The ParABS system is critical for survival and exhibits specific localization patterns in B. bacteriovorus

The B. bacteriovorus genome encodes homologs of ParA and ParB and harbors two parS sites near the chromosomal origin, indicating the presence of a ParABS system for chromosome segregation (Fig 1B). All our attempts to individually delete parA_Bb and parB_Bb were unsuccessful (86 and 90 screened recombinants, respectively, see Methods), underscoring that the ParABS system is likely essential in this organism under the tested conditions. We have shown previously that ParB_Bb labeled with the fluorescent protein mCherry localizes as foci that mark the multiple copies of chromosomal oriC during their partitioning in the growth phase (GP) [47], consistent with the conserved role of ParB in chromosome segregation. However, the localization of ParB_Bb appeared to be cell-cycle regulated in B. bacteriovorus [47], whereas all ParB proteins described in other species permanently cluster at the centromere [7,35,37,55–65]. Using a ParB_Bb-msfGFP fusion produced as a single copy from the native chromosomal locus, we confirmed that ParB_Bb forms foci only during the growth phase of the predator cell cycle (Fig 1C, S1 Video). Conversely, the ParB_Bb-msfGFP fluorescence signal was weak and diffuse in the cytoplasm in non-replicating cells, i.e., in the free attack-phase (AP) cells (Fig 1D) or during the time window corresponding to the G1-S transition upon prey invasion [47] (Fig 1C).

To obtain more insights into the functioning of the ParABS system during the cell cycle of B. bacteriovorus, we constructed a functional ParA_Bb-msfGFP fusion (S1A Fig) produced natively as a single copy and monitored its subcellular localization. We found that ParA_Bb-msfGFP co-localized with the nucleoid in non-replicative cells (Fig 1E), in agreement with the non-specific DNA binding of ParA proteins [16,17]. Unlike in other species with a polarly localized oriC, ParA_Bb-msfGFP localization was not visibly biased towards a particular cell pole, which we were able to distinguish using the invasive cell pole marker RomR-mCherry [47,66] (S1B Fig). During the B. bacteriovorus proliferative phase inside the prey, ParA_Bb-msfGFP displayed a dynamic localization pattern reminiscent of the DNA-bound ParA “cloud” characterized in other organisms (Fig 1F, S2 Video), which pulls apart duplicated ParB·parS partitioning complexes via repeated ParA-ParB interactions [19,67]. In addition, we detected an interaction between ParA_Bb and ParB_Bb in a bacterial two-hybrid assay (S1C Fig) and in a POLAR recruitment assay (S1D–S1F Fig) [68], suggesting a similar interplay between these proteins in B. bacteriovorus. However, in contrast to bacteria that segregate only two copies of their chromosome, several ParA_Bb-msfGFP accumulations were present at the same time and moved dynamically along the filamentous predator cell, shifting from one subcellular region to another (Fig 1F, S2 Video). Using a strain carrying both fluorescently labeled ParA_Bb and ParB_Bb (S1G Fig), we observed that ParB_Bb-mCherry foci were mainly located at the edges of ParA_Bb-msfGFP clouds (Fig 1G, arrowheads, S3 Video). This complex localization pattern is consistent with the idea that the B. bacteriovorus ParABS system is adapted to drive the simultaneous or sequential segregation of multiple chromosome copies.

A fine-tuned balance of parA_Bb and parB_Bb expression underlies proper chromosome segregation and cell cycle progression in B. bacteriovorus

Remarkably, the ParB_Bb-associated fluorescence seemed to drop at the end of the cell cycle, unlike ParA_Bb-msfGFP (see the last time points in Fig 1C, 1F). Thus, our data suggest that these key players of the ParABS system undergo an unknown regulation resulting in distinct changes in protein abundance throughout the predatory cell cycle. To delve deeper into the cell-cycle control of the ParABS system in B. bacteriovorus, we first investigated the regulation of its expression. RT-PCR shows that parA_Bb and parB_Bb are part of the same operon (Figs 2A and S2A). Furthermore, their expression is cell-cycle regulated as the corresponding mRNA was detected mainly in GP and not in AP (Figs 2A and S2A), in agreement with previous reports [69,70].

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Fig 2. Biphasic expression of the parAB_Bb operon during the B. bacteriovorus cell cycle is critical for proper chromosome segregation.

(A) Biphasic expression of the parAB_Bb operon. Top: schematic representation of the B. bacteriovorus parAB and dnaK (housekeeping gene) genomic context; graphical illustrations depict the cell cycle stages corresponding to the indicated time points. The fragments corresponding to the parA-parB junction or inside the dnaK gene amplified by RT-PCR are indicated below (AB, 400 bp; K, 400 bp). RNA samples were isolated from wild-type B. bacteriovorus in AP, 1 h, and 3 h after mixing with prey. Bottom: agarose gel analysis of the indicated fragments amplified by PCR from cDNA (top panel), genomic DNA (middle panel, positive control), and total RNA (bottom panel, negative control, ensuring RT-PCR bands result from the cDNA and not from potential DNA contamination in the RNA sample). Molecular weight markers (in kb) are indicated on the left. (B) Balanced parA_Bb:parB_Bb expression is essential for proper chromosome partitioning. Left: phase contrast and fluorescence images of representative AP cells stained with DAPI. The schematics illustrate the presence of a plasmid allowing the constitutive expression (from the PnptII promoter) of the indicated gene(s) parB_Bb (GL1261), parA_Bb (GL1460), or both (GL1004) in an otherwise WT background. Cells with abnormal nucleoid areas are observed when parA_Bb (parA_Bb⁺⁺) or parB_Bb (parB_Bb⁺⁺) is overexpressed, but to a lower extent when both are overexpressed (parAB_Bb⁺⁺). Scale bar, 1μm. Right: violin plot of cell length, cell area, and nucleoid area distributions in the same cells. The lines indicate the 25, 50, and 75 percent quantiles from bottom to top. Mean, standard deviation and coefficient of variation (CV) values are shown on top of the corresponding plot. n indicate the number of cells analyzed in a representative experiment. All experiments were performed at least twice. See also S2 Fig.

https://doi.org/10.1371/journal.pgen.1010951.g002

Consistent with the significance of the cell cycle-dependent parAB_Bb expression in B. bacteriovorus, we previously found that the constitutive expression of parB_Bb (parB_Bb⁺⁺) negatively impacts cell cycle progression and chromosome segregation [47]. Altering the native parA_Bb expression similarly impaired cell cycle progression and ori partitioning, mimicking the parB_Bb⁺⁺ phenotype (Figs 2B and S2B–S2C). Remarkably, simultaneous constitutive expression of both parA_Bb and parB_Bb (from the same promoter) decreased the severity of the single overexpression phenotypes (Figs 2B and S2B–S2D). Thus, our data show that the ParABS system is under transcriptional or post-transcriptional control during the B. bacteriovorus cell cycle, resulting in phase-dependent abundance of the transcript encoding both parA_Bb and parB_Bb. This level of regulation is crucial for proper chromosome segregation and cell cycle progression, likely by contributing to a fine balance between both partners.

