Cell cycle coordination and regulation of bacterial chromosome segregation dynamics by polarly localized proteins
2010; Springer Nature; Volume: 29; Issue: 18 Linguagem: Inglês
10.1038/emboj.2010.207
ISSN1460-2075
AutoresWhitman B. Schofield, Hoong Chuin Lim, Christine Jacobs‐Wagner,
Tópico(s)Escherichia coli research studies
ResumoArticle27 August 2010free access Cell cycle coordination and regulation of bacterial chromosome segregation dynamics by polarly localized proteins Whitman B Schofield Whitman B Schofield Department of Molecular, Cellular and Developmental Biology, Yale University, New Haven, CT, USA Search for more papers by this author Hoong Chuin Lim Hoong Chuin Lim Department of Molecular Biophysics and Biochemistry, Yale University, New Haven, CT, USA Search for more papers by this author Christine Jacobs-Wagner Corresponding Author Christine Jacobs-Wagner Department of Molecular, Cellular and Developmental Biology, Yale University, New Haven, CT, USA Howard Hughes Medical Institute, Yale University, New Haven, CT, USA Microbial Pathogenesis section, Yale Medical School, New Haven, CT, USA Search for more papers by this author Whitman B Schofield Whitman B Schofield Department of Molecular, Cellular and Developmental Biology, Yale University, New Haven, CT, USA Search for more papers by this author Hoong Chuin Lim Hoong Chuin Lim Department of Molecular Biophysics and Biochemistry, Yale University, New Haven, CT, USA Search for more papers by this author Christine Jacobs-Wagner Corresponding Author Christine Jacobs-Wagner Department of Molecular, Cellular and Developmental Biology, Yale University, New Haven, CT, USA Howard Hughes Medical Institute, Yale University, New Haven, CT, USA Microbial Pathogenesis section, Yale Medical School, New Haven, CT, USA Search for more papers by this author Author Information Whitman B Schofield1, Hoong Chuin Lim2 and Christine Jacobs-Wagner 1,3,4 1Department of Molecular, Cellular and Developmental Biology, Yale University, New Haven, CT, USA 2Department of Molecular Biophysics and Biochemistry, Yale University, New Haven, CT, USA 3Howard Hughes Medical Institute, Yale University, New Haven, CT, USA 4Microbial Pathogenesis section, Yale Medical School, New Haven, CT, USA *Corresponding author. Department of Molecular, Cellular and Developmental Biology, Yale University, KBT 1032, PO Box 208103, New Haven, CT 06520, USA. Tel.: +1 2 03 432 5170; Fax: +2 03 432 6161; E-mail: [email protected] The EMBO Journal (2010)29:3068-3081https://doi.org/10.1038/emboj.2010.207 There is a Have you seen ...? (September 2010) associated with this Article. PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info What regulates chromosome segregation dynamics in bacteria is largely unknown. Here, we show in Caulobacter crescentus that the polarity factor TipN regulates the directional motion and overall translocation speed of the parS/ParB partition complex by interacting with ParA at the new pole. In the absence of TipN, ParA structures can regenerate behind the partition complex, leading to stalls and back-and-forth motions of parS/ParB, reminiscent of plasmid behaviour. This extrinsic regulation of the parS/ParB/ParA system directly affects not only division site selection, but also cell growth. Other mechanisms, including the pole-organizing protein PopZ, compensate for the defect in segregation regulation in ΔtipN cells. Accordingly, synthetic lethality of PopZ and TipN is caused by severe chromosome segregation and cell division defects. Our data suggest a mechanistic framework for adapting a self-organizing oscillator to create motion suitable for chromosome segregation. Introduction Similar to eukaryotes, bacteria use active mechanisms to partition chromosomes equally between daughter cells. Fluorescence microscopy studies in various bacteria have shown that specific centromere-like DNA regions close to the origin of replication are translocated to opposite sides of the cell in one single event during the cell cycle. How this controlled translocation occurs at the mechanistic level is not well understood. Most of our knowledge about DNA segregation in bacteria comes from studies on plasmids, in part because they are not necessary for cell viability. Many