The dimorphic alphaproteobacterium *Caulobacter crescentus* represents an excellent case study in understanding how new morphologies arise from the diversification of the common core peptidoglycan synthesis machinery. *C. crescentus* divides asymmetrically, producing a swarmer cell with a polar flagellum and a DNA replication-competent cell with a polar stalk. This process of division requires strict coordination in time and space with other cell cycle events such as cell growth, chromosome segregation, and differentiation. Therefore it is unsurprising that although *C. crescentus* maintains similar core peptidoglycan synthesis machinery components in its divisome and elongasome, it has evolved discrete scaffolds, regulatory proteins, and cell cycle control mechanisms to adapt the machinery to its specific lifestyle. Determining exactly how *C. crescentus* has repurposed conserved peptidoglycan synthesis components for elongation, division, and stalk synthesis will help to expand our understanding of bacterial morphology.

The diversification of proteins interacting with and regulating FtsZ and MreB allows for strict control of asymmetric division and morphological development in *C. crescentus*. One significant difference between *C. crescentus* and the canonical rod-shaped bacteria *E. coli* and *B. subtilis* is that *C. crescentus* does not utilize a Min system or nucleoid occlusion proteins. Instead, *C. crescentus* uses MipZ gradients to position the FtsZ ring at the division plane (Thanbichler and Shapiro, 2006). MipZ directly interacts with FtsZ to inhibit ring polymerization. When MipZ associates with ParB at the stalk pole prior to S phase, it drives FtsZ monomers to the new, non-stalked pole (**Figure 4B**). During initiation of chromosome replication and origin duplication, some of the MipZ-ParB complex binds the new origin and migrates with it to the new pole, displacing FtsZ from the new pole to the division plane. Although MipZ shares some distant domain similarity with MinD, it likely evolved from ParAlike DNA partitioning proteins and is conserved amongst all Alphaproteobacteria without MinCD orthologues (Thanbichler and Shapiro, 2006).

Other polar regulators have been shown to influence cell morphology and interact with peptidoglycan synthesis machinery (**Figure 4B**). TipN, a polytopic membrane protein with a large coiled-coil cytoplasmic domain, ensures transmission of cell polarity to the daughter cell by marking the new pole and regulating MreB dynamics at the division plane (Lam et al., 2006). PopZ forms oligomeric networks at the poles to form polar ribosome exclusion zones and serves two roles during the cell cycle, switching function during the swarmer to stalk transition: it first interacts with the parS/ParB centromere at the stalked pole prior to the initiation of DNA replication, and later, during polar maturation, it recruits remodeling factors, such as peptidoglycan synthesis machinery, to transform the flagellated pole into a stalked pole (Bowman et al., 2010; Laloux and Jacobs-Wagner, 2013; Ptacin et al., 2014). All of these regulatory polar protein systems serve as examples of alternative networks that regulate the otherwise conserved core peptidoglycan synthesis machinery.

Another difference between *C. crescentus* and canonical rod-shaped bacteria is that it exhibits at least four peptidoglycan synthesis modes over its cell cycle: lateral elongation, elongation at the division plane before division (also called "pre-septal" or "medial" elongation, **Figure 2F**), septation, and stalk synthesis/elongation (**Figure 4A**). As in canonical rod-shaped cells, cell elongation in *C. crescentus* is MreB-dependent. Early studies showed that depletion of MreB disrupts cell morphology and results in lemon-shaped cells (Figge et al., 2004; Gitai et al., 2004). However, the role of MreB has expanded in *C. crescentus* to accommodate the cell cycle and additional growth modes. For example, MreB appears to influence cell polarity and division, as its depletion disturbs the localization of origins of replication as well as the localization of polar regulatory proteins PleC, DivJ, CckA, and DivK (Gitai et al., 2004). Moreover, MreB exhibits dynamic localization throughout the cell cycle, cycling between dispersed helical patterns in swarmer cells (similar to that seen in *E. coli* and *B. subtilis*) and an intense band at the division plane in pre-divisional cells (**Figure 4B**).

