Source: https://jb.asm.org/content/193/7/1504?ijkey=1c59c4949fd269e02751d90ccecb75f42f1b9b89&keytype2=tf_ipsecsha
Timestamp: 2019-04-23 01:40:29+00:00

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Vibrio cholerae carries homologs of plasmid-borne parA and parB genes on both of its chromosomes. The par genes help to segregate many plasmids and chromosomes. Here we have studied the par genes of V. cholerae chromosome I. Earlier studies suggested that ParBI binds to the centromeric site parSI near the origin of replication (oriI), and parSI-ParBI complexes are placed at the cell poles by ParAI. Deletion of parAI and parSI caused the origin-proximal DNA to be less polar. Here we found that deletion of parBI also resulted in a less polar localization of oriI. However, unlike the deletion of parAI, the deletion of parBI increased the oriI number. Replication was normal when both parAI and parBI were deleted, suggesting that ParBI mediates its action through ParAI. Overexpression of ParAI in a parABI-deleted strain also increased the DNA content. The results are similar to those found for Bacillus subtilis, where ParA (Soj) stimulates replication and this activity is repressed by ParB (SpoOJ). As in B. subtilis, the stimulation of replication most likely involves the replication initiator DnaA. Our results indicate that control of chromosomal DNA replication is an additional function of chromosomal par genes conserved across the Gram-positive/Gram-negative divide.
Replication and segregation are the two major processes needed to maintain a chromosome stably. In bacteria, there has been considerable progress in our understanding of the process of chromosome replication. How the replicated sister chromosomes segregate to opposite cell halves is also becoming clear (62, 73). Other than those of Escherichia coli and its close homologs in the gammaproteobacteria, such as Haemophilus influenzae, most sequenced bacterial genomes have homologs of plasmid partition genes, parA and parB (29). Plasmid partitioning also requires a cis-acting sequence, parS (a centromere analog), to which ParB binds. The parS sequence is also well conserved in bacterial chromosomes (49). The par genes have been studied mostly in low-copy-number plasmids, such as P1, F, and R1, where they confer segregational stability (3, 28, 57). ParA is an ATPase and can be of three types (27). The actin-like ATPase belonging to plasmid R1 polymerizes to form filaments that link at each end with ParB-bound plasmid parS sites. Continued polymerization drives the plasmids to opposite cell poles (13). The second type of ATPase, characterized by Walker-box motifs, is more common. Members of this class of ParA also polymerize but appear to function by different mechanisms. The mechanisms involve a dynamic behavior of the ATPase, which seems to depend on the ParB/parS complex, and in some cases chromosomal DNA that somehow distributes the plasmid copies maximally away from each other, thereby increasing the probability of their presence in opposite cell halves (1, 6, 14, 22, 33, 34, 47, 65, 74). Recently, a third type of par system, which uses tubulin-like GTPases (dubbed TubZ) in plasmid segregation, has been discovered (2, 5, 40, 55).
The chromosomally encoded Par proteins appear to be both homologs and analogs of their plasmid counterparts (29, 30, 48). The functional relatedness of the two became evident when the chromosomal par system of Bacillus subtilis was shown to stabilize an unstable Bacillus plasmid (48). More strikingly, the Bacillus system stabilized an unstable E. coli plasmid in E. coli (77). The latter result implied that no host-specific factors are required for plasmid partition, which is still believed to be the case (26).
Chromosomal Par proteins have been studied in many bacteria, including B. subtilis (35, 48), Burkholderia cenocepacia (19), Caulobacter crescentus (12, 21, 53), Helicobacter pylori (42), Mycobacterium smegmatis (36, 37), Pseudomonas aeruginosa (8, 41), Pseudomonas putida (30, 46), Streptomyces coelicolor (37), and Vibrio cholerae (25, 64). These studies revealed several features of the par genes. (1) They map close to the origin of replication but are not essential except in C. crescentus. (2) Their absence or overproduction causes at least some segregation defect, indicated by increased formation of anucleate cells, abnormal nucleoid compactness, and reduced separation of origin-proximal markers. (3) They contribute to other cellular processes, which include chromosome replication, cell division, and sporulation in B. subtilis (35, 54), cell division in C. crescentus and S. coelicolor (12, 21, 37, 52), and cell motility in P. aeruginosa (41).
