Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2013 Nov 1.
Published in final edited form as: Mol Microbiol. 2012 Sep 19;86(3):513–523. doi: 10.1111/mmi.12017

SURFING BIOLOGICAL SURFACES: EXPLOITING THE NUCLEOID FOR PARTITION AND TRANSPORT IN BACTERIA

Anthony G Vecchiarelli 1, Kiyoshi Mizuuchi 1, Barbara E Funnell 2,*
PMCID: PMC3481007  NIHMSID: NIHMS404497  PMID: 22934804

Abstract

The ParA family of ATPases are responsible for transporting bacterial chromosomes, plasmids, and large protein machineries. ParAs pattern the nucleoid in vivo, but how patterning functions or is exploited in transport is of considerable debate. Here we discuss the process of self-organization into patterns on the bacterial nucleoid and explore how it relates to the molecular mechanism of ParA action. We review ParA-mediated DNA partition as a general mechanism of how ATP-driven protein gradients on biological surfaces can result in spatial organization on a mesoscale. We also discuss how the nucleoid acts as a formidable diffusion barrier for large bodies in the cell, and make the case that the ParA family evolved to overcome the barrier by exploiting the nucleoid as a matrix for movement.

Introduction

All organisms must segregate and properly localize DNA to ensure faithful inheritance of genetic material. In prokaryotes, low-copy-number genomes, including chromosomes and plasmids, employ active segregation, or partition (or Par), mechanisms. Sequence analyses show that Par systems are ubiquitous in the microbial world. For plasmids, Par systems are the primary segregation machinery, making them an excellent model for studying this process. In essence, Par is a transportation system for DNA. Indeed, the partition ATPase (usually called ParA, and we use this terminology throughout) that provides the energy for transportation is related to other ATPases that also localize components within a bacterial cell. Here we review these partition systems with particular emphasis on ParAs, which use the bacterial nucleoid as a surface for biological transport. We will argue that they are members of a larger family of proteins that have evolved to position large substrates (DNA, protein machineries or assemblages) and use the nucleoid as a spatial cue within the cell.

Partition loci in bacteria are minimalistic, encoding only three components - a cis-acting partition site and two trans-acting proteins. The partition site marks the DNA target to be segregated, and it is specifically bound by one of the proteins (often called ParB) to form a partition complex. The second protein is an NTPase that uses ATP or GTP binding and hydrolysis to drive the segregation reaction. A useful classification scheme of partition systems was established according to whether the NTPase contains Walker ATP-binding motifs (ParA), or resembles eukaryotic actin (usually called ParM) or tubulin (TubZ) (Reviewed in Gerdes et al., 2010). The ParM system is well understood from studies of R1 plasmid, in which actin-like filaments of ParM push plasmids to opposite cell poles through a mechanism of insertional polymerization. Because of the similarities to tubulin, the TubZ systems are also thought to use a filament-based mechanism, but this has yet to be confirmed.

ParA-mediated segregation systems are the most prevalent class in the microbial world, but their mechanism is not established and is the subject of considerable research and debate. Two competing models have emerged; one where ParA functions as a nucleoid-spanning filament and another where ParA binds the nucleoid as dimers or small oligomers that function as part of a diffusion-ratchet mechanism. One of the central questions is whether ParAs act as cytoskeletal elements as do the actin- or tubulin-like systems. In light of recent evidence that the ‘cytoskeletal’ protein, MreB, does not form a continuous helical filament in vivo, we believe it prudent to re-evaluate observations of ParA behavior in this context. In addition, most previous models for ParA action do not explain the requirement for ParA’s DNA binding activity in the partition reaction. Here we summarize the debate about filamentation and argue that current evidence supports a model where ParA systems pattern via interactions with the bacterial nucleoid. In addition, we explore the role of the nucleoid in both promoting and restricting movement of large DNAs and protein complexes in the cell.

ParAs are members of a larger family of ATPases, defined by the specific variant of their Walker A sequences (Koonin, 1993). It includes MinD, an ATPase required for localizing the E. coli cell division machinery at mid-cell, and a growing list of ATPases involved in intracellular transport (see below). The similarities between ParA and MinD go well beyond the sequence level as suggested previously (Leonard et al., 2005b; Lutkenhaus 2012), however models do not highlight these shared features at the molecular level. We will expand on the common themes because we believe they are critical to understanding both mechanisms.

The debate about helices in vivo and filaments in vitro

MreB was the first actin-like protein reported to form a bacterial cytoskeleton made of long helical filaments that traverse the longitudinal axis of rod-shaped bacteria (Jones et al., 2001). This arrangement was proposed to be responsible for the rod-shaped morphology of several bacterial species. However several recent studies have challenged the contiguous and helical nature of MreB (reviewed in White and Gober, 2012). Using total internal reflection fluorescence microscopy (TIRFM), functional GFP fusions of MreB were observed to form discrete but dynamically rotating patches around the cell circumference (Domínguez-Escobar et al., 2011; Garner et al., 2011). MreB rotation did not depend on its own polymerization, but required the assembly of the peptidoglycan cell wall (van Teefelen et al., 2011). These analyses demonstrated how small MreB foci can be misinterpreted as a long helical filament by conventional epifluorescence microscopy. The helical patterns are likely streaks caused by the rapid movement of small complexes over long exposure times. Furthermore, electron cryotomography of six different rod-shaped bacterial species did not find any long helical filaments (Swulius et al., 2011). The latter approach has recently demonstrated that a YFP-MreB fusion forms filaments but native MreB does not (Swulius and Jensen, 2012). Indeed, Landgraf et al (2012) have shown that some fluorescent tags can produce anomalous aggregation (in this study, as foci) that can be misinterpreted as the pattern of the native proteins. The studies strongly suggest that MreB does not form a cell-spanning cytoskeletal structure and raise the likely possibility that many, or perhaps all, helical patterns (including those formed by some ParAs and MinD; see below) are the result of a protein’s concentrated diffusion within the accessible cytosolic volume between the inner membrane and the nucleoid, which can have a twisted ultrastructure (see below).

The evidence for helical structures of ParAs and MinD has also come from epifluorescence microscopy. ParA coils were limited to the nucleoid region of the cell (Adachi et al., 2006; Ebersbach and Gerdes, 2004; Fogel and Waldor, 2006; Hatano et al., 2007; Pratto et al., 2008; Ringgaard et al., 2009), and MinD coils extended from cell-pole to pole (Shih et al., 2003). These observations led to a “filament-pulling” partition model, in which ATP-bound ParA polymerizes into helical filaments. When a filament end encounters a ParB-bound plasmid, the reaction reverses into filament disassembly, which pulls the plasmid towards the nucleoid pole (Ringgaard et al., 2009). Perpetual polymerization/depolymerization cycles result in continuous relocation of plasmids. However, these activities have yet to be directly demonstrated. Other structures have also been observed by fluorescence microscopy. C. crescentus ParA forms linear, not helical, structures (Ptacin et al, 2010). It remains to be determined whether these accurately reflect the dynamic properties of the ATPase.

