the MinCD complex to the cell poles, with MinE (and phospholipid) stimulating the ATPase activity of MinD (Hu and Lutkenhaus 2001). MinE binds to the membrane at the pole, targeting MinCD complexes, displacing both MinC and MinD and stimulating ATP hydrolysis by MinD (Loose et al. 2011; Park et al. 2011). MinE and MinD set up a high‐speed oscillating system in which MinC is trafficked from pole to pole, on average spending a minimum of time at mid‐cell and most of the time at the poles (Raskin and de Boer 1997; Hu and Lutkenhaus 1999; Hu et al. 2002). It is the relative paucity of MinC at mid‐cell that diminishes the inhibitory influence on FtsZ polymerisation and Z‐ring formation (Hu and Lutkenhaus 1999; Raskin and de Boer 1999a,b). In addition to inhibiting FtsZ polymerisation by protein‐protein interaction, the oscillation of MinC populations from pole to pole has an impact on the distribution of other FtsZ‐interactors. Together with FtsZ itself, the ZapA, ZapB, and ZipA proteins oscillate oppositely to MinC and with a similar dynamic pattern. ZapB does not bind FtsZ directly but through ZapA, which does bind FtsZ. ZipA, with FtsA, connects FtsZ to the cytoplasmic membrane (Pichoff and Lutkenhaus 2005) while ZapA‐ZapB stimulates Z‐ring formation and stabilises it (Buss et al. 2013; Galli and Gerdes 2010; Gueiros‐Filho and Losick 2002). Therefore, the oscillatory movements of MinC proteins probably trigger periodic assembly and disassembly of the Z ring complexes (Bisicchia et al. 2013; Thanedar and Margolin 2004).
B. subtilis possesses the cell‐pole‐targeting protein DivIVA, which is involved both in chromosome attachment at the pole in sporulating cells (Section 1.10) and in directing the cellular localisation of MinC (Cha and Stewart 1997; Edwards and Errington 1997). The utility of DivIVA as a general pole‐targeting protein arises from its ability to sense cell membrane curvature, which is maximal at the poles (Edwards et al. 2000; Lenarcic et al. 2009). The MinC protein is bound by MinD and an adaptor protein, MinJ, connects this complex to DivIVA (Bramkamp et al. 2008; Patrick and Kearns 2008). As the division septum develops, invagination of the membrane, and the associated membrane curvature, recruit DivIVA from the pole to the mid‐cell (which is the soon‐to‐be pole of the new daughter cell) (Gregory et al. 2008; Rodriguez and Harry 2012; van Baarle and Bramkamp 2010).
Genetic elimination of the Min system and of nucleoid occlusion is deleterious for cell growth in rich medium, but the mutants can grow and divide in minimal medium (Bernhardt and de Boer 2005; Yu and Margolin 1999). This suggests that additional systems exist to ensure chromosome segregation and cell division (Bailey et al. 2014; Cambridge et al. 2014). A link between the Ter‐matS‐binding MatP protein and ZapB connects the Ter macrodomain of the chromosome to the divisome's ZapB‐ZapA‐FtsZ complex (Espéli et al. 2012) and this may afford the nucleoid itself a role in determining Z ring placement (Rowlett and Margolin 2015; Yu and Margolin 1999).
The Min system operates on Z ring placement through an inhibitory mechanism. This strategy is not used universally among bacteria. For example, positive placement is used to direct Z ring positioning in Streptomyces spp. Here, the SsgA protein takes up position at mid‐cell and recruits a partner, SsgB, which is thought to promote FtsZ polymerisation and to link the Z rings to the membrane. These rings form the basis of sporulation septa in these actinomycetes (Traag and van Wezel 2008; Willemse et al. 2011). Sporulation takes place in the aerial hyphae where up to one hundred division septa are laid down, dividing the hyphae into compartments that each contain one genome copy (Zhang, L. et al. 2016). The SepG membrane protein recruits SsgB to future septum sites; it is also required for nucleoid compaction indicating that plays a role in coordinating chromosome organisation and placement of the division septum (Zhang et al. 2016). The PomZ protein in Myxococcus spp. is also a positive regulator of Z‐ring positioning; PomZ shares with MinD the property of being a ParA‐like ATPase (Treuner‐Lange et al. 2013). It forms a complex with PomX and PomY that moves over the surface of the nucleoid in a biased, randomised motion that becomes constrained at the mid‐cell. Once at the mid‐cell location, PomXYZ recruits FtsZ (Schumacher et al. 2017).
