Functional dichotomy and distinct nanoscale assemblies of a cell cycle-controlled bipolar zinc-finger regulator

Protein polarization underlies differentiation in metazoans and in bacteria. How symmetric polarization can instate functional asymmetry remains elusive. Here, we show by super-resolution photo-activated localization microscopy and edgetic mutations that the bitopic zinc-finger protein ZitP implements specialized developmental functions – pilus biogenesis and multifactorial swarming motility – while shaping distinct nanoscale (bi)polar architectures in the asymmetric model bacterium Caulobacter crescentus. Polar assemblage and accumulation of ZitP and its effector protein CpaM are orchestrated in time and space by conserved components of the cell cycle circuitry that coordinate polar morphogenesis with cell cycle progression, and also act on the master cell cycle regulator CtrA. Thus, this novel class of potentially widespread multifunctional polarity regulators is deeply embedded in the cell cycle circuitry. DOI: http://dx.doi.org/10.7554/eLife.18647.001


Introduction
Some regulatory proteins that execute important developmental, cytokinetic or morphogenetic functions are localized in monopolar fashion, whereas others are sequestered to both cell poles (Dworkin, 2009;Martin and Goldstein, 2014;Shapiro et al., 2002;St Johnston and Ahringer, 2010). It is unclear if bipolar proteins can confer specialized functions from each polar site, but examples of proteins with a bipolar disposition have been reported for eukaryotes and prokaryotes (Davis et al., 2013;Martin and Berthelot-Grosjean, 2009;Tatebe et al., 2008;Treuner-Lange and Sogaard-Andersen, 2014).
The synchronizable Gram-negative a-proteobacterium Caulobacter crescentus (henceforth Caulobacter) is a simple model system to study pole-specific organization and cell cycle control (Tsokos and Laub, 2012). The Caulobacter predivisional cell is overtly polarized and spawns two morphologically dissimilar and functionally specialized daughter cells, each manifesting characteristic polar appendages ( Figure 1A). The swarmer progeny is a motile and non-replicative dispersal cell that samples the environment in search of food. It harbours adhesive pili and a single flagellum at one pole and is microscopically discernible from the stalked cell progeny, a sessile and replicative cell that features a stalk, a cylindrical extension of the cell envelope, on one cell pole. While the stalked cell resides in S-phase, the swarmer cell is in a quiescent G1-like state from which it only exits concomitant with the differentiation into a stalked cell. During this G1fiS transition, the polar flagellum and pili of the swarmer cell are eliminated and replaced by the stalk that elaborates from the vacated cell pole. Upon sequential transcriptional activation of developmental factors during the cell cycle (Panis et al., 2015), the nascent stalked cell re-establishes polarization and ultimately gives rise to an asymmetric pre-divisional cell that yield a swarmer and a stalked progeny.
The GcrA transcriptional regulator predominates in early S-phase (Holtzendorff et al., 2004) ( Figure 1A-B). It accumulates during the G1fiS transition and activates expression of polarity factors that are required for pilus or flagellum biogenesis and cytokinetic components (Davis et al., 2013;Fioravanti et al., 2013;Murray et al., 2013;Quon et al., 1996;Viollier et al., 2002b) ( Figure 1A-B). Among GcrA target promoters, is the promoter controlling expression of the PodJ polar organizer that localizes to the pole opposite the stalk and directs assembly of the Caulobacter pilus assembly (cpa) machine at that site. In this cascade, PodJ recruits the cytoplasmic CpaE protein that then promotes the localization and assembly of CpaC secretin localization ( Figure 1B) (Viollier, 2002a). Another key promoter controlled by GcrA is the one driving expression of the master cell cycle regulator CtrA that induces the synthesis of a second wave of polar and morphogenesis factors in late S-phase including the cpa operon ( Figure 1B). The abundance of CtrA and GcrA is regulated at the level of synthesis and degradation (Collier et al., 2006;Domian et al., 1997) and as a result, cell division spawns a swarmer and stalked cell progeny containing CtrA and GcrA, respectively.
An important polarity determinant in the a-proteobacteria is the conserved matrix protein PopZ ( Figure 1A) that organizes poles by forming a molecular lattice that traps polar determinants and effectors (Bowman et al., 2008;Deghelt et al., 2014;Ebersbach et al., 2008;Grangeon et al., 2015;Laloux and Jacobs-Wagner, 2013). PopZ is bipolar in the Caulobacter predivisional cell and it interacts directly with numerous cell cycle kinases, the ParAB chromosome segregation proteins and cell fate determinants (Holmes et al., 2016). Here, we dissect at the genetic and cytological level the polar localization and function of two poorly characterized trans-membrane proteins, the zinc-finger protein ZitP and the CpaM effector protein, that are polarly localized and that execute eLife digest Living cells become asymmetric for many different reasons and how they do so has been a long-standing question in biology. In some cells, the asymmetry arises because a given protein accumulates at one side of the cell. In particular, this process happens before some cells divide to produce two non-identical daughter cells that then go on to develop in very different ways -which is vital for the development of almost all multicellular organisms. The single-celled bacterium Caulobacter crescentus also undergoes this type of asymmetric division. The polarized Caulobacter cell produces two very different offsprings -a stationary cell and a nomadic cell that swims using a propeller-like structure, called a flagellum, and has projections called pili on its surface.
Before it divides asymmetrically, the Caulobacter cell must accumulate specific proteins at its extremities, or poles. Two such proteins are ZitP and CpaM, which appear to have multiple roles and are thought to interact with other factors that regulate cell division. However, little is known about how ZitP and CpaM become organized at the poles at the right time and how they interact with these regulators of cell division.
Mignolet et al. explored how ZitP becomes polarized in Caulobacter crescentus using a combination of approaches including biochemical and genetic analyses and very high-resolution microscopy. This revealed that ZitP accumulated via different pathways at the two poles and that it formed distinct structures at each pole. These structures were associated with different roles for ZitP. While ZitP recruited proteins, including CpaM, required for assembly of pili to one of the poles, it acted differently at the opposite pole.
By mutating regions of ZitP, Mignolet et al. went on to show that different regions of the protein carry out these roles. Further experiments demonstrated that regulators of the cell division cycle influenced how ZitP and CpaM accumulated and behaved in cells, ensuring that the proteins carry out their roles at the correct time during division. These findings provide more evidence that proteins can have different roles at distinct sites within a cell, in this case at opposite poles of a cell. Future studies will be needed to determine whether this is seen in cells other than Caulobacter including more complex, non-bacterial cells.  Figure 1 continued on next page multiple regulatory functions. We unearthed two separate localization pathways for each cell pole, one PopZ-dependent and another that is PopZ-independent, and we provide evidence by photoactivated localization microscopy (PALM) and by genetic dissection that each polar cluster has a distinctive architecture and a specialized function.