Several ParAs form linear filaments in vitro, which have been interpreted to represent the in vivo helical structures that wrap around the circumference of the nucleoid (Ebersbach and Gerdes, 2004; Hatano et al., 2007; Ringgaard et al., 2009). We note that MreB also formed long filaments in vitro (Reviewed in Shaevitz and Gitai, 2010). The conditions for polymerization of ParAs vary dramatically among Par systems (see Supplementary Table S1) and a common set of properties has yet to emerge. How the conditions for in vitro filament formation relate to ParA function in vivo remains to be clarified.

Par proteins exploit the nucleoid for plasmid movement

There is compelling evidence that the bacterial nucleoid is important for the partition reaction. ParAs possess non-sequence specific DNA binding activity. In vivo, the vast majority of nonspecific DNA (“nsDNA”) is in the form of the bacterial nucleoid. Several fluorescent versions of ParAs (P1 ParA, F SopA, pB171 ParA, pSM19035 δ, V. cholera ParAI, and B. subtilis Soj) form a variety of dynamic patterns that colocalize with the nucleoid (Ebersbach et al., 2005; Fogel and Waldor, 2006; Hatano and Niki, 2010; Hatano et al., 2007; Lim et al., 2005; Marston and Errington, 1999; Pratto et al., 2008; Quisel et al., 1999; Ringgaard et al., 2009). In the absence of their cognate ParBs, the dynamic patterns are lost but the nucleoid association remains. Plasmid movement is also restricted to within the nucleoid region by ParA (Derman et al., 2008; Ebersbach and Gerdes, 2004; Erdmann et al., 1999; Ringgaard et al., 2009). Finally, mutations that damage ParA’s DNA binding activity are defective for partition in vivo (Hester and Lutkenhaus, 2007; Castaing et al., 2008).

In vitro, ParA binds directly to nsDNA, and this activity requires or is enhanced by ATP for all ParAs studied (Bouet et al., 2007; Castaing et al., 2008; Hester and Lutkenhaus, 2007; Leonard et al., 2005a; Pratto et al., 2008; Ptacin et al., 2010; Vecchiarelli et al., 2010; Hui et al., 2010;). A C-terminal patch of basic residues has been implicated as the nsDNA-binding interface in several ParAs (Castaing et al., 2008; Hester and Lutkenhaus, 2007; Soberón et al., 2011).We have used TIRFM to directly visualize P1 ParA binding to nsDNA (Vecchiarelli et al., 2010). In the presence of ATP, ParA-GFP bound to DNA as many separate clusters. No extended ParA filaments were observed. By comparing the dynamics of nucleotide-induced structural changes in ParA to its DNA-binding kinetics, we identified an ATP-specific conformational change that was coupled to its DNA-binding activity in vitro and to partition activity in vivo. We called the nsDNA-binding form ParA-ATP*, which we believe represents the form of ParA bound to the nucleoid in vivo.

What happens when nucleoid-bound ParA and plasmid-bound ParB meet? Cognate ParBs and their binding sites are required for nucleoid patterning by ParAs, and this activity is driven by ParB stimulation of ParA ATPase activity (Autret et al., 2001; Barilla et al., 2007; Ebersbach et al., 2005; Lim et al., 2005; Marston and Errington, 1999; Pratto et al., 2008; Quisel et al., 1999; Ringgaard et al., 2009). The evidence suggests nucleoid patterning results from ParA release in the vicinity of the plasmid following a ParA-ParB interaction. However little is known about the species of ParA colocalized with its cognate partition complex prior to release. Recent examinations of the Par proteins from P1 (ParA and ParB) and pSM19035 (δ and ω) with nsDNA in vitro have provided clues on how a dynamic plasmid-nucleoid association can be promoted in vivo (Havey et al., 2012; Soberón et al., 2011). For both systems, a large DNA-bridging complex formed, whose properties support the idea that it represents a plasmid-nucleoid interaction. First, although complex assembly occurred on nsDNA, the cognate partition site significantly stabilized the association. Second, complex assembly required ATP binding, and disassembly was coupled to ParB stimulated ATP hydrolysis. In the P1 system, we called this complex the Nucleoid-Adaptor Complex or NAC (Havey et al., 2012). For pSM19035, the bridging-complex was interpreted as being capable of pairing newly replicated plasmids as well as forming a NAC-like complex that bridges plasmid and nucleoid (Soberón et al., 2011).

A direct role for the nucleoid in partition seems to be a feature that has evolved specifically in ParA-mediated partition systems. Actin-like and tubulin-like partition NTPases do not appear to colocalize with the nucleoid (Garner et al., 2004; Moller-Jensen et al., 2002; Salje et al., 2009). In vitro, actin-like ATPases have not been shown to have an affinity for nsDNA (Møller-Jensen et al., 2003) nor do they require an association with DNA for their partition functions in a cell-free system (Garner et al., 2007).

Similarities among Par and Min systems

Nucleoid patterning by ParAs is reminiscent of membrane patterning by the E. coli MinCDE system, which imparts positional information for cell division so that the septum forms at mid-cell (Reviewed in Lutkenhaus, 2007). The remarkable aspect of the Min system is its ability to inhibit division everywhere but at mid-cell as a result of the system oscillations on the membrane. MinD forms a complex with the cell division inhibitor, MinC, and recruits MinC to the membrane. Membrane binding by MinD requires ATP and is controlled by MinE, which stimulates MinD’s ATPase activity and releases MinD from the membrane. In the absence of its ATPase activity or MinE, MinD uniformly binds the membrane (Raskin and deBoer, 1999), akin to how ParAs uniformly bind the nucleoid when its ATPase activity or cognate ParB is removed.

Since ParAs and MinD were first grouped by their ATP-binding motifs, further comparisons yielded a remarkable number of shared features. The structures of MinD and all ParAs to date (MipZ of C. crescentus, Soj of T. thermophilus, pSM19035 δ, P1 ParA, TP228 ParF) show a high degree of similarity (Dunham et al., 2009; Kiekebusch et al., 2012; Leonard et al., 2005a; Pratto et al., 2008; Schumacher et al., 2012; Wu et al., 2011). ParA and MinD have weak ATPase activity that is stimulated by a support “matrix” (DNA and membrane, respectively), and by a stimulator protein (ParB and MinE, respectively). Both bind their respective matrix as ATP-bound dimers, and interaction with the stimulator is likely coupled to matrix release. Both interact with the N-terminus of the stimulator protein, and use C-terminal domains for matrix binding - an amphipathic helix for membrane binding by MinDs (Szeto et al., 2002), and a patch of basic residues for nsDNA binding by ParAs (Castaing et al., 2008; Hester and Lutkenhaus, 2007; Soberón et al., 2011). The last distinction allows MinD to pattern the membrane and ParAs to pattern the nucleoid.

Patterning via a diffusion-ratchet mechanism

The patterning mechanisms for ParA and MinD are likely similar although the support matrices differ. A key question is how the ATPases can dynamically and asymmetrically distribute on the support matrix on a mesoscale? In other words, following release from the matrix after ATP hydrolysis, how do ParA and MinD rebind several microns away instead of in the physical proximity from which dissociation occurred? Current evidence indicates that positional markers (ie a tether) are not necessary for patterning. Consequently, Min and Par systems are referred to as “self-organizing” because the dynamic behavior is solely a result of the interplay among the system components and a support-matrix.