Studies in C. crescentus and B. subtilis have contributed the most to our current understanding of chromosomal par systems. In C. crescentus, replication initiates at the stalked pole, and one of the daughter origins moves rapidly to the opposite pole. All three components of the parABS system contribute to chromosome segregation (60, 61, 66). In B. subtilis, the contribution of par to chromosome segregation is not crucial. Deletion of soj (parA) causes no significant partition defect, and deletion of spo0J (parB) reduces the separation of sister origins from 48 to 40% of cell length only (from cell quarter positions toward the cell center) and causes about 0.3% of cells to be anucleate (35, 43, 44). The nucleoid, however, can be abnormal in 20% of cells grown in rich medium (4). Also, when the B. subtilis parS sites are moved away from their natural origin-proximal positions, they no longer localize to cell quarter positions, implying that the par system might not be sufficient for their localization to cell quarter positions (43). In P. aeruginosa and in P. putida, anucleate cells account for up to 10% of the population in the absence of Par proteins (8, 30, 41, 46). The role of par can be more important under certain growth conditions. In P. putida, anucleate cells were seen at higher frequencies when the chromosome copy number was reduced, such as in minimal media or when cells approach stationary phase (30, 46).
Among the roles of par in cellular processes other than chromosome segregation, there is a considerable understanding of how soj and spoOJ contribute to sporulation and how overproduction of soj leads to overinitiation of replication, a process that is normally regulated by spoOJ (44, 54, 58). The replication stimulation is mediated by Soj complexing with the replication initiator protein DnaA (54). SpoOJ, in addition to controlling replication via Soj, helps to promote chromosome segregation by recruiting the SMC condensin near the origin (31).
Both the chromosomes (chromosome I [chrI] and chrII) of V. cholerae have parAB genes and the cognate parS sites (76), which contribute to segregation in a chromosome-specific manner. The par genes of chrI direct the origin of chrI (oriI) to a cell pole, similar to the situation in C. crescentus, and the par genes of chrII direct the origin of chrII (oriII) to the cell center, similar to the situation in E. coli (23, 24, 64, 69). Separate deletions of parAI and parSI of chrI cause oriI to be less polar (25, 43, 64, 76). A previous attempt to delete parBI or parABI has not been successful (64).
Here we have shown that parBI can be deleted and replaced with a drug cassette. The deletion caused little growth defect and some chrI segregation defect, comparable to the defect observed in parAI and parSI deletion strains. Deletion of parBI, however, showed an additional phenotype: parAI-dependent overinitiation of chrI replication. The results are strikingly similar to those found in B. subtilis and provide another example of chromosomal par genes controlling DNA replication.
Strains, plasmids, and growth conditions.Strains and plasmids used in this study are listed in Table 1. E. coli DH5Δlac (BR2846) was used for standard plasmid manipulations and for plasmid propagation. V. cholerae strains were all derivatives of N16961 (CVC209). Bacterial cells were grown in L broth. Antibiotics were used at the following concentrations: ampicillin, 100 μg/ml; chloramphenicol, 25 μg/ml; kanamycin, 25 μg/ml; spectinomycin, 40 μg/ml; and zeocin, 25 μg/ml.
Deletion of parBI.The deletion was achieved by allele exchange, where the gene was replaced with a zeocin (Zeo) drug cassette from the plasmid pEM7-Zeo (68). Briefly, the PCR primers RG135 and RG136, containing XhoI and XbaI-plus-NheI sites, respectively, were used to amplify the C-terminal region of the parAI open reading frame (ORF) (sequence coordinates 2957716 to 2958175). The PCR product was ligated to NheI- and XhoI-digested pEM7-Zeo to obtain pRKG240. A sequence downstream of parBI (coordinates 2956199 to 2956820) was amplified with the primers RG133 and RG142, containing SalI and EcoRI sites, respectively. The product was digested with SalI and EcoRI and ligated to pRKG240, predigested with the same enzymes. The resultant plasmid, pRKG241, was digested with XbaI and SphI, and the fragment carrying the Zeo cassette with parBI flanking sequences was cloned into pDS132 between the XbaI-SphI sites. The resultant plasmid, pRKG242, was transferred by conjugation to V. cholerae CVC209, using E. coli S17λpir (CVC764) as the donor strain. Recombinants in which parBI was replaced with the Zeo cassette without the pDS132 backbone were obtained by serial subculture of the transconjugant followed by sucrose selection. The deletion was confirmed by PCR using the primers RG122 and RG113, internal to the parBI gene, by DNA sequencing, and by Western blotting using antibody against ParBI (see Fig. S1 in the supplemental material). One such construct, CVC784, was used for all subsequent experiments.
Expression of parAI, parBI, and parABI.To express parAI, the gene was amplified by PCR using the primers RG115 and RG214 with Acc65I and XbaI sites at their 5′ ends, respectively, using N16961 DNA as a template. The fragment was digested with Acc65I and XbaI and cloned into similarly digested pBAD24 to make pRN004. An identical procedure was used to express parBI, except the primers were RG5 (with an Acc65I site) and RG6 (with a HindIII site) and the vector was predigested with Acc65I and HindIII. This resulted in pRKG212. The procedure to express parABI was essentially identical to that used for parAI except that the primers were RG115 and RG113, respectively. This resulted in pRN005. ParAI was also produced from pRN006, which was derived from pRN005 by deleting the parBI gene between the sites BxtXI and HindIII. The former site was present in the beginning of parBI and the latter in the vector backbone. ParAI mutants, K16A and D40A, were produced from pJHB41 and pJHB42, respectively, which were derived from pRN006 by site-directed mutagenesis.
