Prokaryotic Cytokinesis: Little Rings Bring Big Cylindrical Things

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Prokaryotic Cytokinesis: Little Rings Bring Big Cylindrical Things At the division site, most bacteria assemble filaments of the tubulin homolog FtsZ that recruit other proteins into a functional divisome. A recent study describes the in vitro assembly of the divisome component SepF into small rings that organize FtsZ filaments into microtubule-like structures, possibly facilitating efficient septal growth and cytokinesis. Daniel P. Haeusser and William Margolin* The tubulin homolog FtsZ is required for cell division in numerous bacteria and is conserved in most bacteria, euryarchaeota, chloroplasts, and some protist mitochondria [1]. Like tubulin, FtsZ undergoes GTP-dependent assembly into single-stranded protofilaments in vitro, which disassemble upon GTP hydrolysis. Under conditions of relatively low pH and ionic strength, or in the presence of certain cofactors, these FtsZ protofilaments can interact laterally to form bundles and other cross-linked forms [2]. Although FtsZ protofilaments do not inherently assemble into microtubule-like structures, lateral interactions are likely important for assembly of FtsZ polymers into the cellular FtsZ ring (Z ring) [2]. The Z ring recruits a number of membrane-associated proteins, collectively called the divisome, and constricts to regulate septum formation and complete cytokinesis  du et al. [1,2]. New work by Gu¨ndog [3] now suggests that the Bacillus subtilis SepF protein assembles into polymeric rings, which orient FtsZ protofilaments into organized tubules that help the divisome to function properly. Advances in imaging technology have provided continued insights into the structure of the Z ring. Fluorescent protein fusions indicate that FtsZ polymers assemble into a helical structure at the division site and rapidly coalesce into a toroid [4,5]. Cryo-electron tomography of Caulobacter crescentus showed that FtsZ forms numerous, small, and non-overlapping filaments at mid-cell, rather than a continuous ring structure [6]. Most recently, high-resolution imaging of fluorescently labeled FtsZ in Escherichia coli by photo-activated light microscopy (PALM) suggested that the Z ring is composed of a loose

association of bundled protofilaments with a width of w110 nm [7]. Proteins such as ZapA, ZapC, ZipA, FzlA, and SepF likely mediate lateral associations between the FtsZ protofilaments of these in vivo bundles, as these factors promote cross-linking and/or bundling of FtsZ protofilaments in vitro [1,8–11]. Analogous structures may increase the integrity of the Z ring in vivo, making its assembly and eventual constriction more robust. Whereas ZapA is conserved across a wide variety of bacterial species, ZipA, ZapC, FzlA and SepF are more narrowly distributed [1,9–11]. SepF is absent from most Gram-negative species, but is conserved in cyanobacteria and Gram-positive bacteria [1]. Initial studies with Streptococcus pneumoniae and Synechococcus elongatus reported that sepF mutants display cell division defects [12,13]. SepF was subsequently shown to localize to the division site in B. subtilis [14,15] and Synechocystis [16], and to interact directly with FtsZ [14–16]. B. subtilis cells lacking SepF are significantly longer than wild type, and Synechocystis cells depleted of SepF are giant, suggesting an involvement of SepF in septal cross-wall synthesis [14–16]. Consistent with this, B. subtilis sepF null cells show no defects in Z-ring formation or recruitment of divisome components [14,15], but their septa appear deformed and wider than normal [14]. However, when cells lacking SepF were depleted of EzrA or FtsA, which co-purify with SepF and are required for efficient Z-ring assembly [15], they exhibit synthetic defects, including aberrantly dispersed and spiral FtsZ assembly [14,15]. These FtsZ assembly phenotypes mimic those of SepF-depleted Synechocystis cells [16]. Importantly, overproduction of SepF in B. subtilis suppresses the defects of ftsA null cells, suggesting that FtsA and SepF