A key feature of self-organization is energy consumption, such as nucleotide hydrolysis, which allows for dynamic behavior at steady state. For the P1 Par system, a key early step is the ATP-dependent conformational change necessary to convert ParA to its nsDNA binding form (Figure 1A) (Vecchiarelli et al., 2010). This step is very slow, which, in vivo, would create a time delay between the moment ParA releases from the nucleoid following ATP hydrolysis, and the reacquisition of its nucleoid-binding form. This delay is long enough for ParA to diffuse throughout the cell and the biochemical timing mechanism allows ParA to rebind any region of the nucleoid with equal probability rather than rebinding in the physical proximity from which it was released. ParB stimulation of ParA ATPase activity is localized to the vicinity of the plasmid, and coupled with the biochemical time delay in nucleoid rebinding by ParA, establishes mesoscale patterning of ParA on the nucleoid surface. Short-range release and long-range nucleoid rebinding sets up a continual redistribution of ParA, which we have proposed is the ATP-driven force for plasmid movement. In this “diffusion-ratchet” mechanism, the plasmid ratchets along the nucleoid towards a higher local ParA concentration (Figure 1); the plasmid essentially “surfs” across the nucleoid following a wave of ParA. We note that in this model the time delay is key, but it is not necessarily a conformational change in the ParAs of other systems. Any step following ATP hydrolysis and dissociation of ParA from the DNA, if slow, could fulfill this requirement. Finally, the model does not preclude a role for ParA polymerization (just not as a self-supporting filament). For example, the clusters of ParA observed binding to DNA in TIRFM experiments could be dimers, short polymers, or oligomers, and this cooperativity may play a role in the interaction of ParA with the nucleoid.

Figure 1. Diffusion-ratchet models for the Par and Min systems.

Figure 1

Open in a new tab

(A) ParA-mediated nucleoid patterning and plasmid transport. ParA-ATP* binds the nucleoid and ParB dimers load onto the plasmid (black squiggle). Interactions between ParB and nucleoid-bound ParA (ParA-ATPc) bridge the plasmid to the nucleoid. ParA ATPase activity is stimulated by ParB, clearing ParA-ADP from the nucleoid in the vicinity of the plasmid. ParA exchanges ADP for ATP (ParA-ATP) and there is a delay time during the conformational change that creates ParA-ATP*. The delay allows ParA to randomly diffuse before re-associating with the nucleoid. The continual redistribution of nucleoid-bound ParA drives plasmid movement. After replication, the plasmids segregate bi-directionally as they chase high concentrations of nucleoid-bound ParA in opposite directions.

(B) Min-mediated membrane patterning. MinD-ATP* binds the membrane. MinE binds MinD on the membrane (MinD-ATPc), which licenses MinE to also bind the membrane (MinE*). MinD ATPase activity is stimulated by MinE*, clearing MinD-ADP from the membrane. MinE* immediately re-associates with a neighboring MinD dimer. MinD exchanges ADP for ATP, dimerizes and potentially undergoes further conformational changes that allow MinD to diffuse before re-associating with the membrane. At the wave front, the MinE:MinD ratio is low. While MinD is being released, MinE persistently re-associates with adjacent MinD dimers; thus increasing the MinE:MinD ratio towards the wave rear. At the wave rear, there is no available MinD for MinE to re-bind so both proteins release.

(C) Persistent binding model for ParB. The ParA-ParB association produces ParA-ADP, which cannot bind the partition complex or nucleoid, and is consequently released into the cytoplasm. ParB on the other hand remains competent for immediate rebinding to another nucleoid-bound ParA dimer.

MinD/MinE patterning on lipid bilayers in vitro has provided a mechanistic framework to explain Min system oscillations (Ivanov and Mizuuchi, 2010; Loose et al., 2011; Loose et al., 2008). In vivo, MinD-ATP forms a polar zone on the membrane and recruits MinE (Figure 1B). At the cell-pole, the MinE/MinD ratio is low and MinE cannot counteract MinD membrane-binding. At the edge of the MinD polar zone (or the trailing edge of the wave in cell free systems), the MinE/MinD ratio builds to a point where MinE stimulation of MinD release dominates. A key feature in the patterning mechanism lies, as with ParA, in a critical time delay; the rebinding time differential between the ATPase and its matrix. Mathematical models have highlighted the importance of a time delay in MinD membrane binding activity, and the delay was hypothesized to result from nucleotide exchange by MinD (Huang et al., 2003; Meinhardt and deBoer, 2001). However, a biochemical basis for the delay has yet to be found experimentally. While the delay allows MinD to diffuse away, MinE can immediately rebind a nearby membrane-bound MinD (Loose et al., 2011). This “persistent binding” by MinE is consistent with the MinD/MinE co-crystal structure that suggests a “Tarzan” model where MinE swings from one membrane-bound MinD dimer to the next (Park et al., 2011). Persistent binding by the stimulator protein is an attractive addition to the ParA-mediated partition mechanism (Figure 1C). Persistent binding is favored by multiple ParBs bound to the plasmid cargo. Following ParB-stimulated release of ParA-ADP, it is also possible that ParB immediately binds an adjacent ParA-ATP dimer on the nucleoid. The fate of ParB after interaction with ParA is currently unknown, but persistent binding would explain how ParB senses the nucleoid-bound ParA gradient, and maintains a dynamic bridge between nucleoid-bound ParA and plasmid.

Important questions still remain. The filament-pulling and diffusion-ratchet models describe plasmid transport, but neither has been fully developed experimentally to explain how replicated plasmids are separated bidirectionally, or why longitudinal, as opposed to transverse, movement occurs. We anticipate that an improved understanding of the interplay between ParA and ParB at the partition complex will shed light on these aspects of the mechanism. Notably, helical structures are not required for Min patterning on a supported lipid bilayer, nor are they necessary to impart directionality in the diffusion-ratchet model of ParA-mediated partition (Figure 1A). In addition, the bacterial nucleoid is not simply an inert support for ParAs; its structure, surface, and dynamics must be considered in the mechanisms of transport. We next briefly explore some of the properties of the nucleoid that are important for partition and related transport processes.

The nucleoid-cytoplasm interface

Visualizing nucleoid structure and its interface with the cytoplasm under near-native conditions is essential to understanding how it could act as a support matrix for transport. The E. coli nucleoid contains a chromosome that is compacted ~ 1,000-fold or more (Holmes and Cozzarelli, 2000), and confined to only 25% of the intracellular volume of a roughly 1 µm by 3 µm rod-shaped cell (Zimmerman, 2006). Nucleoid morphology has been analyzed using cryoelectron microscopy of vitreous sections for several bacterial species (Reviewed in Eltsov and Zuber, 2006). During active growth, all nucleoids generally show a dense core with multiple arms projecting outward toward the cytoplasmic periphery. More recently, it has been shown that nucleoid ultrastructure can adopt a distinct coiled spiral organization in both gram-negative bacteria by cryo-electron tomography (Butan et al., 2010) and gram-positive bacteria by fluorescence microscopy (Berlatzky et al., 2008). The ordered core of the nucleoid is starting to take shape but highly dynamic regions, namely the nucleoid-cytoplasm interface, remain unresolved.