The protein levels of ParA_Bb and ParB_Bb vary differently during the predatory cell cycle

Since ParA_Bb and ParB_Bb fluorescence profiles hinted at distinct relative protein abundance despite being expressed from the same transcript, we sought to obtain detailed insights into ParA_Bb and ParB_Bb protein levels during the cell cycle. Western blot using an antibody against ParB_Bb shows that the endogenous (untagged) ParB_Bb protein levels vary during the cell cycle, being almost unnoticeable in AP but present in higher amounts in GP (Figs 3A, S3A and S6A), mirroring the fluorescence signal of the natively produced ParB_Bb-msfGFP fusion (Fig 1C). Accordingly, single-cell analysis of the ParB_Bb-msfGFP fluorescence intensity profile over time indicates that ParB_Bb proteins accumulate during cell growth but rapidly drop when B. bacteriovorus divides into multiple daughter cells (Fig 3B, magenta, S3B Fig). Notably, the levels of natively produced ParB_Bb-msfGFP or ParB_Bb-mCherry protein fusions, detected with the same anti-ParB_Bb antibody, fully reflected the endogenous untagged ParB_Bb profile during a synchronized B. bacteriovorus cell cycle (Figs 3A, S3C and S6A), confirming that these fusions are reliable reporters of the native ParB_Bb protein.

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Fig 3. The levels of ParB_Bb and ParA_Bb vary differently during the cell cycle.

(A) ParB_Bb protein levels are cell-cycle dependent. Western blots of whole-cell protein extracts from WT B. bacteriovorus were probed with an anti-ParB_Bb antibody, as represented in the schematics. WT protein samples were isolated at time points throughout the predatory cell cycle: 1h, 2h, 3h, 4h, 5h after mixing with prey (GP), and AP. All samples were run on the same gel; for enhanced figure clarity, sample positions have been rearranged, as indicated by the dashed line. Graphical illustration of the cell cycle stages corresponding to the indicated time points. Arrowhead points to ParB_Bb (detected only during GP). Ponceau staining served as a loading control and quantification from full lanes (displayed in S6A Fig) was used to normalize band intensities. Quantifications are shown as the means of independent triplicates. Error bars represent standard deviations. All replicates are shown in S6A Fig. (B) ParB_Bb-msfGFP fluorescence intensity level drops during cell division while ParA_Bb-msfGFP fluorescence intensity levels remain high. B. bacteriovorus parA_Bb::parA_Bb-msfgfp parB_Bb::parB_Bb-msfgfp (GL2154) was mixed with prey and imaged in time-lapse after 75 min with 8-min intervals. Top: overlay of phase contrast and fluorescence images for ParA_Bb-msfGFP and ParB_Bb-mCherry for selected late time points; Bottom: mean fluorescence intensities of msfGFP (in green) and mCherry (in magenta) plotted over time for the same cell. The time window corresponding to the images displayed on top starts at the dashed line. ParB_Bb-mCherry fluorescence intensity decreases during this interval, unlike ParA_Bb-msfGFP. (C) ParA_Bb protein levels do not decrease in the attack phase. Western blots of whole-cell protein extracts from B. bacteriovorus parA_Bb::parA_Bb-msfgfp (GL2134) probed with an anti-msfGFP antibody, as represented in the schematics. GL2134 protein samples are isolated at time points throughout the predatory cell cycle: 1h, 2h, 3h, 4h, and 5h after mixing with prey (GP), and AP. All samples were run on the same gel; for enhanced figure clarity, sample positions have been rearranged, as indicated by the dashed line. Graphical illustration of the cell cycle stages corresponding to the indicated time points. Arrowhead points to ParA_Bb-msfGFP (detected also in the AP). Ponceau staining served as a loading control and quantification from full lanes (displayed in S6B Fig) was used to normalize band intensities. Quantifications are shown as the means of independent triplicates. Error bars represent standard deviations. All replicates are shown in S6B Fig. (D) Top: Western blots of whole-cell protein extracts from AP cells of B. bacteriovorus WT, parB_Bb::parB_Bb-msfgfp (ParB_Bb, GL1654) and parA_Bb::parA_Bb-msfgfp (ParA_Bb, GL2134) strains were probed with an anti-msfGFP antibody. Only ParA_Bb-msfGFP protein is detected in AP. Bottom: quantifications are shown as the means of independent triplicates, normalized as in A and C. Error bars represent standard deviations. All replicates and Ponceau staining are shown in S6C Fig. (E) Overproduction of ParB_Bb-msfGFP does not lead to the parS-bound focus in AP. Left: representative phase contrast and fluorescence images of AP cells of a WT B. bacteriovorus strain constitutively producing ParB_Bb-msfGFP (GL1003). Right: histogram representing the percentage of cells with zero, one, or two ParB_Bb-msfGFP foci in the same strain; n indicates the number of cells analyzed in a representative experiment. Scale bars are 1 μm. All experiments were performed at least twice. See also S3 Fig.

https://doi.org/10.1371/journal.pgen.1010951.g003

Unlike ParB_Bb, ParA_Bb-msfGFP was detected throughout the cell cycle, including in AP (Figs 3C–3D, S3D and S6B–S6C), consistent with the corresponding fluorescence signal (Fig 1F). ParA_Bb-msfGFP fluorescence intensity remained high during the last cell cycle stages, including cell division (Fig 3B, green, S3E Fig). These results show that, besides cell-cycle regulated expression, the ParABS system in B. bacteriovorus is subjected to an additional level of control that results in distinct ParA_Bb and ParB_Bb protein levels at different cell cycle stages. While in vitro studies showed that the ParA:ParB balance is important for efficient partitioning [71,72], this is the first evidence of cell-cycle-dependent changes in the relative levels of ParA and ParB proteins.

