Paoletti et al., 2007; Yoshimura et al., 2007, and Hara et al., 2008; **Figure 5B**). *In vivo* cross-linking shows that PlsX interacts with cell division proteins (EzrA, DivIVA), cytoskeletal proteins and several metabolic enzymes, in addition to FtsA. PlsX is in a punctuated pattern in the peripheral membrane and is present at potential division sites independently of FtsA and FtsZ. Inactivation of PlsX leads to aberrant Z-ring formation. Thus, the key enzyme for phospholipid metabolism interacts with the cell division machinery in order to complete septum assembly (Takada et al., 2014). Although PlsX interacts with PlsY, many other enzymes involved in phospholipid synthesis, which localize in the septal region (Nishibori et al., 2005), do not interact with it, except MprF that synthesizes lysylPG (Takada et al., 2014). Depletion of PlsX leads to the cessation of both fatty acid and phospholipid biosynthesis (Paoletti et al., 2007; Yao and Rock, 2013).

### Cell Cycle

Transertion may help provide solutions to the problems of coupling growth to the cell cycle, of triggering chromosome replication, and of coupling chromosome segregation spatially and temporally to cell division. The first problem, the way in which growth is coupled to the cell cycle, has been a mystery that has lasted for over half a century (Schaechter et al., 1958; Osella et al., 2014). Many different metabolic processes occur during growth and these processes are needed for the synthesis of RNA, proteins and lipids. These processes come together in transertion, which is therefore well-placed to integrate metabolic information (Fishov and Norris, 2012b). Moreover, transertion is a primary determinant of the structure and composition of the membrane, which is central to the initiation of DNA replication, segregation and cell division, as well as to the structure of the chromosome, which is kept in an expanded state (Woldringh et al., 1995; Libby et al., 2012). Hence transertion is also well-placed to couple growth to the cell cycle. Consistent with this, major effects of transertion on the microviscosity of the membrane have been shown via inhibition of transcription and translation (Binenbaum et al., 1999). The second problem, the nature of the mechanism that initiates chromosome replication, is another long-standing mystery (Eliasson and Nordström, 1997; Wang et al., 2011). This mechanism in *E. coli* is generally attributed to the relative proportions of the DnaA 'initiator' protein in the ATP-DnaA form, which is active in initiation, and the ADP-DnaA, which is inactive (Castuma et al., 1993). The reasons to invoke lipids in a DnaA-based control of initiation include the involvement of unsaturated fatty acids in initiation (Fralick and Lark, 1973), the promotion of formation of ATP-DnaA *in vitro* by anionic phospholipids (Castuma et al., 1993) and the association of 10% of the DnaA in a cell with the membrane (Regev et al., 2012). Given its role in determining membrane dynamics, transertion may well lie at the heart of the initiator mechanism and act via DnaA and, more generally, via promoting strand separation (Norris and Amar, 2012).

The third problem, the coupling between chromosome segregation and cell division, is essentially a membrane problem in the sense that the cell must arrange for the membrane to invaginate and separate segregated chromosomes. One role for transertion may be to create membrane domains in a particular physico-chemical state around the segregating chromosomes such that a different, 'septal' domain forms by default between them (Norris, 1995; Woldringh, 2002; Norris et al., 2004). This septal domain might then generate tubular or other structures that would recruit and activate division proteins. This hypothesis is supported by evidence for cell cycle variations in the lipid composition of *E. coli* (Mozharov et al., 1985) and in the spatial distribution of the lipids of *Micrococcus luteus* (Welby et al., 1996), for the CL-rich domains at the division sites of *E. coli* (Mileykovskaya and Dowhan, 2000; Koppelman et al., 2001) and *B. subtilis* (Kawai et al., 2004), and the septal location of the polyunsaturated fatty acid eicosapentaenoic acid of *Shewanella livingstonensis Ac10* (Sato et al., 2012). Significantly, distinct domains appear around and between the segregating chromosomes at a very early stage of the cell cycle in *E. coli* and these domains disappear when translation – and hence transertion – is abolished (Fishov and Woldringh, 1999). Division is inhibited by the Min system and lipids are again important (Suefuji et al., 2002; Kretschmer and Schwille, 2014). The NH2 terminal domain of MinC perturbs the interactions between FtsZ monomers within an FtsZ polymer, while the COOH-terminal domain perturbs the lateral association of FtsZ protofilaments as well as interacting with MinD, for references see Kretschmer and Schwille (2014); MinD is a peripheral membrane-binding protein with an ATPase activity that is activated by MinE and that triggers the detachment of MinD from the membrane to prevent MinCD from inhibiting division; MinE may self-assemble on the membrane via an N-terminal helix that acts as an MTS. The idea is that, in *E. coli*, MinD binds cooperatively to the membrane at one pole with MinE forming an "E-ring" on the rim of this MinD zone that induces its disassembly leading to the diffusion of MinD and MinE and repetition of this assembly disassembly process at the other pole; MinC follows these oscillations of MinD and MinE to inhibit division (Rowlett and Margolin, 2013; Zheng et al., 2014 and references therein). *In vitro* and *in vivo*, MinD interacts with anionic lipids, which determine its distribution (Mileykovskaya et al., 2003), as does MinE (Vecchiarelli et al., 2014). In the case of the *E. coli* MinD introduced into *B. subtilis*, MinD makes spirals that coincide with the spirals of what are probably anionic phospholipids (Pavlendova et al., 2010). Moreover, MinD can convert phospholipid vesicles into tubes (Hu et al., 2002), consistent with the possible importance of phospholipid structures in nucleating division (Norris et al., 2004; Li et al., 2011). Finally, in spherical *E. coli* cells, FtsZ is associated with intracellular membraneous structures and MinD accumulates and oscillates between the places where these structures form (Bendezú and de Boer, 2008).

The 2 min or *dcw* cluster includes 16 genes, such as *ftsZ*, involved in peptidoglycan synthesis and cell division, and the promoter at the start of the cluster probably contributes to the transcription of the whole cluster, leading to the synthesis of a quite long mRNA (Hara et al., 1997; Mengin-Lecreulx et al., 1998; Vicente et al., 1998). Transertion of this cluster may therefore be an important factor in the spatio-temporal control of division. Transertion structures not only the membrane but also the cytoplasm thereby raising the possibility that chromosome segregation is accompanied by a phase separation in the cytoplasm that affects membrane dynamics and drives cell division. Hence, it is significant that phase separation of dextran and polyethylene glycol within giant vesicles leads to tubes of membrane forming at their interface (Li et al., 2011).

If transertion from the *dcw* cluster is indeed important, what is the consequence of having transertion from two clusters (one per segregating chromosome)? Are the lipid preferences of the secretion machinery different from those of the proteins encoded by this cluster so that the domain-producing potential of transertion is reduced? In bacterial L-forms, which lack a cell wall, it might be supposed that the requirement for many of the *dcw* genes would be reduced and less transertion would occur. This would be consistent with the findings that, in L-forms of *E. coli*, some of the genes were mutated (Siddiqui et al., 2006) and the levels of FtsZ were fivefold lower (Onoda et al., 2000) whilst in an L-form of *B. subtilis* FtsZ could be eliminated altogether (Leaver et al., 2009).

### Discussion

The different ways of making membrane domains include lipid–lipid, lipid–protein, protein–protein, and polyamine–lipid interactions. By integrating these interactions, transertion plays a central role in the organization of the bacterial cell. It might even be argued that transertion helps provide a conceptual context for thinking about a plethora of intracellular events, structures and processes including the regulation and execution of the cell cycle, hyperstructure dynamics and the origins of life. In 1989, the term 'nucleoid occlusion' was coined to describe the inhibitory effect of the nucleoids on cell division (Cook et al., 1989) and it was suggested that nucleoid occlusion "may be related to a transcription or translation activity of the nucleoid" (Mulder and Woldringh, 1989). A few years later, it was proposed that the coupled transcription-translation-insertion of nascent proteins into and through membrane (i.e., transertion) is the mechanism responsible for nucleoid occlusion (Norris, 1995; Norris and Madsen, 1995; Woldringh, 2002). In this proposal, transertion creates membrane domains at the right times and in the right places not only to control chromosome replication, segregation and cell division but also to create the positive feedback needed for differentiation (Norris, 1995; Norris and Madsen, 1995; Binenbaum et al., 1999; Norris et al., 2004; Norris and Amar, 2012). Recently, it has been suggested that the role of transertion in nucleoid occlusion is complemented by the recruitment of Noc nucleoprotein complexes and associated DNA to the membrane (Adams et al., 2015).

Transertion can generate hyperstructures (Norris et al., 2007a; Llopis et al., 2010). Such transertion hyperstructures form a large class that, along with the other classes of hyperstructures, constitute the bacterial cell. The exact conformation of transertion hyperstructures is unknown but could be very important in bacterial cell physiology if, as proposed, interactions between hyperstructures were to be the primary determinants of the phenotype (Norris et al., 2007a). For example, interactions based on the condensation and decondensation of ions (Manning, 2007) might enable transertion hyperstructures containing membrane domains and/or linear assemblies of macromolecules to drive the cell cycle (Norris and Amar, 2012) whilst the kinetics of the reactions in the vicinity of a membrane domains are likely to be different from elsewhere, possibly due to the structuring of water (Wichmann et al., 2003).

Transertion may help explain the controversial existence of many spiral hyperstructures (Errington, 2015). Suppose a particular protein has a tendency to form a spiral via protein–protein and protein–lipid interactions. In the absence of transertion, this tendency may result in a spiral whereas, in the presence of transertion, the high concentration of nascent proteins and associated lipids in a circular membrane domain may compete effectively for the mature proteins and thereby prevent a spiral forming. Alternatively, transertion might help generate a protein spiral if the ensemble of gene, nascent RNA (and perhaps mature RNA) were constrained to be essentially linear via RNA–RNA interactions etc. Given the potential of a transertion hyperstructure for altering the morphology of the membrane or for directing a protein to the wrong location, avoidance of these inappropriate results of transertion may act as a powerful factor in evolution that affects both the position in a protein of membrane-interacting sequences and the type of translocon with which the nascent protein interacts.

Finally, several coupled processes in addition to transertion exist in modern bacteria. These include transcription-translationassembly as in the case of ribosome synthesis (Woldringh and Nanninga, 1985; French and Miller, 1989; Norris et al., 2007a; Cabrera et al., 2009) and transcription-replication as in the case of initiation of DNA replication and DNA repair (Kogoma, 1997; McGlynn, 2010). It might therefore be expected that these processes in their coupled forms would have played a role in the origins of the first cells. This has been suggested for transertion based on its force-generating properties, which are responsible for expanding the nucleoid in modern bacteria (Woldringh et al., 1995; Cabrera et al., 2009; Libby et al., 2012; Bakshi et al., 2014): in this scenario, transertion would have helped maintain membrane integrity in early cells and have been of selective value even without the advantages of coding (Norris et al., 1999).

### Acknowledgments

KM and HH would like to thank professors emeriti Isao Shibuya and Yoshito Sadaie for discussion and encouragement. We would also like to thank the editor, Arieh Zaritsky and the reviewers, Marc Bramkamp, Ding J. Jin, and Germán Rivas for helpful comments and guidance.

## References

role in selection of the mid-cell division site. *Genes Dev.* 12, 3419–3430. doi: 10.1101/gad.12.21.3419

**Conflict of Interest Statement:** The Associate Editor Arieh Zaritsky declares that, despite being affiliated with the same institution as author Itzhak Fishov, the review process was handled objectively and no conflict of interest exists. The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Copyright © 2015 Matsumoto, Hara, Fishov, Mileykovskaya and Norris. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

# Why do bacteria divide?

*Vic Norris\**

*Laboratory of Microbiology Signals and Microenvironment, Theoretical Biology Unit, University of Rouen, Mont Saint Aignan, France*

The problem of not only how but also *why* cells divide can be tackled using recent ideas. One idea from the origins of life – *Life as independent of its constituents* – is that a living entity like a cell is a particular pattern of connectivity between its constituents. This means that if the growing cell were just to get bigger the average connectivity between its constituents per unit mass – its *cellular connectivity* – would decrease and the cell would lose its identity. The solution is division which restores connectivity. The corollary is that the cell senses decreasing cellular connectivity and uses this information to trigger division. A second idea from phenotypic diversity – *Life on the Scales of Equilibria* – is that a bacterium must find strategies that allow it to both survive and grow. This means that it has learnt to reconcile the opposing constraints that these strategies impose. The solution is that the cell cycle generates daughter cells with different phenotypes based on sufficiently complex equilibrium (E) and non-equilibrium (NE) cellular compounds and structures appropriate for survival and growth, respectively, *alias* 'hyperstructures.' The corollary is that the cell senses both the quantity of E material and the intensity of use of NE material and then uses this information to trigger the cell cycle. A third idea from artificial intelligence – *Competitive Coherence* – is that a cell selects the *active* subset of elements that actively determine its phenotype from a much larger set of available elements. This means that the selection of an active subset of a specific size and composition must be done so as to generate both a coherent cell state, in which the cell's contents work together harmoniously, and a coherent sequence of cell states, each coherent with respect to itself and to an unpredictable environment. The solution is the use of a range of mechanisms ranging from hyperstructure dynamics to the cell cycle itself.