Results
ZitP and CpaM are required for pilus biogenesis.
As pili are necessary for infection by the lytic caulophage CbK (jCbK) (Skerker and Shapiro, 2000), we specifically sought mutants in pilus assembly factors encoded outside of the major pilus assembly cpa gene locus (pilA-cpaA-K) (Christen et al., 2016;Skerker and Shapiro, 2000). To this end, we conducted himar1-transposon (Tn) mutagenesis of wild-type (WT) Caulobacter in the presence of jCbK (see Methods) and recovered mutants with Tninsertions in CCNA_02298, renamed here zitP (zinc-finger targeting the poles) because of the pleiotropic roles detailed below, or in cpaM (CCNA_03552) ( Figure 1C) (Marks et al., 2010). While both genes have previously been implicated in polar functions and their transcription is cell cycle-regulated (Christen et al., 2016;Fioravanti et al., 2013;Fumeaux et al., 2014;Hughes et al., 2010;McGrath et al., 2007), they are poorly characterized. The zitP gene is predicted to encode a 311-residue bitopic trans-membrane (TM) protein harbouring a CXXC-(X) 19 -CXXC motif that binds a zinc ion (zinc_ribbon_5 or PF13719 superfamily, residues 1-37) at the cytoplasmic N-terminus (Bergé et al., 2016) and a conserved domain-of-unknown function (DUF3426, residues 128-245) in the C-terminal region that is predicted to reside in the periplasm ( Figure 1C). The cpaM gene codes for a 394-residue protein harbouring a single N-terminal TM domain and a C-terminal CE4_DAC2-like polysaccharide deacetylase domain predicted to be periplasmic ( Figure 1C). ZitP and CpaM are not restricted to the Caulobacter lineage as BLASTP searches revealed orthologs in many a-proteobacterial clades ( Figure 1D). To confirm the phenoytpes of the Tn insertion mutants, we engineered strains with an in-frame deletion in zitP (DzitP) or cpaM (DcpaM) and found that the mutants no longer supported plaque formation (lysis) by the pilus-specific bacteriophage jCbK. By contrast, plaques were still formed by the S-layer specific caulophage jCr30 (Edwards and Smit, 1991) (Figure 2A), showing that mutations in cpaM or zitP prevent infection of jCbK, but not all phages. This defect was corrected upon expression of either ZitP or CpaM from an ectopic locus in DzitP or DcpaM cells, respectively ( Figure 2A).
Next, we conducted time-course adsorption assays and found the adsorption kinetics of DzitP and DcpaM cells to be substantially compromised compared to WT cells ( Figure 2B). The jCbK adsorption kinetics of the mutants closely resemble those for DcpaC cells that cannot assemble pili because they lack the CpaC secretin (Skerker and Shapiro, 2000). Moreover, immunoblotting revealed that DzitP and DcpaM cells do not accumulate the modified form of CpaC, CpaC* ( Figure 2C-D). A comparable reduction in CpaC* abundance has been previously reported for DcpaE, DpodJ and DpleA cells that no longer assemble a polar CpaC pilus channel in the outer membrane and cannot be infected by jCbK (Viollier and Shapiro, 2003;Viollier et al., 2002b). However, CpaC* accumulates in DpilA cells ( Figure 2C), suggesting that the CpaC channel forms independently of PilA. To test whether DzitP and DcpaM cells assemble a pilus filament on the cell surface, we conducted shearing assays followed by immunoblotting using antibodies to the PilA pilin, the subunit of the pilus filament ( Figure 2E) (Skerker and Shapiro, 2000). Whereas PilA was efficiently released from WT cells into the supernatant by shearing, no PilA was detectable in the  Figure 2E), even though PilA is clearly expressed in these cells ( Figure 2C). As the major subunit of the flagellar filament, the FljK flagellin, accumulates in the supernatants in all samples ( Figure 2E), we conclude that ZitP and CpaM are required for the presentation of PilA on the cell surface and, as shown below, that they act in the same pathway ( Figure 1B).
Control of motility, G1-phase and the CtrA regulon.
