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. 2011 Jul 15;25(14):1544–1555. doi: 10.1101/gad.2061811

A family of ParA-like ATPases promotes cell pole maturation by facilitating polar localization of chemotaxis proteins

Simon Ringgaard 1,2,3, Kathrin Schirner 2, Brigid M Davis 1,2,3, Matthew K Waldor 1,2,3,4
PMCID: PMC3143943  PMID: 21764856

Abstract

Stochastic processes are thought to mediate localization of membrane-associated chemotaxis signaling clusters in peritrichous bacteria. Here, we identified a new family of ParA-like ATPases (designated ParC [for partitioning chemotaxis]) encoded within chemotaxis operons of many polar-flagellated γ-proteobacteria that actively promote polar localization of chemotaxis proteins. In Vibrio cholerae, a single ParC focus is found at the flagellated old pole in newborn cells, and later bipolar ParC foci develop as the cell matures. The cell cycle-dependent redistribution of ParC occurs by its release from the old pole and subsequent relocalization at the new pole, consistent with a “diffusion and capture” model for ParC dynamics. Chemotaxis proteins encoded in the same cluster as ParC have a similar unipolar-to-bipolar transition; however, they reach the new pole after the arrival of ParC. Cells lacking ParC exhibit aberrantly localized foci of chemotaxis proteins, reduced chemotaxis, and altered motility, which likely accounts for their enhanced colonization of the proximal small intestine in an animal model of cholera. Collectively, our findings indicate that ParC promotes the efficiency of chemotactic signaling processes. In particular, ParC-facilitated development of a functional chemotaxis apparatus at the new pole readies this site for its development into a functional old pole after cell division.

Keywords: chemotaxis, cell pole maturation, protein localization, bacterial development, ParA, ATPase

Bacteria have complex internal structures, and many important cellular processes are regulated spatially as well as temporally. Consequently, many proteins exhibit distinct patterns of subcellular localization, some of which arise due to active mechanisms (Shapiro et al. 2009). For example, in rod-shaped bacteria, numerous proteins are localized to a cell pole, where they form complexes critical for processes such as chromosome segregation, motility, and chemotaxis (for review, see Gerdes et al. 2010; Sourjik and Armitage 2010; Kirkpatrick and Viollier 2011). At least in some species, the sets of protein complexes found at the two cell poles (the old and new pole, respectively) differ, generating asymmetry between the two ends of the cell (Shapiro et al. 2009). Other proteins, such as those mediating cell division, instead accumulate at bacterial division sites, typically, but not always, restricted to the mid-cell. Notably, as cells proceed through the cell cycle, their subcellular domains adapt as well, so that division sites give rise to new poles, and new poles are converted by division events into old ones. However, the mechanisms by which bacterial subcellular domains initially form and subsequently mature, and the means by which distinct maturation processes are coordinated within the cell, remain incompletely understood.

One comparatively well-studied process that in many bacteria depends on precise positioning of cellular components throughout the cell cycle is that of chromosome segregation. In several rod-shaped bacteria, including Vibrio cholerae and Caulobacter crescentus, cell pole-anchored partitioning (Par) systems facilitate chromosome segregation (Fogel and Waldor 2006; Ptacin et al. 2010). Par systems are comprised of a Walker-type ATPase, ParA; a centromere-binding protein, ParB; and at least one cis-acting centromere site, parS (Gerdes et al. 2010). Interactions between these components are thought to direct movement of chromosomes and plasmids that contain parABS loci, thereby enabling their accurate segregation to daughter cells (Barilla et al. 2005, 2007; Fogel and Waldor 2006; Pratto et al. 2008; Ringgaard et al. 2009; Ptacin et al. 2010). Recently, ParA proteins have also been implicated in the positioning and segregation of other cellular components; for example, the cytosolic chemoreceptors (Tlps) in Rhodobacter sphaeroides (Thompson et al. 2006) and carboxysomes in cyanobacteria (Savage et al. 2010).

Membrane-associated chemotaxis proteins also localize to discrete cellular domains (Maddock and Shapiro 1993; Sourjik and Berg 2000; Thiem et al. 2007; Briegel et al. 2008). These chemotaxis apparati are multiprotein complexes that recognize an extracellular stimulus, transmit it through the bacterial (inner) membrane, and ultimately modulate the direction or extent of flagellar rotation, which governs motility and swimming direction. The sensors of external stimuli are a class of inner membrane proteins, the methyl-accepting chemotaxis proteins (MCPs). They transmit the signal to the cytosolic protein kinase CheA, which is recruited to the MCPs by the adapter protein CheW. When encountering a decrease in attractant or increase in repellant, the MCPs stimulate CheA autophosphorylation. CheA subsequently phosphorylates the response regulator CheY. Phosphorylated CheY can diffuse through the cytoplasm and bind the flagellar switch protein FliM, thereby increasing the probability of a change in motor rotation from the default counterclockwise (CCW, straight swimming) to a clockwise (CW) orientation, resulting in a change of swimming direction (for review, see Sourjik and Armitage 2010).