Does the nucleoid have a definable surface? How far into the cytoplasm can the nucleoid extend? How does DNA metabolism interfere with surface topology of the nucleoid? Ribosomes strongly concentrate in the cytoplasmic periphery, suggesting the nucleoid is a cellular “phase” (Butan et al., 2010; Cabrera and Jin, 2003; Lewis et al., 2000; Mascarenhas et al., 2001; Robinow and Kellenberger, 1994; Wang et al., 2011). Evidence also shows that the cytoplasm-nucleoid boundaries are dynamic (Zimmerman, 2006). For instance, upon transition to stationary phase or when treated with chloramphenicol, the nucleoid rearranges into spherical/toroidal bodies. Visualization of consistent patterns has proven difficult, but recent advances in high-resolution microscopy (Lee et al., 2011; Wang et al., 2011) show promise in resolving nucleoid structure and its interface with the cytoplasm.

Nucleoid exclusion

Passive diffusion is a key contributor to macromolecular movement in bacterial cells. Experimental diffusion coefficients have been measured between 0.2 and 10 µm2/s for proteins (10–100 kDa) in the cytoplasm of E. coli (Reviewed in Mika and Poolman, 2011). A general decrease of mobility with increasing protein size is observed. Macromolecular crowding in bacterial cells is extreme; high crowding lowers mobility, and reduces the reorganization of cellular components, but also favors self- and re-association of molecules. For instance, GFP (~27 kDa) can diffuse from one cell pole to the other in ~ 0.5 s in E. coli. As cytoplasmic volume decreases (for example, by changing salt concentrations), GFP mobility slows by several orders of magnitude and can even become compartmentalized (Konopka et al., 2009; Van Den Bogaart et al., 2007). Under hyperosmotic conditions, it has been postulated that the cell membrane pushes against the compacted nucleoid, forming a barrier in the cell that restricts macromolecular diffusion and results in separated pools of GFP. The high excluded volume of the bacterial cytoplasm in combination with the nucleoid acting as a formidable diffusion barrier are likely contributors to the anomalous diffusion of large intracellular bodies in bacteria. Therefore, understanding plasmid diffusion, especially that of very large plasmids has important implications for the partition mechanism.

Without a Par system, plasmids are mainly at the poles or in the cytosolic space between nucleoids. In other words, the plasmids are “excluded” from the nucleoid, which is the passive effect of the nucleoid as a diffusion barrier. In filamentous cells with increased cytosolic space, plasmids are found throughout the cytoplasm rather than at the poles, which is also consistent with a passive exclusion from the nucleoid (Ringgaard et al., 2009; Erdmann et al., 1999). Similar nucleoid exclusion has been observed for large protein complexes, such as the hybrid polyketide/nonribosomal peptide synthetase in B. subtilis that assembles into a 2.5 MDa organelle-like structure (Straight et al., 2007).

Nucleoid exclusion can passively create polar asymmetry. For instance, nucleoid exclusion of protein aggregates has recently been shown to play a key role in the mechanism of bacterial aging (Winkler et al., 2010). Over time, denatured proteins diffuse freely and associate with a nucleoid-excluded aggregate typically at a cell pole. Following cell division, one cell is burdened with the asymmetric inheritance of the majority of mis-folded proteins, while the other cell is “rejuvenated”. Through successive divisions, the diffusion barrier causes the mother to age and all daughters to be rejuvenated. It is possible that nucleoid exclusion also creates polar asymmetry in the inheritance of large plasmids lacking a Par system. Because most naturally-occurring bacterial plasmids are relatively large, nucleoid exclusion would be expected to exert pressure to evolve an active segregation system. However, a systematic study on the diffusion characteristics of differently sized plasmids, or large protein complexes, has yet to be performed.

Overriding nucleoid exclusion

In addition to DNA cargo, ParA-like proteins have been found to position a growing number of large protein machineries (Table I). Some have been called ParA, although we suggest this nomenclature (ParA) be restricted to “DNA partition” proteins and that other ATPases be named for function and phenotype. In the examples listed in Table I, dynamic patterning of a matrix (nucleoid or membrane) has not typically been examined, but we predict this property will show ATP-dependence as with ParA and MinD. In addition, the identity of a stimulator protein has not always been identified (most are called “orphan ParAs” without an obvious ParB-equivalent). As a recent example, R. sphaeroides PpfA has been shown to use the nucleoid to segregate and position cytoplasmic chemotaxis clusters in a manner that is reminiscent of ParA-mediated plasmid partition (Figure 2A) (Roberts et al., 2012). Intriguingly, the stimulator protein, TlpT, is a physical part of the chemotaxis cluster being transported, and although unrelated to ParBs, its N-terminus is essential for PpfA interaction and cluster transport.

Table I. ParAs involved in macromolecular complex transport.

Partial sequence alignment shows the conserved deviant Walker-box (grey) that defines the family.

Protein Deviant Walker-A Box
KGGXXK[ST]		 Positioned Substrate Intracellular
Position		 Positioning Reference
ParA P1 plasmid graphic file with name nihms404497t1.jpg P1 Plasmid Longitudinally Equidistant (Erdmann et al., 1999)
PldP C. glutamicum graphic file with name nihms404497t2.jpg Divisome Midcell (Donovan et al., 2010)
MipZ C. crescentus graphic file with name nihms404497t3.jpg Divisome Midcell (Thanbichler and Shapiro, 2006)
ParA S. elongatus graphic file with name nihms404497t4.jpg Carboxyzomes Longitudinally Equidistant (Savage et al., 2010)
PpfA R. sphaeroides graphic file with name nihms404497t5.jpg Chemotaxis Machinery Longitudinally Equidistant (Thompson et al., 2006)
BcsQ S. typhimurium graphic file with name nihms404497t6.jpg Cellulose Biosynthesis Complex One polea (Le Quéré and Ghigo, 2009)
VirC1 A. tumefaciens graphic file with name nihms404497t7.jpg T-DNA Nucleoprotein Complex One polea (Atmakuri et al., 2007)
ParC V. cholerae graphic file with name nihms404497t8.jpg Chemotaxis Machinery New Pole (Ringgaard et al., 2011)
FlhGc V. alginolyticus graphic file with name nihms404497t9.jpg Flagellar Machinery Flagellated Pole (Kusumoto et al., 2008)
CpaEd C. crescentus graphic file with name nihms404497t10.jpg Pilus Biogenesis Machinery Piliated Pole (Viollier et al., 2002)
WssJ P. fluorescens graphic file with name nihms404497t11.jpg Cellulose Biosynthesis Complex N/A (Spiers et al., 2003)
AgmE M. xanthus graphic file with name nihms404497t12.jpg The cellb N/A N/A
Open in a new tab
a

Localization towards the old or new pole was not differentiated.

b

Suggested to be involved in “Adventurous” gliding motility of the cell (Youderian et al., 2003)

c

FleN in Pseudomonas spp.

d

TadZ in A. actinomycetemcomitans

Figure 2. Positioning large bodies in bacteria.

Figure 2

Open in a new tab

(A) Without a tether, multiple substrates are separated and positioned longitudinally equidistant to each other. (B) With a polar tether (X), the substrate segregates and one copy is transported across the nucleoid and tethered to the opposing pole. (C) Without active transport, nucleoid exclusion passively maintains the substrate at one pole.