The formation of ParB_Bb·parS complexes is inhibited in non-replicative predator cells

In addition to the sudden decrease of ParB_Bb protein levels at the end of the GP, the subcellular localization of ParB_Bb fluorescent fusions also changed, shifting from clear foci to diffuse localization (Figs 1C and 3B). However, both events could not be easily distinguished due to their occurrence within a relatively short time window and the low fluorescence of the natively produced ParB_Bb fusion at that stage. Therefore, we sought to uncouple the ability of ParB_Bb to form foci (due to parS binding and spreading on adjacent DNA) from ParB_Bb protein levels by constitutively expressing parB_Bb-msfgfp. In these cells, the fluorescence signal (S3F Fig) and protein levels (S3G Fig) of ParB_Bb-msfGFP in AP were stronger than upon expression from the native locus. Still, overproduced ParB_Bb-msfGFP was diffuse in AP cells (Fig 3E), consistent with our previous results with ParB_Bb-mCherry [47]. Furthermore, these cells displayed the same phenotypes observed when the untagged ParB_Bb is constitutively produced (i.e., morphological and chromosome segregation defects; S3H, S2B Figs), further supporting the relevance of these ParB_Bb fusions. Thus, a third level of control is applied to ParB_Bb in B. bacteriovorus to prevent its accumulation at the centromere, regardless of its protein level.

ParB_Bb specifically accumulates on B. bacteriovorus parS in a CTP-dependent manner

The formation of partitioning complexes in other species requires both the initial binding and the spreading of ParB on the DNA [8,10,11,13,14,73]. To investigate ParB_Bb’s ability to perform these two critical functions, we assessed its capacity to bind and spread from parS and to hydrolyze CTP in vitro. The two parS sequences in the proximity of the B. bacteriovorus chromosomal oriC (parS_Bb) are identical and closely resemble the parS consensus (S4A Fig) [2]. In a biolayer interferometry assay, purified ParB_Bb bound a linear DNA substrate carrying parS_Bb, and the addition of CTP triggered ParB_Bb sliding and falling off (marked by the higher Kd in the presence of CTP; Fig 4A), like other characterized ParB homologs [11,13–15,73]. Moreover, ParB_Bb accumulated on a closed DNA loop containing parS_Bb in a CTP-dependent manner, reflecting the capacity of ParB_Bb to bind parS_Bb and escape onto neighboring DNA (Fig 4B). This accumulation was specific to the cognate parS_Bb, as no binding was observed when we used a scrambled parS sequence (nonS; Fig 4B) or parS from Caulobacter crescentus (parS_Cc; Fig 4B). Finally, measurements of inorganic phosphate release showed that ParB_Bb hydrolyses CTP but no other NTPs in the presence of parS_Bb (S4B Fig). Thus, the B. bacteriovorus ParB protein is a bona-fide parS-binding CTPase. Consistent with those in vitro data, ParB_Bb specifically accumulated on parS_Bb in vivo in E. coli (which lacks an endogenous ParABS), used as a heterologous system. Indeed, ParB_Bb-msfGFP formed foci when produced in E. coli, only in the presence of a plasmid carrying parS_Bb and not when the plasmid carried the non-cognate parS_Cc (Figs 4C and S4C). Interestingly, ParB from Caulobacter crescentus (ParB_Cc) also failed to form a focus and was diffuse in the cytoplasm when ectopically produced in AP B. bacteriovorus cells (Fig 4D), even though ParB_Cc can bind parS_Bb in vitro (S4D Fig) and in E. coli (S4E Fig). Therefore, the absence of ParB_Bb·parS_Bb complex in non-replicative B. bacteriovorus cells is unlikely to result from an unusual functionality of the ParB_Bb protein itself.

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Fig 4. ParB_Bb is a CTPase with a specific affinity for its cognate parS_Bb.

(A) CTP reduces the binding of ParB_Bb on parS_Bb on a linear substrate. Left: schematics of the Streptavidin (SA)-coated probe and the linear 40-bp DNA substrate containing the cognate parS_Bb sequence. Right: ParB_Bb titration curves in the presence or absence of CTP obtained from biolayer interferometry (BLI) analysis (see B and Material and Methods). All reactions contained 1 mM CTP (when appropriate), 0.5 μM 40 bp parS_Bb DNA, and an increasing concentration of ParB_Bb. The BLI experiment was done in triplicates; data were fitted to calculate the binding affinity constant Kd (nM); Kd = 134 ± 23 nM for the 40 bp parS_Bb, and Kd = 5500 ± 397 nM for the 40 bp parS_Bb + 1mM CTP. In this assay, ParB_Bb spreading displaces it from the short linear parS fragments by pushing it into the solution. The higher Kd constant (lower parS affinity) in the presence of CTP thus indicates CTP-induced spreading of ParB_Bb. (B) ParB_Bb accumulation at parS_Bb is specific and CTP-dependent. Left: schematics of the SA-coated probe and a closed DNA substrate with a parS sequence, allowing ParB accumulation through parS binding and CTP binding-dependent spreading. CTP hydrolysis removes ParB from the DNA. Right: BLI analysis of the interaction between 1 μM ParB_Bb-6xhis in the presence or absence of 1 mM CTP and a 180 bp DNA substrate containing the cognate parS_Bb (blue), scrambled nonS (black) or non-cognate parS_Cc (grey) sequence. The BLI probe was dipped into a buffer-only solution (base), then into a premix of protein +/- CTP (association), and finally returned to a buffer-only solution (dissociation). Each BLI experiment was done in triplicate, and a representative sensorgram is presented. (C) ParB_Bb protein can bind to its cognate parS sequence in E. coli. Representative phase contrast and fluorescence images of E. coli carrying a plasmid with parS_Bb and allowing constitutive production of ParB_Bb-msfGFP (GL1669; plasmid schematics on the left). The scale bar is 2 μm. See Fig 5D for histogram representing the percentage of cells with zero, one, or two ParB_Bb-msfGFP foci in the same strain. (D) A constitutively produced fusion of Caulobacter crescentus ParB (ParB_Cc) to msfGFP does not form foci in AP B. bacteriovorus cells. Left: representative phase contrast and fluorescence images of AP cells of a WT strain constitutively producing ParB_Cc-msfGFP from a plasmid (GL2108; schematics on the left). The fluorescence signal shows the partial nucleoid exclusion pattern of freely diffusing cytosolic proteins, characteristic of AP B. bacteriovorus cells ¹. Right: histogram of the percentage of cells with zero, one, or two ParB_Cc-msfGFP foci in the same strain; n indicates the number of cells analyzed in a representative experiment. The scale bar is 1 μm. See also S4 Fig.

https://doi.org/10.1371/journal.pgen.1010951.g004

The parS_Bb chromosomal context prevents ParB_Bb accumulation in non-replicative cells