In Escherichia coli, MCPs and chemotaxis proteins have been shown to localize as large protein signaling clusters at the cell poles and along the cell body (Maddock and Shapiro 1993; Sourjik and Berg 2000; Thiem et al. 2007). Recent reports have suggested that these clusters are formed by stochastic self-assembly, which yields a somewhat regular distribution of clusters along the cell length (Thiem and Sourjik 2008). This distribution is similar to the positioning of E. coli's peritrichous flagella and is believed to facilitate a rapid response to chemotactic stimuli because the diffusion time of phosphorylated CheY to a flagellar motor is minimized (Sourjik and Berg 2002; Lipkow et al. 2005). In contrast, V. cholerae, the cause of the diarrheal disease cholera, is monotrichous, and its single flagellum localizes at the old cell pole. To date, the subcellular distribution of V. cholerae chemotaxis proteins has not been reported. The majority of the che genes are encoded in three distinct clusters, while the mcp genes are scattered throughout the genome (Heidelberg et al. 2000; Boin et al. 2004). Thus far, only chemotaxis proteins from cluster II, the main chemotaxis operon, have been shown to be required for chemotaxis (Gosink et al. 2002; Hyakutake et al. 2005). Notably, in addition to the core structural genes of the chemotactic complex, cluster II also encodes a ParA homolog, VC2061. Unlike V. cholerae's other ParA homologs, ParA1 and ParA2, which mediate segregation of the origin regions of chromosome I and II, respectively (Fogel and Waldor 2005, 2006), a role for VC2061 has not been identified.

Here, we explored the intracellular localization of chemotaxis proteins in V. cholerae and how their localization is regulated and coordinated with the cell cycle. We show that CheW1, CheY3, and VC2061, which are all encoded in the main V. cholerae chemotaxis operon, all localize to the old flagellated pole in newborn cells. Then, as cells elongate, VC2061 is recruited to the new pole, where it facilitates the recruitment of CheW1 and CheY3. Thus, VC2061, hereafter called ParC (for partitioning chemotaxis), promotes the maturation of V. cholerae’s new pole, readying this site for its development into a functional old pole. Importantly, ATP hydrolysis is required for ParC function, and thus, in contrast to E. coli, it appears that active rather than stochastic processes mediate the subcellular distribution of chemotactic proteins in V. cholerae.

Results

CheW1 and CheY3 polar localization changes in a cell cycle-dependent fashion

We investigated the subcellular localization of proteins encoded by V. cholerae’s principal chemotaxis operon (cluster II), which has been implicated in the organism's pathogenicity and transmission (Fig. 1A; Lee et al. 2001; Butler and Camilli 2004). Fusions of the adaptor protein CheW1 and the response regulator CheY3 to yellow and cyan fluorescent proteins (Yfp and Cfp, respectively) were ectopically expressed and visualized in V. cholerae. Both proteins localized as distinct foci at one (Fig. 1B,C, white bars) or both (Fig. 1B,C, red bars) cell poles. The majority of young (short) cells contained a single unipolar Yfp-CheW1 or CheY3-Cfp focus, whereas older (longer) cells contained bipolar foci (Fig. 1D), indicating that the second focus develops as cells mature. Time-lapse microscopy experiments confirmed this hypothesis (Fig. 1E). Newborn cells were found to have a single CheW1 focus at one pole; then, as cells elongated, a second CheW1 focus formed at the distal pole (Fig. 1E, red bars), resulting in bipolar localization of CheW1. Subsequently, the cells divided and split into two daughter cells, each with a unipolar CheW1 focus at its old flagellated pole (Fig. 1E, 24′). An identical distribution and progression was observed in cells where the chromosomal cheW1 locus was replaced with a functional yfp-cheW1 (Supplemental Fig. S1A,B), indicating that the observed localization did not result from overexpression of the fusion protein or inactivation of its normal function. The distinct distribution of these chemotaxis proteins indicated that there may be a cell cycle-dependent factor facilitating maturation of the new pole, enabling recruitment of the chemotaxis proteins. The apparent recruitment of CheW1 and CheY3 to the new cell pole in predivisional cells could be a means of ensuring that cell division yields daughter cells with chemotaxis proteins in close proximity to the site of flagellar assembly after cell division.

Figure 1.

Figure 1.

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Cell cycle-dependent changes in CheW1 and CheY3 changes localization. (A) Schematic of the main V. cholerae (cluster II) chemotaxis operon. (B,C) Fluorescence microscopy of Yfp-CheW1 (B) and CheY3-Cfp (C) in wild-type V. cholerae cells. (D) Plot depicting the polar localization pattern (unipolar vs. bipolar) relative to cell length of Yfp-CheW1 and CheY3-Cfp. (E,F) Time-lapse fluorescence microscopy series of wild-type (E) and ΔparC (F) V. cholerae cells expressing Yfp-CheW1. White numbers indicate minutes from the start of the time lapse. Orange numbers and colored bars indicate cells discussed in the text.

Members of a new family of ParA-like proteins are encoded within the chemotaxis operons of many γ-proteobacteria

In addition to chemotaxis proteins, V. cholerae’s cluster II chemotaxis operon (Fig. 1A) also encodes a protein (VC2061, renamed here ParC) that exhibits 31% identity and 52% similarity to V. cholerae’s ParA1 (Supplemental Fig. S2A). A search of the STRING database for ParC homologs and their genetic context revealed that other members of the Vibrio family and several additional γ-proteobacteria, including Pseudomonas, Aeromonas, Pseudoalteromonas, Shewanella, and Xanthomonas, as well as the δ-proteobacterium Desulfuvibrio, also have ParC homologs located within chemotaxis operons (Supplemental Figs. S2B, S3). Phylogenetic comparison (Fig. 2A) between ParA proteins involved in chromosome segregation and in plasmid partitioning and ParA-like proteins encoded in chemotaxis operons showed that these proteins group into three discrete clades. Consistent with previous analyses (Gerdes et al. 2000), clade I is comprised of chromosome-encoded ParA proteins that, in some cases, have been implicated in chromosome segregation (Fig. 2A, green lines), and clade II is largely comprised of ParA proteins encoded by plasmids that, in several cases, have been implicated in plasmid segregation (Fig. 2A, red lines). Remarkably, a group of previously uncharacterized ParA-like proteins encoded within chemotaxis operons, including ParC, form their own distinct clade (clade III) (Fig. 2A, blue lines). In contrast to the parA genes involved in chromosome and plasmid partitioning, which are all encoded adjacent to parB genes, none of the parA-like genes found in clade III are flanked by a parB homolog (Supplemental Fig. S3). Interestingly, all species that encode a clade III ParA-like protein have polar flagella. Clade II also contains ParA-like proteins encoded within chemotaxis operons, including PpfA, which has been shown to be involved in regular positioning of cytoplasmic chemotaxis clusters along the length of the cell in R. sphaeroides (Thompson et al. 2006). Thus, there appear to be two distinct sets of ParA homologs encoded within chemotaxis operons.