Keywords: neural net, FtsZ, connectivity, heterogeneity, molecular assembly, dualism, competitive coherence, hyperstructure

### Introduction

The very fact of growing is a source of many serious problems that bacteria somehow have to solve. These problems include DNA becoming a limiting factor (thereby disrupting patterns of expression and preventing exponential growth), positive feedback (thereby leading to a single phenotype), over-reliance on fragile, non-equilibrium (**NE**) structures such as those created by the coupling of transcription, translation (thereby making the cell vulnerable to changes in its environment), and unbalanced production of RNA, protein and lipid (Norris, 2011). The problems do not stop there. More generally, bacteria must have found ways to avoid incoherence at the level of both the individual cell and the population so as to be able to not only grow but also survive (arguably, the more important of the two behaviors since a dead cell can never grow)

#### *Edited by:*

*Arieh Zaritsky, Ben-Gurion University of the Negev, Israel*

#### *Reviewed by:*

*Luyan Zulie Ma, Institute of Microbiology–Chinese Academy of Sciences, China Charles Edward Helmstetter, Florida Institute of Technology, USA*

#### *\*Correspondence:*

*Vic Norris, Laboratory of Microbiology Signals and Microenvironment, Theoretical Biology Unit, University of Rouen, Mont Saint Aignan 76821, France<EMAIL_ADDRESS>
#### *Specialty section:*

*This article was submitted to Microbial Physiology and Metabolism, a section of the journal Frontiers in Microbiology*

> *Received: 24 February 2015 Accepted: 31 March 2015 Published: 16 April 2015*

#### *Citation:*

*Norris V (2015) Why do bacteria divide? Front. Microbiol. 6:322. doi: 10.3389/fmicb.2015.00322* in what we have called 'Life on the Scales' (Norris and Amar, 2012a). More generally still, bacteria – like all living objects – risk losing the connectivity between their constituents as they

get bigger (Norris, 2015). This is particularly serious because patterns of connectivity are often claimed to define pretty well everything – as testified by the claims of universality made for self-organized criticality (Bak, 1996) and small world networks (Watts and Strogatz, 1999) – so that a bacterium that continued to grow would eventually lose its identity.

The above problems are intimately related to the cell cycle. In non-differentiating bacteria such as *Escherichia coli*, the cell cycle was long considered to have only the function of replicating the hereditary material and distributing it into two cells that would be effectively identical unless driven down the path of differentiation. This fitted with the idea that the laws underlying cell growth – and, in particular, those directly relevant to the cell cycle – could be found not by studying single cells but rather by studying cells as an aggregate (Cooper, 2006). However, it is now clear that even *E. coli* generates substantial phenotypic diversity (Godin et al., 2010; Wang et al., 2010; Kiviet et al., 2014; Osella et al., 2014; Taheri-Araghi et al., 2015) with cell division playing the key role. It is not therefore surprising that connections have been found between the replication of the bacterial chromosome and processes that include central carbon metabolism (Janniere et al., 2007; Baranska et al., 2013), phospholipid synthesis (Sekimizu and Kornberg, 1988; Xia and Dowhan, 1995), respiration, lipoteichoic acid synthesis, and ribosome biosynthesis (Murray and Koh, 2014). Other connections have also been found between cell division and processes that include fatty acid synthesis (Yao et al., 2012), glycolysis and its offshoots (Weart et al., 2007; Hill et al., 2013; Monahan et al., 2014), and polyphosphate metabolism (Rao et al., 2009; Boutte et al., 2012).

In the above context, it is interesting to revisit cell division in terms of concepts, such as *hyperstructures* or *competitive coherence*, that are either new or under development. The concept of *hyperstructures* is directly relevant to the problem of maintaining connectivity. A hyperstructure is an assembly of elements (such as genes, RNA, proteins, small molecules, and ions) that performs a function and that constitutes a substantial proportion of the cellular material (Norris et al., 2007a; Saier, 2013; Meyer et al., 2014). The existence of the hyperstructure level is a partial solution to the problem of reducing the enormity of phenotype space from the combinations of 1000s of types of macromolecules to combinations of a 100 or so hyperstructures, thereby allowing Darwinian selection to operate (Kauffman, 1996; Norris et al., 2014). Hyperstructures exist in **NE** and equilibrium (**E**) forms with the ratio between these forms proposed to control the cell cycle and to generate phenotype diversity by sensing both the intensity with which **NE** hyperstructures are working and the quantity of **E** hyperstructures that has accumulated (Norris, 2011; Norris and Amar, 2012a); the importance of cell division to the related stability of cellular constituents has been demonstrated by simulation (Kamimura and Kaneko, 2014). The concept of *competitive coherence*, which relies on patterns of connections, has been developed to explain the behavior of biological systems, which include, of course, bacteria (Norris et al., 2014). This behavior is determined by a relatively small, phenotypically *active*, subset of all the constituents available to the bacterium; selection of this subset entails competition between (1) those constituents whose activities are coherent with one another and with the environment and (2) those constituents whose activities are coherent with the previous history of the bacterium. Simulation has shown that a system based on competitive coherence can adapt to growth and stress conditions (Norris et al., 2012). Moreover, the concept can be extended to include the role of **NE** and **E** hyperstructures and the role of the cell cycle in the nature and size of the active subset (Norris et al., 2014).

According to Krakauer (2014) at the Santa Fe Institute, this institute "is about circling a phenomenon, considering all the angles and unique perspectives to see a thing in a completely new light." Such circling could be likened to someone preparing to climb a mountain shrouded in fog who tries many routes that may be useful-even if they do not lead to the summit-because they give new views. This is the approach to cell division that I adopt here.

### Maintenance of Connectivity

A bacterium is a physical object but what exactly is a physical object? Put differently, what makes one physical object different from another. A crab-like alien examining the Earth from orbit might mistake me and the chair on which I am sitting for an eight-legged crab. The reason the alien would be wrong is that there is a frontier between my chair and me. My constituent bits or elements are more connected to one another than they are to the chair's bits and *vice versa*. More generally, a physical object can be said to exist in some space if there is a topologically closed (or nearly closed) discontinuity in the connectivity of the elements in that space; this discontinuity constitutes a frontier of connectivity that makes the object different from its environment. The exact nature of this object is defined by the pattern of connectivity of its constituent elements and by the pattern of connectivity of these elements to its environment. In other words, a physical object owes its identity and its very existence to patterns of connectivity. This is one interpretation that can be made of the picture by Mary Cassatt of her dying sister, Lydia (**Figure 1**).

A bacterium is, of course, more than just a physical object, it is a *living* physical object. One of the characteristics of living objects such as bacteria is that at some stage they grow (or, at least, their ancestors grew). Growth entails an increase in mass *via* the addition of new elements or constituents. In the case of a bacterium, these constituents may be the same as those already there (as shown for the constituents numbered 1–6 in **Figure 2A**); hence, the addition of these new constituents may leave the average connectivity unchanged, where connectivity, L/E, equals the number of links or physical associations, L, divided by the number of constituents or elements, **E** (**Figure 2A**). In other words, growth does not necessarily change the number or nature of the connections between the constituents themselves and L/E either remains constant (**Figure 2A**) or even drops (**Figure 2B**). Growth does, of course, mean an increase in the mass of the bacterium and this in turn means that the average connectivity

FIGURE 1 | Profile of Lydia Cassatt (1880). This portrait entitled Autumn of the seriously ill Lydia by her sister, Mary Cassatt, can be interpreted as showing her losing her identity (copied from http://hoocher.com/Mary\_ Cassatt/Mary\_Cassatt.htm original in the Musee du Petit Palais, Paris, France).

*per mass M of the bacterium*, (L/E)/M *alias cellular connectivity*, decreases. To make this clearer, consider a toy cell that comprises six different elements each of which binds to two other elements (such that the connectivity, L/E, is one). Consider too that this cell manages to grow so as to duplicate exactly its six constituents, as shown by the two sets of differently colored constituents in **Figure 2A**. Its L/E either remains one or, if this duplication is not exact, drops below one, as shown by the relative excess of constituents number 1 in **Figure 2B**. However, connectivity divided by mass has halved. Why does this matter?

Scaling up a process or a structure often results in a changed connectivity and a changed behavior. This is one of the dangers posed by growth: identity is changed. Consider a bacterium that has grown much bigger than normal. It would have different mechanical properties (and different resistance to mechanical stresses), a different motility, differences in regulatory networks (the behavior of the ensemble of two identical networks can be qualitatively different from that of one of these networks on its own), possibly differences in the surface/volume ratio (see, Bendezu and de Boer, 2008), etc. Not surprisingly, the changing connections that accompany increased hydration create problems for cells (Lang, 2011). The change in identity that accompanies extensive growth is not always negative. Such growth is, for example, fundamental to the change in the identity of an *E. coli* cell as it becomes a filamentous, multinucleate, hyperflagellate cell that can swarm over the surface of solid media (Harshey and Matsuyama, 1994). That said, the reduction of cellular connectivity that accompanies growth generally constitutes a serious problem constraining the evolution of cells.

It turns out that there are several possible solutions to the problem of how to maintain cellular connectivity despite growth. One of them involves the creation or the expansion of an additional level of organization. Multi-level structuring is a fundamental characteristic of living systems. Each new emergent level of organization in the dynamics of a complex self-organizing system – such as a living system – re-organizes variety on the level below so that it has meaning for the level above (Salthe, 1985; Lemke, 2000). The physical nature of the processes, the spatio-temporal scales, and connections at a particular level may be characteristic of that level (Norris et al., 2007a). In the case of the individual bacterium, I would argue that the level of

(L:E) is 1 and stays 1 in the above case as the cell grows. However, this ratio per mass goes down with growth as (L:E)/M goes to (L:E)/2M. (B) After growth (100% increase in mass), the (L:E)/M has gone down from 1 to (6:12)/2M, i.e., 0.25.

organization that is relevant to the cellular connectivity problem is that of hyperstructures, which is the level intermediate between macromolecules and the bacterial cell itself (Norris et al., 2007a). There are several types of hyperstructure (Norris et al., 2007a). In the case of enzymic hyperstructures, a recent study suggests that a quarter of known cytoplasmic enzymes are in hyperstructures in which they have a higher within-group connectivity than enzymes that appear to be outside hyperstructures (Meyer et al., 2014). The relevance of the hyperstructure concept to growth is that, as the bacterium grows, the dynamics of hyperstructures changes such that some hyperstructures are born and others are consolidated; in other words, connectivity at the level of hyperstructures increases, thereby compensating for the decrease in connectivity at the level of macromolecules and maintaining the average connectivity per mass (i.e., cellular connectivity). A prime candidate for a hyperstructure that is born during growth would be the initiation hyperstructure based on the DnaA protein whilst candidates for hyperstructures that are consolidated during growth would include the ribosomal hyperstructure or microcompartment where ribosomal constituents are made and perhaps assembled (see, Intensity Sensing and Quantity Sensing; Norris and Amar, 2012a). These different hyperstructures and their dynamics may be in part responsible for the relationship between the fluidity of the cytoplasm and metabolism (Parry et al., 2014).

Another possible solution to the cellular connectivity problem involves cell division. To simplify it, cell division produces two daughter cells in which the average connectivity per constituent per mass is restored to that of the parental cell when it was born. A population of bacteria such as *E. coli* usually has a size distribution – characteristic of the rate of growth – that is the culmination of the cell cycle in which mass plays an important regulatory role (see below; Schaechter et al., 1958). It has been proposed that the timing and locating of the division site are the result of what is essentially a local reduction in the connectivity of the membrane *alias* the frontier of discontinuity with the environment (Norris et al., 2004). Evidence for this includes the appearance of membrane domains at the site of division (Mileykovskaya and Dowhan, 2000; Kawai et al., 2004), which constitutes the earliest known step of the formation of the division site in *E. coli* (Fishov and Woldringh, 1999). These anionic lipid-rich domains form in the regions outside (including between) the chromosomes, presumably by default because the membrane around the chromosomes is both enriched in other lipids and structured by the coupling between the processes of transcription, translation, and insertion of nascent proteins into membrane *alias transertion* which increases the microviscosity of the membrane (Binenbaum et al., 1999). Such relatively unconnected domains at the division site may have a propensity to invaginate as shown in phase separation experiments (Li et al., 2011). Moreover, excess production of membrane – which I would argue results in a region of relatively unconnected membrane – results in division in wall-less bacteria (Mercier et al., 2013) and can titrate division enzymes such as FtsZ and MinD in certain shape mutants of *E. coli* (Bendezu and de Boer, 2008). A lower density of connections between envelope proteins, peptidoglycan, and the inner membrane has been proposed to explain the release of membrane vesicles from Gram-negative bacteria and, in particular, from their the division sites (Deatherage et al., 2009). Despite my focus here on the membrane as the important frontier of connectivity, it should be noted that two classes of models of the cell are proposed to exist, one in which the membrane is all-important in defining the cell and a second in which the entire cell is in a different phase relative to its surroundings (Jaeken and Vasilievich Matveev, 2012). Hence, a division site might result not only from a local reduction in the connectivity of the membrane but also from a frontier of discontinuity forming in the interior of the cell (as might be argued in the case of the phragmoplast in plant cells). Indeed, when two liquid phases are in contact with a membrane, their interactions with that membrane can lead to budding (Li et al., 2012).