have partially redundant roles [15]. Unfortunately, the lack of a known specific function of FtsA in B. subtilis makes it difficult to assign any specific role to SepF. The first biochemical clue to SepF function came from experiments with purified B. subtilis SepF and FtsZ, in which increasing amounts of SepF enhanced FtsZ assembly into protofilament bundles. Fluorescently labeled SepF colocalized alongside fluorescent polymers of FtsZ, suggesting that SepF stabilizes FtsZ polymer bundles by binding along their length [17]. Another group reported similar findings using purified Synechocystis components  du [16]. The present work by Gu¨ndog et al. [3] extends these results by visualizing SepF–FtsZ interactions at higher resolution. Using electron microscopy, they found that, in a physiological buffer at pH 7.4, purified SepF spontaneously assembled into ring structures with diameters ranging from w42 to 60 nm. The previous studies may have failed to observe SepF rings as a result of the use of lower pH buffer [17] or GFP-tagged SepF [16] because, in the new study, SepF rings failed to form under these conditions. An observation more startling than the SepF rings themselves was that these SepF structures permitted FtsZ protofilament assembly into apparently hollow tubules ranging from w40 to 55 nm in thickness. These diameters are in the same range as those of SepF rings, consistent with FtsZ protofilaments aligned in parallel around stacks of SepF rings (Figure 1A). Assembly of these FtsZ tubules was dependent on FtsZ, Mg2+, and GTP concentration, but independent of the order of component addition [3]. The precise structural arrangement of proteins in these tubules was not fully resolved, but the SepF rings often appeared to be tilted within the FtsZ tubules, perhaps representing a helix, which could explain why the tubule diameters were somewhat smaller than the diameters of isolated SepF rings. Tracking SepF–FtsZ tubule assembly led to several interesting observations. Over time, tubule length increased to several micrometers, with FtsZ protofilaments protruding straight out from either end of the rigid tubules. As time progressed and GTP levels

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Figure 1. FtsZ filament bundling and SepF rings. (A) The early SepF–FtsZ tubule. A ring of FtsZ protofilaments (red cylinders) assembles around tilted SepF ring platforms (blue rings). In the presence of GTP, FtsZ filaments grow straight from each end of the tubule. An enlargement of one SepF ring depicts SepF molecules (blue tailed triangles), alternately shaded to distinguish individual subunits. The curved tail on each SepF depicts its carboxyl terminus, which may help to force SepF polymerization into a closed toroid. The FtsZ subunits (red circles) of each protofilament bind 1:1 to SepF in the ring. (B) The interaction between FtsZ protofilaments in the SepF–FtsZ tubule is distinct from the bundling of FtsZ protofilaments (chained red circles) by factors such as ZapA (tetramer of blue triangles composed of anti-parallel dimers linked by coiled coils) that promote lateral and longitudinal FtsZ crosslinking [8]. ZapA-mediated bundling produces large increases in 90 angle light scattering and significant decreases in FtsZ’s GTP hydrolysis rate, whereas the arrangement of FtsZ protofilaments by SepF permits normal GTPase activity.

diminished, the emerging protofilaments began to fray and curl, while the tubules began to bend, bifurcate, and form connections resembling traffic roundabouts [3]. This transition from straight to bent tubules correlates with a similar transition that has long been observed for single FtsZ protofilaments following GTP hydrolysis [2]. Notably, while reminiscent of microtubules, FtsZ protofilaments grew equally from either tubule end and did not demonstrate dynamic instability, a hallmark of microtubules [18]. However, the fraying of FtsZ protofilaments at the ends, where SepF rings are no longer able to constrain the protofilaments into a tubule, is analogous to similar spiraling of protofilaments at microtubule ends under certain in vitro conditions [18]. Are these SepF–FtsZ structures present in vivo and relevant to cytokinesis? To address this,  du et al. [3] found that Gu¨ndog a carboxy-terminal truncation (D134) or a G135N point mutant of SepF retained interaction with FtsZ but failed to form SepF rings or induce FtsZ tubule formation. Interestingly, the SepF(G135N) mutant formed polymers, but they were straight, instead of curved into rings. This suggests that replacing the glycine with a larger

residue bends the SepF-dimer interface back to a straight conformation. Cells expressing only mutant SepF(G135N) became dependent on FtsA for survival, suggesting that the ability to form SepF rings is required for SepF function in vivo [3]. However, it remains to be seen whether SepF–FtsZ tubules are assembled in cells, and how the mutant SepF(G135N) affects Z-ring formation or septal shape. An attractive model postulated by the authors is that SepF rings could enhance Z-ring coherence by aligning and restraining FtsZ protofilaments into tubules. In Gram-positive species, the Z ring guides formation of the large division septum [1]. An unnecessarily wide ring might be disadvantageous, as it would result in an inappropriately thick and crooked septum and subsequent problems in cell separation, the primary phenotype observed for sepF null B. subtilis [14]. However, if stable 50-nm wide SepF–FtsZ tubules form in vivo, it is surprising that they have not been seen in electron micrographs of thin sections. Because SepF rings assemble readily in vitro without cofactors, it is likely that SepF–FtsZ tubule generation would be negatively regulated in vivo by other proteins, such as the FtsZ assembly inhibitors

