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The Bacterial Cytoskeleton

Department of Molecular, Microbial and Structural Biology, University of Connecticut Health Center, 263 Farmington Avenue, Farmington, Connecticut 06032

SUMMARY

INTRODUCTION

BACTERIAL CYTOSKELETAL ELEMENTS

Actin Homologs

MreB and MreB homologs.

(i) Cytoskeletal organization of MreB proteins.

(ii) MreB polymerization and depolymerization.

(iii) Cellular functions of MreB and MreB homologs.

Plasmid partitioning by an actin homolog: the ParM system.

(i) ParM polymer assembly and disassembly.

(ii) Mechanism of ParM function.

(iii) Filament disassembly and plasmid migration.

MamK.

Tubulin Homologs

FtsZ.

(i) The FtsZ ring.

(ii) Membrane attachment of the Z-ring.

(iii) FtsZ spiral structures.

(iv) FtsZ polymerization and depolymerization.

(v) Regulation of Z-ring assembly and stability.

BtubA/B.

(i) BtubA/B polymerization.

Microtubule-like structures in Verrucomicrobia.

Intermediate Filament Protein Homologs

Crescentin.

The MinD/ParA Class of Bacterial Cytoskeletal Proteins

Subgroup 1: MinD. (i) The MinCDE system.

(ii) The MinD cytoskeleton.

(iii) MinD structure.

(iv) MinD polymerization.

(v) Membrane targeting of MinD.

(vi) MinD-bilayer interactions.

(vii) Dynamic rearrangements of the MinD cytoskeleton.

Subgroup 2: type I plasmid partitioning proteins.

(i) ParA/B proteins in plasmid partitioning.

(ii) ParA cytoskeletal structures.

(iii) ParA polymerization.

(iv) ParA oscillation.

Soj.

Other Filamentous Intracellular Structures

Spiroplasma melliferum fibrillar structures.

Treponema phagodenis cytoplasmic filaments.

Myxococcus xanthus intracellular filaments.

Mycoplasma pneumoniae filamentous structures.

Miscellaneous intracellular structures.

HELICES, HELICES, AND MORE HELICES

SetB

Sec Proteins

Tar

Outer Membrane Components

Why Is the Helical Distribution Pattern So Popular?

EUKARYOTIC AND PROKARYOTIC CYTOSKELETAL ELEMENTS

Properties and Functional Relationships

Membrane-Associated Cytoskeletal Structures

CONCLUSIONS AND SUMMARY

ACKNOWLEDGMENTS

REFERENCES

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INTRODUCTION

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It is truly breathtaking that within a period of 5 years a longstanding "truth" about one of the major life forms of the planet has been overturned. Although there had been previous indications of organized structures within bacterial cells (18, 139, 209), including the demonstration by Bi and Lutkenhaus that the tubulin homolog FtsZ forms a ring-like structure at the division site (15), the period of rapid advance began with the discovery by Jones et al. in 2001 that bacterial actin homologs in Bacillus subtilis are organized into extended helical structures that play key roles in cell shape regulation (95). Since then, the dictum that bacteria have no long-range internal structure has been replaced by the realization that the interior of the bacterial cell contains a large number of organized cytoskeletal elements, probably with a much larger group of cytoskeleton-associated components still to be discovered. Especially impressive is the large amount of information that has been revealed during this relatively short time span. It is now known that bacterial cytoskeletal homologs exist for all of the main groups of eukaryotic cytoskeletal proteins, i.e., the actin, tubulin, and intermediate filament (IF) groups. However, important differences between the eukaryotic and prokaryotic cytoskeletal systems have also emerged. These include the discovery that there is at least one major group of bacterial cytoskeletal elements, the MinD/ParA group, that has no known eukaryotic counterpart and the finding that analogous functions are frequently carried out by different cytoskeletal classes in eukaryotes and prokaryotes, often using different mechanisms.

The definition of the eukaryotic cytoskeleton has evolved over the past half century. It includes both stable filamentous structures that are composed largely of intermediate filament proteins and dynamic structures such as tubulin-derived microtubular structures and actin filaments that can assemble, disassemble, and redistribute rapidly within the cell in response to signals that regulate cellular functions such as cell cycle progression, intracellular organelle transport, motility, and cell shape. The common characteristics of these systems are their polymeric filamentous nature and their long-range order within the cell. Based on this background, we define bacterial cytoskeletal structures as filamentous structures that are based primarily on polymers of a single class of protein, that show long-range order within the cell, and, where this has been studied, that are capable of self-assembling in vitro into extended polymeric filaments. The bacterial cytoskeleton consists of the several groups of intracellular structures that meet this definition.

In this review we attempt to synthesize the wide range of information that is now available about the bacterial cytoskeleton, concentrating on systems where sufficient information is available to draw significant conclusions. We emphasize the cytoskeletal aspects of these systems and the relation of their cytoskeletal organization to specific cellular functions. We deal relatively briefly with some details of biological function that have recently been reviewed elsewhere or that may not involve the cytoskeleton. Where applicable, we refer the reader to recent reviews on these topics.

BACTERIAL CYTOSKELETAL ELEMENTS

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View larger version (52K): [in this window] [in a new window]   FIG. 1. Structural comparison of eukaryotic and prokaryotic cytoskeletal proteins. Protein structures were downloaded from the Protein Data Bank (PDB) (http://www.rcsb.org/pdb/) and aligned using the EMBL-EBI Dali server (http://www.epi.ac.uk/dali/). Protein structures were then analyzed and displayed using the MolMol program (103). Nucleotides and inorganic molecules that cocrystalized with the proteins are not shown. (A) Actin homologs. From left to right: Saccharomyces cerevisiae actin (PDB entry 1YAGA), ATP bound (218); Thermotoga maritima MreB (PDB entry 1JCG), AMPPNP bound (214); ParM of plasmid R1 (PDB entry 1MWM), ADP bound (216). (B) Tubulin homologs. From left to right: Bos taurus ß-tubulin (PDB entry 1JFFB), GTP/GDP bound (124); Methanococcus jannaschii FtsZ (PDB entry 1FSZ), GDP bound (122); Prosthecobacter dejongeii BtubA (PDB entry 2BTOA), GTP bound (183). (C) MinD (left) and Soj (right) proteins: Archaeoglobus fulgidus MinD (PDB entry 1HYQ), nucleotide free (24); Thermus thermophilus Soj (PDB entry 2BEJ), ADP bound (116). The topological specificity domain of the MinE dimer is shown (residues 31 to 88). The MinD-binding domains of each monomer extend from opposite sides (the right and left sides in this view) of the dimeric topological specificity domain (100).

We discuss here the best-studied bacterial actin-like cytoskeletal proteins, i.e., MreB, ParM, and MamK.

MreB and MreB homologs. MreB and MreB homologs are actin-related cytoskeletal proteins that play an important role in a number of cellular functions in bacteria, including regulation of cell shape, chromosome segregation, cell polarity, and organization of membranous organelles. Some bacterial species, such as Escherichia coli, contain a single MreB protein. Others contain two or more MreB-related proteins, such as the Bacillus subtilis MreB, Mbl (MreB-like), and MreBH (MreB homolog) proteins (1, 117, 217).