Figure 2.

Figure 2.

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Cell cycle-dependent changes in ParC localization. (A) Phylogenetic tree of chromosome-encoded ParA proteins likely to contribute to chromosome segregation (green), plasmid-encoded ParA proteins (red), and ParA-like proteins located on the chromosome within chemotaxis operons (blue). (B) Fluorescence microscopy of Yfp-ParC in wild-type V. cholerae cells. (C) Plot depicting the polar localization pattern (unipolar vs. bipolar) of Yfp-ParC relative to cell length. (D) Time-lapse fluorescence microscopy of Yfp-ParC in wild-type V. cholerae cells. White numbers indicate minutes from the start of the time lapse. White bars indicate cells discussed in the text. (E) Fluorescence microscopy showing the subcellular localization of ParC homologs from V. parahaemolyticus (ParCVp) and A. hydrophila (ParCAh) in V. cholerae and of ParCVp in V. parahaemolyticus.

ParC exhibits a similar pattern of subcellular localization as CheW1 and CheY3

We analyzed the subcellular localization of ParC using a Yfp-ParC fusion protein. The distribution of Yfp-ParC was similar to that of CheW1 and CheY3 (Fig. 2B), with either distinct unipolar (Fig. 2B, white bar) or bipolar (Fig. 2B, red bar) foci observed; nonpolar foci were not detected. Like CheW1 and CheY3, ParC localized to the old flagellated pole in newborn cells. Later in the cell cycle, it was recruited to the new pole, resulting in a bipolar distribution, and after cell division, daughter cells each inherited a single ParC focus at the old pole (Fig. 2C,D). In contrast to ParA proteins involved in DNA segregation (Gerdes et al. 2010), we did not observe localization of ParC to the nucleoid. Furthermore, Yfp-ParC localization was not affected by deletion of the entire cluster II chemotaxis operon (Supplemental Fig. S1D), demonstrating that ParC's polar localization is independent of cluster II chemotaxis proteins. Yfp-ParC expressed from its endogenous promoter was functional (Supplemental Fig. S1A) and its localization identical to that observed when this fusion protein was expressed from a plasmid (Supplemental Fig. S1C). Thus, at least three cluster II-encoded proteins, ParC, CheW1, and CheY3, exhibit a similar dynamic cell cycle-dependent localization.

To investigate whether the pattern of ParC localization was conserved in other species, Yfp fusion proteins of the ParC homologs from Vibrio parahaemolyticus (ParCVp) and Aeromonas hydrophila (ParCAh) were created. Both ParCVp and ParCAh exhibited cell cycle-dependent uni- and bipolar patterns of localization identical to that of ParC when expressed in V. cholerae, as did ParCVp when expressed in its native host (Fig. 2E; Supplemental Fig. S4). Thus, the pattern of cell cycle-dependent polar localization and redistribution is conserved in these ParC homologs, indicating that clade III ParC proteins have similar functions.

ParC is recruited to the new pole earlier in the cell cycle than CheW1 and CheY3

The observation that ParC localized to the pole in the absence of other cluster II proteins suggested that this protein might guide localization of chemotaxis proteins to the cell pole. Analyses of cells coexpressing Cfp-ParC and Yfp-CheW1 were consistent with this idea (Fig. 3). In static images, three distinct localization patterns were observed: (1) unipolar colocalization of ParC and CheW1 foci at the old pole (Fig. 3A, green bar), (2) bipolar ParC and unipolar CheW1 foci (Fig. 3A, white bar), and (3) bipolar colocalization of both ParC and CheW1 foci (Fig. 3A, red bar). More cells had bipolar foci of Cfp-ParC than of Yfp-CheW1 (∼70% and 25%, respectively), and all cells with bipolar CheW1 foci also showed bipolar localization of ParC (Fig. 3B). Identical results were observed for the colocalization of Yfp-ParC and CheY3-Cfp (Supplemental Fig. S5). Plots of the distance of foci from the old cell pole as a function of cell length revealed that bipolar ParC foci become detectable in younger cells than do bipolar CheW1 and CheY3 foci (Fig. 3C; Supplemental Fig S5C), indicating that ParC is recruited to the new pole before CheW1/CheY3. Indeed, this was confirmed by time-lapse microscopy (Fig. 3D), which showed that ParC foci form at the new pole earlier in the cell cycle than CheW1 foci. Collectively, these analyses revealed that recruitment of ParC and chemotaxis proteins from the old to the new pole is at least a two-step cell cycle-dependent process.

Figure 3.

Figure 3.

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ParC localizes at the new cell pole before CheW1. (A) Fluorescence microscopy of wild-type cells coexpressing Cfp-ParC and Yfp-CheW1. Green bars indicate representative cells with unipolar localization of both Cfp-ParC and Yfp-CheW1. White bars indicate representative cells with bipolar localization of Cfp-ParC and unipolar localization of Yfp-CheW1. Red bars indicate representative cells with bipolar localization of both Cfp-ParC and Yfp-CheW1. (B) Graph depicting the percentage of cells with uni- and bipolar localization of Cfp-ParC and Yfp-CheW1, respectively. (C) Distance of Cfp-ParC and Yfp-CheW1 foci from the old pole versus cell length. Dashed lines show the cell length where a focus at the new pole is detected for the first time. (D) Time-lapse series of Cfp-ParC and Yfp-CheW1 in wild-type V. cholerae cells. Colored bars indicate cells discussed in the text.

