again. Similar results were obtained for the pair of markers at ±120°. To relate the dynamic movement of the markers on opposite arms to the replication cycle, we used mCherry–Spo0J (ParB) fusion to localize the origin (14) while simultaneously following the markers at ±87°. When the ±87° markers colocalized, the origin was almost always present at the edge of the nucleoid (Fig. 3C). However, when the ±87° markers were spatially resolved, the replicated origins were either in the middle of the nucleoid between the ±87° markers or had already segregated to the edges of the nucleoid (Fig. 3C). Thus, movement of the duplicated origins to midcell is accompanied by resolution of the left and right arms. This unprecedented choreography can explain some of the unusual localization patterns observed previously (15, 19) and is fully supported by snapshot images in which we visualized fluorescently labeled replisomes and an origin locus (Fig. S6).
Combining the time-lapse microscopy, the three-color images, and the replisome localization relative to the origin, a picture emerges of the replication–segregation cycle in B. subtilis (Fig. 4). At birth, the two origins are located at opposite edges of the nucleoid, with the partially replicated arms lying side by side. At the time of the next round of initiation, replisomes assemble on the polarly localized origins. The newly replicated origins then migrate as a unit to the middle of the nucleoid. At this time, the unreplicated chromosome arms become spatially resolved and the replisomes track independently of each other along them (Fig. S7). The left-ori-right pattern of the template DNA persists even after the origins are segregated to the nucleoid periphery and only returns to the ori-ter–like organization after the arms themselves have been replicated.
Schematic model of the organization–segregation cycle in B. subtilis, and its comparison with C. crescentus and E. coli. Origins are represented as black balls, and termini are represented as brown lines in B. subtilis and E. coli and as a brown oval in C. crescentus. The compacted left and right chromosome arms are shown as thick blue and purple lines (or blobs). Newly replicated DNA is shown with a lighter hue, whereas uncompacted DNA is shown as thin lines. In the B. subtilis, newborn cell, unreplicated DNA is shown as a black cloud. The replication–segregation cycle of C. crescentus and E. coli corresponds to two halves of the cycle in B. subtilis.
We investigated whether the B. subtilis partitioning locus is responsible for maintaining the origins at the nucleoid periphery until the next round of replication. Chromosomally encoded par loci are composed of an ATPase (ParA), a DNA binding protein (ParB), and a centromere-like sequence (parS) (39). All three are critical for origin segregation in C. crescentus (40, 41), Pseudomonas aeruginosa (42), and V. cholerae chromosome II (43). In B. subtilis, cells lacking ParA, called Soj, have a very mild defect in chromosome segregation (23). ParB mutants have a more pronounced phenotype, but this is thought to be a consequence of a failure to enrich SMC complexes at the origin (23, 28, 38). Accordingly, for these experiments, we used a Soj (ParA) mutant that has no apparent impact on origin-localized SMC (28). Analysis of origin dynamics in cells lacking Soj revealed that the directional movement of the newly replicated origins was abolished (Fig. 5 A and C and Fig. S8). Instead of segregating to the edge of the nucleoid (Figs. 3A and 5B and Fig. S8), the origins moved more slowly and in an irregular manner (Fig. 5 A and C and Fig. S8). Although they reached the nucleoid edge, they frequently moved back toward midcell.
Impaired origin segregation and chromosome arm dynamics in the absence of Soj (ParA). (A) Time-lapse progression (5-min intervals) of ∆soj cells. The origin (green) and nucleoid (red) are shown. Representative origin traces in WT (B) and ∆soj (C) are shown. The position of the origin was plotted relative to the total cell length (Fig. S8). (D) Interfocal distances of ±87° loci are analyzed in representative WT (Upper) and ∆soj (Lower) cells in time-lapse progressions (10-min intervals). Each graph plots the distance between a pair of ±87° loci relative to the cell length (L) at indicated times during their replication cycle. For the purpose of this analysis, the replication cycle of these loci begins when both ±87° loci are replicated (defined as frame 1) and finishes when either of the two loci is replicated again. Because each cell has two pairs of ±87° loci in its replication cycle, the maximum distance of a pair of ±87° loci is 50% of the cell length (Fig. S10).