In many organisms, such as E. coli and B. subtilis, mreB is part of an operon that also contains genes coding for the MreC and MreD proteins. MreB, MreC, and MreD are required for cell viability (107, 113, 115). The loss of viability of E. coli mreB cells can be suppressed by modest overexpression of the essential cell division gene FtsZ (107, 188). It was suggested that this reflects a need for more FtsZ because of the larger volume of the spherical MreB-depleted cells (107). It is possible that the viable mreB mutant strains that have been isolated contain elevated cellular FtsZ levels due to secondary suppressor mutations.

(i) Cytoskeletal organization of MreB proteins. The cellular organization of MreB and MreB homologs in Escherichia coli, Bacillus subtilis, Caulobacter crescentus, and Rhodobacter sphaeroides has been described (53, 60, 95, 108, 187, 191). In all cases the proteins are organized into helical filamentous structures that coil around the rod-shaped cell, as shown by immunofluorescence microscopy and fluorescence microscopy of cells expressing fusions of the proteins to green fluorescent protein (GFP) or one of its derivatives, such as yellow fluorescent protein (YFP) (Fig. 2A and B). The extended coiled structures are located on the undersurface of the cytoplasmic membrane and frequently extend along the entire length of the cell. In some cases the structures consist of two intertwined helices (Fig. 2B).

View larger version (35K): [in this window] [in a new window]   FIG. 2. Localization and functions of the actin-like MreB protein. (A to D) Cellular localization of MreB labeled with GFP or YFP. (A to C) E. coli MreB localizes into extended coils (A), intertwined double helices (B), and band-like structures (C). (Reprinted from reference 187 with permission of the publisher. Copyright 2003 National Academy of Sciences, U.S.A.) (D) C. crescentus MreB localizes into a band-like structure at the division site in predivisional cells. (Reprinted from reference 60 with permission of the publisher. Copyright 2004 National Academy of Sciences, U.S.A.) (E) Chromosome segregation in E. coli mreB deletion strain (Y.-L. Shih, unpublished). Defects include multiple disordered chromosomes and unequal chromosome partition into daughter cells (cell 1), production of anucleate cells (cell 2), and chromosome fragmentation presumably due to guillotining of the nucleoid (cell 3). Bacterial DNA (red) is stained with DAPI; membrane (green) is stained with FM4-64 [N-(3-triethylammoniumpropyl)-4-(6-(4-(diethylamino)phenyl)hexatrienyl)pyridinium dibromide]. (F) Suggested organization of cytoskeleton-associated cell wall-synthesizing proteins of E. coli. The cytoplasmic MreB cytoskeleton is linked via MreC and MreD to the PBP murein biosynthetic enzymes (107). RodA may also be a member of the complex, and it is possible that some outer membrane proteins are also part of the MreC-associated structure (35). MreC is shown as a transmembrane protein (107, 113), but, at least in C. crescentus, MreC appears to be a periplasmic protein (35). See text for further details and references. This multiprotein structure may permit the MreB cytoskeleton to regulate the pattern of cell wall biosynthesis by providing spatial information to the murein biosynthetic machinery. OM, outer membrane; PG, peptidoglycan (murein); IM, inner membrane. (G) Pattern of murein assembly in B. subtilis (27). Nascent peptidoglycan was stained with fluorescently labeled vancomycin (see the text). The micrograph contains cells representing different stages of the cell cycle. Helical patterns of labeled vancomycin are indicated by transverse bands (lines) and dots around the cell periphery (arrowheads). The densely stained bands (arrow) in the two right-most cells are located at the cell division sites. (Reprinted from reference 27 with permission from Elsevier.) (H) Pattern of murein insertion into the murein sacculus of E. coli, suggesting a helical mode of murein assembly (arrows indicate two of the helical coils). (Reprinted from reference 32 with permission of the publisher.)

MreB redistribution into a ring structure near midcell in predivisional cells of C. crescentus is impaired, but not eliminated, in a strain that lacks TipN, a protein that normally marks the new cell pole and plays a role in establishment of the axis of cell polarity (83, 111). TipN overproduction leads to its mislocalization to ectopic positions along the cell cylinder, thereby establishing new axes of polarity as shown by outgrowth of rods as branches from the aberrant sites (111). It is possible that this reflects a direct influence of TipN on the directionality of the MreB cytoskeleton.

The MreB and Mbl cytoskeletal elements in B. subtilis are dynamic structures. Rapid movement has been observed within the GFP-labeled MreB and Mbl coiled structures, appearing sometimes to include segmental unidirectional movement detectable on a time scale of several seconds (194). Fluorescence photobleaching studies have also shown that molecules within the Mbl helical structure exchange with Mbl molecules elsewhere in the cell with a half time of approximately 8 min (20).

(ii) MreB polymerization and depolymerization. The MreB protein of Thermotoga maritima self-assembles into long polymeric filaments in vitro (51, 214). Although the cellular abundance of MreB in T. maritima is not known, polymerization of T. maritima MreB is rapid at the normal cellular concentrations of E. coli MreB (30,000 molecules per cell [108]) and B. subtilis MreB (8,000 molecules per cell [95]). Each filament is composed of two side-by-side linear polymers that differ in appearance from the helical double-stranded filaments of F-actin (214). The double-stranded MreB filaments are likely to comprise the helical MreB structures of intact cells.

Polymerization occurs equally well in the presence of ATP and GTP (50, 214), thereby differing from actin polymerization, which occurs only in the presence of ATP. MreB polymerization stimulates ATPase activity, but during the course of MreB polymerization there is a lag between polymerization and phosphate release (50). This implies that ATP hydrolysis occurs after MreB monomers are incorporated into filaments and that ATP binding, rather than hydrolysis, is required for addition of subunits to the growing polymer, thereby resembling actin polymerization (164).

The filaments interact to form bundles that undergo a gelation process, leading to formation of a solid-like structure (51). The bundled MreB structure is more rigid than the equivalent F-actin structure, with properties that are more often associated with eukaryotic intermediate filaments than with actin (51). These include high elasticity, low critical concentrations for polymerization, and a high propensity for bundling. These properties would be useful if the MreB cytoskeleton played a true skeletal role in supporting cell shape. However, the significant changes in cellular distribution of the MreB cytoskeleton that take place within the rod-shaped cells (see above) indicate that the rigidity of organized MreB polymers does not play an essential role in maintaining the rod shape of the cell. This conclusion is supported by studies with the drug A22 [S-(3,4-dichlorobenzyl)isothiourea] (88), which leads to loss of the MreB coiled structures and conversion of the cell from a rod to a sphere. When A22 was added, the rod-like shape of C. crescentus was not altered until long after disappearance of the MreB helical structures (61), arguing against the idea that rigidity of the bundled protofilaments is necessary for maintenance of cell shape.

All current information on the structure and polymerization properties of MreB filaments is based on studies of MreB from the thermophilic organism T. maritima, where the cellular organization of the protein has not yet been studied. Therefore, it will be important to confirm that MreB from organisms such as B. subtilis, E. coli, Caulobacter crescentus, and Rhodobacter sphaeroides, where almost all biological studies have been performed, behaves similarly to the T. maritima protein.

