MOLECULAR CHARACTERIZATION OF CELL DIVISION MACHINERY IN
CAULOBACTER CRESCENTUS
A DISSERTATION
SUMITTED TO THE DEPARTMENT OF CHEMISTRY
AND THE COMMITTEE ON GRADUATE STUDIES
OF STANFORD UNIVERSITY
IN PARTIAL FULFILLMENT OF THE REQUREMENTS
FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY
Yi-Chun Yeh
December 2010
© 2011 by Yi-Chun Yeh. All Rights Reserved. Re-distributed by Stanford University under license with the author.
This work is licensed under a Creative Commons Attribution- Noncommercial 3.0 United States License. http://creativecommons.org/licenses/by-nc/3.0/us/
This dissertation is online at: http://purl.stanford.edu/js518vh3162
ii I certify that I have read this dissertation and that, in my opinion, it is fully adequate in scope and quality as a dissertation for the degree of Doctor of Philosophy.
Harley McAdams, Primary Adviser
I certify that I have read this dissertation and that, in my opinion, it is fully adequate in scope and quality as a dissertation for the degree of Doctor of Philosophy.
William Moerner
I certify that I have read this dissertation and that, in my opinion, it is fully adequate in scope and quality as a dissertation for the degree of Doctor of Philosophy.
Lucille Shapiro
Approved for the Stanford University Committee on Graduate Studies. Patricia J. Gumport, Vice Provost Graduate Education
This signature page was generated electronically upon submission of this dissertation in electronic format. An original signed hard copy of the signature page is on file in University Archives.
iii Abstract
Cell division is a major developmental event in the life cycle of a bacterial cell.
Caulobacter crescentus division is asymmetric, producing daughter cells that differ in morphology and polar features: a sessile stalked cell and a motile swarmer cell that subsequently differentiates into a stalked cell. In this work we investigate the assembly of the Caulobacter cell division machinery (the divisome) using genetics, biochemistry, and microscopy. The bacterial divisome mediates the constriction of the cell membranes and the inward growth of the cell wall in coordination with cell elongation and chromosome segregation.
In Caulobacter , the cell division process requires a set of approximately twenty-three proteins localizing from the cytoplasm to the outer membrane. To understand divisome assembly as a function of the cell cycle, we generated fluorescent fusions to analyze the temporal regulation of 19 divisome and division-site localized proteins. In Chapter 2, we identified a series of stages and transitions in divisome assembly and the associated events yielding a comprehensive temporal picture of the process. First, the FtsZ binding proteins appear at midcell about 10 minutes after the initial assembly of Z ring near midcell. Second, proteins involved in cell growth and morphology specification localize at midcell. Next, an apparent transition to constriction occurs wherein TolQ and FtsA are recruited to the division site.
Subsequently, the arrival of five core divisome proteins to midcell is followed by the appearance of the cell polarity marker protein, TipN. The assembly interdependency for divisome formation in Caulobacter appears to involve cooperative rather than sequential recruitment, suggesting that it is a multiprotein subcomplex model.
iv Cell division in Caulobacter involves constriction and fission of the inner membrane (IM) followed about 20 min later by fission of the outer membrane (OM) and daughter cell separation. In Chapter 3, we describe our investigation of the Tol-Pal complex where we demonstrated that it plays a vital role for membrane integrity maintenance and that it is essential for viability. Cryo-electron microscope images of the Caulobacter cell envelope exhibited outer membrane disruption, and cells failed to complete cell division in TolA, TolB, or Pal mutant strains. In wild type cells, components of the Tol-Pal complex localize to the division plane in early predivisional cells and remain predominantly at the new pole of swarmer and stalked progenies upon completion of division. The Tol-Pal complex is required to maintain the position of the transmembrane TipN polar marker, and indirectly the PleC histidine kinase, at the cell pole, but it is not required for the polar maintenance of other transmembrane and membrane associated polar proteins tested. Co- immunoprecipitation experiments showed that both TolA and Pal interact directly or indirectly with TipN. We propose that disruption of the trans-envelope Tol-Pal complex releases TipN from its subcellular localization. The Caulobacter Tol-Pal complex is thus a key component of cell envelope structure and function, mediating outer membrane constriction at the final step of cell division, as well as the positioning of a protein localization factor.
FtsZ, a relative of tubulin, is the most highly conserved divisome protein. It is a GTPase that polymerizes into a contractile ring near midcell, defining the future site of cell division. Various proteins have been shown to stimulate FtsZ ring assembly while negative regulators dissociate it. In Chapter 4, we describe our examination of
v the FtsZ binding protein, ZapA. We showed that ZapA is required for proper FtsZ ring formation. Using a fluorescent fusion to the zapA gene, we found that ZapA is colocalized with FtsZ to the division plane and new pole. In addition, a ZapA- mCherry fusion is dependent on the localization of FtsZ to the division plane. ZapA is required to maintain a normal cell length, and it acts at midcell to promote Z ring assembly. A ZapA deletion strain is filamentous, showing that ZapA is required for normal divisome assembly. These biochemical and functional studies suggest that
Caulobacter ZapA is a positive regulator of Z ring assembly.
In summary, we have addressed three major stages in developments of the divisome in Caulobacter : Z ring assembly, divisome maturation and outer membrane invagination. These experiments have provided a new understanding of how the
Caulobacter cell temporally executes the cell division program to propagate reliably and how Caulobacter cell division is performed.
vi Acknowledgements
I would like to express my immense gratitude to my advisor, Harley McAdams, for all his support and suggestions over the years. Harley's guidance has been indispensable for my conversion to the biology field without any prior bench experience. I am also grateful to Lucy Shapiro for all her advice and mentoring toward research. Lucy’s enthusiasm and encouragement made the lab an enjoyable place to work. I appreciate the advice given by the other members of my thesis committee, Drs.
W. E. Moener, Robert Simoni and Andrew Spakowitz.
I am grateful to current and former members of the McAdams and Shapiro labs.
Throughout grad school, I have had the pleasure of working with many brilliant, creative, and knowledgeable scientists. I would not be the scientist I am today without the generous help of my mentors and colleagues. In particular, I would like to take this opportunity to thank Erin Goley who collaborated with me on many projects and provided most of the experimental support. The work described in this thesis would not have been possible without her. I am also thankful to Sun-Hae Hung, Martin
Thanbichler, Jay Lesley, Eduardo Abeliuk, Monica Schwartz, Paola Mara, Grant
Bowman, Nathan Hillson, Antonio Iniesta, Virginia Kalogeraki, Masaki Kato and
Yong Jae Chong for their friendship including many delightful discussions and collaborations. I would also like to thank Luis Comolli at LBNL for his help the for cryo-electron microscopy.
I also want to thank my boyfriend Yi-Ju, who has enriched my life. Finally, I want to acknowledge my sister, brother-in-law, my parents and my grandparents for their continual support.
vii Table of Contents
Chapter 1. General introduction ...... 1
1.1 The Caulobacter cell cycle ...... 2
1.2 FtsZ ...... 5
1.3 Regulation of Z-ring formation ...... 6
1.3 Overview of the late cell division proteins ...... 12
1.4 Assembly of the divisome ...... 16
1.5 Overview of other modes of bacterial cell division ...... 18
1.6 Constriction and invagination of the cell envelope ...... 20
Chapter 2. Assembly of a bacterial cell division machine ...... 22
Introduction ...... 23
Materials and methods ...... 28
Results ...... 31
Discussion ...... 57
Chapter 3. The Caulobacter Tol-Pal complex is essential for outer membrane integrity and is required for the completion of cell division and polar protein localization ...... 71
Introduction ...... 72
Materials and methods ...... 75
Resutls ...... 78
Discussion ...... 107
viii Chapter 4. ZapA-FtsZ interaction is required for Caulobacter cell division ...... 117
Introduction ...... 118
Materials and methods ...... 121
Results ...... 123
Discussion ...... 138
Chapter 5. Discussion and Future Directions ...... 141
References ...... 145
ix List of Tables
Table 1. Summary of localization dependency among Caulobacter division-site localized proteins...... 55
Table 2. Summary of features of Caulobacter divisome and division-site localized proteins ...... 56
Table 3. Strains and plamids ...... 62
Table 4. Strains and plasmids ...... 113
Table 5. Strains and plasmids ...... 140
x List of Figures
Figure 1. Caulobacter cell cycle...... 4
Figure 2. Models for division-site selection in Caulobacter (A) and E. coli (B)...... 11
Figure 3. Pathway of divisome assembly hierarchy in E. coli and B. subtilis ...... 15
Figure 4. Schematic diagram of Caulobacter divisome...... 17
Figure 5. Analysis of the dynamics of divisome assembly ...... 35
Figure 6. Time-course of division-site localized proteins using fluorescence microscopy...... 40
Figure 7. Polar localization patterns of division-site localized proteins ...... 41
Figure 8. Assembly of the divisome takes place in a series of stages ...... 43
Figure 9. The timing of midcell localization of endogenously expressed fusion proteins ...... 44
Figure 10. Pairwise comparison among division-site localized proteins ...... 51
Figure 11. Localization of fluorescent fusions in mutant cells ...... 53
Figure 12. Transcriptional regulation of genes encoding division-site localized proteins
...... 54
Figure 13. The Caulobacter crescentus tol-pal gene cluster ...... 80
Figure 14. Subcellular localization of the proteins encoded by the tol-pal cluster...... 84
Figure 15. gfp ‐tolA and pal ‐mCherry can complement deletion mutants of tolA and pal respectively...... 86
Figure 16. Mislocalization of the TipN-GFP and PleC-GFP polar proteins in the absence of the Pal or TolA proteins...... 92
Figure 17. The Tol-Pal complex interacts with TipN-GFP...... 93
xi Figure 18. Construction of Pal depletion strain...... 98
Figure 19. Phenotypes of Pal depletion strains...... 100
Figure 20. Construction of TolA and TolB depletion strains...... 101
Figure 21. Phenotype of TolA and TolB depletion strains...... 104
Figure 22. Structural relationship of the peptidoglycan layer and the IM and OM in
TolA and Pal depletion strains...... 106
Figure 23. Phenotype of ZapA depletion ...... 127
Figure 24. Relative localization of fluorescently labeled ZapA and FtsZ during the cell cycle...... 128
Figure 25. ZapA localization near midcell is dependent on FtsZ...... 134
Figure 26. Coimmunoprecipitation assay shows that ZapA interacts with FtsZ...... 135
Figure 27. ZapA counteracting MipZ to stabilize FtsZ structure...... 137
xii
Chapter 1
General introduction
1 The cell division in every cell cycle is a crucial process that allows continuous propagation of the cells, and allows the faithful transmission of hereditary information from the mother to its daughter progeny. The cell division apparatus (divisome) in bacteria mediates the constriction of the cell membranes and the inward growth of the cell wall in coordination with cell growth and chromosome segregation. In contrast to the well-characterized process of cytokinesis in eukaryotic systems, the molecular details of cell division in prokaryotic system are poorly understood.
The tubulin-like FtsZ is the best characterized and most highly conserved divisome protein (Goehring & Beckwith, 2005; Bi & Lutkenhaus, 1991). It is a
GTPase that polymerizes near the midcell (RayChaudhuri & Park, 1992), defining the site of cell division. FtsZ is the first protein to arrive at the incipient cell division site and is required for the recruitment of every other divisome protein. In addition to acting as a scaffold for assembly of the divisome, FtsZ is hypothesized to generate forces that are important for envelope invagination (Osawa et al. , 2009; Osawa et al. ,
2008). In this thesis, I focus on the characterization of ZapA, a FtsZ binding protein, that contributes to stabilize FtsZ filaments, understanding the role of the Tol-Pal complex in outer membrane invagination, and the dynamics of divisome assembly as a function of the Caulobacter crescentus cell cycle.
1.1 The Caulobacter cell cycle
We use Caulobacter as our model organism to investigate the molecular details of divisome assembly, as it offers two key advantages over the more widely studied γ- proteobacterium Escherichia coli and sporulating Bacillus subtilis. Firstly, it
2 undergoes an asymmetric cell division, where each mother cell produces two morphologically distinct daughter cells: a stalked cell and a swarmer cell that subsequently differentiates into a stalked cell. Hence, cell cycle progression can be easily monitored. Secondly, cell populations can be synchronized by differential centrifugation, which separates swarmer cells from stalked cells. This allows us to grow synchronized cell populations and to track the localization and expression level of any particular protein as cells progress through the cell cycle (Fig. 1).
During each cell cycle, Caulobacter undergoes a series of ordered events (Fig.
1, (McAdams & Shapiro, 1995; Shapiro, 1976; Laub et al. , 2000; Skerker & Laub,
2004)). It begins its life as a motile swarmer cell containing a polar flagellum, multiple polar pili, and a single chromosome (Jenal & Shapiro, 1996). For 25 to 30% of the cell cycle, the swarmer cell is not capable of DNA replication (Hung et al. , 1999). The swarmer cell then differentiates into a stalked cell, whereby the flagellum is ejected, and the pili are retracted. A tubular extension known as the stalk grows out of the cell pole previously containing the flagellum. Following the swarmer-to-stalked transition, the stalked cell is able to replicate and immediately segregate its DNA. Z-ring assembly at midcell is initiated in parallel with the initiation of DNA replication and segregation. During stalked cell elongation, flagellar biogenesis is activated, culminating in the assembly of a polar flagella at the pole opposite of the stalk (Fig. 1).
Also at this time, assembly of the divisome occurs at the Z-ring scaffold and constriction of the cell begins. Complete compartmentalization of the cell does not occur until after the completion of DNA replication and segregation of an entire DNA molecule to each daughter cell. The end result is a polarized predivisional cell, which
3 will divide asymmetrically to produce two genetically identical daughter cells of different cell types and fates – a swarmer cell that re-enters the cell cycle at the beginning, and a stalked cell that is immediately capable of initiating DNA replication.
Figure 1. Caulobacter cell cycle. Schematic diagram of Caulobacter cell in various stages of the cycle. Caulobacter undergoes an asymmetric cell division which yields two morphologically distinct daughter cells: a motile swarmer cell that is replication inert (reminiscent of the eukaryotic G1 phase) and a non-motile stalked cell that is competent to start DNA replication. The period of active DNA synthesis and replication (S) includes the stalked and early predivisional cell. The G2 phase starts in the late predivisional cell when replication is completed and lasts until cell division has occurred. The boxes indicate the approximate time of cell cycle events.
4 1.2 FtsZ
FtsZ is ubiquitous in eubacteria and is also found in archaea and chloroplasts
(de Boer et al. , 1989; Gilson & Beech, 2001). It is a highly conserved tubulin-like
GTPase that forms a ring at the division plane. In the absence of FtsZ, none of the other cell division components are present at midcell. The crystal structure of FtsZ from Methanococcus jannaschii shows great similarity with αβ -tubulin heterodimer structure (Erickson, 1995; Lowe & Amos, 1998; Burns, 1995; Nogales et al. , 1998)
Both proteins consist of two globular domains: the N-terminal GTP-binding domain and the C-terminal domain (Lowe & Amos, 1998). Biochemically, the eukaryotic cytoskeleton tubulin binds to GTP, and GTP binding promotes the assembly of tubulin.
Likewise, FtsZ can polymerize into both single-strand filaments and lateral protofilaments in vitro (Addinall & Lutkenhaus, 1996b; Lowe et al. , 2004).
Osawa et al. recently proposed that the bending of FtsZ protofilaments generates constriction force and demonstrated that the membrane-targeted FtsZ (by splicing an amphipathic helix to its C terminus) could tether itself to the membrane
(Osawa et al. , 2009). Specifically, it was able to assemble into contractile rings when incorporated inside tubular liposomes in vitro . Furthermore, Li et al. used cryo- electron tomography to show that Caulobacter FtsZ filaments appeared to localize as a collection of short arcs near the division site (Li et al. , 2007b). Altogether, bending of protofilaments seems to be a likely mechanism for generating the constriction force, but the detailed relation of bending to GTP hydrolysis remains to be determined.
The functions of FtsZ are not limited to cytokinesis. Recent studies indicate that the machinery is responsible for cell shape and that it directs septal peptidoglycan
5 synthesis. For example, the Caulobacter MreB protein, an actin homologue, which contributes to the rod shape of the cell, contracts towards the midcell at the time of division initiation (Gitai et al. , 2004). This contraction is dependent on FtsZ.
Furthermore, FtsZ also plays a major role in cell wall elongation by spatially and temporally regulating the location of MurG, which produces the essential lipid II peptidoglycan cell wall precursor (Aaron et al. , 2007).
1.3 Regulation of Z-ring formation
The Z-ring is a highly dynamic structure, presumably allowing the cell to rapidly regulate Z-ring formation by modifying the dynamics of assembly and disassembly rate (Anderson et al. , 2004). This dynamism was revealed by studies utilizing fluorescence recovery after photobleaching FtsZ-YFP in live E. coli and B. subtilis cells by Anderson et al. (Anderson et al. , 2004). They found that the half-time of individual subunits of FtsZ in the ring was about 9 s in both E. coli and B. subtilis .
Because Z-ring must be actively maintained, it is influenced by a group of proteins that are modulating the equilibrium of FtsZ between an unassembled cytoplasmic pool and the assembled ring. Some of them serve as anchors for binding FtsZ to the inner membrane or recruit other proteins to the divisome. Indeed, several positive and negative regulators of Z-ring assembly have been identified.
Negative regulator of Z-ring assembly
The role of negative regulators is to ensure that the Z-ring is assembled at the right time and place. Thanbichler and Shapiro demonstrated that the selection of the division in Caulobacter is achieved by a driven by MipZ (Thanbichler & Shapiro,
6 2006). It was shown that MipZ directly interacts with FtsZ in vitro , and stimulates its
GTPase activity which inhibits the polymerization of FtsZ (Thanbichler & Shapiro,
2006). MipZ inhibits the polymerization of FtsZ and since MipZ localizes to the poles by binding to parS (a centromere-like site) via ParB protein (Toro et al. , 2008), FtsZ only assembles near the midcell, which has the lowest concentration of MipZ (Fig.
2A). Therefore, MipZ serves as a spatial regulator coordinating chromosome segregation with cell division in Caulobacter .
Caulobacter cells manage to divide properly without using either the Min system or nucleoid occlusion mechanisms. However, many bacterial species contain the MinCDE system including E. coli and B. subtilis . The absence of either MinC or
MinD in both E. coli and B. subtilis results in the polar formation of Z-rings. In E. coli ,
MinC is the FtsZ assembly inhibitor, and MinD recruits MinC to the membrane, where it is in position to counteract FtsZ assembly. The MinCD complex together oscillates between two ends of the cell (Fig. 2B, (Corbin et al. , 2002; Lowe et al. , 2004; Yu &
Margolin, 1999)). A third protein, MinE, prevents the accumulation of MinCD at midcell (Margolin, 2001). It competes with MinC for MinD binding, and then acts to expel membrane-bound MinD by stimulating its ATPase activity which triggers the release of MinD from the membrane (Hu et al. , 2002; Margolin, 2001; de Boer et al. ,
1989; Marston et al. , 1998). Therefore the dynamic movement of the MinCDE system prevents aberrant cell division by masking the polar sites, thereby leaving the Z-ring assembly at midcell.
Another important spatial regulatory system is nucleoid occlusion. Recent work from both E. coli and B. subtilis has confirmed that nucleoid occlusion is
7 mediated in regulation FtsZ ring assembly (Wu et al. , 2009). During DNA replication and segregation, the nucleoids occupy positions between the pole and midcell that prevents cell division occurring over the highly condensed bacterial chromosome. In E. coli , nucleoid occlusion is mediated by SlmA, which binds nonspecifically to DNA and concomitantly impairs FtsZ assembly (Bernhardt & de Boer, 2005). Similarly, deletion of B. subtilis Noc protein resulted in the aberrant formation of Z ring structures and extensively overlapped nucleoids (Wu et al. , 2009).
Positive regulator of Z-ring assembly
The positive regulators stabilize FtsZ assembly while negative regulators dissociate it, thus establishing equilibrium. Various proteins have been shown to stimulate formation of FtsZ rings. In E. coli, FtsA and ZipA both bind to a conserved
C-terminal sequence of FtsZ (Hale et al. , 2000; Wang et al. , 1997; Pichoff &
Lutkenhaus, 2005). ZipA is a membrane protein that binds to FtsZ through the interaction with its C-terminal domain (Din et al. , 1998; Johnson et al. , 2004; Addinall
& Lutkenhaus, 1996a). In addition, RayChaudhuri has shown that ZipA promotes the lateral association of FtsZ protofilaments in vitro (RayChaudhuri, 1999). FtsA is a homologue of actin possessing an amphipathic helix responsible for interaction with the cytoplasmic membrane (Addinall & Lutkenhaus, 1996a). FtsA and ZipA play important roles in anchoring the midcell location of Z ring, but each partially masks the contribution of the other. Only when both are inactivated is a severe defect in Z ring formation observed. ZipA has been identified in only a subset of Gram-negative bacteria (RayChaudhuri, 1999; Johnson et al. , 2004), whereas FtsA is much more widely conserved in the bacterial kingdom. This finding raises the possibility that FtsA
8 is the primary factor responsible for anchoring FtsZ filaments to the membrane in most bacteria. Recent studies in B. subtilis have shown that SepF is a newly identified divisome component (Hamoen et al. , 2006). It is conserved among Gram-positive bacteria and was shown to interact directly with FtsZ by two-hybrid analysis (Singh et al. , 2008; Hamoen et al. , 2006).
Another positive regulator, ZapA, is widely conserved across bacteria. ZapA was found to promote FtsZ polymerization and bundling in vitro , and deletion of zapA led to death of cells with abnormally low FtsZ levels (Gueiros-Filho & Losick, 2002;
Small et al. , 2007). In contrast to FtsA and ZipA, ZapA is not essential for cell viability of E. coli or B. subtilis under normal laboratory conditions. We show in
Chapter 4 that Caulobacter ZapA counteracts MipZ, a cell division inhibitor, in stabilizing assembly of FtsZ filaments in vitro .
To identify unknown FtsZ-binding proteins in Caulobacter , Goley et al. developed an assay involving overproduction of a GTPase-defective mutant of FtsZ, causing cells to exhibit a distinct morphology: long, slender constrictions containing
FtsZ separating the cell bodies. Fluorescent fusions to all known Caulobacter FtsZ- binding proteins were shown to colocalize with FtsZ in the constriction sites of mutant cells. From this screen, two more novel FtsZ-binding proteins, FzlA and FzlC, were identified. These two proteins are both conserved in α-proteobacteria. Specifically,
FzlA modifies the conformation of FtsZ filaments into helical bundle structures (Goley et al ., 2010b). These finding show that molecular differences exist in the composition of the divisome across different bacterial families.
9
10 Figure 2. Models for division-site selection in Caulobacter (A) and E. coli (B).
Shown are different stages of the swarmer cell cycle, beginning with a newborn swarmer cell and finishing with cell division that produces two daughter cells. (A) In
Caulobacter , division site placement by the MipZ and ParB. MipZ forms a complex with the ParB protein to a centromere-like site, parS . Chromosome replication generates two copies of the origin region, which immediately move together with
MipZ and ParB toward the cell poles. As a consequence, a gradient of MipZ is established. Due to the inhibitory effect of MipZ on FtsZ polymerization, the Z-ring therefore assembles near midcell with the lowest concentration of MipZ. (B) MinE is in yellow, FtsZ in orange, MinD in blue, and MinC in purple. In E. coli , MinE localizes to a ring-like structure at or near the middle of the cell early in the division cycle. MinD accumulates alternately at the membrane periphery on either side of the
MinE ring. MinCD complex oscillates between two cell poles. Therefore cooperation of the nucleoid occlusion and MinCDE system prevents aberrant cell division.
11 1.3 Overview of the late cell division proteins
The cell division process requires a set of approximately twenty proteins localizing from the cytoplasm to the outer membrane in Gram-negative organisms.
Most of the late cell division proteins contain large extracytoplasmic domains, suggesting that they might function in the construction of septal peptidoglycan structure modifications. Although physiological roles for a few cell division proteins have been identified in both E. coli and B. subtilis , the function of the majority of the division proteins is still unknown. It remains to be clarified to what extent the results obtained in these model systems can be applied to Caulobacter and other bacteria.