The jCbK adsorption kinetics hinted that motility might be altered in DzitP and DcpaM cells. This hypothesis is based on the comparison of the jCbK adsorption kinetics to WT, DpilA and Dfljx6 (lacking all six flagellin genes: fljJ/K/L/M/N/O) cells to DzitP and DcpaM cells. While pililess DpilA cells assemble a flagellum and are motile ( Figure 2F), Dfljx6 cells are flagellumless, but piliated (jCbK sensitive) (Guerrero-Ferreira et al., 2011). The kinetics of adsorption of jCbK to DzitP and DcpaM cells was strongly reduced compared to WT, fitting halfway between the adsorption curves of jCbK to DpilA and Dfljx6 cells ( Figure 2B). Since it is known that jCbK first reversibly adsorbs to the flagellar filament rotating counter-clockwise, before the irreversibly attachment to the pilus portal is established for productive infection (Guerrero-Ferreira et al., 2011), we wondered whether there are fewer motile cells in the DzitP and DcpaM populations than in WT or if motility in these mutants is altered in other ways. In fact, motility tests on swarm (0.3%) agar revealed a mild reduction in motility of DcpaM cells and a severe reduction of DzitP cells compared to WT ( Figure 2F). However, DzitP cells still have residual motility that allows them to spread in swarm agar compared to Dfljx6 cells ( Figure 2F). Expression of Dendra2-ZitP from an ectopic locus confers near WT motility to DzitP cells ( Figure 2G), showing that this deficiency in motility is indeed due to the absence of ZitP.
As Caulobacter divides into a motile G1-phase cell and a sessile S-phase cell, mutants accumulating fewer G1-phase cells in the population can exhibit reduced motility on soft agar . To test if ZitP controls the G1 cell number, we used flow cytometry to quantify the number of G1 cells and indeed observed fewer G1 cells in the DzitP population compared to WT ( Figure 2H). Knowing that the master cell cycle transcriptional regulator CtrA retains cells in G1-phase and activates many cell cycle-regulated promoters that fire in G1-phase (Domian et al., 1997;Fumeaux et al., 2014;Quon et al., 1996), we then conducted promoter-probe assays using several CtrA-activated promoters fused to the promoterless lacZ gene and quantified CtrA-dependent promoter activity in WT and DzitP cells (Figure 2-figure supplement 1). While all such promoter-probe reporters for the CtrA regulon exhibited a decrease in activity by 30-40% in DzitP versus WT cells, promoter-probe reporters for the GcrA regulon or other reporters were unaffected. Thus, ZitP is required for optimal CtrA activity and G1 cell accumulation.
The reduction in CtrA-dependent transcription does not appear to be solely responsible for the motility defect of DzitP cells. First, promoter-probe assays revealed that DcpaM cells also suffer from reduced CtrA-dependent activation (  exceeds that of DzitP cells ( Figure 2F). Second, we were able to mitigate the defect in CtrA-dependent transcription by ectopic expression of the (p)ppGpp alarmone, a signalling molecule that enhances CtrA function and stability via a poorly understood mechanism (Gonzalez and Collier, 2014). We accomplished this by heterologously expressing the truncated version of the E. coli (p) ppGpp-synthase RelA (RelA') from the xylose-inducible promoter at the xylX locus in WT and DzitP cells. LacZ-based promoter-probe assays revealed that ectopic induction of (p)ppGpp restores CtrAdependent promoter activity to near WT levels (Figure 2-figure supplement 2B). However, the motility of DzitP cells ectopically producing (p)ppGpp is still substantially lower than that of WT cells ( Figure 2-figure supplement 2C-D), indicating that ZitP also promotes motility through a CtrAand (p)ppGpp-independent pathway.
To reinforce this conclusion, we isolated a spontaneous motile suppressor of DzitP cells (see Materials and Methods, Figure 2I) with a single point mutation in the fliG flagellar gene (fliG D306G ) that neither corrects the pilus assembly defect (jCbK-resistance, Figure 2A), nor the reduction in G1 cell number of the DzitP mutant ( Figure 2H). As FliG encodes a component of the flagellar motor that is associated with the cytoplasmic membrane (Macnab, 2003), we conclude that ZitP controls pilus biogenesis and a multifactorial motility phenotype, with a minor contribution from a CtrAdependent pathway and a major one from a CtrA-independent pathway(s) that can be bypassed by a mutant variant of FliG.