(iii) Cellular functions of MreB and MreB homologs. (a) Cell shape determination. Since the original discovery of the mreB (murein cluster B) gene in a search for mutants that are sensitive to amdinocillin (220), genes coding for MreB and MreB-related actin homologs have been shown to be present in almost all rod-shaped species and absent from species that grow as cocci (27). MreB-depleted cells usually grow as spheres, suggesting that MreB may play a role in cell shape control in rod-shaped bacteria.

It has long been known that the primary determinant of bacterial cell shape is the murein (peptidoglycan) exoskeleton, which is located outside of the plasma membrane. The murein sacculus retains the shape of bacterial cells even when purified away from other cellular components, and in the absence of cell wall murein, rod-shaped cells become spherical. This makes it clear that the sacculus is the shape-determining structure of the cell (184). The mreB operon is part of a large cluster of genes involved in murein synthesis, implying a possible relation between MreB and biosynthesis of the rigid murein exoskeleton.

The shape of rod-shaped cells is dependent on enzymes responsible for longitudinal murein growth. The rod shape is also dependent on the presence of MreB or MreB homologs, since depletion of these proteins leads to loss of the normal rod shape, with formation of spherical cells, or, in the case of loss of Mbl, to markedly deformed cells with large bulges and irregular increases in cell width (95). It has been suggested that Mbl controls cylindrical cell wall synthesis (27). Several other cellular proteins are also implicated in establishment of the rod shape, since loss of these proteins also leads a rod-to-sphere transition. Among others, these include the murein biosynthetic enzyme penicillin-binding protein 2 (PBP2) and the RodA protein of E. coli, which are discussed below.

It is likely that MreB and its homologs regulate the shape of rod-shaped cells by organizing murein biosynthetic enzymes into a helical pattern that is oriented along the long axis of the cell, leading to the pattern of murein synthesis that is responsible for the rod shape. This was initially suggested by studies with fluorescein-labeled vancomycin. Vancomycin blocks the cross-linking of newly synthesized glycan-pentapeptide chains into the murein sacculus by covalently attaching to the terminal D-alanine of the pentapeptide murein biosynthetic precursors (12, 17, 27). Fluorescein-labeled vancomycin therefore has been used as a marker for the cellular pattern of murein biosynthesis. The vancomycin studies showed that the immediate precursors of mature murein in B. subtilis (27) are organized in a coiled pattern that extends along the long axis of the cell and resembles the distribution patterns of MreB and Mbl (Fig. 2G). The coiled vancomycin pattern was dependent on the presence of the MreB homolog Mbl (27). In species that lack an mbl gene, it is likely that MreB or another MreB homolog carries out this function. Consistent with these results, studies of murein deposition in E. coli cells (32) also suggest a coiled pattern of new murein incorporation into the sacculus (Fig. 2H).

The following experiments indicate that the Mbl-dependent helical pattern of murein synthesis reflects a helical organization of the murein biosynthetic enzymes needed for longitudinal cell growth. First, the biosynthetic murein transpeptidase PBP2, which is required for rod shape, is distributed in a coiled pattern along the cell cylinder, similar to the distribution patterns of MreB and Mbl. This has been shown in C. crescentus (35, 38, 53) and is likely also true in E. coli (30). Second, the coiled distribution pattern of PBP2 is dependent on the presence of MreB and MreC (35, 53), implying that the helical MreB and MreC cytoskeletal structures (discussed further below) play an essential role in determining the cellular organization of PBP2.

MreC appears to act as a bridge between the MreB cytoskeleton and the murein biosynthetic machinery, as shown in studies of E. coli and B. subtilis (107, 113). The MreD protein is also likely to be a component of the bridging complex in organisms that contain an mreD gene, such as E. coli and B. subtilis. Thus, E. coli MreC interacts with MreB and MreD in bacterial two-hybrid assays, whereas MreB does not interact with MreD (107). This suggests that MreC may be intercalated between MreB and MreD in a putative multiprotein complex. In addition, several lines of evidence suggest that MreB, MreC, MreD, PBP2, and perhaps other murein biosynthetic enzymes are components of a structure that mediates the effects of the MreB cytoskeleton on the topology of murein synthesis (Fig. 2F). First, formation of normal MreB helical cytoskeletal structures requires the presence of mreC and mreD in E. coli and B. subtilis (29, 107). Second, several PBPs which code for murein biosynthetic enzymes were recovered by affinity chromatography of a C. crescentus cell extract on an MreC column, suggesting a link between MreC and multiple elements of the murein biosynthetic machinery in this organism (35). Third, C. crescentus MreC and PBP2 (35, 38) and B. subtilis MreC and MreD (113) are present in helical patterns that resemble those of the MreB and Mbl coiled elements. It has been suggested that RodA, which is required for maintenance of the rod shape and for the enzymatic activity of PBP2 (87, 196), may be another component of the MreBC complex (107). MreC has been thought to be the transmembrane link in this complex (107), because sequence analysis predicts a transmembrane organization. However, it has recently been reported that cell fractionation studies indicate a periplasmic location for the C. crescentus MreC protein (35). Further studies will be needed to clarify the cellular location of MreC in the various organisms under study.

The interaction between MreB and the MreC-based structure in vivo appears to be transient, since, in contrast to MreB, the distribution of the MreC helical pattern does not significantly change during the cell cycle (35). The observation that disruption of the MreB helical structures by A22 did not have immediate effects on the localization of MreC and PBP2 also supports the idea that MreB is not an essential part of the basic MreC-PBP complex (35, 38).

MreC affinity chromatography identified approximately 19 candidates to be MreC-associated proteins (35). These included eight presumed outer membrane proteins, nine cytoplasmic proteins, and small amounts of several PBPs. Studies of GFP derivatives of five of the outer membrane proteins showed clusters of labeled protein that were interpreted to indicate a spiral, punctate, or banding distribution pattern similar to that seen with MreC and PBP2 (35). Based on these results, it was suggested that the PBPs and outer membrane proteins might be part of an MreC-based complex anchored in the inner membrane that could provide a link between the internal MreB cytoskeleton and the outer layers of the cell envelope. If this is correct, the failure to recover MreB from the MreC affinity column might be attributed to problems in solubilization of the MreB cytoskeletal structures or to the presence of intermediate linking proteins between MreB and MreC. Further work will be needed to fully interpret these observations.

Interestingly, loss of either MreB or MreC causes PBP2 to lose its helical organization and instead to localize near midcell in C. crescentus (38). This requires the essential division protein FtsZ, suggesting that the cellular localization of PBP2, and presumably also its site of action, may be regulated by an interplay between the cell division machinery and the MreB/MreC cytoskeleton. This is consistent with the observation that PBP2 localizes to midcell at the time of septation in E. coli (30). The unrelated PBP2 of Staphylococcus aureus also localizes to midcell (161).

A full understanding of the role of the cytoskeleton in organizing the murein biosynthetic machinery is complicated by observations such as the following. (i) Localization studies of all 11 vegetative PBPs of B. subtilis failed to show a helical distribution (182). Unless these results reflect technical limitations, this implies that the association of murein biosynthetic enzymes with the cytoskeleton may vary from species to species or may be limited to a small subset of the enzymes. (ii) Growth of B. subtilis in the presence of 25 mM Mg²⁺ restored a normal rod morphology and normal helical distribution of nascent murein to mreB cells (54). Therefore, although there is inferential evidence that the MreB cytoskeleton may participate in determination of cell shape by providing a scaffold for the helical distribution of murein biosynthetic enzymes along the length of the cell, this effect either is indirect or operates through another scaffolding protein, perhaps MreC. High Mg²⁺ levels could stabilize the scaffolding partner to permit it to function in the absence of MreB. These observations show that the MreB cytoskeleton is not essential for cell shape determination in B. subtilis despite the fact that depletion of MreB results in a change of cell shape.