Many of the essential components of the E. coli divisome have been identified in a variety of bacterial families (den Blaauwen et al. , 2008), suggesting that the core divisome machinery is generally conserved. Since molecular differences exist in the composition and the regulation of the divisome across different bacterial lineages, I sought to understand the minimal set of proteins belonging to this core division apparatus by studying the Caulobacter divisome in the context of the functional analysis. A summary of the features of the core late division components is as follows.
FtsK is a bifunctional protein that functions in chromosome segregation and cell division in E. coli , B. subtilis (SpoIIIE) and Caulobacter (Wang et al. , 2006; Wu
& Errington, 1998). FtsK comprises three distinct domains, including the multispanning membrane domain at the N-terminus, a central domain, and the C- terminal domain (Wang et al. , 2006). The N-terminal domain is essential to target
FtsK to the midcell and the C-terminus functions as a DNA translocase that likely mediates clearance of the terminal regions of the chromosomes from the division site.
12 Interestingly, the C-terminal domain of FtsK is essential in Caulobacter , which is in direct contrast to what has been observed in E. coli , suggesting that chromosome segregation may be more tightly coordinated with cell division in Caulobacter. The N- and C-terminal domains of FtsK are linked by a proline-glutamine rich linker, which was shown to be involved in stabilizing the interaction of FtsK with other division proteins (Lau et al. , 2003; Wang et al. , 2006).
The FtsB, FtsL, and FtsQ proteins in E. coli and their homologs DivIB, FtsL, and DivIC in B. subtilis form a conserved subcomplex within the divisome. This complex is necessary for linking the upstream division proteins, which are predominantly cytoplasmic, with the downstream division proteins, which are predominantly periplasmic. Other than their structural role, no function has been attributed to these proteins so far.
FtsI is a division-site specific transpeptidase required for peptidoglycan synthesis. Peptidoglycan synthesis is abolished during division but still supports cell elongation in the absence of FtsI (Weiss et al. , 1997). A number of studies have suggested that the interactions between FtsI with other divisomal proteins are critical for its localization at midcell (Weiss et al. , 1997; Costa et al. , 2008).
FtsN is not a well-conserved protein, and it is generally thought to be limited to enteric bacteria and their close relatives (Moll & Thanbichler, 2009). FtsN shows weak sequence similarity to cell wall amidases, suggesting a possible role for cell wall hydrolysis. Recently, Moll and Thanbichler identified Caulobacter FtsN based on structural and functional similarity to E. coli FtsN (Addinall et al. , 1997; Moll &
Thanbichler, 2009). They further identified FtsN-like cell division proteins in β- and δ-
13 proteobacteria, suggesting that FtsN is widely conserved among Gram-negative bacteria.
Since protein–protein interaction is critical for the assembly of the divisomal proteins, a number of studies have analyzed interactions between divisomal proteins
(Goehring et al. , 2005; Maggi et al. , 2008; Lowe et al. , 2004). Two-hybrid analyses indicate a complex network of interactions among proteins of the divisome, and some of these predicted interactions have been corroborated by co-immunoprecipation or by in vitro analysis of purified proteins. In E. coli , studies on the localization dependency of cell division proteins in conditional mutants revealed that the components of the divisome are recruited to midcell according to a linear hierarchy pathway (Fig. 3).
Interestingly, the assembly interdependency for divisome formation in B. subtilis seems to suggest a multiprotein subcomplex. Several lines of evidence suggest that their recruitment to the midcell is fully independent (Fig. 3). Recent studies have suggested, however, that the hierarchical order does not always reflect localization dependency or timing of the assembly. The following section details current knowledge of the dynamics of the divisome assembly.
14
Figure 3. Pathway of divisome assembly hierarchy in E. coli and B. subtilis . In each of these pathways, assembly of a given protein is dependent on all upstream proteins (to the left) to localize and is, in turn, required for the localization of proteins downstream (to the right) (adapted from (Goehring & Beckwith, 2005)).
15 1.4 Assembly of the divisome
An important question in bacterial division is to define how the multiprotein complex of the divisome is assembled. Prior models for the structure and assembly of the bacterial divisome have been derived mostly from the studies in E. coli and B. subtilis . A current model proposes that the temporal assembly and maturation of the divisome in E. coli and B. subtilis occurs in two well-defined steps (Aarsman et al. ,
2005; Gamba et al. , 2009). In E. coli , the average cell age at which a number of cell division proteins arrival at midcell was analyzed by immunofluorescence and GFP- fluorescence microscopy (Aarsman et al. , 2005). It was shown that the Z-ring localizes at midcell, in parallel with arrive of FtsZ-interacting proteins. After approximately 17 min, the second group of divisome proteins assembles. To follow the assembly of B. subtilis divisome, Gamba et al. examined the localization pattern of five division proteins of outgrowing cells after spore germination (Gamba et al. , 2009). A two-step assembly model appears to be also present in B. subtilis . Furthermore, the delay appeared strikingly constant and comparable to the delay in E. coli . Knowledge of the order of events and the localization dependencies along the pathway of cytokinetic ring assembly and constriction are prerequisites for deciphering the underlying molecular mechanisms. Since Caulobacter is easily synchronized with respect to cell cycle progression, I took advantage of this feature to analyze assembly of the divisome over time and gain insight into the temporal progression of cell division.
Besides FtsZ, the cell division machinery in Caulobacter includes ZapA, FtsA,
FzlA, FzlC, DipM, MurG, MreB, FtsQ, FtsE, FtsX, FtsK, FtsB, FtsL, FtsW, FtsI, FtsN,
AmiC, Tol-Pal, TipN and KidO (Fig. 4). Until recently, little was known about the
16 dynamics of cell division in Caulobacter , although the position of FtsZ, MurG, MreB,
FtsN, FtsA, FtsI and FtsK in the assembly hierarchy of the Coulobacter divisome was examined from previous studies (Aaron et al. , 2007; Moll & Thanbichler, 2009; Costa et al. , 2008). There are still many unanswered questions about the dynamics and hierarchy of the divisome. To gain comprehensive insight into the temporal recruitment of all components of divisome proteins, I synchronized different bacterial strains, each expressing a different fluorescently tagged cell division protein, and imaged them throughout the cell cycle. The details of these studies are in Chapter 2.
Figure 4. Schematic diagram of Caulobacter divisome. The divisome is an assembly of at least 24 proteins that localize to the division plane and function together to drive invagination and fission of the inner and outer membranes and synthesis and remodeling of the peptidoglycan cell wall in coordination with chromosome segregation and other cell cycle events.
17 1.5 Overview of other modes of bacterial cell division
Cell division in Caulobacter differs from that of E. coli or B. subtilis in several ways. Notably, cytokinesis in the Caulobacter is achieved by constriction of the inner membrane and peptidoglycan, whereas cytokinesis in E. coli (or septum forming bacteria) appears to occur by simultaneous formation of a septum and cell envelope constriction. A summary of the features of other bacterial cell division mechanisms is as follows.
Sporulation of B. subtilis
In B. subtilis , cell division normally occurs at the midcell during vegetative growth. However, upon nutrient deprivation, cells initiate a developmental program that culminates in the formation of a spore, a cell type capable of resisting various environmental stresses for long periods of time. One of the distinct steps in this developmental program is the switch from medial to polar cell division. The smaller cell will become the spore, and the larger cell is the mother cell. Distinct programs of gene expression regulate the cell differentiation of spore and mother cell compartments. During the spore formation, the same cell division apparatus is responsible for the polar septum formation, whereas the alteration in Z ring localization from midcell to the poles is controlled by the master response regulator
Spo0A and the alternative sigma factor σH (Fujita & Losick, 2003; Lu et al. , 2003;
Barak & Wilkinson, 2005). It is not yet known what mechanisms directly contribute the change in preference for division site. Ben-Yehuda and Losick recently showed that SpoIIE, a membrane-bound phosphatase, and elevated levels of FtsZ may play
18 important roles in effecting the switch to polar Z ring formation (Ben-Yehuda &
Losick, 2002; Guberman et al. , 2008).
Cell division of L-form bacteria
L-form bacteria, cell wall deficient bacteria, were recently shown to be capable of division in the absence of FtsZ (Leaver et al. , 2009). Firstly, the cells increased in size, and started forming blunt protrusions on the surface. Subsequently, the protrusion resolved by cleaving into several daughter cells (Leaver et al. , 2009). Although the molecular details underlying this process remain unclear, the ability of L-form bacteria to grow and divide without the presence of either cell wall or FtsZ may represent a form of cell division in early forms of life.
Cell division of Cocci
The molecular mechanism of cell division in cocci bacteria is not well characterized.
In most rod-shaped cells, FtsZ assembles a ring-like structure at the midcell to divide the daughter progenies. However, it is more complex in cocci bacteria, which have an very large number of alternating cell division planes that can give rise to the problem of identifying future division site. The division site selection of enterococci cells is differentiated by identification of new cell wall synthesis (Zapun et al. , 2008).
However, other cocci, such as Deinococcus radiodurans and Neisseria gonorrhoeae, appear to divide in alternating planes (Chou & Tan, 1991; Westling-Haggstrom et al. ,
1977).
19 1.6 Constriction and invagination of the cell envelope
After the divisome is fully assembled, the invagination of the cell envelope begins. The divisome regulates constriction of the cell envelope: it synchronizes inner membrane fusion, coordinates septal peptidoglycan synthesis and invaginates the outer membrane (OM). In E. coli and B. subtilis , the inner membrane (IM) and the cell wall invaginate together, forming the septum, and the outer membrane constricts later
(Rothfield et al. , 1999). However, Caulobacter OM invagination is temporally and spatially separated from peptidoglycan and IM invagination. In the early stages of cell division, the IM and OM are constricted simultaneously. Later, the IM and peptidoglycan layers constrict faster, creating a separation of the IM and OM near the division plane about 17 min before cell separation (Judd et al. , 2005). The mechanism that implements the delayed constriction of the OM layer of the cell envelope is poorly understood.
Curiously, none of the known essential cell division proteins are located in the
OM, thereby raising the question of how the OM is able to invaginate and separate the two daughter cells. A recent paper provides some evidence that the Tol-Pal complex in
E. coli may be involved in this process (Gerding et al. , 2007).
The Tol-Pal complex is widely conserved among Gram-negative bacteria and plays multiple physiological roles. In E. coli , it is specifically involved in maintaining
OM interaction with the peptidoglycan, expressing lipopolysaccharide surface antigens and virulence factors, facilitating infection by filamentous DNA phage and reducing sensitivity to detergents (Lloubes et al. , 2001; Deprez et al. , 2005; Paterson et al. , 2009; Llamas et al. , 2003b). Cell chains with lateral membrane blebs were
20 formed in E. coli in low osmolarity medium when Tol-Pal complex was inactivated
(Gerding et al. , 2007). The results suggest that Tol-Pal plays a role in completing cell division under conditions of membrane stress in these organisms. We addressed in
Chapter 3 the function of the Tol-Pal complex in Caulobacter .
Collaborative parts of this thesis
Some of the work described in this thesis was carried out in collaboration with other researchers. In chapter 2, construction and characterization of the divisome assembly was a collaboration with Dr. Erin Goley in Lucy Shapiro’s laboratory. Dr.
Goley and I worked jointly to construct all the strains, collected the microscopy images, and analyzed the results.
In chapter 3, cryo electron microscopy (CryoEM) was performed in collaboration with Dr. Comolli in Ken Downing’s laboratory. I constructed strains and performed genetic analysis. Dr. Comolli collected all of the cryoEM images used in the final analysis.
21
Chapter 2
Assembly of a bacterial cell division machine
Note: The majority of the information presented in this chapter was included in the publication: Goley EG*, Yeh YC*, Hong SH, Fero MJ, Abeliuk E, McAdams HH, and Shapiro L.
(in preparation) Assembly of a bacterial cell division machine
* These authors contributed equally to this work.
22 Introduction
Progression of the cell cycle culminates in the physical separation of the mother cell into two daughters through the process of cytokinesis. In the vast majority of bacteria, cytokinesis is mediated by the divisome, a multiprotein complex that assembles near midcell and drives invagination of the inner membrane (IM), synthesis and remodeling of the peptidoglycan (PG) cell wall and, in Gram-negative organisms, invagination of the outer membrane (OM). The tubulin-like GTPase, FtsZ, forms the structural basis of the divisome, acting as a scaffold for assembly of the rest of the division machinery (Margolin, 2005; Goehring & Beckwith, 2005) and generating constrictive force (Osawa et al. , 2008; Osawa et al. , 2009). Over a dozen proteins assemble downstream of FtsZ and are essential for cytokinesis. The precise roles of these factors in executing division are mostly unknown, however existing evidence supports the following general functions: interaction with and stabilization of the FtsZ ring (FtsA, ZapA, FzlA) (Addinall & Lutkenhaus, 1996a; Beall & Lutkenhaus, 1992;
Goley et al. , 2010b; Gueiros-Filho & Losick, 2002; Martin et al. , 2004; Ohta et al. ,
1997; Sackett et al. , 1998), synthesis and remodeling of peptidoglycan (DipM, FtsW,
FtsI/Pbp2B, AmiC) (Bernhardt & de Boer, 2003; Boyle et al. , 1997; Costa et al. , 2008;
Daniel et al. , 2000; Goley et al. , 2010a; Henriques et al. , 1992; Moll et al. , 2010a;
Poggio et al. , 2010b; Weiss et al. , 1997), coordination of division with chromosome segregation (FtsK/ SpoIIIE) (Wang et al. , 2006; Wu & Errington, 1994; Yu et al. ,
1998), outer membrane invagination (Tol-Pal complex) (Gerding et al. , 2007; Yeh et al. , 2010), and stabilization of interactions within the divisome (FtsQ/DivIB, FtsL,
FtsB/DivIC, FtsN) (Addinall et al. , 1997; Buddelmeijer et al. , 2002; Chen et al. , 1999;
23 Ghigo & Beckwith, 2000; Katis et al. , 1997; Martin et al. , 2004; Moll & Thanbichler,
2009; Rowland et al. , 1997; Sackett et al. , 1998; Sievers & Errington, 2000).
The physical process of cell division must be tightly coordinated in time and space with other cell cycle events, such as cell growth, segregation of the genome, and cellular development. The need for this coordination is particularly evident in the dimorphic α-proteobacterium, Caulobacter crescentus , in which the cell division site has emerged as a critical spatial landmark that receives signals from the cell poles and relays them to downstream cell cycle events. Caulobacter begins its cell cycle as a motile swarmer cell in which FtsZ is localized at the new cell pole opposite the chromosomal centromere that is anchored at the old, flagellated pole (Thanbichler &
Shapiro, 2006). The swarmer cell undergoes a developmental transition to a stalked cell, shedding its polar flagellum, building a stalk in its place, and initiating replication of the single, circular chromosome. Upon duplication, one copy of the chromosomal centromere is quickly segregated to the opposite cell pole, bringing with it a bound complex of the ParB partitioning protein and the MipZ inhibitor of FtsZ polymerization (Viollier et al. , 2004; Thanbichler & Shapiro, 2006; Toro et al. , 2008).
The completion of centromere segregation results in bipolar localization of MipZ. The ability of MipZ to promote depolymerization of FtsZ displaces FtsZ from the new cell pole, and it reassembles into a polymeric ring structure (the Z ring) at the site of lowest MipZ concentration in the cell: roughly midcell (Thanbichler & Shapiro, 2006).
In this way, the site of Z ring assembly, and therefore the future division site, is specified in coordination with chromosome segregation.
24 In addition to its central role in cell division, Caulobacter FtsZ also recruits proteins that direct cell elongation and cellular polarity. Prior to directing inward growth of PG during division, FtsZ organizes midcell-localized PG synthesis for the elongation phase of growth (Aaron et al. , 2007). This is mediated, at least in part, by the midcell recruitment of MurG, which catalyzes the last step in lipid II (PG precursor) synthesis. The actin homolog, MreB, is also localized to midcell in an FtsZ-dependent manner, and is functionally implicated in PG synthesis and cell shape maintenance
(Gitai et al. , 2004; Figge et al. , 2004). Subsequently, through an unknown mechanism, there is a switch from elongation phase PG synthesis to division. IM fission and the resulting compartmentalization of the cytoplasm late in the division process is functionally linked to specifying the developmental fate of the two daughters, since it yields biochemically distinct compartments that contain different concentrations and/or post-translational modifications of critical regulatory proteins (Ausmees &
Jacobs-Wagner, 2003; McAdams & Shapiro, 2003). Very late in the cell cycle, the new pole marker, TipN, is recruited to the division site, and it remains at the new pole after cell separation to establish its identity and direct polar development (Lam et al. , 2006;
Huitema et al. , 2006).
Understanding how the divisome functions to implement cell division, and how its activity is regulated and integrated into the cell cycle, requires detailed knowledge of its assembly mechanism in time relative to cell cycle progression.
Comprehensive genetic analyses in Escherichia coli suggest a linear hierarchy of divisome assembly that might indicate a simple series of binary interactions that lead to formation of a functional divisome (Goehring & Beckwith, 2005). However,
25 studies aimed at identifying protein-protein interactions among divisome components in both E. coli and Bacillus subtilis indicate the existence of a highly complicated network of interactions (Goehring et al. , 2005; Buddelmeijer & Beckwith, 2004; Di
Lallo et al. , 2003). Microscopy experiments aimed at following the midcell localization of a subset of divisome proteins over time in E. coli and B. subtilis revealed a two-step process: FtsZ and its associated proteins localize to midcell first, and late divisome proteins assemble after a substantial maturation period (Aarsman et al. , 2005; Gamba et al. , 2009). Not all divisome proteins were included in those studies, however, and they were not performed using synchronized vegetatively growing cells.
Caulobacter is more easily synchronized than E. coli or B. subtilis , and is thus ideally suited to high-resolution temporal studies. Moreover, its cell cycle and developmental program is well-characterized, allowing us to place the arrival of each protein into functional context with respect to cell cycle events. Several cell division proteins, including FtsZ, FtsA, FtsQ, DipM, and FzlA, have been reported to be cell cycle regulated at the transcript and/or protein levels in Caulobacter , indicating that divisome assembly and function is tightly regulated in time (Goley et al. , 2010a;
Goley et al. , 2010b; Martin et al. , 2004; Quardokus et al. , 1996). Scattered reports of the temporal pattern of localization of a number of Caulobacter divisome proteins have been published and are mostly consistent with the two-step assembly pathway reported in other organisms (Addinall et al. , 1997; Aaron et al. , 2007). To date, however, a comprehensive analysis of the assembly of all divisome and division-site localized proteins in this organism has not been reported. Our goal in the current
26 study was to systematically analyze the temporal regulation of 19 divisome and division-site-localized proteins over the course of the cell cycle in Caulobacter . To this end, we analyzed their localization, transcript levels, and protein levels over time in the cell cycle, and performed extensive analysis of their genetic dependency of localization. Our results allow us to present a model for divisome assembly with previously unattained temporal resolution that leads to significant insights into divisome regulation and function.
27 Materials and methods
Bacterial strains, synchronization, and growth conditions.
All Caulobacter strains were derived from CB15N and grown at 28°C in peptone yeast extract (PYE) or M2-glucose minimal medium (M2G) with select antibiotics, if any.
All experiments were performed with cells in log phase of growth in PYE with the exception of synchronization of CB15N (for determination of M2 protein levels over the course of the cell cycle), which were grown in M2G. Small-scale synchrony for microscopy was performed as described previously (Tsai & Alley, 2001). For the depletion strains, cells were grown in PYE medium containing 0.3% xylose, washed with plain PYE medium three times, and then re-suspended in PYE medium containing 0.2% glucose or 0.3% xylose. Samples were taken and analyzed by phase contrast and fluorescence microscopy. The construction of bacterial strains is detailed in the Table SI.
Microscopy and image analysis
For localization studies, 0.3% xylose and 0.5mM vanillic acid (pH 7.5) as appropriate were used to induce expression of protein fusions. For time-course experiments, cells were grown in PYE medium, synchronized when appropriate, and viewed on 1% agarose in M2G media. Phase contrast and fluorescence microscopy images were obtained using a Leica DM 6000 B microscope with a HCX PL APO 100x/1.40 Oil
PH3 CS objective, Hamamatsu EM-CCD C9100 camera, and a custom-designed microscope control and image analysis software KAMS (Christen et al. , 2010).
Images were processed with Adobe Photoshop.
The quantitative measurements of localization pattern of fluorescence images were performed manually in ImageJ. The quantitative measurements of size, membrane
28 boundary and degree of invagination as a function of time were performed automatically in MATLAB. The invagination measurements were fit with a sigmoid function and the first derivative was used to characterize the time to invagination. The histogram of time to half invagination shows a narrow distribution of characteristic times with a mean and standard deviation of 66.2+/-8.1 minutes. The quantitative measurements of unipolar and biopolar MipZ peaks were performed automatically in
MATLAB. The cells are located by applying calibrated threshold to the phase contrast images. Then the corresponding cell regions of fluorescence images were segmented out. The cells were aligned along their long axis by regionprops function in MATLAB.
We modified pkfnd from SPtrack for our purpose and used it to analyze the number of
MipZ peaks.
Curve fitting
The half-maximum time of the appearance of fusion proteins at midcell were analyzed using the sigmoidal dose response function of graphical program Prism (GraphPad,
San Diego, CA).
Transcriptional regulation analysis
For each profile, we performed a cubic-spline interpolation followed by an affine transformation so that the expression values of each profile ranged between 0 and
100%. We then ordered these 10 genes according to the time-points at which each one of these profiles reached 50% expression, relative to their peak expression minus basal expression values (bottom gene, ftsZ, is the first one to reach 50% expression).
Immunoblotting
For a given experiment, equivalent OD units of cell lysate were loaded. Western blotting to monitor Flag fusions, HU and FtsZ was performed with standard
29 procedures. Samples were probed with primary antibodies at the following concentrations: α-Flag (1:1000), α-FtsZ (1:10 000) (Thanbichler & Shapiro, 2006), α-
ZapA (1:4000), and α-HU2 (1:5000). Secondary horse radish peroxidase-conjugated goat anti-rabbit or donkey anti-mouse antibodies were used at a 1:10 000 dilutions and chemiluminescent substrate was applied prior to exposure to film.
30 Results
Strategy for determining the timing of divisome assembly
We selected 19 proteins that localize to the cell division site in Caulobacter and participate in cell growth and morphology specification (MurG, MreB, DipM), cell division (ZapA, FzlA, DipM, FtsZ, A, E/X, K, Q, L, B, W, I, N, Tol-Pal complex), or cell polarity specification (TipN, MreB), or that have unknown functions (FzlC, KidO)
(Goley et al. , 2010b; Radhakrishnan et al. , 2010) (Fig. 5A). For each, we aimed to determine the time at which they first appear at the incipient division site. In order to compare localization timing between strains, we integrated an inducible copy of the fluorescent fusion in question at the chromosomal vanA or xylX locus for vanillate or xylose-inducible expression, respectively, in a strain background bearing mipZ- cerulean as the only copy of mipZ at its native chromosomal locus (Fig. 5B). The dynamics of MipZ localization are well-characterized (Thanbichler & Shapiro, 2006) and served as a marker for comparing cell cycle progression between strains. We synchronized each strain, suspended the isolated swarmer cells in rich PYE liquid media for growth at 28 oC, took samples at 10-minute intervals, and imaged phase contrast, MipZ-Cerulean, and the fluorescent fusion of interest at each time point.
Under these conditions, the cell cycle was completed in ~90 minutes. At each time point in each strain we quantified: 1) the localization of MipZ (i.e. percentage of cells with monopolar vs bipolar MipZ), 2) the localization of the protein in question (i.e. percentage of cells with polar, diffuse, midcell, or other localization) 3) and the percentage of cells with an invagination at the incipient division site visible by phase contrast and/or the degree of envelope invagination (Fig. 5C). Each strain was
31 independently synchronized and analyzed at least twice.
