The Tubulin Ancester, FtsZ, Draughtsman, Designer and Driving Force for Bacterial Cytokinesis

doi: 10.1016/S0022-2836(02)00024-4 available online at http://www.idealibrary.com on

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J. Mol. Biol. (2002) 318, 219–236

REVIEW ARTICLE

The Tubulin Ancester, FtsZ, Draughtsman, Designer and Driving Force for Bacterial Cytokinesis Stephen G. Addinall1 and Barry Holland2* 1 School of Biological Sciences University Manchester 2.205 Stopford Building Oxford Road, Manchester M13 9PT, UK 2

Institut de Ge´ne´tique et de Microbiologie Universite´ de Paris-Sud Baˆt 409, 91405 Orsay France *Corresponding author

We discuss in this review the regulation of synthesis and action of FtsZ, its structure in relation to tubulin and microtubules, and the mechanism of polymerization and disassembly (contraction) of FtsZ rings from a specific nucleation site (NS) at mid cell. These topics are considered in the light of recent immunocytological studies, high resolution structures of some division proteins and results indicating how bacteria may measure their mid cell point. q 2002 Elsevier Science Ltd. All rights reserved

Keywords: bacterial cytokinesis; Z ring; Min oscillation; FtsZ polymerization; division site selection

Introduction Both rod and coccal forms of bacteria grow and divide at varying rates and volumes according to growth conditions. In Salmonella typhinurium at least there is a clear relationship between length, surface area and growth rate.1 In Escherichia coli different strains may change their diameter or length according to growth rate.2 Nevertheless, the division process remains tightly regulated with a small coefficient of variation in size at division for a given growth rate.3 Moreover, division is coordinated with a highly efficient nucleoid (chromosome) segregation mechanism ensuring that daughter cells lacking DNA occur at a frequency less than , 0.03%4 per generation. Helmstetter & Cooper5 demonstrated in 1968 that E. coli B/r, following termination of a round of DNA replication, abruptly divided at a constant, D, 20 minutes later, over a wide range of growth rates. The nature of this constant D period has, however, remained obscure. How do cells therefore measure a doubling in mass whose absolute value differs with growth rate, assemble a division apparatus rather precisely at mid cell and then divide rapidly, producing daughter cells with new poles and an identical genetic complement? Although many of these questions still remain unanswered, it is clear that the key element in this Abbreviations used: NS, nucleation site. E-mail address of the corresponding author: [email protected]

division process is FtsZ, found in virtually all eubacteria, archae, in chloroplasts, and in some mitochondria. This protein has remarkable properties and surprisingly is a GTPase which is very closely related to the eukaryote cytoskeletal tubulin proteins (see below). Bacterial cell division research was dramatically revitalized in 1991 with the discovery by Bi and Lutkenhaus that FtsZ forms a cytokinetic ring at mid cell in E. coli.6 Earlier genetic studies had indicated that FtsZ was uniquely required for cell division and appeared to be involved in the “initiation” of this event.7 The Lutkenhaus laboratory (and subsequently many others) then used immunoflourescence techniques, pioneered in bacteria by Harry et al.,8 to study the properties of FtsZ and other cell division proteins. In this way, predictions about the cell biology of bacterial cell division arising from genetic experiments could be tested. From experiments like these combined with structural studies, fundamental elements of the bacterial cell cycle have been shown to be highly conserved. We shall review the properties and mechanism of action of FtsZ and how the assembly and disassembly of the FtsZ ring might be regulated. Regulating the action of FtsZ A major question concerning the regulation of cell division is whether FtsZ synthesis occurs throughout the cell cycle or in a burst at a

0022-2836/02/$ - see front matter q 2002 Elsevier Science Ltd. All rights reserved

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particular point in the cycle. It is also important to know when and how assembly and contraction of the cytokinetic Z ring is regulated. The amount of FtsZ in E. coli has been estimated to be about 15,000 molecules/cell,9 an amount which may be relatively constant irrespective of growth rate (and therefore size10). Division at a particular growth rate is delayed when the level of FtsZ is artificially reduced.11 – 13 However, when FtsZ is moderately overexpressed,14 division at mid cell does not apparently occur much earlier but rather occurs aberrantly at previously used division sites close to cell poles, over-riding the mechanism dependent on MinCD (see later), which normally forbids this. These results indicate that the supply of FtsZ can be limiting but that FtsZ concentration may not be a major factor in control of division. Thus, another factor(s), regulating the timing of the assembly or contraction of FtsZ rings may control the division process. Finally, it is interesting to note that calculations indicate that the 15,000 molecules of FtsZ could form up to 15 rings around the circumference of E. coli. However, it is not known whether all the FtsZ molecules are necessarily deployed at a division site. We shall consider first how the synthesis of FtsZ appears to be regulated quite differently in E. coli and Caulobacter crescentus, two closely related bacteria. Regulation of synthesis of FtsZ in E. coli The most detailed studies on transcriptional control have been performed in E. coli where the upstream fts genes together with ftsZ, form a highly conserved cluster of genes at two minutes on the E. coli chromosome. At least six promoters have been identified for ftsZ in the upstream genes.15 Various transcriptional regulators have been implicated in the control of these promoters, primarily located in the most proximal genes ftsA (pZ1,2,4), ftsQ (pR) and ddlB (pQ1,2), respectively. These regulators include SdiA (a possible quorum sensing system, operating on pQ2), RcsB (a signal transduction response regulator acting at the pZ promoters) and sigma S (a stationary phase regulator at pQ1). Most interestingly, pQ1,2, pZ2,4 have been shown to be more strongly expressed at slower growth rates and in stationary phase. Such promoters have been named gear box promoters with proportionately reducing expression as growth rate increases, and therefore are predicted to generate approximately constant amounts of FtsZ per cell, independent of growth rate. Consequently just enough FtsZ is produced per generation to make one division event.10,16 We have no idea at the moment of the nature of the signal which co-ordinates the synthesis of FtsZ, or FtsA which is required in a specific fixed ratio with respect to FtsZ,17,18 in this strict relation to growth. Interestingly, this mode of gear box regulation is precisely mirrored by that of the major porin in the outer membrane in E. coli.19 This ensures that porin remains at a constant

Review: FtsZ Regulation, Structure and Function

proportion of the surface membrane irrespective of growth rate since the surface area to mass of the bacteria is known to decrease in exactly the same way at faster growth rates. Regulation of FtsZ production may therefore be linked in some way, as is porin synthesis, to surface growth. There have been two reports of an increased (at least twofold) rate of ftsZ mRNA synthesis during the cell cycle. However, it has not been demonstrated that this translates into an increased rate of synthesis of FtsZ protein. Moreover, since FtsZ is reportedly quite stable in E. coli20 such a fluctuation may not affect the concentration of FtsZ significantly. These two studies unfortunately provided contradictory evidence for the origin of this mRNA, primarily pZ2 in one study20 or from promoters further upstream in the other.21 In fact, division still occurs when ftsZ is placed under the control of the constitutive tac promotor, although under these conditions division at the normal size apparently required 40% more FtsZ protein per cell.20 This suggests that the timing of division was affected in some perhaps subtle way just by changing the pattern of expression of FtsZ. FtsZ synthesis is normally dependent upon the housekeeping sigma70 factor although as cells enter stationary phase, when cells generally divide more frequently, sigmaS (RpoS) appears to be responsible for increased ftsZ expression from pQ1.22,23 In summary, despite extensive studies of the regulation of transcription of ftsZ, in the absence of any other evidence (in E. coli at least), we have to conclude that cells accumulate FtsZ molecules constitutively, with the concentration of FtsZ remaining constant. Alternative mechanisms have therefore to be considered to explain how the action of FtsZ is regulated through the cell cycle. These might include post translational modifications, localized sequestration of FtsZ to produce concentration changes, or fluctuations in the concentration of effector molecules. However, the available evidence24 – 26 indicates that there are no fluctuations (oscillations) in the concentration of any phosphorylated nucleotides, alarmones (like ppGpp), gene regulators (like cAMP) or enzyme co-factors (like ATP, GTP). Regulation of ftsZ expression in Caulobacter crescentus In C. crescentus, cell division normally takes place, dependent upon FtsZ, from a position near to mid cell. However, in this case division is considered asymmetric because this results in functionally different cells, a smaller swarmer cell and a larger, sendentary stalked cell. We invisage that in the normal environment the flagellated swarmer might spend an extensive period with a marauding existence without further growth until, presumably in response to extracellular signals, it undergoes a developmental change from swarmer to stalk cell, and re-enters the cell cycle. Thus, DNA replication

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Figure 1. Localisation of FtsZ in E. coli. W3110 wildtype (a) or ftsA12 ts (b) cells were processed for immunolocalisation of FtsZ (red) using sheep anti-FtsZ primary anti-bodies and donkey anti-sheep IgG (conjugated to Alexa594 (Molecular Probes)) secondary antibodies. DNA is stained blue with DAPI.

is then triggered and a new asymmetric division cycle initiated. New born stalk cells in contrast can immediately re-enter the growth cycle, triggering a new round of DNA replication and division to produce new stalk cells.27 However, whilst swarner cells have been analysed in some detail, information concerning the cell cycle in stalked cells is more limited. FtsZ, remarkably, is absent from swarmer cells with its synthesis switched off by binding of the transcriptional regulator, CtrA, to a ctrA box in the single promoter upstream of ftsZ.28 CtrA is a member of the super family of response regulators which normally function with a cognate membrane sensor kinase. In this case the action of CtrA is in fact controlled by phosphorylation involving at least three histidine kinases, DivL, CckA and the DivJ – DivK couple. DivJ, DivL and CckA localize to the membrane with two or more predicted transmembrane domains and presumably act as sensor kinases in signal transduction. However, the presumed extracellular cues being detected by these sensors are unknown. CtrA is in fact a master regulator, also negatively regulating DNA replication by binding to oriC and positively activating expression of the upstream ftsQ and ftsA, whilst repressing ftsZ itself.27 CtrA is specifically degraded during the transition from swarmer to stalk cell allowing re-entry of the stalk cells into the cell cycle (the replication cycle followed by division). CtrA is also degraded in the stalk precursor in the pre-divisional cell. In this way, particularly in swarmer cells, FtsZ protein is provided when and where it is required for division. Thus finally, FtsZ is no longer synthesized in the swarmer cell and the FtsZ protein is degraded in the final stages of division, accompanying contraction of the cytokinetic ring.28 The mechanism of cell cycle-dependent proteolysis of