(b) Cell polarity. The poles of rod-shaped cells differ in several respects from the remainder of the cell body. These differences include the specific polar localization of a number of membrane-associated proteins (90), the presence of polar flagella or pili in certain species (185), the lack of turnover of murein and of externally labeled surface proteins at the cell poles (31, 33), the absence of zones of adhesion between the inner membrane and the murein-outer membrane layer at the poles (23, 131), and anatomic changes (the bacterial birth scar) at the newly formed cell pole (130). MreB has been implicated in one of these aspects of cell polarity, the localization of specific proteins to one or both cell poles.

Proteins that play a role in regulating the C. crescentus differentiation cycle are differentially targeted to one or both cell poles (reviewed in reference 59). These include the membrane histidine kinases, PleC, DivJ, and CckA. MreB is required for the polar targeting of these proteins (60). After depletion of MreB, polar localization of the proteins is lost and they become diffusely distributed within the cell.

MreB is also required for the polar localization of proteins in other organisms. In E. coli and related gram-negative bacteria, membrane-associated proteins involved in chemotaxis, motility, secretion, and virulence are normally targeted to one or both cell poles (185). When several of these proteins were expressed in E. coli, depletion of MreB led to a change of polar targeting. The proteins include the E. coli aspartate chemoreceptor (188), the Shigella flexneri virulence protein IcsA (151, 188), and the Vibrio cholerae type II secretion protein EpsM (151). It is likely that MreB also will be shown to play a role in the polar targeting of other proteins.

However, not all polar proteins require MreB for their localization. Thus, the assembly of the E. coli Min proteins into membrane-associated polar zones (188, 205) and the polar localization of the C. crescentus TipN protein (111) appear to be independent of MreB. These may be exceptions to the general rule that MreB is involved in localization of polar proteins, reflecting the special role that the Min proteins play in establishing the position of the cellular division site (175) and the special role of TipN in marking the cell pole and in determination of cell polarity (83, 111). It is not known how MreB accomplishes the polar targeting of proteins. The MreB helical filaments could participate directly in moving the proteins to the poles by providing tracks for the active translocation of specific cargo proteins, probably in collaboration with substrate-specific carrier and/or motor proteins. Segments of the MreB filaments might themselves move toward the poles (194) as part of the translocation process. Alternatively, MreB might act indirectly by positioning polar targets for protein localization. For example, the polar end of the MreB cytoskeleton might initiate the localization or organization of other polar components that would then act as polar binding sites for a family of substrate proteins. The targets need not be proteins, since both the specialized membrane lipid composition of the poles (141) and the presence of a segregated polar murein compartment (33) could contribute to the target sites.

(c) Chromosome segregation. During the normal cell cycle, daughter nucleoids move rapidly to opposite ends of the cell, leading to their equipartition into the two daughter cells (74). This process is perturbed when MreB is depleted in E. coli and C. crescentus (60, 108). This is manifested by the production of cells in which multiple nucleoids are irregularly distributed within the cytoplasm (Fig. 2E, cell 1) (108) and of anucleate cells (Fig. 2E, cell 2) or cells in which an incompletely partitioned nucleoid is guillotined by the septum (2E, cell 3). The possibility that the nucleoid segregation defects are caused by the spherical shape of the MreB-depleted cells has been excluded by the observation that expression of certain mutant alleles of mreB that do not interfere with the rod shape of the cell is still associated with nucleoid segregation defects (106, 108).

Further evidence that MreB is needed for normal chromosome partition came from studies of the segregation of the origin and terminus regions of newly replicated chromosomes of E. coli and C. crescentus (61, 108). In wild-type cells, the newly replicated oriC regions rapidly move to opposite ends of the cell. Separation of the terminus regions takes place later. In contrast, in MreB-depleted cells, the normal movement of the newly replicated oriC regions to opposite ends of the cell does not occur, and the terminus regions appear to adhere together (106, 108).

Biochemical evidence that the oriC region is the chromosomal target of MreB came from studies of C. crescentus cell extracts by Gitai and coworkers showing that MreB and DNA from the origin-proximal region could be chemically cross-linked and were coimmunoprecipitated with anti-MreB antiserum (61). This strongly implies that MreB is associated with the origin-proximal region of the chromosome, either directly or via other proteins that correspond to the centromere-binding kinetochore proteins of eukaryotic cells. These results imply that the association of the MreB cytoskeleton with the oriC region plays an important role in moving the daughter chromosomes towards opposite cell poles.

The mechanism responsible for the chromosomal movement is not known. Based on the present evidence, it appears likely that the oriC region directly or indirectly interacts with the MreB cytoskeleton and is then translocated along the helical MreB structures from midcell to the cell poles, perhaps accompanied by movement of segments of the MreB filaments (194). The energy requirement for translocation could be met by a separate motor protein, by coupling the ATPase activity of MreB to movement of the DNA, or by changes in chromosome folding. It is not known whether active movement of other chromosomal regions by a similar translocation mechanism is also involved in the segregation process. This may not be needed, since the active compaction of chromosomal DNA after oriC is moved to the poles could complete the process of nucleoid separation (147, 181). It also is not known whether the redistribution of MreB that occurs during the cell cycle is related to the oriC translocation events (53, 191).

The previous suggestion that RNA polymerase (RNAP) contributes to the motive force for chromosome separation (37) is supported by the recent finding that inactivation of RNAP in E. coli leads to a defect in nucleoid separation in DAPI (4',6'-diamidino-2-phenylindole)-stained cells, similar to the effect of MreB depletion or inactivation (106). This appears to be primarily associated with a failure of the terminus regions to segregate, although there may also be some effect on origin segregation. Of special interest, immunoprecipitation experiments using anti-MreB antibody indicated that MreB and RNAP are physically associated in cell extracts and in in vitro reconstitution experiments (106). These results suggest that RNAP is associated with the MreB cytoskeleton and works together with MreB in the chromosomal segregation process. It has been pointed out that a significant force can be generated by a stationary RNAP molecule during transcription, and this could contribute to moving DNA during the segregation process (37, 56).

A mechanism must also exist to explain the vectorial nature of the translocation process, in which the two newly replicated oriC regions are moved to opposite ends of the cell. The directionality could be provided by a double-helical MreB cytoskeletal structure (Fig. 2B) if the helical strands were of opposite polarity. This would provide two tracks that point in opposite directions, each providing a one-way highway for the oriC cargo. In another model the directionality would be imparted by the orientation of the newly replicated oriC regions as they exit from the replication apparatus (37). The two models are not mutually exclusive, and other mechanisms are also possible. A fuller understanding of the role of the MreB cytoskeleton in chromosome segregation will require a more detailed understanding of the macromolecular organization of the MreB helical arrays, the polarity of filament assembly and disassembly, and the basis of the MreB-oriC interaction. In contrast to the results for E. coli and C. crescentus, it is not clear whether MreB or one of the other actin homologs, Mbl or MreBH, is required for chromosome segregation in B. subtilis (54, 193). Thus, depletion of MreB in B. subtilis in the absence of polar effects on downstream genes did not lead to significant defects in chromosome segregation (54), and depletion of MreB or Mbl led to only mild segregation defects (194). Whether the third B. subtilis actin homolog, MreBH, influences chromosome segregation remains to be determined. It also will be of interest to see what mechanism is used for chromosome segregation in coccal species, which lack MreB homologs.