### Dualism

An apparently intractable dilemma often confronts bacteria: either to interact, take risks and grow, or to minimize interactions, avoid risks, shut down, and survive. A bacterium that tries to grow forever will eventually run out of resources and to survive must abandon this strategy. Conversely, a bacterium that refuses to grow whilst others grow successfully must abandon this strategy if it is not to be out-competed. A strong evolutionary pressure therefore exists to compel bacteria to choose between at least two apparently incompatible strategies or, to put it differently, to navigate in phenotype space between the two basins of growth and survival. This is the problem of *Life on the Scales* (Norris and Amar, 2012b).

The solution adopted by bacteria is to generate a phenotypically diverse population (Smits et al., 2006; Maisonneuve and Gerdes, 2014; Serra et al., 2014). This diversity extends to the growth rate itself such that, within a growing population of *E. coli*, non-growing persisters are produced at a low frequency (Balaban et al., 2004). Growth rate diversity is, however, much greater than this. Measurements of the buoyant mass of individual *E. coli*reveal a considerable variation in 'instantaneous' growth rates (Godin et al., 2010) as do measurements of the elongation rates of individual *E. coli* (Wang et al., 2010; Kiviet et al., 2014; Osella et al., 2014).

The many ways to achieve a phenotypically diverse population include multi-stationarity in networks (Thomas, 1980; Dubnau and Losick, 2006), noise (Silva-Rocha and de Lorenzo, 2010), spontaneous gene amplification (Anderson and Roth, 1977), and chemical communication (Vega et al., 2012). These mechanisms may not necessarily lead to the satisfaction of two requirements: the phenotypes generated need to be both (1) internally coherent and (2) coherent with respect to the other phenotypes in the population. In the case of the former requirement, if an evolutionarily useful, phenotypic diversity is to be generated, each phenotype must be coherent with respect to the set of genes expressed (Norris and Amar, 2012a). This requirement for coherence is strong in the case of the growth rate since growth and survival strategies are incompatible at the level of the individual cell.

An evident mechanism to generate coherent phenotypic diversity is the cell cycle itself. Each division in differentiating, bacterial species such as *Caulobacter crescentus* generates two cells with very different metabolisms (Bowman et al., 2013). It is therefore conceivable that even in non-differentiating bacteria such as *E. coli*, the cell cycle is the primary mechanism for generating daughter cells with different growth rates. Indeed, the authors' interpretation of the results of a microfluidics-based experiment on the growth rate distribution of *E. coli* is that the cell "forgets" on division its growth rate in the previous cell cycle (Wang et al., 2010); further analysis of these results confirms the importance of cell division in generating diversity (Osella et al., 2014). The question is how.

The answer is that differentiation far from being difficult to achieve is difficult to avoid. Differentiation is almost inescapable in systems where negative feedback operates globally and positive feedback operates locally (Norris, 1995b; Norris and Madsen, 1995). Consider two similar research laboratories competing with others for limited funds (i.e., a globally negative feedback); if one of these laboratories is actually funded and the other is not, in the next competition, the previously funded laboratory is at an advantage (i.e., a locally positive feedback). During the cell cycle of modern bacteria, two – what are for my purpose here – essentially identical chromosomes compete for limiting factors like RNA polymerases and ribosomes; this constitutes negative regulation *in trans*. At the same time, a region of one chromosome that is being expressed by polymerases is pulled by these polymerases out of the nucleoid (where it would otherwise be buried and relatively inaccessible) and this expression increases the region's accessibility to more polymerases; this constitutes positive regulation *in cis*; in addition, transertion allows ribosomal subunits to penetrate the nucleoid and to perform cotranscriptional translation (Bakshi et al., 2014), which in turn would protect nascent mRNA (Deana and Belasco, 2005; Deneke et al., 2013), limit the formation of the RNA degradosome (Strahl et al., 2015) and stop RNA polymerases backtracking (McGary and Nudler, 2013). The two chromosomes therefore spontaneously have different patterns of expression (Norris, 1995b; Norris and Madsen, 1995). This 'globally negative, locally positive' mechanism might explain the generation of daughters with different phenotypes but not the coherence of these phenotypes.

There are many ways to achieve a diversity that is coherent (Norris et al., 2014). One of them is *dualism* (Norris, 2011). It works like this: the cell cycle generates two daughter cells with different ratios of **NE** and **E** hyperstructures; these different ratios confer different properties on the daughter cells such that those with higher **NE**:**E** ratios grow fast to profit from nutrient availability whilst those with lower **NE**:**E** ratios grow slowly or cease growth so as to resist stresses. These ratios give rise to coherent phenotypes because one of the parental strands of DNA is physically associated with the **NE** hyperstructures appropriate for growth whilst the other strand is physically associated with the **E** hyperstructures appropriate for survival (Rocha et al., 2003). The logic here is that, in a competitive cytoplasm, an old strand associated with a particular hyperstructure (e.g., *via* coupled transcription–translation) has more chance of retaining that hyperstructure than a chemically identical, newly replicated (and therefore unexpressed) strand has of acquiring that hyperstructure (**Figure 3**). Hence, the older strands have the advantage in terms of making the major contribution to the phenotypes of the daughter cells. Evidence consistent with dualism includes the different locations of the leading and lagging strands (White et al., 2008) and the colocation of genes and their products (Llopis et al., 2010). In the dualism hypothesis, it is essential to achieving a phenotypically diverse population that the replication of the chromosome be followed by cell division since this puts these chromosomes into separate cells. Coherent separation leading to division can, of course, be achieved by several, complementary, physical mechanisms as shown by lipid vesicles that encapsulate an aqueous two-phase system of polyethylene glycol and dextran within a membrane of liquid-disordered and liquid-ordered domains (Andes-Koback and Keating, 2011). The philosophical rationale behind this physical separation is that the regulatory and metabolic networks that depend on these chromosomes can disrupt one another if they are not separated, as illustrated by a simple model based on artificial chemistry (Demarty et al., 2003).

In discussing the maintenance of cellular connectivity, I proposed above that cell division maintains identity or phenotype by regenerating the original patterns of connectivity. This can now be seen to be an over-simplification. More exactly, division is fundamental to generating the two classes of patterns of connectivity that allow a population to live on the scales. In other words, even in steady-state conditions, growth, and division create a bacterial population with a constant distribution of varied but coherent phenotypes that interchange over successive generations.

### Intensity Sensing and Quantity Sensing

The cell cycle is also the solution to another two fundamental problems. The *intensity* problem that confronts bacteria during growth is that the **NE** constituents that do the work, like enzymes, eventually work with such intensity that they can do no more; the cell can then do no better than grow linearly. The *quantity* problem that also confronts bacteria during growth is that the quantity of unused, **E** material – be it lipids or macromolecules – accumulates; this material risks being a waste of resources. By sensing both intensity and quantity, a bacterium could decide when it was time to increase the number of enzymes and to convert the unused macromolecules into another form. In the case of *E. coli*, we have proposed that this combination of intensity and quantity sensing reaches a threshold that triggers the cell cycle and it is then the events of the cell cycle that solve both intensity and quantity problems (Norris, 2011; Norris and Amar, 2012a; Norris et al., 2014). The idea is that the state of one or more **NE** hyperstructures reflects the intensity of metabolism at that *particular moment in time* and that when this intensity is sufficiently high for growth to risk growth being limited, one or more signals are emitted by these hyperstructures to initiate chromosome replication. In addition, the state of one or more **E** hyperstructures senses the quantity of accumulated material and when this is likely to be sufficient for daughter cells to be generated, a complementary, initiation signal is emitted. These signals trigger the cell cycle which has the function of converting **E** into **NE** material and redistributing this material into the daughter cells.

There are several mechanisms that might be responsible for intensity sensing; one attractive mechanism could be based on interactions between cytoskeletal hyperstructures and metabolic enzymes such that, for example, an enzyme that is catalyzing its reaction has a greater affinity for the replication and/or cytoskeletal hyperstructures and helps to stabilize/destabilize them (Norris et al., 2013). This actually occurs in *Bacillus subtilis* where assembly of FtsZ into an effective division structure is inhibited by interaction between FtsZ and the glucosyltransferase UgtP, which depends on the concentration of UDP-glucose (Weart et al., 2007); a similar system exists in *E. coli* where the interaction is between FtsZ and the functional homolog of UgtP, OpgH (Hill et al., 2013). FtsZ dynamics are also affected by the location of pyruvate dehydrogenase E1α, which depends on the concentration of pyruvate (Monahan et al., 2014). Another mechanism might involve sensing the density of RNA polymerases transcribing genes and of ribosomes translating mRNA (Norris, 1995c); in line with this, a correlation exists between attaining a particular growth rate per unit mass in mouse lymphoblasts and their entry into S phase (Son et al., 2012). There are also several mechanisms that might be responsible for quantity sensing and that, for example, might be based on the possible accumulation of proteins and lipids (Norris and Amar, 2012a). In the former case, examples might include the wall synthesis enzyme, MurG, which is stored in an inactive form in the cell poles (Michaelis and Gitai, 2010) and the CTP synthetase, CtpS, which is stored in an inactive form as a polymer (Barry et al., 2014). In the latter case, accumulation of cardiolipin would be an attractive candidate given its roles in initiation of replication (Sekimizu and Kornberg, 1988; Makise et al., 2002) and in cell division (Mileykovskaya and Dowhan, 2000; Koppelman et al., 2001; Kawai et al., 2004). Finally, there are molecules of major physiological importance that are often overlooked (Norris et al., 2015); these include polyphosphate which, in some bacteria, has been shown to play a role in the cell cycle (Boutte et al., 2012) and to exist in both soluble and insoluble forms (Klauth et al., 2006).

### Competitive Coherence

*Competitive coherence* is a particular way of choosing a temporal series of subsets of elements from a much larger set (Norris et al., 2012). These subsets are *active* subsets insofar as a subset actively determines the behavior of the entire system at a particular time. Such a subset (*alias* the phenotype) is selected by a competition between (1) those elements that have a relationship with the elements in the previous subset and (2) those elements that have a relationship with the elements already selected to be part of the new, emerging subset. At the level of a bacterium, growth and survival require selection of an active subset of macromolecules in response to external and internal conditions; such responses entail both the generation of a coherent cell state, in which the cell's contents work together efficiently and harmoniously, and the generation of a coherent sequence of cell states. Incoherence within a cell state is punished since, for example, a cell that simultaneously induces the expression of genes for growth at high temperature and at low temperature is likely to be outcompeted by rival cells that induce each set of genes only when needed. Incoherence in the succession of cell states is also punished since a cell that goes from one cell state to another very different one (without good environmental reason) is wasting resources. A strong selective pressure, therefore, exists to generate active subsets of elements to provide both coherent cell states and a coherent sequence of such states (Norris et al., 2014). In response to this pressure, the process of competitive coherence has evolved to reconcile two sorts of coherence in generating the active subset. Note that the active subset is not the same as the bacterium's entire set of genes, set of mRNAs and/or set of proteins: the active subset is the set of elements that determines the behavior of the cell at a particular time. Hence, a gene that is not expressed or an mRNA that is not translated or an enzyme that is not in the right hyperstructure might have no effect on the phenotype at a particular time and therefore would not be part of the active subset. The question therefore arises as to whether the concept of competitive coherence can provide an insight into the cell cycle.

One of the important parameters in competitive coherence is the nature and size of the active subset. It is far from clear, however, how the size of the active subset should vary with conditions. In a bacterium growing rapidly, the need for it to be ready to adapt to new opportunities or stresses might be better served by a large active subset containing mainly **NE** elements whilst in a non-growing bacterium the need for it to maintain its structures (rather than abandon them too readily so as to profit from a growth opportunity) may require a small active subset containing mainly **E** elements. It might be argued that in stress conditions, the active subset may be smaller in order to conserve energy and to lose diversity as the system concentrates on a limited number of **E** hyperstructures in which the elements are tightly connected with one another (Norris et al., 2014). In changing conditions, however, we predict that the active subset would also change and go through a period when it changes in size, as proposed for *Streptomyces coelicolor* as it goes through changes in nutrient availability and population density (Vohradsky and Ramsden, 2001).

Changes in the active subset is where the cell cycle comes in. Cell division permits activity subsets not just with different contents but also of different sizes. One question here is what would happen to the active subset if a bacterium were to grow without going through the cell cycle. Presumably, RNA and protein production would eventually become limited by the relative scarcity of DNA so that exponential growth could not be sustained (Zaritsky, 1975; Norris, 2011). It is tempting to think that, since many different genes compete for an increasing number of RNA polymerases (Stickle et al., 1994), continuation of growth in the absence of a cell cycle would lead to the inappropriate transcription of an increasing number of genes and to an incoherent phenotype, but this must depend to some extent on how the regulatory network would change. Assuming an increase in transcriptional and translational machinery relative to unit DNA, growth without cell cycle progress should lead to a gene encoding a repressor reaching a maximum level of expression and the repressor eventually being diluted by growth and becoming insufficient to repress its targets. It is therefore conceivable that the absence of the cell cycle would result in an increase in the size of the active subset such that all the phenotypes in the population converged onto a single phenotype. Inhibition of chromosome replication in *B. subtilis*, accompanied by continued growth led to significant alterations in the expression of over a 100 genes, around a half being regulated by DnaA, which is both an initiator of replication and a transcription factor (Goranov et al., 2005). In this experiment, the period of unbalanced growth lasted for a maximum of 90 min and only the mRNA levels of the entire population could be measured. To explore the ideas outlined above would require following the global patterns of transcription and translation in individual cells growing over several hours without cell cycle progress. Unfortunately, this is at present impossible. In summary, once again, the cell cycle is about the preservation of the coherence of individual phenotypes and the coherence of the population of phenotypes.