ParC promotes proper polar localization of CheW1 and CheY3

To investigate whether ParC aids recruitment of CheW1/CheY3 to the new pole, Yfp-CheW1 (Fig. 4A, panel b) and CheY3-Cfp (Fig. 4B, panel b) localizations were analyzed in a strain lacking parC (strain ΔparC). In the ΔparC background, ∼25% of cells contained mislocalized (nonpolar) Yfp-CheW1 or CheY3-Cfp foci (Fig. 4A [panel b], B [panel b, white bars],C), which were positioned randomly relative to the cell pole (Fig. 4D). In contrast, <2% of wild-type cells showed nonpolar foci (Fig. 4C). The mislocalization of chemotaxis proteins in ΔparC cells was due to the absence of ParC (Fig. 4C), as this phenotype could be complemented to almost wild-type levels by expressing ParC in trans (Fig. 4C). Imaging of Yfp-CheW1 and CheY3-Cfp in the ΔparC strain revealed that both polar and nonpolar Yfp-CheW1 and CheY3-Cfp foci colocalized (Fig. 4E), suggesting that the entire chemotaxis apparatus is mislocalized in the absence of parC. In some cells, only aberrant Yfp-CheW1 and CheY3-Cfp foci were observed, whereas in other cells, unipolar or bipolar foci along with aberrant foci were seen. Furthermore, we observed cells with no foci at all (Fig. 4A,B, red bars). Time-lapse series showed that, in the ΔparC background, cell division can occur before bipolar localization of CheW1 and CheY3 has been established (Fig. 1F, cells 2 and 3). Thus, ParC enables accurate inheritance of the chemotactic machinery by promoting recruitment of chemotaxis proteins to the new pole before cell division. In cell 2′, an aberrant CheW1 focus was subsequently formed near the mid-cell (Fig. 1F, 36′, red bar), demonstrating that aberrant CheW1 foci need not migrate to their positions from a cell pole. In cell 3′, a Yfp-CheW1 focus subsequently formed at the new pole without CheW1 localized at its old pole (Fig. 1F, 42′, orange bar). Thus, with respect to CheW1 localization, cell 3′ appears to have lost pole identity and is unable to distinguish between its old and new pole. Finally, cell 1 in Figure 1F displays wild-type localization of Yfp-CheW1, showing that localization of CheW1 is not absolutely dependent on ParC.

Figure 4.

Figure 4.

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Localization of CheW1 and CheY3 in a ΔparC strain. (A) Localization of Yfp-CheW1 in wild-type V. cholerae and ΔparC. (B) Localization of CheY3-Cfp in wild-type V. cholerae and in ΔparC. (C) Percentage of cells with aberrant (nonpolar) Yfp-CheW1 and CheY3-Cfp foci in wild-type V. cholerae, ΔparC, and ΔparC complemented with Cfp- or Yfp-tagged ParC, ParCG11V, and ParCK15Q. (D) Plot showing the distance of polar and nonpolar Yfp-CheW1 and CheY3-Cfp foci to the closest cell pole in V. cholerae ΔparC coexpressing Yfp-CheW1 and CheY3-Cfp. (E) Localization of Yfp-CheW1 and CheY3-Cfp in ΔparC.

To begin to address whether ParC activity is restricted to recruiting cluster II chemotaxis proteins to the cell poles, we examined the localization of several additional proteins in the ΔparC mutant. The localization of ParA1K11E-Yfp, a protein that is ordinarily found only at the cell poles, was not altered in the absence of ParC (Supplemental Fig. S6). Furthermore, the polar localization of CheW3, which is in V. cholerae chemotaxis cluster III, was unchanged in the ΔparC mutant (data not shown). Thus, ParC appears to be specific for the polar localization of cluster II chemotaxis proteins.

ParC is recruited from the old pole to the new pole as the cell cycle progresses

The cell cycle-dependent localization of ParC at the new pole could result from redistribution of ParC from the old pole or by de novo accumulation. In time-lapse series, the formation of the Yfp-ParC focus at the new pole was accompanied by a decrease in the intensity of the fluorescence of the focus at the old pole (Fig. 5A,B). When the polar fluorescence intensities of Yfp-ParC (15 cells) were plotted as a function of time and relative to the initial intensity at the old pole, it was evident that, as the fluorescence intensity of Yfp-ParC at the old pole gradually decreased, there was a concomitant gradual increase in intensity at the new pole (Fig. 5B). Together, these observations suggested that, as the cell cycle progresses, ParC is released from the old pole and relocalizes at the new pole.

Figure 5.

Figure 5.

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Dynamic localization and redistribution of ParC during the cell cycle. (A, panel a) Time-lapse series of Yfp-ParC in a single cell. Numbers indicate the time elapsed after the first image in minutes. (Panel b) Three-dimensional surface intensity plot of the corresponding time frames in panel a, revealing that, as fluorescence intensity decreases at the old pole, it increases at the new pole. (B) Graph depicting the fluorescence intensity of Yfp-ParC at the cell poles relative to the initial intensity at old pole during time-lapse series. The averages of 15 cells are shown along with standard error mean. (C, panel a) Photoactivation of PAmCherry-ParC in wild-type V. cholerae. Numbers indicate minutes pre- and post-activation. Dashed circles show activated region, the red arrow indicates the signal from PAmCherry-ParC localized to the old pole after activation, and the green arrow indicates PAmCherry-ParC that has relocalized from the old pole as a focus at the new pole. (Panel b) Graph depicting the fluorescence intensities at the old and new pole pre- and post-activation in the time lapse shown in panel a. (D, panel a) FRAP microscopy of unipolar Yfp-ParC in wild-type V. cholerae showing that bleached regions of ParC at the cell pole recover post-bleaching. Numbers indicate minutes pre- and post-FRAP. Dashed circles show bleached regions, and arrowheads indicate areas of Yfp-ParC recovery post-bleaching. (Panel b) Graph depicting fluorescence intensity of the red dashed circle in panel a pre- and post-bleaching.