Representative images from the synchrony of strain EG490, bearing mipZ- cerulean at the mipZ locus and ftsZ-yfp at the vanA locus are presented in Fig. 5D and quantification of the synchronies of that strain is presented in Fig. 5E. By 10 minutes post-synchrony, most cells exhibited bipolar MipZ localization, indicating that the chromosomal centromere was duplicated and segregated, and by 30 minutes essentially all cells had bipolar MipZ. MipZ remained bipolar until cell separation at
~90 minutes, when we observed a marked decrease in bipolar MipZ localization owing to the presence of daughter cells that had not yet duplicated and/or segregated their centromeres. We frequently observed very late predivisional cells that had 3
MipZ foci: one each at the extreme cell poles, and one near midcell (18.7 ± 3.9% of cells at t = 80 min, n = 6 synchronies, 210-557 cells each). We interpret these to be cells wherein the cytoplasm has been compartmentalized, allowing DNA replication and centromere segregation in the stalked compartment prior to the completion of splitting of the PG and OM fission. We observed visible invagination of the envelope, marking the switch from the elongation mode of growth to the division mode, in the majority of cells by 60 minutes post-synchrony. This, too, decreased upon cell separation at ~90 minutes, when daughter swarmer and stalked cells lacking visible invaginations were plentiful.
In newborn swarmer cells of strain EG490, FtsZ-YFP was observed at the new cell pole opposite the MipZ-Cerulean focus, and assembled into a loose band at midcell shortly after bipolarization of MipZ, as reported previously (Fig. 5D, E). The
Z ring became more focused by 30 minutes post-synchrony, and FtsZ remained at
32 midcell until cell separation. At 90 minutes, we observed cells with monopolar MipZ and an FtsZ focus at the opposite pole, as well as daughter stalked cells that exhibited bipolar MipZ and a loose band of FtsZ at midcell.
Analysis of the localization of inducible fluorescent fusions of division-site localized proteins
We completed the analysis described above for FtsZ-YFP for an additional 18 division-site localized proteins. Quantification of the timing of bipolar localization of
MipZ, of initiation of invagination (Fig. 5F) and of half-maximal invagination (data not shown) in each strain analyzed revealed remarkably constant timing of these events. This indicated that the timing of cell cycle progression of each strain was similar and enabled direct comparison of the timing of midcell localization of division-site localized proteins across strains.
Each division-site localized protein exhibited a characteristic localization pattern (Fig. 6). Of the 19 proteins, 14 began the cell cycle at the new cell pole, while the rest were diffuse in swarmer cells (Fig. 6, 7). Of the polar proteins, 8 were displaced from the cell pole prior to accumulating at the division plane and 6 localized to both the pole(s) and midcell (Fig. 6, 7). Each protein arrived at midcell with characteristic timing (ranging from ~10 to 70 minutes in a 90 minute cell cycle), and most of them remained at midcell until the completion of cell division. A clear exception to that rule, however, is MreB, which was dispersed from midcell into a patchy distribution well before cell separation (Fig. 6).
33
34 Figure 5. Analysis of the dynamics of divisome assembly
(A) Schematic diagram of the Caulobacter divisome proteins and their predicted membrane topologies. IM: inner membrane; OM: outer membrane; PG: peptidoglycan;
Fts protein names have been abbreviated by excluding “Fts” from them. (B) Schematic representation of the strategy for generating fluorescent fusion proteins of interest.
(C) Illustration of the localization of MipZ (blue) and protein of interest (red) over the cell cycle. Arrows point to the key events we examined: appearance of MipZ bipolarization, initiation of invagination and cell separation. (D) Phase contrast and fluorescence micrographs of cells expressing MipZ-CER and FtsZ-YFP in strain
EG490 over the cell cycle. Cells were grown in PYE media with 0.5 mM vanillate for
1 h prior to synchrony, and subsequently applied to M2G agarose pads for imaging.
Cells were withdrawn from cultures at 10 min intervals and visualized by phase contrast and fluorescence microscopy to determine the localizations of fluorescently labeled proteins. (E) Percentage of cells with visible constrictions, bipolar MipZ-CER, and FtsZ-YFP localizations near midcell or at the pole were plotted as a function of time during a cell cycle of about 90 min. Protein localization and constriction patterns were examined using synchronized population of strain EG490 as (D). At least two independent time-course experiments were performed and over 500 cells were considered for each time point. (F) Temporal montage represents MipZ-CER bipolarization and the initiation of invagination as a function of time during the cell cycle of different strains. The mean appearance time of MipZ bipolarization and initiation of invagination are plotted in red and blue, respectively. The timing of MipZ bipolarization and initiation of invagination in all strains behaved similarly.
35 To assign an order of assembly of division-site localized proteins, we calculated the percentage of cells with a midcell band or focus of each protein at each time point (for representative data, see Fig. 8A, top). We then fit a curve to those data and found the time at which half-maximal midcell localization was achieved for each
(Fig. 8A, bottom). We performed similar curve fitting to calculate the timing of bipolar MipZ localization, initiation of invagination, and half-maximal degree of invagination. This analysis revealed a series of stages and transitions in divisome assembly and associated events (Fig. 8B). First, MipZ becomes bipolar, followed closely by the assembly of a loose band of FtsZ near midcell. Roughly 10 minutes later, the FtsZ binding proteins ZapA, FzlC, FtsE, and FzlA first appear at midcell.
Concomitant with FzlA localization at midcell, about 25 minutes into the cell cycle, proteins involved in cell growth and morphology specification (MreB, MurG, and
DipM) localize at midcell. Next, an apparent transition to division occurs wherein
TolQ (representing the Tol-Pal complex) and FtsA are recruited to the division site between 30 and 40 minutes post-synchrony. Subsequently, most of the divisome proteins (FtsN, FtsQ, FtsL, FtsI, and FtsK) arrive. FtsW and FtsB arrive 5-10 minutes after the core set of divisome proteins, at approximately the same time as initial invagination of the cell envelope is apparent. KidO then arrives at midcell, followed by dispersal of MreB into a patchy distribution, and midcell localization of TipN.
Fluorescent fusions expressed from native loci exhibit similar dynamics of assembly as inducible fluorescent fusions
Transcript levels of hundreds of genes in Caulobacter are known to vary over the course of the cell cycle and transcriptional regulation can be critical in specifying
36 the timing of cell cycle events. The approach detailed above circumvents transcriptional regulation since the genes encoding the fluorescent fusions were expressed constitutively from inducible promoters. To ensure that our expression strategy did not alter the normal timing of division site protein localization, we attempted to generate strains bearing fluorescent fusions to many of the genes encoding division-site localized proteins at their chromosomal loci under the control of their own promoters in the mipZ-cerulean background. We were able to do this for
ZapA, FzlA, FzlC, FtsE, DipM, MurG, FtsK, FtsI, and FtsW. We were unable to recover strains bearing fluorescent fusions of ftsZ , ftsA , ftsL, or ftsQ as the only copy of the gene at its chromosomal locus, indicating that the tags interfere with the essential functions of these proteins. We assessed cell cycle progression of each of the native tagged strains as described above and found that, although viable, the strains with mCherry-FzlC, DipM-mCherry, and Venus-FtsI produced as the only copy from the native gene locus grew more slowly than the others, so we did not include them in our analysis.
We determined the timing of midcell localization of ZapA, FzlA, FtsE, MurG,
FtsK, and FtsW produced from their native loci and compared them to their timing of localization when expressed from an inducible locus. The cell cycle dependent localization and arrival times of the native and inducible versions were indistinguishable for MurG, FtsK, and FtsW (Fig. 9). Native Venus-FtsE localized to the division site with at the same time as the inducible version, however the protein produced by native expression localized to the cell pole in swarmer cells, whereas the induced version did not (Fig. 6, 9). The native tagged versions of FzlA and ZapA
37 localized slightly earlier than the inducible versions, but in the same time window (i.e. between initial assembly of FtsZ at midcell and recruitment of proteins involved in PG synthesis). These results validate the use of inducible fluorescent fusions for determining the timing of divisome assembly.
38
39 Figure 6. Time-course of division-site localized proteins using fluorescence microscopy.
Protein localization of indicated fluorescent fusions to the Caulobacter division site.
Cells expressing YFP, Venus, mCherry and CER fusion proteins were grown in PYE media containing 0.3% xylose for 2 h or 0.5 mM vanillate for 1 h prior to synchrony.
The time-course montage of the representative images of nineteen strains over the cell cycle was shown at intervals of 10 min. At least 500 cells were analyzed at every time point. Dotted lines represent the outline of the cells for clarity when the background signal is low. The phase contrast and MipZ-CFP images correspond to the FtsZ-YFP images.
40
Figure 7. Polar localization patterns of division-site localized proteins
Bar graph depicting percentages of synchronized swarmer cells of indicated fluorescent fusions with characteristic polar localization pattern of Fig 7.
41
42 Figure 8. Assembly of the divisome takes place in a series of stages
(A) The upper panel shows the representative data of indicated proteins. Cells of strains EG420 ( PvanA-ftsZ–venus mipZ –cerulean ), EG503 ( PxylX –venus–zapA mipZ –cerulean ), EG519 ( PxylX–venus–mreB mipZ –cerulean ), EG494 ( PxylX–yfp– ftsA mipZ –cerulean ), EG493 ( PxylX–venus–ftsL mipZ –cerulean ), EG501 ( PxylX– venus–ftsW mipZ –cerulean ), and EG746 ( PxylX–tipN–mCherry mipZ –cerulean ) were grown and imaged as above. Percent of cells with visible midcell localizations of the fusion proteins were plotted as a function of time during a cell cycle. The lower panel shows the fitting curves by applying a sigmoidal dose response to calculate the half point of the signal change. Dashed line indicates 50% of cells with midcell localization. (B) Summary of time of occurrence at half-maximum for midcell assembly of division-site localized proteins, bipolar MipZ, dispersal of MreB, initiation of invagination and half-maximum of invagination. Each point represents individual time-course experiments. Proteins are grouped upon their functions and shown in different colors.
43
Figure 9. The timing of midcell localization of endogenously expressed fusion proteins
(A) Time-course montage of representative cells from indicated strains over the cell cycle is shown at intervals of 10 min. At least two independent time-course experiments were performed and over 500 cells were considered for each time points.
(B) Summary of time at half-maximum for seven endogenously expressed fusion proteins at midcell is shown.
44 The order of assembly observed between strains is maintained when pairwise assembly is followed in a single strain
Our analysis thus far indicates a specific order of assembly of proteins at the division site. To assess the robustness of that order and confirm the surprising results from that analysis, we selected pairs of proteins to image in the same cells. First, we selected three pairs of proteins that our analysis indicates arrive approximately simultaneously: ZapA with FtsE, ZapA with FzlA, and FtsI with FtsK. We generated strains bearing native tagged versions of each, synchronized them, and determined the percentage of cells with neither protein at midcell, both proteins at midcell, and either one or the other at midcell at a time point wherein about half the cells had both proteins at midcell. For ZapA/FtsE, we observed a small percentage of cells (1.5%) with an apparent enrichment of FtsE at midcell but no ZapA, and a slightly higher percentage (5%) with ZapA at midcell but no FtsE. This is consistent with our determination that native tagged ZapA arrives at midcell slightly ahead of native tagged FtsE (Fig. 10A). In the case of ZapA/FzlA double labeling, we never observed cells that had only one of these proteins at midcell (Fig. 10B), indicating that they do indeed arrive simultaneously, as suggested from our previous analysis. The same was true of FtsI/FtsK, although in that strain we found approximately equal percentages of cells that had FtsI at midcell but not FtsK, and FtsK at midcell but not FtsI (Fig. 10C).
Again, this suggests that these proteins arrive at approximately the same time.
Our temporal analysis revealed a few surprises in light of previous work.
Specifically, in E. coli , FtsW was reported to be required for recruitment of FtsI to the division site, but we find that FtsI arrives first of the two. We imaged native Venus-
45 FtsI with vanillate-induced mCherry-FtsW (the double native-tagged strains was viable, but filamentous) and found that, as suggested by the analysis of separate strains,
FtsI was often observed at midcell when FtsW was still absent (28.5% of cells 50 minutes post-synchrony) (Fig. 10D). FtsW was never observed to arrive prior to FtsI, confirming our previous results. Contrary to studies in E. coli and B. subtilis , which found that FtsA arrives at midcell approximately at the same time as FtsZ and other
FtsZ-binding proteins, we observed a significant delay in its recruitment. To verify this, we imaged double labeled strains bearing native ZapA-mCherry (representing the early-arriving FtsZ-binding proteins) with xylose-induced YFP-FtsA and mCherry-
FtsW (a late-arriving divisome protein) with xylose-induced YFP-FtsA. We observed a midcell band of ZapA without midcell FtsA in a significant fraction (17.5%) of cells, and never observed FtsA at midcell before ZapA, indicating that it does arrive well after initial Z ring assembly (Fig. 10F). Additionally, we found that a small, but significant, fraction of cells (4.5%) had FtsA at midcell before FtsW, whereas FtsW was almost never observed at midcell with FtsA there (Fig. 10E). Collectively, these data confirm the order of assembly determined by our initial analysis.
FtsZ and FtsL are key factors in assembly of the division machinery
The temporal assembly of divisome proteins we observed might indicate a series of protein-protein interactions wherein each protein recruits the next to arrive in time. Extensive genetic probing of the dependency relationships among division-site localized proteins in E. coli support the existence of such a serial recruitment mechanism. For example, in E. coli , FtsZ is required to recruit FtsA which is required to recruit FtsK, which is required to recruit FtsQ, etc. To test whether this is the case
46 in Caulobacter , we generated a set of strains with deletions of genes encoding non- essential divisome proteins and inducer-dependent expression of genes encoding essential divisome proteins (Table 2). We then introduced fluorescent fusions of other division-site localized proteins into these deletion or depletion strains and asked whether the fluorescent fusion was still localized to midcell foci in the absence of the deleted or depleted protein. Table 1 summarizes these results along with those of previously published experiments addressing the genetic dependency of division-site protein localization in Caulobacter .
We were surprised to find that, in contrast to the scenario in E. coli , very few proteins were strictly required for the localization of any others. The exceptions were
1) FtsZ, which was required for the localization of every other protein we tested (Table
1 and Fig. 11A), 2) components of the Tol/Pal complex, which were previously shown to be required for normal localization of TipN, 3) FtsX, which was required for the localization of the FtsE protein with which it forms a heterodimeric ABC transporter complex (Table 1 and Fig. 11D), and 4) FtsL, which was absolutely required for localization of FtsB and FtsQ and without which FtsI and FtsW were localized only weakly to midcell foci (Table 1 and Fig. 11E-I). We were notably surprised that none of the fluorescently tagged division-site localized proteins we tested required the presence of FtsA to localize to midcell foci, (Table 1 and Fig. 11B), since FtsA is placed very early in the hierarchical dependency for divisome protein localization in E. coli . Moreover, FtsW was not absolutely required for FtsI to localize to midcell foci in
Caulobacter (Table 1 and Fig. 11J). This differs from the scenario in E. coli where
FtsW is required for localization of FtsI to the division plane, but is consistent with
47 our observation that FtsI localizes prior to FtsW in Caulobacter . In sum, these results indicate that of the cell division proteins we were able to delete or deplete, FtsZ and
FtsL perform key roles in recruiting other factors to midcell and that the assembly of the division machinery does not depend upon a strictly linear array of protein-protein interactions.
Transcript and protein levels of a subset of division-site localized proteins are cell cycle regulated
To further address the mechanisms underlying the time-dependent recruitment of proteins to the division site, we asked whether any division-site localized factors are cell cycle regulated at the transcipt and/or protein level. Analysis of Affymetrix data from synchronized cells grown in minimal M2G media indicated that transcript levels of 10 of the 19 genes are strongly cell cycle regulated (Fig. 12A). ftsZ transcript levels peak in stalked cells, whereas ftsW , murG , ftsI , ftsQ , fzlA , ftsB , ftsA , ftsK , and kidO peak in pre-divisional cells (Fig. 12A and (McGrath et al. , 2007)). Protein levels of
FtsZ, FzlA, FtsA, FtsQ, DipM, and KidO have previously been reported to vary over the cell cycle, reaching a maximum shortly after the peak in transcription in each case.
We attempted to follow protein levels of FtsW, FtsI, FtsB, and FtsK by Flag-M2 epitope tagging the native copy of each gene. Given the apparent importance of FtsL in recruiting other factors (Table 1), we included FtsL-M2 in our analysis even though its transcript was apparently not strongly cell cycle regulated. We were unable to detect FtsW-M2, FtsI-M2 or FtsB-M2 by immunoblotting of whole cell lysates, so we could not follow their levels over the cell cycle. Immunoblot analysis of FtsK-M2 and
FtsL-M2 over the course of the cell cycle indicate that they are mildly cell cycle
48 regulated, reaching their highest levels in late pre-divisional cells. We conclude that a subset of division-site localized proteins are strongly regulated at the transcript and protein levels, notably FtsZ, FtsA, FtsQ, and KidO. FtsL, FtsK, FzlA, and DipM exhibit weak but reproducible cell cycle variation in protein levels.
49
50 Figure 10. Pairwise comparison among division-site localized proteins
Representative images of phase contrast and fluorescence microscopy of strains (A)
EG699 ( zapA-mCherry yfp-ftsE ), (B) Strain EG650 ( zapA-cfp mCherry-fzlA ), (C)
Strain EG658 ( venus-ftsI ftsK-mCherry ), (D) Strain EG689 ( venus-ftsI Pvan-mCherry- ftsW ), (E) Strain EG690 (P xylX–yfp–ftsA mCherry –ftsW ), (F) Strain EG692 (P xylX– yfp–ftsA zapA-mCherry ) at the indicated time after synchrony are shown. Percentage of cells with neither protein at midcell, both proteins at midcell, and either one or the other at midcell were quantified. (D) was grown in PYE media containing 0.5mM vanillate for 2 h prior synchrony. (E) and (F) were grown in PYE media containing
0.3% xylose for 2 h prior synchrony. In (A)–(F), at least 200 cells from two separate time-course experiments were analyzed. The accumulation of fluorescent fusions at midcell is indicated by arrows. SD represents standard deviations.
51
52 Figure 11. Localization of fluorescent fusions in mutant cells
Representative images of phase contrast and fluorescence microscopy of localization of indicated fluorescent fusions in indicated mutant cells. (A)-(J) Cells grown for the indicated amount of time in PYE with or without xylose inducer were examined. Two to four hours before analysis, expression of the fluorescent fusion of interest was induced by addition of 0.5mM vanillate. Arrows indicated the localized protein fusions in mutant cells. Double arrows indicated the partially localized protein fusions. Bar, 1
µm.
53
Figure 12. Transcriptional regulation of genes encoding division-site localized proteins
(A) Normalized abundance of transcript levels of the indicated genes over the cell cycle. (B) Immunoblots using antibodies against the indicated proteins of cell lysates from synchronized cells of the indicated strains grown in M2G.
54 Table 1. Summary of localization dependency among Caulobacter division-site localized proteins.
Localization of: Depletion FtsZ FtsA ZapA FzlA DipM FzlC MurG MreB TolQ FtsE FtsX FtsK FtsQ FtsL FtsB FtsI FtsW TipN FtsZ X D D D(8) D(3,5,6) D(8) D(7) D(10) D(4) D D D D D D(9) D D(11,12) FtsA LXL L(5,6) LL LLLLLLL ZapA LXLLL FzlA L(8)L XL L L LLLLL DipM LLXLLLL L TolA L(4) L(4) L L L*(4) Pal L(4) L(4) L L L(4) L L L*(4) FtsE XLLLL FtsX DXL FtsK L(2) L LLXLLL FtsL LL L L L D X D L*L* FtsN L(1) L(1) L*(3) L(4) L L(1) L L L(1) L FtsB LLLL FtsW L LLLLLLLX
D: diffused ; L: localized. L*: partially localized. (1) (Moll & Thanbichler, 2009); (2) (Wang et al. , 2006); (3) (Moll et al. , 2010a); (4) (Yeh et al. , 2010); (5) (Goley et al. , 2010a); (6) (Poggio et al. , 2010b); (7) (Aaron et al. , 2007); (8) (Goley et al. , 2010b); (9) (Costa et al. , 2008); (10) (Figge et al. , 2004); (11) (Lam et al. , 2006); (12) (Huitema et al. , 2006)
55 Table 2. Summary of features of Caulobacter divisome and division-site localized proteins
Gene Essential Mutant strains/Phenotype ftsZ Yes ftsZ depletion/smooth filaments (Quardokus et al. , 1996) ftsA Yes ftsA depletion, ftsA temperature-sensitive/filamentous cells with deep constrictions (Sackett et al. , 1998; Ohta et al. , 1997) zapA No zapA deletion/heterogeneous phenotype: slightly elongated, normal morphology cells fzlA Yes fzlA depletion/smooth filaments, ectopic poles (Goley et al. , 2010b) dipM No dipM deletion, dipM depletion/heterogeneous phenotype: filamentous, chaining, wider cells , outer membrane blebs (Goley et al. , 2010a; Poggio et al. , 2010b; Moll et al. , 2010a) fzlC No fzlC deletion/normal morphology (Goley et al. , 2010b) mreB Yes mreB depletion/lemon-shaped cells, outer membrane blebs (Gitai et al. , 2004; Figge et al. , 2004) tolA Yes tolA depletion/formation of chains of cells, outer membrane blebs (Yeh et al. , 2010) pal Yes pal depletion/formation of chains of cells, outer membrane blebs (Yeh et al. , 2010) ftsE Yes ftsE depletion/formation of chains of cells ftsX Yes ftsX depletion/formation of chains of cells ftsK Yes ftsK depletion/filamentous cells with slight constrictions (Wang et al. , 2006) ftsL Yes ftsL depletion/smooth filaments, rarely shallow constrictions ftsN Yes ftsN depletion/filamentous cells with slight constrictions (Moll & Thanbichler, 2009) ftsW Yes ftsW depletion/smooth filaments, rarely shallow constrictions ftsI Yes ftsI temperature-sensitive/filamentous cells with slight constrictions (Costa et al. , 2008; Ohta et al. , 1997) ftsB No ftsB deletion/normal morphology tipN No tipN deletion/pleiotropic polarity defects (Huitema et al. , 2006; Lam et al. , 2006)
56 Discussion
By systematically analyzing the temporal regulation of each divisomal component, we have shown that the assembly of divisome and division-site localized proteins in Caulobacter occurs in a series of stages (Fig. 8B). In combination with the results from the localization dependency along the pathway of cytokinetic ring assembly, we have established a framework to investigate the underlying molecular mechanisms of midcell recruitment of the divisome proteins during cellular growth and division.
The assembly of the divisome in Caulobacter occurs in a series of stages
Previous studies have shown that the timing of divisome assembly appears to follow a two-step assembly model in E. coli and B. subtilis (Gamba et al. ; Aarsman et al. ). In Caulobacter , high-resolution temporal studies allow us to refine the timing of at least seven stages of recruitment (Fig. 8B). First, FtsZ localizes to the incipient division plane at the time of chromosomal origin duplication and segregation to the cell poles. FtsZ binding proteins appear at midcell after the initial assembly of Z-ring near midcell. Next, the arrival of peptidoglycan remodeling proteins parallels the timing of medial peptidoglycan elongation, suggesting this group of proteins is involved in redirecting the location of peptidoglycan synthesis near midcell. The recruitment of TolQ to the division plane followed the arrival of peptidoglycan remodeling proteins, consistent with the role of Tol-Pal complex mediating outer membrane-peptidoglycan contacts during growth and division (Yeh et al. , 2010;
Anwari et al. , 2010). In contrast to other bacteria (Aarsman et al. , 2005; Gamba et al. ,
2009), however, we observed a significant delay in FtsA’s recruitment. Subsequently,
57 the arrival of five core divisome proteins to midcell followed the appearance of the
FtsA. Interestingly, visible invagination of the envelope, the switch from elongation to constriction phase, is coincident with the arrival of FtsW, suggesting that FtsW may have a role in localizing peptidoglyan remodeling machinery during cell constriction.
FtsW has been postulated as a lipid II flippase (Boyle et al. , 1997; Ikeda et al. , 1989), but its role during septal peptidoglycan synthesis is unclear.