FtsZ is unclear but was speculated by Kelly et al.28 to be linked to the oligomerization of FtsZ providing protection against protease attack. Whilst all these data demonstrate the importance of temporal control of FtsZ synthesis and instability in the development of swarm cells, it is not yet clear to what extent this also pertains to the stalk cell division cycle. In a more recent study, Quardokus et al.29 artificially regulated the expression of ftsZ in C. crescentus to show that even high levels of FtsZ in swarmer cells, failed to assemble into rings. This shows that additional controls must exist to block assembly. This study also indicated that under these conditions when swarmer cells finally re-entered the replication cycle and constricted to separate new born swarmer and stalk cells, these constrictions only occurred at the normal site, never at the poles to produce mini cells as in E. coli, when FtsZ is overproduced. Visualisation of the FtsZ ring in vivo Experiments using both indirect immunofluorescence30 (Figure 1) and GFP –FtsZ fusion proteins,31 have indicated that FtsZ ring formation is initiated at a single site on one side of the bacterium and appears to grow bidirectionally. It has also been demonstrated that assembly of Z rings in E. coli can occur in less than one minute,32 with a rate therefore corresponding to about 2 mm/ minute, similar to the rate of microtubule polymer growth in vitro.33 Microtubules, like actin filaments, assemble and disassemble as linear polymers, with only a strict head to tail association of subunits. The formation of in vivo “rings” of FtsZ formed by bidirectional growth and similar structures detected in vitro (see later), is therefore surprising. From the known structure of FtsZ subunits it is difficult to imagine how two monomers at the growing ends of two polymers could truly bond face to face. Such ends may therefore have to be stabilized by other means. Alternatively, Justice et al.34 have suggested the interesting idea that several protofilaments of FtsZ, stabilized by lateral cross-links between strands rather than forming closed circles, may constitute the actual ring-like structure observed in the microscope. Control and timing of FtsZ ring formation during the cell cycle From the earliest cytological studies it was presumed that FtsZ molecules (probably monomers, see Rivas et al.35), remained cytoplasmic until late in the cell cycle. However, several studies have indicated (based on immunofluorescence staining showing up to 90% of cells with a Z ring in rich media) that completed rings may be present for a considerable time before they are used in cytokinesis. This has been confirmed in a recent definitive study by Den Blaauwen et al.,33 who used immunofluorescence, combined with length

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Review: FtsZ Regulation, Structure and Function

Table 1. Modulators of FtsZ action Polymerization/ring stability

Synthesis of FtsZ SdiA22,144 RcsBp147 YycF (RR)a150 SigmaS (RpoS)15 Sigma-7015

þ þ þ þ þ

FtsA17,18,67 ZipA148 FtsW101 EzrAa45 SfiA47,51 Ca2þ 156 GTP138 PE (Post Ring)159 Ap4A149,161

Unknown point of action

Positioning 2 þ þ 2 2 þ þ þ /

CpxAa145 MinCDE55,67 Nucleoidb64 MinCD/DIV IVAa152

2 2 2 2

ppGpp15,146 gly tRNAsyn149 SAM151 H-NS/GalU130 tRNALeu6,3153 – 155 tRNASer1157 tRNASer2158 tRNAArg4160 cys tRNA synthetasea44 FlhD162 Acetyl phosphate163 CedA164 Peptide termination165 Fatty acids166

þ / 2 þ þ þ þ þþ 2 2 þ þ þ þ

Many factors influencing cell division in E. coli have been identified mainly through phenotypic effects of various mutations causing cell filamentation. Only a few of these are discussed in the text, of the others, YycF is an essential resonse regulator; CpxA is part of a signal transduction stress response system; SAM, S-adenosyl methionine; HNS-GalU, global regulators; ppGpp, Ap4A, alarmones; FlhD, transcription regulator; CedA, division regulator of unknown action. PE, phosphatidylethanolamine; þ or 2 indicates whether the effect is positive or negative on FtsZ; / indicates effect not characterized; a, refers to B. subtilis; b, nucleoid occlusion hypothesis, mutations, eg par, affecting the organization/packaging of the nucleoid or otherwise causing a delay in segregation of nucleoids may block or delay division through interference with FtsZ assembly or ring “contraction”.

distribution analysis at different growth rates, in two strains of E. coli where the timing of cell cycle events has previously been characterized in detail. In cells with a doubling time of 85 minutes, completed FtsZ rings were detected in approximately mid cycle, close to the time of termination of DNA replication and therefore co-incident with the commencement of the D-period under these conditions. The results clearly showed that FtsZ rings were always assembled well before nucleoids had fully separated and apparently 10– 20 minutes before constrictions could be detected. Interestingly, extrapolation to some earlier studies also indicated that the assembly of Z rings coincided with the previously observed increase in peptidoglycan synthesis required for the production of new hemispherical poles,36 – 38 an increase in cell length extension39 and a doubling in the rate of synthesis of outer membrane proteins.19 In C. crescentus, constriction of the FtsZ ring is similarly uncoupled from assembly and may take 30 minutes to occur after initial formation of FtsZ rings.29 These studies confirm therefore that we have to consider two distinct timing events: the triggering of assembly (polymerization) of Z rings and the much later activation of contraction of the ring through bending and disassembly or depolymerization (see below). So far we have no clues as to what these controls might be, but it is hard to believe that these involve fluctuations in the level of cellular GTP:GDP during the cell cycle. In searching for such control steps, many efforts have been exerted in attempts to tie the timing of the division cycle positively to the termination of the DNA replication cycle, but these have all failed to provide any convincing evidence for such a coupling. Indeed several findings clearly indicate

that this is not the case and the generally held view, summarized recently by Helmstetter40 still remains, as proposed by Jones & Donachie,41 supported by studies of Nordstrom et al.,42 that the DNA replication and division cycles overlap and run in parallel, with a veto generated in some way to prevent division before DNA replication is completed. On the other hand this issue of the coupling of replication to division was reopened recently,43 with the observation that inhibition of DNA synthesis by nalidixic acid or thymine starvation in a recA mutant (and therefore no induction of the division check point inhibitor, as part of the SOS response; see below), not only blocked division but was accompanied by a marked reduction in FtsZ synthesis. This is intriguing, but it should be noted that both these treatments may have many other secondary effects on cellular processes. Moreover, previous studies using UV-irradiation or mitomycin, for example, to inhibit DNA synthesis, have not detected any inhibition of cell division which is independent of the recA-controlled SOS system discussed in the next section.

Other possible factors affecting the frequency of division In E. coli in particular, many factors, including mutations in many genes not involved directly in the mechanism of cytokinesis have been shown to affect cell division. As shown in Table 1, mutations in many genes whose products include several proteins affecting segregation and or the packaging of nucleoids, molecular chaperones, elements of the protein synthesizing machinery, global regulators or enzymes involved in the synthesis of various phosphorylated nucleotides, all affect cell

Review: FtsZ Regulation, Structure and Function

division, giving rise to mutants with short or longer filaments as division is delayed, or in rare cases where division is actually brought forward.44 The possibility of the induced expression of the endogenous FtsZ inhibitor SulA (SfiA) (see below), resulting from perturbations linked to the mutations or other factors listed in Table 1, has been excluded in most cases. However, whether the disturbance of division in these cases affects the synthesis, assembly or contraction of FtsZ rings was usually not tested and even effects on later stages of cytokinesis, involving FtsA or other components of the septalsome (see below), have not been excluded. One exception is the protein EzrA in Bacillus subtilis, which negatively regulates FtsZ ring formation. In the absence of EzrA, extra FtsZ rings form at the poles without affecting the synthesis of FtsZ, whilst in depletion experiments, less FtsZ was apparently required to form a mid cell ring when EzrA was absent. EzrA co-localizes with the FtsZ ring, but the EzrA rings are completely dependent on the presence of FtsZ. Therefore, Levin et al.45 have postulated that EzrA may destabilize FtsZ polymers in some way, acting as a modulator of Fts function by controlling the rate of ring formation. However, taking appropriate note of genomic analysis, as well we should, it is unlikely that EzrA plays a very fundamental role in regulation of division, since its presence is restricted to a small minority of bacteria in the Bacillus/Clostridium group. Specific endogenous inhibitors of FtsZ In E. coli, SulA (SfiA) was identified as a specific inhibitor of FtsZ function, mediating a stressdependent division check point, as part of the SOS response, when DNA damage occurs. SulA is an 18 kDa protein, with an extremely short half-life when not complexed with FtsZ.46,47 The synthesis of SulA is tightly regulated, being normally repressed by LexA, and plays no role in the normal division cycle.48 DNA damage activates RecA, which in turn stimulates LexA autoprotease activity and the SOS regulon is induced.49 In vivo, synthesis of SulA, induced by UV-irradiation to inhibit DNA replication, immediately blocks division up to very late stages just preceding separation of daughter cells.50 This inhibition results in the failure of FtsZ to localize to rings and several studies, genetic and biochemical have confirmed that SulA interacts directly with FtsZ.51,52 Moreover, in vitro experiments have shown that SulA inhibits both the GTPase activity of FtsZ and the ability of FtsZ to form polymers.53,54 A recent study34 confirmed that the SulA protein inhibits dimerization and then oligomerization of FtsZ, both in vitro and apparently in vivo. MinCD forms a complex in E. coli which also inhibits the formation of FtsZ rings. However, the action of the MinC inhibitor (targeted to the membrane by MinD) is modulated by the topology factor MinE, which appears, through a protection

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mechanism, to permit FtsZ assembly at mid cell, whilst the MinCD complex is free to prevent division at other sites, in particular at the cell poles.55 Early genetic studies indicated that MinCD also interacted directly with FtsZ to inhibit its polymerization and one report indicates that a MalE – MinC fusion inhibits FtsZ oligomerization in vitro.56 However, another study34 comparing the action of SulA and MinCD in vivo, found distinctive mechanisms of action, SulA affecting polymerization of FtsZ whilst MinCD prevented the recruitment of FtsA into FtsZ rings. Interestingly, the high resolution structure of MinC57 revealed some structural similarity to FtsA, providing a basis for speculation that MinC and FtsA compete for binding to FtsZ. However, a recent study58 specifically re-investigating the effect of MinCD on Z ring formation could not confirm the results of Justice et al. In fact, Pichoff and Lutkenhaus provided further evidence to the contrary, showing that MinCD inhibits the in vivo assembly of Z rings.58 The MinC structure is composed of head to head molecules forming a dimer and Cordell et al.57 proposed that the dimer bound to a MinD monomer provides the specificity necessary for MinC to recognize FtsZ polymers and not monomers. More detailed consideration of the significance of the Min system is left to later. Other endogenous inhibitors of FtsZ are present in E. coli. SfiC is a close homologue of SfiA (SulA) and is encoded by a cryptic prophage.59 DicB is also encoded by a cryptic prophage and can substitute for MinC in inhibiting mini cell formation by blocking FtsZ-dependent division at the poles.60 How is the assembly of the FtsZ ring localized to mid cell? Cell division in E. coli is a remarkably accurate process, both with respect to average cell size in a given medium3,61 and localization of the mid cell division plane, deviating by less than 1%.62 As would be expected from these data, positioning of FtsZ rings at mid cell is also very precise.63 At least two distinct types of mechanism can now be envisaged for localizing the FtsZ ring to mid cell, although only one of these is apparently consistent with the high precision of mid cell selection normally observed. These mechanisms are not mutually exclusive. Woldringh and co-workers64 have proposed the nucleoid occlusion model, in which FtsZ ring formation over the nucleoid is somehow precluded, being permitted only after nucleoids have segregated and consequently vacates mid cell to allow FtsZ assembly. A second possible mechanism, more speculative, envisages the employment of a “measuring stick”, which measures lengths from each pole. Donachie and colleagues39 first provided evidence that in E. coli cell length is closely correlated with the onset of the D-period, leading to division. Koch65 proposed that the nucleoid, tethered to each end of the cell by newly replicated origins, formed such a