Having established that ParB_Bb activities are not per se drastically different than previously characterized ParB homologs, we investigated the possibility that in AP cells, the chromosomal context of the centromere is incompatible with ParB_Bb accumulation. Moving one or both parS_Bb sites to a remote location on the chromosome is expected to result in pleiotropic effects due to a major perturbation of segregation, which would prevent unambiguous interpretation. Instead, we addressed whether the genomic context impacts ParB_Bb localization by cloning a parS_Bb site on a replicative plasmid and constitutively producing ParB_Bb-msfGFP from the same vector in B. bacteriovorus (Fig 5A). Remarkably, we found that ParB_Bb-msfGFP localizes as clear foci in AP cells only when the plasmid carries the intact parS_Bb. No focus was observed when we used a mutated parS_Bb* as a control (S5A Fig). These results indicate that in AP cells, ParB_Bb has the capacity to bind parS_Bb specifically but only when parS_Bb is out of its native chromosomal context. Moreover, AP B. bacteriovorus cells carrying a parS_Bb + parB_Bb-msfgfp⁺⁺ vector did not exhibit the sick phenotype observed in parB_Bb⁺⁺ cells (S5B Fig, compared with S2C Fig), presumably because the plasmidic parS_Bb titrates the excess ParB_Bb present in the cytoplasm, thereby alleviating the negative effect associated with its overproduction. Hence, our data show that the chromosomal context is an essential factor underlying the unique subcellular distribution of ParB_Bb in B. bacteriovorus.

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Fig 5. The genomic context of parS_Bb plays a role in ParB_Bb localization.

(A) Changing the native context of the parS_Bb enables ParB_Bb binding. Left: representative phase contrast and fluorescence images of AP cells of WT B. bacteriovorus strain constitutively producing ParB_Bb-msfGFP from a plasmid carrying a copy of the parS_Bb sequence (GL1751). Right: histogram representing the percentage of cells with zero, one, or two ParB_Bb-msfGFP foci in the same strain. (B-C) Short chromosomal regions around each parS_Bb site are enough to prevent ParB_Bb from clustering on the centromere. Top: schematic representation of the parAB operon in B. bacteriovorus and the upstream parS sequences; chromosomal regions around parS_Bb sequences (“extended parS”) cloned in the plasmids are indicated; plasmid schematics are shown on the right. Bottom: representative phase contrast and fluorescence images of AP cells of WT strain constitutively producing ParB_Bb-msfGFP from a plasmid carrying parS_1extended (B, GL1749) or parS_2extended (C, GL1925); histogram representing the percentage of cells with zero, one, or two ParB_Bb-msfGFP foci in the same strain. Scale bars are 1 μm. (D) The genomic context of the parS_Bb prevents ParB_Bb focus formation only in B. bacteriovorus. Top: representative phase contrast and fluorescence images of E. coli strains carrying plasmids as in Figs 3D, 5A–5C, constitutively producing ParB_Bb-msfGFP in the absence of parS (GL1661), or in the presence of parS_Bb (GL1669), parS_1extended (GL1737) or parS_2extended (GL1899), respectively. Schematics illustrate the parB_Bb–msfgfp expression plasmid with or without (extended) parS sequences. Bottom: histogram representing the percentage of cells with zero, one, or two ParB_Bb-msfGFP foci in the same strains. The scale bar is 1 μm. n indicates the number of cells analyzed in a representative experiment; all experiments were performed at least twice. See also S5 Fig.

https://doi.org/10.1371/journal.pgen.1010951.g005

To obtain clues on the difference between the chromosomal and plasmidic parS_Bb leading to distinct ParB_Bb localization patterns, we tested the impact of the parS_Bb surrounding regions. We constructed two plasmids carrying the parS_Bb sequence flanked by their short upstream and downstream chromosomal regions (named here parS1 extended and parS2 extended, spanning 242 bp and 330 bp, respectively, Fig 5B–5C). Strikingly, the overproduced ParB_Bb-msfGFP failed to form a fluorescent focus in both cases (Fig 5B–5C). Instead, the signal was diffuse in AP cells, which exhibited the characteristic parB_Bb⁺⁺ phenotype (S5B Fig, compared with S2C Fig). On the contrary, ParB_Bb could form a fluorescent focus in the same strains later during growth (S5D–S5E Fig). The effect of the DNA adjacent to parS sites was limited to B. bacteriovorus since in E. coli, ParB_Bb-msfGFP localized as foci despite the addition of parS1 or parS2 extended regions (Fig 5D). Therefore, our results suggest that a Bdellovibrio-specific factor makes the chromosomal environment of parS_Bb sites incompatible with ParB_Bb clustering during the non-proliferative phase of the cell cycle.

Intricate subcellular dynamics of ParA_Bb and ParB_Bb during the replicative stage of the cell cycle

We investigated the localization of ParA_Bb in B. bacteriovorus during the predatory cell cycle using a functional, natively produced fluorescent fusion. The subcellular localization of ParA has been observed in Streptomyces species, which also feature a polyploid growth stage [74,75]. While the chromosome is copied multiple times during vegetative growth in these bacteria, the ParABS system only acts later in a distinct developmental stage to separate nucleoids in future spores [76]. Thus, our data provide the first visualization of the ParA protein in a bacterium in which segregating multiple chromosome copies is an integral part of the cell cycle, rather than a conditional process as in Streptomyces. In non-replicative cells, ParA_Bb is uniformly distributed on the nucleoid, in contrast with the ParA gradient observed in other species in which oriC is localized at one cell pole [3]. The absence of a ParB_Bb·parS nucleoprotein complex at the centromere-like region (see below) might explain this non-polarized ParA_Bb pattern in AP B. bacteriovorus cells. Furthermore, our observations of ParA_Bb and ParB_Bb during chromosome replication and partitioning in the filamentous predator cell revealed several dynamic ParA_Bb “clouds” between segregating ParB_Bb foci, demonstrating that the ParABS system segregates multiple chromosomal copies simultaneously in the B. bacteriovorus cell.

In contrast to binary-dividing species like C. crescentus, the partitioning complexes do not move from one pole to another across the entire predator cell. Instead, ParB_Bb foci adopt a clear “beads-on-a-string” pattern ([47] and this study), reminiscent of the ParABS-mediated positioning of low-copy plasmids at non-polar locations [77]. It is still unclear how the segregation of each chromosome is spatially confined within the filamenting B. bacteriovorus cell body (which is filled with other replicating and segregating copies of the chromosome) to achieve this regular distribution [47]. We found that perturbations in the ParABS system, such as alterations of the ParA_Bb:ParB_Bb balance, prevent this exquisite arrangement of newly synthesized centromeres. In other bacteria, specific hub proteins at the cell poles polarize the choreography of ParABS-dependent segregation [29], but no homologs of these proteins were identified in B. bacteriovorus. Future research will determine whether this bacterium uses polar landmarks or other mechanisms to spatially organize chromosome partitioning along the mother cell.