Plasmid partitioning by an actin homolog: the ParM system. Stable inheritance of low-copy plasmids is carried out by active partitioning systems that include a partitioning protein with ATPase activity. In type I partitioning systems these proteins belong to the Walker-type ATPase superfamily, and these are discussed later in this review. In type II systems the partitioning protein is an ATPase belonging to the actin protein superfamily, exemplified by the ParM protein of E. coli plasmid R1.

The par locus of plasmid R1 is the best-understood example of a bacterial system that fulfills the role of the mitotic apparatus of eukaryotic cells. The locus includes three genes: parM codes for the actin-like ParM protein (M for motor); parC is the cis-acting centromeric DNA site (C for centromere); and parR (R for repressor) codes for the ParR protein, which binds to parC sequences and acts both to autoregulate transcription of the par genes and to link the ParM motor protein to the plasmid DNA. The parC site contains 10 iterons of 11-base-pair repeats that act as sites for ParR binding (93).

In plasmid-containing cells, ParM forms linear filamentous structures that extend along the long axis of the cell, where they can be visualized by immunofluorescence microscopy (Fig. 3A, panel 1) (145, 146). The high cellular abundance of ParM (18,000 molecules per cell) suggests that each intracellular filament consists of a number of protofilaments (145, 146). Growth of the ParM filaments pushes the progeny plasmids to opposite ends of the cell (Fig. 3B), as discussed below.

View larger version (16K): [in this window] [in a new window]   FIG. 3. ParM-mediated partition of plasmid R1. (A) Fluorescence micrographs showing the ParM filament (green) and plasmid DNA (red). Panel 1 shows a ParM filament extending between two plasmids that are located at the ends of the cell. Panel 2 shows a plasmid at each cell pole, attached to a short ParM filament that extends toward midcell; this presumably represents an intermediate stage of filament disassembly after the plasmids have been delivered to the ends of the cell. (Reprinted from reference 145 with permission from Elsevier.) (B) After plasmid replication, growth of a ParM polymeric filament (yellow) pushes apart the two progeny R1 plasmids, moving them from midcell to the two poles. (C) Cartoon representing the boxed region in panel B, showing details of the addition of new subunits at the two ends of the ParM filament. The ParM-ADP filament is capped by ParM-ATP subunits. The capped end is attached to the parC centromeric region of the plasmid via the ParR protein. ParM filament growth occurs by insertion of ParM-ATP subunits at the interface between the end of the ParM filament and the ParR/parC complex. This is associated with hydrolysis of the ATP moiety of the previous ParM-ATP subunit, thereby converting the penultimate subunit to ParM-ADP.

ParM filament assembly and disassembly are regulated by ATP binding and hydrolysis (55, 145, 146). Filament assembly requires either ATP or a nonhydrolyzable ATP analog, indicating that the ATP-bound form of ParM is competent to polymerize (55, 93, 146). Unlike actin filaments, in which subunits are added only at one end of the filament, ParM filaments grow by addition of subunits at both ends of the growing polymer (55).

Polymer growth occurs by addition of a ParM-ATP subunit to the end of the ParM chain. This is associated with hydrolysis of the ATP moiety of the previously incorporated terminal ParM-ATP, explaining the polymerization-dependent ATPase activity of the system. This generates ParM-ADP as the new penultimate residue of the polymer (Fig. 3C). Thus, ParM filaments consist of chains of ParM-ADP, capped by the most recently added ParM-ATP subunits. ParM-ADP polymers are very unstable and have a high probability of disassembly unless capped by ParM-ATP (55).

A dramatic feature of the in vitro ParM polymerization system in the absence of ParR-parC is the sudden switch after a variable period of bidirectional elongation to a stage of rapid catastrophic disassembly from one end of the filament, presumably triggered by an increase in the rate of hydrolysis of the terminal ATP of the polymer (55). This reflects the fact that although the rate of subunit addition at the ends of the ParM filament is similar to the rate of growth of actin filaments, the rate of disassembly is 100 times higher than that of F-actin (55). A similar pattern of dynamic instability is characteristic of eukaryotic microtubules. The dynamic instability is thought to be necessary to prevent intracellular ParM filaments from growing to significant lengths before they interact with their plasmid targets (55).

(ii) Mechanism of ParM function. A substantial body of evidence indicates that ParR acts as a bridge between the ends of the ParM filament and the plasmid DNA that will be moved to the ends of the cell (Fig. 3B and C) (92, 94, 145, 146). Plasmid R1, like other unit copy plasmids, is replicated near midcell. ParR protein dimers then interact with the 10 iterons in the parC domains of the two daughter plasmids, leading to formation of plasmid pairs held together by multiple copies of ParR (94). The ends of a ParM oligomer or short polymer (55) then interact with the ParR-parC complex, separating the plasmid pair and leaving each end of the filament bound via ParR to a molecule of plasmid DNA (Fig. 3B). The polymer then grows bidirectionally by addition of new ParM subunits that are inserted between the end of the ParM filament and its ParR-parC cap, thereby pushing the two attached plasmids toward opposite ends of the cell (Fig. 3B and C). Addition of each new subunit is thought to be accompanied by hydrolysis of the ATP of the terminal ParM-ATP cap (Fig. 3C), as discussed above.

The presence of the ParR-parC complexes at the two ends of the ParM filament inhibits the dissociation of subunits during the period of polymer growth, preventing the catastrophic disassembly that otherwise would occur after a relatively limited period of polymer growth (55). Polymer growth stops when the ends of the filament approach the ends of the cell. The expected intermediate structure, a filament extending along the long axis of the cell with plasmid DNA attached at both ends, has been visualized in elegant double-label studies by Møller-Jensen et al. (Fig. 3A, panel 1) (145). The ParM polymers then disassemble, providing ParM subunits for initiating the cycle of filament assembly that will occur after plasmid replication at the midcell replication site in the daughter cell.

(iii) Filament disassembly and plasmid migration. The mechanism responsible for triggering disassembly of the ParM filament is not yet understood. We consider two possible modes of filament disassembly. (i) Disassembly proceeds bidirectionally from the two ends of the intact filament in response to a signal related to the arrival of the plasmids at the cell poles. To achieve this, the assembly/disassembly balance must be changed to favor disassembly from the ends of the filament. The switch from polymer growth to polymer disassembly may reflect a decrease in the rate of subunit addition as a consequence of low cytoplasmic ParM concentration resulting from its incorporation into the growing polymer as well as a diminution in synthesis of ParM because of the transcriptional autoregulation of the operon (41). This is likely to be associated with release of the ParR-plasmid complex, which normally inhibits subunit loss. It should generate cells containing a single ParM filament with two free ends. The filament will become progressively shorter as subunits are released from both ends. Although the predicted intermediate structures have not been detected, this could be explained if disassembly was too rapid for easy detection or if the stochastic disassembly of protofilaments left some individual pole-to-pole protofilaments intact through much of the disassembly period. (ii) In a second model, disassembly proceeds from midcell to each of the cell poles, initiated by internal cleavage of the filament. This would generate two filaments, each with a free end located somewhere near midcell and a plasmid-ParR complex at its polar end. Because the free ends lack a ParR-parC cap, the two filaments will disassemble rapidly from the free ends toward the cell poles. Cleavage might be carried out by proteins similar to eukaryotic factors that sever microtubules and actin filaments, such as katanins and cofilins, respectively (132, 137). Consistent with this model, the predicted intermediate structures, short ParM filaments each extending toward midcell from a plasmid located at a cell pole, have been visualized in double-label experiments (Fig. 3A, panel 2). At this point it is not possible to make a definitive choice between these two alternative disassembly mechanisms, each of which would presumably leave the plasmids in opposite ends of the cell.