Distinct polar ZitP assemblies control CpaM localization
To investigate if ZitP also controls its polar functions from the cell pole, we resorted to live-cell fluorescence imaging by epifluorescence microscopy (

Localization and functional determinants in ZitP
To identify the determinants within ZitP governing the differential polar localization and to test if they support specific functions, we first constructed a mutant variant of ZitP in which all four zinccoordinating cysteine residues in the zinc-finger domain (Bergé et al., 2016) are replaced by serine residues (henceforth ZitP CS , Figure 1C). The motility of DzitP cells expressing ZitP CS or Dendra2-ZitP CS is reduced compared to those expressing the WT version of ZitP (ZitP or Dendra2-ZitP; Figure 2G and K). While Dendra2-ZitP CS exclusively localizes to the pole opposite the stalk in DzitP cells ( Figure  By contrast, the opposite effect was seen when ZitP 1-133 , a ZitP variant that lacks the periplasmic DUF3426 but retains the cytoplasmic and TM domains (residues 1-133, Figure 1C), is expressed in DzitP cells. ZitP 1-133 supports efficient motility and is polarly localized, but no longer supports pilus function (i.e. plaque formation by jCbK), CpaM localization and efficient CtrA-activated transcription ( Figure 2G and J, Figure 3-figure supplement 8A-B). Thus, the periplasmic DUF3426 plays a critical role in promoting pilus assembly through the polar recruitment of CpaM. Support for the notion that DUF3426 function is regulated from sequences N-terminal to the DUF3426 came from a forward genetic screen (see Materials and Methods) that led to the identification of ZitP GAP ( Figure 1C), a mutant variant in which residues Arg93 and Ala94 preceding the TM domain are deleted. ZitP GAP supports motility ( Figure 2K), but neither plaque formation by jCbK, nor CpaC* production (Figure 2A and D). As ZitP GAP still localizes to the cell poles, interacts with CpaM and recruits Dendra2-CpaM (Figure 3-figure supplements 7A-D and 8C), ZitP also acts on pilus biogenesis independently of CpaM localization.
Taken together our experiments indicate that function and localization of ZitP can be genetically uncoupled. The periplasmic DUF3426 region is required for pilus biogenesis and CtrA-dependent transcription and it implements these functions via the recruitment of CpaM to the pole opposite the stalk. The zinc_ribbon_5 domain promotes PopZ-dependent localization of ZitP to the stalked pole and efficient swarming motility by an unknown mechanism. Interestingly, in a related study, we recently found that ZitP controls PopZ localization independently of the DUF3426 (Bergé et al., 2016).

Cell cycle control of ZitP and CpaM assemblies
Synchronization studies and genetic experiments with cell cycle mutants showed that ZitP and CpaM polarization is temporally and functionally coordinated with cell cycle progression. Immunoblotting revealed the steady-state levels of ZitP and CpaM to fluctuate during the cell cycle ( Figure 4A), exhibiting a trough during the G1fiS transition and concomitant loss of polar fluorescence at this time ( Figure 4B-C). Consistent with the genetic and cytological hierarchy, ChIP-Seq data shows that the early S-phase regulator GcrA directly promotes ZitP and CtrA expression, while the late S-phase regulator CtrA activates expression of CpaM (Fiebig et al., 2014;Fioravanti et al., 2013;Fumeaux et al., 2014;Murray et al., 2013). Moreover, ZitP, CtrA and CpaM abundance is reduced when GcrA is depleted or inactivated ( Figure 4D). ZitP expression is also strongly reduced in the absence of the CcrM adenine methyltransferase that methylates adenines at the N6-position in the context of 5'-GANTC-3' sequences. GANTC methylation is required for efficient recruitment of GcrA to its target promoters (Fioravanti et al., 2013;Murray et al., 2013).
Additionally, we found that the DivJ-PleC-DivK (kinase-phosphatase-substrate) system that regulates cell cycle progression and polar development influences the appearance of polar Dendra2-ZitP and Dendra2-CpaM ( Figure 3F

Discussion
The pole-specific and distinctly shaped assemblies of ZitP are governed via independent localization pathways and linked with functional specialization ( Figure 4E). While ZitP acts on pilus assembly by recruiting CpaM and, subsequently, the CpaC pilus channel to the pole opposite the stalk (1B and 4E), CpaM is also required for efficient activation of CtrA-dependent promoters by an unknown mechanism. A similar reduction in CtrA-dependent transcription occurs in DzitP cells that are unable to localize CpaM. While diminished CtrA activity can undermine motility by reducing the number of motile G1-phase cells in the population , ZitP affects motility in another way, since DzitP cells are diminished in motility compared to DcpaM cells. Moreover, ectopic induction of the alarmone (p)ppGpp reinforces CtrA abundance and activity (Boutte et al., 2012;Gonzalez and Collier, 2014;Lesley and Shapiro, 2008;Ronneau et al., 2016;, but only modestly improves the motility of DzitP cells. Such a motility defect also manifests when ZitP CS , a variant that no longer localizes to the stalked pole, is expressed in DzitP cells. How ZitP promotes swarming motility from the stalked pole is unclear, but there is precedence of other regulators (SpmX/Y and CpdR) that localize exclusively to the stalked pole and affect Caulobacter motility indirectly by regulating cell cycle factors (Janakiraman et al., 2016;McGrath et al., 2006;Radhakrishnan et al., 2008). Moreover, SpmX and CpdR interact with PopZ directly and their localization is compromised in the absence of PopZ (Bowman et al., 2010;Holmes et al., 2016). It is therefore conceivable that ZitP also affects motility indirectly from the stalked pole, possibly via cell cycle regulation, flagellar performance and/or polarity. The fact that the motility defect of DzitP cells can be restored by compensatory mutations in a switch component (FliG) of the flagellar motor (Kojima and Blair, 2004), suggests that flagellar performance, reversals or timing (i.e. the length of flagellation in the cell cycle) could be altered by the DzitP mutation.
Zinc-finger domain proteins other than ZitP may be implicated in linking motility and polarity. The gliding motility protein AgmX confers a flagellum-and pilus-independent form of surface motility in Myxococcus xanthus (Nan et al., 2010), a d-proteobacterium that periodically reverses the polarity of movement. Since AgmX also harbors a related N-terminal Zinc-finger domain, at least two related zinc-finger domains control different types of motility. This is intriguing and hints at a potentially important and conserved role of such zinc-finger domain proteins in developmental processes that rely on protein polarization in bacteria and polar matrix proteins such as PopZ to interact with them. In a complementary study, we additionally show in vitro and in vivo that zinc-bound ZitP binds PopZ directly and regulates PopZ localization without the periplasmic DUF3426 domain (Bergé et al., 2016), suggesting that this activity in ZitP may underlie the aforementioned CtrA-independent role in motility.
The conservation of ZitP, CpaM ( Figure 1D) and PopZ orthologs (Bowman et al., 2010) in distant a-proteobacterial lineages that reside in different ecological niches hints that the functions that these proteins control are not unique to the Caulobacter branch. Indeed, we describe an interaction between ZitP and PopZ in several distinct a-proteobacterial lineages (Bergé et al., 2016). On a more general scale, our work suggests that pole-specific functions conferred by bipolar regulators may be commonly used in bacteria and possibly eukaryotes. Such mechanisms could be relevant for toggle proteins, moonlighting/trigger enzymes (Commichau and Stulke, 2015) and other bifunctional regulators (Radhakrishnan and Viollier, 2012) that have more than one biochemical activity and function, for example kinase-phosphatases or synthase-hydrolases of cyclic-di-GMP sequestered to both cell poles (Boyd, 2000;Kazmierczak et al., 2006;Tsokos and Laub, 2012).
In sum, the functional and topological versatility of ZitP illustrates how a conserved regulator is used to coordinate multiple functions from different locations and structures in the same cell, relying on distinct protein domains and partners to control localization or to implement function. As these functions and polar remodelling events are coordinated with cell cycle progression in Caulobacter via conserved cell cycle proteins, it is likely that superimposed temporal layers similarly act on ZitP and CpaM orthologs in other a-proteobacterial cell cycles.