### Discussion

It is somewhat surprising that after decades of intense research into the cell cycle of some of biology's best understood model organisms, the fundamental nature of the control over the cell cycle remains elusive and, in particular, the coupling between the cell cycle and the growth rate (Marshall et al., 2012; Campos et al., 2014; Taheri-Araghi et al., 2015). One explanation is that the paradigm that has guided this research is wrong. If so, a new paradigm is required. This is easier said than done. The approach to generating new paradigms that I advocate here is to ask the question "why do bacteria divide?" Such 'existentialist' questions are fundamental and attempts to answer them can lead to new insights. Some of us have considered explanations for cell division in terms of the maintenance of cellular connectivity, the reconciliation of growth and survival strategies, and combining present coherence with past coherence. Could these ideas lead to a new paradigm and, if so, would we know if it were a good one?

A new paradigm might be evaluated on grounds that include its esthetic qualities, its testability, its difference with the current paradigm, and the breadth of its fields of application (Norris, 1995a). In other words, does the new paradigm result in a rapid advance in a specific field of science, and does it help us understand other fields? One approach is to focus on those *unifying* problems that recur in different fields, for example, how to limit the enormity of phenotype space so that Darwinian selection can act (Kauffman, 1996). To solve this problem, we have invoked a key role for simple, universal molecules and inorganic ions, termed SUMIs, which include polyphosphate, polyhydroxybutyrate, polyamines, and calcium (Norris et al., 2015). A complementary approach is to ask whether the paradigm would help elucidate another field, for example, could a hypothesis developed for the origins of life help with cell division? This may be the case for the SUMIs, all of which have major roles in the growth, cell cycle, and differentiation of bacteria (Huang and Reusch, 1996; Igarashi and Kashiwagi, 2006; Naseem et al., 2009; Rao et al., 2009; Boutte et al., 2012). It may also be that case for **NE** and **E** hyperstructures, dualism, and life on the scales, which we would argue are as important to the division of modern cells as they were to the origins of protocells in the prebiotic ecology (Norris and Raine, 1998; Hunding et al., 2006; Norris et al., 2007b). Indeed, concepts such as life on the scales and competitive coherence are essentially scale-free and substance-free, and are applicable to life elsewhere in the cosmos (Norris, 2015).

In discussing the maintenance of cellular connectivity, we proposed that cell division maintains identity by regenerating the original patterns of connectivity. This is clearly an oversimplification since division is fundamental to generating the two patterns of connectivity that allow a population to live on the scales. The point is that growth and division create a population of individuals with a constant distribution of phenotypes or identities that interchange over successive generations. Ironically, this fundamental role for the cell cycle may have been obscured by the importance – admittedly understandable – attributed to only studying the cell cycle of bacteria growing in steady-state conditions which minimize differentiation (Schaechter et al., 1958; Fishov et al., 1995).

### Acknowledgments

I thank Andreas Dress, Alex Grossman, Marie Beurton-Aimar and the reviewers for helpful comments.

### References

**Conflict of Interest Statement:** The author declares that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Copyright © 2015 Norris. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

# **How much territory can a single** *E. coli* **cell control?**

### *Ziad W. El-Hajj\* and Elaine B. Newman*

*Department of Biology, Concordia University, Montreal, QC, Canada*

Bacteria have been traditionally classified in terms of size and shape and are best known for their very small size. *Escherichia coli* cells in particular are small rods, each 1–2 µ. However, the size varies with the medium, and faster growing cells are larger because they must have more ribosomes to make more protoplasm per unit time, and ribosomes take up space. Indeed, Maaløe's experiments on how *E. coli* establishes its size began with shifts between rich and poor media. Recently much larger bacteria have been described, including *Epulopiscium fishelsoni* at 700 µm and *Thiomargarita namibiensis* at 750 µm. These are not only much longer than *E. coli* cells but also much wider, necessitating considerable intracellular organization. *Epulopiscium* cells for instance, at 80 µm wide, enclose a large enough volume of cytoplasm to present it with major transport problems. This review surveys *E. coli* cells much longer than those which grow in nature and in usual lab cultures. These include cells mutated in a single gene (*metK*) which are 2–4*×* longer than their non-mutated parent. This *metK* mutant stops dividing when slowly starved of *S*-adenosylmethionine but continues to elongate to 50 µm and more. FtsZ mutants have been routinely isolated as long cells which form during growth at 42°C. The SOS response is a well-characterized regulatory network that is activated in response to DNA damage and also results in cell elongation. Our champion elongated *E. coli* is a *metK* strain with a further, as yet unidentified mutation, which reaches 750 µm with no internal divisions and no increase in width.

#### **Keywords:** *E. coli***, cell length, cell division, giant bacteria, metabolism**

## **Introduction**

*Escherichia coli* has astonished investigators with its remarkable metabolic efficiency packed into such a small size. In its 0.5–2 µ length, it packs its genetic material, its metabolic machinery, and an impressive variety of adaptive strategies. It can make a new cell as fast as every 30 min with scarcely an error. The brilliant analysis of *E. coli* function by Jacob, Monod, and Lwoff excited the entire field and led to the amazing detail with which *E. coli* is now understood (Cohn, 2014).

In order to produce a new cell, *E. coli* must approximately double its cell contents and distribute them between 2 daughter cells. It must exactly duplicate and segregate its DNA, and it must double its length and divide itself at midcell. It becomes longer using a cell wall synthesizing system based on penicillin binding protein 2 (PBP2) to elongate. This elongation is the result of the combined activity of peptidoglycan synthesis and hydrolysis enzymes, which constantly remodel the cell wall, but the net result is an increase in cell length (Johnson et al., 2013). The direction of cell wall synthesis changes when the length has doubled, uses a different enzyme system based on PBP3, and coincides with synthesis of a septum at midcell. This system, known as binary fission, is thus an alternation between elongation via a PBP2 complex and division via a PBP3 complex (Lutkenhaus et al., 2012).

#### *Edited by:*

*Arieh Zaritsky, Ben-Gurion University of the Negev, Israel*

#### *Reviewed by:*

*Moselio Schaechter, San Diego State University, USA William Margolin, University of Texas Health Science Center at Houston, USA James R. Walker, University of Texas at Austin, USA*

#### *\*Correspondence:*

*Ziad W. El-Hajj, Department of Biology, Concordia University, 1455 De Maisonneuve Boulevard, Montreal, QC H3G 1M8, Canada<EMAIL_ADDRESS>
#### *Specialty section:*

*This article was submitted to Microbial Physiology and Metabolism, a section of the journal Frontiers in Microbiology*

> *Received: 24 February 2015 Accepted: 29 March 2015 Published: 21 April 2015*

#### *Citation:*

*El-Hajj ZW and Newman EB (2015) How much territory can a single E. coli cell control? Front. Microbiol. 6:309. doi: 10.3389/fmicb.2015.00309* El-Hajj and Newman Review on very long bacteria

Initiation of the septum at midcell involves spatial inhibitors that prevent septum formation elsewhere, such as SlmA involved in nucleoid occlusion (Du and Lutkenhaus, 2014), and the wellknown MinCD complex (Ghosal et al., 2014). Although the mechanisms by which they inhibit division are relatively well understood, how *E. coli* finds its mid-point in the first place has been a long standing problem. This was settled very recently by the lab of Suckjoon Jun, who showed that it divides when it has added a constant volume, the rate depending on how fast its environment allows it to do so (Taheri-Araghi et al., 2015). The cell alters its volume and length according to its environment. However in whatever conditions it can grow, it makes viable cells and wastes nothing, i.e., except for the end products of metabolism, it does not overproduce and excrete metabolic products.

### **Various Ways to Grow Very Long** *E. coli*

It would make sense that if one were to inhibit the activation of the PBP3 divisome in such a way that everything else functions, the cell would not divide, nor would it stop growing. It would continue its various metabolic functions and become very long. This is indeed what seems to occur in our strain MNR2 (see Extremely Elongated *E. coli* Cells). In this section we will discuss the problems that such an elongating cell might face, and assess the role of two amino acids (methionine and L-serine) and of alternative cell envelope components in promoting elongation.

### **Potential Problems in Elongation**

The elongating cell has a number of problems, and some might be expected to increase in severity the longer it gets. Among these, it has to add peptidoglycan to the wall, it has to synthesize and distribute DNA, it has to transcribe from the new and old DNA, make new ribosomes and distribute those, and use them to make new proteins and enzymes, and form new enzyme complexes and distribute them.

As the cell lengthens, its mass clearly must increase and each addition it makes to its protoplasm and its peptidoglycan must be pushing against more mass and more wall. If this caused it problems, one might expect it to grow more slowly than a cell which divides every time it doubles, and perhaps more important, to slow down as it becomes longer. This effect might be lessened by the saving in time and energy occasioned by not making the vertical cell wall.

However, a cell elongates by adding many short lengths of new peptidoglycan at many points along its surface, thus providing new space for the macromolecules it is making, and for the duplicating DNA to move into. Because the additions are many and short, rather than few and long, elongation may require less forces. If the sites are numerous enough, they may not even have to have a fixed pattern to maintain a linear form overall. Otherwise there must be some pattern to where PBP2 acts. In any case this forms an unobstructed tube within which cytoplasmic components can move.

Many physical chemists, and others, suggest that *E. coli* cytoplasm is extremely crowded, and this is often supposed to interfere with diffusion and distribution of cell contents. By one estimate, its cytoplasm contains 200 mg/ml protein, 11–18 mg/ml DNA and 75–120 mg/ml RNA occupying 20–40% of the cell volume and affecting transit though much more "excluded volume" (Hasnain et al., 2014). They suggest this implies that it is not easy for even small molecules to make their way through the cytoplasm, and even harder for large molecules.

The effects of crowding are considered in interesting detail in a 1999 review (Hoppert and Mayer, 1999). These authors suggest that bacterial cells have functional compartments but they are not bounded by membranes. Instead they are formed from the nucleoid, from multienzyme complexes, from storage granules, and cytoskeletal elements all of which affect the properties of water in the cell (Hoppert and Mayer, 1999).

These papers, and many others, make it clear that the cell is crowded with respect to macromolecules. However, the rapidity of elongation suggests that this apparent crowding does not actually slow synthesis much or inhibit function of newly synthesized and distributed material, no doubt because of the continuous elongation of the tube. This is perhaps another example of *E. coli* being able to solve its problems better than we can understand them.

Indeed one of the problems of "crowded cytoplasm" considerations is that it does not allow for the nature of growing cells and turbulence within expanding cytoplasm. While the wall is being extended by hundreds of insertions, the DNA is being synthesized and segregated, and causing turbulence in the cytoplasm around it, which must aid the motion of other molecules in the cytoplasm. Small molecules can probably move with very little obstruction.

### **Involvement of Methionine**

We have found several cases in which production of long cells in *E. coli* is associated with alterations in methionine and *S*-adenosylmethionine (SAM) metabolism. The first long *E. coli* isolated in the Newman lab was derived from a supposed *metK* mutant described in R. C. Greene's lab (Greene et al., 1973). This carried a point mutation in *metK* together with an *lrp* mutation which decreased Lrp expression and increased *metK* expression just enough to permit growth (Newman et al., 1998). We showed that a strain carrying only the *metK* mutation required leucine for growth, as an inducer of the *lrp* gene. When grown overnight with limiting leucine, e.g., 5 µg/ml, the cells did not divide but elongated up to 100-fold with imperfectly segregated nuclei and no visible constriction (Newman et al., 1998). Whereas other cell division mutants (such as strain MNR2 or *ftsZ84* mutants) made no septum at all, the SAM-starved cells incorporated three proteins into the septum, FtsZ, ZipA, and FtsA, but continued to elongate with these partial septa (Wang et al., 2005).

These cultures probably did not represent the full potential for elongation of the *metK* strain. The cells could elongate only once they reduced the leucine sufficiently to inhibit division, but then quickly ran out of leucine for protein synthesis. The 100-fold elongation is thus quite prodigious. We conclude that a supply of SAM is needed for complete activation of septum formation, and suggest that this is due to the need for one or more methyltransferase reactions in activating division.

We could not make a strain carrying a deletion of *metK* because *E. coli* cannot transport SAM, but we did construct a deletion compensated by a plasmid-carried inducible *metK* (Wei and Newman, 2002). However, the group of D. O. Wood created a SAM-transporting *E. coli* strain by isolating a SAM transporter from *Rickettsia prowazekii* (Tucker et al., 2003) and cloning it into an *E. coli* plasmid, allowing a mutant deficient in *metK* to grow in rich medium with an exogenous SAM supply (Driskell et al., 2005). They kindly allowed us to use this transport gene for all our work on SAM utilization including strain MNR2.