The mechanism of movement of the segregated plasmids from the ends of the cell to their new replication sites has not been defined. After the ParM filament is disassembled, each released plasmid must move from the end of the cell to the new replication site, which will be located at midcell in the postdivision cell. During this process the freed plasmid must be prevented from diffusing back into the other half of the cell prior to septal closure. Although it is not known how this is accomplished, there are several possibilities. (i) The initiation of polymer disassembly could be coupled to an event in the cell cycle to ensure that the ParM filaments would not start to disassemble before septation occurred. (ii) A segregation trap could be present at the cell quarters to trap plasmids released at the proximal cell pole. The trap could, for example, be provided by the newly assembled plasmid replication machinery. This would require the replication apparatus to assemble at the cell quarters prior to septum formation at midcell, perhaps at the same positions where the cell division machinery assembles in rapidly growing cells (176). A plasmid diffusing from the proximal pole would encounter and engage with this site before crossing midcell. (iii) Cleavage of the ParM filament and subsequent disassembly of the filament (according to the second model in the paragraph above) could be directly or indirectly dependent on septation, consistent with the observation that ParM filaments do not disassemble when septation is blocked by cephalexin treatment (Fig. 2Ar in reference 146). This would ensure that plasmid release and diffusion from the pole would not occur until after septation occurred. In a simple model, filament cleavage could be a direct result of septum formation. Cleavage proteins might even be present at the leading edge of the ingrowing septum, where other functional proteins are located (48). (iv) The plasmids could associate with specific polar binding sites until a signal for their release is received. All of the above mechanisms, except for mechanical disruption of the ParM filament by the ingrowing septum, would require host cell factors.

MamK. MamK is an actin homolog that plays a role in the subcellular organization of magnetosomal membranes. This defines a new function of actin-like proteins, the positioning of cellular organelles in bacterial cells (101).

Magnetosomes are membrane-bounded organelles of Magnetospirillum magneticum sp. strain AMB01 that contain iron crystals within the membrane-bounded structures. Electron cryotomography has shown that the magnetosomal membranes represent membrane invaginations of the cytoplasmic membrane which are organized into linear arrays that are generally oriented along the long axis of the cell (101). The magnetosomes are bordered by filamentous MamK structures that extend along the length of the organellar structures near the inner curvature of the cell (Fig. 4A and B) (101). They have been visualized by electron cryotomography and by labeling cells with MamK fused to GFP. Deletion of mamK leads to both disappearance of the filaments and disappearance of the ordered magnetosome membrane arrays, confirming that the MamK cytoskeleton-like structures are required for magnetosome organization within the cell.

View larger version (51K): [in this window] [in a new window]   FIG. 4. Intracellular cytoskeletal structures formed by the actin-like MamK protein (A and B) and the intermediate filament homolog crescentin (C). (A and B) Electron cryotomography (A) and cartoon illustration (B) of magnetosomes located under the surface of the cytoplasmic membrane of Magnetospirillum magneticum. MamK filaments extend along the surface of the linear array of membrane-bounded magnetosome structures (white arrows in panel A). (The electron micrograph was reprinted from reference 101 with permission of AAAS.) (C) The intermediate filament-like protein crescentin (green), visualized by expressing GFP-tagged crescentin in wild-type cells, forms filamentous structures (white arrows) along the concave edge of C. crescentus. Membranes (in red) are stained with dye FM 4-64. (Reprinted from reference 8 with permission from Elsevier.)

Phylogenetic analysis suggests that FtsZ and tubulin are likely to have evolved from a common ancestor and to have diverged at an early stage of evolution, whereas BtubA and BtubB have evolved more recently following horizontal gene transfer of a tubulin or tubulin-like gene from a eukaryotic parent (91, 183, 192, 195). Consistent with this suggestion, BtubA and BtubB are more similar to tubulin than to FtsZ in amino acid sequence and protein structure, whereas other P. dejongeii genes are considered phylogenetically closer to their prokaryotic counterparts (91, 195). BtubA contains a carboxyl-terminal domain that resembles the carboxyl terminus of tubulin and is missing in FtsZ. This domain is important for the interaction of tubulin with motor proteins and tubulin-associated proteins.

FtsZ. FtsZ is essential for bacterial cytokinesis, and highly conserved FtsZ homologs are present in almost all bacteria and archaea. FtsZ homologs are also present in eukaryotic organelles such as plastids, which are believed to be derived from bacterial endosymbionts (for a review, see reference 5). A plasmid-borne FtsZ variant that plays a role in plasmid replication is also found in virulence plasmid pXO1 of Bacillus anthracis (155, 156) and in other megaplasmids (207). The role of FtsZ in cellular and organelle division has recently been reviewed (47, 133, 172).

(i) The FtsZ ring. FtsZ is an essential cell division protein in most bacteria. It is the earliest known component of the division machinery to be targeted to the cell division site, where it assembles into a circumferential ring, the Z-ring, located at the inner surface of the cytoplasmic membrane (Fig. 5D). The Z-ring has been visualized by immunoelectron microscopy (15) and by fluorescence microscopy using anti-FtsZ antibody or FtsZ linked to GFP or one of its derivatives (Fig. 5A) (3, 129). The Z-ring is believed to consist a number of polymeric FtsZ protofilaments, but it is not known whether individual protofilaments extend completely around the cell circumference or whether there are larger numbers of shorter polymers that extend around the cell in some type of staggered configuration.

View larger version (45K): [in this window] [in a new window]   FIG. 5. Cellular organization of FtsZ. (A to C) Immunofluorescence micrographs showing cellular localization of FtsZ in B. subtilis cells. (Reprinted from reference 13 with permission from Elsevier.) (A) FtsZ ring at midcell during vegetative growth; (B) extended FtsZ helical structures during the transition from vegetative growth to sporulation; (C) bipolar FtsZ rings at a later stage of the transition. (D) Diagrammatic representation of a portion of the cytokinetic ring (septasome) of E. coli. FtsZ is anchored to the cell membrane by FtsA and ZipA, and the other septasomal proteins are then added to the complex (63). The symbols representing the proteins are arranged for clarity, and the cartoon does not accurately represent the details of their interactions. OM, outer membrane; PG, peptidoglycan (murein); IM, inner membrane.