Chromosomal integration of P1parS.P1parS as a parS-Kn cassette was inserted at approximately +135 kb in the intergenic region between VC0142 and VC0143. First, the PCR primer pair PS11 and PS14, both containing a PstI site at their 5′ ends, were used to amplify sequences between 133582 and 135584, containing VC0142 and VC0143. The product was cloned into pDS132, predigested with PstI, to generate pPS4. To clone the P1parS-Kn cassette, pPS4 was digested with XmnI and ligated to the parS-Kn cassette obtained from pPS45 by digestion with BamHI and blunting with the Klenow enzyme followed by digestion with SspI. The resultant plasmid, pPS47, was transferred to V. cholerae cells by conjugation, and the transconjugants (CVC851 for wild-type [WT], CVC899 for ΔparAI, CVC853 for the ΔparBI, and CVC1126 for ΔparABI cells) with parS-Kn inserted in chrI were used to localize oriI. To localize oriII, the P1parS-Kn cassette was integrated in the intergenic region between VCA032 and VCA033, both coding hypothetical proteins, as described previously (70). The resultant strains were CVC827 for the WT, CVC1130 for the ΔparAI strain, CVC787 for the ΔparBI strain, and CVC1128 for the ΔparABI strain.
Natural transformation of Vibrio cholerae.A hapR+ derivative of N16961 (CVC1118) was grown to log phase (optical density at 600 nm [OD600] between 0.1 and 0.2) in L broth, and the cells were resuspended in defined artificial seawater (DASW) medium (11). The cells were added to chitin beads (New England BioLabs). After culturing overnight at 30°C to allow biofilm formation, the planktonic bacteria together with medium were removed and replaced with fresh DASW medium. Immediately after the medium exchange, 2 μg PCR DNA fragment from the donor strain was added, and the incubation continued at 30°C overnight. The cells were detached from the beads by vortexing and plated to select for antibiotic-resistant transformants.
Fluorescence microscopy.Cells were cultured in L broth at 37°C to log phase (OD600 between 0.1 and 0.2) and concentrated 100-fold by centrifugation and resuspension in the same medium. They were stained for DNA with Hoechst-33342 at a final concentration of 0.5 μg/ml for 30 s at room temperature and observed under a fluorescence microscope (E1000; Nikon) fitted with a Hamamatsu EM-CCD digital camera (Photonics KK) and controlled by the MetaMorph Premier software program (Molecular Devices, Sunnyvale, CA). Focus locations were measured using the Image J software program (http://rsb.info.nih.gov/ij/index.html).
Flow cytometry.Chloramphenicol (25 μg/ml) and cephalexin (10 μg/ml) were added to exponentially growing cells (OD600 ≈ 0.1) to inhibit new rounds of replication and cell division, respectively. Incubation was continued for an additional 5 h to ensure completion of all rounds initiated prior to addition of the drug. Cells were further treated for flow cytometry, and the number of replication origins in individual cells was determined with a flow cytometer, as described previously (71).
Bacterial two-hybrid assay.A bacterial adenylate cyclase two-hybrid (BACTH) system (EUK001; Euromedex, France) was used according to the manufacturer's protocol. The parAI, parBI, and dnaA genes of V. cholerae N16961 were cloned into pKT25 and pUT18C in frame with the T25 and T18 fragments to make bait and prey plasmids, respectively (Table 1). The PCR primers BJH1 (containing a PstI site at the 5′ end) and BJH2 (containing a BamHI site at the 5′ end), BJH3 (containing a PstI site at the 5′ end) and BJH4 (containing a BamHI site at the 5′ end), and BJH12 (containing a SalI site at the 5′ end) and BJH13 (containing a BamHI site at the 5′ end) (see Table S1 in the supplemental material) were used to amplify the parAI, parBI, and dnaA genes, respectively. The PCR products were digested with PstI and BamHI for parA1 and parB1 or with SalI and BamHI for dnaA and cloned into pKT25 and pUT18C. The pKT25 derivatives carrying parA1, parB1, and dnaA were pBJH3, pBJH4, and pBJH10, respectively, and the corresponding pUT18C derivatives were pBJH5, pBJH6, and pBJH, respectively. Pairs of bait and prey plasmids were used to transform E. coli BTH101 (CVC1837) cells, and the transformants were grown overnight at 30°C in L broth with appropriate antibiotics in the presence of 0.5 mM isopropyl-β-d-thiogalactopyranoside (IPTG). β-Galactosidase activity was measured to determine the functional complementation between the fusion proteins (51).