The arrival of KidO at the midcell is slightly later than visible invagination of the envelope, consistent with the role of KidO contributing Z-ring disassembly during constriction (Radhakrishnan et al. , 2010). Lastly, TipN, a cell polarity marker protein, is recruited to the division site. Notably, the arrival of TipN at the midcell is concomitant with the dispersal of MreB from midcell. Previous studies have shown that TipN is required for localization of MreB to the cell division site (Lam et al. ,
2006). This result suggests that dynamic localization of TipN might play a role to trigger the switch from midcell to dispersed localization of MreB. In addition, it was recently reported that TipN affects the timing and position of FtsZ ring formation at the midcell by affecting the dynamics of MipZ-associated parS /ParB complex. TipN might play a key role in maintaining the directionality of chromosome segregation by interacting with ParA at the pole (Ptacin et al. , 2010; Schofield et al. , 2010). Thus, the timing of chromosome segregation and cell division is tightly coordinated by both
MipZ and TipN.
Transcriptional regulation
In Caulobacter , the delay between FtsZ-binding protein recruitment and core divisome assembly was quantified to be 25% of the cell cycle (Fig. 8B). The delay of
58 these two clusters are comparable to the ones found in E. coli and B. subtilis (Gamba et al. ; Aarsman et al. ), indicating that the assembly of core divisome components is triggered by a conserved mechanism. One distinctive feature of Caulobacter is that transcriptional regulation is governed by a complex transcriptional circuitry tightly linked to the developmental programs of regulatory proteins. This is true of cell division genes, and may play a key role in regulating divisome assembly and function.
Interestingly, FtsZ peaks earliest among all cell cycle-dependent genes we examined, whereas other divisome components reach their maximum expression between 40-
80% of the cell cycle. In addition, immunoblot analysis revealed that the levels of
FtsA (Martin et al. , 2004), FtsQ (Martin et al. , 2004), DipM (Goley et al. , 2010a),
FzlA (Goley et al. , 2010b), FtsL and FtsK (Fig. 12) reached their highest concentration in early or late predivisional cells, consistent with their functional role at the time of cell division.
Notably, expression of ftsA and ftsQ peaks closely in the pre-divisional cells
(Fig. 12). These findings raise the possibility that FtsA or FtsQ triggers the assembly of the core divisome components. It has previously been shown that FtsQ, FtsL and
FtsB interact in vivo (Buddelmeijer & Beckwith, 2004; Daniel et al. , 2006).
Furthermore, FtsQ was shown to be involved in maintaining the stability of FtsL
(Daniel & Errington, 2000). FtsQ is is an attractive candidate substrate, since none of the division-site localized proteins we tested required the presence of FtsA to localize to midcell (Table 1). However, we cannot rule out that the arrival of FtsA at the midcell is the key step in assembly of core cell division proteins, as the arrival of FtsA to midcell is right before the appearance of core divisome components.
59 Comparison with cytokinesis in other organisms
Pathways of localization dependency have been examined extensively in E. coli and B. subtilis ; however, these remain unclear in Caulobacter . From our localization dependency analysis, it appears that FtsL plays a key role in recruitment of FtsQ and FtsB to midcell and that FtsX is required to position FtsE to the division site. These results indicated that FtsQ/L/B and FtsE/X might function as complexes inside the cells, consistent with other bacterial divisomes (Buddelmeijer & Beckwith,
2004; Schmidt et al. , 2004; Garti-Levi et al. , 2008; Arends et al. , 2009). Notably, there are a number of differences in the localization dependencies of Caulobacter cells and mutants in E. coli . First, we have observed previously that DipM and Tol-Pal complex do not require FtsA for their localization to the division site (Goley et al. ,
2010a; Yeh et al. , 2010). Surprisingly, our analysis of the localization dependency showed that none of the divisome proteins recruited to the division plane was directly dependent on FtsA. In contrast to E. coli , however, cells resulting from FtsA depletion show constrictions. It is possible that assembly reactions downstream of FtsA can still operate in its absence, albeit imperfectly, leading to the formation of deeply constricted filamentous cells (Fig. 11B). Since FtsA is the primary factor for anchoring FtsZ filaments to the membrane in most bacteria, we hypothesized that an unknown protein might act redundantly with FtsA to anchor FtsZ to the membrane in
Caulobacter . Another key difference between Caulobacter and E. coli is the relationship between FtsI and FtsW. FtsI depends on FtsW for recruitment in E. coli , whereas FtsI arrives prior to FtsW at the division site in Caulobacter . Further analysis in Caulobacter of protein complexes and their localization dependencies showed that
60 FtsI does not require FtsW to localize to the midcell (Fig. 11J). It seems that
Caulobacter does not have a linear hierarchy of assembly of the divisome proteins, as in E. coli (Goehring & Beckwith, 2005). Although, further work is needed to identify protein-protein interactions among divisome components, these discrepancies represent the differences between distantly related species. Hence, it is important that several model species are investigated in parallel.
61 Table 3. Strains and plasmids
Strain or plasmid Relevant genotype/description Construction or Source CB15N synchronizable derivative of Caulobacter crescentus (Evinger & Agabian, 1977) CB15 EG208 CB15N mipZ::mipZ –cerulean Electroporation of pEG266 into CB15N EG420 CB15N mipZ::mipZ –cerulean vanA:: Pvan –ftsZ– Transduction of yfp vanA:: Pvan –ftsZ–yfp Kan R from MT196 (Thanbichler & Shapiro, 2006) into EG208 EG503 CB15N mipZ::mipZ –cerulean xylX:: PxylX –venus– Transduction of zapA xylX:: PxylX –venus–zapA Kan R from EG199 (Goley et al. , 2010b) into EG208 EG518 CB15N mipZ::mipZ –cerulean xylX:: PxylX– Transduction of mCherry–fzlA xylX:: PxylX–mCherry–fzlA Kan R from EG425 (Goley et al. , 2010b) into EG208 EG554 CB15N mipZ::mipZ –cerulean xylX:: PxylX–venus– Electroporation of pEG478 fzlC into EG208 EG492 CB15N mipZ::mipZ –cerulean xylX:: PxylX–venus– Transduction of ftsE xylX:: PxylX– venus–ftsE Kan R from EG744 into EG208 EG509 CB15N mipZ::mipZ –cerulean xylX:: PxylX–dipM– Transduction of mCherry xylX:: PxylX–dipM– mCherry Kan R EG261 (Werner et al. , 2009) into EG208 EG555 CB15N mipZ::mipZ –cerulean xylX:: PxylX–venus– Electroporation of pEG480 mreB into EG208 EG507 CB15N mipZ::mipZ –cerulean xylX:: PxylX– murG– Transduction of mCherry xylX:: PxylX– murG – mCherry Kan R from EG258 into EG208 EG517 CB15N mipZ::mipZ –cerulean xylX:: PxylX–tolQ– Transduction of yfp xylX:: PxylX–tolQ– venus Kan R from LS4517 (Yeh et al. , 2010) into EG208 EG495 CB15N mipZ::mipZ –cerulean xylX:: PxylX– yfp– Electroporation of pEG032 ftsA into EG208 EG651 CB15N mipZ::mipZ –cerulean xylX:: PxylX– venus– Electroporation of ftsN pAM041(Moll & Thanbichler, 2009) into EG208 EG508 CB15N mipZ::mipZ –cerulean xylX:: PxylX– ftsK– Electroporation of mCherry pGB022(Bowman GR , unpublished ) into EG208 EG497 CB15N mipZ::mipZ –cerulean xylX:: PxylX– yfp– Transduction of ftsQ xylX:: PxylX–yfp–ftsQ Kan R
62 Strain or plasmid Relevant genotype/description Construction or Source from EG085 into EG208 EG493 CB15N mipZ::mipZ –cerulean xylX:: PxylX– venus– Transduction of ftsL xylX:: PxylX– venus–ftsL Kan R from EG750 into EG208 EG502 CB15N mipZ::mipZ –cerulean xylX:: PxylX– venus– Transduction of ftsI xylX:: PxylX–venus–ftsI Kan R from EG051 into EG208 EG501 CB15N mipZ::mipZ –cerulean xylX:: PxylX– venus– Transduction of ftsW xylX:: PxylX–venus–ftsW Kan R from EG432 (Goley et al. , 2010b) into EG208 EG494 CB15N mipZ::mipZ –cerulean xylX:: PxylX– venus– Transduction of ftsB xylX:: PxylX– venus–ftsB Kan R from EG741 into EG208 EG647 CB15N mipZ::mipZ –cerulean xylX:: PxylX– kidO– Electroporation of pEG528 yfp into EG208 EG746 CB15N mipZ::mipZ –cerulean xylX:: PxylX– tipN– Electroporation of mCherry pJP173(Ptacin JL, unpublished ) into EG208 EG697 CB15N mipZ::mipZ –cerulean zapA::zapA– Electroporation of pEG555 mCherry into EG208 EG648 CB15N mipZ::mipZ –cerulean fzlA:: mCherry–fzlA Electroporation of pEG408 into EG208 EG652 CB15N mipZ::mipZ –cerulean ftsE:: venus–ftsE Electroporation of pEG556 into EG208 EG636 CB15N mipZ::mipZ –cerulean murG::murG – Transduction of mCherry murG::murG –mCherry Gent R from EG383 into EG208 EG698 CB15N mipZ::mipZ –cerulean ftsK::ftsK –mCherry Electroporation of pEG557 into EG208 EG638 CB15N mipZ::mipZ –cerulean ftsW::mCherry –ftsW Transduction of ftsW::mCherry –ftsW from EG123 into EG208 EG699 CB15N zapA::zapA–mCherry ftsE:: venus–ftsE Electroporation of pEG556 into EG645 EG654 CB15N zapA::zapA–cfp fzlA:: mCherry–fzlA Electroporation of pEG553 into EG384 EG658 CB15N ftsK::ftsK –mCherry ftsI:: venus–ftsI Electroporation of pEG557 into EG120 EG689 CB15N xylX:: PvanA–mCherry–ftsW ftsI::venus–ftsI Electroporation of pEG550 into EG120 EG692 CB15N xylX:: PxylX–yfp–ftsA zapA::zapA–mCherry Electroporation of pEG032 into EG645 EG690 CB15N xylX:: PxylX–yfp–ftsA ftsW::mCherry –ftsW Electroporation of pEG032 into EG123 LS3702 CB15N ∆ftsZ xylX ::P xylX –ftsZ (Wang et al. , 2001) EG666 CB15N ∆ftsZ xylX ::P xylX –ftsZ vanA:: Pvan– Electroporation of pEG543 mCherry–ftsA into LS3702
63 Strain or plasmid Relevant genotype/description Construction or Source EG700 CB15N ∆ftsZ xylX ::P xylX –ftsZ zapA::zapA– Electroporation of pEG555 mCherry into LS3702 EG701 CB15N ∆ftsZ xylX ::P xylX –ftsZ pPftsE –venus –ftsE Electroporation of pEG558 into LS3702 EG702 CB15N ∆ftsZ xylX ::P xylX –ftsZ pPftsL –venus –ftsL Electroporation of pEG559 into LS3702 EG703 CB15N ∆ftsZ xylX ::P xylX –ftsZ pPftsB –venus –ftsB Electroporation of pEG560 into LS3702 EG668 CB15N ∆ftsZ xylX ::P xylX –ftsZ vanA:: Pvan– Electroporation of pEG547 mCherry–ftsQ into LS3702 EG667 CB15N ∆ftsZ xylX ::P xylX –ftsZ vanA:: Pvan– Electroporation of pEG545 mCherry –ftsI into LS3702 EG669 CB15N ∆ftsZ xylX ::P xylX –ftsZ vanA:: Pvan– Electroporation of pEG549 mCherry –ftsW into LS3702 EG021 CB15N ∆ftsA xylX ::P xylX –ftsA Electroporation of pEG017 into EG005
EG085 CB15N xylX:: PxylX– yfp–ftsQ pP xylx -ftsZ-G109S (Goley et al. , 2010b) EG083 CB15N ∆ftsA xylX ::P xylX –ftsA Kan R Transduction of xylX ::P xylX –ftsA Kan R from EG150 into EG021 EG704 CB15N ∆ftsA xylX ::P xylX –ftsA zapA::zapA– Electroporation of pEG555 mCherry into EG021 EG705 CB15N ∆ftsA xylX ::P xylX –ftsA ftsK::ftsK –mCherry Electroporation of pEG557 into EG083 EG706 CB15N ∆ftsA xylX ::P xylX –ftsA pPftsE –venus –ftsE Electroporation of pEG558 into EG083 EG707 CB15N ∆ftsA xylX ::P xylX –ftsA pPftsL –venus –ftsL Electroporation of pEG559 into EG083 EG708 CB15N ∆ftsA xylX ::P xylX –ftsA pPftsB –venus –ftsB Electroporation of pEG560 into EG083 EG674 CB15N ∆ftsA xylX ::P xylX –ftsA vanA:: Pvan– Electroporation of pEG548 mCherry–ftsQ into EG021 EG673 CB15N ∆ftsA xylX ::P xylX –ftsA vanA:: Pvan– Electroporation of pEG546 mCherry–ftsI into EG021 EG675 CB15N ∆ftsA xylX ::P xylX –ftsA vanA:: Pvan– Electroporation of pEG550 mCherry–ftsW into EG021 EG456 CB15N ∆ftsA xylX ::P xylX –ftsA tipN::tipN –gfp Transduction of LS4527(Huitema et al. , 2006) into EG021 EG709 CB15N ∆zapA Electroporation of pEG562 into CB15N EG710 CB15N ∆zapA vanA::PvanA –ftsZ –cfp Electroporation of pEG094 into EG709 EG411 CB15N ∆zapA dipM::dipM –mCherry Transduction of dipM::dipM –mCherry Gent R from EG326 into EG709 EG711 CB15N ∆zapA murG::murG –mCherry Transduction of murG::murG –mCherry Gent R from EG383 into EG709 EG412 CB15N ∆zapA vanA::PvanA –mCherry–mreB Transduction of
64 Strain or plasmid Relevant genotype/description Construction or Source vanA::PvanA –mCherry– mreB Gent R from EG328 into EG709 EG312 CB15N ∆fzlA xylX ::P xylX –fzlA (Goley et al. , 2010b) EG662 CB15N ∆fzlA xylX ::P xylX –fzlA vanA::PvanA – Electroporation of pEG544 mCherry –ftsA into EG312 EG679 CB15N ∆fzlA xylX ::P xylX –fzlA murG::murG – Transduction of mCherry murG::murG –mCherry Gent R from EG383 into EG312 EG680 CB15N ∆fzlA xylX ::P xylX –fzlA dipM::dipM – Transduction of mCherry dipM::dipM –mCherry Gent R from EG326 into EG312 EG437 CB15N ∆fzlA xylX ::P xylX –fzlA pPftsE –venus –ftsE Electroporation of pEG558 into EG312 EG664 CB15N ∆fzlA xylX ::P xylX –fzlA vanA:: Pvan– Electroporation of pEG548 mCherry–ftsQ into EG312 EG663 CB15N ∆fzlA xylX ::P xylX –fzlA vanA:: Pvan– Electroporation of pEG546 mCherry–ftsI into EG312 EG665 CB15N ∆fzlA xylX ::P xylX –fzlA vanA:: Pvan– Electroporation of pEG550 mCherry–ftsW into EG312 EG353 CB15N ∆dipM xylX ::P xylX –dipM (Goley et al. , 2010a) EG364 CB15N ∆dipM xylX ::P xylX –dipM vanA:: Pvan–ftsZ– Electroporation of pEG094 cfp into EG353 EG402 CB15N ∆dipM xylX ::P xylX –dipM pP ftsQA–venus– Electroporation of pEG339 ftsA into EG353 EG470 CB15N ∆dipM xylX ::P xylX –dipM vanA:: Pvan– Electroporation of pEG392 mCherry–fzlA into EG353 EG483 CB15N ∆dipM xylX ::P xylX –dipM vanA:: Pvan– Electroporation of pEG385 mCherry– fzlC into EG353 EG485 CB15N ∆dipM xylX ::P xylX –dipM vanA:: Pvan– Electroporation of pEG388 murG –mCherry into EG353 EG542 CB15N ∆dipM xylX ::P xylX –dipM vanA:: Pvan– Transduction of mCherry–mreB vanA::PvanA –mCherry– mreB Gent R from EG328 into EG353 EG524 CB15N ∆dipM xylX ::P xylX –dipM pPtolQ –tolQ–yfp Electroporation of pEG561 into EG353 EG454 CB15N ∆dipM xylX ::P xylX –dipM vanA:: Pvan– Electroporation of pEG471 tipN –mCherry into EG353 LS4525 CB15N ∆tolA xylX ::P xylX-tolA (Yeh et al. , 2010) EG712 CB15N ∆tolA xylX ::P xylX-tolA zapA::zapA– Electroporation of pEG555 mCherry into LS4525 EG713 CB15N ∆tolA xylX ::P xylX-tolA vanA:: Pvan–dipM – Electroporation of pEG394 mCherry (Goley et al. , 2010a) into LS4525 LS4524 CB15N ∆pal xylX ::P xylX-pal (Yeh et al. , 2010) EG714 CB15N ∆pal xylX ::P xylX-pal zapA::zapA–mCherry Electroporation of pEG555 into LS4524 EG715 CB15N ∆pal xylX ::P xylX-pal vanA:: Pvan–dipM – Electroporation of pEG394 mCherry (Goley et al. , 2010a) into
65 Strain or plasmid Relevant genotype/description Construction or Source LS4524 EG716 CB15N ∆pal xylX ::P xylX-pal pPftsL –venus –ftsL Electroporation of pEG559 into LS4524 EG717 CB15N ∆pal xylX ::P xylX-pal pPftsB –venus –ftsB Electroporation of pEG560 into LS4524 EG718 CB15N ∆ftsE xylX ::P xylX-ftsE Electroporation of pEG563 into EG721 EG749 CB15N ∆ftsE xylX ::P xylX-ftsE vanA:: Pvan–ftsX – Electroporation of pEG564 mCherry into EG718 EG719 CB15N ∆ftsE xylX ::P xylX-ftsE pPftsL –venus –ftsL Electroporation of pEG559 into EG718 EG753 CB15N ∆ftsE xylX ::P xylX-ftsE vanA:: Pvan– Electroporation of pEG545 mCherry–ftsI into EG718 EG754 CB15N ∆ftsE xylX ::P xylX-ftsE vanA:: Pvan– Electroporation of pEG550 mCherry–ftsW into EG718 EG745 CB15N ∆ftsX xylX ::P xylX-ftsX Electroporation of pEG565 into EG751 EG747 CB15N ∆ftsX xylX ::P xylX-ftsX pPftsE –venus –ftsE Electroporation of pEG558 into EG751 EG748 CB15N ∆ftsX xylX ::P xylX-ftsX pPftsL –venus –ftsL Electroporation of pEG559 into EG751 LS4206 CB15N xylX::ftsK; ftsK:: ΩaacC4 (Wang et al. , 2006) EG558 CB15N xylX::ftsK; ftsK:: ΩaacC4 pPftsAQ- ftsA– Electroporation of pEG301 venus into into LS4206 EG722 CB15N xylX::ftsK; ftsK:: ΩaacC4 pPftsE –venus – Electroporation of pEG558 ftsE into LS4206 EG723 CB15N xylX::ftsK; ftsK:: ΩaacC4 pPftsL –venus – Electroporation of pEG559 ftsL into LS4206 EG724 CB15N xylX::ftsK; ftsK:: ΩaacC4 pPftsB –venus – Electroporation of pEG560 ftsB into LS4206 EG725 CB15N xylX::ftsK; ftsK:: ΩaacC4 pPtolQ –tolQ–yfp Electroporation of pEG561 into LS4206 EG726 CB15N xylX::ftsK; ftsK:: ΩaacC4 pPftsAQ –yfp– Electroporation of pEG397 ftsQ into LS4206 EG727 CB15N ∆ftsL xylX ::P xylX-ftsL Electroporation of pEG566 into EG752 EG728 CB15N ∆ftsL xylX ::P xylX–ftsL vanA:: Pvan–ftsZ-yfp Transduction of vanA:: Pvan –ftsZ–yfp Kan R from MT196 (Thanbichler & Shapiro, 2006) into EG727 EG670 CB15N ∆ftsL xylX ::P xylX–ftsL vanA:: Pvan– Electroporation of pEG544 mCherry–ftsA into EG727 EG720 CB15N ∆ftsL xylX ::P xylX-ftsL ftsK::ftsK –mCherry Electroporation of pEG557 into EG727 EG729 CB15N ∆ftsL xylX ::P xylX-ftsL pPtolQ –tolQ–yfp Electroporation of pEG561 into EG727 EG730 CB15N ∆ftsL xylX ::P xylX-ftsL pPftsE –venus –ftsE Electroporation of pEG558 into EG727 EG731 CB15N ∆ftsL xylX ::P xylX-ftsL pPftsB –venus –ftsB Electroporation of pEG560 into EG727 EG732 CB15N ∆ftsL xylX ::P xylX-ftsL pPftsAQ –yfp–ftsQ Electroporation of pEG397
66 Strain or plasmid Relevant genotype/description Construction or Source into EG727 EG671 CB15N ∆ftsL xylX ::P xylX-ftsL vanA:: Pvan– Electroporation of pEG545 mCherry–ftsI into EG727 EG672 CB15N ∆ftsL xylX ::P xylX-ftsL vanA:: Pvan– Electroporation of pEG550 mCherry–ftsW into EG727 AM52 CB15N ∆ftsN vanA ::P van-ftsN (Moll & Thanbichler, 2009) EG696 CB15N ∆ftsN vanA ::P van-ftsN murG::murG – Electroporation of pEG534 mCherry into AM52 EG683 CB15N ∆ftsN vanA ::P van-ftsN xylX ::P xylX –venus – Electroporation of pEG480 mreB into AM52 EG733 CB15N ∆ftsN vanA ::P van-ftsN pPftsE –venus –ftsE Electroporation of pEG558 into AM52 EG734 CB15N ∆ftsN vanA ::P van-ftsN pPftsL –venus –ftsL Electroporation of pEG559 into AM52 EG735 CB15N ∆ftsN vanA ::P van-ftsN pPftsB –venus –ftsB Electroporation of pEG560 into AM52 EG179 CB15N ∆ftsW xylX ::P xylX-ftsW Electroporation of pEG127 into EG010 EG546 CB15N ∆ftsW xylX ::P xylX-ftsW vanA:: Pvan–ftsZ-yfp Transduction of vanA:: Pvan–ftsZ-yfp Kan R from MT196 (Thanbichler & Shapiro, 2006) into EG179 EG659 CB15N ∆ftsW xylX ::P xylX-ftsW vanA:: Pvan– Electroporation of pEG543 mCherry– ftsA into EG179 EG736 CB15N ∆ftsW xylX ::P xylX-ftsW pPtolQ –tolQ–yfp Electroporation of pEG561 into EG179 EG737 CB15N ∆ftsW xylX ::P xylX-ftsW pPftsE –venus –ftsE Electroporation of pEG558 into EG179 EG738 CB15N ∆ftsW xylX ::P xylX-ftsW pPftsL –venus –ftsL Electroporation of pEG559 into EG179 EG739 CB15N ∆ftsW xylX ::P xylX-ftsW pPftsB –venus –ftsB Electroporation of pEG560 into EG179 EG661 CB15N ∆ftsW xylX ::P xylX-ftsW vanA:: Pvan– Electroporation of pEG547 mCherry–ftsQ into EG179 EG740 CB15N ∆ftsW xylX ::P xylX-ftsW ftsK::ftsK –mCherry Electroporation of pEG557 into EG179 EG660 CB15N ∆ftsW xylX ::P xylX-ftsW vanA:: Pvan– Electroporation of pEG545 mCherry–ftsI into EG179 EG682 CB15N ∆ftsB Electroporation of pEG567 into CB15N EG694 CB15N ∆ftsB vanA:: Pvan–mCherry–ftsA Electroporation of pEG543 into EG682 EG693 CB15N ∆ftsB pPftsL –venus –ftsL Electroporation of pEG559 into EG682 EG695 CB15N ∆ftsB vanA:: Pvan–mCherry–ftsI Electroporation of pEG545 into EG682 EG742 CB15N ftsK::ftsK –m2 Electroporation of pEG568 into CB15N EG743 CB15N ftsL::ftsL –m2 Electroporation of pEG569 into CB15N EG744 CB15N xylX:: PxylX–venus–ftsE Electroporation of pEG570
67 Strain or plasmid Relevant genotype/description Construction or Source into CB15N
EG258 CB15N xylX:: PxylX– murG–mCherry pP xylx -ftsZ- (Goley et al. , 2010b) G109S EG006 CB15N xylX:: PxylX– yfp–ftsA Electroporation of pEG032 into CB15N EG081 CB15N xylX:: PxylX– yfp–ftsQ Electroporation of pEG148 into CB15N EG750 CB15N xylX:: PxylX– venus–ftsL Electroporation of pEG571 into CB15N EG051 CB15N xylX:: PxylX– venus–ftsI Electroporation of pEG101 into CB15N EG741 CB15N xylX:: PxylX– venus–ftsB Electroporation of pEG572 into CB15N EG383 CB15N murG::murG –mCherry Electroporation of pEG410 into CB15N EG645 CB15N zapA::zapA–mCherry Electroporation of pEG555 into CB15N EG384 CB15N fzlA:: mCherry–fzlA Electroporation of pEG408 into CB15N EG123 CB15N ftsW::mCherry –ftsW Electroporation of pEG205 into CB15N EG005 CB15N xylX ::P xylX –ftsA Electroporation of pEG021 into CB15N EG326 CB15N dipM::dipM –mCherry Electroporation of pEG369 into CB15N EG328 CB15N vanA::PvanA –mCherry–mreB Electroporation of pEG386 into CB15N EG721 CB15N xylX ::P xylX –ftsE Electroporation of pEG573 into CB15N EG751 CB15N xylX ::P xylX-ftsX Electroporation of pEG574 into CB15N EG752 CB15N xylX ::P xylX-ftsL Electroporation of pEG575 into CB15N EG010 CB15N xylX ::P xylX-ftsW Electroporation of pEG037 into CB15N pChyC-4 For generation of C-terminally mCherry (Thanbichler et al. , 2007) tagged genes integrated at native gene locus pXVENN-2 For integration of N-terminal Venus fusions at xylX (Thanbichler et al. , 2007) locus pXYFPC-2 For integration of C-terminal YFP fusions at xylX (Thanbichler et al. , 2007) locus pEG478 fzlC inserted into pXVENN-2 This study pEG528 kidO inserted into pXYFPC-2 This study pEG555 zapA inserted into pChyC-4 This study pNPTS138 For gene replacement by double M.R.K. Alley, unpublished homologous recombination pEG266 mipZ –cerulean –mipZdownstream inserted into This study pNPTS138 pEG556 ftsE:: venus–ftsE inserted into pNPTS138 This study pEG408 fzlA:: mCherry–fzlA inserted into pNPTS138 This study pEG557 ftsK inserted into pChyC-4 This study pEG205 ftsW::mCherry –ftsW inserted into pNPTS138 This study