FtsZ is a high-abundance protein (reported to be present at 3,400 to 20,000 copies per E. coli cell) (26, 125, 149, 162, 178, 197). Of the other components of the cytokinetic ring, FtsA and ZipA are present at approximately 740 and 1,250 copies per cell, respectively (66, 178), whereas the other ring components are much less abundant, at 30 to 50 molecules per cell (176). The components of the septasome carry out a variety of functions in the septation process, most of which are poorly understood.

Interestingly, studies using fluorescence recovery after photobleaching have shown that the Z-ring is a highly dynamic structure, with FtsZ molecules exchanging with FtsZ molecules elsewhere in the cell with a half time of 30 seconds, depending upon experimental conditions (6, 197). It is not known whether the exchange of FtsZ molecules occurs along the length of the polymer or only at the ends of FtsZ polymers. Atomic force microscopy has indicated that FtsZ polymers undergo fragmentation and reannealing at internal locations (143). Therefore, it is possible that the exchange occurs at internal sites within FtsZ polymers, accompanied by the repeated breaking and resealing at internal sites within polymer strands. The potential for breaking and resealing of the polymer could also provide a mechanism for extrusion of FtsZ molecules during the shrinkage of the Z ring that occurs during septal constriction.

(ii) Membrane attachment of the Z-ring. Assembly of the Z-ring of E. coli requires the FtsA or ZipA protein to anchor the Z-ring to the membrane and possibly also to promote ring assembly or stability. Either protein is sufficient to support membrane association of FtsZ and formation of the Z-ring. Interestingly, although either FtsA or ZipA is capable of supporting Z-ring formation, both proteins are required for the subsequent entry of the other division proteins into the ring (63), and FtsZ, FtsA, and ZipA all remain as permanent components of the cytokinetic ring. FtsA homologs are found in many bacterial species, whereas ZipA is restricted to a small number of organisms. It is not known whether different proteins in other species fulfill the role of the E. coli ZipA protein.

ZipA contains a single hydrophobic transmembrane domain that anchors the Z-ring to the membrane (160). In contrast, FtsA anchors the Z-ring to the membrane via an amphipathic helix at the carboxy terminus of the FtsA protein, with the hydrophobic amino acid side chains on the nonpolar face of the helix inserting into the interior of the membrane bilayer (159). A similar amphipathic helix is present at the carboxyl termini of FtsA proteins of other species. The amphipathic helical membrane-binding domain of FtsA is similar in structure to and is interchangeable with the membrane-binding domain of MinD (159) (discussed below), demonstrating the relatively nonspecific nature of the membrane anchor. It is not known whether the Z-ring formed in the absence of one of the two anchoring proteins is structurally identical to the Z-ring formed when both membrane-binding proteins are present or whether FtsA and ZipA perform roles in the division process other than their roles in Z-ring anchoring and septasome assembly.

FtsA is an actin homolog, as shown by sequence and structural similarities (16, 215). The Streptococcus pneumoniae FtsA protein polymerizes in vitro in the presence of ATP (112), and removal of the E. coli membrane-targeting sequence is associated with formation of FtsA filaments within the cytoplasm (159). The ability of cytoplasmic FtsA to form polymeric filaments in vitro and the formation of FtsA filaments in E. coli suggest that FtsA may play a cytoskeleton-like role in the structure of the cytokinetic ring or in the septation process itself, in addition to its role in anchoring FtsZ to the membrane.

(iii) FtsZ spiral structures. In addition to constituting the Z-ring at division sites, FtsZ also forms transient helical arrays that coil around the long axis of the cell, resembling the helical structures formed by cytoskeletal proteins MreB, Mbl, MreBH, and MinD.

Evidence suggesting that the FtsZ spiral structures may be intermediates in formation of the FtsZ ring has come from studies of sporulating and vegetative B. subtilis cells (13). During sporogenesis the septation site switches from midcell to a site near one pole, requiring repositioning of the Z-ring from its normal midcell location. During this process, disappearance of the midcell ring is followed by the appearance of new Z-rings at both cell poles (Fig. 5C). One ring then disappears, and the other progresses to assemble the division machinery for formation of the polar spore septum (119). Ben Yehuda and Losick have shown that spiral FtsZ structures extend along the long axis of the cell during the change from medial to polar Z-rings (Fig. 5B), suggesting that these represent intermediates in the FtsZ redistribution process during sporogenesis (13).

Studies of a thermosensitive FtsZ mutant suggested that FtsZ spiral structures may be intermediates in formation of the FtsZ ring in vegetatively growing as well as sporulating B. subtilis cells (140). When grown at the nonpermissive temperature, FtsZ rings did not form and were replaced by short FtsZ spiral structures located in internucleoid regions along the cell. On shift back to permissive temperature, time-lapse microscopy of FtsZ-YFP showed rapid conversion of the spirals to Z-rings. On the basis of these observations it was suggested that the mutation causes a block in conversion of FtsZ spirals to FtsZ rings, implying that the spiral structures are precursors of the Z-ring.

The structural relationship between FtsZ spirals and Z-rings has not been established. The two structures may be independent, with FtsZ molecules leaving the spiral to form a ring structure at the division site. On the other hand, it is quite possible that the Z-ring is not a true ring but rather a tightly compressed spiral structure derived from the more extended FtsZ helical structures discussed above. A choice between these alternatives will require higher-resolution studies of Z-rings within cells and/or the isolation and characterization of the Z-ring itself.

It also has been shown that FtsZ helical structures are present in E. coli cells that overproduce FtsZ or FtsZ-GFP, a protein that is not fully functional but that is used as a marker for Z-rings and other FtsZ structures in living cells (129, 199). The coiled FtsZ arrays show periodic waves of oscillation with a periodicity of 30 to 60 s (205). This oscillatory behavior was unaffected by loss of the MreB helical cytoskeleton, but the periodicity was interrupted in a minCDE deletion mutant, in which division site placement is perturbed (205). The relationship of these observations to the behavior of the Min proteins, which are required for identification of the division site and which also are organized into oscillating helical structures (discussed below), remains to be defined.

(iv) FtsZ polymerization and depolymerization. The three-dimensional structure of FtsZ from Methanococcus jannaschii resembles the structures of - and ß-tubulins (Fig. 1B). There also are similarities in the polymerization characteristics of FtsZ and tubulins. In both cases the proteins polymerize unidirectionally into linear protofilaments in a GTP-dependent manner. Under appropriate conditions, the FtsZ and tubulin filaments both form bundles and sheets (45, 123). Rheometric measurements have indicated that bundles of FtsZ polymers form highly elastic structures, and it was suggested that this could be useful in maintenance of the Z-ring under the pressures generated during septal constriction (52).

However, there is no evidence that FtsZ forms microtubular structures in vitro or in vivo, and there are significant differences in the kinetics of nucleotide exchange between FtsZ and tubulin filaments. There also is no convincing evidence that FtsZ filaments are composed of FtsZ-GDP polymers capped with an FtsZ-GTP subunit whose hydrolysis leads to rapid polymer disassembly, as is the case for tubulin polymers. Instead, the polymer appears to consist of FtsZ-GTP subunits, and FtsZ polymer disassembly in vitro appears to be regulated by a balance between the rates of GTP hydrolysis and GTP rebinding to FtsZ along the length of the filament (82, 133, 142, 157, 172). This conclusion is based on the observation that hydrolysis of bound GTP leads to disassembly of FtsZ filaments unless sufficient GTP is available to exchange with the GDP product (reviewed in reference 172). Although changes in GTP/GDP ratio affect the rate of polymer disassembly, it is unlikely that this could explain the loss of FtsZ subunits from the Z-ring during septal constriction, because the high GTP/GDP ratio in the cytosol should provide sufficient GTP to prevent spontaneous filament disassembly. The important question of the mechanism whereby the Z-ring grows smaller during septal ingrowth remains open.