parBI is dispensable in V. cholerae.Although parAI and parSI have been deleted without any significant growth defect, deletion of parBI has not been successful (64). In the present study, the gene was replaced with a Zeo cassette by an allele exchange method (Materials and Methods). Sequences between coordinates 2956820 and 2957716 covering the entire parBI ORF (2956823 to 2957704) were deleted. The deletion was obtained with similar frequency whether or not a parBI-complementing plasmid (pRKG212) was present in the recipient cells during conjugation and subsequently in transconjugant cells. The parBI gene was also deleted by natural transformation by introducing a PCR fragment (∼2.5 kb) carrying the Zeo cassette and the parBI flanking regions into CVC1118 (N16961 hapR+) and selecting for zeocin resistance. The hapR+ allele was required to make the strain competent for natural transformation (50). The template DNA for PCR was either the plasmid pRKG241 or the genomic DNA of the parBI-deleted strain, CVC784. Similarly, both the parAI and parBI genes were deleted (coordinates 2956820 to 2958505) and replaced with the Zeo cassette by natural transformation using pRKG252 as the template for generating the PCR fragment (∼2.5 kb). The resulting strain was called CVC1124. These results show that ParBI is not essential for V. cholerae growth in L broth.
In an earlier study, codons 11 to 244 of the 258-codon parAI ORF were deleted in frame, and the growth of the mutant was found to be indistinguishable from that of the WT in L broth (64). The doubling time, morphology, and frequency of anucleate cell formation of ΔparBI and ΔparABI cells in L broth were essentially identical to those of the WT and ΔparAI cells, described above. The generation times were approximately 19.5 min, and anucleate cells, analyzed after Hoechst staining, accounted for <1% of the population (varying from 0.1 to 0.8% in the four cases). Nucleoid condensation was also observed at similar low frequencies (0.6 to 0.8%) in all four strains. The parABI genes thus do not contribute significantly to chromosome stability.
oriI segregation is compromised in ΔparBI cells.oriI of V. cholerae preferentially locates to cell poles, but in ΔparAI and ΔparSI cells, the origin is found to be less polar (25, 64). ParBI binds to parSI sites, which are located about +65 kb from the origin of chrI, and ParAI is believed to interact with the ParBI-parSI complex, directing parSI and the nearby DNA, including oriI, to the pole. In this scenario, oriI is expected to be less polar in ΔparBI cells. To test this, WT, ΔparAI, ΔparBI, and ΔparABI cells were marked identically with the plasmid P1parS near oriI, at about the +135 kb position of chrI. These cells were then transformed with pALA2705 to supply GFP-P1ParB under the control of the inducer, IPTG. In this system, GFP-P1ParB binds to P1parS and spreads to nearby DNA to give a bright fluorescent focus. The system has proven to be a reliable indicator of origin number and position in earlier studies (56, 69).
V. cholerae cells grown in L broth at 37°C showed two oriI foci in newborn cells and up to four foci in longer cells (Fig. 1A and B). In WT cells, the focus closest to a pole was on average about 10.8% of the cell length away from the pole; the other origin was at various intermediate positions all the way to the opposite pole (Fig. 1B and 2A). These results are consistent with earlier results obtained with similar and other origin labeling systems (23, 24, 64, 69). In ΔparAI cells, the origin foci were less polar, the mean position of the focus closest to the pole being 17.3% removed from the pole (Fig. 1B and 2A). These results are consistent with earlier findings that suggested a ParAI requirement at the final stages of oriI localization to the pole (25, 64). In parBI and ΔparABI cells, the oriI foci were located 18.9% and 17.3% away from the pole, respectively, similar to the situation in ΔparAI cells (Fig. 1B and 2A). Nucleoid sizes (along the long axes of the cells) were nearly the same (between 98.8 and 99.4% of cell length) in the WT and the mutants. These results support the notion that ParAI and ParBI participate in the pathway that leads oriI to the pole, and nonpolar localization of oriI in the mutants is unlikely to be due to gross changes in nucleoid structure in the mutants.