plasmids use the Par Type 1 system for partitioning (Gerdes et al, 2010). This system has three components, here referred to as parS, ParB and ParA. Interestingly, orthologues of these components are commonly found on bacterial chromosomes (Livny et al, 2007). Although these components can have a function in the regulation of DNA replication in some bacteria (Murray and Errington, 2008), the parS/ParB/ParA system has also been shown to be involved in chromosome segregation (Gerdes et al, 2010). Plasmid and chromosome-encoded ParB proteins are known to bind to cognate, centromere-like parS DNA sequences. ParA proteins are deviant Walker-type ATPases whose weak ATPase activity is stimulated by an interaction with ParB either alone or when bound to parS (Radnedge et al, 1998; Easter and Gober, 2002; Leonard et al, 2005; Barillà et al, 2007; Bouet et al, 2007; Pratto et al, 2008). In vitro, ParA-ATP dimers bind to DNA non-specifically and in a cooperative manner, whereas ADP-bound forms do not (Leonard et al, 2005; Pratto et al, 2008; Ringgaard et al, 2009). Recently, in vivo mechanistic insight into chromosome segregation was provided for chromosome I of Vibrio cholerae (Fogel and Waldor, 2006). Here, segregation starts at the old cell pole and occurs unidirectionally with a duplicated parSI/ParBI partition complex moving to the opposite, new pole (Fogel and Waldor, 2006). ParAI localizes in a cloud-like structure that extends from the new pole to near the parSI/ParBI focus at the old pole. Shrinking of the ParAI structure towards the new pole is accompanied with the translocation of parSI/ParBI, suggestive of a so-called 'pulling' mechanism (Fogel and Waldor, 2006). Such a mechanism has also been invoked for plasmid partitioning. For instance, ParA of plasmid pB171 forms DNA-bound structures that perpetually extend and shrink over the nucleoid and in-between plasmids (Ebersbach et al, 2006; Ringgaard et al, 2009). These continuous cycles are correlated with back-and-forth motions of plasmids, which result in time-averaged equidistribution of plasmids along the cell length. In the case of both plasmid pB171 and V. cholerae chromosome I, it has been proposed that parS/ParB interaction with the edge of ParA nucleoprotein structures stimulates ParA-ATPase activity, leading to disassembly of ParA subunits and hence retraction of the ParA structure (Fogel and Waldor, 2006; Ringgaard et al, 2009). However, the continuous rounds of ParA assembly/disassembly producing perpetual plasmid movement and the single DNA translocation event per cell cycle of bacterial chromosomes are clearly distinct. What generates this difference is not clear. Another important aspect of chromosome segregation that is difficult to address with plasmid models is its necessary temporal and spatial coordination with other cell cycle events. How this cell cycle coordination can occur is poorly understood. Some major advances have been achieved in Caulobacter crescentus, in which segregation of the parS/ParB partition complex is linked to cell division (Mohl et al, 2001) through MipZ, a cell division inhibitor that associates with parS/ParB partition complexes (Thanbichler and Shapiro, 2006). In C. crescentus, the DNA spreads throughout the cell, and as in V. cholerae, segregation is unidirectional (Jensen and Shapiro, 1999; Viollier et al, 2004). After replication of the parS locus at the old pole, one parS/ParB copy is rapidly translocated to the new pole (Viollier et al, 2004; Thanbichler and Shapiro, 2006; Toro et al, 2008). MipZ, which inhibits the cell division protein FtsZ, forms polar gradients through an association with ParB (Thanbichler and Shapiro, 2006). Hence, segregation of parS/ParB triggers the formation of a bipolar MipZ gradient, which has been proposed to result in the preferential assembly of the FtsZ cytokinetic ring at midcell in which the MipZ concentration is thought to be lowest (Thanbichler and Shapiro, 2006). Although MreB has been proposed to affect parS/ParB segregation (Gitai et al, 2005), data obtained from a mutagenized form of ParA suggests that ParA has a greater function in this process (Toro et al, 2008). The dynamics of ParA are, however, unknown in this organism. Division has also been