68 Strain or plasmid Relevant genotype/description Construction or Source pEG553 zapA::zapA–cfp (Hillson NJ, unpublished ) pVCHYN- For integration of N-terminal mCherry fusions at (Thanbichler et al. , 2007) 2 vanA locus pEG550 ftsW inserted into pVCHYN-2 This study pXYFPN-2 For integration of N-terminal YFP fusions at xylX (Thanbichler et al. , 2007) locus Kan R pEG032 ftsA inserted into pXYFPN-2 This study pVCHYN- For integration of N-terminal mCherry fusions at (Thanbichler et al. , 2007) 1 vanA locus Spec R pEG543 ftsA inserted into pVCHYN-1 This study pRVVENC- replicating plasmid for vanllilate-inducible C- (Thanbichler et al. , 2007) 4 terminal YFP fusions pEG558 pPftsE –venus –ftsE inserted into pRVVENC-4 This study pEG559 pPftsL –venus –ftsL inserted into p RVVENC-4 This study pEG560 pPftsB –venus –ftsB inserted into p RVVENC-4 This study pEG561 pPtolQ –tolQ–yfp inserted into pRVVENC-4 This study pEG547 ftsQ inserted into pVCHYN-1 This study pEG545 ftsI inserted into pVCHYN-1 This study pEG549 ftsW inserted into pVCHYN-1 This study pEG017 pNPTS138 ∆ftsA This study pXMCS-2 For generation of xylose dependent depletion strains (Thanbichler et al. , 2007) pEG150 ftsA inserted into pXMCS-2 This study pEG548 ftsQ inserted into pVCHYN-2 This study pEG546 ftsI inserted into pVCHYN-2 This study pEG562 pNPTS138 ∆zapA This study pVCFPC-1 For integration of C-terminal CFP fusions at vanA Thanbichler et al. , 2007) locus pEG094 ftsZ inserted into pVCFPC-1 This study pEG544 ftsA inserted into pVCHYN-2 This study pEG339 pP ftsQA–venus–ftsA Chlor R This study pEG385 fzlC inserted into pVCHYN-4 This study pEG388 murG inserted into pVCHYN-4 This study pEG471 tipN inserted into pVCHYC-4 This study pEG563 pNPTS138 ∆ftsE This study pVCHYC-2 For integration of C-terminal mCherry fusions at (Thanbichler et al. , 2007) vanA locus pEG564 ftsX inserted into pVmChyC-2 This study pEG565 pNPTS138 ∆ftsX This study pEG301 pPftsAQ- ftsA–venus inserted into pRVVENC-2 This study pEG397 Pqa-yfp-ftsQ inserted into pRVVENC-6 This study pEG566 pNPTS138 ∆ftsL This study pEG534 murG inserted into pCHYC-2 This study pEG480 xylX ::P xylX –venus – mreB inserted into pXVenN-2 This study pEG127 pNPTS138 ∆ftsW This study pEG567 pNPTS138 ∆ftsB This study pJM21 For integration of C-terminal M2 tag fusions (Alley et al. , 1993) pEG568 ftsK inserted into pJM21 This study pEG569 ftsL inserted into pJM21 This study pXVENC-2 For integration of C-terminal Venus fusions at xylX (Thanbichler et al. , 2007) locus
69 Strain or plasmid Relevant genotype/description Construction or Source pEG148 ftsQ inserted into pXYFPN-1 This study pEG101 ftsI inserted into pXVENN-2 This study pEG410 murG inserted into pVCHYC-4 This study pEG021 xylXupstream-ftsA-xylXdownstream inserted into This study pNPTS138 pEG369 dipM inserted into pCHYC-4 This study pEG386 mreB inserted into pVCHYN-4 This study pEG037 xylXupstream-ftsW-xylXdownstream inserted into This study pNPTS138 pEG570 ftsE inserted into pXVENC-2 This study pEG571 ftsL nserted into pXVENC-2 This study pEG572 ftsB nserted into pXVENC-2 This study pMT69 For integration of genes at the xylX locus (Thanbichler & Shapiro, 2006) pEG573 ftsE inserted into pMT69 This study pEG574 ftsX inserted into pMT69 This study pEG575 ftsL inserted into pMT69 This study
70
Chapter 3
The Caulobacter Tol-Pal complex is essential for outer membrane integrity and
is required for the completion of cell division and polar protein localization
Note: The majority of the information presented in this chapter was included in the publication:
Yeh YC , Comolli LR, Downing KH, Shapiro L, and McAdams HH. (2010) The Caulobacter
Tol-Pal complex is essential for outer membrane integrity and is required for the completion of cell division and polar protein localization. J. Bacteriol., 192(19):4847-58
71 Introduction
The cell envelope of Caulobacter crescentus and other Gram-negative bacteria consists of a peptidoglycan layer positioned between the inner membrane (IM) and the outer membrane (OM). Caulobacter cell division is implemented by the constrictive
IM-associated Z-ring, a polymeric structure of the highly conserved tubulin-like FtsZ protein positioned at the division plane. In Caulobacter , FtsZ localizes to the incipient division plane at the time of chromosomal origin duplication and segregation to the cell poles (Thanbichler & Shapiro, 2006; Moll & Thanbichler, 2009), well before a cell constriction is visible in the light microscope (Thanbichler & Shapiro, 2006). In the early stages of cell division, the inner and outer membranes are constricted simultaneously. However, late in the cell division process, the IM and peptidoglycan layers constrict faster, creating a separation of the inner and outer membranes near the division plane (Judd et al. , 2005). Fission of the IM and the peptidoglycan layer occurs about 20 minutes before cell division creating a cell containing two inner membrane and peptidoglycan-bound cytoplasmic compartments surrounded by a single continuous outer membrane (Judd et al. , 2005). Since Caulobacter OM invagination is temporally and spatially separated from peptidoglycan and IM invagination, separate mechanisms must drive the two processes. Hydrolysis of short membrane-bound FtsZ filaments that affects their curvature has been suggested as the mechanism for generation of the constrictive force for invagination of the IM (Osawa et al. , 2009; Li et al. , 2007a). However, the mechanism that implements the delayed constriction of the OM layer of the cell envelope is poorly understood.
72 The Tol-Pal complex of Gram-negative bacteria is widely conserved and plays multiple physiological roles, including maintaining OM interaction with the peptidoglycan, expressing lipopolysaccharide surface antigens and virulence factors, facilitating infection by filamentous DNA phage and reducing sensitivity to detergents (Davies & Reeves, 1975; Bernadac et al. , 1998; Gaspar et al. , 2000; Click
& Webster, 1997; Llamas et al. , 2003a; Llamas et al. , 2003b). In many bacteria, tol- pal mutants form cell chains with lateral membrane blebs in low osmolarity or high ionic strength medium, suggesting that Tol-Pal plays a role in completing cell division under conditions of membrane stress in these organisms (Bernadac et al. , 1998; Clavel et al. , 1996; Vianney et al. , 1994). In E. coli , TolA, TolQ and TolR are inner membrane proteins (Fig. 13A) and the TolA transmembrane domain interacts with the transmembrane domain of TolQ and TolR (Germon et al. , 1998; Derouiche et al. ,
1995). Pal, an abundant outer membrane lipoprotein, is thought to interact with the peptidoglycan layer through a conserved α-helical motif (Bouveret et al. , 1999;
Koebnik, 1995; Leduc et al. , 1992), while TolB is a periplasmic protein that interacts with Pal, the Lpp murein lipoprotein, and OmpA (Bouveret et al. , 1995; Ray et al. ,
2000; Clavel et al. , 1998; Weigand et al. , 1976). Thus, the Tol-Pal system bridges the three layers of the cell envelope via multiple interactions, including the interaction of the C-terminal periplasmic domain of TolA with Pal and TolB (Cascales et al. , 2001;
Walburger et al. , 2002; Dubuisson et al. , 2002; Germon et al. , 2001). In E. coli , the peptidoglycan-associated Lpp protein is a structural protein that is involved in maintaining the integrity of the cell envelope structure. Caulobacter does not have an
Lpp homolog. The Lpp protein is found only in enteric and endosymbiont bacteria
(string-db.org), suggesting that adaptation to survival in a high-osmolarity
73 environment may explain the significant differences between the E. coli and
Caulobacter Tol-Pal systems.
Here, we report that the Caulobacter Tol-Pal complex is concentrated at the division plane and following cell division it remains at the new poles, where it plays a role in maintaining the subcellular positioning of the TipN polar localization factor.
Caulobacter strains depleted of TolA, TolB, or Pal components of the essential Tol-
Pal complex exhibited surface bleb formation at both the division plane and the cell poles, and defects in invagination during the final stages of Caulobacter cell division.
74 Materials and methods
Bacterial strains, synchronization, growth conditions and cloning.
All Caulobacter strains were derived from CB15N and grown at 28°C in peptone yeast extract (PYE) with selected antibiotics. Strains and plasmids are listed in Table 4 and details on their construction are given in Table 3. For the Pal, TolA and TolB depletion constructs, cells were grown overnight in PYE medium containing 0.3% xylose (PYEX), washed with PYE medium, and then re-suspended in PYE medium containing 0.2% glucose (PYEG).
Fluorescence Microscopy.
Cells were immobilized using a thin layer of agarose in M2G medium. For localization studies, 0.3% xylose or 0.5mM vanillate (pH 7.5) were used to induce expression of fluorescent protein fusions from the xylX or vanA promoters for at least
2 h, respectively. 2 µg/ml N-(3-triethylammoniumpropyl)-4-(6-(4-
(diethylamino)phenyl)hexatrienyl)pyridinium dibromide (FM4-64) was added to the agarose to fluorescently label membranes (Molecular Probes). Differential interference contrast (DIC) and fluorescence microscopy images were obtained using a Leica DM
6000 B microscope with a HCX PL APO 100x/1.40 Oil PH3 CS objective,
Hamamatsu 16 EM-CCD C9100 camera, and a custom-designed microscope control and image analysis software package called KAMS (Christen et al. , 2010).
Scanning electron microscopy.
Samples were fixed in 2% glutaraldehyde and 4% formaldehyde in a 0.1M sodium cacodylate buffer (pH 7.3) overnight, allowed to adhere onto poly-L-lysine coated 12 mm coverslips, and moved to 4 oC. Subsequently, samples were dehydrated with 50%,
75 70%, 95% and 100% ethanol, for 10 min each, and then coated with gold-palladium.
All SEM images were taken on a Hitachi S-3400N VP-SEM.
CryoEM.
Cells were grown overnight in liquid PYE media containing 0.3% xylose at 28°C, washed with un-supplemented PYE medium, and then re-suspended in PYE medium containing 0.2% glucose for 11 h until reaching an OD 660 of approximately 0.4.
Aliquots of 5 µl were then taken directly from the culture and placed onto glow- discharged carbon grids (Ted Pella 01881). The grids were blotted, and plunged as described previously (Judd et al. , 2005). All cryoEM images were acquired with a
JEOL-3100-FEF electron microscope at the Lawrence Berkeley National Laboratory.
Co-immunoprecipitation experiments.
Co-immunoprecipitation was performed as previously described (Iniesta et al. , 2006).
Strains LS4536 and LS4529 were grown in PYE media containing 0.3% xylose and were harvested in log phase. LS4538 and wild type strains were induced with 0.3% xylose for 3 h before being harvested. 1 liter of cell culture in PYE medium was washed in Co-IP buffer (20 mM Hepes pH 7.5, 100 mM NaCl and 20% glycerol), treated with the membrane permeable cross linking agent, formaldehyde (Fisher
Biotech), at a final concentration of 1%, and then quenched with 0.125M glycine.
Cells were lysed by passage through a French Press at 15000 psi three times. The cell lysate was then incubated with anti-FLAG conjugated beads (FLAGIPT-1 kit, Sigma) overnight. Subsequently, the beads were washed, bound proteins were eluted by incubating with 3 x FLAG peptides, and the supernatant was collected.
76 Immunoblot analysis.
Immunoblots were performed as previously described (Bowman et al. , 2008; Wheeler
& Shapiro, 1999) and the blots were developed chemiluminescently. Western blotting to monitor DivJ, PleC, FtsZ and PopZ was performed as described previously
(Bowman et al. , 2008; Mohl et al. , 2001; Wheeler & Shapiro, 1999). Antibody dilutions: GFP (1/1000; Roche), Pal (1/50,000) and Flag monoclonal (1/10,000;
Sigma).
77 Results
Genomic organization of the Caulobacter tol-pal gene cluster
The cell envelope configuration of the protein components of the E. coli and
Caulobacter Tol-Pal complexes are shown schematically in Fig. 13A, and the genomic organization of Caulobacter tol-pal homologs (CC3233-CC3229) is shown in Fig.
13B. The Caulobacter gene CC3231, originally predicted to encode a hypothetical protein (Nierman et al. , 2001), shares only 27% identity with the tolA from E. coli.
However, tolA represents a highly divergent gene family with a strongly species- dependent sequence (Sturgis, 2001). The predicted TolA protein contains a signature single transmembrane helix at the N-terminus and a large periplasmic domain at the C- terminus. As observed in other Gram-negative bacteria, the tolA gene is adjacent to the predicted tolQ, tolR, and tolB homologues. Based on its chromosome location and conserved structure features, we designated Caulobacter gene CC3231 as tolA .
Microarray assays of the gene expression temporal pattern over the cell cycle indicate that tolA, tolB, tolR , and tolQ are not cell cycle regulated (Fig. 13C) (McGrath et al. ,
2007). In contrast, pal is cell cycle regulated with high transcript levels in swarmer cells that drop thereafter (Fig. 13C) (McGrath et al. , 2007). To determine if the Pal protein level is also cell cycle dependent, we raised a polyclonal antibody against purified recombinant Pal and used the antibody to probe synchronized wild type
CB15N cell lysate samples taken at various times after synchronization. This immunoblot analysis revealed that the level of Pal was roughly constant over the cell cycle (Fig. 13D and 13E). The quality of the synchrony was monitored by immunoblot analysis of the FtsZ protein (Quardokus et al. , 1996; Thanbichler & Shapiro, 2006)
78 which was low in swarmer cells, increased in stalked cells, and peaked at the onset of cell division as expected (Fig. 13D and 13E).
Proteins of the Tol-Pal cluster accumulate at the incipient division site
To examine the cellular localization of the proteins in the Tol-Pal complex, we constructed fusions of each gene to a fluorescent protein coding sequence and placed the fusion construct under the control of a xylose-inducible promoter at the chromosomal xylX locus. In each case, the wild type copy of the gene was retained at its native chromosomal site, creating a merodiploid strain. We constructed a yfp fusion to the 3’ end of tolQ , gfp fusions to the 5’ end of tolR and tolA , and mCherry fusions to the 5’ end of tolB and to the 3’ end of pal . Individual strains, each containing one of the five fusion constructs, were incubated in the presence of 0.3% xylose to induce the expression of the fluorescently-tagged proteins. All of the fusion proteins exhibited significant accumulation at the division plane in predivisional cells and were retained at the new poles of the daughter swarmer and stalked cells (Fig. 14A). In contrast to E. coli (Gerding et al. , 2007), however, a significant fraction of the components of the
Tol-Pal complex also displayed a punctate (TolQ, TolR and TolA) or peripheral localization pattern (TolB and Pal) throughout the cell envelope, including the stalk.
To investigate the localization dynamics of TolA, we followed the subcellular positioning of GFP-TolA over the course of the cell cycle. We isolated swarmer cells of strain LS4519 (P xyl-gfp-tolA ) and imaged the cells by fluorescent microscopy as they grew synchronously on agarose pads supplemented with 0.3% xylose (Fig. 14B).
79
Figure 13. The Caulobacter crescentus tol-pal gene cluster (A) Predicted organization of the components of the Tol-Pal complex in the E. coli (Lloubes et al. ,
2001) and Caulobacter cell envelopes. TolQ, TolR and TolA are integral IM proteins, and Pal is an OM protein that interacts with the periplasmic TolB protein. The
Caulobacter TipN polar marker is an inner membrane (IM) protein (Lam et al. , 2006;
Huitema et al. , 2006). (B) Schematic of the gene organization of the Caulobacter tol- pal components. Arrows indicate the direction of transcription and putative position of promoters. The +1 transcriptional start site is at -49 of the tolQ coding sequence, as indicated (McGrath et al. , 2007). (C) mRNA expression patterns of genes encoding the components of the Tol-Pal complex over the course of a cell cycle. pal expression peaks in the swarmer cell and drops thereafter, while the expression of the other genes is not cell cycle dependent. (D) Western blot analysis of relative Pal protein levels
80 (upper panel, Pal protein indicated by arrows) during the cell cycle starting with a synchronous population of wild type swarmer cells. The FtsZ protein (lower panel) is shown as a loading control and quality control for the synchrony. (E) Normalized abundance of Pal and FtsZ protein level over the course of the cell cycle. The Pal protein level does not exhibit significant changes during the course of the cell cycle.
81 Following separation of the two daughter cells, GFP-TolA remained at the new poles. When the swarmer cell differentiated into a stalked cell, an additional focus appeared at midcell. Later, predivisional cells had a single focus at the cell division site. We demonstrated that the GFP-TolA fusion protein was functional by transducing the P xyl-gfp-tolA allele into the TolA depletion strain, LS4525, replacing the wild type tolA gene at the xylX locus with gfp-tolA to create strain LS4522. This strain has the native tolA gene deleted, and the only full length copy of the gene is gfp-tolA . Upon induction with xylose, the localization of GFP-TolA was similar to that observed in wild type cells (Fig. 15A), and the cellular morphology and growth rate was indistinguishable from wild-type. Thus, the GFP-TolA fusion protein alone complemented the TolA deletion phenotype. The Pal-mCherry fusion also fully complemented the Pal depletion phenotype (Fig. 15A). To verify that the expected fusion proteins were produced, cell lysates from the above strains were probed with antibodies that recognize GFP and Pal (Fig. 15B and 15C).
To determine the temporal order of the Tol-Pal complex and divisome assembly, we compared the localization dynamics of TolA and TolQ with those of
FtsZ, FtsA and FtsI, using in vivo time-course microscopy. The localization timing of
FtsZ–Venus, Venus–FtsA, Venus–FtsI, GFP–TolA and TolQ–YFP near midcell (Fig.
14C) shows that both TolQ and TolA localize to midcell after FtsZ is assembled at the division plane, well before Venus–FtsA and Venus–FtsI.
82
83 Figure 14. Subcellular localization of the proteins encoded by the tol-pal cluster.
(A) Fusions of TolQ, TolR, TolA, TolB and Pal to GFP, YFP or mCherry, at either N- or C-termini, were generated and their cellular location was observed by fluorescence microscopy. LS4517 (P xylX -tolQ-yfp ), LS4518 (P xylX -gfp-tolR ), LS4519 (P xylX - gfp-tolA ), LS4520 (P xylX -mCherry-tolB ) and LS4521 (P xylX -pal-mCherry ) strains were each incubated in the presence of 0.3% xylose for 2 h to induce the expression of the fluorescently-tagged fusions in a merodiploid containing the wild type gene. All of the fusion proteins tested localized to both the cell pole (upper panel) and to the division plane (lower panel). (B) A synchronized LS4519 swarmer cell population, producing GFP-TolA, was suspended in PYE media containing 0.3% xylose and allowed to proceed through the cell cycle. Samples were visualized at 20 min intervals by phase contrast and fluorescence microscopy. A schematic of Caulobacte r cells showing the dynamic localization of TolA as a function of the cell cycle is shown below the micrographs. The accumulation of GFP-TolA at the division plane is indicated by arrows. (C) Cells of strains EG052 ( PxylX-ftsZ–venus ), EG055 ( PxylX– venus–ftsA ), EG051 ( PxylX–venus–ftsI ), LS4517 ( PxylX–tolQ–venus ) and LS4519
(P xylX–gfp–tolA ) were grown to exponential phase in PYE medium, induced for 1 h with 0.3% xylose, synchronized and re-suspended in PYE medium supplemented with
0.3% xylose. Cells were withdrawn from the cultures at 10 min intervals and visualized by phase contrast and fluorescence microscopy to determine if the fluorescently labeled proteins were positioned at the incipient division plane. At least
150 cells were analyzed per time point. (D) Localization of YFP-TolA in strain
LS4546 depleted of FtsZ for 3 h (-FtsZ) and with FtsZ depleted for 3 h then with FtsZ added back by incubation in the presence of the xylose inducer for 0.5 h (add FtsZ
84 back). In both cases, cultures were incubated in the presence of 0.5mM vanillate to induce the expression of y fp-tolA. Localization of YFP-TolA in strain LS4547 depleted of FtsA after growth in PYE medium with glucose (PYEG) for 8 h. White arrows indicate foci of YFP-TolA. YFP-TolA requires FtsZ, but not FtsA, to form fluorescent foci. (E) Fluorescence microscopy of Pal depletion strains carrying plasmid-borne P van -tolQ-yfp, Pvan-yfp-tolR or Pvan-yfp-tolA incubated in the absence of xylose (-Pal). Cells were grown in PYE medium with xylose (PYEX), washed and incubated in PYEG for 11 h to deplete Pal. TolA depletion strains bearing the plasmids P van -tolQ-yfp or Pvan-yfp-tolR were incubated in the absence of xylose (-
TolA) for 11 h. A TolA depletion strain, with P van-pal-mCherry integrated at the chromosomal vanA locus, was incubated in the presence of 0.5mM vanillate to induce
Pal-mCherry and in the absence of xylose for 11 h (-TolA). Cells were imaged using phase contrast and fluorescence microscopy.