The dynamic localization of MreB drives a shift in growth mode from dispersed, lateral elongation mediated by MreB to zonal, medial elongation mediated by both MreB and FtsZ. The condensation of MreB at the division plane requires assembly of the FtsZ ring (Figge et al., 2004; Aaron et al., 2007), and high-resolution temporal studies have shown that MreB recruitment to the FtsZ ring coincides with medial elongation (Goley et al., 2011). *E. coli* also exhibits a transient pre-septal elongation mode (de Pedro et al., 1997; Aarsman et al., 2005; Potluri et al., 2012), in which FtsZ and MreB interact directly to transfer peptidoglycan synthesis machinery from the elongasome to the divisome (Fenton and Gerdes, 2013). However, in *C. crescentus,* this growth mode predominates a portion of the cell cycle. Because the FtsZ quickly reassembles after division in stalked cells, stalked cells likely elongate primarily via medial elongation (Aaron et al., 2007). Therefore, a similar medial elongation growth mode has been extended in *C. crescentus* to accommodate the asymmetric cell cycle.

MreB appears to have species-specific functions in *C. crescentus*. For example, it appears to interact directly or indirectly with CreS, and is required for proper attachment of the CreS filament to the membrane (Ausmees et al., 2003; Cabeen et al., 2009). Mutations along one surface of MreB alter cell curvature and disrupt the association of the CreS filament with the membrane (Charbon et al., 2009; Dye et al., 2011). Therefore MreB may organize CreS during cell growth, facilitating the attachment of the filament in a stretched conformation (Charbon et al., 2009). Another set of MreB mutations, which occur within the nucleotide-binding pocket, result in morphological defects and affect the localization pattern and dynamics of MreB (Dye et al., 2011). The nucleotide binding site mutants suggest that the ATP hydrolysis cycle of MreB could be coupled to the *C. crescentus* cell cycle to spatiotemporally regulate specific growth modes. The regulation of MreB by ATP could drive its relocation from the sides of the elongating cell to the division plane during the cell cycle.

Some MreB mutants have interesting implications for understanding how *C. crescentus* may remodel polar peptidoglycan. For a subset of the nucleotide binding site MreB mutants (E213G, D16G, N21D, and A325P), cells exhibit a variable width phenotype in which the ends of the cells become extremely tapered and pointed (Dye et al., 2011; Harris et al., 2014). In these mutants, although MreB still exhibits wild type function, it localizes to the cell poles instead of dispersing or condensing at the division plane (Harris et al., 2014). This behavior presumably drives aberrant peptidoglycan synthesis to create elongated cell poles. These MreB mutants may be trapped in a phase of the ATP cycle in which MreB localizes to the poles for a specific polar growth mode in wild type *C. crescentus* cells (Dye et al., 2011). Previous observations that the tapered shape of the wild type *C. crescentus* cell pole develops during the next cell cycle, not during septation or medial elongation (Aaron et al., 2007), corroborates the idea that MreB could participate in remodeling the cell wall at the poles. That the polar cell wall in *C. crescentus* undergoes remodeling instead of remaining entirely inert contradicts what is typically assumed for rod-shaped bacteria. Given that *C. crescentus* belongs to the Alphaproteobacteria class, many members of which exhibit polar growth (Brown et al., 2011, 2012), it may not be surprising if its poles do not follow the same rules as *E. coli* and *B. subtilis*.