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symmetrical measuring device with the termini at “dead centre”. Note that this model does not escape the need for a specific “site”, in this case at the poles, for fixing each oriC region of the chromosome. A popular idea of a measuring stick for detecting mid cell, until recently the most speculative, envisages the synthesis of a microtubularlike structure from each pole, defining mid cell upon meeting. A variation on this (see Cook & Rothfield66) foresees that cell poles periodically produce an electrical or chemical signal transmitted along the membrane or through the cytoplasm, again meeting at mid cell. As indicated above such models simply displace the problem from delineating a specific site at mid cell to some form of unique site at the cell poles from which “signals” are generated. Although previously thought rather unlikely, as discussed below, such mechanisms may be the reality even if we do not understand their molecular basis. The nucleoid occlusion mechanism does not appear to have the necessary precision to account for the observed accuracy of mid cell selection in E. coli (see Koch,65 Margolin67 and Harry68). In addition, several studies have detected septa forming over the nucleoid, which ultimately guillotine the nucleoid in E. coli under certain conditions. In a study creating filaments with large nucleoid free zones, utilizing ftsA, dnaA ts double mutants, Cook & Rothfield66 found no correlation between septal localization and nucleoid position when division was restored at permissive temperature. Notably in this study the resuming division sites at permissive temperature were located a constant distance, approximately 5 mm, from a pole, irrespective of the distance to the nearest nucleoid. In particular, none were seen at the old division site at the pole. As pointed out by Cook & Rothfield66 many studies have also indicated the formation of FtsZ rings at mid cell or cell quarters, early in the cell cycle (see also Den Blaauwen et al.33), prior to nucleoid segregation. In B. subtilis when replication in outgrowing spores is blocked by thymine starvation, mid cell Z rings were observed by Regamey et al.69 to form over nucleoids. These authors, like Cook & Rothfield, favour a specific mid cell selection mechanism which normally provides for the nucleation of FtsZ assembly at the appropriate time. In these thymine-starved cells no replication of oriC occurs and the Z rings are in fact formed much earlier than normal in the cycle. This has led to the suggestion that a unique, mid cell site in either B. subtilis,69 or S. crescentus,29 designated NS (nucleation site) is initially masked by its prior occupation by the stationary replication complex which replicates the chromosome by a “spooling” mechanism, coupled to segregation and refolding of the nucleoid.70,71 The barrier to developing ideas and experimental approaches to the riddle of the mid cell FtsZ nucleation site, was recently finally breached by dramatic discoveries in fluorescence microscopy on the mechanism of action of the E. coli MinCDE

Review: FtsZ Regulation, Structure and Function

system in relation to regulation of FtsZ function. As described below, whilst different authors have interpreted these new results for or against a specific, unique assembly site at mid cell, all agree that the new findings on the astonishing dynamic (oscillating) cellular localization of MinD and MinE, have clearly established the idea that concentration gradients in principle can be the measuring stick that we have been searching for. MinCD proteins display an extraordinary oscillatory behaviour capable of “marking” a mid-cell site In E. coli, MinC, D encoded by the MinB locus, form a complex which appears to block the formation of FtsZ rings at the cell poles, at the ancient mid cell division sites, whilst MinE, encoded at the same locus, specifically prevents the action of MinCD at mid cell.55 MinD is a membrane-associated ATPase, related to the Par family of proteins involved in plasmid segregation, suggesting that MinD could be an ATP-dependent motor. MinD binds and targets MinC to the membrane. MinD enhances the anti FtsZ function of MinC by a mechanism dependent upon an active ATPase, which in turn is required for MinCD complex formation. Recent rather extraordinary developments in molecular cytology, utilizing MinDC or E fused to GFP, have shown that MinD and therefore its passenger MinC, repetitively oscillates from one end of the cell to the other, with a frequency of approximately 45 seconds at 30 8C (summarized by Hale et al.72). This oscillation depends upon MinE, which itself has now been shown to oscillate from one end of the cell to the other, paradoxically by a mechanism which depends in turn upon the presence of MinD.72,73 Initial studies of MinCD – Gfp fusions appeared to indicate that MinCD simply “accumulated” transiently at one cell pole before rapidly reappearing at high concentration at the opposite pole. Initial microscope studies of MinE –Gfp (without time lapse) on the other hand indicated that MinE formed a static ring, independently of FtsZ but close to mid cell. However, the most recent studies72 – 74 can be best interpreted as the formation of a dynamic MinD tubular structure close to the membrane, if not directly interacting with the membrane. At its maximum, this structure extends from one pole practically to mid cell before retracting/dissassembling back to the pole, followed by reassembly and extension to mid cell from the other pole. Min E does indeed primarily form a diffuse ring-like structure. However, as shown by time lapse studies the greater proportion of MinE appears to be closely associated with the rim of the MinD tubular structure, moving towards mid cell, then towards the poles, as the MinD tube assembles or retracts alternatively in each half of the cell. Although these data emphasize the seeming importance of cell poles, these apparently mobile structures are also seen at different positions, quite separate from the poles

Review: FtsZ Regulation, Structure and Function

when long filaments (blocked in FtsZ function) are analysed. Whilst these studies immediately raise many obvious questions concerning the mechanism of repetitive retraction and reassembly of these structures and the interdependence of MinD and MinE in their respective functions, the results have wider implications (developed below) for understanding the fundamental mechanism of mid cell site selection. First however, it is important to take a closer look at these MinD, MinE “structures”. Rothfield and co-workers have previously reported that E. coli contains approximately 3000 and 200 molecules, respectively, of MinD (30 kDa) and MinE (10 kDa).75,76 Clearly in this case neither molecule is sufficiently abundant alone to form the observed structures. Moreover, from studies of MinD, including a recent structural analysis,77,78 there is no indication of any capacity to oligomerize or indeed to interact with membrane. MinE on the other hand does form dimers79 but higher oligomers have not been reported. Obviously the small size and very low abundance of this protein precludes the formation of even a single protofilament capable of encompassing the cell diameter. One possibility is that what we actually see in the microscope are clusters of the dynamic Min proteins decorating an as yet unidentified cytoskeletal protein structure. After many years of unnecessary denial we have now to accept that rod-shaped bacteria do indeed possess a cytoskeleton, following the recent revelation that the conserved proteins MreB and Mbl in B. subtilis,80 form complex cytoplasmic macro-structures. Moreover, the high resolution structure of a homologue of MreB has demonstrated its remarkable resemblance to actin and to actin filaments.81 These findings now raise the possibility of these or other underlying structures providing the matrix for Min and other proteins specifically involved in cytokinesis. On the other hand the underlying matrix for binding dispersed clusters of Min proteins, which are then visible in the fluorescence microscope, might simply be the inner surface of the cytoplasmic membrane. In either case, the observed dynamics might be a property of the Min proteins themselves or the postulated matrix or both. Interestingly, a recent study by Hu and Lutkenhaus82 has shown that MinE in the presence of phospholipid vesicles stimulates the ATPase activity of purified MinD several fold. Moreover, with mutants of MinE defective in this stimulatory effect, there was a good correlation both with loss of function in vivo, and most importantly, with the total abolition or an increased period of the cellular oscillation of MinD structures. These results clearly link the hydrolysis of ATP by MinD to its oscillatory behaviour. Nevertheless, the mechanism which promotes disassembly and then reassembly in the opposite half of the cell remains an enigma.

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MinCD oscillation: a mechanism for identifying the site of FtsZ assembly? Returning to the broader significance of the oscillatory behaviour of MinCDE, de Boer and co-workers83,84 in particular, have proposed that the oscillation of MinCD could itself provide the measuring device for the differentiation of the mid cell point. This follows since the time integrated concentration of MinCD should be the lowest at mid cell. Thus, this by itself could be sufficient to permit FtsZ ring assembly at mid cell by creation of the NS. Attractive as this idea is there are some difficulties in accepting it. First, such a process of division control would be expected to be fundamental and therefore highly conserved. However, MinE is in fact absent from many bacteria, including B. subtilis. MinD moreover, has not been observed to oscillate from pole to pole in this organism (J. Errington, personal communication). Nevertheless, MinD itself is widely conserved in eubacteria and Archae and has been reported to be involved in division of chloroplasts in plants and algae85 and perhaps in some mitochondria,86 attesting to an indispensible role in regulating division at some level. Secondly, it is by no means clear that E. coli deleted for the min locus is defective in forming a unique mid cell NS for FtsZ assembly. One major study clearly reported that min 2 cells continued with equal probability to divide precisely at mid cell or at the poles, the presumed ancient mid cell sites.87 In some contrast, a subsequent detailed pedigree study found that min mutants divided much more randomly in time and position compared to the wild-type. Moreover, in this case even the resulting mini cells varied greatly in size;88 see also Yu & Margolin.63 However, the same study by Akerlund et al.88 also revealed considerable disturbance of the structural organization and distribution of nucleoids in min mutants, a pleiotrophic effect which could complicate the interpretation. More recently, Margolin and co-workers63,67 observed highly accurate mid cell placement of FtsZ rings in many cells in a minCDE deletion. Any disturbance of FtsZ positioning was often associated with irregular nucleoid positioning, suggesting that disturbance of mid cell location in the mutants may be a secondary effect of an altered nucleoid structure leading to irregular segregation. When the latter was exaggerated by combining the min deletion with parC 2 (temperature-sensitive for topoisomerase IV), and an ftsA mutation (affecting invagination at the constriction site, see below), large nucleoid free zones were generated in the resulting filaments, in which multiples of up to six FtsZ rings were detected. Overall these results indicated that at least under these conditions FtsZ rings can form in many places along the cell membrane, suggesting no requirement for a specific nucleation centre at a mid cell site. Thus, the authors concluded that FtsZ assembly is

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Figure 2. Dual function of the Z ring. Cartoon illustrating a cross-section through the division septum, highlighting functions of FtsZ. The cytoplasmic membrane is represented as a thick black line, GTP-bound FtsZ as grey spheres and GDP-bound FtsZ as green spheres. Cell division proteins recruited to the site of constriction by interaction with the Z ring into multiprotein complexes, include FtsI/PBP3 (green), FtsQ/FtsN (pink), FtsL (light blue), FtsK (orange), ZipA (yellow), FtsW (red) and FtsA (dark blue triangle). A newly formed Z ring (above) constricts (below) due to hydrolysis of GTP, leading to loss of subunits and perhaps bending of protofilaments and concomitant deepening of the division constricton.