85
Figure 15. gfp ‐‐‐tolA and pal ‐‐‐mCherry can complement deletion mutants of tolA and pal respectively. (A)When strain LS4522 was grown in the presence of xylose, inducing expression of gfp-tolA , localization of the GFP-TolA protein was similar to that of wild-type. The viability of the strain demonstrates that the GFP-TolA fusion protein alone complemented the TolA depletion phenotype. The Pal-mCherry fusion also fully complemented the Pal depletion phenotype. (B) The GFP-TolA fusion proteins were characterized by Western blot analysis using an anti-GFP antibody.
There was no significant clipping of the fusion releases observed. (C) The Pal- mCherry fusion proteins were characterized by Western blot analysis using an anti-Pal antibody. There is a small amount of degradation that results in clipping of the Pal- mCherry, while full-length Pal-mCherry is the major form.
86 Thetime of the localization pattern at the division plane
Since the division plane localization of GFP-TolA and TolQ-YFP occurs later than FtsZ, we asked if the localization of the components of the Tol-Pal complex is dependent on the prior localization of FtsZ. Accordingly, we constructed a strain
(LS4546) with P van -yfp-tolA on a high copy plasmid in a background in which the only functional copy of ftsZ is under the control of the xylose-inducible promoter.
Cells grown in the absence of xylose were depleted of FtsZ (data not shown) and formed smooth filamentous cells. In these cells, YFP-TolA was diffuse (Fig. 14D).
FtsZ was then allowed to accumulate in these cells by incubating them in PYE containing 0.3% xylose for 0.5 h. The localization of TolA to the division plane was restored upon FtsZ accumulation (arrows, Fig. 14D), indicating that TolA requires
FtsZ for its positioning at the division site. As the recruitment of FtsA to the division site is significantly later than TolA and TolQ (Fig. 14C), we examined the localization patterns of YFP–TolA in cells that had been depleted of FtsA. Interestingly, the recruitment of YFP-TolA to the divisome was independent of FtsA, a protein reported to be immediately downstream of FtsZ in the E. coli assembly hierarchy (Goehring &
Beckwith, 2005). YFP-TolA localized to discrete bands in filamentous cells depleted of FtsA (arrows, Fig. 14D). Similar to the localization pattern observed for YFP-TolA, all other proteins of Tol-Pal complex were also robustly recruited to the division site in the absence of FtsA (data not shown), arguing that FtsA is not required for Tol-Pal localization in Caulobacter . We also examined the localization pattern of YFP–TolA in cells depleted of FtsN, the last known essential protein recruited to divisome in the
E. coli assembly hierarchy (Addinall et al. , 1997). YFP-TolA foci were observed at
87 several sites in the FtsN-depleted filamentous cells (arrows, Fig. 14D), suggesting that the recruitment of TolA to the divisome occurs independently of FtsN.
In E. coli , division site localization of TolQ and TolA is independent of any of the other four Tol–Pal proteins, while TolR requires TolQ, and possibly TolB, and Pal requires TolA to accumulate at the division plane (Gerding et al. , 2007). To determine which of the remaining components of the Tol-Pal complex were able to localize to the division site in the Caulobacter tolA and pal depletion strains, we examined the localization patterns of TolQ–YFP, YFP–TolR, Pal-mCherry and YFP–TolA in the tolA and pal mutant strains. In the absence of Pal, TolQ–YFP, YFP–TolR and YFP–
TolA were localized to the division plane (arrows, Fig. 14E), indicating that Pal is not required to position the other proteins of the Tol-Pal complex to the division site. Both
TolQ–YFP and YFP–TolR were localized to the division site in the absence of TolA
(arrows, Fig. 14E), but Pal-mCherry failed to localize to a subcellular site (Fig. 14E).
Thus, Pal requires TolA for its localization to the division plane.
TolA and Pal are required for the polar positioning of TipN and PleC
Since the Tol-Pal complex remains at the new poles in swarmer and stalked daughter cells after cell division, we asked if transmembrane polar proteins are mislocalized in mutant strains lacking components of the Tol-Pal system. We constructed Pal or TolA depletion strains containing either tipN-gfp , pleC-gfp , divJ-yfp , or Pvan::popZ-yfp in place of each wild-type gene. In wild type cells, TipN-GFP accumulates at the cell division plane and then, following division, it remains as a single focus at the new poles of the daughter cells (Lam et al. , 2006; Huitema et al. ,
2006). In cells depleted of Pal by growth in PYEG for 11 h, 46% of the cells (n = 297)
88 exhibited TipN-GFP foci that were aberrantly placed throughout the cell (Fig. 16A).
Similarly, when TolA was depleted by growth in PYEG for 11 h, we observed an aberrant placement of TipN-GFP (Fig. 16B). Since TipN contributes to the polar placement of the PleC histidine kinase (Lam et al. , 2006), we examined the localization of PleC in the absence of either Pal or TolA. In a localization pattern similar to the pattern observed for TipN-GFP in cells depleted of either Pal or TolA,
PleC-GFP foci were aberrantly positioned throughout the cell (Fig. 16A and 16B).
The cellular localization of two additional polar proteins, the DivJ-YFP histidine kinase (Wheeler & Shapiro, 1999) and the membrane associated PopZ-YFP centromere anchor (Bowman et al. , 2008; Ebersbach et al. , 2008a), was maintained in depletion strains of Pal and TolA (Fig. 16A and 16B). Thus, membrane integrity mediated by the Tol-Pal complex is specifically required to localize a subset of polar proteins. Of the polar proteins imaged, only the TipN protein first accumulates at the division plane and then subsequently at the new cell poles in wild type strains, suggesting that in the TolA and Pal depletion strains TipN is not properly positioned at the division plane and consequently is mislocalized at the cell poles. After 11 h of either TolA or Pal depletion, 80% of the cells remained viable, and in the TolA and
Pal depletion strains, there were no significant changes in the levels of either TipN or
PleC after 11 h of growth in the absence of xylose (Fig. 16C). Thus, the mislocalization of TipN in the depletion strains was not due to effects in dying cells or elimination of TipN. We examined the localization of FtsZ and divisome components
ZapA and FtsL in cells depleted of TolA or Pal, and all localized normally in these strains (data not shown).
89 To determine if TipN interacts with TolA and Pal, we used strain LS4536 ( tipN-gfp
∆tolA PxylX-tolA-m2 ) in which the native tolA coding sequence was deleted and replaced with a chromosomal xylose inducible tolA gene carrying a C-terminal Flag
M2 tag. When strain LS4536 was grown in the presence of xylose, inducing expression of tolA-m2 , cell morphology was similar to that of the wild type (data not shown). As a control, an isogenic strain tipN-gfp ∆tolA PxylX-tolA (LS4529) was generated. Formaldehyde, a membrane permeable cross-linking agent, was added prior to cell lysis, then cultures of LS4529, without the M2 epitope tag and LS4536 carrying
TolA-M2, were immunoprecipitated with anti-M2 coupled beads. Analysis of the purified products by western blot showed that TipN-GFP, FtsZ and Pal interacted directly or indirectly with the TolA complex, while neither DivJ, PleC, nor PopZ interacted with TolA-M2 (Fig. 17A). PleC was found not to interact with TolA-M2
(Fig. 17A), suggesting the observed mislocalization of PleC in Pal and TolA mutants is likely an indirect effect. To obtain additional evidence for an interaction between
TipN and components of the Tol-Pal complex, we immunoprecipitated TipN-M2 from a strain bearing xylose-inducible tipN-m2 at the chromosomal xylX locus (LS4538).
Immunoblots of the immunoprecipitated sample were then probed with anti-Pal or anti-PopZ antibodies (Fig. 17B). A band corresponding to the size of Pal was detected in the TipN-M2 immunoprecipitated sample, while a control protein, PopZ, was not.
These results, together with the localization data, support the conclusion that TipN directly or indirectly interacts with both TolA and Pal.
The localization dependencies described in this and the preceding section are diagrammed in Figure 16D.
90 91 Figure 16. Mislocalization of the TipN-GFP and PleC-GFP polar proteins in the absence of the Pal or TolA proteins. (A) Fluorescence microscopy of Pal depletion strains bearing TipN-GFP, PleC-GFP, DivJ-YFP or PopZ-YFP grown in the presence of xylose to induce Pal accumulation or in the presence of glucose to deplete Pal.
Strain LS4528 ( tipN-gfp ; ∆pal PxylX-pal ) was imaged after either 9 or 11 h of incubation in the absence of xylose to deplete Pal. The upper panels show the sub- cellular location of TipN-GFP, PleC-GFP, and DivJ-YFP in cells grown in the presence of xylose to induce Pal, as compared to cells shown in the lower panels incubated with glucose to deplete Pal. Strain LS4534 ( ∆pal PxylX-pal ; Pvan-popZ-yfp ) was incubated for 1 h in the presence of 0.5 mM vanillate to induce PopZ-YFP in the presence of either xylose (+Pal) or glucose (- Pal). (B) The same fusion proteins were examined by fluorescence microscopy in TolA-depletion strains LS4529 ( tipN-gfp ;
∆tolA PxylX-tolA ), LS4531 ( pleC-gfp ; ∆tolA PxylX-tolA ), LS4533 ( divJ-yfp ; ∆tolA
PxylX-tolA ), LS4535 ( ∆tolA PxylX-tolA Pvan-popZ-yfp ) incubated for 11 h in the presence of either xylose (+TolA) or glucose (-TolA). Arrows indicate polar localization of DivJ-YFP and PopZ-YFP in the absence of either Pal or TolA. (C)
Immunoblot analysis of Pal, TipN-GFP and PleC levels in cell extracts of TolA depletion and Pal depletion cultures. Both TipN-GFP and PleC levels were not affected after 11 h of growth in the absence of xylose. (D) The predicted localization dependency pathway derived from experiments shown in this Figure and in Figure 14.
92
Figure 17. The Tol-Pal complex interacts with TipN-GFP. (A) In vivo coimmunoprecipitation of TolA-M2 in strain LS4536 (tipN-gfp tolA PxylX-tolA-m2 ) and LS4529 ( tipN-gfp tolA PxylX-tolA ). Western blots of whole-cell extracts (lysate) and eluted samples were probed with the indicated antibodies (anti-M2 to probe TolA-
M2 and anti-GFP to probe TipN-GFP, anti-Pal, anti-FtsZ, anti-DivJ, anti-PleC, and anti-PopZ). TipN, FtsZ, and Pal, but not DivJ, PleC, or PopZ, were observed to interact, directly or indirectly, with TolA-M2. (B) Coimmunoprecipitation of Pal with
TipN-M2 in strain LS4538 (P xylX-tipN-m2 ) and in wild-type cells. Western blots of eluted samples were probed with anti-M2 to probe TipN-M2, anti-Pal, and anti-PopZ.
Pal but not PopZ was observed to interact with TipN-M2.
93 The Pal, TolA and TolB components of the Tol-Pal complex are essential, and are required for OM invagination during cell division
Phenotype of a Pal depletion strain .
It was not possible to disrupt the pal gene without complementing with a copy of the gene, suggesting that pal is essential in Caulobacter, as recently reported
(Anwari et al. , 2010). We constructed a pal depletion strain, LS4524, by deleting the majority of the native pal coding region (aa 13 to177) and placing a full length pal gene under the control of a xylose inducible promoter at the chromosomal xylX locus
(Fig. 18A). Immunoblot analysis showed that the levels of Pal were reduced in Pal depletion strain LS4524, after 9 h of growth in the absence of xylose (Fig. 18B), and this coincided with a decline in colony forming unit (CFU).
Depletion of Pal by growth of strain LS4524 in PYEG for 10 h resulted in the accumulation of chains of cells (Fig. 19A, right), unlike strain LS4524 grown in the presence of xylose to induce pal expression (Fig. 19A, left), suggesting that Pal is required for the completion of cell separation. To determine the effect of Pal depletion on membrane invagination, LS4524 cells were stained with the lipophilic fluorescent styryl dye, FM4-64, and examined by fluorescence microscopy. Strain LS4524 grown in the presence of xylose appeared similar to wild type (Fig. 19A, left), but those incubated with glucose to deplete Pal initially grew as chains and then exhibited extensive membrane blebs (blebbing phenotype: 88.6%, n=120 cells), predominantly at division sites and cell poles (Fig. 19A, right).
94 Scanning electron microscopy (SEM) images of cells depleted of Pal (grown in
PYEG for 12 h) had large, irregular blebs both laterally and at the cell poles which were not observed in wild type cells or the Pal depletion strain grown in the presence of the xylose inducer (Fig. 19C). In the absence of the xylose inducer, cell growth arrested and a portion of the cells lysed, while in the presence of inducer, both growth rate and cell morphology were indistinguishable from wild type (Fig. 19B).
To visualize the IM and OM in the presence and absence of Pal, plunge-frozen cells of wild type and the Pal depletion strain LS4524 were observed by cryo electron microscopy (cryoEM). A cryoEM image of a dividing wild type cell clearly shows the
IM, peptidoglycan (PG), OM, and S-layer, and the fissioned IM and PG layers that create two cellular compartments within an OM envelope (Fig. 19C-left). When strain
LS4524 was grown in xylose to induce Pal, the constriction and the space between the
IM and OM in the region of the division plane was indistinguishable from wild type cells (data not shown). However, in the absence of Pal, OM invagination was disrupted and daughter cell separation did not complete in many cases yielding chains of cells. In these cells, large OM membrane extrusions were observed at the division site (Fig. 19C-center). Notably, the OM was also abnormally separated from the IM at the cell poles (Fig. 19C - right). Cumulatively, these results demonstrate that
Caulobacter Pal is required for the maintenance of OM-peptidoglycan integrity and for mediating invagination of the OM during cytokinesis.
95 Phenotypes of TolA and TolB depletion strains .
We constructed a tolA depletion strain, LS4525, with a chromosomal deletion of tolA (Fig. 20A), by deleting the native tolA coding region (aa 13 to 259) and integrating a full-length tolA gene at the chromosomal xylX locus. A tolB depletion strain was constructed by incorporating a truncated version of tolB at the native chromosomal site and a second, full-length copy under control of the xylose inducible promoter, creating the strain LS4526 (Fig. 20A). Strains LS4525 and LS4526 exhibited impaired cell separation and OM defects (Fig. 20C and 20D) and remained viable only in the presence of the xylose inducer (Fig. 20B). Thus, TolA and TolB are essential for viability. It was reported previously that genes encoding two other components of the Caulobacter Tol-Pal complex, tolQ and tolR , are essential
(Eisenbeis et al. , 2008).
Scanning electron microscopy of strains LS4525 and LS4526 incubated in the presence (+ TolA or + TolB) or absence (- TolA or - TolB) of the xylose inducer showed that in the depletion of either TolA or TolB, envelope blebs appeared at the poles of both strains, and cells exhibited cell division and polar structural defects (Fig.
21A and 21B). Although cells depleted of TolB exhibited outer membrane blebs primarily at the site of cell division and the cell poles, cells depleted of TolA also exhibited extensive OM blebs at lateral surfaces of the cell. CryoEM images of these strains, grown in the presence of glucose for 11 h to deplete TolA or TolB, exhibited disruptions of the OM (Fig. 21C and 21D). OM invagination was impaired with large
OM bulges visible at the cell division plane and also at the cell poles. Notably, depletion of either TolA or TolB caused the outer membrane to bleb outwards, while
96 the cytoplasmic membrane remained intact. The absence of TolA induced more extensive lateral OM defects (over 82% of the cells, N=35 cells) than in either Pal or
TolB depletion strains (Fig. 19C). Cumulatively, the phenotypes of strains depleted of either Pal, TolA or TolB suggest that these proteins all participate in both cell division and polar membrane integrity.
97
Figure 18. Construction of Pal depletion strain. (A) A schematic diagram of the construction of Pal depletion strain, LS4524. An in frame deletion of pal was constructed by deleting base pairs 37-531, out of a total of 567 bp of coding sequence, while the full-length pal gene was integrated at the chromosomal xylX locus. (B) CFU for strain LS4524 grown in PYEX, washed and then incubated in the presence of either PYEG or PYEX. Cell viability decreased after 9 h of growth in PYEG
(indicated by arrows). LS4524 was grown in PYEX, washed and then suspended in
PYEG (t = 0 h). Protein levels were measured by western blot analysis of samples taken at the indicated times during the course of Pal depletion.
98
99 Figure 19. Phenotypes of Pal depletion strains. (A) To visualize cell morphology in cells depleted of Pal (right panels), compared to cells containing Pal (left panels), strain LS4524 was grown in PYEX to induce pal expression. Cells were then washed and grown in PYEG for 10 h to deplete Pal. Cells grown in either PYEX or PYEG were stained with FM4-64 and visualized by light microscopy. Bar, 2 µm. When strain
LS4524 was grown in PYEG for 10 h, the cells exhibited a late-stage cell division defect and polar blebs (white arrows). (B) Scanning EM images of the Pal depletion strain LS4524 incubated in the presence of either xylose or glucose for 12 h. Wild- type cells and strain LS4524 grown in the presence of xylose to induce expression of
Pal were similar. However, in the absence of Pal, surface blebs were visible across the lateral cell surface, at the cell poles, and at the division plane (indicated by arrows). (C)
A cryo-EM image of wild-type cells exhibiting a well-defined IM, PG layer, OM, and
S-layer. Visualization of strain LS4524 after Pal depletion by growth in PYEG for 10 h showed aberrant OM structures at the cell pole and at the site of division.
100
Figure 20. Construction of TolA and TolB depletion strains. (A) A schematic diagram of the construction of TolA depletion strain, LS4525. An in frame deletion of tolA was constructed by deleting base pairs 37-777, out of a total of 813 bp of coding sequence, while the full-length tolA gene was integrated at the chromosomal xylX locus. A tolB depletion strain was constructed by incorporating a truncated version of tolB at the native chromosomal site and a second, full-length copy under control of the xylose inducible promoter, creating the strain LS4526. (B) CFU for strain LS4525 and
LS4526 grown in PYEX, washed and then incubated in the presence of either PYEX or PYEG. Cell viability decreased after 12 h of growth in PYEG (indicated by arrow).
(C) To visualize cell morphology in cells depleted of TolA, as compared to cells containing TolA, strain LS4443 was grown in PYEX to induce tolA expression. Cells were then washed and grown in PYEG for 12 h to deplete TolA. When strain LS4525 was grown in PYEG for 12 h, cells exhibited a late-stage cell division defect and polar
101 blebs (indicated by arrows). (D) To visualize cell morphology in cells depleted of
TolB, strain LS4526 was grown in PYEX, as described for LS4525. When strain
LS4526 was grown in PYEG for 12 h, cells exhibited cell division and membrane integrity phenotypes (indicated by arrows).
102 103 Figure 21. Phenotype of TolA and TolB depletion strains. (A) SEM images of the
TolA depletion strain LS4525 grown in the presence of either xylose or glucose for 12 h (to deplete TolA). (B) SEM images of the TolB depletion strain LS4526 grown for
12 h in the presence of either xylose or glucose (to deplete TolB). Depletion of either
TolA or TolB yielded cells with extensive surface blebs. Bar (A and B), 1 µm. (C)
Cryo-EM images of strain LS4525 grown for 11 h in glucose to deplete TolA, showing significant OM disruptions both laterally and at the cell poles (arrows). (D)
Cryo-EM images of strain LS4526 grown for 11 h in glucose to deplete TolB show
OM defects at both the cell poles and the division plane (arrows). Bar (C and D,) 100 nm.
104 The Tol-Pal complex attaches IM and OM to the peptidoglycan layer
Since the Tol-Pal complex in E. coli is known to span the periplasmic space
(Levengood et al. , 1991), and Caulobacter depletion mutants of both TolA and Pal have large OM regions that are substantially separated from the IM (Fig. 19 and 21), we used cryoEM to observe the position of the peptidoglycan layer with respect to each membrane layer in both TolA and Pal depletion strains (Fig. 22). In the Pal depletion strain, the peptidoglycan layer (arrows, Fig. 22A) adhered solely to the IM over large areas of the OM blebs at the cell pole (13/14 cells examined). In contrast, in the TolA depletion strain, the peptidoglycan layer (arrows, Fig. 22B) adhered to the
OM, and did not attach to the IM over large areas at the cell pole in 35% of cells
(n=20 cells). The same phenomenon was observed for the peptidoglycan layer along the lateral cell surface (Fig. 22C and 22D).
105
Figure 22. Structural relationship of the peptidoglycan layer and the IM and OM in TolA and Pal depletion strains. (A) Cryo-EM of strain LS4524 after Pal depletion by growth in PYEG for 10 h showed an OM polar bleb with the peptidoglycan layer
(arrows) dissociated from the OM but adhering to the IM. (B) Cryo-EM of strain
LS4525 grown for 11 h in glucose to deplete TolA showed an OM polar bleb with the peptidoglycan layer (arrows) adhering to the OM but separated from the IM. (C)
Lateral cell envelope of strain LS4524 depleted of Pal with the OM separated from the peptidoglycan layer (arrows). (D) Lateral cell envelope of strain LS4525 depleted of
TolA, with the peptidoglycan layer separated from the IM.
106 Discussion
The Caulobacter crescentus Tol-Pal complex is required for proper construction of the cell envelope and for completion of OM invagination in the late stage of cell division, as is the case in E. coli (Gerding et al. , 2007). Unlike E. coli , however, the Caulobacter Tol-Pal complex is essential for viability. Although the Tol-
Pal complex is not essential in E. coli , it contributes to the invagination of OM during cell division and is localized to the division site (Gerding et al. , 2007), and it also helps to maintain the integrity of the cell wall by connecting the OM and the peptidoglycan network (Cascales et al. , 2002). E. coli primarily employs Lpp, a lipoprotein, to maintain the cell envelope integrity via generally distributed linkages between the OM and peptidoglycan layer (Weigand et al. , 1976). Caulobacter does not have a Lpp homolog. Instead, we suggest that Caulobacter uses Tol-Pal to maintain cell envelope integrity performing an analogous function to Lpp in E. coli . In support of this, we observed that all of the fusion proteins of the Tol-Pal complex localize to the division site and a significant fraction accumulate throughout the cell envelope (Fig. 19A). The overproduction of Pal or TolA in E. coli complements the
OM integrity defect of an lpp mutant strain (Cascales et al. , 2002), suggesting that Pal and TolA play an analogous though less important role than Lpp in E. coli . In
Caulobacter , both the TolA and Pal proteins are required to maintain the subcellular location of the TipN polar marker (Lam et al. , 2006; Huitema et al. , 2006) that plays a critical function in cell asymmetry and polar development.
107 TolA and Pal accumulation at the incipient division site is dependent on FtsZ assembly
The TipN polar localization protein and the TolA, Pal, TolB, TolR and TolQ components of the Tol-Pal complex (Fig. 19) localize to the cell division plane and then remain at the new poles upon completion of cell division. TolA localizes to the division plane well after the accumulation of the FtsZ protein at the incipient cell division site, and TolA is dependent on FtsZ for its positioning at that site. We found that the recruitment of FtsA to the division plane followed arrival of TolA and, as expected, the recruitment of TolA, and all the other components of the Tol-Pal complex, to the division plane was independent of FtsA. The finding parallels that of the FtsZ-dependent and FtsA-independent midcell localization of DipM, which is a putative peptidoglycan endopeptidase required for peptidoglycan remodeling during cell division (Moll et al. , 2010b; Poggio et al. , 2010a; Goley et al. , 2010c). We found, however, that the recruitment of Pal to the division plane required TolA (Fig. 19E), as is the case in E. coli (Gerding et al. , 2007), while the recruitment of the other components of the Tol-Pal complex was independent of both TolA and Pal. Thus, the mid-cell localization of the TolA and Pal components of the Tol-Pal complex depends on the formation of the Z-ring, as does the mid-cell positioning of the TipN protein
(Huitema et al. , 2006; Lam et al. , 2006).