Using the *metK* deletion strain of Wei Yuhong, and the Wood plasmid, with some modifications, we were able to determine the growth requirements of an *E. coli metK* mutant (El-Hajj et al., 2013). The mutant in fact had two requirements, SAM because of the deletion, and methionine because in the presence of SAM, methionine synthesis is inhibited.

As we discovered with considerable difficulty and chagrin, using externally provided SAM is a matter of real complexity. We have described these problems in DETAIL (El-Hajj et al., 2013) and anyone wishing to use commercial SAM should read that description. We used commercial SAM, purified of course, for our experiments (before understanding all these complexities) but would make our own enzymatically for any further experiments.

After a year on the lab shelf, commercial SAM is a 1:1 mixture of its enantiomers *(R)*-SAM and *(S)*-SAM, even if it was provided as pure *(S)*-SAM. Enzymatic SAM is purely *(S)*-SAM when first made but isomerizes relatively quickly. Only *(S)*-SAM is a methyl donor for most *E. coli* methyltransferases. However,*(R)*-SAM provided exogenously is a methyl donor for one reaction, the methylation of homocysteine to methionine by the enzyme MmuM, a third methionine synthase.

We found then that the SAM deletion strain requires both SAM and methionine, but the amount of methionine it requires depends on the proportion of SAM which is in the form of *(R)*-SAM and that proportion varies greatly. However with more *(R)*-SAM than it needs, the cell still needs methionine though very little—about 100 ng/ml rather than 20–40 µg usually provided to a methionine-requiring mutant such as a *metB*. We do not understand what the cell needs the small amount of methionine for, when it is getting so much methionine from *(R)*-SAM and is not actually blocked in methionine biosynthesis.

We conclude that methionine metabolism is less well understood than generally thought, and that*(R)*-SAM can serve *E. coli* as a methyl donor for conversion of homocysteine to methionine via the enzyme MmuM. We suggest further that the switch from PBP2 elongation to PBP3 division requires the availability of methyl donors.

#### **Involvement of L-Serine**

Among the oddities of L-serine metabolism is that *E. coli* codes for three different very specific high K*<sup>m</sup>* L-serine deaminases (L-SDs) but cannot use L-serine as carbon source. These enzymes, SdaA, SdaB, and TdcG (Su and Newman, 1991; Shao and Newman, 1993; Burman et al., 2004), use a [4Fe-4S] cluster to catalyze the deamination of serine to pyruvate (Cicchillo et al., 2004). They are so highly specific for L-serine that even TdcG, which is coded by a gene in an anaerobic threonine utilization operon, does not deaminate threonine but is a dedicated L-SD (Burman et al., 2004). However, they only work on high concentrations of L-serine, preventing interference with peptidoglycan synthesis (Zhang et al., 2010).

Deleting all three genes from *E. coli* of course greatly increased the availability of L-serine and it also severely impeded the ability of the mutant to grow in minimal medium with amino acids, where the deletion mutant made very large, abnormally shaped cells (Zhang and Newman, 2008). In Luria broth, the triple mutant made long filaments—not as long as strain MNR2 but nonetheless an average of 10*×* normal length and up to 80*×*. Most surprising, even though SAM normally cannot enter the *E. coli* cell, it prevents filament formation in the triple mutant equipped with the Rickettsial transporter (Zhang and Newman, 2008).

We suggest that high serine interferes with the serine/ glycine/C1 balance of the cell and decreases the availability of C1 units and SAM for the cell division specific methylation resulting in production of long cells. On the other hand, overproduction of L-SD also produced long cells. The mutant with the highest serine deaminase activity recorded, the *ssd* mutant now shown to be a subclass of *cpxA* mutants (Rainwater and Silverman, 1990; Monette, 2006), grows in glucose minimal medium expressing high L-SD, with 50% of its cells as small rods and the other 50% as a few enormously long rods. This suggests that every now and then the cell cannot divide and does not recover from this but instead goes on elongating.

Neither of these cases is well understood. However, we suggest that when the balance of serine, glycine and C1 is not maintained, some methylation associated with cell division does not occur.

### **Interference with Cell Division by Provision of Alternative Cell Wall Components**

If the cell is provided with the means to make an entirely different cell wall, e.g., not peptidoglycan, it may go ahead and do it, and then find that it has problems breaking the alternative wall down into smaller cells. Like so many of us, Lederer et al. (2011) used *E. coli* to clone and express proteins of interest to them in *E. coli*. In this case, they cloned S-layer proteins from *Lysinibacillus sphaericus* JG-A12 into *E. coli* BL21(DE3). These are mostly protein and glycoprotein, forming the S-layer envelope of various organisms. *E. coli* can express them, grow well and produce viable cultures. However, its morphology is entirely changed (Lederer et al., 2011).

In early exponential phase these cells form filaments 100 µm long or more. Later these appear as transparent filaments with small *E. coli* rods lined up inside them. For the moment, there is not enough information to analyze these in detail. Nonetheless it is obvious that adding alternative cell wall components to the cell's possibilities has interfered with peptidoglycan synthesis and allowed it to construct an alternative cell envelope of considerable length.

### **Extremely Elongated** *E. coli* **Cells**

#### **Isolation of a Mutant that Does Not Activate PBP3**

*Escherichia coli* divides every 0.5 h or so in Luria broth with or without added sodium chloride (LB, LBNoSalt), and about every 58 min in minimal medium with glucose, all at 37°C. In our attempts to understand the requirement for methionine in the *metK* deletion mutant (see Involvement of Methionine), we have recently isolated a mutant which divides every 80 min in minimal medium but divides rarely in LB and almost not at all in LBNS. That is, it can elongate via PBP2 and divide with PBP3 when growing in minimal medium, but in LB or LBNS it can only use PBP2 and thus continues to elongate indefinitely. The longest cell we have seen so far was is 750 µ long (El-Hajj and Newman, 2015, Figure 1).

When the mutant, known as MNR2, is grown in minimal medium it makes the rods one expects, but they are considerably longer than the parent cell. When the population is subcultured into LBNS, every cell continues to elongate and cannot divide, so that they gradually grow longer over the next 15–24 h. This elongation occurs at many points along the cell (Woldringh et al., 1987). The exact pattern of elongation points is not known (den Blaauwen, 2013). However, even our longest cells show no distortion, so that points of elongation must be chosen according to some pattern. A biophysical simulation by Sean Sun and colleagues suggests that MreB stabilizes the peptidoglycan tube, and provides a framework for the localization of the PBP2 complex and its function (Jiang et al., 2011).

We attributed the elongation in MNR2 to the much lower osmotic pressure of LB and LBNS (El-Hajj and Newman, 2015). This was based on the fact that adding the components of LB to minimal medium produced a medium which had a high osmolality (349 vs. 342 mOSm/kg) and did not prevent division. Also, cells were much longer in LBNS than in LB (93 vs. 256 mOsm/kg). This may indicate that the elongation rate is faster at low osmolality. However, it is also possible that the elongation rate does not change but the probability of transient PBP3 activation is higher at the osmolality of LB.

The size of wild-type *E. coli* is larger the faster it grows (Chien et al., 2012). The larger size is required in order for the cell to contain all the machinery it needs to make cell components, among others, ribosomes to make protein at an increased rate. Thus when *E. coli* is taken from minimal to rich medium, it responds by growing wider and longer (Grover and Woldringh, 2001). Strain MNR2 of course grows longer, but the mechanism for this may be different since its width did not change in any of the thousands of cells we examined.

Cells subcultured in LBNS all elongate for the first few hours, with some extremely elongated (eel) cells reaching lengths up to 200 µm. These cells could be made to grow even longer by plating them on LBNS agar slide cultures, which provides them with fresh nutrients and is how we isolated the longest cell we have seen, 750 µm long. However, when plating these cells on the agar slides and examining them by time-lapse photography, a few produced a new rod at the end of the long cell (El-Hajj and Newman, 2015). These rods are about the size of the usual MNR2 rods in minimal medium, but they immediately begin to elongate again. This predilection for division near the ends is also seen in filaments formed in other ways (Dajkovic et al., 2010, movie S1 early frames). We conclude that PBP3 is normally inactive in the mutants in Luria media, and if it happens to function, it is quickly re-inactivated.

When we instead plated the eel cells on minimal medium slide cultures, we expected that PBP3 would immediately reactivate and they would start dividing promptly. Much to our surprise, the long cells continued elongating at multiple points along the cells, forming loops that pushed outward from the long axis of the cell (El-Hajj and Newman, 2015). This suggests that there are pre-requisites for PBP3 activation even when cells are over the length required to divide. How PBP3 function is activated is currently unknown. However, two separate studies have suggested that peptidoglycan synthesis is altered at the division point even before PBP3 activity begins. One group suggest that there may be PBP3 independent synthesis at midcell of a narrow band of new peptidoglycan, a process involving only two very early septation proteins, FtsZ and ZipA (Potluri et al., 2012). A second group shows that PBP2 and PBP3 interact directly for a short time, and suggests an association between them to synthesize a new area of peptidoglycan prior to the actual septation, so that both enzymes work concurrently for a short time as peptidoglycan synthesis shifts from PBP2 to PBP3 (van der Ploeg et al., 2013). The loops we have seen could be this new peptidoglycan being synthesized. In a small rod-shaped cell, this synthesis would simply push the poles apart and the cell would elongate in a straight line. However, the eel cells are much longer, and the adherence of the cell to the agar on the slide culture would require enough force to push away the entire length of the cell on either side, which could be up to 100 µm of cellular content. Peptidoglycan synthesis does not produce sufficient force to do this, so instead the force pushes in a direction perpendicular to the long axis of the cell and the area where it occurs loops outward.

The cell wall of *E. coli* is usually considered to protect the cell against osmotic pressure by virtue of its rigidity, and this applies to the usual small rods. The peptidoglycan at the poles (which is always the result of a former PBP3-driven lateral peptidoglycan synthesis during division) is thought to be inert. That is, old peptidoglycan is not degraded and replaced by newly-synthesized peptidoglycan (Koch and Woldringh, 1994). This is in contrast to the longitudinal peptidoglycan synthesized by PBP2, which undergoes constant recycling as older peptidoglycan is degraded by and newer peptidoglycan is incorporated in its place (reviewed in Johnson et al., 2013). In a small *E. coli cell* 1 or 2 µ long, the rigidity of the inert polar peptidoglycan maintains the characteristic unbending rod shape. However in the much longer MNR2 eel cells, the rigid poles are frequently separated by over 100 µm and as a result the cells are remarkably flexible. In fact we found cells can be intertwined even by the forces of pipetting a drop of a diluted culture onto a slide! This can be seen in **Figure 1A**, where the free ends of cells are aligned by the flow caused by the falling cover slip. The flexibility can also be seen in a single cell twisting around itself into corkscrew shapes (**Figure 1B**). Long chains of small *Bacillus subtilis* cells form similar corkscrew macrofibers of astounding flexibility, and their elasticity generates enough torque and force to allow self-propulsion when their natural unwinding is hindered (Mendelson et al., 2000). Neil Mendelson recorded a remarkable video of these bacterial macrofibers on his website (www.neilhmendelson.com), which unfortunately seems unavailable at the time of writing.

Whether the high flexibility of the MNR2 eel cells makes it more difficult to produce a crosswall away from the poles, or whether the inability of the cells to form the crosswall in the first place is 10 µm.

what lends them this remarkable flexibility, seems to be a chickenor-egg question. What is clear is that the ends of the cell somehow alleviate this problem, as seen by the transient activation of PBP3 on LBNS slide cultures discussed earlier. At the pole, the cell can take advantage of the rigidity conferred by the end wall, and using that, it requires constriction at only one point one normal cell length away to form a rod of the usual *E. coli* size. This would also allow for the MinCD complex to shuttle within its usual length (Lutkenhaus et al., 2012) and help in accurate placement of the division site, as well as subsequent formation of a functional divisome.

### **Characteristics of Long Cells**

We did not attempt to find conditions which allowed the cells to grow particularly long. The 750 µ cell we photographed (El-Hajj and Newman, 2015) extended over three microscope fields, and was simply the longest we happened to find with our usual growth media. There is no reason to suppose that this is the longest possible, though physical problems may eventually make lengthening more difficult.

It is odd that the diameter of the cell does not increase on subculture into LB or LBNS. One of the earliest findings in studies of *E. coli* adaptation is that wild type *E. coli* transferred from minimal to rich medium, gets both longer and wider (Schaechter et al., 1958). Though there are many circumstances in which *E. coli* can get wider, reviewed in a fascinating review of bacterial shape (Young, 2010), this widening is not seen with this mutant when grown on enriched medium. Strains derived from the same parent do become very much wider when overexposed to L-serine (Zhang et al., 2010) so this does not seem to be simply a strain difference.

The long cells are remarkably flexible, as seen in the corkscrew pictured here (**Figure 1B**). They nonetheless occasionally lyse, as is particularly visible on solid medium. Lysis could be due to adherence to or to other cells pulling them in a different direction from the one in which they elongate. When they lyse, the entire contents pour out leaving an empty tube. In one of our photographs (El-Hajj and Newman, 2015, Figure 2B), the point of lysis fell at a point where the cell formed a closed loop on an agar surface, and one could see the cell contents within that loop. Clearly we could adapt this to collect the entire content of a single cell, perhaps by micromanipulation with a microsyringe.