There is an energy barrier for the initiation and early steps of tubulin polymerization, providing a barrier to nucleation of new filaments (172). If this was true for FtsZ polymerization, it could provide a point for regulating protofilament formation during assembly of the Z-ring or the FtsZ spiral structures. It also is not known whether assembly or disassembly of FtsZ polymers within intact cells is modified by auxiliary proteins, as is true for microtubules in eukaryotic cells.

(v) Regulation of Z-ring assembly and stability. The assembly, stability, and function of the Z-ring and cytokinetic ring must be regulated both spatially and temporally during the normal division cycle. Spatial regulation restricts Z-ring formation to the desired site for later septum formation (reviewed in reference 175). This is accomplished primarily by the Min site selection proteins, which are organized into a cytoskeletal system that is discussed later in this review. Temporal regulation is required to ensure that septum formation occurs at the correct time in the cell cycle, after chromosome replication and segregation are completed. Control of cellular FtsZ protein concentration by regulation of transcription, translation, and/or degradation may play a role in regulating Z-ring formation in some cases, as, for example, during the cell cycle of C. crescentus (97, 165, 179). However, in most cases regulation is accomplished by the use of positive or negative effector proteins that regulate Z-ring assembly or stability. A number of effector proteins have been identified (Table 1) (reviewed in references 133 and 172), and others probably remain to be discovered. Thus far it is not known how these or other regulatory factors determine the timing of septation in the cell.

BtubA/B. A second group of bacterial tubulin homologs is exemplified by the BtubA and BtubB (bacterial tubulin) proteins. Unlike FtsZ, which is present in almost all bacterial species, BtubA and BtubB have thus far only been identified in the genus Prosthecobacter in the division Verrucomicrobia. The proteins from Prosthecobacter dejongeii have been studied in some detail. BtubA and BtubB are the only tubulin homologs in this organism, and electron crystallographic studies indicate that the three-dimensional structures of the BtubA monomer and BtubA/B heterodimer resemble those of tubulin (Fig. 1B) (183). Several lines of evidence, including similarities in polymerization properties, suggest that the BtubA/B system may provide a good model system for studying certain aspects of tubulin behavior and microtubule assembly.

(i) BtubA/B polymerization. BtubA and BtubB copolymerize in the presence of GTP into double-helical protofilaments and small rings (183, 192). Filament formation requires both proteins, although BtubB can self-assemble into 35-nm-wide rings in the absence of BtubA, whereas BtubA cannot polymerize by itself. The BtubA/B protofilaments self-associate into bundles that are sometimes organized around a central cavity, generating a microtubule-like structure. The distance between helical turns of the BtubA/B protofilament is similar to that of tubulin and FtsZ protofilaments (183). The cooperative mode of polymerization and the self-assembly of the polymers into tubular structures (192) are reminiscent of the assembly of microtubular structures from eukaryotic tubulin, which also are composed of heterodimer subunits (ß-tubulins). BtubA/B polymerization is also associated with an increase in GTPase activity, similar to the polymerization-dependent GTPase activation that occurs with both tubulin and FtsZ polymerization. The BtubA/B tubular structures are thicker than eukaryotic microtubules (40-nm versus 25-nm outside diameters) and are composed of two layers of protofilaments instead of a single layer. Stable polymers were also formed when BtubA and BtubB were coexpressed in E. coli cells, as shown by the formation of long, straight filaments that reacted with anti-BtubB antibody (192). Similar localization studies have not yet been done in P. dejongeii to confirm the likely supposition that the intracellular tubular structures are composed of BtubA/B. Since P. dejongeii does not contain an equivalent of the tubulin homolog FtsZ in the 95% of the genome that has been completed, it is conceivable that BtubA/B might also play a role in cytokinesis.

Microtubule-like structures in Verrucomicrobia. Ectosymbionts of marine hypotrich ciliates (Euplotidium species) have been characterized as Verrucomicrobia, the division that also includes P. dejongeii (see above), although they are distinct from P. dejongeii (158). During one stage of its complex life cycle, the ectosymbiont develops an extrusive apparatus designed to protect against predators. The apparatus is surrounded by a basket consisting of bundles of tubular structures which remain within the cell after the apparatus is extruded. The outside diameter of the tubules is 22 nm, and the diameter of the lumen is 13 nm (cited in reference 158), similar to the dimensions of eukaryotic microtubules (206). The tubules react with several monoclonal antitubulin antibodies and disassemble in the presence of the microtubule inhibitor nocodazole, supporting the idea that they are counterparts of eukaryotic microtubules (158, 173). The constituents of these tubular structures have not yet been identified, and it is not yet known whether the organisms contain Btub homologs.

The fact that microtubular structures are present in these organisms and the demonstration that the BtubA/B proteins of P. dejongeii resemble ß-tubulins in their ability to polymerize into tubular structures suggest that the verrucomicrobial systems may prove useful for the study of aspects of tubulin and microtubule behavior, especially if appropriate genetic systems can be developed.

Crescentin. Crescentin is the only cytoskeletal IF homolog thus far identified in prokaryotic cells. It is responsible for the shape of the comma-shaped organism C. crescentus (8) and was identified in a screen for C. crescentus transposon insertion mutations that affected cell shape. Loss of the structural gene for crescentin, creS, leads to a change in cell shape from comma to rod. The assignment of crescentin as an IF protein homolog is based primarily on its predicted protein domain organization. Crescentin has approximately 25% sequence identity and 40% similarity to eukaryotic IF proteins. However, it was pointed out that there are similar degrees of sequence similarity to non-IF proteins that contain extensive coiled-coil motifs (8). More specifically, crescentin and IF proteins share a predicted domain organization of four central coiled-coil structures with a characteristic discontinuity in the heptad repeat structure within the last coiled-coil segment (72).

Evidence that crescentin forms a cytoskeletal structure within the cell came from immunofluorescence studies showing that crescentin is present as an extended filamentous structure along the concave side of the C. crescentus cell (Fig. 4C) (8). This was confirmed in living cells by study of nonfunctional crescentin-GFP fusions which were coexpressed with untagged crescentin and by the observation that the curved shape of the cell is lost when crescentin is absent (8). These results suggest that the filamentous structure along the inner curve of the cell is a crescentin-based cytoskeletal element that establishes or maintains the comma shape of the cell.

The presumption that the curved cellular filamentous structures are composed of crescentin is supported by in vitro studies showing that purified crescentin rapidly self-assembles into long filaments. The self-assembly process does not require nucleotides or other cofactors, thereby resembling IF protein polymerization and differing from the polymerization of bacterial or eukaryotic tubulin or actin homologs (8). The crescentin filaments also resemble eukaryotic IFs in their fast assembly and viscoelastic properties, indicating that the crescentin network is solid-like and resistant to mechanical strains (Osigwe Esue and Denis Wirtz, personal communication). However, crescentin filaments diffe