At first glance, few similarities among the positioned substrates are apparent: they can be DNA or protein, the biological processes in which they participate are diverse, and their final position site can be mid-cell, the poles, or longitudinally equidistant to each other. The one unifying feature however is size; they are all very large macromolecular complexes. We propose ParAs facilitate the transport of large bodies over and/or across the nucleoid using the nucleoid as a support matrix, and by a common type of patterning mechanism, as discussed above.

Recently, it has been proposed that ParAs that localize their cargo to the poles use a “landmark” mechanism that is different than the mechanism that positions cargo longitudinally equidistant to each other (Lutkenhaus, 2012). However we do not see these processes as mutually exclusive, and suggest that the basic ParA/MinD-driven motion can be similar for both cases. When a polar tether is present, the processes of segregation and positioning are separable, that is, the ATPase transports the substrate and the tether maintains position (Figure 2B). The clearest and most recent examples come from chromosome segregation studies in C. crescentus. After DNA replication initiates at the cell pole, one ParB-parS complex comes into contact with nucleoid-bound ParA (Ptacin et al., 2010). ParB stimulates ATP-hydrolysis, which releases ParA from the nucleoid while the ParB-parS complex ratchets across the nucleoid to the other pole. The ParB-parS complex and ParA are maintained at the new pole by the proteins PopZ and TipN, respectively (Schofield et al., 2010). Strikingly, when polar associations are damaged, ParA can redistribute on the nucleoid leading to system oscillations not unlike those observed for ParA-mediated plasmid partition. Removal of the ATPase, with or without a polar tether, results in nucleoid exclusion of the cargo (Figure 2C).

Summary

The increasing scope of positioned substrates directed by the ParA/MinD superfamily of ATPases has raised interest in understanding the novel mechanism of transport. The “surfing” mechanism is inherently different from the classical motor protein or the actin/microtubule-types of transport. Although we have focused on the ParA/MinD family, mechanistically related systems may involve unrelated ATPases. Indeed, ATP-driven DNA transposition systems have recently been proposed to position transposon DNA onto its target DNA via a diffusion-ratchet mechanism (Han and Mizuuchi, 2010). Current models recognize patterning on biological surfaces as a critical feature, but the question of how the ATPase interacts with its matrix to generate a pulling force for cargo movement will be a major subject for future study. In vitro reconstitution of bacterial DNA segregation will be vital in settling whether continuous ParA polymers and/or helical structures are indeed involved in the system dynamics. Most bacterial chromosomes encode par systems and virtually all have their cognate partition sites located near the origin of replication. It is attractive to speculate that origin segregation is similar to plasmid partition whereby the replicated origins “surf” on the nucleoid in a manner analogous to plasmids. Segregation of the remaining chromosome may follow the origin by other mechanisms.

Not all large intracellular bodies have dedicated mechanisms for active transport and, to our knowledge, a systematic study on the diffusion characteristics of “massive” bodies (Megadaltons or greater) in bacteria has not been performed. Recent studies on the mitochondrial nucleoid and its influence on the positioning of large complexes have also asked if size matters (Reviewed in Bogenhagen, 2011). We believe it does, and it is our expectation that surface-mediated biomolecular patterning will become an emerging theme for intracellular transport of large bodies in cells of all kingdoms of life.

Supplementary Material

Supp Table S1

Acknowledgements

We thank James Havey and Ling Chin Hwang for critical reading of this paper. This work was supported by grant 37997 from the Canadian Institutes of Health Research (to B.E.F.), and by intramural research fund for NIDDK, NIH, HHS, US Government (to K.M.).

Footnotes

Author Contributions

A.G.V., K.M., and B.E.F. wrote the paper.

Conflict of Interest

Authors declare that they have no conflict of interest.