We assume that as elongation goes on, new DNA is made and transcribed throughout the length of the cell (El-Hajj and Newman, 2015). As an example of "games" which become possible with cells of this size, it might be interesting to add an inducer like X-gal at different times and see if it can induce at all parts of the cell, or to streak inducer on the plate and let cells grow toward it. Similarly we plated cells on slides coated with turmeric powder and found that the cells appeared much thicker due to the negative staining, which would allow us to search for even longer cells at lower magnifications.

What is clear, and surprising, is how well these cells grow and metabolize even after many hours. They do not show signs of metabolic problems and the majority of the long cells can give rise to a colony (El-Hajj and Newman, 2015). This does not inform us as to how many of the DNA molecules in a long cell can give rise to colonies. It does tell us that the long cells remain alive and their metabolism lets them continue to elongate—i.e., their metabolic capacity is remarkable. As long as they are in LB or LBNS, we do not see any localized defect other than occasional lysis. As seen below, this is no longer the case when we transfer the cells to glucose minimal medium where they might be able to subdivide and multiply thereafter as rods.

### **The Return of Elongated Cells to Minimal Medium**

The elongated cells can reach the size of the largest bacteria known (see Giant Bacteria). As long as they are kept in enriched medium, the entire cell appears intact and functional. When they occasionally form rod-shaped cells at the ends of the long cells, these rods maintain the same characteristics, i.e., they elongate once again if left in rich medium. The remaining long cell also continues to elongate. There is no indication that there are nonfunctional areas in these long cells, with the possible exception that DNA does not segregate perfectly in all cells. However, even cells with smears of DNA elongate.

The situation is very different when the elongated cells are plated on minimal medium, where most of the cells give rise to a colony. However, this new colony is not derived from all parts of the elongated cells. Some parts do not grow at all, and some grow and die off, but each long cell has at least one area in which cells grow into a microcolony of rod shaped cells and ultimately into a colony with the same characteristics as MNR2.

We did not follow these in the detail they deserve but intend to do so. When plated on minimal plates, the cells did not immediately produce rods. As seen in **Figure 2**, the eel cell began to elongate, but by 2.5 h incubation, showed the problems we expected but did not see in rich medium. The new lengths of cells could not push the old cell mass to the sides but instead grew outward making irregularly spaced loops. These loops were randomly placed but the same process was seen in all dividing cells.

We have not found other mentions of these loops in *E. coli*. This is the first evidence that the long cells are not entirely homogeneous, and this becomes obvious by 4.5 h when the cells

**FIGURE 2 | Loop formation in the elongated cells after shift down.** MNR2 eel cells from a 16h LBNS culture were plated on minimal medium slide cultures and photographed after 2.5 h. Scale

**FIGURE 3 | Heterogeneity in the MNR2 elongated cells.** A 16 h culture of *E. coli* MNR2 in LBNS plated on a glucose minimal medium slide culture at 37°C was photographed after 6.5 h **(A)**, then left at room temperature for 18 h and the same field was re-photographed **(B)**.

produce a few rods, and break up into segments of various lengths, and some even lyse. An example of this extreme heterogeneity is seen in **Figure 3**. However, the design of this experiment actually promotes heterogeneity. This experiment is indeed a return to conditions which allow division, but it is also a major metabolic shift down. While the cells can suddenly begin to activate PBP3 and divide, they are also suddenly deprived of nutrients and have to begin making all the compounds they need. They can do this at a few points, but not at many. This experiment is worth repeating with a shift from LBNS to LBmin, i.e., restoring high osmolality in the presence of all the nutrients in LB.

### **Filament Formation During Cell Division Arrest**

Although the MNR2 eel cells we saw at low osmolality are the longest *E. coli* observed, conditional elongation in *E. coli* had been observed during the 1960s and 1970s, when it was characterized in mutants that were impaired in cell division.

Cell division requires the assembly of a complex division machinery. The process is initiated when FtsZ, the most well studied of the division proteins, polymerizes into a discrete structure, called the FtsZ ring or Z-ring, but likely much more complex than currently thought (Fu et al., 2010). Assembly of the Z-ring is essential for the recruitment of subsequent members of the division complex, including PBP3 (Errington et al., 2003). In conditions under which cells cannot form the Z-ring, PBP3 is not activated and they cannot form a crosswall, but they do grow into relatively short-lived elongated cells called filaments.

Many factors can disrupt the elongation-division cycle of *E. coli* and lead to formation of filaments. These include alterations in the stability or availability of the essential division proteins, especially FtsZ. Mutations in *ftsZ* or depletion of the protein due to DNA damage and subsequent triggering of the SOS response are both well-studied conditions that cause filamentation (Jones and Holland, 1984; Addinall et al., 1996). These differ from MNR2 eel cells, which are also blocked in cell division and elongate under adverse environmental conditions (in this case, low osmolality), but are viable and become much longer.

### *E. coli* **Filaments in** *fts* **Mutants**

The *fts* (filamentation temperature sensitive) series of mutants were isolated by virtue of their inability to divide when grown on LBNS at 42°C and have allowed the elucidation of many of the steps in Z-ring formation (Lutkenhaus et al., 2012). Such temperature-sensitive (Ts) mutations have been isolated in many of the essential cell division genes, which gave rise to the nomenclature of *fts* for these genes. Most of these mutants share common traits when shifted to 42°C, in that they begin cell division, and reach different stages depending on the mutation and the gene affected, but none of them can complete the process (Taschner et al., 1988). Because FtsZ is the earliest division protein, *ftsZ* mutants abort division very early on, which typically results in socalled "smooth" filaments. Disrupting the later division proteins, such as in *ftsA*, *ftsQ*, or*ftsE* mutants, allows the cells to begin septation. Most such mutants can polymerize FtsZ but cannot assemble the full complex required for successful division, and these cells assemble many partial, usually regularly-spaced Z-rings along the length of the filaments (Addinall et al., 1996). This results in "rough" filaments that show clear, marked constrictions where these incomplete rings form (Taschner et al., 1988, Figure 3). Regardless of what stage is inhibited, the filaments remain viable for only a few hours. The *ftsZ84* mutant, one of the earliest and best studied, lyses and sees a drop in viability 3 h after the temperature shift (Ricard and Hirota, 1973).

Much work was done with *ftsZ84* over the decades, taking advantage of the conditional mutation to study and understand FtsZ function and the essential role the protein plays in cell division (Lutkenhaus et al., 2012). Although most of these studies focused on FtsZ itself and its molecular role in cell division, they did in the process reveal a lot of information about how *E. coli* functions (or does not) when it reaches much longer lengths than usual. Most of the *ftsZ*(Ts) mutants can function at permissive temperatures such as 30°C. Under these conditions, the *ftsZ84* mutant synthesizes DNA and segregates nucleoids regularly (Ma and Margolin, 1999). The cells grow as rods and divide normally, although a significant number of cells (about one-third) lack Zrings entirely (Addinall and Lutkenhaus, 1996). Upon shifting to 42°C, regular nucleoid segregation continues, but now occurs over the entire length of the filament (Bi and Lutkenhaus, 1992; Ma and Margolin, 1999). However, cell division is blocked with remarkable speed. FtsZ rings in an *ftsZ84* mutant seem to disappear as early as 2 min after the temperature shift, even though some of these cells retain a sharply-demarcated septum when viewed by electron microscopy (Addinall et al., 1997, Figure 2). After 10 min, these are replaced by deeper, blunted constrictions, which never complete the process. This is consistent with division sites beginning to form, but due to the mutated FtsZ the cells cannot complete the process at 42°C and septation aborts prematurely, leaving these blunted constrictions to mark the site of their failure.

As *ftsZ84* filaments are left at 42°C, they continue elongating and show no signs of rings (Addinall et al., 1996). If left for several hours they eventually lyse and die (Ricard and Hirota, 1973), but this fate can be avoided by shifting the filaments back to a permissive temperature (such as 30°C) before enough damage has been done. When cells of an *ftsZ84* mutant are upshifted to 42°C for 2 min, then downshifted back to 30°C, they start forming new

rings at unconstricted, future division sites, but they seem unable to reform a ring at a previously-constricted site that was aborted during the upshift (Addinall et al., 1997). The cell "remembers" that it chose a division site and started to constrict, but it either cannot remember that the constriction aborted prematurely, or it cannot promptly resume or restart the constriction, presumably because the topology of an aborted site is different and a partial septum is already there and prevents reassembly of the Z-ring complex. Unfortunately no study followed these filaments for a longer time after the downshift, and whether the cells are eventually capable of fully recovering and re-targeting a previous division site if given enough time remains unknown. The MNR2 eel cells, with their increased viability and lengths, could provide a useful tool to settle this question.

*In vitro* studies of the mutated FtsZ84 protein have revealed defects in its biochemical function even at 30°C, but these do not seem to affect its ability to support cell division under permissive conditions *in vivo*. FtsZ is a GTPase and a tubulin homolog (De Boer et al., 1992; RayChaudhuri and Park, 1992). Like most members of this superfamily, it polymerizes by binding GTP and immediately hydrolyzing it, then depolymerizes when GDP is released (Scheffers and Driessen, 2002). The polymer structure is very dynamic and the turnover of FtsZ is high as individual molecules are cycled in and out of the polymer (Sun and Margolin, 1998). FtsZ84 has greatly reduced GTPase activity as seen by biochemical assays, even at 30°C, yet complete Z-rings are readily seen in *ftsZ84* mutants and the cells can divide (Mukherjee et al., 2001). Likewise, the Z-ring of an *ftsZ84* mutant has a much slower turnover rate than wild-type, but does not affect its doubling time at 30°C (Stricker et al., 2002). The biochemical defects of the mutant protein, which are measurable under all conditions, only manifest in the cell itself under specific environmental conditions. Unfortunately the exact players that compensate for these defects at 30°C yet allow the formation of the elongated filaments at 42°C remain unknown.

However, a couple of factors that can affect filamentation in *ftsZ84* have been well characterized. The first is that mild overexpression (twofold to threefold) of *ftsZ84* can suppress filamentation and allow *ftsZ84* mutants to divide at 42°C, which indicates that an increase in the level of the mutated protein can compensate for its functional deficit (Phoenix and Drapeau, 1988; Lu et al., 2001). The second is that many of the *fts*(Ts) mutations do not filament when grown in medium of higher osmolality. The *ftsZ84* mutant filaments in LBNS, but when grown on LB (i.e., with sodium chloride) and at 42°C, *ftsZ84* cells divide, are alive and form colonies (Ricard and Hirota, 1973). The inhibition of cell division in the MNR2 eel cells is similarly dependant on low osmolality, though unlike the *fts*(Ts) mutants they elongate without needing a temperature shift (El-Hajj and Newman, 2015). Not all the temperature-sensitive mutants are affected by osmolality or salt; for example *ftsZ26* filament at 42°C in both LB and LBNS (Bi and Lutkenhaus, 1992).

Why an increase in osmotic pressure restores division in some mutants but not others is not fully understood. In the MNR2 eel cells, elongation correlates with a decrease in the cellular levels of FtsZ, which we have suggested is the reason these cells cannot form a functional Z-ring (El-Hajj and Newman, 2015), but no such correlation has been drawn in any of the *fts*(Ts) mutants that we know of. On the contrary, in a number of *ftsZ*(Ts) mutants (including *ftsZ84*) grown in LBNS, the levels of FtsZ were *higher* at 42°C, when the cells filament, than they were at 30°C where these mutants could divide as the usual short rods (Addinall et al., 2005). *E. coli* grows within a large range of osmotic pressure but functions optimally between 300 to 500 mOsm (Cayley and Record, 2004). A change in osmotic pressure affects the cell at just about every level, from the composition of the lipid bilayer to proteinnucleic acid interactions and most enzymatic reactions (Kornblatt and Kornblatt, 2002). Low osmolality affects MNR2 and *ftsZ84* similarly, in that they both elongate, but the precise mechanism disrupted by this change in osmotic pressure is different and the cells become either filaments destined to die or eel cells that can keep elongating.

### **Filament Formation During the SOS Response**

As we mentioned in the introduction to this review, one of the remarkable features of *E. coli* (as well as many other bacteria) is that its metabolism is very efficient. New cells are produced rapidly with no wasting of metabolites and with very few mutants. The SOS response is a general stress response that many bacteria undergo when exposed to environmental or chemical factors that lead to DNA damage, such as radiation or some antibiotics (Walker, 1996). It involves a global and comprehensive regulatory network aimed at temporarily halting replication and division while repairing the DNA lesions, which ensures that the mutations that led to triggering the response do not get passed on to daughter cells.

The mechanism underlying induction of the SOS response is well described (Erill et al., 2007). Most of the genes involved in triggering the response are regulated by the LexA repressor, which is normally bound to their promoter region and blocks their expression (d'Ari, 1985). Conditions that induce DNA damage (such as UV irradiation) lead to the formation of lesions in the DNA. During DNA replication, the DNA polymerase complex inevitably encounters these lesions and cannot get past them, which stalls the replication fork. This leads to the recruitment of RecA to the site of the lesion and its activation when it binds to the single-strand DNA generated by the interruption in replication (Sassanfar and Roberts, 1990). Activated RecA can promote the autolytic cleave of LexA (Horii et al., 1981), which dissociates it from the promoter region of the SOS genes and enables their active transcription. A host of genes are then expressed (Janion, 2008), many of which are involved in stabilizing the stalled replication fork or in repairing the lesion (Baharoglu and Mazel, 2014). In order to buy time for repairing the DNA damage, the cell delays septation through the SOS-induced cell division inhibitor SulA (Huisman et al., 1984).