Location of oriI in WT cells or in parAI-, parBI-, and parABI-deleted cells growing exponentially in L broth. (A) Localization was done by using the GFP-P1ParB/P1parS system by inserting P1parS about 135 kb away (clockwise) from the origin. Cells normally showed up to four foci except in the case of ΔparBI cells, where the number was higher (arrow). These cells accounted for about 7% of the population and were not included in the plot. (B) Plots show the positions of oriI foci in different cells. The cells were arranged according to increasing lengths along the abscissa and focus positions along the ordinate as fractional cell lengths. Focus positions were measured from the pole from which the nearest focus was closer than the nearest focus of the opposite pole. The pole from which measurements were made was placed on the abscissa. Small cells usually showed two foci (light-blue symbols) and larger cells from three to five foci (purple, red, and green symbols, respectively). Black dashed lines show the mean position of the foci nearest to the abscissa from all cells, and it is at 0.108 for the WT, 0.175 for ΔparAI cells, 0.189 for ΔparBI cells, and 0.173 for ΔparABI cells.
(A) Distribution of polar distances (x) of the focus nearest to a pole in the WT and in par mutants. The nearest focus to a pole is shown schematically (black circles) in a two-focus cell and in a four-focus cell. The distribution of the polar distances from different cells is shown after binning of the distances into equal 0.05 fractional cell length intervals and using the mean lengths of the binning distances as the x values (0.025, 0.075, and so on). For ease of presentation, the data were fit by the Kaleidagraph 4.03 software program (Synergy Software, Reading, PA) using unweighted least squares and the fitting function y(x) = c1 xm [exp(−c2x[r]) + c3 exp(−c4 x)], where x denotes the focus position as defined above, y denotes the % of cells, and Ci and m are fitting parameters. r values were 0.998, 0.973, 0.976, and 0.951 for the WT, ΔparAI, ΔparBI, and ΔparABI cases, respectively, and the corresponding locations of the peak maximums were 0.052, 0.118, 0.126, and 0.124. (B) Distribution of cell numbers with different numbers of origin foci. To simplify the distribution for oriI, cells with either one or two foci or three or four foci were grouped before plotting. For oriII, cells had either one or two foci, precluding any need to simplify the plotting further.
Because the segregation defect was modest in the absence of parABI in L broth, we wanted to test whether the defect gets more pronounced during slower growth, as has been reported in P. putida (30). To slow down the growth rate, we used a less rich medium (M63 salts plus 0.2% fructose plus 0.1% Casamino Acids), which increased the generation time of the WT from about 19.5 to 28.5 min. The increases were similar for the three mutants. In all mutants, the oriI foci were removed from the pole compared to their positions in L broth (Fig. 3). In L broth, compared to its position in the WT, oriI in the mutants was on average about 7% more removed from the pole. In the new medium, the corresponding value was 18%. The basis for this growth rate-dependent requirement of par function remains to be understood.
Location of oriI in WT and in parAI-, parBI-, and parABI-deleted cells grown in a defined medium (M63 + 0.2% fructose + 0.1% Casamino Acids). (A) In this medium, cells usually showed from one to two foci (dark- and light-blue symbols, respectively) except in the case of parBI-deleted cells, which showed 12 three-focus cells and 1 four-focus cell (red symbols: about 11% of total cells). Black dashed lines show the mean position of the foci nearest to the abscissa from all cells, and it is at 0.059 for WT, 0.252 for ΔparAI, 0.210 for ΔparBI, and 0.267 for ΔparABI cells. Other details are the same as in Fig. 1B. (B) Distribution of cell numbers with different numbers of origin foci.
oriII locates differently from oriI.In L broth at 37°C, usually one focus is seen near the cell center in smaller cells and up to two foci approaching cell quarter positions in larger cells. These replication and segregation patterns appeared unchanged in the mutants (see Fig. S2 in the supplemental material). We note that because oriII foci do not normally separate as much as the oriI foci, a small perturbation of the segregation pattern in the mutants, to the extent seen for oriI, might not have been apparent in the background of inherent noise in oriII positions. Others have tested oriII positioning and found it to remain unaltered in ΔparAI cells (25, 64).
Chromosome I overinitiates in ΔparBI cells.In the above experiments, although the mean location of origins did not differ significantly between ΔparAI and ΔparBI cells, the distribution of cells with different numbers of oriI foci did vary in the two mutants. In the ΔparBI mutant, there was an increase in the number of cells with three or four oriI foci and a corresponding decrease in the number of cells with one or two oriI foci (Fig. 2B). If it is assumed that origin separation follows soon after replication, this indicates that replication of chrI is initiated earlier in the cell cycle in the ΔparBI mutant than in the ΔparAI mutant. The distributions of the WT and the ΔparABI strain were similar to that of the ΔparAI strain. The increase in the number of cells with more replication origins was not simply due to cell elongation but more likely was due to earlier initiation because the number of origins per cell length increased in ΔparBI cells (Table 2). The distribution of cells with different numbers of oriII foci was similar in all four strains (Fig. 2B).