negatively regulated both by the presence of nucleoids (as in the nucleoid occlusion model) and by the inhibitory effect of MinCD, preventing ring formation at all sites except mid cell. However, as the authors themselves point out, with the use of these complex combinations of mutations, several alternative explanations of the data are available, not least that many lower affinity sites for FtsZ ring formation become available under these unusual conditions, thereby masking the presence of a single specific, mid cell site, normally targeted by FtsZ for assembly. In reality a number of factors may present technical difficulties in detecting specific mid cell sites for FtsZ assembly. In particular, as now pointed out by a number of authors67,89 such a site for much of the cell cycle may be blocked/masked by the presence of the replication apparatus at the identical or an overlapping site. This may only become available at later stages of replication when segregation of “organizing centres” for the nucleoid, involving proteins like SeqA90 in E. coli or Soj in B. subtilis91 has occurred. In our view, the idea that bacteria require a genetically programmed mechanism, dependent upon the temporal action and spatial distribution of dedicated proteins, for precise determination of a mid cell site, uniquely endowed with the property to recruit the division protein FtsZ and perhaps also the replication machinery, is still an attractive one. Whilst the Min proteins may not themselves fulfil this role, the mechanism which generates the gradient of Min proteins from poles to mid cell, involving perhaps an underlying as

yet unknown structure, could clearly represent the elusive measuring stick that is required. What might be the nature of the mid cell site? As discussed above many authors still prefer the idea of a specific site at mid cell which acts as the nucleation centre for FtsZ assembly and ultimately as the site of constriction and biogenesis of new hemispherical poles. Such a site, or something equally specific, may also serve earlier in the cell cycle as the assembly point for the DNA replication machinery, as also discussed above. Most likely such a site will contain specific membraneassociated proteins which then anchor the replication apparatus and subsequently the initial FtsZ molecules for further polymerization. An oscillating device, as discussed in the previous section, may be the basis for concentrating specifically at mid cell such anchor proteins directly, or at the behest of small ions such as Ca2þ or effector nucleotides. Alternatively (or in addition), the membrane bilayer itself may be differentially structured such that specific lipids or formations of lipids may be induced uniquely at mid cell, in response to an oscillator, providing a recognition feature for molecules92 such as FtsZ. Evidence for distinguishable domains in membranes at mid cell and cell poles has been reported for E. coli with fluorescent dyes.93,94 Such domains could be established by an oscillatory mechanism considered above or by the biosynthetic activities and segregation of the nucleoid, as a variation on the nucleoid occlusion model (summarized by

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elements of peptidoglycan synthesis, form several complexes, septalsomes98 or divisomes.99 Assembly into observable ring structures for the division proteins absolutely depends upon FtsZ, although FtsA, ZipA and possibly FtsW, in turn may contribute to the stability of the FtsZ ring.84,100,101 FtsA, also with some structural similarity to actin monomers,102 may contribute to the structural or force-producing properties of the FtsZ ring, whilst ZipA and FtsW may link the FtsZ ring tightly to the membrane. Properties of FtsZ

Figure 3. Polymers of FtsZ generated in vitro. Cartoon illustrating different FtsZ polymers observed in vitro. FtsZ monomers are represented as grey spheres (below) and a three-dimensional representation of the overall structure is indicated above. (a) protofilament, bar represents 5 nm, (b) thick filament, (c) sheet formed in the presence of Ca2þ, DEAE dextran or GMPCPP, (d) tubule formed in the presence of Ca2þ, bar represents 23 nm. Both (c) and (d) are made up of anti-parallel thick filaments. Tubes with larger diameters have been observed using the non-hydrolysable GTP analogue, GMPCPP.121

Norris & Fishov95). For reasons stated above this latter mechanism may be insufficiently precise for the observed accuracy of mid cell localization of FtsZ. Nevertheless, it would be surprising if mid site selection and activity did not in some way involve a specific role for the membrane lipids, and this possibility should be explored. Dual function for the FtsZ ring: a framework for the septalsome and for cytokinesis The dual functions of FtsZ are illustrated in the cartoon in Figure 2. First, as will be considered in more detail in the next section, the structure and the properties of FtsZ clearly provide it with the capacity for the cytoskeletal, perhaps motor role, necessary for “contraction” along the division plane. In addition, however, the FtsZ ring structure provides the framework for the recruitment or assembly of the ten or so membrane and cytoplasmic proteins, uniquely required for cell division in E. coli or B. subtilis, some of which are required for biogenesis of the new hemispherical poles of the two daughter cells. In this way FtsZ is a specific morphological determinant, determining the shape of the hemispherical poles.30,96 For more comprehensive reviews of the role of some of these other division proteins, including the synthesis of new peptidoglycan, the reader is referred to Nanninga,37 Justice & Rothfield97 and Margolin.67 It has been proposed that some of these division proteins, for example FtsI or FtsQ, together with enzymes essential for common

Biochemical and structural characterization of FtsZ has significantly advanced the bacterial cell division field. Like tubulin, FtsZ has GTPase activity and forms a number of different polymers in vitro. Three-dimensional structures of tubulin and FtsZ show remarkable similarity, providing even stronger evidence for an evolutionary relationship between these two proteins. GTPase activity in vitro The GTPase specific activity of FtsZ from seven organisms (six species of eubacteria and one archaebacteria) has been measured in vitro and is up to 100-fold higher than that of tubulin.9,103 – 109 The in vivo significance of this high GTPase has yet to be established however.110 For example, the FtsZ84 mutation in E. coli results in a protein which has a much reduced GTPase activity in vitro105,106 but can support Z ring formation and cell division in vivo (albeit in a temperature-sensitive fashion).32 However, whether the maximal FtsZ GTPase activity detected in vitro correlates with the GTPase activity manifested in vivo remains to be established. Reports describing the GTPase activity of FtsZ as being co-operative105 – 108 were consistent with early suspicions that FtsZ may be able to oligomerize and that this might stimulate the GTPase activity.6,111 Although oligomerization of FtsZ is now well established in the literature (see below), contradictory reports have either confirmed112 or questioned113 whether the GTPase (and the associated polymerization) behaves co-operatively. These contradictions are likely to reflect the nature of the polymers being studied. For example, Romberg and co-workers detected no co-operativity in polymerization of FtsZ, under conditions where only single protofilaments (see below) are formed.113 They and others have detected similar but shorter polymers in the presence of GDP, which are also formed in a non-co-operative manner, best described as isodesmic;35,113 that is, where addition of each monomer is as likely as the last, without any nucleation event or a critical concentration being required. In contrast, Mukherjee & White and co-workers104,114 demonstrated a dependence of both the GTPase activity and polymerization on FtsZ concentration, under

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conditions where slightly more complex polymers are formed. These are 7–20 nm filaments which probably correspond to a mixture of protofilaments, thick filaments and protofilament bundles (see Figure 3). Isodesmic protein polymers have not yet been observed in vivo,113 however, it is possible that Romberg and co-workers have described the initial stages of FtsZ polymerization in vivo, which in turn leads to formation of the more complex FtsZ polymers described by Mukherjee et al.114 FtsZ polymerization in vitro Polymerization of FtsZ in vitro was initially detected and characterized using electron microscopy and centrifugation.115 – 117 More recently, techniques such as light-scattering35,104,113,114 and immunofluorescence microscopy of reactions spiked with GFP – FtsZ fusion protein118 have been used to observe real-time dynamics of FtsZ-polymerization. FtsZ can polymerize into various structures with the major ones illustrated in Figure 3. The simplest of these is a single linear polymer of FtsZ monomers, called a protofilament. Protofilaments can associate laterally to form pairs (sometimes called thick filaments119), bundles (ill-defined linear associations of multiple protofilaments or thick filaments115,116,120), sheets (parallel or anti-parallel two-dimensional associations of thick filaments117 – 119) and tubes (anti-parallel associations of thick filaments in a circular fashion to form a tubular structure) (see Lowe & Amos121 and references therein). In addition, small circles of FtsZ monomers (a short protofilament bent around to join itself, apparently head to tail) have been observed and termed mini-rings.117 Formation (or stability) of tubes and sheets is promoted by the presence of GMPCPP, a non-(or slowly) hydrolysable GTP analogue,121,122 Ca2þ,118,119,121 the polycation DEAE dextran, or cationic phospholipids.117 It has been suggested therefore84 that formation of complex polymers in vitro by the anionic FtsZ protein is promoted by many cationic substances, although it is not known whether any of these polymers are formed in vivo (see below). Simple protofilaments 5– 7 nm in diameter, or thick filaments, appear to be the basis for all of the more complex polymers of FtsZ. Protofilament formation requires the presence of nucleotide, however, their precise form depends on the type of nucleotide. In the presence of GDP, the protofilaments are short and curved (, 10mers) whilst in the presence of GTP, they are longer and straighter.35,113,122 GTP hydrolysis is not required for polymerization but is required for turnover of protofilaments formed in the presence of GTP.114,123 Nature of the Z ring, its assembly and constriction As a result of the in vitro studies, many challenging questions face the bacterial cell division field:

Review: FtsZ Regulation, Structure and Function

for example, which of these in vitro FtsZ polymers are present in the functional Z ring? It seems unlikely that tubes such as those formed in the presence of GMPCPP or Ca2þ have been overlooked, for example, at the leading edge of the division septum in electron-microscopic sections of bacteria. Although tubes of similar diameter to those observed in vitro have been detected in some plastids, it is not known if these are made of FtsZ.124 – 126 Even if tubes can be ruled out, the Z ring could still potentially be formed from several single protofilaments, thick filaments or sheets. A second group of questions relate to formation and regulation of the Z ring and the role of the FtsZ GTPase in these processes. Formation of the Z ring in vivo has been observed to be initiated by a single nucleation event and then to expand bidirectionally,30,31 and it seems reasonable to assume, on the basis of the in vitro studies, that nucleotide-binding is required for this step. The nature of this nucleation event is completely unknown, however, it could represent an important regulatory step in cell division (see earlier). Moreover, as described earlier the Z ring forms early in the E. coli cell cycle, and waits for a time until cell division begins.33,127 This represents a second potential point of cell division regulation. One possibility is that stimulation of the FtsZ GTPase at this stage could cause the Z ring to begin constricting through disassembly and “bending” of the polymers. Erickson128 has proposed that bending results from changing/increasing the angle between monomers by as much as 228. A role for calcium ions in polymerization of the Z ring has been proposed based on evidence that 10 mM Ca2þ stimulates FtsZ polymerization in vitro.118 The concentration of free Ca2þ in the E. coli cytoplasm is known to be orders of magnitude lower than 10 mM,129 however, it has been calculated that transient mobilization of all Ca2þ (bound, for example, to DNA, membranes, etc.) inside E. coli cells could raise the Ca2þ concentration close to 10 mM (see Holland et al.130). Alternatively, temporally and/or spatially regulated import of Ca2þ could transiently raise the Ca2þ concentration in order to stimulate polymerization of the Z ring. However, there is currently no experimental data to support these models, and the role of Ca2þ in Z ring formation in vivo remains unproven. A further critical question is how does the Z ring drive constriction of the septum? Does FtsZ provide the energy for constriction itself, or does the Z ring act passively, as a track for other energy-producing molecules? One hypothesis comes from the observation that GTP-bound polymers are long and straight whilst GDP-bound polymers are short and bent. Polymers within the Z ring could undergo a transition from GTP-bound to GDP-bound as hydrolysis occurs. Shortening and bending of the polymers would exert an inward force on the cytoplasmic membrane causing invagination of the surface