REFERENCES

  1. Adachi S, Hori K, Hiraga S. Subcellular positioning of F plasmid mediated by dynamic localization of SopA and SopB. J Mol Biol. 2006;356:850–863. doi: 10.1016/j.jmb.2005.11.088. [DOI] [PubMed] [Google Scholar]
  2. Atmakuri K, Cascales E, Burton OT, Banta LM, Christie PJ. Agrobacterium ParA/MinD-like VirC1 spatially coordinates early conjugative DNA transfer reactions. EMBO J. 2007;26:2540–2551. doi: 10.1038/sj.emboj.7601696. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Autret S, Nair R, Errington J. Genetic analysis of the chromosome segregation protein Spo0J of Bacillus subtilis: evidence for separate domains involved in DNA binding and interactions with Soj protein. Mol Microbiol. 2001;41:743–755. doi: 10.1046/j.1365-2958.2001.02551.x. [DOI] [PubMed] [Google Scholar]
  4. Berlatzky IA, Rouvinski A, Ben-Yehuda S. Spatial organization of a replicating bacterial chromosome. Proc Natl Acad Sci USA. 2008;105:14136–14140. doi: 10.1073/pnas.0804982105. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Bogenhagen DF. Mitochondrial DNA nucleoid structure. Biochim Biophys Acta. 2012;1819:914–920. doi: 10.1016/j.bbagrm.2011.11.005. [DOI] [PubMed] [Google Scholar]
  6. Bouet J-Y, Ah-Seng Y, Benmeradi N, Lane D. Polymerization of SopA partition ATPase: regulation by DNA binding and SopB. Mol Microbiol. 2007;63:468–481. doi: 10.1111/j.1365-2958.2006.05537.x. [DOI] [PubMed] [Google Scholar]
  7. Butan C, Hartnell LM, Fenton AK, Bliss D, Sockett RE, Subramaniam S, Milne JLS. Spiral architecture of the nucleoid in Bdellovibrio bacteriovorus. J Bacteriol. 2010;193:1341–1350. doi: 10.1128/JB.01061-10. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Cabrera JE, Jin DJ. The distribution of RNA polymerase in Escherichia coli is dynamic and sensitive to environmental cues. MolMicrobiol. 2003;50:1493–1505. doi: 10.1046/j.1365-2958.2003.03805.x. [DOI] [PubMed] [Google Scholar]
  9. Castaing J-P, Bouet JY, Lane D. F plasmid partition depends on interaction of SopA with non-specific DNA. Mol Microbiol. 2008;70:1000–1011. doi: 10.1111/j.1365-2958.2008.06465.x. [DOI] [PubMed] [Google Scholar]
  10. Derman AI, Lim-Fong G, Pogliano J. Intracellular mobility of plasmid DNA is limited by the ParA family of partitioning systems. Mol Microbiol. 2008;67:935–946. doi: 10.1111/j.1365-2958.2007.06066.x. [DOI] [PubMed] [Google Scholar]
  11. Domínguez-Escobar J, Chastanet A, Crevenna AH, Fromion V, Wedlich-Söldner R, Carballido-López R. Processive movement of MreB-associated cell wall biosynthetic complexes in bacteria. Science. 2011;333:225–228. doi: 10.1126/science.1203466. [DOI] [PubMed] [Google Scholar]
  12. Dunham TD, Xu W, Funnell BE, Schumacher MA. Structural basis for ADP-mediated transcriptional regulation by P1 and P7 ParA. EMBO J. 2009;28:1792–1802. doi: 10.1038/emboj.2009.120. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Ebersbach G, Gerdes K. Bacterial mitosis: partitioning protein ParA oscillates in spiral-shaped structures and positions plasmids at mid-cell. Mol Microbiol. 2004;52:385–398. doi: 10.1111/j.1365-2958.2004.04002.x. [DOI] [PubMed] [Google Scholar]
  14. Ebersbach G, Ringgaard S, Moller-Jensen J, Wang Q, Sherratt DJ, Gerdes K. Regular cellular distribution of plasmids by oscillating and filament-forming ParA ATPase of plasmid pB171. Mol Microbiol. 2006;61:1428–1442. doi: 10.1111/j.1365-2958.2006.05322.x. [DOI] [PubMed] [Google Scholar]
  15. Ebersbach G, Sherratt DJ, Gerdes K. Partition-associated incompatibility caused by random assortment of pure plasmid clusters. Mol Microbiol. 2005;56:1430–1440. doi: 10.1111/j.1365-2958.2005.04643.x. [DOI] [PubMed] [Google Scholar]
  16. Eltsov M, Zuber Bt. Transmission electron microscopy of the bacterial nucleoid. J Struct Biol. 2006;156:246–254. doi: 10.1016/j.jsb.2006.07.007. [DOI] [PubMed] [Google Scholar]
  17. Erdmann N, Petroff T, Funnell BE. Intracellular localization of P1 ParB protein depends on ParA and parS. Proc Natl Acad Sci USA. 1999;96:14905–14910. doi: 10.1073/pnas.96.26.14905. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Fogel MA, Waldor MK. A dynamic, mitotic-like mechanism for bacterial chromosome segregation. Genes Dev. 2006;20:3269–3282. doi: 10.1101/gad.1496506. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Garner EC, Bernard R, Wang W, Zhuang X, Rudner DZ, Mitchison T. Coupled, circumferential motions of the cell wall synthesis machinery and MreB filaments in B. subtilis. Science. 2011;333:222–225. doi: 10.1126/science.1203285. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Garner EC, Campbell CS, Mullins RD. Dynamic instability in a DNA-segregating prokaryotic actin homolog. Science. 2004;306:1021–1025. doi: 10.1126/science.1101313. [DOI] [PubMed] [Google Scholar]
  21. Garner EC, Campbell CS, Weibel DB, Mullins RD. Reconstitution of DNA segregation driven by assembly of a prokaryotic actin homolog. Science. 2007;315:1270–1274. doi: 10.1126/science.1138527. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Gerdes K, Howard M, Szardenings F. Pushing and pulling in prokaryotic DNA segregation. Cell. 2010;141:927–942. doi: 10.1016/j.cell.2010.05.033. [DOI] [PubMed] [Google Scholar]
  23. Han Y-W, Mizuuchi K. Phage Mu transposition immunity: Protein pattern formation along DNA by a diffusion-ratchet mechanism. Mol Cell. 2010;39:48–58. doi: 10.1016/j.molcel.2010.06.013. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Hatano T, Niki H. Partitioning of P1 plasmids by gradual distribution of the ATPase ParA. Mol Microbiol. 2010;78:1182–1198. doi: 10.1111/j.1365-2958.2010.07398.x. [DOI] [PubMed] [Google Scholar]
  25. Hatano T, Yamaichi Y, Niki H. Oscillating focus of SopA associated with filamentous structure guides partitioning of F plasmid. Mol Microbiol. 2007;64:1198–1213. doi: 10.1111/j.1365-2958.2007.05728.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Havey JC, Vecchiarelli AG, Funnell BE. ATP-regulated interactions between P1 ParA, ParB and non-specific DNA that are stabilized by the plasmid partition site, parS. Nucl Acids Res. 2012;40:801–812. doi: 10.1093/nar/gkr747. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Hester CM, Lutkenhaus J. Soj (ParA) DNA binding is mediated by conserved arginines and is essential for plasmid segregation. Proc Natl Acad Sci USA. 2007;104:20326–20331. doi: 10.1073/pnas.0705196105. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Holmes VF, Cozzarelli NR. Closing the ring: Links between SMC proteins and chromosome partitioning, condensation, and supercoiling. Proc Nat Acad Sci USA. 2000;97:1322–1324. doi: 10.1073/pnas.040576797. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Huang KC, Meir Y, Wingreen NS. Dynamic structures in Escherichia coli: Spontaneous formation of MinE rings and MinD polar zones. Proc Natl Acad Sci USA. 2003;100:12724–12728. doi: 10.1073/pnas.2135445100. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Hui MP, Galkin VE, Yu X, Stasiak AZ, Stasiak A, Waldor MK, Egelman EH. ParA2, a Vibrio cholerae chromosome partitioning protein, forms left-handed helical filaments on DNA. Proc Natl Acad Sci USA. 2010;107:4590–4595. doi: 10.1073/pnas.0913060107. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Ivanov V, Mizuuchi K. Multiple modes of interconverting dynamic pattern formation by bacterial cell division proteins. Proc Natl Acad Sci USA. 2010;107:8071–8078. doi: 10.1073/pnas.0911036107. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Jones LJF, Carballido-López R, Errington J. Control of cell shape in bacteria: helical, actin-like filaments in Bacillus subtilis. Cell. 2001;104:913–922. doi: 10.1016/s0092-8674(01)00287-2. [DOI] [PubMed] [Google Scholar]