Bacterial strains, plasmids, and oligonucleotides
Bacterial strains, plasmids, and oligonucleotides used in this study are listed and described in supplementary tables.
b-Galactosidase assays b-Galactosidase assays were performed at 30˚C as described previously (Huitema et al., 2006;Viollier and Shapiro, 2003). Experimental values represent the averages (standard error of the mean, SEM) of at least three independent experiments.

PALM imaging conditions
To image C. crescentus, overnight cultures were diluted in fresh PYE, xylose was added (0.3% final concentration), and the cells were grown for 3 hours to mid-exponential phase (OD (660)~0.4). Two uL of culture was placed on a agarose pad containing PYE. The agarose pad was mounted in a silicone gasket (Grace Biolabs 103280) sandwiched between two microscope coverslips to minimize shrinkage of the agarose. The temperature of the microscope enclosure during experiments was 24˚C. Images were acquired using a previously described custom built PALM microscope (Holden et al., 2014). Fluorescent proteins were excited at 560 nm, and photoactivation was induced at 405 nm at~0-16 W/cm 2 . For PALM images of Dendra2-ZitP in C. crescentus, cells were imaged at an exposure time of 10 milliseconds for 10,000 frames, and an excitation intensity of~4 kW/cm 2 . For stroboscopic single particle tracking PALM measurement of ZitP binding time, cells were imaged at an exposure time of 30 milliseconds, with a variable interval between each frame, at an excitation intensity of~1 kW/cm 2 . PALM localizations were accumulated in a 2D histogram; the resulting image was blurred with a 2D Gaussian of radius 15 nm to reflect the localization uncertainty of the measurement. The image was gamma adjusted to 0.5 to compensate for the large dynamic range of the image, and the 'Red Hot' ImageJ colormap was applied.