Next, we investigated whether Soj helps maintain the two chromosomal arms adjacent to each other in an ori-ter pattern. Time-lapse imaging of markers at ±87° in the ∆soj mutant revealed that these loci remain resolved for the majority (62 ± 8%) of the replication cycle (n = 100) (Fig. 5D and Figs. S9 and S10B). By contrast, in WT cells, these loci were resolved for only 37 ± 7% (n = 100) of the replication cycle (Figs. 3B and 5D and Figs. S5 and S10A). Furthermore, instead of alternating between arm colocalization and arm resolution as observed in WT cells (Figs. 3B and 5D and Figs. S5 and S10A), the ∆soj mutant had more dynamic arm movements, with irregular patterns of arm colocalization and resolution (Fig. 5D and Figs. S9 and S10B). These data indicate that Soj, and presumably the partitioning locus, facilitates bidirectional origin segregation and plays a central role in both establishing and maintaining the ori-ter pattern.
Our data support a model in which origin-localized condensin complexes and the partitioning system are the principal driving forces that underlie the two patterns described here, whereas replication initiation serves as the switch that triggers the oscillation between them. In this model, recruitment of SMC complexes to the origin by ParB (28, 38) sets up the left-ori-right pattern. SMC resolves the replicated origins (44, 45), and by analogy to the stiffening of eukaryotic centromeres by condensin (46), we hypothesize that SMC rigidifies the origin region constraining the left and right arms on either side of the origin through lengthwise condensation (47). Because SMC complexes are present at the origin throughout the cell cycle, force must be applied to the chromosome to generate the ori-ter pattern. We have shown that Soj (ParA) actively segregates origins to the nucleoid periphery and plays an important role in both establishing and maintaining this ori-ter organization. By analogy to chromosome and plasmid segregation in other bacteria (48, 49), we hypothesize that Soj acts by exerting pulling forces on Spo0J (ParB) bound to parS adjacent to the origin. Finally, we propose that replication initiation triggers the change in organization pattern by transiently inactivating the forces exerted on the origins. One possibility is that Soj’s role in replication initiation (50) could temporarily relax its segregation function. Alternatively or in addition, replication of the origin region could inactivate the partitioning system by transiently disrupting the Spo0J–parS complexes. When force on the origins is lost, the chromosome resolves into its SMC-mediated left-ori-right pattern. Par-mediated segregation of the newly replicated origins initiates a new round of this oscillatory cycle. Intriguingly, in cells that are blocked for replication initiation, the partitioning system appears to become inactivated, leading to a stable left-ori-right configuration (Fig. 2). Chronic inhibition of replication initiation could maintain Soj in its replication-active state (50), resulting in a sustained loss of its segregation function. Alternatively, the loss of partitioning activity could be due to a change in the dynamic properties of the partitioning system (8, 11, 39, 48, 49) resulting from a reduced chromosome content in an enlarged cellular compartment. Systematic analysis of chromosome organization in P. aeruginosa (42) suggests that its replication–segregation cycle could follow an alternating pattern similar to the one described here. Interestingly, this bacterium contains both condensin complexes and the Par system, and it does not anchor its origins at the cell poles. Furthermore, recent quantitative analysis of snapshot images suggests an ori-ter–like chromosome organization in fast-growing E. coli (51). The localization of the replisome to midnucleoid zones in these fast-growing cells (51) raises the possibility that a transient left-ori-right intermediate might exist during these overlapping replication cycles. Time-lapse imaging of loci on both replication arms or simultaneous imaging of three loci will be required to establish whether or not this is the case.
In the cycle we report here, the template DNA adopts a left-ori-right organization, whereas the newly replicated DNA is segregated and organized in an ori-ter pattern. We hypothesize that this oscillating pattern enhances the efficiency of replication and segregation of the bacterial chromosome. The left-ori-right pattern ensures that the two replisomes track on the template DNA in opposite cell halves (Figs. S6 and S7), providing spatial resolution to the complex topologies that inevitably arise at the replication forks. The ori-ter pattern of the newly replicated DNA ensures that the sister chromosomes are segregated as far as possible from each other. Finally, in the specific case of B. subtilis, the partitioning system also ensures that the origins reside close to the poles, where they can be readily anchored if the cell enters the sporulation pathway (24, 26). Initiation of a new round of replication transiently blocks entry into sporulation (52), and our data suggest that it also releases the origins from their polar position, allowing the replication–segregation cycle to begin again.