  33. Kiekebusch D, Michie KA, Essen L-O, Löwe J, Thanbichler M. Localized dimerization and nucleoid binding drive gradient formation by the bacterial cell division inhibitor MipZ. Mol Cell. 2012;46:245–259. doi: 10.1016/j.molcel.2012.03.004. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Konopka MC, Sochacki KA, Bratton BP, Shkel IA, Record MT, Weisshaar JC. Cytoplasmic protein mobility in osmotically stressed Escherichia coli. J Bacteriol. 2009;191:231–237. doi: 10.1128/JB.00536-08. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Koonin EV. A superfamily of ATPases with diverse functions containing either classical or deviant ATP-binding motif. J Mol Biol. 1993;229:1165–1174. doi: 10.1006/jmbi.1993.1115. [DOI] [PubMed] [Google Scholar]
  36. Donovan C, Schwaiger A, Kramer R, Bramkamp M. Subcellular localization and characterization of the ParAB system from Corynebacterium glutamicum. J Bacteriol. 2010;192:3441–3451. doi: 10.1128/JB.00214-10. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Kusumoto A, Shinohara A, Terashima H, Kojima S, Yakushi T, Homma M. Collaboration of FlhF and FlhG to regulate polar-flagella number and localization in Vibrio alginolyticus. Microbiol. 2008;154:1390–1399. doi: 10.1099/mic.0.2007/012641-0. [DOI] [PubMed] [Google Scholar]
  38. Landgraf D, Okumus B, Chien P, Baker TA, Paulsson J. Segregation of molecules at cell division reveals native protein localization. Nat Meth. 2012;9:480–482. doi: 10.1038/nmeth.1955. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Le Quéré B, Ghigo JM. BcsQ is an essential component of the Escherichia coli cellulose biosynthesis apparatus that localizes at the bacterial cell pole. Mol Microbiol. 2009;72:724–794. doi: 10.1111/j.1365-2958.2009.06678.x. [DOI] [PubMed] [Google Scholar]
  40. Lee SF, Thompson MA, Schwartz MA, Shapiro L, Moerner WE. Super-resolution imaging of the nucleoid-associated protein HU in Caulobacter crescentus. Biophys J. 2011;100:L31–L33. doi: 10.1016/j.bpj.2011.02.022. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Leonard TA, Butler PJ, Löwe J. Bacterial chromosome segregation: structure and DNA binding of the Soj dimer – a conserved biological switch. EMBO J. 2005a;24:270–282. doi: 10.1038/sj.emboj.7600530. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Leonard TA, Møller-Jensen J, Löwe J. Towards understanding the molecular basis of bacterial DNA segregation. Philos T Roy Soc B. 2005b;360:523–535. doi: 10.1098/rstb.2004.1608. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Lim GE, Derman AI, Pogliano J. Bacterial DNA segregation by dynamic SopA polymers. Proc Natl Acad Sci USA. 2005;102:17658–17663. doi: 10.1073/pnas.0507222102. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Loose M, Fischer-Friedrich E, Herold C, Kruse K, Schwille P. Min protein patterns emerge from rapid rebinding and membrane interaction of MinE. Nat Struct Mol Biol. 2011;18:577–583. doi: 10.1038/nsmb.2037. [DOI] [PubMed] [Google Scholar]
  45. Loose M, Fischer-Friedrich E, Ries J, Kruse K, Schwille P. Spatial regulators for bacterial cell division self-organize into surface waves in vitro. Science. 2008;320:789–792. doi: 10.1126/science.1154413. [DOI] [PubMed] [Google Scholar]
  46. Lutkenhaus J. Assembly dynamics of the bacterial MinCDE system and spatial regulation of the Z ring. Annu. Rev. Biochem. 2007;76:539–562. doi: 10.1146/annurev.biochem.75.103004.142652. [DOI] [PubMed] [Google Scholar]
  47. Lutkenhaus J. The ParA/MinD family puts things in their place. Trends Microbiol. 2012 doi: 10.1016/j.tim.2012.05.002. http://dx.doi.org/10.1016/j.tim.2012.05.002. [DOI] [PMC free article] [PubMed]
  48. Marston AL, Errington J. Dynamic movement of the ParA-like Soj protein of B. subtilis and its dual role in nucleoid organization and developmental regulation. Mol Cell. 1999;4:673–682. doi: 10.1016/s1097-2765(00)80378-0. [DOI] [PubMed] [Google Scholar]
  49. Mascarenhas J, Weber MHW, Graumann PL. Specific polar localization of ribosomes in Bacillus subtilis depends on active transcription. EMBO J. 2001;2:685–689. doi: 10.1093/embo-reports/kve160. [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Meinhardt H, de Boer PAJ. Pattern formation in Escherichia coli: A model for the pole-to-pole oscillations of Min proteins and the localization of the division site. Proc Natl Acad Sci USA. 2001;98:14202–14207. doi: 10.1073/pnas.251216598. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Mika JT, Poolman B. Macromolecule diffusion and confinement in prokaryotic cells. Curr Opin Biotech. 2011;22:117–126. doi: 10.1016/j.copbio.2010.09.009. [DOI] [PubMed] [Google Scholar]
  52. Møller-Jensen J, Borch J, Dam M, Jensen RB, Roepstorff P, Gerdes K. Bacterial mitosis: ParM of plasmid R1 moves plasmid DNA by an actin-like insertional polymerization mechanism. Mol Cell. 2003;12:1477–1487. doi: 10.1016/s1097-2765(03)00451-9. [DOI] [PubMed] [Google Scholar]
  53. Moller-Jensen J, Jensen RB, Lowe J, Gerdes K. Prokaryotic DNA segregation by an actin-like filament. EMBO J. 2002;21:3119–3127. doi: 10.1093/emboj/cdf320. [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Park K-T, Wu W, Battaile KP, Lovell S, Holyoak T, Lutkenhaus J. The Min oscillator uses MinD-dependent conformational changes in MinE to spatially regulate cytokinesis. Cell. 2011;146:396–407. doi: 10.1016/j.cell.2011.06.042. [DOI] [PMC free article] [PubMed] [Google Scholar]
  55. Pratto F, Cicek A, Weihofen WA, Lurz R, Saenger W, Alonso JC. Streptococcus pyogenes pSM19035 requires dynamic assembly of ATP-bound ParA and ParB on parS DNA during plasmid segregation. Nucl Acids Res. 2008;36:3676–3689. doi: 10.1093/nar/gkn170. [DOI] [PMC free article] [PubMed] [Google Scholar]
  56. Ptacin JL, Lee SF, Garner EC, Toro E, Eckart M, Comolli LR, Moerner WE, Shapiro L. A spindle-like apparatus guides bacterial chromosome segregation. Nat Cell Biol. 2010;12:791–798. doi: 10.1038/ncb2083. [DOI] [PMC free article] [PubMed] [Google Scholar]
  57. Quisel JD, Lin DCH, Grossman AD. Control of development by altered localization of a transcription factor in B. subtilis . Mol Cell. 1999;4:665–672. doi: 10.1016/s1097-2765(00)80377-9. [DOI] [PubMed] [Google Scholar]
  58. Raskin DM, deBoer PAJ. Rapid pole-to-pole oscillation of a protein required for directing division to the middle of Escherichia coli. . Proc Natl Acad Sci USA. 1999;96:4971–4976. doi: 10.1073/pnas.96.9.4971. [DOI] [PMC free article] [PubMed] [Google Scholar]