MinC or EzrA [1]. High-resolution microscopy techniques such as PALM or cryo-electron tomography will be crucial for visualizing SepF rings and/or SepF–FtsZ tubules in vivo during cell division, and for comparison with other organization states of FtsZ such as ZapA-formed bundles (Figure 1B) and SlmA- or FzlA-formed coils [8,10,19]. Understanding SepF function may also help elucidate the function of other cell division proteins. For example, it is not clear how the loss of FtsA can be largely suppressed by excess SepF [15]. If FtsZ–SepF tubules mimic FtsA function, then overproduction of the SepF(G135N) mutant protein should probably not be able to suppress ftsA null defects. As transitions from straight to curved FtsZ filaments are considered important for Z-ring constriction forces [2,6], perhaps excess SepF promotes FtsZ protofilament curving, which is a postulated function of FtsA and FzlA [10,20]. Another possibility is that excess SepF can compete for FtsZ binding with negative-acting factors such as EzrA [1]. Given the interplay between FtsZ, SepF, FtsA, and EzrA [15], additional filament arrangements or tubule structures may exist in the context of a complete divisome complex in vivo.

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Important questions about SepF remain. For example, does SepF play a role in septation in divergent species such as cyanobacteria? Can purified SepF from other species also spontaneously self-assemble into rings and orient FtsZ protofilaments into tubules? SepF may be essential for cell division of cyanobacteria because they lack FtsA and/or EzrA homologs [13,16]. Other Gram-negative bacteria, which lack SepF, must also maintain Z-ring integrity to coordinate constriction, septum formation, and outer membrane invagination. For the g-proteobacteria, evidence suggests that ZipA and the well-conserved FtsA mediate this coordination [1] and it is likely that other bacteria have as yet unidentified, functionally related factors. Although the basic theme of cell division is becoming clear, unraveling the plethora of variations in the most diverse group of organisms on Earth remains a challenge.

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References 1. Adams, D.W., and Errington, J. (2009). Bacterial cell division: assembly, maintenance and disassembly of the Z ring. Nat. Rev. Microbiol. 7, 642–653. 2. Erickson, H.P., Anderson, D.E., and Osawa, M. (2010). FtsZ in bacterial cytokinesis: cytoskeleton and force generator all in one. Microbiol. Mol. Biol. Rev. 74, 504–528.  du, M.E., Kawai, Y., Pavlendova, N., 3. Gu¨ndog Ogasawara, N., Errington, J., Scheffers, D.J., and Hamoen, L.W. (2011). Large ring polymers

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align FtsZ polymers for normal septum formation. EMBO J. 30, 617–626. Peters, P.C., Migocki, M.D., Thoni, C., and Harry, E.J. (2007). A new assembly pathway for the cytokinetic Z ring from a dynamic helical structure in vegetatively growing cells of Bacillus subtilis. Mol. Microbiol. 64, 487–499. Thanedar, S., and Margolin, W. (2004). FtsZ exhibits rapid movement and oscillation waves in helix-like patterns in Escherichia coli. Curr. Biol. 14, 1167–1173. Li, Z., Trimble, M.J., Brun, Y.V., and Jensen, G.J. (2007). The structure of FtsZ filaments in vivo suggests a force-generating role in cell division. EMBO J. 26, 4694–4708. Fu, G., Huang, T., Buss, J., Coltharp, C., Hensel, Z., and Xiao, J. (2010). In vivo structure of the E. coli FtsZ-ring revealed by photoactivated localization microscopy (PALM). PLoS One 5, e12682. Dajkovic, A., Pichoff, S., Lutkenhaus, J., and Wirtz, D. (2010). Cross-linking FtsZ polymers into coherent Z rings. Mol. Microbiol. 78, 651–668. Durand-Heredia, J.M., Yu, H.H., De Carlo, S., Lesser, C.F., and Janakiraman, A. (2011). Identification and characterization of ZapC, a stabilizer of the FtsZ-ring in Escherichia coli. J. Bacteriol. 193, 1405–1413. Goley, E.D., Dye, N.A., Werner, J.N., Gitai, Z., and Shapiro, L. (2011). Imaging-based identification of a critical regulator of FtsZ protofilament curvature in Caulobacter. Mol. Cell 39, 975–987. Hale, C.A., Shiomi, D., Liu, B., Bernhardt, T.G., Margolin, W., Niki, H., and de Boer, P.A. (2011). Identification of Escherichia coli ZapC (YcbW) as a component of the division apparatus that binds and bundles FtsZ polymers. J. Bacteriol. 193, 1393–1404. Fadda, D., Pischedda, C., Caldara, F., Whalen, M.B., Anderluzzi, D., Domenici, E., and Massidda, O. (2003). Characterization of divIVA and other genes located in the chromosomal region downstream of the dcw cluster in Streptococcus pneumoniae. J. Bacteriol. 185, 6209–6214. Miyagishima, S.Y., Wolk, C.P., and Osteryoung, K.W. (2005). Identification of cyanobacterial cell division genes by