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Figure 4. Important amino acid residues for FtsZ function. Methanococcus janaschii FtsZ three-dimensional structure135 is illustrated on the left in grey. Amino (N) and carboxy (C) termini are indicated. Bound GDP is coloured in pink. Residues highlighted in blue represent the FtsZ/tubulin signature motif.105 – 107 Positions of five FtsZ mutations characterised in E. coli, which result in significant loss of GTPase activity (summarised by Erickson138) are illustrated. The mutated residues have been highlighted at corresponding residues in the M. jannaschii sequence, calculated from primary sequence and secondary structure alignments (M. M. Khattar, personal communication). Those highlighted in yellow are close to the nucleotide binding pocket. Those highlighted in green contribute to the other side of the binding pocket in a region called the synergy loop.138 It should be noted that two other FtsZ mutations, known to cause significant loss of GTPase activity, the Z3 and Z84 mutations, are located in the FtsZ/tubulin signature motif (blue) and the site of the FtsZ26 mutation (insertion of six amino-acids close to the N terminus96) which results in (mis) assembly of FtsZ into spirals instead of rings30 is indicated in red. On the right, outlines of the FtsZ structure are used to show the orientation of monomers in the protofilament, as viewed from the equivalent of the inside of a tubulin microtubule. Highlighted residues provide a simplified illustration of how the regions of adjacent monomers come together to form the nucleotide binding pocket (see Lo¨we & Amos119). ZipA and FtsA bind to the C-terminal, highly conserved 15 – 20 residues of FtsZ, a region which is absent from the M. jannaschii protein and hence not illustrated here. Erickson128 has proposed that ZipA binding effectively allows the protein to staple the Z ring to the membrane.

layers.122 In support of this model Mingorance et al.131 have recently suggested that FtsZ polymers can readily exchange GDP for GTP throughout their length, in vitro, rather than solely at the ends of polymers as is the case for microtubules (although this interpretation of the results has been disputed132). Thus, in theory, large changes in polymer shape could occur quite quickly in response to changes in GTP/GDP ratios, however, it is not clear how this could be controlled or indeed whether fluctuations in GTP/GDP levels actually occur see (see above). Interaction of FtsZ with bacterial motor proteins analagous to microtubule-dependent motors such as kinesins and dyneins (as yet undiscovered in prokaryotes) has been proposed as another potential mechanism for Z ring contraction (e.g. Bramhill and Thomson116) and therefore cellular constriction. Interestingly, microtubules themselves (and other cytoskeletal

proteins such as actin) can indeed exert force in either of two ways;133,134 that is, as a result of polymerization/depolymerization or mediated by motor proteins. These hypotheses pose other questions, for example, how does the Z ring attach to the inner face of the cytoplasmic membrane and how strong is this interaction? If this involves interaction with membrane-spanning proteins such as FtsW and ZipA, which have been implicated in stabilizing FtsZ rings,84,100,101 then what happens in organisms which lack these proteins? ZipA is present only in some gamma-proteobacteria and although FtsW is more widespread, some firmicutes and apparently all euryarchaeota lack both proteins.67 In these species therefore, perhaps the Z ring does not require stabilisation, or the roles of FtsW and ZipA are carried out by other as yet uncharacterised proteins.

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Figure 5. Division site selection in E. coli. Replicating chromosomes (blue) and the Z ring (purple) are indicated inside a growing and dividing bacterium. The action of the oscillatory Min system for mid-cell selection is illustrated using a gradient of red and white on the cytoplasmic membrane. Red indicates a high probability of the MinCD division inhibitor being present and white indicates a low probability. The nucleoid occlusion model for inhibition of division is illustrated by an area of green surrounding nucleoids which represents, until segregation has occurred, an inhibitory influence on division. The nucleation site (NS)68 for Z ring assembly is illustrated in yellow.

The high resolution structure of FtsZ The structure of Methanococcus jannaschii FtsZ was solved by X-ray crystallography in 1998.135 The structure of the alpha/beta tubulin dimer, solved by electron crystallography and published at the same time, shows remarkable similarity to FtsZ,136 indeed, the helices and strands of the N-terminal GTPase domain and the C-terminal domain of FtsZ are virtually superimposable upon the equivalent domains in the tubulin structure. Using information from the FtsZ structure, analysis

Review: FtsZ Regulation, Structure and Function

of FtsZ sheets formed in the presence of Ca2þ and comparison with the alpha/beta tubulin dimer structure, Lo¨we & Amos produced a model of the longitudinal alignment of FtsZ monomers in a protofilament.119 This illustrated a potential mechanism for the co-operativity of the FtsZ GTPase, since the active site is formed in between two subunits. As shown in Figure 4 bound nucleotide sits between individual monomers in the axis of the protofilament, as it does between the alpha and beta subunits of tubulin. The availability of the tertiary structure of the FtsZ molecule has given added meaning to the wealth of genetic information on FtsZ. Thus, with the E. coli FtsZ sequence superimposed upon the M. jannaschii structure, the positions of E. coli FtsZ mutations affecting function have been analysed (e.g. Erickson137). For example, the tubulin signature sequence proposed to be involved in nucleotide-binding105 – 107 is situated in direct contact with the bound nucleotide.135 Also, a cluster of mutations distant from the tubulin signature sequence, correspond to residues which participate in forming the “other side” of the nucleotidebinding pocket. That is, they lie at the base of the next subunit in an FtsZ protofilament119 (see Figure 4). A comprehensive list of FtsZ mutations and their phenotypes has been published elsewhere.138 Interactions of the E. coli FtsZ with two other cell division proteins have been mapped onto the tertiary structure. The essential cell division proteins ZipA and FtsA both interact with a similar 15 –20 residue peptide at the FtsZ C terminus (which is situated at the side of the protofilament, equivalent to the outside of a microtubule).84,139 – 141 Due to the relatively low abundance of ZipA and FtsA (at least in E. coli: see Feucht et al.142) it seems unlikely that they are in competition for FtsZ binding, however, this could explain the lethal effects of over-expression of either of the two proteins.17,18,143 Perspectives The multiple functions of FtsZ, the key molecule in bacterial cytokinesis, have to be expressed at different times during the cell division program. Surprisingly perhaps, at least in E. coli, none of these appear to be regulated by control of the synthesis or stability of FtsZ. Rather, the results of genetic and biochemical analysis, structural studies and cell biology, suggest that interactions with multiple proteins and small effector molecules (for example, nucleotide triphosphates) are the likely modes of regulation. In that case it is not excluded that fluctuations in the local (or general) concentration of small molecules might control the temporal action of FtsZ. Strikingly, even early in the cell cycle, the Z rings form very precisely around mid cell. How this is achieved remains a mystery. The underlying mechanism which determines the fascinating oscillatory behaviour of the MinCDE proteins, although equally mysterious, has revealed a system that does allow a cell to

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accurately find its centre, as illustrated in Figure 5. Nevertheless, the Min proteins themselves are probably not essential for FtsZ localization at mid cell; some baceria even lack Min E. However the timing and polymerization of the Z ring from FtsZ monomers is triggered in vivo, this most likely involves GTP-binding and perhaps GTP-hydrolysis. Indeed, in vitro studies have already provided valuable insights into the structures which might be involved in the assembly and dynamics of Z rings, including GTP hydrolysis-driven bending of the FtsZ polymers. Some proteins uniquely required for cell division (e.g. FtsA), may function to stabilize Z rings whilst others (e.g. FtsI, FtsL), which may promote and co-ordinate ingrowing of the septal cell wall with invagination of the cellular membranes, are specifically recruited to the cell centre by FtsZ, acting as a template for multiprotein assemblies. The Z ring, in response to another unknown signal, then drives septation as it decreases in diameter. The “contraction” of the ring probably involves GTP hydrolysis and ultimately the return of FtsZ monomers to the cytoplasm. Importantly, FtsZ is not merely the assembly point for cell wall enzymes, since the structure of FtsZ itself determines the shape, the architecture, of the ingrowing septum, and thus the shape of the actual poles of the new born cells. We suspect that the correct positioning of the division proteins and the timing of septation events are controlled by a plethora of positive and negative effectors, linked to growth and macromolecular synthesis, as well as intra and extracellular signals (for example, quorum regulators) determining appropriate adaptatory responses. Such effectors are expected to vary between species, as the division process is fine-tuned to ensure its high fidelity in diverse ecological niches. Despite the exhilarating advances in bacterial cell cycle studies in general and the role of FtsZ in particular, important fundamental aspects of bacterial division remain to be characterized. These include the nature of the elusive nucleation site on the cytoplasmic membrane at mid cell which seeds FtsZ polymerization, the precise structure of FtsZ polymers and their lateral contacts and the intriguing mechanism by which FtsZ drives constriction of the septum. These fundamental properties are likely to be conserved in the different contexts in which FtsZ plays a role, for example, in both normal and asymmetric, sporulationrelated division events, and the division of archaebacteria, chloroplasts and mitochondria.

Acknowledgments I.B.H. acknowledges the support of CNRS and Universite´ de Paris-Sud, and also the Human Frontier Science Program (Contract, RG-386/ASM). S.G.A. acknowledges the support of the Wellcome Trust and the University of

Manchester. We are grateful to Vic Norris and Elaine Small for critical reading and discussions of the manuscript. We are also grateful to many colleagues who have contributed to the exciting and highly stimulating developments in this field in the last few years, but can only regret that we had to make selective and therefore difficult choices concerning what to include.

References 1. Schaechter, M., Maaloe, O. & Kjeldgaard, N. O. (1958). Dependency on medium and temperature of cell size and chemical composition during balanced growth of Salmonella typhimurium. J. Gen. Microbiol. 29, 596– 606. 2. Nanninga, N. & Woldringh, C. L. (1985). Cell growth, genome duplication and cell division in Escherichia coli. In Molecular Cytology of E. coli, pp. 260–318, Academic Press, London. 3. Koch, A. L. & Schaechter, M. (1962). A model for the statistics of the cell division process. J. Gen. Microbiol. 29, 435– 454. 4. Hiraga, S., Niki, H., Ogura, T., Ichinose, C., Mori, H., Ezaki, B. & Jaffe, A. (1989). Chromosome partitioning in Escherichia coli: novel mutants producing anucleate cells. J. Bacteriol. 171, 1496– 1505. 5. Helmstetter, C., Cooper, S., Pierucci, O. & Revelas, E. (1968). On the bacterial life sequence. Cold Spring Harbor Symp. Quant. Biol. 33, 809– 822. 6. Bi, E. F. & Lutkenhaus, J. (1991). FtsZ ring structure associated with division in Escherichia coli. Nature, 354, 161– 164. 7. Walker, J. R., Kovarik, A., Allen, J. S. & Gustafson, R. A. (1975). Regulation of bacterial cell division: temperature-sensitive mutants of Escherichia coli that are defective in septum formation. J. Bacteriol. 123, 693– 703. 8. Harry, E. J., Pogliano, K. & Losick, R. (1995). Use of immunofluorescence to visualize cell-specific gene expression during sporulation in Bacillus subtilis. J. Bacteriol. 177, 3386– 3393. 9. Lu, C., Stricker, J. & Erickson, H. P. (1998). FtsZ from Escherichia coli, Azotobacter vinelandii, and Thermotoga maritima: quantitation, GTP hydrolysis, and assembly. Cell Motil. Cytoskel. 40, 71 – 86. 10. Aldea, M., Garrido, T., Pla, J. & Vicente, M. (1990). Division genes in Escherichia coli are expressed coordinately to cell septum requirements by gearbox promoters. EMBO J. 9, 3787– 3794. 11. Dai, K. & Lutkenhaus, J. (1991). ftsZ is an essential cell division gene in Escherichia coli. J. Bacteriol. 173, 3500– 3506. 12. Palacios, P., Vicente, M. & Sanchez, M. (1996). Dependency of Escherichia coli cell-division size, and independency of nucleoid segregation on the mode and level of ftsZ expression. Mol. Microbiol. 20, 1093– 1098. 13. Tetart, F. & Bouche, J. P. (1992). Regulation of the expression of the cell-cycle gene ftsZ by DicF antisense RNA. Division does not require a fixed number of FtsZ molecules. Mol. Microbiol. 6, 615–620. 14. Ward, J. E., Jr. & Lutkenhaus, J. (1985). Overproduction of FtsZ induces minicell formation in E. coli. Cell, 42, 941– 949. 15. Joseleau-Petit, D., Vinella, D. & D’Ari, R. (1999). Metabolic alarms and cell division in Escherichia coli. J. Bacteriol. 181, 9 – 14.