An independent measure of the origin number was obtained by flow cytometry after replication runout. Replication initiation and cell division were blocked by chloramphenicol and cephalexin, and sufficient time (5 h) was allowed to complete replication elongation (replication runout) in cells that had already initiated replication at the time of drug addition. In the case of the ΔparBI mutant, cells with four genome equivalents of DNA dominated over cells with two genome equivalents, whereas the opposite was true in the other three strains (Fig. 4A). These results suggest that at the time of drug addition, more of the ΔparBI cells had already initiated replication, supporting the microscopy results that suggested earlier replication initiation in ΔparBI cells.
Distribution of DNA contents of cells by flow cytometry after replication runout. (A) DNA contents in WT, ΔparAI, ΔparBI, and ΔparABI cells. (B and C) DNA contents under protein overproduction conditions. In panel B, WT cells either had the vector plasmid pBAD or the parAI, parBI, or parABI genes under the control of the PBAD promoter of the vector. In panel C, the plasmids were the same but the host was ΔparABI cells. One hundred thousand cells were analyzed in all cases.
Chromosome I overinitiation requires ParAI.As shown in Fig. 4A, the overinitiation phenotype of ΔparBI cells was not seen in the ΔparABI double mutant, indicating that ParAI could be required for the phenotype. To test this, we overproduced ParAI, ParBI, and the two together in WT and in ΔparABI cells. Overproduction was achieved by cloning the genes in a pBAD vector, under the control of an arabinose-inducible promoter, PBAD, and an increase in the level of proteins of interest was confirmed by Western blotting (see Fig. S1B and C in the supplemental material). In the WT background, the profiles of fluorescence intensity were comparable between the cells with pBAD and cells with pBADparBI (Fig. 4B). In contrast, when the cells had the pBADparAI or pBADparABI overproducing plasmid, there were more cells showing higher fluorescence, i.e., there was a decrease of cells with two genome equivalents and an increase of cells with four genome equivalents. In the ΔparABI background, the results were essentially the same, with the profiles of cells with pBAD and pBADparBI being comparable to each other and the profiles of cells with pBADparAI and pBADparABI being comparable to each other. A common feature of these results is that overinitiation could be seen under all ParAI overproduction conditions, whether or not ParBI was also overproduced. The role of ParBI in this process is most likely that of an inhibitor of ParAI activity rather than a transcriptional repressor of the parAI gene, since overinitiation was achieved in ΔparBI cells without significant overproduction of ParAI (see Fig. S1A). The overinitiation activity of ParAI apparently dominated over the inhibitory activity of ParBI when both proteins were overproduced. This could be because in these experiments ParAI was overproduced about 8-fold more than ParBI (∼160- versus ∼20-fold, respectively; see Fig. S1). When these findings are taken together with the microscopy results, it is clear that ParAI is required to increase initiation from chrI. This can happen at near-physiological levels of ParAI, but the level needs to be significantly increased when ParBI is also present. When ParA1 was overproduced about 30-fold (using 0.02% arabinose), the replication enhancement was not apparent (data not shown). We note that even in the absence of ParBI, a considerable excess of ParAI over the physiological level was needed for overinitiation when ParAI was supplied from a plasmid rather from its natural chromosomal locus. The basis of this cis-trans difference in the efficacy of ParAI remains to be understood.
From studies in B. subtilis, it was expected that the nucleotide-bound state of ParAI could be important for its role in replication initiation (20, 54). As a preliminary test of this notion, parAI was mutated at conserved positions that resulted in K16A and D40A changes in the protein. The K16A change is expected to be defective in ATP binding and the D40A change in ATP hydrolysis (54). As in B. subtilis, only the latter protein was proficient in replication promotion (Fig. 5). In summary, both the amount and form of ParAI appear important for the proper control of replication initiation of V. cholerae chrI.
Overreplication phenotypes of ParAI mutants with K16A and D40A changes in Walker-box motifs. (A) Replication was determined by flow cytometry as with Fig. 4. (B) The protein level after overproduction was determined by Western blotting, as with Fig. S1 in the supplemental material, and the levels in the mutants were comparable to that of WT ParAI.
ParAI overproduction causes nucleoid condensation.Cells overproducing ParAI, ParBI, or both were also examined by microscopy. In WT cells without the overproducing plasmids, the nulceoid boundaries spanned 91% ± 4% of the cell length (Fig. 6). The results were essentially similar in the ΔparAI cells, where the value was 94% ± 4%. In contrast, under ParAI-overproducing conditions, the nucleoid appeared more compact, since the value was 62% ± 16%. When ParBI or both ParAI and ParBI were overproduced, the compaction was again not significantly different from that of the WT (96% ± 4%, 90% ± 11%, and 91% ± 4%, respectively). It appears that ParBI can effectively suppress the nucleoid compaction phenotype of ParAI.