Macrodomains play important roles in determining the choreography of the daughter chromosomes, as these segregate prior to cell division (Espéli et al. 2008). They also correlate with global gene expression patterns, suggesting that the overall gene expression programme of the cell is written into the architecture of the nucleoid (Cameron et al. 2017; Sobetzko 2016; Sobetzko et al. 2012). To appreciate the significance of the connections between nucleoid structure and gene expression, it will be necessary to consider the contributions made to both by variable DNA structure and NAPs.
1.26 DNA in the Bacterial Nucleoid
DNA in bacterial cells is maintained in an underwound state and this affects the shape that the DNA duplex adopts as it seeks to adopt a minimal energy conformation. The underwinding arises due to a deficit in helical turns, i.e. the number of times the two DNA strands twist around the duplex axis. The twist deficiency imparts torsional stress to the duplex, which is relieved by allowing the duplex to adopt a writhed confirmation in which the helical axis coils around itself. This coiling of the already coiled DNA duplex creates supercoiling and has the effect of making the DNA molecule more compact. In the context of the nucleoid, such compaction assists with solving the problem of packaging the genetic material within the cell. The most supercoiled parts of the chromosome form branches, facilitating further compaction.
1.27 DNA Topology
The topological state of a DNA molecule is described by three parameters: the linking number (the number of times one DNA strand crosses the other in the duplex); the twist (the number of complete turns made by the strands around one another along the length of the duplex); and the writhe (the number of writhing turns made by the duplex axis around itself) (Bauer et al. 1980; Boles et al. 1990; Vinograd et al. 1965). Changes to the linking number can be made by breaking one or both strands of the duplex, twisting the DNA with or against the sense of the double helix, and then resealing the strand breaks (Higgins and Vologodskii 2015). Twisting the DNA in keeping with the sense of the double helix tightens the duplex and imparts positive writhe, resulting in positive supercoiling. Twisting the DNA against the sense of the double helix underwinds the molecule, imposing negative writhe and therefore negative supercoiling (Sinden 1994). In nature, negative supercoiling is the norm although positive supercoils do arise naturally, especially during the movement of the DNA replication fork (Figure 1.14) and during transcription (Liu and Wang 1987) (Figure 1.15). In contrast to mesophilic bacteria like E. coli, hyperthermophilic archaea that live in environments that are characterised by very high temperatures possess a reverse gyrase activity and maintain their DNA in a positively supercoiled state (Couturier et al. 2014; Lipscomb et al. 2017). Reverse gyrase, first described in Sulfolobus acidocaldarius in 1984, uses ATP to introduce positive supercoils into DNA (Kikuchi and Asai 1984; Ogawa et al. 2015).
Figure 1.14 The topological consequences of replisome activity. The replication fork, moving from left to right, over winds the DNA ahead to create a domain of positive supercoiling. Unless this is relaxed, the replisome will be unable to advance further. Relaxation of the positively supercoiled domain is carried out by DNA gyrase: it neutralises the positive supercoils by introducing negative ones. In C. crescentus, the GapR protein binds to positively supercoiled DNA and stimulates its relaxation by type II topoisomerases (see Guo et al. 2018). The newly synthesised DNA behind the replisome becomes catenated. These interwound, double‐stranded DNA molecules are decatenated by DNA topoisomerase IV, allowing the daughter chromosomes to be segregated at cell division.
For further information, see Lopez et al. (2012) and Postow et al. (2001).