The sub-cellular positioning of TipN is dependent on, and TipN forms a complex with, the TolA and Pal proteins
In wild type cells, TipN, a polar protein with two transmembrane domains and a short periplasmic domain, transiently localizes to the division plane significantly
108 later than the appearance of TolA at midcell. TipN is then maintained at the new cell poles following cell separation (Huitema et al. , 2006; Lam et al. , 2006) at the same position as polar TolA and Pal. We found that TipN mislocalized in either Pal or TolA depletion strains. Although the retention of TipN at the division plane and at the cell poles required both the TolA and Pal proteins, it is possible that cell envelope defects induced by inactivation of the Tol-Pal complex might disrupt the interaction between
TipN and a possible unknown positioning factor. However, we showed that TipN interacts, directly or indirectly, with both TolA and Pal in co-immunoprecipitation assays (Fig. 17), suggesting that complex formation with TolA and Pal contributes to the subcellular localization of TipN. Moreover, it was recently reported that FtsN can interact with both TolR and TipN in a bacterial adenylate cyclase two-hybrid assay
(Moll et al. , 2010b). Our observation that the TipN polar landmark protein appears to interact with Pal in the outer membrane, and that it also associates with the TolA inner membrane protein, suggests trans-envelope subcellular localization of the
TolA/Pal/TipN complex at the cell pole.
The PleC histidine kinase was also partially mislocalized in cells depleted of
TolA and Pal, but it was not found to be part of TolA complex, indicating that the mislocalization of PleC is probably a downstream effect of TipN mislocalization.
Notably, other than the PleC, whose polar positioning is dependent on TipN (Lam et al. , 2006), the polar localization of other polar transmembrane proteins, including DivJ and PopZ, was unaffected by depletion of the TolA or Pal proteins. We cannot exclude the possibility that mislocalization of PleC in the TolA/Pal depletion depends on PodJ, since PodJ is also required for the polar localization of PleC. Cumulatively, these observations suggest the localization dependency pathway shown in Fig. 16D.
109 The Tol-Pal complex is necessary for OM-peptidoglycan integrity
In TolA, TolB and Pal depletion strains, the OM extrudes outward to form widespread blebs (Fig. 19C, 21C and 21D), suggesting that these OM blebs probably formed due to a deficiency of Tol-Pal connections between the OM and the peptidoglycan layer in the mutant strains. Interestingly, depletion of dipM , causes peptidoglycan thickening and the formation of cell surface blebs similar to that seen in
Tol-Pal depletion strains (Goley et al., 2010c). The physical contact between TolA and Pal may be disrupted in DipM-depleted cells due to peptidoglycan thickening.
CryoEM images revealed that in cells depleted of TolA, the peptidoglycan layer sometimes failed to attach to the IM, while adhering to the OM (Fig. 22B and
22D). Similar disruptions in envelope organization were not detected in either Pal or
TolB depletions. Instead, the peptidoglycan layer adhered only to the IM in the blebs of those mutant strains (Fig. 22A and 17C). The depletion of tolA also caused the formation of OM blebs along the cell sidewalls. In accordance with phenotypes exhibited by E. coli TolA mutants (Lloubes et al. , 2001), the absence of TolA in
Caulobacter induced more extensive OM defects than in either pal or tolB mutant strains. Thus, these observations suggest TolA plays the larger role in maintaining general cell envelope integrity.
In wild type Caulobacter , there is a short stretch of unsupported OM that maintains its integrity and withstands internal pressure during the late stages of cell division when the IM and peptidoglycan layer are constricting and ultimately fissioned to form two cell compartments before completion of OM invagination (See WT cell in
Fig. 19C) (Judd et al. , 2005). In either Pal or TolB depletion strains, the OM-
110 peptidoglycan integrity is disrupted, predominately at newly synthesized areas of the cell wall, such as the division plane and recently created poles. Thus, the OM in these areas appears to be poorly anchored, resulting in localized OM bulging (Fig. 19C, middle). Newly synthesized Pal is inserted at the site where new peptidoglycan is incorporated (Anwari et al. , 2010), consistent with the Tol-Pal complex playing a vital role in OM membrane integrity by mediating OM-peptidoglycan contacts during growth and division.
Owing to the temporal separation of the constriction of the IM/peptidoglycan and the OM in Caulobacter , there is a physical tension between the OM and the peptidoglycan layer as the IM/peptidoglycan invagination approaches the terminal stage of fission. Upon fission of the IM and peptidoglycan layer, the OM is left unsupported, as shown in Figure 19C (WT cell). The final stages of IM fission leading to cell compartmentalization probably occur extremely rapidly (Judd et al. , 2005).
Thereafter, over a period of about 20 minutes, the periplasmic gap between the IM and the peptidoglycan layer is closed by progressive attachment of the OM to the peptidoglycan layer, presumably by action of Tol-Pal complex molecules, at the point of OM and peptidoglycan layer divergence so that the region of untethered OM becomes smaller and smaller (Judd et al. , 2005). At some point, when the unattached region is small enough that the OM curvature reaches a critical point, the OM also fissions and reforms over the new cell poles of the daughter cells. In Pal depletion strains, the region of OM-peptidoglycan disconnection appears to be significantly larger and the subsequent reconnection of OM and peptidoglycan fails to occur, leading to the blown out sections as in Figure 19C (middle). Some of the cell poles in the Pal depletion strain exhibit the peptidoglycan layer adhered to the IM and the OM
111 blown out in a large bleb-like structure, unable to make contact with the PG and IM
(Fig. 22A). These cells are probably those that completed cell division earlier in the depletion process when there was still a partially functional Tol-Pal complex.
The fitness advantage to Caulobacter of separating the time of IM fission from final OM constriction and cell separation is a matter of conjecture. In the extended time interval when the cell is compartmentalized, but not separated, free diffusion can occur through continuous periplasmic space surrounding both compartments. The stalk acts to collect phosphorus efficiently in low-phosphate environments (Gonin et al. , 2000; Wagner et al. , 2006), so a possible fitness benefit is that this cell division method provides more reliable development of the nascent swarmer daughter cell in low environmental phosphate situations. A time interval between compartmentalization and cell separation may also assure the assembly of the chemotaxis control system before the swarmer cell is physically separated from the stalked cell. The delayed completion of OM constriction would be a necessary feature of late cell division in this scenario.
112 Table 4: Strains and plasmids
Strain or Construction or plasmid Relevant genotype/description Source CB15N synchronizable derivative of Caulobacter crescentus CB15 (Evinger & Agabian, 1977) LS4517 CB15N xylX:: PxylX -tolQ-yfp Electroporation of pYC103 into CB15N LS4518 CB15N xylX:: PxylX -gfp-tolR Electroporation of pYC104 into CB15N LS4519 CB15N xylX:: PxylX -gfp-tolA Electroporation of pYC105 into CB15N LS4520 CB15N xylX:: PxylX -mCherry-tolB Electroporation of pYC106 into CB15N LS4521 CB15N xylX:: PxylX -pal-mCherry Electroporation of pYC107 into CB15N LS4522 CB15N ∆tolA xylX ::P xylX-gfp-tolA LS4525X Φ (LS4519) LS4523 CB15N ∆pal xylX ::P xylX-pal-mCherry LS4524X Φ (LS4521) EG 052 CB15N xylX:: PxylX -ftsZ–venus (Goley, unpublished ) EG 055 CB15N xylX:: PxylX –venus-ftsA (Goley, unpublished) EG 051 CB15N xylX:: PxylX–venus–ftsI (Goley, unpublished ) LS3702 CB15N ∆ftsZ xylX ::P xylX-ftsZ (Wang et al. , 2001) EG 083 CB15N ∆ftsA xylX ::P xylX-ftsA (Goley, unpublished) LS4524 CB15N ∆pal xylX ::P xylX-pal Constructed using pYC102 and pYC200 LS4525 CB15N ∆tolA xylX ::P xylX-tolA Constructed using pYC101 and pYC201 LS4526 CB15N ∆tolB tolB:: PxylX-tolB Electroporation of pYC103 into CB15N LS4527 CB15N tipN::tipN-gfp (Huitema et al. , 2006) LS3785 CB15N pleC::pleC-gfp (Viollier et al. , 2002) LS4199 CB15N divJ::divJ-yfp (Matroule et al. , 2004) GB175 CB15N vanA ::P van-popZ-yfp (Bowman et al. , 2008) LS4528 CB15N tipN :: tipN-gfp ∆pal xylX ::P xylX-pal LS4524X Φ (LS4527) LS4529 CB15N tipN :: tipN-gfp ∆tolA xylX ::P xylX-tolA LS4525X Φ (LS4527) LS4530 CB15N pleC :: pleC-gfp ∆pal xylX ::P xylX-pal LS4524X Φ
113 Strain or Construction or plasmid Relevant genotype/description Source (LS3785) LS4531 CB15N pleC :: pleC-gfp ∆tolA xylX ::P xylX-tolA LS4525X Φ (LS3785) LS4532 CB15N divJ :: divJ-yfp ∆pal xylX ::P xylX-pal LS4524X Φ (LS4199) LS4533 CB15N divJ :: divJ-yfp ∆tolA xylX ::P xylX-tolA LS4525X Φ (LS4199) LS4534 CB15N ∆pal PxylX-pal vanA ::P van-popZ-yfp LS4523X Φ (GB175) LS4535 CB15N ∆tolA PxylX-tolA vanA ::P van-popZ-yfp LS4524X Φ (GB175) LS4536 CB15N tipN :: tipN-gfp ∆tolA xylX ::P xylX-tolA-m2 LS4537 X Φ (LS4527) LS4537 CB15N ∆tolA xylX ::P xylX-tolA-m2 LS4525X Φ (LS4545) LS4538 CB15N xylX ::P xylX-tipN-m2 Electroporation of pYC119 into CB15N LS4539 CB15N ∆tolA xylX :: PxylX-tolA vanA ::P van-pal-mCherry Electroporation of pYC113 into LS4525 LS4540 CB15N ∆tolA xylX ::P xylX-tolA pP van-tolQ-yfp Electroporation of pYC110 into LS4525 LS4541 CB15N ∆tolA xylX ::P xylX-tolA pP van-yfp-tolR Electroporation of into pYC111 LS4525 LS4542 CB15N ∆pal xylX ::P xylX-pal pP van-tolQ-yfp Electroporation of pYC110into LS4524 LS4543 CB15N ∆pal xylX ::P xylX-pal pP van-yfp-tolR Electroporation of pYC111into LS4524 LS4544 CB15N ∆pal xylX ::P xylX-pal pP van-yfp-tolA Electroporation of pYC112into LS4524 LS4545 CB15N xylX ::P xylX-tolA-m2 Electroporation of pYC114 into LS4524 LS4546 CB15N ∆ftsZ xylX ::P xylX-ftsZ pP van-yfp-tolA Electroporation of pYC112 into LS3702 LS4547 CB15N ∆ftsA xylX ::P xylX-ftsA pP van-yfp-tolA Electroporation of pYC112 into EG083 AM52 CB15N ∆ftsN vanA ::P van-ftsN (Moll & Thanbichler, 2009) LS4626 CB15N ∆ftsN vanA ::P van-ftsN xylX ::P xylX-yfp-tolA Electroporation of pYC117 into AM52 pXYFPN-2 For integration of N-terminal YFP fusions at xylX locus (Thanbichler et al. , 2007) pXGFPN-2 For integration of N-terminal GFP fusions at xylX locus (Thanbichler et al. , 2007) pXmChyN-2 For integration of N-terminal mCherry fusions at xylX locus (Thanbichler et al. ,
114 Strain or Construction or plasmid Relevant genotype/description Source 2007) pXmChyC-2 For integration of C-terminal mCherry fusions at xylX locus (Thanbichler et al. , 2007) pYC103 A PCR fragment of the tolQ gene (without stop codon) This study obtained with primers TTTTGGTACCatggacgccgcggccgcc and TTTTGAATTCCCgacccgctcggccaggcgacg was digested with KpnI and EcoRI and cloned into pXYFPC-2 pYC104 A PCR fragment of the tolR (with stop codon) obtained with This study primers TTTTGGTACCatggcgatgtcctccaacgacgcctt and TTTTGAATTCctactgcgcaggccgcaggtcgg was digested with KpnI and EcoRI and cloned into pXGFPN-2 pYC105 A PCR fragment of the tolA (with stop codon) obtained with This study primers TTTTGGTACCatgagcgctcgccgcgaacagactct and TTTTGAATTCtcaacgcgaacaggcctgttttgcat was digested with KpnI and EcoRI and cloned into pXGFPN-2 pYC106 A PCR fragment of the tolB (with stop codon) obtained with This study primers TTTTGGTACCatgcgccttagagccctgctgctgat and TTTTGAATTCctagtccagaaggggcgaccaggcgg was digested with KpnI and EcoRI and cloned into pXmChyN-2 pYC107 A PCR fragment of the pal (without stop codon) obtained This study with primers TTTTGGTACCttgaggagaaactggatgagcttcgacaccc and TTTTGAATTCCCgcgagcgccgtccgtgatggc was digested with KpnI and EcoRI and cloned into pXmChyC-2 pNPTS138 For gene replacement by double homologous recombination M.R.K. Alley, unpublished pYC101 pNPTS138 carrying tolA upstream and downstream 500 bp. This study The flanking regions of tolA were amplified using CB15N chromosomal DNA as template and the primer pairs ttttaagcttcgcgacgggcgggcgc with TTTTGGATCCggccggagacagagtctgttcgcg and TTTTGGATCCctcaacttcaatgcaaaacaggcctgttcg with TTTTGAATTCggtcaggcgctcatagaccgcgtcg. The PCR fragments were digested with HindIII and BamHI and with BamHI and EcoRI, respectively. The two PCRs were triple ligated into pNPTS138 digest with HindIII and EcoRI. pYC102 pNPTS138 carrying pal upstream and downstream 500 bp. This study The flanking regions of pal were amplified using CB15N chromosomal DNA as template and the primer pairs TTTTCTGCAGtggccgttacaccacgccggtctgg with TTTTGGTACCgcgctgggtgtcgaagctcatccagt and TTTTGGTACCaacggccgcacggccatcacgg with TTTTGAATTCcaggtcggccagcgtcttctcaatcg. The PCR fragments were digested with PstI and KpnI and with KpnI and EcoRI, respectively. The two PCRs were triple ligated into pNPTS138 digest with PstI and EcoRI. pMT69 For integration of genes at the xylX locus (Thanbichler & Shapiro, 2006) pYC200 tolA inserted between NdeI and EcoRI sites into pMT69 This study pYC201 A PCR fragment of the pal obtained with primers This study TTTTCATATGaggagaaactggatgagcttcgacaccc and TTTTGAATTCtcagcgagcgccgtccgtgatggc was digested with NdeI and EcoRI and cloned into pMT69
115 Strain or Construction or plasmid Relevant genotype/description Source pXMCS-2 For generation xylose dependent depletion strains (Thanbichler et al. , 2007) pYC103 A PCR fragment of the tolB (1-400 bp) obtained with primers This study TTTTCATatgaacaaggagaccccgatgcgcctta and TTTTGGTACCcccttctcgccggtcaggcgct was digested with NdeI and KpnI and cloned into pXMCS-2 pVCHYC-2 For integration of C-terminal mCherry fusions at vanA locus (Thanbichler et al. , 2007) pYC107 The KpnI and EcoRI digested fragment from pYC107 was This study cloned into pVCHYC-2 pRVYFPC-5 replicating plasmid for vanillate-inducible C-terminal YFP (Thanbichler et al. , fusions 2007) pRVYFPN-5 replicating plasmid for vanllilate-inducible N-terminal YFP (Thanbichler et al. , fusions 2007) pYC110 The KpnI and EcoRI digested fragment from pYC103 was This study cloned into pRVYFPN-5 pYC111 The KpnI and EcoRI digested fragment from pYC104 was This study cloned into pRVYFPC-5 pYC112 The KpnI and EcoRI digested fragment from pYC105 was This study cloned into pRVYFPN-5 pXFLGC-2 For integration of C-terminal M2 tag fusions at xylX locus (Thanbichler et al. , 2007) pYC119 A PCR fragment of the tipN (without stop codon) obtained This study with primers TTTTGGTACCcgttcacgcgccgcgccg and TTTTGAATTCCCggccagatcgccgctcgccg was digested with KpnI and EcoRI and inserted into pXFLGC-2 pVCHYC-4 For integration of C-terminal mCherry fusions at vanA locus (Thanbichler et al. , 2007) pYC113 A PCR fragment of the pal (without stop codon) obtained This study with primers TTTTGGTACCttgaggagaaactggatgagcttcgacaccc and TTTTGAATTCCCgcgagcgccgtccgtgatggc was digested with KpnI and EcoRI and cloned into pVCHYC-4 pXGFPC-4 For integration of C-terminal GFP fusions at xylX locus (Thanbichler et al. , 2007) pYC115 A PCR fragment of the tolA-m2 (with stop codon) obtained This study with primers TTTTGGTACCatgagcgctcgccgcgaacagactct and TTTTgctagcttacttgtcatcgtcatccttgtagtcggaccggtgacgcgtaacgttc gaattcggacgcgaacaggcctgttttgcatt was digested with KpnI and NheI and cloned into pXGFPC-4 digest with KpnI and NheI pXYFPN-2 For integration of N-terminal YFP fusions at xylX locus (Thanbichler et al. , 2007) pYC117 The KpnI and EcoRI digested fragment from pYC105 was This study cloned into pXYFPN-2
Φ indicates generalized transduction, as mediated by bacteriophage ΦCr30. For example, CB15N x Φ(LS3785) means that a bacteriophage lysate made from LS3785 was used to infect CB15N.
116
Chapter 4
ZapA-FtsZ interaction is required for Caulobacter cell division
117 Introduction
Cell division is a complex process that requires the temporal and spatial coordination with the chromosome segregation and cell growth to ensure that each daughter progeny receives a complete genetic complement. Bacterial cells divide by placing a cytokinetic ring, a polymeric structure of the conserved tubulin-like FtsZ protein, at the future site of division. The Z-ring serves as the scaffolding network that recruits other components of the divisome (the cell division machinery) to the division plane (Goehring & Beckwith, 2005). FtsZ is a GTPase that undergoes GTP-dependent polymerization in vitro. Recently, short membrane-bound FtsZ filaments have been suggested as the mechanism for generation of the constrictive force to drive the cell division, but the detailed mechanism is unknown. Assembly of the Z-ring is a primary point of control over timing and positioning of the cell division plane.
The current view is that the precise selection of the division site at the midcell is controlled by confining the robustness of Z ring establishment through two mechanisms. These two mechanisms are the Min system and nucleoid occlusion factors (Wu et al. , 2009). Acting negatively, the Min system involves accumulation of division inhibitors MinC and MinD at the cell poles. This mechanism is widely used among bacteria, via modified forms such as B. subtilis DivIVA and E. coli MinE contributing the spatial tuning of the MinCD complex at the poles. EzrA is another B. subtilis membrane protein that localizes to the Z-ring but also inhibits FtsZ assembly throughout the cell, probably counteracting positive assembly factors. Also acting negatively, SlmA from E. coli and Noc from B. subtilis interact nonspecifically with chromosomal DNA and therefore prevent Z-ring assembly. Caulobacter does not have
118 either the Min system or nucleoid occlusion. Instead, it imposes a mode of regulation that integrates spatial information from both the cell poles and the chromosome via a factor called MipZ. It was previously shown that MipZ directly interacts with FtsZ in vitro and stimulates its GTPase activity, which inhibits the polymerization of FtsZ
(Thanbichler & Shapiro, 2006). MipZ inhibits the polymerization of FtsZ and since
MipZ localizes to the poles by binding to parS via ParB, the FtsZ can only assemble near the midcell, which has the lowest concentration of MipZ.
In contrast, there are antagonistic proteins that stimulate formation of FtsZ rings. Various positive regulators of FtsZ polymerization or Z-ring formation are identified (Goehring et al. , 2005; RayChaudhuri, 1999; Goley et al.; Ishikawa et al. ,
2006; Goley et al. , 2010b), including ZipA and FtsA of E. coli and SepF of B. subtilis
(Hamoen et al. , 2006). In E. coli, ZipA and FtsA both bind to FtsZ through a conserved sequence at the C-terminus (Wang et al. , 1997), and both are essential for cell division. Interestingly, Z-rings are able to assemble in the absence of either FtsA or ZipA (RayChaudhuri, 1999; Addinall & Lutkenhaus, 1996a). Geissler et. al have shown that FtsA*(R286W) is sufficient to bypass the need for ZipA (Geissler et al. ,
2007). Therefore, this result suggests that FtsA is the primary factor for anchoring
FtsZ filaments to the membrane in most bacteria. Recent studies in B. subtilis have shown that SepF is conserved among Gram-positive bacteria and interacts directly with FtsZ by two-hybrid analysis (Singh et al. , 2008).
Another positive stabilizing agent has recently been discovered with wide conservation amongst Gram-positive and Gram-negative bacteria. ZapA protein was first identified in B. subtilis as a protein whose overexpression can overcome the
119 division defects produced by overexpression of the MinD (Hu et al. , 2002). ZapA is recruited to the divisome at an early stage in its assembly through a direct interaction with FtsZ. ZapA is not essential for cell division both in B. subtilis and E. coli , but several lines of evidence indicate that ZapA is important for promoting the bundling of
FtsZ protofilaments in vitro . The crystal structure of ZapA from Pseudomonas aeruginosa was recently solved (Low et al. 2004). Despite the availability of protein crystal structure for both ZapA and FtsZ, the detailed interaction mechanism remains unclear.
In this study, we report that the Caulobacter ZapA is concentrated at the division plane and following cell division it remains at the new poles, colocalizing and interacting with FtsZ. Cells bearing a null mutation in zapA are viable, but they are slightly elongated and tend to form filamentous cells. We showed that the interactions between ZapA and FtsZ are corroborated by co-immunoprecipation and by in vitro analysis of purified proteins. Furthermore, Caulobacter ZapA counteracted MipZ, a cell division inhibitor, in stabilizing assembly of FtsZ filaments in vitro .
120 Materials and methods
Bacterial strains, synchronization, growth conditions and cloning. All Caulobacter strains were derived from CB15N and grown at 28°C in M2-glucose minimal medium
(M2G) or peptone yeast extract (PYE) with select antibiotics. For the ZapA depletion construct, cells were grown overnight in PYE medium containing 0.3% xylose, washed with plain PYE medium, and then re-suspended in PYE containing 0.2% glucose (t = 0 h). Samples were taken every three hours and analyzed by western blotting and light microscopy. The construction of bacterial strains is in the table 4.
EM assay. FtsZ (1.5 µM) was incubated with ZapA (1.5 or 6 µM) and BSA for 30 min at 25°C in buffer P (50mM Hepes/NaOH, pH 7.2, 50mM KCl, 10mM MgCl 2, 1mM β- mercaptoethanol). Subsequently, 10 µl aliquots were applied on the copper grid for analysis. 2% uranyl acetate was used to stain the filaments for 1min. Images were taken on a JEOL TEM1230 transmission electron microscope.
Fluorescence Microscopy. Cells were immobilized using a thin layer of agarose in
M2G medium. For localization and labeling studies, 0.3% xylose and 0.5mM vanillic acid (pH 7.5) were used to induce expression of cfp , venus or mCherry fusions from the xylX and vanA promoters for at least 1 h, respectively. Differential interference contrast (DIC) and fluorescence microscopy images were obtained using a Leica DM
6000 B microscope with a HCX PL APO 100x/1.40 Oil PH3 CS objective,
Hamamatsu EM-CCD C9100 camera, and a custom-designed microscope control and image analysis software package KAMS (Christen et al. , 2010).
Co-immunoprecipitation. TAP tag coimmunoprecipitation was performed as described in (Puig et al. , 2001). TAP strain was induced with 0.3% xylose for 3 h.
121 Cells were lysed by passage through the French Press at 15000 psi twice. The cell lysate was then incubated with IgG sepharose overnight. Subsequently, the IgG sepharose was washed and cleaved with TEV protease overnight; then the supernatant was collected and incubated with S protein agarose for 6 h. The S protein agarose was boiled for western blot analysis.