linked to cell polarity in C. crescentus through the landmark polarity factor TipN (Lam et al, 2006). C. crescentus is a highly polarized bacterium, forming a predivisional cell with a flagellum and a stalk (a thin extension of the cell body) at opposite poles. Asymmetric division yields a flagellated 'swarmer' cell slightly shorter than its sibling, the 'stalked' daughter cell. TipN, through its localization at the new pole, ensures proper positioning of new-pole markers such as the flagellum, whereas it has no effect on the positioning of old-pole markers (such as the stalk) or on the asymmetric localization of the daughter cell fate determinant CtrA (Huitema et al, 2006; Lam et al, 2006). Surprisingly, TipN appears to affect cell division placement, as ΔtipN cells, unlike wild type (Terrana and Newton, 1975; Quardokus and Brun, 2002), often constrict closer to the old pole, generating a reversed asymmetry in daughter cell size (Lam et al, 2006). How the polarity factor TipN spatially affects cell division is puzzling given the distance between the site of division and the new-pole location of TipN. Here, we present evidence suggesting that TipN exerts its long-distance effect on cell division positioning by regulating ParA and parS/ParB segregation dynamics. Our study provides mechanistic insights into how rapid and directional DNA segregation can be achieved and regulated. Moreover, our findings suggest that extrinsic regulation of the parS/ParB/ParA system has a profound effect on several aspects of cell cycle coordination. Results and discussion TipN alters the timing and positioning of FtsZ ring formation Our study was initially motivated by the intriguing observation that unlike wild-type cells, ΔtipN cells frequently appear to divide closer to the old pole (Lam et al, 2006). To quantify the distribution of the division defect within the ΔtipN population, we measured the division ratios of constricting cells, defined by the length between the old pole and the site of cell constriction divided by the total cell length (Supplementary Figure S1A) using a DivJ–CFP fusion as an old-pole marker (Wheeler and Shapiro, 1999). Most wild-type cells had an average division ratio of 0.537 (with a standard error of the mean of ±0.001, n=1433; Supplementary Figure S1B) that is consistent with a previous electron microscopy study (Terrana and Newton, 1975). In contrast, the majority of ΔtipN cells had a division ratio of 0.468 (±0.001, n=1766; Supplementary Figure S1B). Thus, TipN clearly affects where division occurs. How C. crescentus selects its division site is not completely understood. MipZ is thought to restrict FtsZ polymerization to the region of lowest MipZ concentration (Thanbichler and Shapiro, 2006). As the bipolar MipZ gradients appear symmetric, it was suggested that FtsZ ring assembly may occur at midcell and that unequal growth rate between the sides flanking the FtsZ ring may cause the asymmetric division. An asymmetry in FtsZ ring positioning had been measured in dividing C. crescentus cells (Quardokus and Brun, 2002), but the assembly of the FtsZ ring occurs well before cell constriction is initiated (Kelly et al, 1998; Aaron et al, 2007). Therefore, it was conceivable that the FtsZ ring would form at midcell, but that its position would become progressively asymmetric over time because of unequal growth between sides. To test this idea, we quantified the temporal and spatial distribution of FtsZ by time-lapse microscopy starting with synchronized swarmer cell populations producing FtsZ–YFP. The relative position of FtsZ–YFP along the long cell axis in individual cells was plotted as a function of time after cell cycle synchronization (Figure 1A). In wild-type cells, FtsZ–YFP moved from the new pole to a central region and soon formed a ring (band) at a 0.536±0.003 position (n=190 cells), and this asymmetric position did not significantly change for the remainder of the cell cycle (Figure 1A; Supplementary Movie S1). Thus, it is the asymmetric location of the FtsZ ring, rather than an asymmetry in growth rate, that dictates the asymmetric division. Supporting this notion, in ΔtipN cells, the FtsZ ring stabilized on average at a 0.445±0.003 position (n=114 cells) relative to the long