ParAI overproduction causes nucleoid condensation and positions oriI at the pole-ward edges of the nucleoid. The nucleoids were visualized by Hoechst staining, and oriI was visualized by marking with GFP, as in Fig. 1. Overlays of the two are shown in the last column. The positions of nucleoid boundaries and oriI were measured independently and are shown together in the same plots to visualize the correspondence between the two positions on a population basis, not on a cell-to-cell basis. The color coding is the same as in Fig. 1, except the nucleoid boundaries are shown in dark blue. The results are summarized in a schematic where the nucleoid is shown as a blue oval and oriI positions as red circles.
We also checked oriI localization in each of the overproducing strains. The position of the origin was less polar in ParAI-overproducing cells, comparable to the situation in ΔparAI cells (Fig. 6). In the former case, it is likely that the origins failed to reach the pole or stay at the pole because of nucleoid compaction. If there exists an anchoring mechanism to hold the origins to the pole, as has been found in C. crescentus and sporulating B. subtilis cells, it might not have been strong enough to withstand the force of compaction or was made ineffective by an excess of ParAI (9, 12, 21). It is intriguing that both the absence of a par system (ΔparABI and ΔparSI) and overproduction of ParAI lead to the same oriI localization defect, although the reasons in the two cases need not be the same.
In B. subtilis, increased expression of Soj makes the nucleoid less compact, the opposite of the results presented here (58). Compaction was seen when both Soj and SpoOJ were overproduced together. Under these conditions, the SpoOJ-to-Soj ratio was higher than their ratio in the WT, which was interpreted to mean that the excess of SpoOJ could be responsible for the compaction. SpoOJ has been found to recruit the SMC protein, a condensin (31, 72). In C. crescentus, constitutive overexpression of either ParA or ParB causes cell division defects that are less severe when both proteins are overexpressed together (53). These results suggest that proper stoichiometry of the two Par proteins is important for normal cell physiology. The importance of concentration is also suggested by the observation that Soj can be either a positive or negative regulator of replication, depending upon its level of production (54).
Interactions of DnaA with ParBI and ParAI.The DnaA protein is considered to be the limiting factor for chromosomal replication initiation (63). Therefore, it is expected that ParAI-mediated stimulation of replication could be mediated through DnaA. Such has been the observation for B. subtilis (54). To determine whether this could be the case in V. cholerae, we carried out protein-protein interaction studies using ParAI, ParBI, and DnaA. The interactions were assayed by the bacterial two-hybrid system. The system is based on the reconstituted activity of Bordetella pertussis adenylate cyclase (cya) from its two complementary fragments, T25 and T18 (38). The full-length ParAI, ParBI, and DnaA proteins from V. cholerae were fused at the C terminus of either the T25 or T18 fragment. As a positive control, a 35-amino-acid (aa)-long leucine zipper belonging to the yeast transcriptional activator GCN4 was fused to both T18 and T25. Considering the activity obtained with two empty vectors (about 20 Miller units) as the background level, 10/10 randomly picked colonies containing the leucine zippers as interacting partners showed at least 2-orders-of-magnitude-higher activity. In the experimental samples, 2 to 5 colonies were considered significantly positive, whereas in strains containing one empty vector plus one with a fusion protein, none of the colonies showed activity significantly above the background level (Table 3). The assay thus indicates interactions between ParAI and ParAI, ParAI and ParBI, and ParBI and ParBI, although the interactions were much weaker than those between the leucine zippers. In the case of DnaA, an indication of interactions with both ParAI and ParBI could be found when DnaA was fused to T18 (T18-DnaA). However, with T25-DnaA, the results were positive with T18-ParAI only. T25-DnaA fusion, however, complemented to some extent the growth defect of a dnaA46(Ts) strain at the nonpermissive temperature (see Fig. S3 in the supplemental material). It is possible that the conformation of DnaA upon fusion to T25 could have changed, so that it was no longer active for interaction with itself and ParBI. Alternatively, the concentration of T25-DnaA could be lower than optimal because of the plasmid producing the fusion protein having a lower copy number than the one producing T18-DnaA.