Measurement of ZitP binding time by PALM
Binding time, t off, of ZitP to the C. crescentus poles was determined via stroboscopic single particle tracking PALM (Gebhardt et al., 2013;Manley et al., 2008). Under these conditions, Dendra2 bleached under continuous illumination with a photobleaching lifetime, t b , on the order of 50 milliseconds. Since rapid diffusion means that Dendra2-ZitP is only visible when bound to the membrane, and since photobleaching will shorten the observed binding time, the effective on-time of a single Dendra2-ZitP molecule, t eff , will be the convolution of the photobleaching lifetime, t b , and the binding lifetime t off , Effective on-time was measured by combining individual Dendra2-ZitP localizations in adjacent frames into tracks (Crocker and Grier, 1996), and fitting a single exponential model to the observed the track length distribution (Figure 3-figure supplement 6A). In order to measure binding times longer than the photobleaching lifetime, the photobleaching lifetime of the fluorescent protein may be artificially extended by using stroboscopic illumination, introducing large gaps between short periods of illumination. This increases the effective bleaching lifetime to: where t tl is duration of the gap (time lapse/strobe interval), t int is camera integration time. By measuring the effective on-time for multiple different stroboscopic illumination times, t tl , and performing a fit of: to the data, both the binding time and photobleaching lifetime may be calculated (Gebhardt et al., 2013) (Figure 3-figure supplement 6B and C Model 1). We performed non-linear least squares fitting of the raw t eff data directly to Eq. 3, instead of calculating the quantity t tl /t eff and performing a linear fit as proposed by Gebhardt and coworkers (Gebhardt et al., 2013), since the inverse transform proposed results in a non-linear transformation of the sample error distribution incompatible with least squares fitting. We observed that for stroboscopic illumination times significantly greater than the binding time, the data appeared to transition from the hyperbolic relationship predicted by Eq. 3 to a zero-gradient plateau (Figure 3-figure supplement 6B), giving very poor fits between Eq 3 and the data, especially for the DpopZ strain which appeared to have a shorter Dendra2-ZitP binding lifetime (Figure 3-figure supplement 6B). We hypothesized that this was due to an inability to accurately estimate effective on-time when molecules bind and unbind in a time significantly less than the duration of a single strobing interval (since the observed track length will almost always equal 1 frame). We confirmed this hypothesis by performing the stroboscopic tracking analysis on simulated data (Figure 3-figure supplement 6C). We simulated timetraces of molecules binding/ unbinding with finite bleaching lifetimes, and measured the observed on-time for each simulated molecule by fitting a single exponential to the on-time histogram as above. We observed as hypothesized that the observed off-times showed a sharp plateau for long-strobe intervals due to the finite integration time of the measurement, giving a poor fit of Eq 3 to the data (Figure 3-figure supplement 6C). In order to correct for this, we modified the fitting model to include a minimum measurable on-time plateau: Use of the modified model allowed us to obtain accurate fits to the entire simulated dataset (Figure 3-figure supplement 6C; Eq 4).
We therefore used our updated model to fit the experimental data ( Figure 3E and Figure 3-figure supplement 6B) and to calculate the observed binding times (Figure 3-figure supplement  6D). This gave a much better fit to the data, both at late and early strobe intervals. Notably, independent fits to the WT and DpopZ datasets gave similar observed t min tl of~0.4 frames, supporting the use of the updated model.

Measurement of ZitP cluster area and shape by PALM
In order to estimate the area of Dendra2-ZitP polar complexes, observed localizations were clustered based on local density using DBSCAN (Endesfelder et al., 2013;Ester et al., 1996). Identified clusters were converted to PALM images binarized, and morphologically closed (Figure 3figure supplement 5Bi-iii). By performing morphological closing on the binary image, we obtained segmented clusters (Figure 3-figure supplement 5Biii) which were less sensitive to noise and better reflected the visually estimated extent of the non-segmented cluster. For each identified cluster, the area of the segmented cluster was calculated.
For the NA1000 xylX::P xyl -dendra2-zitP strain, clusters were visually identified as belonging to the stalked or flagellar poles based on the PALM and phase contrast images of the region. For the DpopZ::W xylX::P xyl -dendra2-zitP strain, there was no clear difference in pole morphology, so the cluster area for cells was calculated without discriminating poles. Measurement noise means that the measured area of even a zero-area cluster will be larger than zero (and approximately proportional to the localization uncertainty). To test whether Dendra2-ZitP formed an extended polar complex, we compared the area of ZitP clusters to the measured area of simulated zero-area clusters by generating simulated datasets containing localizations coming from a point source, with photon count, background noise and total number of localizations equal to the median values of either the WT or DpopZ::W datasets (Figure 3-figure supplement 5D). The cluster area of the simulated datasets was then calculated as above.
We also calculated the following shape based metrics to further quantify the differences in pole shape: circularity, solidity and eccentricity (Figure 3-figure supplement 5C).
Circularity measures similarity of a shape to a circle, C ¼ 4pA p 2 ; where A is shape area and p is perimeter. Solidity measures the extent to which a shape is convex or concave, S ¼ A H where A is shape area and H is the convex hull area of the shape. Eccentricity measures how elongated a shape is, E ¼ a b where a is the length of the minor axis and b is the length of the major axis. Since the observed distributions showed significant non-normality, statistical significance was assessed by the non-parametric test, Mood's median test. Stars on Figure 3D and Figure 3-figure supplement 5C indicate statistical significance: n.s, p>0.05; *p<0.05; **p<0.01; ***p<0.001.
The stalked and the other (swamer) pole foci in WT showed statistically significant differences (p<0.001) in area, circularity and solidity, supporting the conclusion that ZitP forms distinct polar assemblies.
The WT stalked pole showed statistically significant differences (p<0.001) to the DpopZ::W mutant foci for area, circularity, solidity and eccentricity, supporting the conclusion that PopZ specifically promotes the formation of large polar assemblies at the stalked pole.

Isolation of 'CbK resistant mutants
A himar1-based transposon mutagenesis of the NA1000 (wild-type, WT) strain was done using the E. coli S17-1 lpir strain containing the himar1-delivery plasmid pHPV414 . The mutant library was selected on plates containing nalidixic acid and kanamycin embedded in top agar containing jCbK. Colonies emerging from this selection were pooled. We then created generalized transducing lysate from this pool using phage jCr30 and transduced it into strain PV14 DpilA-cpaF::Waac3 (conferring resistance to aparamycin), selecting for apramycin and kanamycin resistant transductants to eliminate himar1 insertions in the pilA-cpaF locus. The transductants were pooled and a generalized transducing lysate was prepared from this pool using jCr30. This new lysate was then used to transduce NA1000 to kanamycin resistance and the resulting clones were individually tested for resistance to jCbK. The himar1 insertion site mapping of jCbK-resistant himar1 mutants was done as described before .
To isolate the zitP GAP mutation, we generated a mutant library of zitP alleles by electroporating pMT335-zitP into the mutator E. coli XL1-Red strain. We collected and pooled over 20,000 clones for plasmid extraction and we electroporated the plasmid library into the DzitP mutant. We incubated the electroporated cells during two hours for regeneration and next added jCbK for one hour in order to eradicate clones that bear a mutated zitP allele restoring effective phage infection. Finally, we plated cells on soft (0.3% swarming) agar to evaluate the motility properties. We picked and streaked out motile clones for amplification and plasmid extraction and introduced the plasmids back into a DzitP background in the perspective to confirm the motility-proficient and jCbK resistant phenotypes. We isolated a unique plasmid, pMT335-zitP GAP , which bears the zitP GAP allele (deletion of the Arg93 and Ala94 in the ZitP protein).