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16. Donachie, W. D. (1993). The cell cycle of Escherichia coli. Annu. Rev. Microbiol. 47, 199– 230. 17. Dewar, S. J., Begg, K. J. & Donachie, W. D. (1992). Inhibition of cell division initiation by an imbalance in the ratio of FtsA to FtsZ. J. Bacteriol. 174, 6314– 6316. 18. Dai, K. & Lutkenhaus, J. (1992). The proper ratio of FtsZ to FtsA is required for cell division to occur in Escherichia coli. J. Bacteriol. 174, 6145– 6151. 19. Boyd, A. & Holland, I. B. (1979). Regulation of the synthesis of surface protein in the cell cycle of E. coli B/r. Cell, 18, 287–296. 20. Garrido, T., Sanchez, M., Palacios, P., Aldea, M. & Vicente, M. (1993). Transcription of ftsZ oscillates during the cell cycle of Escherichia coli. EMBO J. 12, 3957– 3965. 21. Zhou, P. & Helmstetter, C. E. (1994). Relationship between ftsZ gene expression and chromosome replication in Escherichia coli. J. Bacteriol. 176, 6100– 6106. 22. Sitnikov, D. M., Schineller, J. B. & Baldwin, T. O. (1996). Control of cell division in Escherichia coli: regulation of transcription of ftsQA involves both rpoS and SdiA-mediated autoinduction. Proc. Natl Acad. Sci. USA, 93, 336– 341. 23. Ballesteros, M., Kusano, S., Ishihama, A. & Vicente, M. (1998). The ftsQ1p gearbox promoter of Escherichia coli is a major sigma S-dependent promoter in the ddlB – ftsA region. Mol. Microbiol. 30, 419– 430. 24. Plateau, P., Fromant, M., Kepes, F. & Blanquet, S. (1987). Intracellular 50 ,50 -dinucleoside polyphosphate levels remain constant during the Escherichia coli cell cycle. J. Bacteriol. 169, 419– 422. 25. Holtje, J. V. & Nanninga, N. (1984). The intracellular concentration of cyclic adenosine 30 ,50 -monophosphate is constant throughout the cell cycle of Escherichia coli. FEMS Microbiol. Letters, 22, 189– 192. 26. Segal, E. & Le Pecq, J. B. (1986). Relationship between cellular diadenosine 50 ,5000 -P1,P4-tetraphosphate level, cell density, cell growth stimulation and toxic stresses. Exp. Cell Res. 167, 119 – 126. 27. Jenal, U. (2000). Signal transduction mechanisms in Caulobacter crescentus development and cell cycle control. FEMS Microbiol. Rev. 24, 177–191. 28. Kelly, A. J., Sackett, M. J., Din, N., Quardokus, E. & Brun, Y. V. (1998). Cell cycle-dependent transcriptional and proteolytic regulation of FtsZ in Caulobacter. Genes Dev. 12, 880– 893. 29. Quardokus, E. M., Din, N. & Brun, Y. V. (2001). Cell cycle and positional constraints on FtsZ localization and the initiation of cell division in Caulobacter crescentus. Mol. Microbiol. 39, 949– 959. 30. Addinall, S. G. & Lutkenhaus, J. (1996). FtsZ-spirals and -arcs determine the shape of the invaginating septa in some mutants of Escherichia coli. Mol. Microbiol. 22, 231– 237. 31. Sun, Q. & Margolin, W. (1998). FtsZ dynamics during the division cycle of live Escherichia coli cells. J. Bacteriol. 180, 2050–2056. 32. Addinall, S. G., Cao, C. & Lutkenhaus, J. (1997). Temperature shift experiments with an ftsZ84(Ts) strain reveal rapid dynamics of FtsZ localization and indicate that the Z ring is required throughout septation and cannot reoccupy division sites once constriction has initiated. J. Bacteriol. 179, 4277– 4284. 33. Den Blaauwen, T., Buddelmeijer, N., Aarsman, M. E., Hameete, C. M. & Nanninga, N. (1999).

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34.

35.

36. 37. 38.

39. 40.

41.

42.

43.

44.

45.

46.

47.

48.

49.

Timing of FtsZ assembly in Escherichia coli. J. Bacteriol. 181, 5167– 5175. Justice, S. S., Garcia-Lara, J. & Rothfield, L. I. (2000). Cell division inhibitors SulA and MinC/MinD block septum formation at different steps in the assembly of the Escherichia coli division machinery. Mol. Microbiol. 37, 410– 423. Rivas, G., Lopez, A., Mingorance, J., Ferrandiz, M. J., Zorrilla, S., Minton, A. P. et al. (2000). Magnesiuminduced linear self-association of the FtsZ bacterial cell division protein monomer. The primary steps for FtsZ assembly. J. Biol. Chem. 275, 11740–11749. Hoffman, B., Messer, W. & Schwarz, U. (1972). Regulation of polar cap formation in the life cycle of Escherichia coli. J. Supramol. Struct. 1, 29 – 37. Nanninga, N. (1998). Morphogenesis of Escherichia coli. Microbiol. Mol. Biol. Rev. 62, 110 – 129. Wientjes, F. B. & Nanninga, N. (1989). Rate and topography of peptidoglycan synthesis during cell division in Escherichia coli: concept of a leading edge. J. Bacteriol. 171, 3412– 3419. Donachie, W. D., Begg, K. J. & Vicente, M. (1976). Cell length, cell growth and cell division. Nature, 264, 328– 333. Helmstetter, C. E. (1996). Timing of synthetic activities in the cell cycle. In Escherichia coli and Salmonella typhimurium: Cellular and Molecular Biology (Neidhardt, F. C., ed.), 2nd edit., pp. 1627– 1639, American Society for Microbiology, Washington, DC. Jones, N. C. & Donachie, W. D. (1973). Chromosome replication, transcription and control of cell division in Escherichia coli. Nature New Biol. 243, 100– 103. Nordstrom, K., Bernander, R. & Dasgupta, S. (1991). The Escherichia coli cell cycle: one cycle or multiple independent processes that are co-ordinated? Mol. Microbiol. 5, 769– 774. Liu, G. W., Begg, K., Geddes, A. & Donachie, W. D. (2001). Transcription of essential cell division genes is linked to chromosome replication in Escherichia coli. Mol. Microbiol. 40, 909– 916. Seror, S. J., Casaregola, S., Vannier, F., Zouari, N., Dahl, M. & Boye, E. (1994). A mutant cysteinyltRNA synthetase affecting timing of chromosomal replication initiation in B. subtilis and conferring resistance to a protein kinase C inhibitor. EMBO J. 13, 2472– 2480. Levin, P. A., Kurtser, I. G. & Grossman, A. D. (1999). Identification and characterization of a negative regulator of FtsZ ring formation in Bacillus subtilis. Proc. Natl Acad. Sci. USA, 96, 9642–9647. Mizusawa, S. & Gottesman, S. (1983). Protein degradation in Escherichia coli: the lon gene controls the stability of sulA protein. Proc. Natl Acad. Sci. USA, 80, 358– 362. Jones, C. & Holland, I. B. (1985). Role of the SulB (FtsZ) protein in division inhibition during the SOS response in Escherichia coli: FtsZ stabilizes the inhibitor SulA in maxicells. Proc. Natl Acad. Sci. USA, 82, 6045– 6049. Huisman, O. & D’Ari, R. (1983). Effect of suppressors of SOS-mediated filamentation on sfiA operon expression in Escherichia coli. J. Bacteriol. 153, 169– 175. Smith, B. T. & Walker, G. C. (1998). Mutagenesis and more: umuDC and the Escherichia coli SOS response. Genetics, 148, 1599– 1610.

Review: FtsZ Regulation, Structure and Function

50. Burton, P. & Holland, I. B. (1983). Two pathways of division inhibition in UV-irradiated E. coli. Mol. Gen. Genet. 190, 309– 314. 51. Huang, J., Cao, C. & Lutkenhaus, J. (1996). Interaction between FtsZ and inhibitors of cell division. J. Bacteriol. 178, 5080– 5085. 52. Higashitani, A., Higashitani, N. & Horiuchi, K. (1995). A cell division inhibitor SulA of Escherichia coli directly interacts with FtsZ through GTP hydrolysis. Biochem. Biophys. Res. Commun. 209, 198–204. 53. Mukherjee, A., Cao, C. & Lutkenhaus, J. (1998). Inhibition of FtsZ polymerization by SulA, an inhibitor of septation in Escherichia coli. Proc. Natl Acad. Sci. USA, 95, 2885– 2890. 54. Trusca, D., Scott, S., Thompson, C. & Bramhill, D. (1998). Bacterial SOS checkpoint protein SulA inhibits polymerization of purified FtsZ cell division protein. J. Bacteriol. 180, 3946– 3953. 55. de Boer, P. A., Crossley, R. E. & Rothfield, L. I. (1989). A division inhibitor and a topological specificity factor coded for by the minicell locus determine proper placement of the division septum in E. coli. Cell, 56, 641– 649. 56. Hu, Z., Mukherjee, A., Pichoff, S. & Lutkenhaus, J. (1999). The MinC component of the division site selection system in Escherichia coli interacts with FtsZ to prevent polymerization. Proc. Natl Acad. Sci. USA, 96, 14819– 14824. 57. Cordell, S. C., Anderson, R. E. & Lowe, J. (2001). Crystal structure of the bacterial cell division inhibitor MinC. EMBO J. 20, 2454– 2461. 58. Pichoff, S. & Lutkenhaus, J. (2001). Escherichia coli division inhibitor MinCD blocks septation by preventing Z ring formation. J. Bacteriol. 183, 6630– 6635. 59. D’Ari, R. & Huisman, O. (1983). Novel mechanism of cell division inhibition associated with the SOS response in Escherichia coli. J. Bacteriol. 156, 243– 250. 60. Labie, C., Bouche, F. & Bouche, J. P. (1990). Minicellforming mutants of Escherichia coli: suppression of both DicB- and MinD-dependent division inhibition by inactivation of the minC gene product. J. Bacteriol. 172, 5852– 5855. 61. Koppes, L. H., Woldringh, C. L. & Nanninga, N. (1978). Size variations and correlation of different cell cycle events in slow-growing Escherichia coli. J. Bacteriol. 134, 423– 433. 62. Trueba, F. J. (1982). On the precision and accuracy achieved by Escherichia coli cells at fission about their middle. Arch. Microbiol. 131, 55– 59. 63. Yu, X. C. & Margolin, W. (1999). FtsZ ring clusters in min and partition mutants: role of both the Min system and the nucleoid in regulating FtsZ ring localization. Mol. Microbiol. 32, 315– 326. 64. Woldringh, C. L., Mulder, E., Valkenburg, J. A., Wientjes, F. B., Zaritsky, A. & Nanninga, N. (1990). Role of the nucleoid in the toporegulation of division. Res. Microbiol. 141, 39 – 49. 65. Koch, A. L. (1990). Cell shape and division in E. coli. Res. Microbiol. 141, 136– 139. 66. Cook, W. R. & Rothfield, L. I. (1999). Nucleoid-independent identification of cell division sites in Escherichia coli. J. Bacteriol. 181, 1900– 1905. 67. Margolin, W. (2000). Themes and variations in prokaryotic cell division. FEMS Microbiol. Rev. 24, 531–548. 68. Harry, E. J. (2001). Bacterial cell division: regulating Z ring formation. Mol. Microbiol. 40, 795– 803.