cell axis (Figure 1A; Supplementary Movie S2), which corresponds well to the average position of the division site of this mutant (Supplementary Figure S1B). Figure 1.FtsZ and MipZ dynamics in wild-type and ΔtipN cells. (A) Time-lapse microscopy of FtsZ–YFP in wild-type (MT199) and ΔtipN (CJW2563) cells after synchrony. The expression of ftsZ–yfp was induced with 0.5 mM vanillic acid 2.5 h before synchronization and imaging. Images were acquired every 1.5 min and the cells were identified using MicrobeTracker. The FtsZ–YFP signal in representative wild-type and ΔtipN cells is shown for selected time points as an overlay with the MicrobeTracker cell outline (the old pole is marked by the arrow). The graphs show the trace of the relative FtsZ–YFP position along the cell length over time. (B) Time-lapse microscopy of MipZ–mCFP and FtsZ–YFP in a wild-type background (strain CJW3455). FtsZ–YFP expression was induced with 0.5 mM vanillic acid 1 h before cell synchronization. Time-lapse results from a representative cell are shown as an overlay with MicrobeTracker cell outlines in white (the old pole is marked by the arrow.). The relative position of FtsZ–YFP (red trace) is indicated over a kymograph of the MipZ–mCFP signal profile (green) along the cell length as a function of time after synchrony. Yellow arrows show first instance of MipZ–mCFP and FtsZ–YFP colocalization at the new pole. (C) Same as (B) except in a ΔtipN background (strain CJW3612). Purple arrowheads show backwards motion of MipZ–mCFP. (D) Time-lapse recordings of MipZ–YFP in wild-type (CJW2022) and ΔtipN (CJW3366) cells after synchrony. Images were acquired every 1.5 min and the cells were identified using MicrobeTracker. Shown are profiles of the mean MipZ–YFP signal along the cell long axis of 94 wild-type cells (green) and 63 ΔtipN cells (blue) from the time point when MipZ–YFP becomes bipolar (yellow arrow in (B) and (C) to the onset of cell constriction). The arrows show the minima in MipZ–YFP intensity for each strain. Download figure Download PowerPoint Another striking difference between wild-type and ΔtipN backgrounds was that FtsZ–YFP remained at the new pole for a longer time in ΔtipN cells, and consequently, FtsZ ring formation (represented by the stabilization of FtsZ–YFP localization at an off-centre position) was considerably delayed (Figure 1A). Under our experimental conditions, FtsZ–YFP ring formation occurred on average 11.5±0.7 min after cell synchronization in wild-type cells (n=190), whereas in ΔtipN cells (n=114), FtsZ–YFP ring formation was observed at about 42.6±1.7 min after cell synchronization. Thus, the ΔtipN mutation causes a significant delay in FtsZ ring formation (Kolmogorov Smirnov (K-S) test, P<0.0001), while having no effect on the timing of cell constriction or cell separation (see below). TipN affects parS/ParB/MipZ segregation dynamics MipZ regulates the timing of FtsZ ring formation during the cell cycle (Thanbichler and Shapiro, 2006), raising the possibility that the delay in FtsZ ring formation in ΔtipN cells is due to abnormal MipZ dynamics. Kymographs made from time-lapse sequences of wild-type cells producing MipZ–mCFP (Figure 1B) displayed the expected dynamics during the cell cycle (Thanbichler and Shapiro, 2006). Before DNA replication, MipZ–mCFP localized at the old pole with the single parS/ParB partitioning complex. Following initiation of DNA replication and duplication of parS, a second MipZ–YFP signal rapidly moved to the new pole through its interaction with the segregating parS/ParB complex (Thanbichler and Shapiro, 2006). When MipZ–mCFP reached the new pole, it displaced FtsZ–YFP from the new pole (Figure 1B, yellow arrow). FtsZ–YFP then moved to its off-centre position where it formed a ring. In ΔtipN cells, it took much longer for MipZ–mCFP to reach the new pole (Figure 1C), explaining the delay in FtsZ ring assembly in these mutant cells. In addition to being slower in ΔtipN cells, the motion of MipZ–mCFP was erratic with many changes in direction. Even when MipZ–CFP reached the new pole (Figure 1C, yellow arrow), it usually temporarily moved back to the cell interior, often multiple times (Figure 1C, purple arrowheads). To examine the spatial distribution of the MipZ gradients along the cell length, we used MipZ–YFP-expressing cells and averaged their fluorescent profiles from the first time MipZ–YFP reached the new pole to the onset of cell constriction. In wild-type cells (n=94), the minimum of the MipZ–YFP signal was slightly biased towards the new pole with a mean value of 0.530±0.004 (Figure 1D, green arrow), whereas in ΔtipN cells (n=63), this minimum was shifted towards the old pole with a mean value of 0.460±0.008 (Figure 1D, blue arrow), most likely because of the common backwards motions of MipZ after FtsZ displacement from the new pole (Figure 1C, see images at time points 42 and 45 min for examples). This likely contributes to the difference in relative FtsZ–YFP ring positioning along the cell length between wild-type and ΔtipN cells. As MipZ is associated with the parS/ParB complex, we examined the segregation dynamics of this complex (Thanbichler and Shapiro, 2006). Without TipN, translocation of the parS/CFP–ParB complex from one pole to the other took longer with an average speed of 0.038±0.001 μm/min in ΔtipN cells (n=79) compared with 0.109±0.006 μm/min in wild-type cells (n=94). Translocation was also more erratic, displaying many directional changes (Supplementary Figure S2A). The origin of replication had a similarly erratic motion in ΔtipN cells (Supplementary Figure S2B), consistent with its proximity to the parS locus. DAPI staining revealed no major defect in overall DNA organization at a gross level (Supplementary Figure S2C). The MipZ segregation and FtsZ localization defects in ΔtipN cells were complemented by expression of plasmid-encoded TipN (Supplementary Figure S2D). Our data thus argue that TipN affects the timing and positioning of FtsZ ring formation by affecting the segregation of the MipZ-associated parS/ParB partition complex. TipN regulates ParA localization dynamics How can TipN, a protein that localizes at the new pole (Huitema et al, 2006; Lam et al, 2006), affect parS/ParB segregation? To address this question, we examined the spatial and temporal localization of ParA. In C. crescentus, parA is part of an operon that includes gidA, gidB, parA and parB. We generated strains in which the chromosomal parA open reading frame was substituted with a parA–yfp fusion while maintaining the native promoter, the operon structure and the upstream parS locus. Maintaining endogenous expression was important to determine the correct localization pattern as expression from heterologous promoters can lead to abnormal localization pattern as shown for the ParA homologue Soj in Bacillus subtilis (Murray and Errington, 2008). Our ParA–YFP fusion was functional (Supplementary data; Supplementary Figure S3A–C). Time-lapse microscopy and kymograph analysis of the average ParA–YFP signal over time in synchronized wild-type cells (n=65) revealed a dynamic and consistent pattern (Figure 2A). During the beginning of the cell cycle, ParA–YFP formed a 'cloud' that extended from the new pole towards the old pole. About 10–20 min into the cell cycle, the ParA–YFP cloud rapidly condensed into a focus at the new pole (Figure 2A, arrowhead) and this new-pole accumulation persisted for the rest of the cell cycle. The ParA–YFP signal was constant during the cell cycle (Supplementary Figure S3D), consistent with western blot analysis of ParA levels (Mohl and Gober, 1997). Thus, the rapid condensation of the ParA–YFP cloud observed in wild-type cells is caused by a change in protein localization as opposed to degradation. The level of ParA–YFP signal was similar in the ΔtipN background (Supplementary Figure S3D), but its localization pattern exhibited two important differences. First, the retraction of the ParA–YFP cloud was slow (Figure 2B) and the protein accumulated at the old pole (Figure 2B, pink arrowhead). Second, unlike in wild-type cells, the ParA–YFP signal failed to accumulate at the new pole in ΔtipN cells (Figure 2B). Figure 2.ParA dynamics in wild-type, ΔtipN and parA-overexpressing cells. (A) Kymograph of the average ParA–YFP signal intensity along the cell length as a function of time after synchrony. Wild-type cells (CJW3010) were imaged every 1.5 min by time-lapse microscopy and analysed using MicrobeTracker. (B) Same as (A) except