Coupling centromere organization to cell cycle progression

Our previous work uncovered a striking difference between the ParB protein in B. bacteriovorus and those characterized in other species. Whereas ParB homologs always localize at the parS site [7,35,37,55–65], ParB_Bb is unable to accumulate at the chromosomal centromere in non-replicative cells, independently of its protein levels [47]. This study offers unambiguous support to this finding by showing that (i) the unique on-off localization pattern of ParB_Bb is independent of the fluorescent tag, and (ii) the protein fusions used to monitor ParB_Bb subcellular localization and abundance represent reliable reporters of the native protein during the entire B. bacteriovorus cell cycle. Furthermore, we identified ParB_Bb as a bona-fide CTPase like several other ParB homologs [10,13,73,78], which strongly suggests that ParB_Bb accumulation at the centromere results from its binding on parS and CTP-dependent sliding on adjacent DNA, while subsequent CTP hydrolysis unloads the protein.

Remarkably, we found that the absence of ParB_Bb·parS complex depends on the chromosomal context of parS, as ParB_Bb formed a focus in B. bacteriovorus AP cells when parS was present on a plasmid. This result indicates that fluctuations in the cellular concentration of CTP, if any, are not responsible for the cell cycle-dependent localization of ParB_Bb. It also hints that ParB_Bb has the capacity to bind and spread from parS in the absence of ongoing chromosome replication, even though the first ParB_Bb focus systematically appears after DNA replication initiation in GP cells [47]. Moreover, our data reveal that the direct chromosomal context of parS sites has a major role in preventing ParB_Bb complex formation, as ParB_Bb lost its ability to accumulate on the plasmidic parS when this sequence was flanked by short DNA stretches adjacent to parS1 or parS2 on the chromosome. This intriguing effect was only observed in B. bacteriovorus, and not in E. coli cells. Therefore, we propose that the local topology of the DNA in the immediate vicinity of the parS sequences, possibly induced by B. bacteriovorus-specific proteins during the GP-AP transition, might interfere with parS binding and/or the spreading of ParB_Bb. Identifying the specific inhibition that prevents ParB_Bb from accumulating at the centromere in a cell-cycle and chromosomal context-dependent manner will certainly reveal additional complexity in bacterial centromere organization.

Cell cycle-dependent fluctuations of the ParA and ParB protein balance

The ParA_Bb:ParB_Bb balance appears to be crucial for B. bacteriovorus cell cycle progression, as constitutive production of ParA_Bb or ParB_Bb throughout the cell cycle led to cellular dysfunction, unless both proteins are overproduced. It is likely that the excess of either protein interferes with their fine-tuned interplay during the chromosome segregation process, resulting in the observed phenotypes. The compensatory effect of the double ParA_Bb and ParB_Bb overproduction is consistent with this idea.

Intriguingly, our results also hint that ParA_Bb and ParB_Bb protein levels are differentially controlled in the non-replicative stage of the B. bacteriovorus cell cycle and suggest that ParB_Bb is specifically degraded in a cell cycle-dependent manner. Indeed, although the co-expression of parA_Bb and parB_Bb is turned off during AP, protein concentration is not expected to change between the mother cell and the newborn cells upon cell division. Yet, ParB_Bb protein levels dropped within minutes at the GP-AP transition, becoming undetectable in the AP, unlike ParA_Bb. Whereas there has been no report of a mechanism regulating ParB degradation in other bacteria, a recent study identified ParB as a candidate substrate of the conserved Lon protease in C. crescentus [79]. Future investigation will be needed to reveal the potential proteolytic mechanism that depletes ParB_Bb at the end of the B. bacteriovorus growth phase.

While it is clear that ParB_Bb levels are tightly regulated during the B. bacteriovorus cell cycle, the physiological role of the disappearance of ParB_Bb at the GP-AP transition remains to be determined. We hypothesize that in addition to the importance of proper ParA_Bb and ParB_Bb levels during growth to ensure accurate chromosome segregation, B. bacteriovorus benefits from clearing the cell from ParB_Bb in the AP. This idea is rationalized by the incapacity of ParB_Bb to cluster at the chromosomal oriC during that stage, which would result in the entire ParB_Bb pool being free in the cytoplasm in the absence of degradation. We propose that such excess of cytosolic ParB_Bb in AP cells introduces unknown deleterious effects, since we observed that the ParB_Bb overproduction phenotype was largely alleviated when the protein could form a parS-bound cluster on a plasmid in AP cells.

Although studies in Streptomyces coelicolor have shown developmental stage-specific modulation of ParA and ParB protein levels resulting from the differential control of parAB transcription [41], to the best of our knowledge, this is the first report of fluctuating ParA:ParB protein ratios across the lifecycle of a bacterium. Altogether, our study sheds new light on the intricate dynamics and adaptation of the conserved ParABS system in bacteria. Our work hints that investigating the ParABS system in organisms with distinctive lifestyles will unveil new levels of complexity in centromere organization, chromosome segregation, and cell cycle control.

Strains

The strains and plasmids used in this study are listed in S1 Table, along with their respective construction methods (S2 Table). Standard molecular cloning techniques were employed, and DNA assembly was conducted using the NEBuilder HiFi mix from New England Biolabs. All oligos used in the study can be found in S3 Table. The B. bacteriovorus strains were produced from the wild-type HD100 strain, while the E. coli strains employed as prey were generated from MG1655. Microscopy, conjugation, bacterial two-hybrid, and POLAR assay were conducted utilizing E. coli MG1655, S17-λpir, BL21, BTH101, and TOP10, CC118 λpir, TB28, respectively. All plasmids were introduced into B. bacteriovorus through mating, following the methodology described below. Scarless allelic replacements into the HD100 chromosome were achieved via a two-step recombination approach using a pK18mobsacB-derived suicide vector. The overexpression experiments were performed using the low-copy plasmid pTNV215 (RSF1010 replicon) as a backbone [47]. Chromosomal modifications were screened by PCR and verified by Sanger DNA sequencing, following the protocol detailed in [47].

Routine culturing of B. bacteriovorus and E. coli

E. coli strains were grown in LB medium. For imaging experiments, overnight starter cultures from single colonies were diluted at least 1:500 in fresh medium until exponential phase. B. bacteriovorus strains were grown as described in DNB medium (Dilute Nutrient Broth, Becton, Dickinson, and Company) supplemented with 2 mM CaCl₂ and 3 mM MgCl₂ salts in the presence of E. coli prey at 30°C with continuous shaking [80]. When appropriate, antibiotic-resistant E. coli strains were used as prey for overnight culturing of the corresponding antibiotic-resistant B. bacteriovorus. When required, kanamycin, gentamycin, ampicillin, chloramphenicol, or tetracycline was added in liquid and solid media at 50 μg/ml, 10 μg/ml, 50 μg/ml, 15 μg/ml, and 7.5 μg/ml respectively.

Plasmid conjugation by mating

Mating was carried out between the E. coli S17-λpir donor strain carrying the plasmid to be conjugated and the B. bacteriovorus receiver strain, following the protocol detailed in [47]. The exponentially growing E. coli donor strains were harvested and washed twice in DNB medium before resuspending in DNB salts at a 1:10 ratio of the initial volume. This donor suspension was combined with an equal volume of fresh overnight lysate of a receiver HD100 strain. The mating mixture was then incubated for a minimum of 4 hours at 30°C with shaking before being plated on a selective medium using the double-layer technique [80]. Single plaques were isolated, and transconjugants were validated by microscopy (when applicable), PCR, and sequencing.

RT-PCR

A predator: prey mixture (using a 1:1 volume ratio, which minimizes the excess of free attack phase cells) was prepared and incubated for several hours with shaking at 30°C. RNA samples were collected at different times during synchronous predation to track the expression of parA_Bb (bd3906), parAB_Bb (bd3905-bd3906), parB_Bb (bd3905) and dnaK (bd1298) throughout the predatory cycle of B. bacteriovorus on E. coli prey. A NucleoSpin RNA kit (MACHEREY-NAGEL) or SV Total RNA Isolation System (Promega) was used to isolate RNA, following the manufacturer’s instructions. An additional TURBO DNase (Invitrogen) treatment was performed for 1 hour at 37°C following manufacturer’s instructions. RNA quality was assessed by measuring the 260/280 nm and 260/230 nm absorbance ratios. Reverse transcription (RT) and PCR were carried out utilizing the QIAGEN OneStep RT-PCR kit with the following thermocycling parameters: 50°C for 30 minutes, 94°C for 15 minutes, followed by 30 cycles of 94°C for 1 minute, 50°C for 1 minute, and 72°C for 1 minute, with a final step of 72°C for 10 minutes.

Bacterial-two hybrid assay (BACTH)

Interaction between protein pairs was assessed by the adenylate cyclase-based bacterial two-hybrid technique, as detailed in [81]. Briefly, the proteins of interest were merged with the isolated T18 and T25 catalytic domains of the Bordetella pertussis adenylate cyclase. The two plasmids producing the fusion proteins were introduced into the BTH101 reporter strain, and co-transformants were incubated on a selective medium overnight at 30°C. A single colony from each co-transformation was inoculated into 400 μl LB medium supplemented with ampicillin (200 μg/ml), kanamycin (50 μg/ml), and IPTG (0.5 mM). After incubation overnight at 30°C, 5 μl of each culture was placed onto LB plates supplemented with ampicillin, kanamycin, IPTG (using the abovementioned concentrations), and X-gal (40 ng/μl) and incubated at 30°C. An interaction assay with pKT25-Zip and pUT18C-Zip, two Zip protein domains, was included as a positive control. The experiments were conducted in triplicate, and representative results are presented.

PopZ-linked apical recruitment (POLAR) assay

Protein-protein interaction between ParA_Bb and ParB_Bb was assessed using the POLAR assay, similarly as in [68]. Briefly, proteins of interest were fused to msfGFP-H3H4 or mScarlet to generate the bait and prey protein fusions, respectively. Bait- and prey-encoding plasmids, cloned in E.coli TOP10 and CC118 λpir, respectively, were introduced in electrocompetent E. coli TB28 cells containing the temperature-sensitive helper plasmid pAH69. Selection of co-transformants and loss of pAH69 were allowed by overnight incubation on LB plates supplemented with chloramphenicol and tetracycline at 37°C. An overnight starter was prepared by inoculating LB medium supplemented with appropriate antibiotics with a single colony and grown at 37°C. A 6 ml fresh co-culture was prepared in LB medium with antibiotics using a 1:100 dilution and incubated at 37°C. Once OD₆₀₀ reached 0.2, cells were collected by centrifugation (5000 rpm for 2 min) and resuspended in 6 ml 1x M9 minimal medium with salts supplemented with 0.2% Casaminoacids. The 6 ml culture was then split into two 3 ml cultures, one supplemented with 100 μM IPTG for prey protein induction only and the other with both 100 μM IPTG and 0.2% arabinose to induce the expression of the prey protein, PopZ_Cc, and the bait protein. Snapshots were performed following a 2 h induction at 37°C.

Live-cell imaging

B. bacteriovorus were grown overnight with the appropriate E. coli prey, and antibiotics were used when needed. Subsequently, they were grown on wild-type MG1655 for at least one generation without antibiotics before the beginning of the imaging experiment. For snapshots of fresh AP B. bacteriovorus, the cells were deposited on 1.2% agarose pads prepared with DNB-salt media. For the time-lapse imaging of synchronous predation cycles, MG1655 E. coli cells were grown in 2TYE medium to the exponential phase (OD₆₀₀ = 0.4-0.6), harvested at 5000 x g at room temperature for 5 minutes, washed twice, and resuspended in DNB medium. E. coli and B. bacteriovorus were then combined in a 1:3 to 1:5 volume ratio to allow the majority of prey cells to be infected. This setup ensured infection of all E. coli cells, which was optimal for time-lapse imaging. The remaining free Bdellovibrio cells did not pose any issue in that setup as they neither proliferate nor can re-infect existing bdelloplasts. In all synchronous predation imaging experiments, the prey-predator mixing step was indicated as time 0. The cells were either directly deposited on DNB-agarose pads for imaging or left shaking at 30°C before imaging for the indicated durations. In time-lapse experiments, the same fields of view on the pad were imaged at regular intervals, with the enclosure temperature set to 27°C. When appropriate, prior to imaging, cells were incubated for 5 minutes with DAPI (Life Technologies) at a final concentration of 5 μg/ml for nucleoid staining experiments. Regarding snapshots of E. coli strains, overnight cultures were diluted at least 1:500 and allowed to grow to the exponential phase before being deposited on 1.2% agarose pads prepared with PBS buffer supplemented with 0.2% glucose, 0.2% casamino acids, 1 μg/ml thiamine, 2 mM MgSO₄, and 0.1 mM CaCl2), except for POLAR strains which were diluted 1:100 and imaged on 1.2% agarose pads prepared with 1x M9 minimal medium with salts following induction as described above.