We also tried to validate these interactions by the yeast two-hybrid system. Here, the evidence for positive interactions was obtained among the three proteins in all possible combinations (see Table S2 in the supplemental material). However, the results were again significantly above the background level in only some of the colonies. At least in the bacterial system used, the all-or-none response is expected if the proteins interact weakly. The proteins are expressed from the lac promoter, which itself depends upon cAMP. If the basal level of expression of the bait and prey proteins is low and the proteins interact weakly, this might not lead to enough cAMP production, and a negative result follows. When the basal level of cAMP crosses a threshold, more bait and prey proteins are made, which in turn make more cAMP, leading to an autocatalytic reaction and a robust positive response. Another inherent weakness of the system is that it is based on multicopy vectors, and plasmid copy number fluctuation could also be a source of heterogeneity. In our case, in the extract from the cya mutant host, the fusion proteins were barely detected by Western blot analysis, although all the fusion constructs were transcribed (see Fig. S4A). The ParAI and ParBI fusion proteins were detected using a cya+ strain, which was also deleted for the ClpXP and Lon proteases (see Fig. S4B). Unfortunately, DnaA fusion proteins could not be detected using antibodies against E. coli DnaA. Thus, both low expression and instability of the fusion proteins could be responsible for the all-or-none response in the results of the bacterial two-hybrid system.
In summary, the results from bacterial and yeast two-hybrid systems indicate that V. cholerae ParAI, ParBI, and DnaA can interact with each other. Interactions between the ParA and ParB proteins have been found in other bacteria (7, 8, 10). DnaA interaction with ParA (Soj) has been found in B. subtilis (54). To the best of our knowledge, it has not been reported that a ParB protein (ParBI) can also interact with DnaA.
Concluding remarks.V. cholerae provides a second example of participation of chromosomal Par proteins in chromosomal replication, which was first observed in B. subtilis (44, 54, 58). Conservation of Par functions in different bacterial phyla was evident from sequence conservation, stabilization of a Bacillus nato plasmid by the B. subtilis par system, and even stabilization of an E. coli plasmid by the B. subtilis par system in E. coli (29, 48, 77). The Par proteins, however, are not required for chromosomal replication initiation in either B. subtilis or V. cholerae and normally do not appear to contribute to replication significantly under the conditions tested. In E. coli, the regulators of oriC include DnaA, SeqA, Dam, pgsA, DnaK, relA, Hda, IHF, HU, FIS, IciA, datA, DiaA, nrdAB, fre, and DARS1 and -2 (39). Most of these regulators can be deleted at least individually with minor effects. In a system with multiple regulators, the effect of any one may not be expected to be great. Even then, it is remarkable that the apparent ancillary role of ParA in stimulatnig replication has been retained in two bacteria that are believed to have diverged more than a billion years ago.
Replication and segregation modules in plasmids are usually found next to each other. However, it is not known even for well-characterized systems whether the two modules can function mechanistically independently of each other in their natural context (27). In only a few cases, the modules are not discrete and some links between the two systems have been found at the level of transcription. For example, in the broad-host-range plasmid RK2, the replication initiator gene trfA is controlled by the ParB homolog KorB, which also serves as a general transcription factor in that plasmid (67). In the repABC plasmid family, the partition and replication genes are present in the same operon (15). In a Streptococcus pyogenes plasmid pSM19035, the replication and partition genes are under a common regulator, ω2 (16). In the linear phage-plasmid N15, one of the two promoters of the partition operon is repressed by protelomerase, the enzyme that completes prophage replication (18). Beyond transcription, however, there is no evidence yet as to whether the processes of replication and partition influence each other. The situation seems to be different for chromosomes. The act of replication has been invoked to provide the driving force for segregation (45). A negative regulator of replication, SeqA, has been proposed to help in proper segregation of daughter chromosomes (75). Here we show the opposite in that ParB contributes to chromosome segregation but can be a negative regulator of replication as well (Fig. 1 and 4). In B. subtilis, SpoOJ helps to regulate chromosome replication via Soj and chromosome segregation by SMC condensin recruitment near the origin of replication (31, 72). V. cholerae encodes a MukBEF complex, analogous to SMC. It remains to be seen whether ParBI helps recruit the complex near oriI. Together these results indicate that interactions between replication and segregation systems might be more common than is currently known. Finally, we note that V. cholerae ParAI could not stimulate replication in E. coli or in an engineered E. coli strain where oriC has been replaced with V. cholerae oriI (R. Kadoya, unpublished results) (17). Thus, even in closely related strains, such as E. coli and V. cholerae, there seems to have been considerable adaptation between the replication and segregation proteins.
We are grateful to Melanie Blokesch, Ranajit Ghosh, Lyn Sue Kahng, Jon Kaguni, Mike Maurizi, Ole Skovgaard, Preeti Srivastavaand, and Matt Waldor for strains, reagents, and advice. Special thanks go to Aurelia Battesti and Tapan Som for help with two-hybrid systems and to Jean-Philippe Castaing, Grazyna Jagura-Burdzy, David Lane, Michael Yarmolinsky, and the anonymous reviewers for critical reading of the manuscript.
This work was supported by the Intramural Research Program, Center for Cancer Research, National Cancer Institute.
Accepted manuscript posted online 21 January 2011.
† Supplemental material for this article may be found at http://dx.doi.org/10.1128/JB.01067-10.
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