  59. Ringgaard S, Schirner K, Davis BM, Waldor MK. A family of ParA-like ATPases promotes cell pole maturation by facilitating polar localization of chemotaxis proteins. Genes Dev. 2011;25:1544–1555. doi: 10.1101/gad.2061811. [DOI] [PMC free article] [PubMed] [Google Scholar]
  60. Ringgaard S, Van Zon J, Howard M, Gerdes K. Movement and equipositioning of plasmids by ParA filament disassembly. Proc Natl Acad Sci USA. 2009;106:19369–19374. doi: 10.1073/pnas.0908347106. [DOI] [PMC free article] [PubMed] [Google Scholar]
  61. Roberts MAJ, Wadhams GH, Hadfield KA, Tickner S, Armitage JP. ParA-like protein uses nonspecific chromosomal DNA binding to partition protein complexes. Proc. Natl. Acad. Sci. USA. 2012;109:6698–6703. doi: 10.1073/pnas.1114000109. [DOI] [PMC free article] [PubMed] [Google Scholar]
  62. Robinow C, Kellenberger E. The bacterial nucleoid revisited. Microbiol Rev. 1994;58:211–232. doi: 10.1128/mr.58.2.211-232.1994. [DOI] [PMC free article] [PubMed] [Google Scholar]
  63. Salje J, Zuber B, Lowe J. Electron cryomicroscopy of E. coli reveals filament bundles involved in plasmid DNA segregation. Science. 2009;323:509–512. doi: 10.1126/science.1164346. [DOI] [PubMed] [Google Scholar]
  64. Savage DF, Afonso B, Chen AH, Silver PA. Spatially ordered dynamics of the bacterial carbon fixation machinery. Science. 2010;327:1258–1261. doi: 10.1126/science.1186090. [DOI] [PubMed] [Google Scholar]
  65. Schofield WB, Lim HC, Jacobs-Wagner C. Cell cycle coordination and regulation of bacterial chromosome segregation dynamics by polarly localized proteins. EMBO J. 2010;29:3068–3081. doi: 10.1038/emboj.2010.207. [DOI] [PMC free article] [PubMed] [Google Scholar]
  66. Shaevitz JW, Gitai Z. The structure and function of bacterial actin homologs. Cold Spring Harb Perspect Biol. 2010;2:a000364. doi: 10.1101/cshperspect.a000364. [DOI] [PMC free article] [PubMed] [Google Scholar]
  67. Shih Y-L, Le T, Rothfield L. Division site selection in Escherichia coli involves dynamic redistribution of Min proteins within coiled structures that extend between the two cell poles. Proc Natl Acad Sci USA. 2003;100:7865–7870. doi: 10.1073/pnas.1232225100. [DOI] [PMC free article] [PubMed] [Google Scholar]
  68. Schumacher MA, Ye Q, Barge MT, Zampini M, Barilla D, Hayes F. Structural mechanism of ATP induced polymerization of the partition factor ParF: implications for DNA segregation. J Biol Chem. 2012 doi: 10.1074/jbc.M112.373696. [DOI] [PMC free article] [PubMed] [Google Scholar]
  69. Soberón NE, Lioy VS, Pratto F, Volante A, Alonso JC. Molecular anatomy of the Streptococcus pyogenes pSM19035 partition and segrosome complexes. Nucl Acids Res. 2011;39:2624–2637. doi: 10.1093/nar/gkq1245. [DOI] [PMC free article] [PubMed] [Google Scholar]
  70. Spiers AJ, Bohannon J, Gehrig SM, Rainey PB. Biofilm formation at the air–liquid interface by the Pseudomonas fluorescens SBW25 wrinkly spreader requires an acetylated form of cellulose. Mol Microbiol. 2003;50:15–27. doi: 10.1046/j.1365-2958.2003.03670.x. [DOI] [PubMed] [Google Scholar]
  71. Straight PD, Fischbach MA, Walsh CT, Rudner DZ, Kolter R. A singular enzymatic megacomplex from Bacillus subtilis. Proc Natl Acad Sci USA. 2007;104:305–310. doi: 10.1073/pnas.0609073103. [DOI] [PMC free article] [PubMed] [Google Scholar]
  72. Swulius MT, Chen S, Jane Ding H, Li Z, Briegel A, Pilhofer M, Tocheva EI, Lybarger SR, Johnson TL, Sandkvist M, Jensen GJ. Long helical filaments are not seen encircling cells in electron cryotomograms of rod-shaped bacteria. Biochem Biophys Res Commun. 2011;407:650–655. doi: 10.1016/j.bbrc.2011.03.062. [DOI] [PMC free article] [PubMed] [Google Scholar]
  73. Swulius MT, Jensen GJ. The helical MreB cytoskeleton in E. coli MC1000/pLE7 is an artifact of the N-terminal YFP tag. J Bacteriol. 2012 doi: 10.1128/JB.00505-12. in press. [DOI] [PMC free article] [PubMed] [Google Scholar]
  74. Szeto TH, Rowland SL, Rothfield LI, King GF. Membrane localization of MinD is mediated by a C-terminal motif that is conserved across eubacteria, archaea, and chloroplasts. Proc Natl Acad Sci USA. 2002;99:15693–15698. doi: 10.1073/pnas.232590599. [DOI] [PMC free article] [PubMed] [Google Scholar]
  75. Thanbichler M, Shapiro L. MipZ, a spatial regulator coordinating chromosome segregation with cell division in Caulobacter. Cell. 2006;126:147–162. doi: 10.1016/j.cell.2006.05.038. [DOI] [PubMed] [Google Scholar]
  76. Thompson SR, Wadhams GH, Armitage JP. The positioning of cytoplasmic protein clusters in bacteria. Proc Natl Acad Sci USA. 2006;103:8209–8214. doi: 10.1073/pnas.0600919103. [DOI] [PMC free article] [PubMed] [Google Scholar]
  77. Van Den Bogaart G, Hermans N, Krasnikov V, Poolman B. Protein mobility and diffusive barriers in Escherichia coli: consequences of osmotic stress. Mol Microbiol. 2007;64:858–871. doi: 10.1111/j.1365-2958.2007.05705.x. [DOI] [PubMed] [Google Scholar]
  78. Vecchiarelli AG, Han Y-W, Tan X, Mizuuchi M, Ghirlando R, Biertümpfel C, Funnell BE, Mizuuchi K. ATP control of dynamic P1 ParA–DNA interactions: a key role for the nucleoid in plasmid partition. Mol Microbiol. 2010;78:78–91. doi: 10.1111/j.1365-2958.2010.07314.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  79. Viollier PH, Sternheim N, Shapiro L. A dynamically localized histidine kinase controls the asymmetric distribution of polar pili proteins. EMBO J. 2002;21:4420–4428. doi: 10.1093/emboj/cdf454. [DOI] [PMC free article] [PubMed] [Google Scholar]
  80. Wang W, Li G-W, Chen C, Xie XS, Zhuang X. Chromosome organization by a nucleoid-associated protein in live bacteria. Science. 2011;333:1445–1449. doi: 10.1126/science.1204697. [DOI] [PMC free article] [PubMed] [Google Scholar]
  81. White CL, Gober JW. MreB: pilot or passenger of cell wall synthesis? Trends Microbiol. 2012;20:74–79. doi: 10.1016/j.tim.2011.11.004. [DOI] [PubMed] [Google Scholar]
  82. Winkler J, Seybert A, Konig L, Pruggnaller S, Haselmann U, Sourjik V, Weiss M, Frangakis AS, Mogk A, Bukau B. Quantitative and spatio-temporal features of protein aggregation in Escherichia coli and consequences on protein quality control and cellular ageing. EMBO J. 2010;29:910–923. doi: 10.1038/emboj.2009.412. [DOI] [PMC free article] [PubMed] [Google Scholar]
  83. Wu W, Park K-T, Holyoak T, Lutkenhaus J. Determination of the structure of the MinD–ATP complex reveals the orientation of MinD on the membrane and the relative location of the binding sites for MinE and MinC. Mol Microbiol. 2011;79:1515–1528. doi: 10.1111/j.1365-2958.2010.07536.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  84. Youderian P, Burke N, White DJ, Hartzell PL. Identification of genes required for adventurous gliding motility in Myxococcus xanthus with the transposable element mariner. Mol Microbiol. 2003;49:555–570. doi: 10.1046/j.1365-2958.2003.03582.x. [DOI] [PubMed] [Google Scholar]
  85. Zimmerman SB. Shape and compaction of Escherichia coli nucleoids. J Struct Biol. 2006;156:255–261. doi: 10.1016/j.jsb.2006.03.022. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supp Table S1
NIHMS404497-supplement-Supp_Table_S1.pdf (68.1KB, pdf)

RESOURCES