that the average spatial distribution of ParA–YFP over time was obtained from imaging ΔtipN cells (CJW3011). (C) Time-lapse microscopy of ParA–YFP and CFP–ParB in a wild-type background (strain CJW3367). The results from a representative cell are shown as an overlay between the relative position of CFP–ParB (green trace) and a kymographic representation of the ParA–YFP signal profile (red) along the cell length as a function of time after synchrony. CFP–ParB synthesis was induced with 0.03% xylose for 1 h before synchronization and imaging. (D) Profiles of the mean CFP–ParB and ParA–YFP intensity profiles of wild-type cells (n=72) from time-lapse sequences described in (C). (E) Same as (C) except in a ΔtipN background (strain CJW3376). (F) Time-lapse microscopy of YFP–ParA and MipZ–mCFP in cells slightly overproducing YFP–ParA (strain CJW3373). YFP–ParA overproduction was induced with 0.03% xylose for 2 h before cell synchronization and imaging. The results from a representative cell are shown as an overlay between the relative position of MipZ–mCFP (green trace) and a kymographic representation of the ParA–YFP signal profile (red) along the cell length as a function of time after synchrony. (G) Same as (F) except that the overproduction of YFP–ParA was induced with 10 times more xylose (0.3%) for 2 h before cell synchronization and imaging. Download figure Download PowerPoint To understand how ParA dynamics may affect parS/ParB segregation, we covisualized ParA–YFP and CFP–ParB during the cell cycle. Kymograph analysis of time-lapse sequences from synchronized cell populations revealed that the condensation of the ParA–YFP cloud towards the new pole was temporally and spatially correlated with CFP–ParB segregation (Figure 2C; Supplementary Movie S3). When the duplicated CFP–ParB focus (Figure 2C, green trace) reached the ParA–YFP signal, the ParA–YFP cloud rapidly retracted to form a focus at the new pole (Figure 2C, red, 31:30 to 37:30 min; Supplementary Movie S3). Averaging the ParA–YFP and CFP–ParB signals from multiple cells showed that the translocating CFP–ParB focus follows in the wake of a retracting ParA–YFP gradient (Figure 2D). In ΔtipN cells, rapid shifts in ParA–YFP localization between poles correlated with back-and-forth movements and stalls of the segregating CFP–ParB focus (Figure 2E; Supplementary Movie S4), as if the back-and-forth motion of ParA was attracting the partitioning complex in opposite directions. Transient localizations of ParA–YFP behind the translocating CFP–ParB focus typically preceded the direction reversal of CFP–ParB towards the old pole (Figure 2E; Supplementary Movie S4). These phenotypes were complemented by producing TipN in trans (Supplementary Figure S3G). Altogether, the data suggest that the localization defect of ParA–YFP in ΔtipN cells causes the slower, erratic translocation of the parS/ParB complex. Further evidence that a change in ParA dynamics affects parS/ParB/MipZ segregation came from examining DNA segregation when ParA was slightly overproduced. Overexpression of a yfp–parA fusion from the xylose-inducible promoter (Pxyl) from a second locus had considerable effects on several aspects of ParA dynamics and localization (Figure 2F). First, YFP–ParA tended to form larger cloud structures in swarmer cells, often expanding all the way to the old pole. Second, the condensation of the YFP–ParA cloud was slower, accompanied with a correspondingly slower segregation of the parS/ParB/MipZ–mCFP complex. Third, ParA–YFP occasionally exhibited oscillations between new and old poles, which were associated with the reversal of parS/ParB/MipZ–mCFP movement (Figure 2F, 90–110 min). TipN interacts with ParA at the new pole In the absence of TipN, ParA clouds failed to condense at the new pole and new ParA structures frequently formed at the opposite end behind the partitioning complex, which was correlated with stalls or backwards movement of this complex (Figure 2E). As TipN specifically localizes at the new pole (Huitema et al, 2006; Lam et al, 2006), we hypothesized that TipN may interact with ParA at that location. Consistent with this hypothesis, a TipN-FLAG fusion (produced from the native chromosomal tipN promoter in place of wild-type TipN) was