Western blotting. Western blotting to monitor ZapA, IgG, and FtsZ was performed as standard procedures. ZapA and FtsZ polyclonal antibodies (1/2000), IgG monoclonal antibody (1/20,000), and HRP conjugated-anti-rabbit secondary antibody (1/20,000) were used.
122 Results
ZapA acts as an important factor in cell division of Caulobacter
The representation of Caulobacter zapA and surrounding genes on the chromosome was shown schematically in Fig. 23A. Sequence analysis of CC3247 indicates that it shares 21.9% identity and 39.6% similarity with zapA from E. coli .
The predicted ZapA protein structure contains a signature coiled-coil structure at the carboxyl terminus. Based on the sequence and structure homology, we designated
Caulobacter gene CC3247 as zapA .
To explore the physiological role of the Caulobacter ZapA protein upon cytokinesis, we generated an in-frame zapA deletion strain, YC40, by deleting the majority of the native zapA coding region (amino acid 13 to 97) (Fig. 23B). We were able to obtain ∆zapA cells, indicating that it is not essential for viability. However, examination of cells lacking ZapA by phase contrast microscopy revealed that cells display varying degrees of filamentation, indicating that ZapA has a critical role in supporting efficient cell division. We therefore generated a complementing vector with the goal of rescuing mutant phenotypes. The complementing vector, which carries a full-length zapA gene under the control of a xylose-inducible promoter, was integrated at the chromosomal xylX locus in the zapA strain background generating strain
YC202.
In the depletion strain, cells looked morphologically wild type in the presence of xylose, but became elongated by 8 h of depletion in the absence of inducer. Similar to the phenotype of zapA strain, cells displayed heterogeneity in cell length
123 distribution even though they did not exhibit a significant difference of doubling time compared with wild-type cells when grown in rich media (data not shown). Cells lacking ZapA varied in cell lengths (Fig. 23C). Specifically, we noted aberrant division site placement within elongated cells. In order to assess ZapA protein levels upon its depletion, polyclonal antibodies were raised against purified recombinant ZapA and used to probe Caulobacter cell lysates. Immunoblot analysis with anti-ZapA antibodies showed that ZapA was undetectable in the deletion strain and that it was depleted to trace amount between 6 and 9 h in the absence of xylose in the depletion strain (Fig. 23C). In the ZapA depletion strain, there was no significant change in the levels of FtsZ after 9 h of growth in the absence of xylose (Fig. 23C). Thus, the filamentous phenotype in the depletion strains was not due to effects in elimination of
FtsZ. This result suggests that ZapA plays an important role in cytokinesis of
Caulobacter cells.
ZapA-mCherry accumulates at the incipient cell division site
To gain insight into the timing of ZapA accumulation, we assessed zapA transcript and ZapA protein levels over the cell cycle. Microarray analysis of the temporal gene expression pattern indicated that zapA transcript is not cell cycle regulated (McGrath et al. , 2007). In order to examine ZapA protein level over the course of the cell cycle, anti-ZapA was used to probe Caulobacter cell lysates taken at various time points post synchronization (Fig. 24A). This immunoblot analysis revealed that the level of ZapA was low in swarmer cells but increased in stalked cells and were high throughout the remainder of the cell cycle, consistent with it acting at the time of cell division (Fig. 24A). The quality of the synchrony was monitored by
124 immunoblot analysis of the FtsZ protein. After one cell cycle, a second synchrony was performed. It was found that upon cell division, ZapA is compartmentalized to the stalked cells, while the swarmer cells inherited relatively low amount of the protein
(data not shown). With the compartmentalization of the predivisional cell, both FtsZ and ZapA were down-regulated in swarmer cells, and reaccumulated in stalked cell.
Several cell division proteins in Caulobacter are post-translationally regulated over the course of the cell cycle and peak in abundance when they are functional (McGrath et al. , 2007). Therefore, this result is consistent with ZapA functioning at the time of cell division.
In order to investigate the subcellular localization of ZapA, we replaced the native zapA gene of wild-type CB15N with a zapA-mCherry fusion, resulting in strain
YC225. The cellular morphology and growth rate of strain YC225 was indistinguishable from wild-type, thus the ZapA-mCherry fusion protein is functional.
To compare the localization dynamics of ZapA with FtsZ, we therefore investigated the temporal order of ZapA recruitment to the midcell. A strain was made by transducing the P van-ftsZ-yfp allele into strain YC225. We followed the localization of
ZapA-mCherry and FtsZ-YFP simultaneously over the course of the cell cycle by isolating swarmer cells and imaging them as they grew synchronously on agarose pads
(Fig. 24A). As previously described (Thanbichler & Shapiro, 2006), FtsZ-YFP began as a focus at the pole of swarmer cells, and appeared at the incipient division site.
Interestingly, ZapA-mCherry was found to colocalize with FtsZ-YFP at the pole of swarmer cells and the division site of stalked and predivisional cells. Following separation of the two daughter cells, FtsZ-YFP and ZapA-mCherry remained at the
125 new pole of swarmer and stalked cells.
ZapA requires FtsZ for its division site localization
Since ZapA-mCherry was found to colocalize with FtsZ-YFP at the division site, we ask if the localization of the ZapA is dependent on the localization of the FtsZ.
To determine this, strain YC226 was made by transducing P zapA -zapA-mCherry allele in a background where expression of the only functional copy of ftsZ is under the control of the xylose-inducible promoter. Synchronized swarmer cells grown in the absence of xylose were depleted of FtsZ for 3 h and became filamentous, and ZapA- mCherry was observed in a diffuse peripheral pattern in these cells (Fig. 25B). FtsZ was then allowed to accumulate in these cells by adding xylose on the agarose pad.
Repletion of FtsZ restored the localization of ZapA to midcell bands (Fig. 25B). Thus,
ZapA requires FtsZ for its positioning at the division site.
To ascertain whether the fairly diffused pattern of ZapA-mCherry was due to the loss of FtsZ and not cell elongation, we treated YC226 swarmer cells with cephalexin to block late cell division by inhibiting FtsI activity. After 4 h of cephalexin treatment, cells were slightly filamentous. ZapA-mCherry was found to colocalize with FtsZ within the elongated cells (Fig. 25C), suggesting that ZapA is an early recruit to the cell division apparatus and its localization is linked to and dependent on the FtsZ localization.
126
Figure 23. Phenotype of ZapA depletion (A) Schematic of the gene organization of the Caulobacter zapA . Arrows indicate the direction of transcription and putative position of promoters. (B) A schematic diagram of the construction of ZapA depletion strain, YC202. An in frame deletion of zapA was constructed by deleting base pairs
37-288, out of a total of 324 bp of coding sequence, while the full-length zapA gene was integrated at the chromosomal xylX locus. (C) YC202 was grown in PYEX, washed and then suspended in PYEG (t = 0 h). Cells were visualized by phase- contrast microscopy at the indicated time points. (D) ZapA and FtsZ protein levels were measured by western blot analysis of samples taken at the indicated times during the course of ZapA depletion.
127
Figure 24. Relative localization of fluorescently labeled ZapA and FtsZ during the cell cycle. (A) Western blot analysis of relative ZapA protein level (upper panel; arrows indicate the ZapA protein) during the cell cycle starting with a synchronous population of wild-type swarmer cells. The FtsZ protein (lower panel) is shown as a loading control and quality control for the synchrony. (B) Synchronized YC227 swarmer cell populations producing ZapA-mCherry and FtsZ-YFP were transferred onto an M2G-agarose pad ( t = 0 min) and visualized at 20-min intervals by phase- contrast and fluorescence microscopy. Schematics showing the temporal sequence of protein localization and cell constriction are given. Red: ZapA-mCherry; Green: FtsZ-
YFP.
128 Since zapA mutant shows morphological defects at the division site, this raised the possibility that ZapA plays a role in regulating Z-ring assembly. Thus, we asked whether the FtsZ localization is affected in the absence of ZapA. To test this, a strain bearing vanillate-inducible ftsZ-cfp fusion at the chromosomal vanA locus as well as xylose-inducible mipZ-yfp at the chromosomal xylX locus in the zapA background (YC99) was generated. Interestingly, as opposed to the single Z ring observed in predivisional cells of wild-type cells, multiple aggregates of FtsZ appeared throughout the cell upon ZapA depletion. The cellular localization of MipZ-YFP remained normal in zapA cells. Thus, the aberrant FtsZ rings in the ZapA deletion strain was not due to effects in mislocalization of MipZ. This in vivo result suggests that ZapA might be a regulator for Z-ring assembly.
ZapA physically interacts with FtsZ in Caulobacter
To obtain additional evidence for an interaction between ZapA and FtsZ, we immunoprecipitated ZapA-TAP from a strain bearing xylose-inducible zapA-tap at the chromosomal xylX locus (YC114). The advantage of this method is that tandem affinity peptide purification (TAP) offers an effective means to purify target protein by performing two successive affinity purifications to reduce the contaminations. When strain YC114 was grown in the presence of xylose, inducing expression of zapA-tap for 3 h, cell morphology was similar to that of the wild type (data not shown). As controls, ftsA-tap (YC248) and popZ-tap (YC225) were generated. Formaldehyde, a membrane permeable cross-linking agent, was added prior to cell lysis. We immunoprecipitated cultures of YC114, YC248 and YC225 with anti-ProteinA coupled beads, and added TEV protease to release the bound material. The eluted
129 samples were incubated with calmodulin-coated beads. Analysis of the purified products by western blot showed that FtsZ was found to immuoprecipitate by FtsA-
TAP and ZapA-TAP strains but not PopZ-TAP (Fig. 26). FtsZ was examined in whole cell lysates from three strains to ensure that the protein abundance remained the same in the xylose-induced cells. Moreover, FtsA was not probed in the ZapA-TAP strain and vice versa, suggesting that ZapA-TAP interacts with FtsZ but not with FtsA. These results, together with the localization data, support the conclusion that ZapA directly or indirectly interacts with FtsZ.
Overexpression of ZapA can rescue the cell filamentation of MipZ overexpression
Since MipZ directly interferes with FtsZ assembly, accumulation of MipZ during overproduction causes cell filamentation and minicelling in Caulobacter
(Thanbichler & Shapiro, 2006). In E. coli and B. subtilis , ZapA has been shown in vitro to promote assembly of FtsZ bundling (Gueiros-Filho & Losick, 2002; Small et al. , 2007). To test whether ZapA might compete with the inhibitory activity of MipZ, we created the strain YC213 that highly overexpressed zapA from a high-copy-number plasmid upon induction with xylose and mildly overexpressed mipZ from a high-copy- number plasmid upon induction with vanillate (weaker promoter than xylX ). As a control, an isogenic strain YC214 was generated in which only mipZ is overexpressed from a high-copy-number plasmid upon induction with vanillate. In the absence of both inducers, strain YC213 and YC214 cells grew with normal rates and morphology as compared to wild type. As shown previously, strain YC213 and YC214 cells overexpressing mipZ alone exhibited cell filamentation. Surprisingly, YC213 cells that simultaneously overproduced MipZ and ZapA protein showed a substantial decrease
130 in the filamentous morphology (Fig. 27A), suggesting that overexpression of ZapA can rescue the phenotype of the MipZ overexpression. In the YC213 and YC214 strains, immunoblot analysis with anti-ZapA, anti-FtsZ and anti-MipZ antibodies confirmed that there were no significant changes in the levels of either MipZ or FtsZ after 6 h of growth in the presence of xylose and vanillate. In YC213, complete rescue of the morphological defects associated with zapA overproduction in the presence of xylose confirmed that they were directly attributable to overproduction of zapA.
Our findings thus far implicate ZapA counteracting MipZ to stabilize FtsZ structure and/or dynamics during cell division. We examined this possibility more directly by using in vitro assays of purified components that allow us to assess FtsZ polymerization in the presence and absence of MipZ or ZapA. To test biochemical activity of Caulobacter ZapA, we cloned zapA gene into pET28a vector (Novagen).
This clone was designed to enable the expression of N-terminal hexahistidine-tagged recombinant ZapA in E. coli . The protein was purified by affinity chromatography.
Both Caulobacter native FtsZ and MipZ-His6 were purified as described by
Thanbichler and Shapiro (Thanbichler & Shapiro, 2006).
Since it has previously been showed that MipZ alters the conformation of FtsZ filaments (Thanbichler & Shapiro, 2006), we ask whether ZapA was able to recover
FtsZ filaments under similar conditions. First, we introduced ZapA into FtsZ pelleting reactions in the presence of MipZ at increasing concentrations and found that as ZapA concentration increased, the proportion of FtsZ recovered in the pellet increased. We further explored the effect of ZapA on FtsZ polymers by electron microscopic analyses. As shown previously (Thanbichler & Shapiro, 2006), FtsZ formed short and
131 highly curved polymers in the presence of MipZ (Fig. 27B). However, if inclusion of
ZapA at a 5:1 molar ratio to MipZ, these characteristic structures were essentially undetectable (Fig. 27B). In the presence of ZapA, however, striking bundles were observed instead of short protofilaments which were morphologically different from those formed by MipZ with FtsZ alone. We propose that the inhibitory activity of
MipZ on FtsZ assembly could be restored by adding ZapA protein.
132
133 Figure 25. ZapA localization near midcell is dependent on FtsZ. (A) Fluorescence microscopy of ZapA deletion strains bearing MipZ-YFP and FtsZ-CFP. Strain YC99
(mipZ-cfp vanA :: P van-ftsZ-cfp ∆zapA ) was imaged. The upper panels show the sub- cellular location of MipZ-YFP and FtsZ-CFP in wild type cells, as compared to cells shown in the lower panels in zapA mutant. (B) Synchronized YC226 ( zapA ::P zapA- zapA-mCherry vanA ::P van-ftsZ-yfp ) swarmer cells grown in the absence of xylose were depleted of FtsZ for 3 h and became filamentous, and the localization of ZapA- mCherry was observed in a diffuse pattern in these cells. ZapA-mCheery foci were restored after 2 h grow on the pad in the presence of xylose to induce FtsZ expression.
White arrows indicate foci of ZapA-mCherry. ZapA-mCherry requires FtsZ to form fluorescent foci near the midcell. (C) Synchronized YC226 swarmer cells grown in the presence of cephalexin for 4 h to block late cell division. ZapA-mCherry was found to colocalize with FtsZ-YFP within the elongated cells.
134
Figure 26. Coimmunoprecipitation assay shows that ZapA interacts with FtsZ. (A)
In vivo coimmunoprecipitation of ZapA-TAP in strain YC114 (P xylX-zapA-tap ),
PopZ-TAP in strain YC225 (P xylX-popZ-tap ), and FtsA-TAP in strain YC248 (P xylX- ftsA-tap ). Western blots of whole-cell extracts (Cell lysate) and eluted samples (IO-
TAP) were probed with anti-FtsZ, anti-FtsA, anti-PopZ and anti-ZapA antibodies. FtsZ but not PopZ was observed to interact directly or indirectly with ZapA-TAP and FtsA-
TAP. (B)
135
136 Figure 27. ZapA counteracting MipZ to stabilize FtsZ structure. (A) Strain YC214 was grown to exponential phase in PYE medium. Vanillate was added to a final concentration 0.5mM to induce overexpression of MipZ for 6 h. Cells were withdrawn and analysed by DIC microscopy and by immunoblotting using anti-MipZ, anti-FtsZ and anti-ZapA antiserum. Strain YC213 was grown as described above. Vanillate and xylose were added to co-induce overexpression of both MipZ and ZapA for 6 h. Cells were withdrawn and analysed in the same way as described above. (B) Effect of ZapA on the structure of FtsZ polymers in the presence of MipZ. 2 µM FtsZ and 2 µM MipZ were incubated for 30 min with 2 mM GTP and 1 mM ATP in the absence or presence of 10 µM MipZ. Subsequently, the mixtures were applied to carbon-coated grids. The proteins were stained and examined by transmission electon microscopy at 100,000- fold (left panels) or 300,000-fold (right panels) magnification.
137 Discussion
Overall, the results shown here demonstrate that ZapA of Caulobacter is required for proper FtsZ ring formation. Unlike E. coli , however, the Caulobacter
ZapA is required for normal cell division. The appearance of helical Z rings in ZapA deletion cells shows that ZapA is required for normal Z-ring assembly. In addition,
ZapA is required to maintain a normal cell length and acts at midcell to promote Z ring assembly. A ZapA-mCherry fusion is functional and depended on the localization of
FtsZ to division plane. The Caulobacter ZapA represents a type of FtsZ-modulating protein based on biochemical and functional studies.
The ability of ZapA to promote the formation of FtsZ bundles in the presence of MipZ implicates ZapA as a regulator of FtsZ filament assembly. Various proteins have been shown to stimulate formation of FtsZ rings, and Caulobacter ZapA shares some common features with these factors. Like ZipA, ZapB, SepF and FzlA (Gueiros-
Filho & Losick, 2002; Goley et al.; RayChaudhuri, 1999; Singh et al. , 2008;
Ebersbach et al. , 2008b; Goley et al. , 2010b), ZapA reduces the GTPase activity of
FtsZ (data not shown). However, it is unclear if Caulobacter ZapA promotes formation of bundled FtsZ structures in a similar pattern as that observed for ZapA in
B. subtilis , since no curved FtsZ conformation promoted by ZapA could be detected, at least in the conditions we tested. Specifically, Goley et al. showed that FzlA stimulates the formation of highly curved FtsZ filaments in vitro , suggesting FzlA as a main regulator of FtsZ filament bending in Caulobacter . This indicates that the ability of cytoskeletal regulators to organize FtsZ polymers into distinct superstructures might show significant differences in different species.
138 In addition to ZapA and FzlA, Caulobacter has yet two more non-essential cell division factors called FzlC (Goley et al. , 2010b) and CCNA3357. FzlC is recruited to the divisome in an FtsZ-dependent manner (Goley et al. , 2010b). However, cells grew with normal rates and morphology in the absence of FzlC under normal laboratory conditions. On the other hand, the ccna3357 gene is usually present immediately upstream of zapA on the chromosome, and CCNA3357 protein localizes to midcell.
Preliminary data suggests that CCNA3357 might interact with FtsZ directly based on a microscopy-based assay for identification of Caulobacter FtsZ-binding proteins
(Goley et al. , 2010b).
Caulobacter is the only bacterium for which there is clear evidence for cell cycle control of FtsZ level. How do regulatory mechanisms operate to ensure that FtsZ will assemble and function at the right time during cell cycle progression? In order to answer this question, we should take into account the known factors that manipulate
FtsZ dynamics and structure operating in growing cells. Even though we know a number of proteins that affect FtsZ assembly, the details of how they function physiologically remain largely unclear. Future work will address the regulation of FtsZ conformation, and the question of how FtsZ structure relates to the constrictive force generation. It is important to characterize the full complement of FtsZ regulatory proteins in a given organism. Ultimately, by understanding the factors that manipulate
FtsZ dynamics, it may help to piece together the exact mechanism driving FtsZ assembly dynamics in the cell.
139 Table 5: Strains and plasmids
Strain or Construction or plasmid Relevant genotype/description Source CB15N synchronizable derivative of Caulobacter crescentus (Evinger & Agabian, 1977) CB15 YC040 CB15N ∆zapA Electroporation of pYC15 into CB15N YC202 CB15N ∆zapA xylX:: PxylX -zapA Electroporation of pYC16 into YC040 YC225 CB15N zapA:: PzapA -zapA-mCherry Electroporation of pYC17 into CB15N YC226 CB15N ∆ftsZ xylX:: PxylX -ftsZ zapA:: PzapA -zapA- Electroporation of pYC17 mCherry into LS3702 YC227 CB15N zapA:: PzapA -zapA-mCherry Electroporation of pYC107 vanA:: PvanA –-ftsZ-yfp into YC215 YC099 CB15N ∆zapA vanA:: PvanA –-ftsZ-cfp xylX:: Pxyl –- Electroporation of pYC107 mipZ-yfp and pYC108 into YC040 YC114 CB15N xylX:: PxylX –zapA-tap Electroporation of pYC107 into CB15N YC115 CB15N xylX:: PxylX –ftsA-tap Electroporation of pYC107 into CB15N YC225 CB15N xylX:: PxylX –popZ-tap Electroporation of pYC107 into CB15N YC213 CB15N pP van-mipZ-Pxyl-zapA Electroporation of pYC107 into CB15N YC214 CB15N pP van-mipZ Electroporation of pYC107 into CB15N YC215 CB15N vanA:: PvanA –-ftsZ-yfp Electroporation of pYC112 into CB15N LS3702 CB15N ∆ftsZ xylX ::P xylX-ftsZ (Wang et al. , 2001)
140
Chapter 5
Discussion and Future Directions
141 Much of what we previously knew about cell division in bacteria has come from extensive studies with E. coli or B. subtilis that showed that molecular differences exist in the composition of the divisome across different bacterial families.
Hence, it is important that several model species are investigated in parallel. Our extensive knowledge of the Caulobacter cell cycle and and its developmental regulation and several properties of the cell, such as its inherent asymmetry, single
DNA replication per cell cycle, and the tight coordination between chromosome segregation with cell division make it an excellent system to study cell division.
The results presented in this thesis have furthered our understanding of how the activity of the Caulobacter divisome is regulated and integrated into the cell cycle.
Our results show that there are a series of stages and transitions in divisome assembly.
Knowledge of the temporal and spatial pathway for divisome assembly in Caulobacter confronts us with many questions about the process. An important future direction is to understand some of these questions to gain more insight into the mechanisms of cytokinesis.
For example, the mechanisms controlling the dynamics and disassembly of various divisome proteins have not been completely studied yet. Most cell division proteins play unidentified physiological roles during cytokinesis. Of particular interest are FtsE and FtsX, which are widely conserved homologs of the putative ABC transporters and FtsE is the cytoplasmic partner of FtsX. Furthermore, FtsE has been reported to interact with FtsZ directly in E. coli , although this remains to be confirmed in vitro . Using the TAP tag, I have shown that FtsZ co-immunoprecipitated with the
Caulobacter homolog of FtsE. Interestingly, overexpression of FtsEX simultaneously
142 is lethal for cells. Understanding the structure and underlying molecular details of the
FtsEX complex will yield information about how it binds to and regulates FtsZ.
Efforts to reconstitute the division machinery and its regulators with purified proteins in vitro will also increase our understanding of how the cell division is performed. Osawa et al. have demonstrated that the membrane-targeted FtsZ could tether itself to the liposome wall in vitro . These experiments have demonstrated the possibility for understanding of divisome function in this manner. In vitro reconstitution of “functional cell division machinery” could provide a complementary approach to live-cell studies by using cell-free extracts or purified proteins, leading to a direct understanding of biological processes.
Interestingly, the outer membrane composition of Caulobacter yields a highly negatively-charged membrane due to the absence of the phospholipids and the consequently high percentage of phosphatidylglycerol and cardiolipin (Contreras et al. ,
1978). Detailed protein truncation tests would reveal whether interactions within Tol-
Pal complex or with membrane phosphatidylglycerol and cardiolipin are disrupted.
Cascales et al . has found that the proton motive force was coupled to the interactions between TolA and Pal (Cascales et al. , 2000). Additional studies will be needed to test how energized TolA molecules reach and interact with Pal in the OM to drive the invagination of OM.
We phenotypically assessed cell division in each of the mutant strains we created by imaging cells expressing the mutant proteins by cryo-EM. These results have provided preliminary morphological results and allowed us to resolve at what
143 step in division is the process aberrant. Furthermore, photoactivated localization microcopy (PALM) or stochastic optical reconstruction microscopy (STORM) techniques (Betzig et al. , 2006; Hess et al. , 2006; Rust et al. , 2006), and three- dimensional superresolution microscopy (Pavani et al. , 2009; Shtengel et al. , 2009), have begun to expand our knowledge of the bacterial cell structure. Fu recently has investigated the spatial organization of the Z-ring, structure-function relationships, and cell-cycle-dependent regulation of the Z-ring by PALM (Fu et al. , 2010).
The work reported in this thesis has uncovered many elements of the mechanisms of divisome assembly process. However, the detailed understanding of how the multiprotein complex of the divisome is assembled needs further elucidation.
Ultimately, knowledge of the molecular details of the divisome and the functional roles of the individual protein elements will produce a more precise model of the division machinery
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