233

69. Regamey, A., Harry, E. J. & Wake, R. G. (2000). Midcell Z ring assembly in the absence of entry into the elongation phase of the round of replication in bacteria: co-ordinating chromosome replication with cell division. Mol. Microbiol. 38, 423–434. 70. Lemon, K. P. & Grossman, A. D. (2000). Movement of replicating DNA through a stationary replisome. Mol. Cell, 6, 1321– 1330. 71. Sawitzke, J. & Austin, S. (2001). An analysis of the factory model for chromosome replication and segregation in bacteria. Mol. Microbiol. 40, 786–794. 72. Hale, C. A., Meinhardt, H. & de Boer, P. A. (2001). Dynamic localization cycle of the cell division regulator MinE in Escherichia coli. EMBO J. 20, 1563– 1572. 73. Fu, X., Shih, Y. L., Zhang, Y. & Rothfield, L. I. (2001). The MinE ring required for proper placement of the division site is a mobile structure that changes its cellular location during the Escherichia coli division cycle. Proc. Natl Acad. Sci. USA, 98, 980– 985. 74. Rowland, S. L., Fu, X., Sayed, M. A., Zhang, Y., Cook, W. R. & Rothfield, L. I. (2000). Membrane redistribution of the Escherichia coli MinD protein induced by MinE. J. Bacteriol. 182, 613– 619. 75. de Boer, P. A., Crossley, R. E., Hand, A. R. & Rothfield, L. I. (1991). The MinD protein is a membrane ATPase required for the correct placement of the Escherichia coli division site. EMBO J. 10, 4371– 4380. 76. Zhao, C. R., de Boer, P. A. & Rothfield, L. I. (1995). Proper placement of the Escherichia coli division site requires two functions that are associated with different domains of the MinE protein. Proc. Natl Acad. Sci. USA, 92, 4313– 4317. 77. Hayashi, I., Oyama, T. & Morikawa, K. (2001). Structural and functional studies of MinD ATPase: implications for the molecular recognition of the bacterial cell division apparatus. EMBO J. 20, 1819– 1828. 78. Cordell, S. C. & Lowe, J. (2001). Crystal structure of the bacterial cell division regulator MinD. FEBS Letters, 492, 160– 165. 79. King, G. F., Shih, Y. L., Maciejewski, M. W., Bains, N. P., Pan, B., Rowland, S. L. et al. (2000). Structural basis for the topological specificity function of minE. Nature Struct Biol. 7, 1013– 1017. [In Process Citation.] 80. Jones, L. J., Carballido-Lopez, R. & Errington, J. (2001). Control of cell shape in bacteria: helical, actin-like filaments in Bacillus subtilis. Cell, 104, 913–922. 81. van Den Ent, F., Amos, L. A. & Lowe, J. (2001). Prokaryotic origin of the actin cytoskeleton. Nature, 413, 39 – 44. 82. Hu, Z. & Lutkenhaus, J. (2001). Topological regulation of cell division in E. coli spatiotemporal oscillation of MinD requires stimulation of its ATPase by MinE and phospholipid. Mol. Cell, 7, 1337– 1343. 83. Raskin, D. M. & de Boer, P. A. (1999). MinDEdependent pole-to-pole oscillation of division inhibitor MinC in Escherichia coli. J. Bacteriol. 181, 6419– 6424. 84. Hale, C. A., Rhee, A. C. & de Boer, P. A. (2000). ZipA-induced bundling of FtsZ polymers mediated by an interaction between C-terminal domains. J. Bacteriol. 182, 5153– 5166. 85. Beech, P. L. & Gilson, P. R. (2000). FtsZ and organelle division in Protists. Protist, 151, 11 –16.

234

86. Beech, P. L., Nheu, T., Schultz, T., Herbert, S., Lithgow, T., Gilson, P. R. & McFadden, G. I. (2000). Mitochondrial FtsZ in a chromophyte alga. Science, 287, 1276– 1279. 87. Teather, R. M., Collins, J. F. & Donachie, W. D. (1974). Quantal behavior of a diffusible factor which initiates septum formation at potential division sites in Escherichia coli. J. Bacteriol. 118, 407– 413. 88. Akerlund, T., Bernander, R. & Nordstrom, K. (1992). Cell division in Escherichia coli minB mutants. Mol. Microbiol. 6, 2073– 2083. 89. Harry, E. J. (2001). Coordinating DNA replication with cell division: lessons from outgrowing spores. Biochimie, 83, 75 – 81. 90. Hiraga, S., Ichinose, C., Niki, H. & Yamazoe, M. (1998). Cell cycle-dependent duplication and bidirectional migration of SeqA-associated DNAprotein complexes in E. coli. Mol. Cell, 1, 381– 387. 91. Marston, A. L. & Errington, J. (1999). Selection of the midcell division site in Bacillus subtilis through MinD-dependent polar localization and activation of MinC. Mol. Microbiol. 33, 84 – 96. 92. Norris, V. (1992). Phospholipid domains determine the spatial organization of the Escherichia coli cell cycle: the membrane tectonics model. J. Theoret. Biol. 154, 91 – 107. 93. Fishov, I. & Woldringh, C. L. (1999). Visualization of membrane domains in Escherichia coli. Mol. Microbiol. 32, 1166– 1172. 94. Mileykovskaya, E. & Dowhan, W. (2000). Visualization of phospholipid domains in Escherichia coli by using the cardiolipin-specific fluorescent dye 10-N-nonyl acridine orange. J. Bacteriol. 182, 1172–1175. 95. Norris, V. & Fishov, I. (2001). Hypothesis: membrane domains and hyperstructures control bacterial division. Biochimie, 83, 91 – 97. 96. Bi, E. & Lutkenhaus, J. (1992). Isolation and characterization of ftsZ alleles that affect septal morphology. J. Bacteriol. 174, 5414– 5423. 97. Rothfield, L. I. & Justice, S. S. (1997). Bacterial cell division: the cycle of the ring. Cell, 88, 581– 584. 98. Holland, I. B. (1987). Genetic analysis of the E. coli division clock. Cell, 48, 361– 362. 99. Nanninga, N. (1991). Cell division and peptidoglycan assembly in Escherichia coli. Mol. Microbiol. 5, 791– 795. 100. Hale, C. A. & de Boer, P. A. (1999). Recruitment of ZipA to the septal ring of Escherichia coli is dependent on FtsZ and independent of FtsA. J. Bacteriol. 181, 167– 176. 101. Khattar, M. M., Addinall, S. G., Stedul, K. H., Boyle, D. S., Lutkenhaus, J. & Donachie, W. D. (1997). Two polypeptide products of the Escherichia coli cell division gene ftsW and a possible role for FtsW in FtsZ function. J. Bacteriol. 179, 784– 793. 102. van den Ent, F. & Lowe, J. (2000). Crystal structure of the cell division protein FtsA from Thermotoga maritima. EMBO J. 19, 5300– 5307. 103. Mikulik, K., Zhulanova, E., Kratky, M., Kofronova, O. & Benada, O. (2000). Isolation and characterization of dcw cluster from Streptomyces collinus producing kirromycin. Biochem. Biophys. Res. Commun. 268, 282– 288. 104. White, E. L., Ross, L. J., Reynolds, R. C., Seitz, L. E., Moore, G. D. & Borhani, D. W. (2000). Slow polymerization of Mycobacterium tuberculosis FtsZ. J. Bacteriol. 182, 4028– 4034.

Review: FtsZ Regulation, Structure and Function

105. RayChaudhuri, D. & Park, J. T. (1992). Escherichia coli cell-division gene ftsZ encodes a novel GTPbinding protein. Nature, 359, 251– 254. 106. de Boer, P., Crossley, R. & Rothfield, L. (1992). The essential bacterial cell-division protein FtsZ is a GTPase. Nature, 359, 254– 256. 107. Mukherjee, A., Dai, K. & Lutkenhaus, J. (1993). Escherichia coli cell division protein FtsZ is a guanine nucleotide binding protein. Proc. Natl Acad. Sci. USA, 90, 1053– 1057. 108. Wang, X. & Lutkenhaus, J. (1993). The FtsZ protein of Bacillus subtilis is localized at the division site and has GTPase activity that is dependent upon FtsZ concentration. Mol. Microbiol. 9, 435– 442. 109. Wang, X. & Lutkenhaus, J. (1996). FtsZ ring: the eubacterial division apparatus conserved in archaebacteria. Mol. Microbiol. 21, 313– 319. 110. Lu, C., Stricker, J. & Erickson, H. P. (2001). Sitespecific mutations of FtsZ: effects on GTPase and in vitro assembly. BMC Microbiol. 1, 7. 111. Lutkenhaus, J. (1993). FtsZ ring in bacterial cytokinesis. Mol. Microbiol. 9, 403– 409. 112. Sossong, T. M., Jr., Brigham-Burke, M. R., Hensley, P. & Pearce, K. H., Jr (1999). Self-activation of guanosine triphosphatase activity by oligomerization of the bacterial cell division protein FtsZ. Biochemistry, 38, 14843– 14850. 113. Romberg, L., Simon, M. & Erickson, H. P. (2001). Polymerization of FtsZ, a bacterial homolog of tubulin: is assembly cooperative? J. Biol. Chem. 276, 11743– 11753. 114. Mukherjee, A. & Lutkenhaus, J. (1999). Analysis of FtsZ assembly by light scattering and determination of the role of divalent metal cations. J. Bacteriol. 181, 823– 832. 115. Mukherjee, A. & Lutkenhaus, J. (1994). Guanine nucleotide-dependent assembly of FtsZ into filaments. J. Bacteriol. 176, 2754 –2758. 116. Bramhill, D. & Thompson, C. M. (1994). GTPdependent polymerization of Escherichia coli FtsZ protein to form tubules. Proc. Natl Acad. Sci. USA, 91, 5813– 5817. 117. Erickson, H. P., Taylor, D. W., Taylor, K. A. & Bramhill, D. (1996). Bacterial cell division protein FtsZ assembles into protofilament sheets and minirings, structural homologs of tubulin polymers. Proc. Natl Acad. Sci. USA, 93, 519– 523. 118. Yu, X. C. & Margolin, W. (1997). Ca2þ -mediated GTP-dependent dynamic assembly of bacterial cell division protein FtsZ into asters and polymer networks in vitro. EMBO J. 16, 5455– 5463. 119. Lowe, J. & Amos, L. A. (1999). Tubulin-like protofilaments in Ca2þ -induced FtsZ sheets. EMBO J. 18, 2364– 2371. 120. Erickson, H. P. (1995). FtsZ, a prokaryotic homolog of tubulin? Cell, 80, 367– 370. 121. Lowe, J. & Amos, L. A. (2000). Helical tubes of FtsZ from Methanococcus jannaschii. Biol. Chem. 381, 993– 999. 122. Lu, C., Reedy, M. & Erickson, H. P. (2000). Straight and curved conformations of FtsZ are regulated by GTP hydrolysis. J. Bacteriol. 182, 164– 170. 123. Mukherjee, A. & Lutkenhaus, J. (1998). Dynamic assembly of FtsZ regulated by GTP hydrolysis. EMBO J. 17, 462– 469. 124. Sprey, B. (1968). Zur Feinstruktur des Plastidenstromas von Hordeum vulgare L. Protoplasma, 66, 469– 479.