Indeed, photoactivation of PAmCherry-ParC showed that ParC redistributes from the old to the new pole as the cell cycle progresses. Laser activation of PAmCherry-ParC at the old pole yielded polar ParC foci (Fig. 5C, panel a, red arrow). Within 10 min after activation, fluorescence was also detected in the cytoplasm (Fig. 5C, panel a), and fluorescence intensity at the old pole decreased simultaneously (Fig. 5C, panel b), showing that a portion of PAmCherry-ParC is released from the pole into the cytoplasm over time. By 30 min after activation, a PAmCherry-ParC focus was detected at the new pole (Fig. 5C, panels a,b, green arrow). Thus, some ParC molecules that were originally located at the old pole are released and relocalize as a focus at the new pole.

FRAP (fluorescent recovery after photobleaching) experiments confirmed that there is an exchange between polar and cytoplasmic ParC. When unipolar Yfp-ParC foci were photobleached, gradual recovery of polar fluorescence was observed (Fig. 5D, panels a,b), showing an exchange between the pool of diffuse cytosolic ParC with polarly localized ParC. No exchange of ParC between the cytosol and the new pole was detected during the time course of the FRAP experiments, further establishing the specificity of ParC for the old pole during the early parts of the cell cycle. These results, coupled with those presented above, show that, in young cells, ParC can be either in the cytoplasm, where it appears to diffuse freely, or localized as a focus at the old pole. Then, late in the cell cycle, a portion of ParC is recruited to the new pole, perhaps because a cell cycle-dependent anchor that enables ParC to be captured and accumulated develops at this site.

The ParB1/parS1 complex arrives at the new pole before ParC

The cell cycle-coupled temporal redistribution of ParC is reminiscent of that of ParB1, which binds to origin-proximal parS1 sites (Fogel and Waldor 2006). After replication of the oriCI region, one ParB1/parS1 complex remains at the old pole, while the other moves across the cell to the new pole in a ParA1-dependent fashion (Fig. 6A; Fogel and Waldor 2005, 2006). We used fluorescently tagged ParB1 (Cfp-ParB1) in conjunction with Yfp-ParC to determine whether there is any correlation between the movement of the ParB1/parS1 complex and ParC (Fig. 6A,B). In young cells, ParC and ParB1 foci were both found at the old pole (Fig. 6A [panel a], B). However, in cells with two ParB1 foci, only one of which was polar (i.e., when one of the replicated Cfp-ParB1/parS1 complexes was migrating to the new pole), YFP-ParC exclusively colocalized with the ParB1 focus that remained at the old pole (Fig. 6A, panel b). The arrival of the ParB1/parS1 complex at the new pole always preceded the appearance of ParC at this site (Fig. 6A [panel c], B). Thus, these two proteins do not appear to move in tandem to the new pole, an idea consistent with the observation that a subset of ParC moves gradually from the old pole to the new pole (Fig. 5). Furthermore, ParC movement to the new pole is not dependent on prior arrival of ParB1, as ParC still formed bipolar foci in a parA1 mutant (Supplemental Fig. S7), in which ParB1 is not targeted to the cell poles (Fogel and Waldor 2006; Yamaichi et al. 2007). Together, these data indicate that new pole maturation has at least three discrete stages: (1) arrival of the segregating ParB1/parS1 complex at the new pole, (2) recruitment of ParC, and (3) formation of chemotaxis protein clusters before cell division. Thus, new pole maturation appears to be an ordered process marked by the successive formation of polar protein complexes that control chromosome segregation and chemotaxis.

Figure 6.

Figure 6.

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The Walker motif of ParC is required for its ATPase activity and proper polar localization. (A,B) Spatiotemporal relationship between ParB1 localization and recruitment of ParC to the new pole. (A) Fluorescence microscopy of Cfp-ParB1 and Yfp-ParC in wild-type V. cholerae. (B) The distance of Yfp-ParC and Cfp-ParB1 foci from the old cell pole in wild-type V. cholerae plotted as a function of cell length. (C,D) ATPase activities of His-ParC and His-ParCG11V as a function of : ATP concentration (C), and protein concentration (D). (C) Protein was added to a final concentration of 10 μM. ATP was added in 5 mM, 2.5 mM, 1.25 mM, 0.63 mM, 0.31 mM, 0.16 mM, and 0.08 mM. (D) ATP-hydrolysis as function of protein concentration using 2 mM ATP. Protein was added in 5 μM, 2.5 μM, 1.25 μM, 0.63 μM, 0.31 μM, and 0.16 μM. Fluorescence microscopy of Yfp-ParCK15Q (E) and Yfp-ParCG11V (F) in V. cholerae ΔparC.