Finally, our data raise the possibility that the disparate patterns of chromosome organization observed in bacteria arise from modifications to a common organization–segregation mechanism. In the case of E. coli, which lacks a partitioning locus, the condensin-mediated left-ori-right pattern is established shortly after origin segregation and this ground state is maintained throughout the replication–segregation cycle (Fig. 4). In the case of C. crescentus, V. cholerae chromosome I, and sporulating B. subtilis, distinct polar anchoring mechanisms “lock” chromosome organization in the ori-ter pattern, half of the full cycle (Fig. 4). Although the removal of the polar anchoring mechanism or addition of a par locus is unlikely to be sufficient to recover the second half of the respective cycles, we propose that these factors are the principal drivers in evolving distinct organization–segregation cycles.
B. subtilis strains were derived from the prototrophic strain PY79 . Cells were grown in defined rich casein hydrolysate medium (53) or minimal medium (S750) (54) supplemented with 1% glucose, sorbitol, or succinate as specified. To monitor chromosome organization during sporulation, we used a mutant of the SpoIIIE DNA translocase (spoIIIE36) that engages the forespore chromosome after polar division but is blocked in DNA transport (28, 55). Sporulation was induced by resuspension at 37 °C according to the method of Sterlini–Mandelstam (53). Images were taken 2 h after the initiation of sporulation. E. coli SspB protein was induced with 0.5% xylose for degradation of SsrA-tagged proteins (56). Strains, plasmids, and oligonucleotides used in this study are listed in Tables S1–S3. Strain and plasmid constructions are described in SI Materials and Methods.
Fluorescence microscopy was performed with an Olympus BX61 microscope equipped with an UPLFLN 100×/1.3-N.A. phase-contrast oil objective and a CoolSnapHQ cooled CCD camera (Photometrics) or a Nikon Ti microscope equipped with Plan Apo 100×/1.4-N.A. phase-contrast oil objective and a CoolSnapHQ2 camera. Membranes were stained with trimethylammonium diphenylhexatriene (Molecular Probes) at 0.01 mM or with FM4-64 [N-(3-triethylammoniumpropyl)-4-(6-(4-(diethylamino) phenyl) hexatrienyl), pyridinium dibromide; Molecular Probes] at 3 μg/mL. DNA was stained with DAPI (Molecular Probes) at 2 μg/mL. Images were cropped and adjusted using MetaMorph software (Molecular Devices). Final figure preparation was performed in Adobe Illustrator (Adobe Systems).
For snapshot imaging, cells were immobilized using 2% (wt/vol) agarose pads containing growth media. For time-lapse imaging, a glass-bottomed dish (Willco dish HBSt-5040; Willco Wells) was used as a coverslip. Exponentially growing cells were concentrated at 3,300 × g for 30 s. After removal of 90% of the supernatant, 2 μL of the culture was spotted onto the glass-bottomed dish. A 2% (wt/vol) agarose pad containing growth media was then laid on top of the bacteria. These cells were imaged on a Well Plate Holder stage (TI-SH-W; Nikon) equipped with a humid, temperature-controlled incubator (TC-MIS; Bioscience Tools). The upper face of the pad was fully exposed, allowing adequate oxygen for growth. The objective was heated using a Bioptechs objective heater system. Images were acquired every 5 or 10 min as specified. Image analyses were performed using the MathWorks MATLAB-based program MicrobeTracker (27). Details of image analysis can be found in SI Materials and Methods.
We thank members of the Bernhardt laboratory and the laboratory of D.Z.R. as well as Viknesh Sivanathan for stimulating discussions and support, and we thank Alan Grossman, Kevin Griffith, Dirk Landgraf, David Sherratt, and Rodrigo Reyes-Lamothe for plasmids. Support for this work comes from National Institutes of Health Grants GM086466 and GM073831 (to D.Z.R.). X.W. was a long-term fellow of the Human Frontier Science Program. P.M.L. is a Helen Hay Whitney postdoctoral fellow.
Author contributions: X.W. and D.Z.R. designed research; X.W. performed research; X.W. and P.M.L. contributed new reagents/analytic tools; X.W. and D.Z.R. analyzed data; and X.W. and D.Z.R. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
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