Review: FtsZ Regulation, Structure and Function

125. Lawrence, M. E. & Possingham, J. V. (1984). Observations of microtubule-like structures within spinach plastids. Biol. Cell, 52, 77 – 82. 126. Miyagishima, S., Takahara, M. & Kuroiwa, T. (2001). Novel filaments 5 nm in diameter constitute the cytosolic ring of the plastid division apparatus. Plant Cell, 13, 707– 721. 127. Addinall, S. G., Bi, E. & Lutkenhaus, J. (1996). FtsZ ring formation in fts mutants. J. Bacteriol. 178, 3877– 3884. 128. Erickson, H. P. (2001). The FtsZ protofilament and attachment of ZipA: structural constraints on the FtsZ power stroke. Curr. Opin. Cell Biol. 13, 55– 60. 129. Jones, H. E., Holland, I. B., Baker, H. L. & Campbell, A. K. (1999). Slow changes in cytosolic free Ca2þ in Escherichia coli highlight two putative influx mechanisms in response to changes in extracellular calcium. Cell Calcium, 25, 265– 274. 130. Holland, I. B., Jones, H. E., Campbell, A. K. & Jacq, A. (1999). An assessment of the role of intracellular free Ca2þ in E. coli. Biochimie, 81, 901–907. 131. Mingorance, J., Rueda, S., Gomez-Puertas, P., Valencia, A. & Vicente, M. (2001). Escherichia coli FtsZ polymers contain mostly GTP and have a high nucleotide turnover. Mol. Microbiol. 41, 83 – 91. 132. Scheffers, D. & Driessen, A. J. (2001). The polymerization mechanism of the bacterial cell division protein FtsZ. FEBS Letters, 506, 6 –10. 133. Inoue, S. & Salmon, E. D. (1995). Force generation by microtubule assembly/disassembly in mitosis and related movements. Mol. Biol. Cell, 6, 1619– 1640. 134. Theriot, J. A. (2000). The polymerization motor. Traffic, 1, 19 – 28. 135. Lowe, J. & Amos, L. A. (1998). Crystal structure of the bacterial cell-division protein FtsZ. Nature, 391, 203–206. 136. Nogales, E., Wolf, S. G. & Downing, K. H. (1998). Structure of the alpha beta tubulin dimer by electron crystallography. Nature, 391, 199– 203. 137. Erickson, H. P. (1998). Atomic structures of tubulin and FtsZ. Trends Cell Biol. 8, 133– 137. 138. Erickson, H. & Stricker, J. (1998). In Guidebook to the Cytoskeletal and Motor Proteins (Kreis, T. & Vale, R., eds), pp. 352– 356, Oxford University Press, Oxford. 139. Haney, S. A., Glasfeld, E., Hale, C., Keeney, D., He, Z. Z. & de Boer, P. (2001). Genetic analysis of the Escherichia coli FtsZ·ZipA interaction in the yeast two-hybrid system: characterization of FtsZ residues essential for the interactions with ZipA and with FtsA. J. Biol. Chem. 276, 11980– 11987. 140. Ma, X. & Margolin, W. (1999). Genetic and functional analyses of the conserved C-terminal core domain of Escherichia coli FtsZ. J. Bacteriol. 181, 7531– 7544. 141. Yan, K., Pearce, K. H. & Payne, D. J. (2000). A conserved residue at the extreme C terminus of FtsZ is critical for the FtsA– FtsZ interaction in Staphylococcus aureus. Biochem. Biophys. Res. Commun. 270, 387– 392. 142. Feucht, A., Lucet, I., Yudkin, M. D. & Errington, J. (2001). Cytological and biochemical characterization of the FtsA cell division protein of Bacillus subtilis. Mol. Microbiol. 40, 115 –125. 143. Hale, C. A. & de Boer, P. A. (1997). Direct binding of FtsZ to ZipA, an essential component of the septal ring structure that mediates cell division in E. coli. Cell, 88, 175– 185.

235

144. Wang, X. D., de Boer, P. A. & Rothfield, L. I. (1991). A factor that positively regulates cell division by activating transcription of the major cluster of essential cell division genes of Escherichia coli. EMBO J. 10, 3363– 3372. 145. Pogliano, J., Dong, J. M., De Wulf, P., Furlong, D., Boyd, D., Losick, R. et al. (1998). Aberrant cell division and random FtsZ ring positioning in Escherichia coli cpxA* mutants. J. Bacteriol. 180, 3486– 3490. 146. MacVanin, M., Johanson, U., Ehrenberg, M. & Hughes, D. (2000). Fusidic acid-resistant EF-G perturbs the accumulation of ppGpp. Mol. Microbiol. 37, 98 –107. 147. Carballes, F., Bertrand, C., Bouche, J. P. & Cam, K. (1999). Regulation of Escherichia coli cell division genes ftsA and ftsZ by the two-component system rcsC – rcsB. Mol. Microbiol. 34, 442– 450. 148. RayChaudhuri, D. (1999). ZipA is a MAP-Tau homolog and is essential for structural integrity of the cytokinetic FtsZ ring during bacterial cell division. EMBO J. 18, 2372 –2383. 149. Nishimura, A., Moriya, S., Ukai, H., Nagai, K., Wachi, M. & Yamada, Y. (1997). Diadenosine 50 ,5000 -P1,P4-tetraphosphate (Ap4A) controls the timing of cell division in Escherichia coli. Genes Cells, 2, 401– 413. 150. Fukuchi, K., Kasahara, Y., Asai, K., Kobayashi, K., Moriya, S. & Ogasawara, N. (2000). The essential two-component regulatory system encoded by yycF and yycG modulates expression of the ftsAZ operon in Bacillus subtilis. Microbiology, 146, 1573– 1583. 151. Newman, E. B., Budman, L. I., Chan, E. C., Greene, R. C., Lin, R. T., Woldringh, C. L. & D’Ari, R. (1998). Lack of S-adenosylmethionine results in a cell division defect in Escherichia coli. J. Bacteriol. 180, 3614– 3619. 152. Edwards, D. H. & Errington, J. (1997). The Bacillus subtilis DivIVA protein targets to the division septum and controls the site specificity of cell division. Mol. Microbiol. 24, 905– 915. 153. Chen, M. X., Bouquin, N., Norris, V., Casaregola, S., Seror, S. J. & Holland, I. B. (1991). A single base change in the acceptor stem of tRNA(3Leu) confers resistance upon Escherichia coli to the calmodulin inhibitor, 48/80. EMBO J. 10, 3113 – 3122. 154. Bouquin, N., Chen, M. X., Kim, S., Vannier, F., Bernard, S., Holland, I. B. & Seror, S. J. (1996). Characterization of an Escherichia coli mutant, feeA, displaying resistance to the calmodulin inhibitor 48/80 and reduced expression of the rare tRNA3Leu. Mol. Microbiol. 20, 853– 865. 155. Nakayashiki, T. & Inokuchi, H. (1998). Novel temperature-sensitive mutants of Escherichia coli that are unable to grow in the absence of wild-type tRNA6Leu. J. Bacteriol. 180, 2931– 2935. 156. Onoda, T., Enokizono, J., Kaya, H., Oshima, A., Freestone, P. & Norris, V. (2000). Effects of calcium and calcium chelators on growth and morphology of Escherichia coli L-form NC-7. J. Bacteriol. 182, 1419– 1422. 157. Tamura, F., Nishimura, S. & Ohki, M. (1984). The E. coli divE mutation, which differentially inhibits synthesis of certain proteins, is in tRNASer1. EMBO J. 3, 1103– 1107. 158. Leclerc, G., Sirard, C. & Drapeau, G. R. (1989). The Escherichia coli cell division mutation ftsM1 is in serU. J. Bacteriol. 171, 2090– 2095.

236

Review: FtsZ Regulation, Structure and Function

159. Mileykovskaya, E., Sun, Q., Margolin, W. & Dowhan, W. (1998). Localization and function of early cell division proteins in filamentous Escherichia coli cells lacking phosphatidylethanolamine. J. Bacteriol. 180, 4252– 4257. 160. Garcia, G. M., Mar, P. K., Mullin, D. A., Walker, J. R. & Prather, N. E. (1986). The E. coli dnaY gene encodes an arginine transfer RNA. Cell, 45, 453– 459. 161. Nishimura, A. (1998). The timing of cell division: Ap4A as a signal. Trends Biochem. Sci, 23, 157– 159. 162. Pruss, B. M., Markovic, D. & Matsumura, P. (1997). The Escherichia coli flagellar transcriptional activator flhD regulates cell division through induction of the acid response gene cadA. J. Bacteriol. 179, 3818– 3821.

163. Pruss, B. M. (1998). Acetyl phosphate and the phosphorylation of OmpR are involved in the regulation of the cell division rate in Escherichia coli. Arch. Microbiol. 170, 141– 146. 164. Katayama, T., Takata, M. & Sekimizu, K. (1997). CedA is a novel Escherichia coli protein that activates the cell division inhibited by chromosomal DNA over-replication. Mol. Microbiol. 26, 687– 697. 165. Olafsson, O., Ericson, J. U., VanBogelen, R. & Bjork, G. R. (1996). Mutation in the structural gene for release factor 1 (RF-1) of Salmonella typhimurium inhibits cell division. J. Bacteriol. 178, 3829 –3839. 166. Bouquin, N., Tempete, M., Holland, I. B. & Seror, S. J. (1995). Resistance to trifluoroperazine, a calmodulin inhibitor, maps to the fabD locus in Escherichia coli. Mol. Gen. Genet. 246, 628– 637.

Edited by P. E. Wright (Received 8 November 2001; received in revised form 22 January 2002; accepted 28 January 2002)