Image acquisition

Phase contrast and fluorescence images were obtained using a Nikon Ti2-E fully-motorized inverted epifluorescence microscope (Nikon) equipped with a CFI Plan Apochromat l DM 100x 1.45/0.13 mm Ph3 oil objective (Nikon), a Sola SEII FISH illuminator (Lumencor), a Prime95B camera (Photometrics), a temperature-controlled light-protected enclosure (Okolab), and filter-cubes for DAPI (32 mm, excitation 377/50, dichroic 409, emission 447/60; Nikon), mCherry (32 mm, excitation 562/40, dichroic 593, emission 640/75; Nikon), and GFP (32 mm, excitation 466/40, dichroic 495, emission 525/50; Nikon). Multi-dimensional image acquisition was supervised using the NIS-Ar software (Nikon). The pixel size was 0.074 μm using the 1.5X built-in zoom lens of the Ti2-E microscope. The same LED illumination and exposure times were applied when capturing images of various strains and/or conditions in one experiment and were set to a minimum for time-lapse acquisitions to limit phototoxicity.

Image processing

To prepare figures, images were processed using FIJI [82], with contrast and brightness settings being kept consistent for all regions of interest in each figure unless stated otherwise. Denoising (Denoise.ai, Nikon) was applied to all phase contrast and fluorescence channels for Fig 1C, 1F, 1G to improve the display of time-lapse images captured with low exposure (necessary to preserve cell viability). Figures were constructed and labeled using Adobe Illustrator. B. bacteriovorus outlines were obtained with Oufti [83] for AP cells, or drawn manually in Adobe Illustrator for GP cells within bdelloplasts.

Protein overexpression and purification

Expression and purification of ParB_Bb with a C-terminal His-tag were conducted as follows. E. coli BL21(DE3) cells containing pET21, which expresses C-terminal His-tagged ParB_Bb, were cultivated at 37°C in autoinducing media supplemented with ampicillin (200 μg/ml). Following a 5-hour induction, cells were harvested by centrifugation and suspended in 35 ml lysis buffer (100 mM Tris-HCl [pH8], 300 mM NaCl, 5% glycerol (v/v), supplemented with a protease inhibitor cocktail (Complete, Roche)). The suspended cells were stored at 20°C. The frozen cells were thawed on ice and lysed using three passages through a French pressure cell at 1500 psi. The suspension was then centrifuged for 15 minutes at 40 000 x g at 4°C, and the supernatant was filtered through 0.45 μm filters before loading onto a 1 ml HisTrap column (GE Healthcare) pre-equilibrated with buffer A (100 mM Tris-HCl [pH8], 300 mM NaCl, and 5% [v/v] glycerol). The protein was eluted from the column using an increasing imidazole gradient (15–300 mM) in the same buffer. As a final purification step, size-exclusion chromatography was performed using a HiLoad 16/60 Superdex 75 column (GE Healthcare) with buffer B (100 mM Tris-HCl [pH8], 300 mM NaCl, and 5% [v/v] glycerol). The ParB_Bb-containing fractions were pooled and analyzed for purity using SDS-PAGE. Glycerol was added to the ParB_Bb fractions to a final volume of 10% (v/v), followed by 10 mM EDTA and 1 mM DTT. The purified ParB_Bb was aliquoted, snap-frozen in liquid nitrogen, and stored at –80°C.

Construction of DNA substrates for BLI assays

All DNA constructs were designed using VectorNTI (ThermoFisher) and synthesized chemically (gBlocks dsDNA fragments, IDT). To produce a linear biotinylated 40 bp DNA substrate, complementary oligos (with and without biotin) were heated at 98°C for 5 min before being left to cool down to room temperature overnight to form 50 mM double-stranded DNA. A 180 bp DNA construct was created with M13F and M13R homologous regions at each end. To produce a dual biotin-labeled DNA substrate, PCR reactions were carried out using a 2x GoTaq PCR master mix (Promega), biotin-labeled M13F and biotin-labeled M13R primers, and gBlocks fragments as a template. PCR products were separated by electrophoresis and then purified from the gel.

Measurement of protein-DNA interaction by bio-layer interferometry (BLI)

Bio-layer interferometry experiments were performed using a BLItz system equipped with Dip-and-Read Streptavidin Biosensors (Molecular Devices) as previously described [15]. The streptavidin biosensor was first equilibrated in a low-salt binding buffer (100 mM Tris-HCl (pH 8), 100 mM NaCl, 1 mM MgCl2, and 0.005% Tween 20) for at least 10 min before each experiment. Biotinylated double-stranded DNA (dsDNA) was then immobilized onto the surface of the biosensor through a cycle of baseline (30 s), association (120 s), and dissociation (120 s). During the association phase, different concentrations of ParB_Bb dimers, with or without NTPs at varying concentrations, were added to the binding buffer and allowed to associate with the immobilized DNA for 120 s. Finally, the sensor was transferred into a protein-free binding buffer to monitor the dissociation kinetics for 120 s. The sensor was recycled by dipping in a high-salt buffer (100 mM Tris-HCl (pH 8), 1000 mM NaCl, 1 mM MgCl₂, and 0.005% Tween 20) for 5 min to remove bound ParB_Bb. All sensorgrams were recorded and analyzed using the BLItz analysis software (BLItz Pro version 1.2, Molecular Devices) and replotted using DataGraph for presentation. The data were fitted using a one-site-specific binding model in GraphPad Prism Version 5.04 to calculate the binding affinity constant Kd. All experiments were conducted at least in triplicate, and a representative sensorgram is presented in each figure.

DNA preparation for EnzCheck phosphate assay

A 20 bp palindromic single-stranded DNA fragment (parS_Bb: GGATGTTCCACGTGGAACATCC or parS_Cc: GGATGTTTCACGTGAAACATCC; 100 mM in 1 mM Tris-HCl pH 8.0, 5 mM NaCl buffer) was heated at 98°C for 5 min before being left to cool down to room temperature overnight to form 50 μM double-stranded DNA. The parS_Bb and parS_Cc sequences are underlined.