SulA blocks cell division during the SOS response by interacting directly with FtsZ in a 1:1 (Higashitani et al., 1995). This forms a stable complex that prevents FtsZ subunits from polymerizing and effectively inhibits septation at its earliest stage (Trusca et al., 1998; Cordell et al., 2003). SulA inhibition of FtsZ polymerization is reversible (Maguin et al., 1986), which allows the cell to exit the SOS phase and resume normal division once the DNA damage has been dealt with. This is controlled by the Lon protease, which plays a role in the degradation of SulA, freeing the FtsZ monomers to polymerize and initiate a new division cycle (Schoemaker et al., 1984).

If Lon function is inhibited, SulA is no longer degraded and can continue to accumulate and to sequester FtsZ. This extends the division block of the SOS response, and as the cells continue to elongate they form aseptate, multinucleate filaments (Adler and Hardigree, 1965). These filaments are not viable and the cell division block eventually becomes irreversible; within 4 or 5 h of their formation, only 2% of the filaments can recover and form colonies (Adler and Hardigree, 1964). However, the block can be overcome by reactivating Lon in the filaments (Walker and Pardee, 1967), since SulA would then be degraded, freeing FtsZ to polymerize and reinitiate cell division (Schoemaker et al., 1984).

Early during their characterization, SOS-induced filaments were noted for being metabolically competent despite being inhibited in cell division. They continue to elongate, increase mass, and synthesize nucleic acids and proteins (Walker and Pardee, 1967). Despite triggering the SOS state in response to DNA damage, they replicate their chromosomes and distribute DNA continuously and regularly throughout their length (Adler and Hardigree, 1965). Why then do they lyse and die within a few hours, while the metabolically competent MNR2 eel cells survive and continue to elongate well past this time? The SOS response triggers the expression of diverse genes, many of which are not necessarily involved in cell division but in regulating other aspects of *E. coli* metabolism and physiology (Janion, 2008). This probably affects the filaments in ways that were not measured or observed but that disrupt their metabolism and eventually leads to their death. This would be advantageous from an evolutionary perspective—a filament where the SOS response stays induced for so long is one that cannot repair the DNA lesions that triggered it in the first place and its death ensures the mutations do not get propagated to daughter cells. One feature they still share with the MNR2 eel cells is that in both cases elongation is initiated by a decrease in the availability of FtsZ.

The cellular concentration of FtsZ is constant throughout the division cycle, and is set at a critical concentration below which division is hindered (Rueda et al., 2003; Weart and Levin, 2003). Division begins not by synthesizing more FtsZ but by assembling the heretofore soluble FtsZ into the Z-ring (Sun and Margolin, 1998), with eventual activation of PBP3. A decrease in FtsZ can prevent PBP3 activation and onset of division and lead to the formation of elongated cells, as seen when cells are partially or completely depleted of FtsZ (Dai and Lutkenhaus, 1991). In a typical *ftsZ*(Ts) mutant such as *ftsZ84*, the cells form filaments with no constrictions after transfer from 30°to 42°C (Addinall et al., 1996), where FtsZ84 function is inhibited. The filaments produce increased levels of FtsZ84, perhaps in an attempt to compensate for the inhibition in the activity of the mutant protein (Addinall et al., 2005). During the SOS response, the FtsZ molecule itself is intact, but its sequestration by SulA decreases the pool of available, active FtsZ, which also leads to filamentation (Lutkenhaus, 1983; Chen et al., 2012). A similar mechanism is used by OpgH, a moonlighting enzyme that acts as a nutrient-dependant regulator of cell size by sequestering FtsZ under nutrient-rich conditions (Hill et al., 2013). In the case of the MNR2 eel cells, FtsZ activity is reduced because the levels of the proteins are much lower in medium of low osmolality, presumably lower than the critical concentration required for division (El-Hajj and Newman, 2015). A reduction in FtsZ function is therefore clearly correlated with division inhibition and elongation, but the differences underlying the exact mechanisms in each case also lead to different fates for the elongated cells.

## **Giant Bacteria**

Although MNR2 grows as long as any bacterium when it is cultured in low-osmolality LBNS, it has no way of dividing in this medium. This distinguishes the *E. coli* long cells from naturallyoccuring giant bacteria, which can divide despite reaching similar lengths. The giant bacteria have evolved complex mechanisms to survive and take advantage of this enormous size, which is indeed the norm for them.

When we think of bacteria, we tend to think of unicellular microorganisms 1–5 µm in length or diameter. Many of the "traditional" bacteria fall within this length, such as *E. coli*, *Salmonella*, *Vibrio, Bacillus*, and *Staphylococcus*. However, bacterial sizes encompass a much wider range. *Mycoplasma pneumoniae*, one of the smallest known bacteria, is only 0.2 µm in diameter. These cells are so small that, over the course of its evolution, the bacterium drastically reduced the size of its genome by loss of most of its metabolic pathways, locking it into an obligate parasitic lifestyle for the acquisition of most metabolites (Himmelreich et al., 1996). At the opposite end of the spectrum, *Epulopiscium fishelsoni* cells can reach lengths of 700 µm and widths of 80 µm (Angert et al., 1993). The largest bacterium known, *Thiomargarita namibiensis*, is a giant sulfur bacterium that forms linear chains of spherical-shaped cells with diameters of up to 750 µm, so large that individual cells can be seen without any optical enhancements (Schulz et al., 1999).

### *Thiomargarita* **and Giant Sulfur Bacteria**

Giant bacteria need to deal with very different surface/volume considerations and have evolved elaborate transport and division systems in order to survive at sizes much larger than the more typical bacteria. Koch attempted to estimate the maximum size a cell could be if it relied exclusively on diffusion, and calculated this maximum as 300 µm (Koch, 1996). The diameter of *T. namibiensis* cocci frequently exceeds this and can reach values as high as 750 µm. They overcomes transport problems by organizing their cytoplasm around a large storage vacuole that takes up almost 98% of the cell's volume (Schulz et al., 1999). The actual cytoplasm is a thin spherical shell that wraps around this liquid vacuole and contains a large number of sulfur globules. This arrangement allows all points of the cytoplasm close proximity to both the extracellular space and the vacuole, which overcomes the diffusion size limit. Another giant sulfur bacterium was isolated in the Gulf of Mexico, with large spherical cells up to 375 µm in diameter. This *Thiomargarita*-like bacterium had a similar cellular structure consisting of a central storage vacuole surrounded by a spherical shell of cytoplasm, which also contained sulfur globules (Kalanetra et al., 2005).

Why do the *Thiomargarita* bacteria organize their cytoplasm in such a way, and why do they need such a large central vacuole? A very long cell with a small diameter, such as the MNR2 eel cells, can overcome surface/volume limitations while maintaining a much smaller biovolume. The central vacuole seems to be a feature of giant sulfur bacteria and had already been described in *Beggiatoa* (Nelson et al., 1989). *Beggiatoa* is motile and usually uses oxygen as an electron acceptor in its sulfur oxidation, but it can store nitrate in the vacuole and use it as a temporary alternative electron acceptor when oxygen is low and it needs to move to another, more oxygen-rich environment (Dunker et al., 2011). Both of the identified *Thiomargarita* bacteria are non-motile and their vacuoles are much larger than in *Beggiatoa*. They may have evolved these giant vacuoles as nitrate storage tanks to survive long periods of anoxia and of low environmental nitrate. Schulz and Jorgensen (2001) calculated that the vacuole of *T. namibiensis* could store enough nitrate for a cell to survive almost 2 months without an external supply of either oxygen or nitrate, but under laboratory conditions a cell can survive for well over a year.

### **Epulopiscium fishelsoni**

Before the identification of *Thiomargarita*, the previous contender for largest bacterium was *Epulopiscium fishelsoni*, a symbiotic microorganism that spends the entirety of its life cycle within the gut of the surgeonfish (Fishelson et al., 1985). Reaching a length up to 700 µm and a width of 80 µm, the cell encloses a large volume of continuous cytoplasm, resulting in transport problems from outside and within the cell (Angert et al., 1993). *Epulopiscium fishelsoni* overcomes surface/volume limitations using extreme polyploidy; each cell has between 50,000 and 120,000 copies of the chromosome, with the exact number being directly proportional to the size of the cell (Mendell et al., 2008). As *Epulopiscium fishelsoni* cells get larger they replicate their chromosome to such high numbers to allow extremely high expression of transport pumps, as well as ribosomes, many of which are concentrated toward the center of the cell, away from the cell membrane. This is coupled to a large number of infoldings of the cell membrane, which is thought to enhance transport across the membrane and further into the center of the cell (Clements and Bullivant, 1991). Transport pumps on the cell membrane are similar to those in other bacteria, but they are much more kinetically active and have unusually high transport rate (Bresler and Fishelson, 2006), presumably to compensate for *Epulopiscium fishelsoni*'s much lower surface/volume ratio.

*Epulopiscium* does not divide like the typical *E. coli* using FtsZ and PBP3 as we described earlier. Instead, two daughter cells form inside the parent cell, each with its own cell membrane, and both completely enclosed within the cytoplasm of the parent (Angert, 2012). FtsZ does polymerize into a ring structure, not at midcell, but at the poles of the daughter cells instead (Angert and Clements, 2004). Eventually the daughter cells take over most of the cytoplasm of the parent and burst out, killing the parent in the process (Montgomery and Pollak, 1988). The growth of these daughter cells inside the parental cell shares some similarities with endospore formation in bacteria such as *Bacillus* (Errington, 1993), with one major difference: *Epulopiscium fishelsoni* daughter cells are not quiescent but fully active. This makes *Epulopiscium* and its relatives unique in that they rely on viviparous reproduction with the formation of multiple internal offsprings, which are released by destroying the parental cell.

### **Desulfobulbaceae and Cable Filaments**

Recently, a relative of *Desulfobulbus* was identified that blurs the line of what defines an individual bacterial cell. These Desulfobulbaceae form extremely long cable-like filaments in marine sediments, up to 1.5 cm long, twice that of the longest MNR2 eel cell (Pfeffer et al., 2012). Although an individual eel cell has no compartmentalization and a continuous cytoplasm, the Desulfobulbaceae filament consists of a linear chain of hundreds of cells, each only 3 µm long and each with its own inner membrane. However, all the cells within the filament are encased within the same outer membrane, which spans the entire 1.5 cm of length uninterrupted. Whether each filament is a single cell or a colony of hundreds of cells depends on whether we define the inner membrane or the outer membrane as enclosing a single cell. The cytoplasm of each short cell within the filament is unconnected to that of the adjacent cells, but each cell is connected to its neighbors by a filling that bridges the gap (Pfeffer et al., 2012). This organization allows high electron conductivity, which gives the Desulfobulbaceae filament the remarkable property of acting like an electric cable capable of conducting current across the layers of sediment where it normally grows. When followed over several weeks, the filaments grow and expand deeper into the sediments (Larsen et al., 2014; Schauer et al., 2014). The remarkable length of these filaments is thought to help them couple oxygen consumption, which requires contact with surface sediments, and the sulfide oxidation that occurs in deeper, anoxic layers.

### **Concluding Remarks**

*Escherichia coli* normally exists as a small rod, but we have discussed many conditions that cause it to elongate when cell division is inhibited. Some of these are related to lowered activity of FtsZ,

### **References**

such as in *ftsZ*(Ts) mutants, or when FtsZ is sequestered by SulA during the SOS response. Many metabolic defects also interfere with cell division, including metabolism of amino acids such as serine and methionine, as well as disruptions SAM synthesis, which is required for most methylations. In the MNR2 eel cells, disruption of SAM metabolism linked to one or more unknown mutations has resulted in the longest *E. coli* cells observed. This might be due to the methylation of a key cell division activator being disrupted in this mutant.

We conclude from these and other examples that as long as it remains narrow, allowing efficient uptake and excretion of material, *E. coli* can arrange its metabolism to function and grow over large lengths of undivided protoplasm. It can place its DNA and ribosomes so as to produce mRNA and proteins, and organize its enzymes spatially to function efficiently. In these ways it is as efficient as the giant bacteria like *Thiomargarita*, which have evolved as very large cells and have specific mechanisms to live at these sizes. *E. coli* lacks the specific adaptation to divide but has no trouble surviving at such sizes.

What an ultra elongated *E. coli* cannot do that *Thiomargarita* can is divide! We suggest that long length presents no particular metabolic problems by itself. However, the lack of an efficient division system for its long variants keeps *E. coli* short.

### **Acknowledgments**

We would like to thank Giri Narasinham for teaching us to analyze whole genome sequence data. ZE is grateful to Pat Gulick and Justin Powlowski for their financial support at Concordia University. EN wishes to thank Kalai Mathee for transferring the Newman lab from the downtown campus to the west end and organizing the new lab, Noel Guyves, Dean Solin, Jean Rosa, Marianne Chancerel, Jack Kornblatt, and Conrad Woldringh. ZE dedicates this review toWalid, Carmen, and Shadi El-Hajj for their continued support and love. EN dedicates this review to Harold Newman and Vijay Mathur without whom, nothing.