ParC ATPase activity is required for its localization and capacity to recruit chemotaxis proteins

ParC, like other members of the ParA protein family, contains a Walker-type ATPase motif (Supplemental Fig. S2). Purified His-tagged ParC possessed intrinsic ATPase activity, with a Vmax of 3.3 mM ATP hydrolyzed per minute per mole ParC and a KM of 150 μM ATP (Fig. 6C,D), values similar to those reported for other ParA family members (Barilla et al. 2005). Based on previously characterized mutations within ParA family proteins (Leonard et al. 2005), we introduced single amino acid substitutions into the putative Walker-A ATP-binding pocket of ParC. These substitutions are expected either to prevent ATP binding (ParCK15Q) or to prevent ATP hydrolysis while still permitting binding (ParCG11V) (Leonard et al. 2005; Murray and Errington 2008). As predicted, His-tagged ParCG11V lacked ATPase activity (Fig. 6C,D). Neither ParCG11V nor ParCK15Q displayed the typical localization pattern of wild-type ParC. Yfp-ParCK15Q was distributed diffusely throughout the cytoplasm (Fig. 6E); in contrast, Yfp-ParCG11V largely formed uni- and bipolar foci, but nonpolar foci were also observed (Fig. 6E, white bars). Neither ParC point mutant restored wild-type localization to CheY3 in the ΔparC background (Fig. 4C). The stability of the ParC point mutants was not significantly altered by the amino acid substitutions (data not shown). Therefore, the lack of function of the mutant proteins is likely attributable to loss of function rather than their instability. Thus, the capacity of ParC to bind and hydrolyze ATP not only regulates its localization, but is essential for its capacity to promote the polar localization of chemotaxis proteins.

A parC mutant is impaired in chemotaxis

Given the role of ParC in positioning cluster II chemotaxis proteins at the V. cholerae poles, we tested whether absence of parC alters V. cholerae’s chemotactic capacity. Initially, we assessed the ability of wild-type V. cholerae, the ΔparC mutant, and a strain lacking the entire cluster II chemotaxis operon (SR28) to swarm in soft agar. As expected, SR28 failed to swarm on these plates. In contrast, the ΔparC mutant exhibited a 10% reduction in swarming ability (Supplemental Fig. S8A,B). This relatively minor phenotype may reflect the fact that a significant proportion of cells lacking parC still have correctly positioned, and presumably functional, chemotactic apparati.

To further explore the contribution of parC to V. cholerae chemotaxis, we tested whether deletion of this gene alters the bacteria's propensity for linear motility. We tracked swimming cells (Fig. 7A) to determine the frequency with which they reverse their direction. The ΔparC mutant reversed direction much less frequently (0.28 s−1) than did the wild-type V. cholerae (0.8 s−1) (Fig. 7B), showing that ΔparC displays CCW-biased flagellar rotation (Fig. 7A,B), presumably due to impaired chemotactic signaling. For a subset of the ΔparC mutant cells, no reversals were detected within the time of recording (Fig. 7C, dashed circle), indicating an extreme CCW bias; however, all other cells showed a reduced (relative to wild-type cells) but still detectable frequency of reversal (Fig. 7C, columns 3,4). It is possible that this duality reflects the two populations of parC mutants noted in Figure 4; namely, those lacking or with nonpolar foci and those with polar chemotaxis protein foci.

Figure 7.

Figure 7.

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V. cholerae ΔparC has lower swimming reversal rate and elevated intestinal colonization versus wild type. When present, error bars show standard error mean. (A–C) Positional tracking of swimming cells and reversal rates showing that ΔparC exhibits a decreased reversal rate compared with wild type. (A) Representative images of positional tracking of swimming cells. Each colored line represents a single cell's swimming path. (B) Average reversal rate in swimming direction of wild-type, ΔparC, and ΔparC expressing Yfp-ParC in three independent experiments. Three asterisks indicate a difference with a P < 0.0001; the single asterisk indicates P < 0.05. (C) Scatter plot of the reversal rates for all tracked cells of wild-type and ΔparC V. cholerae. The dashed circle indicates the group of ΔparC cells that do not reverse in swimming direction during the time period in which they were tracked. Three asterisks indicate a difference with a P < 0.0001, and two asterisks indicate a difference of P < 0.001, compared with wild type. (D,E) Intestinal colonization of infant mice with wild type and ΔparC. (D) Competition experiment between wild type and ΔparC in the small intestine (SI) of infant mice and during in vitro growth in LB. Asterisks indicate a difference with a P < 0.0001, compared with in vitro competition. (E) A model depicting cell pole maturation and the specific spatiotemporal recruitment of proteins to the new pole during the V. cholerae cell cycle.

The ΔparC mutant outcompetes wild-type V. cholerae in intestinal colonization

Previous analyses have demonstrated that disruption of V. cholerae’s chemotactic response can alter its capacity to colonize the small intestine of suckling mice (Lee et al. 2001; Butler and Camilli 2004), a widely used model to study cholera (Angelichio et al. 1999). In this model, wild-type V. cholerae mainly colonizes the middle and distal parts of the small intestine (Angelichio et al. 1999). Notably, in vivo competition assays revealed that the ΔparC mutant is more proficient at colonization than wild-type V. cholerae, with an especially pronounced competitive advantage in the proximal part of the small intestine (Fig. 7D). The ΔparC mutant does not exhibit enhanced growth in general, since no competitive advantage over a wild-type strain was observed in an in vitro competition assay (Fig. 7D). A cheY3 mutant, which, like the ΔparC mutant, exhibits a CCW swimming bias, was also found to have an enhanced capacity to colonize the proximal small intestine (Butler and Camilli 2004). Thus, the ectopic colonization of the upper small intestine by the ΔparC mutant likely relates to ParC's role in promoting the polar localization of cluster II chemotaxis proteins and the impairment in chemotaxis (i.e., CCW-biased swimming) (Fig. 7A–C) that accompanies their mislocalization.

Discussion

Together, our findings show that V. cholerae’s ParA-like protein, ParC, modulates the subcellular distribution of chemotaxis proteins in a cell cycle-dependent fashion, and thereby promotes the development of the new cell pole. ParC is a member of a new family (clade III) of ParA homologs that are more closely related to each other than to homologs that mediate chromosome and plasmid segregation. All clade III ParC homologs are encoded within chemotaxis operons in organisms that produce a polar flagellum. Thus, like V. cholerae, these organisms have a biological rationale for polar targeting of chemotactic machinery. By restricting the subcellular distribution of chemotaxis proteins in young cells to the old pole, ParC likely facilitates efficient transmission of the CheY-P signal to its target at the flagellum. Additionally, ParC-mediated localization of chemotaxis proteins at the new pole as the cell matures readies this site for transformation into a functional old pole following completion of cell division.