able to pull down ParA–YFP from C. crescentus cells using the anti-FLAG antibody, whereas no ParA–YFP was pulled down in the control strain producing untagged TipN (Figure 3A). We confirmed that ParA and TipN interact inside cells using fluorescence resonance energy transfer (FRET) microscopy (Verveer et al, 2006). After correcting for the bleed through from strains expressing TipN–CFP and ParA–YFP alone (Supplementary Figure S4A), we obtained a mean apparent energy transfer efficiency (nFRET/YFP) of 0.21±0.03 (n=77) between TipN–CFP and ParA–YFP (Figure 3B), indicative of an interaction. Consistent with this, TipN–CFP and mYFP-PopZ, which colocalize at the new pole, but do not physically interact (Bowman et al, 2010), displayed a mean nFRET/YFP of about zero (0.014±0.005; n=141; Figure 3B; we obtained similar results when PopZ was fused to YFP and produced from its native chromosomal locus in place of PopZ; data not shown). The difference in distributions of nFRET/YFP values for TipN/ParA and TipN/PopZ fusions were highly significant (K-S test, P<0.00001). Figure 3.Interaction between ParA and TipN. (A) Western blotting of pull-down eluates from lysates of cells carrying ParA–YFP and TipN-FLAG (TipN-FLAG; strain CJW3359) and lysates of control cells carrying ParA–YFP and untagged TipN (TipN; strain CJW3010). Elution was carried out in the presence (FLAG) or absence (Mock) of FLAG peptide. (B) Distributions of nFRET/YFP values for TipN–CFP and ParA–YFP (strain CJW3406) and for TipN–CFP and mYFP-PopZ (strain CJW3614) at the new pole. mYFP-PopZ was induced with 0.03% xylose for 2 h before imaging. The red line is the Gaussian fit of the distributions (see Supplementary data). (C) Overlays of phase-contrast images with fluorescent images of MipZ–mCFP, ParA–YFP and TipN–mCherry localization in CJW3407 cells in which TipN–mCherry overproduction was either uninduced (normal TipN) or induced with 0.5 mM vanillic acid for 6 h (TipN–mCherry overproduced). (D) Time-lapse sequence of MipZ–mCFP (green) and ParA–YFP (red) signals in CJW3408 cells during TipN overproduction, which was induced with 0.25 mM vanillic acid for 3 h before imaging on agarose pads containing vanillic acid. Download figure Download PowerPoint Overproduction of TipN can cause aberrant accumulation of TipN at the old pole and occasionally along lateral sides of the cell (Lam et al, 2006). ParA–YFP and partition complexes (visualized with MipZ–mCFP) were recruited to these abnormal TipN–mCherry localization sites (Figure 3C, arrows). Aberrant old-pole localization of ParA–YFP resulted in a slow and erratic segregation of the MipZ–mCFP-labelled partition complex characterized by frequent direction reversals and stalls (Figure 3D), similarly to what happens in ΔtipN cells (Figure 2E). Thus, collectively, our findings suggest that TipN is required for ParA recruitment to the new pole during condensation of the ParA structure, which in turn enables rapid and unidirectional segregation of the partition complex. Extrinsic regulation of the parS/ParB/ParA system We envision a model in which the ParA cloud observed before DNA replication is primarily made of DNA-bound ParA-ATP dimers forming nucleoprotein structures. The weak intrinsic ATPase activity of ParA (Easter and Gober, 2002) would maintain this steady-state until replication is initiated. The replication process itself (Lemon and Grossman, 2001; Dworkin and Losick, 2002) and/or entropy would presumably push duplicated parS/ParB complexes apart and help one of them come in contact with the edge of the nucleoprotein structure. This interaction would lead to ParB-dependent stimulation of ParA-ATPase activity and release of ParA from the nucleoprotein structure into the cytoplasm, resulting in the shrinkage of the ParA-ATP structure (see model in Figure 7A). Space-constrained Brownian motion of chromosomal loci results in some space exploration (Elmore et al, 2005; Ebersbach et al, 2008), which presumably would allow the parS/ParB complex to interact again with the edge of the slightly smaller DNA-bound ParA-ATP structure. Repetition of this process would cause the parS/ParB complex to move closer and closer to the new pole in the wake of a 'shrinking' ParA-ATP nucleoprotei
Referência(s)