Epi-fluorescence microscopy
PYE or M2G cultivated cells in exponential growth phase were immobilized using a thin layer of 1% agarose. Fluorescence and DIC images were taken with an Alpha Plan-Apochromatic 100x/1.46 DIC (UV) VIS-IR oil objective on an Axio Imager M2 microscope (Zeiss) with 488 nm laser (Visitron Systems GmbH, Puchheim, Germany) and a CoolSnap HQ (Boutte et al., 2012) camera (Photometrics) controlled through Metamorph V7.5 (Universal Imaging). Images were processed using Image J software. Quantifications were done by manually numbering cells in the diffuse, monopolar or bipolar state.

Protein purification and production of antibodies
The PCR-amplified zitP Cterm and cpaM DTM genes were cloned into the pET28a vector (Novagen). The His 6 -ZitP Cterm and His 6 -CpaM DTM recombinant proteins were overexpressed in E. coli strain Rosetta and purified in standard native conditions on Ni 2+ -NTA agarose as described previously to raise rabbit polyclonal IgGs in New Zealand White rabbits (Josman LLC, Napa, CA).

Tandem affinity purification (TAP) and mass spectrometry
We followed the TAP procedure as was previously described (Puig et al., 2001). When a 1 L-culture reached OD660 between 0.4 and 0.6 in the presence of 50 mM vanillate, cells were harvested by centrifugation (6000xg, 10 min). We washed the pellet in 50 mL of buffer I (50mM sodium phosphate pH 7.4, 50 mM NaCl, 1 mM EDTA) and lysed for 15 minutes at room temperature in 10 mL of buffer II (buffer I + 0.5% n-dodecyl-b-D-maltoside, 10mM MgCl 2 , two protease inhibitor tablets [Complete EDTA-free, Roche] per 50 mL of buffer II, 1x Ready-Lyse lysozyme [Epicentre], 500U of DNase I [Roche]). Cellular debris was removed by centrifugation (7000xg, 20 minutes, 4˚C). The supernatant was incubated for 2 hours at 4˚C with IgG Sepharose beads (GE Healthcare Biosciences) that had been washed once with IPP150 buffer (10 mM Tris-HCl pH 8, 150 mM NaCl, 0.1% NP40). After incubation, the beads were washed at 4˚C three times with 10 mL of IPP150 buffer and once with 10 mL of TEV cleavage buffer (10 mM Tris-HCl pH 8, 150 mM NaCl, 0.1% NP40, 0.5 mM EDTA, 1 mM DTT). The beads were then incubated overnight at 4˚C with 1 mL of TEV solution (TEV cleavage buffer with 100 U of TEV protease per ml [Promega]) to release the tagged complex. 3 mM CaCl 2 was then added to the solution. The sample with 3 mL of calmodulin-binding buffer (10 mM b-mercaptoethanol, 10 mM Tris-HCl pH 8, 150 mM NaCl, 1 mM magnesium acetate, 1 mM imidazole, 2 mM CaCl 2 , 0.1% NP40) was incubated for 1 hour at 4˚C with calmodulin beads (GE Healthcare Biosciences) that previously had been washed once with calmodulin-binding buffer. After incubation, the beads were washed three times with 10 mL of calmodulin-binding buffer and eluted five times with 200 mL IPP150 calmodulin elution buffer (calmodulin-binding buffer substituted with 2 mM EGTA instead of CaCl 2 ). Amicon Ultra-4 spin columns (Ambion) were used to concentrate eluates. Eluates were analyzed by SDS-PAGE and stained with silver using the Biorad Silver Stain Plus kit (Biorad, USA). We then cut specific bands and directly sent the gel slices to the Taplin Biological Mass Spectrometry Facility (Harvard Medical School, Boston, USA) for mass spectrometric analyses.