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Copyright © 2015 El-Hajj and Newman. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

# **The stoichiometric divisome: a hypothesis**

### *Alexander J. F. Egan\* and Waldemar Vollmer\**

*The Centre for Bacterial Cell Biology, Institute for Cell and Molecular Biosciences, Newcastle University, Newcastle upon Tyne, UK*

Dividing *Escherichia coli* cells simultaneously constrict the inner membrane, peptidoglycan layer, and outer membrane to synthesize the new poles of the daughter cells. For this, more than 30 proteins localize to mid-cell where they form a large, ring-like assembly, the divisome, facilitating division. Although the precise function of most divisome proteins is unknown, it became apparent in recent years that dynamic protein–protein interactions are essential for divisome assembly and function. However, little is known about the nature of the interactions involved and the stoichiometry of the proteins within the divisome. A recent study (Li et al., 2014) used ribosome profiling to measure the absolute protein synthesis rates in *E. coli*. Interestingly, they observed that most proteins which participate in known multiprotein complexes are synthesized proportional to their stoichiometry. Based on this principle we present a hypothesis for the stoichiometry of the core of the divisome, taking into account known protein–protein interactions. From this hypothesis we infer a possible mechanism for peptidoglycan synthesis during division.

#### *Edited by:*

*Arieh Zaritsky, Ben-Gurion University of the Negev, Israel*

#### *Reviewed by:*

*William Doerrler, Louisiana State University, USA Miguel A. De Pedro, Consejo Superior de Investigaciones Científicas, Spain Manjula Reddy, Centre for Cellular and Molecular Biology, India*

#### *\*Correspondence:*

*Alexander J. F. Egan and Waldemar Vollmer, The Centre for Bacterial Cell Biology, Institute for Cell and Molecular Biosciences, Newcastle University, Richardson Road, Newcastle upon Tyne NE2 4AX, UK<EMAIL_ADDRESS><EMAIL_ADDRESS>
#### *Specialty section:*

*This article was submitted to Microbial Physiology and Metabolism, a section of the journal Frontiers in Microbiology*

> *Received: 25 March 2015 Accepted: 27 April 2015 Published: 12 May 2015*

#### *Citation:*

*Egan AJF and Vollmer W (2015) The stoichiometric divisome: a hypothesis. Front. Microbiol. 6:455. doi: 10.3389/fmicb.2015.00455* **Keywords: bacterial cell division, peptidoglycan, peptidoglycan synthesis, divisome and multiprotein complex**

### **Introduction**

The peptidoglycan cell wall is an essential component of most bacteria, required for the maintenance of cell morphology and structural integrity (Weidel and Pelzer, 1964). Peptidoglycan (PG) forms a net-like, continuous layer, called the sacculus, surrounding the cytoplasmic membrane. Thus, a bacterial cell needs to increase the surface of its sacculus in order to grow and divide. Sacculus growth has to be well-controlled, because the accumulation of defects may lead to cell lysis; this is the case when antibiotics like β-lactams inhibit peptidoglycan synthesis. Remarkably, the growth of the single-layered sacculus in *E. coli* is accompanied by the release of as much as 50% of the total peptidoglycan material per generation by hydrolases (Goodell, 1985). What are the molecular mechanisms of sacculus growth? Höltje (1998) proposed that multi-enzyme complexes made of peptidoglycan synthases and hydrolases simultaneously synthesize new peptidoglycan and incorporate it into the sacculus, and remove old material, by a so-called 3-for-1 mechanism. Subsequent experimental evidence supported the multi-enzyme complex hypothesis. Genetic and biochemical data have demonstrated many protein–protein interactions between various peptidoglycan synthases, hydrolases, regulatory proteins, and cytoskeletal elements (reviewed in Typas et al., 2012; Egan and Vollmer, 2013). These have been observed in several species including, but not limited to, the Grampositive *Bacillus subtilis*, *Staphylococcus aureus* and *Streptococcus pneumoniae* and the Gram-negative *Caulobacter crescentus* and *E. coli*, which will be the focus of this article.

In *E. coli*, cell division involves over 30 proteins, with twelve of these (the Fts proteins and ZipA) absolutely required for the process (Margolin, 2005). The assembly of essential proteins occurs in two steps; FtsZ, FtsA, ZipA, Zap proteins (A–E) and FtsEX assemble early at the future division

**TABLE 1 | Synthesis rates of the late division proteins (and FtsA) and their ratios relative to PBP3 in MOPS minimal and complete media according to Li et al. (2014).**

*<sup>a</sup>We have assumed that previously observed homodimerizations (e.g., PBP3, PBP1B) occur within the divisome.*

*<sup>b</sup>Despite the apparent excess of LpoB, we have assumed two molecules per complex given its stoichiometry with PBP1B is 1:1 (Egan et al., 2014).*

site, before any constriction is visible. Immediately before the onset of constriction the divisome matures through the incorporation of FtsK, FtsQ, FtsL, FtsB, FtsW, PBP3 (FtsI), and FtsN (Aarsman et al., 2005). By a largely unknown process these proteins, along with other accessory proteins, then facilitate the synthesis of the new cell poles of each daughter cell (reviewed in Typas et al., 2012; Egan and Vollmer, 2013). Although different methodologies have identified a large number of interactions between divisome proteins (Egan and Vollmer, 2013), for most of these the precise interaction sites are not known. Moreover, while the cellular copy number of some but not all divisome proteins have been reported over the years, the stoichiometry of proteins within the divisome is not known.

A recent study by Li et al. (2014) used ribosome profiling/ footprinting to evaluate the genome-wide absolute protein synthesis rates and protein copy numbers in *E. coli*. Interestingly, they observed proportional synthesis of proteins present in multiprotein complexes. Proteins of 59 out of 64 cytosolic and membrane complexes (92%) with known stoichiometry were found to be synthesized proportionally to their stoichiometry. Based on this principle of proportional synthesis of proteins participating in complexes, and using the protein synthesis rates from Li et al. (2014), we suggest a hypothetical model for the core divisome complex in *E. coli* factoring in known protein–protein interactions. From this hypothetical stoichiometry model we suggest possible aspects of the mechanisms for PG synthesis during division.

## **Using the Ratios of Division Proteins to Suggest a Stoichiometry for the Complex**

Our aim here was to model the complex formed by the late division proteins and FtsA, without FtsZ and other cytoplasmic or accessory components. PBP3 is the monofunctional peptidoglycan transpeptidase (TPase) essential for cell division in *E. coli* (Weiss et al., 1997), and there is no evidence that PBP3 is active elsewhere in the cell other than the division site. We therefore surmised PBP3 is a reasonable choice for use as the reference point for our considerations on divisome stoichiometry. To this end we calculated the ratios of absolute synthesis rates of each of the *E. coli* cell division proteins (FtsA, FtsK, FtsQ, FtsL, FtsB, PBP3 (*ftsI*), FtsW, FtsN, and PBP1B; Li et al., 2014) to the synthesis rate of PBP3 for cells grown in both minimal and complete media (MOPS media with either full or minimal supplement; **Table 1**). The ratios are similar at both conditions except for PBP1B and its interacting protein FtsN (Müller et al., 2007) which are both slightly more abundant in complete media for unknown reason. This increased abundance may reflect the fact that PBP1B is able to function outside of the divisome, presumably in cell elongation. PBP1B has been shown to be functionally redundant with the other major PG synthase of *E. coli*, PBP1A (Yousif et al., 1985; Denome et al., 1999). FtsN interacts with PBP1B in non-dividing cells (Müller et al., 2007) and hence might be associated with PBP1B at all times. An apparent disparity in synthesis rate was seen with the OM lipoprotein regulator of PBP1B, LpoB, which is in sixfold or fourfold excess over its cognate synthase in minimal and compete media, respectively. It was previously shown that PBP1B and LpoB proteins interact with a 1:1 stoichiometry (Egan et al., 2014). The reason for the excess of LpoB is unknown, but may be of regulatory consequence. LpoB is absolutely essential for PBP1B function in the cell (Paradis-Bleau et al., 2010; Typas et al., 2010), therefore, an excess of LpoB may increase the likelihood that PBP1B can be activated. However, we cannot exclude other mechanisms including, for example, enhanced turnover of LpoB in the cell, necessitating higher synthesis.

PBP3 interacts with PBP1B and both form, and are likely functional as, homodimers (Zijderveld et al., 1991; Bertsche et al., 2005, 2006; Sauvage et al., 2014). We therefore assume that there are two molecules of PBP3 and two molecules of PBP1B present in a functional peptidoglycan synthesis unit (or complex). Interestingly, most other late division proteins (except FtsK, FtsL, and FtsN) had ratios of approximately 1:1 with PBP3 and PBP1B. FtsL, whose ratio to PBP3 is 1.4:1, is part of the FtsQLB complex within the divisome (Buddelmeijer and Beckwith, 2004). FtsL is unstable in the absence of FtsB in both *B. subtilis* and *E. coli* (Daniel and Errington, 2000; Buddelmeijer et al., 2002), and FtsQ, FtsL, and FtsB appear to interact in equimolar stoichiometry; different models suggest a 1:1:1 or a 2:2:2 complex (Masson et al., 2009; Villanelo et al., 2011). We therefore assume two molecules FtsL per complex, equivalent to FtsQ and FtsB.

FtsN has been previously shown to self-associate (Di Lallo et al., 1999), but there are no published data on the stoichiometry of its interactions with PBP1B and PBP3 (Wissel and Weiss, 2004; Müller et al., 2007). Given its ratio of 2:1 to PBP3 (and PBP1B) FtsN may exist in the divisome as a tetramer, or as two separate dimers. FtsN is the last essential protein recruited to mid-cell, and its absence leads to the delocalization of the already assembled divisome components (Rico et al., 2010). Recent evidence suggests that FtsN is the ultimate regulator of septal PG synthesis. In addition to its interactions with PBP3 and PBP1B, FtsN interacts directly with the early divisome protein FtsA (Busiek et al., 2012). This interaction is thought to provide the final signal for the beginning of constrictive PG synthesis through both FtsA and the FtsQLB complex. It was recently shown that point mutations in *ftsA*, *ftsL*, and *ftsB* can bypass the need for FtsN and the altered proteins acted synergistically to restore cell division in the absence of FtsN, suggesting that FtsN signals for constriction via these proteins (Liu et al., 2015; Weiss, 2015). Of note, the copy number of *∼*4500 FtsN molecules per cell determined previously by immunodetection (Ursinus et al., 2004) is significantly higher than the 260 (MOPS minimal)/870 (MOPS complete) molecules per cell determined by ribosome profiling (Li et al., 2014), suggesting that FtsN numbers might vary with strain and growth conditions, which were different in both studies.

FtsK is the first of the "late" divisome proteins, bridging the PG synthetic and cytoskeletal parts of the complex through interactions with FtsA and FtsQ (Chen and Beckwith, 2001; Buddelmeijer and Beckwith, 2004; Aarsman et al., 2005). FtsK has two domains, an N-terminal domain anchored in the inner membrane (IM) which is essential for cell division (Grenga et al., 2008; Dubarry et al., 2010) and a cytoplasmic C-terminal domain which hexamerizes to form a directional DNA pump to resolve chromosome dimers, which is essential in cells with catenated sister chromosomes (Aussel et al., 2002). These domains are linked by a 600 residue long flexible region (Löwe et al., 2008). A specific mutation in *ftsA* or overproduction of *ftsQ, ftsA*, and *ftsZ* can partially compensate for the loss of *ftsK*, suggesting that the essential role of FtsK in cell division may be ensuring divisome stability or spatial regulation (Geissler and Margolin, 2005). The N-terminal domain of FtsK was shown to form hexamers independently of the C-terminal domain and exist as such at midcell (Bisicchia et al., 2013). The mid-cell of predivisional or dividing cells contained 1–8 (average of 7) hexamers of FtsK (Bisicchia et al., 2013). The ratio of FtsK to PBP3 is 1.5:1, suggesting that there are three molecules of FtsK for every two of PBP3 and thus three FtsK molecules per complex. Therefore, the FtsK homohexamer appears to interact with two complexes. We therefore suggest that FtsK spatially co-ordinates two of the peptidoglycan synthesis complexes, which we term "synthesis nodes," with the FtsZ-FtsA cytoskeletal structures in the cytoplasm.

**Figure 1A** shows our hypothetical model for core divisome stoichiometry based on the determined protein ratios. The protein ratios suggest that there are two peptidoglycan synthesis nodes associated with an FtsK hexamer, when three may well be able to associate. The complex is likely highly dynamic, coalescing and dissociating repeatedly in the cell for separate rounds of PG synthesis. Remarkably, such a complex has a total of 42 membrane proteins together containing as many as 92 transmembrane helices. Hence, the impact of these on membrane properties is likely to be profound. Moreover, the divisome complex is likely to be even larger because we have not included accessory divisome components such as FtsEX, FtsP, ZipA or the peptidoglycan hydrolases.

### **Constrictive PG Synthesis by the Divisome**