Measurement of NTPase activity by EnzCheck phosphate assay

The EnzCheck Phosphate Assay Kit from Thermo Fisher was used to measure NTP hydrolysis. A reaction buffer containing 1 mM NTP and 1 μM of ParB_Bb was analyzed in a Biotek EON plate reader at 25°C 15 hours with readings taken every minute. The reaction buffer consisted of 740 μl of ultrapure water, 50 μl of a 20x customized reaction buffer (100 mM Tris pH 8.0, 2 M NaCl, and 20 mM MgCl₂), 200 μl of MESG substrate solution, and 10 μL of purine nucleoside phosphorylase (1 unit). Control reactions were performed with buffer only, buffer plus protein, or buffer plus NTP only. The plates were shaken continuously at 280 rpm for 15 hours at 25°C. Each assay was performed at least in triplicate. Data analysis was done using DataGraph, and the NTPase rates were calculated by fitting a linear regression in DataGraph.

Western blot analysis

For Western blot analysis, the AP samples were prepared following the method used in [47], starting from 1.5 ml cleared predation lysates. For the growth phase samples, a 1:1 volume ratio of E. coli prey to predator mixture was established, minimizing the excess of attack phase cells. The presence of uninfected, free E. coli cells did not interfere with downstream Western blot analysis as antibodies recognized proteins present only in B. bacteriovorus, and normalization by total protein estimates was not done between AP and GP samples (see below). Samples were collected at indicated time intervals for the time course analysis. NuPage Bis-Tris SDS precast polyacrylamide gels (Invitrogen) were used to load the samples and were run at 190 V for 50 minutes in the NuPAGE MOPS SDS running buffer. The assessment of equal total protein loading was carried out using PonceauS staining. The resulting stained membrane was imaged using an Image Quant LAS 500 camera (GE Healthcare). Standard Western blotting procedures were followed using primary antibodies against ParB_Bb (rabbit serum obtained after injecting purified ParB_Bb-6xhis protein; CER Group, Marloie, Belgium), GFP (mouse monoclonal JL-8 antibody, Takara), ParA_Cc (rabbit polyclonal antibody, Le lab) and mCherry (polyclonal antibody, Thermo Fisher, product # PA5-34974). Secondary antibodies were goat anti-mouse IgG-peroxidase antibody (Sigma) for JL-8 and goat anti-rabbit IgG-peroxidase antibody (Sigma) for mCherry, ParA_Cc and ParB_Bb. Detection of antibody binding was performed by visualizing chemiluminescence from the reaction of horseradish peroxidase with luminol, imaged with an Image Quant LAS 500 camera (GE Healthcare). The quantification of band intensities was done separately for the AP and GP samples using the ImageQuantTL software. Normalization of band intensities to obtain relative protein levels was done by the total PonceauS intensity in the corresponding lane, used as loading and transfer control. Means and standard deviations were calculated and plotted using GraphPad Prism. Image processing was done with ImageJ. The assembly and annotation of figures were performed using Adobe Illustrator. Replicates of corresponding experiments presented in Fig 3A, 3C–3D are provided in S6A–S6C Fig, respectively.

Cell, nucleoid, and spot detection from microscopy images

The automated cell detection tool Oufti [83] was used to detect outlines of AP B. bacteriovorus cells, uninfected E. coli cells, or entire bdelloplasts with subpixel precision from phase contrast images. Fluorescence signals were added to cell meshes after background subtraction. Oufti was also used to detect diffraction-limited fluorescent foci and nucleoids with subpixel precision from fluorescence images, using the spotDetection and objectDetection modules. The detected spots and objects were added to the corresponding cell in the Oufti cell lists, including features related to coordinates, morphology, and intensity. The same optimized nucleoid detection parameters were used to ensure consistency and comparisons as done previously [47]. Parameters for spot detection were optimized for each dataset or control set of images.

Quantitative image analysis from cell meshes

Fluorescence-related analysis, nucleoids, spots-related information, and other properties of individual cells based on microscopy images were extracted from Oufti cellLists data and plotted using custom codes in MATLAB (Mathworks), described below.

Fluorescence profiles

The custom Matlab script MeanIntProfile.m was used to obtain mean relative fluorescence profiles. Briefly, the fluorescence profile of each cell (corresponding to the array of fluorescence intensity per cell segment provided by the relevant signal field in the Oufti cellList) was first normalized by the corresponding array of steparea values (corresponding to the area of each segment of the cell), then divided by their sum to obtain relative fluorescence values for each cell (to account for potential concentration differences between cells). When needed, arrays of relative fluorescence were oriented based on the position of the maximal fluorescence intensity of the indicated signal in each cell half. Cell length vectors were normalized from 0 to 1, and the corresponding relative fluorescence profiles from individual cells were interpolated to a fixed-dimension vector and concatenated before averaging.

Fluorescence intensity analysis of time-lapse experiments

Bdelloplasts outlines were detected in the first frame of the time-lapse using Oufti and copied to all following frames using MATLAB. The resulting time-lapse cell list was reused in Oufti after background subtraction to add the fluorescence signal(s) to the bdelloplasts. The mean fluorescence profiles per cell over time were computed using a custom Matlab script (MeanFluoIntPerCell_TimeLapse), which extracts and plots the total fluorescence per cell (here bdelloplast) divided by the area of the corresponding cell (bdelloplast) over time.

Kymographs and demographs

Demographs of relative fluorescence intensity in cells sorted by length were plotted as in [47,84]. When needed, arrays of relative fluorescence intensity values were oriented based on the position of the maximal fluorescence intensity of the indicated signal in each cell half. Kymographs were obtained using the built-in kymograph function in Oufti [83].

Nucleoid size measurement

To measure nucleoid size, we used the objectDetection module in Oufti [83]. We considered the nucleoid area as a proxy for nucleoid size, based on previous work [85], which demonstrated that variations do not influence nucleoid area measurements in DAPI signal intensity. The nucleoid area was obtained from the nucleoid area field in the Oufti cell lists and exported to MATLAB for all cells with a single nucleoid. The nucleoid area values were then converted to μm² and used to generate violin plots of nucleoid area distributions using R [86].

Statistical analyses

The sample sizes and the number of repeats are included in the figure legends. Means, standard deviations, and coefficients of variation (CV) were calculated in GraphPad Prism, MATLAB (Mathworks), R, or Microsoft Excel.

Data, resources, and software availability

Matlab and R codes and datasets have been deposited at Zenodo (doi: 10.5281/zenodo.8325025).

Funding sources

We thank Charles de Pierpont for excellent technical support, Kilian Dekoninck and Laurie Thouvenel for advice on protein purification, Ngat Tran for advice with RT-PCR and Western blot, Francesco Caligiore for advice with ImageQuantTL analysis, Thomas G. Bernhardt for sharing strains for the POLAR assay, Michaël Deghelt for reviewing the draft, and all members of the Laloux lab for stimulating discussions and critical reading of the manuscript.