Chromatin: Bind at Your Own RSC Recent work has identified a novel RSC–nucleosome complex that both strongly phases flanking nucleosomes and presents regulatory sites for ready access. These results challenge several widely held views. Nicolas E. Buchler1,2,3,* and Lu Bai4,5 Genome-wide experiments in yeast, fly and mammalian cells have identified the existence of nucleosome-depleted regions in promoters and enhancers [1–4]. Transcription factors are thought to bind to their cognate sites located in these nucleosome-depleted regions, subsequently recruit nucleosomeremodeling and modifying complexes, and evict or reposition flanking nucleosomes that block RNA polymerase assembly at the promoter. By using a novel, quantitative assay, recent work from the Ptashne lab has uncovered several striking insights into

nucleosome occupancy at the GAL1/10 promoter of budding yeast [5–7]. These results challenge current ideas of whether nucleosome-depleted regions are completely nucleosome-free, whether strongly positioned nucleosomes are always incompatible with the binding of regulatory proteins, and whether the occupancy of a DNA fragment by a nucleosome is mostly determined by its sequence. Nucleosome occupancy at a particular genomic location is measured by assessing nucleosome-mediated ‘protection’ (often assumed to be the canonical, mono-nucleosome size of 147 bp) of

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comparative and mutational analyses. Mol. Microbiol. 56, 126–143. Hamoen, L.W., Meile, J.C., de Jong, W., Noirot, P., and Errington, J. (2006). SepF, a novel FtsZ-interacting protein required for a late step in cell division. Mol. Microbiol. 59, 989–999. Ishikawa, S., Kawai, Y., Hiramatsu, K., Kuwano, M., and Ogasawara, N. (2006). A new FtsZ-interacting protein, YlmF, complements the activity of FtsA during progression of cell division in Bacillus subtilis. Mol. Microbiol. 60, 1364–1380. Marbouty, M., Saguez, C., Cassier-Chauvat, C., and Chauvat, F. (2009). Characterization of the FtsZ-interacting septal proteins SepF and Ftn6 in the spherical-celled cyanobacterium Synechocystis strain PCC 6803. J. Bacteriol. 191, 6178–6185. Singh, J.K., Makde, R.D., Kumar, V., and Panda, D. (2008). SepF increases the assembly and bundling of FtsZ polymers and stabilizes FtsZ protofilaments by binding along its length. J. Biol. Chem. 283, 31116–31124. Desai, A., and Mitchison, T.J. (1997). Microtubule polymerization dynamics. Annu. Rev. Cell Dev. Biol. 13, 83–117. Tonthat, N.K., Arold, S.T., Pickering, B.F., Van Dyke, M.W., Liang, S., Lu, Y., Beuria, T.K., Margolin, W., and Schumacher, M.A. (2010). Molecular mechanism by which the nucleoid occlusion factor, SlmA, keeps cytokinesis in check. EMBO J. 30, 154–164. Beuria, T.K., Mullapudi, S., Mileykovskaya, E., Sadasivam, M., Dowhan, W., and Margolin, W. (2009). Adenine nucleotide-dependent regulation of assembly of bacterial tubulin-like FtsZ by a hypermorph of bacterial actin-like FtsA. J. Biol. Chem. 284, 14079–14086.

Department of Microbiology and Molecular Genetics, University of Texas Medical School at Houston, 6431 Fannin St, Houston, TX 77030, USA. *E-mail: [email protected]

DOI: 10.1016/j.cub.2011.02.006

that sequence from digestion by micrococcal nuclease (MNase). Typical nucleosome occupancy assays fix chromatin in cells, lightly digest chromatin at a single concentration of MNase, and quantify protected DNA fragments by quantitative PCR (qPCR), tiling microarrays, or next-generation sequencing. Unfortunately, DNA sequence itself influences digestion efficiency of MNase, a bias that can create a false apparent protection of ‘naked’ genomic DNA. Strikingly, recent papers show that MNase digestion of naked genomic DNA infers similar nucleosome occupancies to that obtained by MNase digestion of chromatin DNA [8,9]. Bryant et al. [5] developed a quantitative MNase protection assay that normalizes against such variability. The assay digests naked genomic DNA and fixed chromatin DNA over a wide range of MNase concentrations,