Not all ParA homologs encoded within chemotaxis operons cluster with ParC. Instead, a subset of these proteins (including the R. sphaeroides protein PpfA) appears more closely related to the ParAs that mediate plasmid segregation (clade II). PpfA is required for regular localization of cytoplasmic chemotaxis clusters (Wadhams et al. 2003; Thompson et al. 2006). It was previously proposed that ParA homologs in chemotaxis gene clusters exclusively aid localization of such cytoplasmic chemotaxis complexes (Hamer et al. 2010). However, our data indicate that only the subgroup of these proteins that belong to clade II (e.g., PpfA) are involved in segregation of cytoplasmic chemotaxis clusters. It seems likely that clade II and clade III ParA homologs use distinct mechanisms to modulate the subcellular distribution of cytoplasmic and membrane-associated chemotaxis proteins, respectively.

Several cellular distributions have previously been observed for ParA homologs (Ebersbach et al. 2006; Fogel and Waldor 2006; Hatano et al. 2007; Ringgaard et al. 2009; Ptacin et al. 2010); however, the localization of ParC does not appear concordant with any of them. Collectively, our observations indicate that a “diffusion and capture” model (Rudner et al. 2002) can explain the cell cycle-dependent changes in the subcellular distribution of ParC. In such a model, early in the cell cycle there is a continuous exchange between ParC at the old pole and the cytoplasm. Then, later in the cell cycle, an as yet to be discovered ParC anchor develops at the new pole, which allows ParC to be captured there. Consequently, ParC released from the old pole will diffuse through the cytoplasm and relocalize at the new pole (Fig. 7E), where it accumulates. The temporal progression of ParC's subcellular distribution indicates that the polar anchoring mechanism only becomes active at the new pole late in the cell cycle.

Correct positioning of membrane-associated chemotaxis proteins has not previously been shown to result from an active process. In E. coli, clustering and localization of chemotaxis proteins is believed to be a largely stochastic process, resulting from insertion of chemotactic receptor proteins at sites throughout the cell membrane, followed by diffusion-mediated cluster formation that is independent of cytoskeletal guidance or a dedicated targeting mechanism (Thiem and Sourjik 2008; Greenfield et al. 2009). MCP receptor proteins are sufficient to mediate cluster formation (Kentner et al. 2006); however, CheA and CheW are both thought to stabilize receptor complexes (Skidmore et al. 2000; Kentner et al. 2006). In contrast, V. cholerae is dependent on ParC for precise localization of chemotaxis proteins, presumably because its chemotactic apparati are to be limited to the cell pole(s) rather than regularly distributed along the cell periphery. Given what is known about assembly of chemotactic complexes in E. coli, one possible role for ParC is to promote polar localization and/or retention of MCP receptors, either via interacting with them directly or through modulating their interactions with other factors, such as CheW and/or CheA. Although the precise mechanism of targeting by ParC remains to be elucidated, it is clearly an active process that requires nucleotide binding and hydrolysis.

We presume that the underlying biological imperative for targeting of V. cholerae’s chemotaxis apparatus to the cell pole is to couple it spatially with the bacterium's single polar flagellum. Such optimization is not needed in peritrichously flagellated organisms such as E. coli, in which passive processes are sufficient to ensure coordinated positioning. In both organisms, colocalization of chemotaxis clusters and flagella minimize the need for diffusion of CheY between the two apparati, and hence facilitate a rapid response to environmental stimuli. The observed reduced chemotaxis and altered swimming behavior of the ΔparC mutant, coupled with its mislocalization of chemotaxis proteins, is consistent with this hypothesis. The mutant's bias toward straight swimming (i.e., CCW flagellar rotation) likely reflects altered transmission of a chemotaxis-based signal to the flagellar motor, perhaps due to imprecise protein positioning, and also probably accounts for its hypercolonization of infant mice. Hypercolonization was also observed using specific cheW and cheY mutants with CCW-biased flagellar rotation, whose prolonged periods of straight (CCW-biased) swimming is thought to enable expanded host tissue range (Butler and Camilli 2004).

Unlike the observed bipolar distribution of chemotactic apparati in mature cells, bipolar flagella have not been reported. Furthermore, the cell cycle-dependent polar localization of ParC and chemotaxis proteins is independent of FlhF, a protein required for polar positioning of the flagellum (Green et al. 2009) and of the flagellar MS ring component FliF (Supplemental Fig. S9), suggesting that the positioning of the flagellar and chemotactic machineries occur through independent pathways. Thus, the late targeting of chemotaxis proteins to the new pole is unlikely to influence cellular motility prior to cell division. Why, then, are they needed at this site? Presumably, the preplacement of chemotaxis proteins at the maturing new pole by ParC ensures that all bacteria inherit a chemotactic cluster localized at the old cell pole after cell division. This chemotactic cluster enables the newborn cell to respond to its environment as soon as it completes flagellar assembly. By promoting the inheritance of the chemotaxis cluster, ParC function is analogous to that of ParA proteins that promote inheritance of chromosomal and plasmid DNA. However, the independence of ParC- and ParA1-mediated recognition of the new pole (marked by emergence of bipolar ParC/chemotaxis and ParB1 foci, respectively) (Fig. 7E) demonstrates that multiple distinct processes underlie recognition and maturation of this cellular domain, which collectively culminate with its transformation into a functional old pole after completion of cell division.