Co-immunoprecipitation
Cells were harvested from a 50 mL-culture (OD (660 nm) between 0.4-0.6) by centrifugation at 5000xg for 10 minutes. We washed the cell pellet in 10 mL of buffer I (50mM Tris-HCl (pH 7.5); 50 mM NaCl; 1mM EDTA), centrifuged the cell again and resuspended in 1 mL of buffer II (buffer I plus 0.5% n-dodecy-b-D-maltoside; 10 mM MgCl 2 ; EDTA-free protease inhibitors). We incubated the mixture for 15 minutes with 5000 units of Ready-Lyse lysozyme (Epicentre), and 30 units of DNase I (Roche). Cellular debris were removed by centrifugation at 10,000xg for 3 minutes at 4˚C. We cleared the supernatant by incubation for 1 hour at 4˚C with Protein-A agarose beads (Roche) previously washed three times with 500 mL of buffer II. We removed agarose beads by centrifugation and added to the pre-cleared solution polyclonal IgG rabbit serum for 90 min at 4˚C (dilution 1:500). Next, we trapped for 1 hour at 4˚C the antibodies-proteins complexes with the addition of Protein-A agarose beads (Roche) previously washed three times with 500 mL of buffer II. The samples were then centrifuged at 3000xg for 1 minute at 4˚C and the supernatant was removed. The beads were washed once with buffer I plus 0.5% n-dodecy-b-D-maltoside, three times with 500 mL of wash buffer (10 mM Tris-HCl at pH 7.5; 150 mM NaCl; 0.1% n-dodecy-b-D-maltoside) and finally resuspended in 70 ml SDS sample buffer (50 mM Tris-HCl at pH 6.8), 2% SDS, 10% glycerol, 1% b-mercaptoethanol, 12.5 mM EDTA, 0.02% Bromophenol Blue), heated to 95˚C for 10 minutes and stored at À20˚C.

Motility assays and phage infectivity tests
Swarming properties were assessed with 5 ml-drops of overnight culture spotted on PYE soft agar plates (0.3% agar) and grown for 24 hours. Phage susceptibility assays were conducted by mixing 500 mL of overnight culture in 6 mL soft PYE agar and overlaid on a PYE agar plate. Upon solidification of the soft (top) agar, we spotted 5 mL-drops of serial dilution of phages (jCbK or jCr30) and scored for plaques after one day incubation at 4˚C.

Shearing experiments
We centrifuged 5 mL mid-log phase cultures of WT or mutant strains and resuspended them in 700 ml of PYE. Then, we pumped in and out (10x) the cells into a syringe endowed with a thin diameter needle. We centrifuged the shear-stressed cells to remove cells debris and collected 200 mL of each supernatant. We added SDS-blue straining and loaded samples on SDS-PAGE gels.

'CbK adsorption assay
To determine the adsorption rate of jCbK, Caulobacter crescentus NA1000 and derivatives were first grown overnight in M2G medium at 30˚C and then re-started in fresh M2G at 30˚C with shaking until the bacterial cell culture reached an OD660 value of 0.4 (0.4 Â 10 8 CFU/ml). Then cell cultures were infected by 0.02 multiplicity of jCbK infection (MOI: ratio of phage to bacteria number). The mixtures were incubated at 30˚C without shaking for phage adsorption, followed by separation of unbound phages by centrifugation at 13,000 rpm in specified time points up to 30 minutes. Supernatants were immediately supplemented by the addition of chloroform (1/20 of cell culture volume) and mixed vigorously to kill remaining bacterial cells. A control tube containing only jCbK (equivalent to 0.02 MOI) was maintained in parallel for the duration of the experiment and used as reference to control the initial phage plaque-forming units (pfu) titer. A 50 mL of the phage supernatant from each tube was mixed with 200 mL of Caulobacter crescentus NA1000 culture at log phase and incubated without shaking at room temperature for 10 minutes to allow adsorption. Infected cells were added to 6 mL of soft PYE agar (0.5%) and immediately overlaid on 1.5% PYE agar plates. Plates were incubated at 30˚C for 24 hours, when pfu were visible. The jCbK adsorption value (in% of the initial phage pfu titer) was calculated. Values are the mean of three biological replicates; error bars represent data ranges.
Flow cytometry (Fluorescence-activated cell sorting, FACS) Cells in exponential growth phase (OD660nm=0.3-0.6) cultivated in PYE, were fixed in ice-cold 77% ethanol solution. Fixed cells were re-suspended in FACS staining buffer, pH 7.2 (10 mM Tris-HCl, 1 mM EDTA, 50 mM NaCitrate, 0.01% Triton X-100) and then treated with RNase A (Roche) at 0.1 mg mL À1 for 30 minutes at room temperature. Cells were stained in FACS staining buffer containing 0.5 mM of SYTOX Green nucleic acid stain solution (Invitrogen) and then analysed using a BD Accuri C6 flow cytometer instrument (BD Biosciences). Flow cytometry data were acquired and analysed using the CFlow Plus V1.0.264.15 software (Accuri Cytometers Inc.). 20,000 cells were analysed from each biological sample. Experimental values represent the averages of three independent experiments. fliG D306G swarming pseudo-revertant isolation and backcrossing We spotted several 5 mL-drops of DzitP overnight culture on soft agar plates and waited for flares spreading out the bulk of cells. Flares were peaked out and streaked on fresh agar plates for amplification and subsequently challenged for motility in comparison to WT and DzitP strains. Motility-proficient clones were sent for Illumina HiSeQ 2000 sequencing (Fasteris, www.fasteris.com/). Genomes were compared to NA1000 genome and we identified a single mutation in the fliG gene (D306G).
In order to backcross the fliG D306G allele in different backgrounds, the suppressor strain was electrotransformed with the suicide vector pNTPS138-hook and selected on kanamycin-supplemented plates for single crossing-over in close vicinity of the fliG locus. We prepared lysate of this strain, transduced the fliG D306G -linked pNTPS138 into WT and DzitP cells and screen by sequencing for clones harbouring the fliG D306G allele. Finally, we grew up the strain without any antibiotic and selected for plasmid excision by plating an overnight culture on sucrose.