Materials and methods

Growth conditions and media

In most experiments, V. cholerae, V. parahaemolyticus, A. hydrophila, and E. coli were grown in LB medium or on LB agar plates at 37°C containing antibiotics in the following concentrations: 200 μg/mL streptomycin, 50 μg/mL kanamycin, 100 μg/mL ampicillin, 50 μg/mL carbenicillin, and 20 μg/mL chloramphenicol for E. coli; and 5 μg/mL for V. cholerae, V. parahaemolyticus, and A. hydrophila.

Strains and plasmids

The strains and plasmids used in this study are listed in Supplemental Tables S1 and S2, respectively. Primers used are listed in Supplemental Table S3. All V. cholerae strains used were derived from the El Tor clinical isolate C6706. E. coli strains DH5αλpir and SM10λpir were used for cloning. E. coli strain SM10λpir was used to transfer DNA into V. cholerae by conjugation (Miller and Mekalanos 1988). Construction of V. cholerae deletion mutants was performed with standard allele exchange techniques using derivatives of plasmid pCVD442 (Donnenberg and Kaper 1991). Strains ΔparC, SR6, SR14, and SR28 were created using pCVD442 derivative plasmids pCVD442-ΔVC2061, pSR1010, pSR1009, and pSR1020, respectively. See the Supplemental Material for details of plasmid and strain construction.

Measuring the reversal frequency in swimming direction

To measure the reversal frequency in swimming direction of V. cholerae cells, 10 μL of an overnight culture grown in LB was transferred to 5 mL of fresh LB. Cells were grown to OD600 = 0.4. Then, 2 μL was spotted on an eight-well printed microscope slide, covered with a coverslip, and immediately used for microscopy. Images were recorded every 112 msec using the streaming acquisition function in the Metamorph software. Three individual experiments were performed, and in each experiment, 20–30 cells were analyzed. Individual cells were tracked using the MTrackJ plug-in for ImageJ imaging software. Cells were tracked for as long as possible, the number of reversals was counted, and the reversal frequency was calculated as the number of reversals per second per cell. Then, for each experiment, the average reversal frequency was calculated.

Fluorescence and time-lapse microscopy

Five milliliters cultures of cells harboring the relevant plasmid was grown in LB medium to OD600 ≈ 0.1, and expression of fluorescent fusion proteins was induced by addition of L-arabinose to a final concentration of 0.2%. The cultures were incubated for an additional 2 h. Cells were then mounted on 1% agarose in PBS buffer on microscope slides, and microscopy was performed. For time-lapse microscopy, cells were mounted on 1% agarose in PBS buffer containing 10% LB medium. For colocalization and time-lapse microscopy of Yfp-ParC and Cfp-ParB1, strain ΔparC harboring pSR1028 was inoculated in M9 minimal medium supplemented with 0.02% casamino acids, 0.2% glycerol, and 1 μg/mL thiamine. Cells were grown for 3 h before adding L-arabinose to a final concentration of 0.2%. After incubating the cultures for an additional 2 h, samples were mounted on 1% agarose in the same medium and incubated 30 min at ambient temperature before time-lapse microscopy was performed. Microscopy images were analyzed using ImageJ imaging software (http://rsbweb.nih.gov/ij).

For single-labeling fluorescence microscopy of Yfp-ParC, Cfp-ParC, PAmCherry-ParC, Yfp-ParCG11V, Yfp-ParCK15Q, Yfp-CheW1, CheY3-Cfp, Yfp-AHA1389, Yfp-VP2227, and Yfp-PA1462, the fusion proteins were expressed ectopically from plasmids pSR1024, pSR1027, pSR1037, pSR1025, pSR1026, pSR1033, pBAD33-parA1K11E-yfp-cheY3-cfp, pSR1034, pSR1035, and pSR1036, respectively. For double labeling of Cfp-ParC/Yfp-CheW1, Yfp-ParC/CheY3-Cfp, Yfp-ParC/Cfp-ParB1, ParA1K11E-Yfp/CheY3-Cfp, Yfp-CheW1/CheY3-Cfp proteins were expressed ectopically from plasmids pSR1028, pSR1029, pSR1032, and pBAD33-parA1K11E-yfp-cheY3-cfp, pSR1038, respectively.

FRAP and photoactivation microscopy

Cells were treated and mounted on agarose pads as described for time-lapse fluorescence microscopy. Microscopy was performed using a Nikon Ti motorized inverted microscope with Perfect Focus System and images obtained with a Hamamatsu ORCA-R2 cooled CCD camera. FRAP and photoactivation were performed using Photonic Instruments MicroPoint laser targeting system. For FRAP experiments, areas of interest were bleached with five pulses using a 405-nm laser at 50% intensity. For photoactivation, areas of interest were activated with five pulses using a 405-nm laser at 35% intensity.

Swarming assay

V. cholerae cells were grown in 5 mL of LB overnight at 37°C. A toothpick was dipped in the overnight culture and pricked into semisolid LB agar plates solidified with 0.3% agar. The plates were incubated overnight at 30°C, and the swarm diameter of each strain was measured and compared with wild type.

Infant mouse infection assay

Competition and single-infection experiments in suckling mice were carried out as described in Angelichio et al. (1999).

Acknowledgments

We are grateful to Professor Suzanne Walker for use of equipment and materials. We thank Felipe Cava for helpful discussions and construction of strain V. cholerae ΔparC and for plasmid pCVD442-ΔVC2061. We are grateful to the M.K.W. laboratory members and Professor David Rudner at Harvard Medical School for helpful comments on the manuscript. We thank the Nikon Imaging Center at Harvard Medical School for help with light microscopy; in particular, Wendy C. Salmon. This work was funded by grants from NIH (R37AI-42347) and HHMI to M.K.W. S.R. was funded with a post-doctoral fellowship from the Villum Kann Rasmussen Foundation, and K.S. was supported with a post-doctoral fellowship by the Deutsche Forschungsgemeinschaft (DFG).

Footnotes

Supplemental material is available for this article.

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