In addition to its diversified polar regulators and various growth modes, *C. crescentus* hosts an expanded repertoire of bifunctional peptidoglycan synthases (penicillin-binding proteins, or PBPs). *E. coli* has three bifunctional PBPs. Although PBP1A and PBP1B exhibit semi-redundancy in function, PBP1A appears to preferentially associate with the elongasome (Banzhaf et al., 2012) and PBP1B with the divisome (Bertsche et al., 2006; Müller et al., 2007). The function of PBP1C remains unknown (Schiffer and Höltje, 1999). In contrast, *C. crescentus* has five predicted bifunctional PBPs (PBP1A, PbpC, PbpX, PbpY, PbpZ) which appear to be largely redundant, with any of them capable of functioning as the sole bifunctional PBP with the exception of PbpZ (Yakhnina and Gitai, 2013; Strobel et al., 2014). The same four enzymes were shown to be capable of interacting with divisome components FtsN, FtsL, and DipM (Strobel et al., 2014). Despite this redundancy, each PBP has a specific localization pattern during the cell cycle, suggesting specific cellular functions and roles: Pbp1A and PbpZ localize to the cell periphery; PbpY and PbpX to the periphery, division plane, and stalk; and PbpC to the stalk. Therefore, with the exception of PbpZ, these bifunctional PBPs have retained the ability to interact with both the elongasome and divisome while also evolving distinct affinities for specific localization factors.

*Caulobacter crescentus* exemplifies how the core peptidoglycan synthesis machinery can be adapted to create its specific morphology and to fit its lifestyle. While it retains an MreBdependent elongasome and an FtsZ-dependent divisome, it has diversified its growth modes by adding what appears to be a dual MreB- and FtsZ-mediated zonal growth mode at the division plane. These growth modes are strictly regulated in order to synchronize them with the cell cycle and to accommodate asymmetric division. The vibrioid shape arises from diversifying cytoplasmic scaffolds (CreS) and changing the interacting partners of MreB to achieve cell cycle-dependent dynamics. In addition, *C. crescentus* has expanded its suite of bifunctional PBPs and may have additional peptidoglycan-modifying enzymes. Finally, *C. crescentus* has evolved a specialized growth mode, stalk synthesis, which is described in the next section.

### **The Stalk as a Model System for Understanding Zonal Growth**

Stalk synthesis and elongation constitutes one growth mode unique to *C. crescentus* among model rod-shaped bacteria (although not among Alphaproteobacteria). A thin extension of the inner membrane, peptidoglycan, and outer membrane layers (Poindexter, 1964), the stalk is a morphologically distinct organelle thought to improve the nutrient scavenging ability of the cell (Poindexter, 1978; Ireland et al., 2002; Wagner et al., 2006). The narrow cytoplasm of the stalk is free of DNA, ribosomes and most cytoplasmic proteins (Poindexter and Cohen-Bazire, 1964; Ireland et al., 2002; Wagner et al., 2006) and compartmentalized by cross-bands, disk-like, proteinaceous structures that intersect the width of the stalk perpendicular to the long axis of the cell (Jones and Schmidt, 1973; Schlimpert et al., 2012). Cross-bands prevent exchange of membrane and soluble proteins between the stalk and cell body (Schlimpert et al., 2012). As a nonessential organelle, the stalk serves as a convenient model for zonal, or targeted, peptidoglycan growth (Wagner and Brun, 2007).

Stalk synthesis initiates at the cell pole during the swarmer to stalked cell transition phase of the cell cycle, and elongation occurs at the cell-stalk junction during pre-divisional cell elongation (Schmidt and Stanier, 1966; Aaron et al., 2007). MreB and RodA (an MreB-associated protein) depletion, or treatment with the PBP2 inhibitor mecillinam (Seitz and Brun, 1998), result in the loss or shortening of stalks, and recovery from depletion leads to the growth of ectopic stalked poles, suggesting that MreB, in concert with RodA and a PBP2 homolog, plays a role in recruiting peptidoglycan synthesis machinery for stalk synthesis (Wagner et al., 2005). Overexpression of RodZ (another MreB-associated protein) resulted in multiple stalks forming at the same pole, opposite pole, or on the cell body (Alyahya et al., 2009). RodZ localization dynamics coincided with FtsZ during the cell cycle but appeared to depend on MreB. Overall, these data suggest that stalk synthesis and elongation in *C. crescentus* potentially depends on a similar core apparatus as elongation, utilizing MreB as a scaffold. The mechanism for cross-band formation may have interesting intersections with the processes of stalk synthesis and elongation. Early on, electron micrographs of *C. crescentus* stalks showed that cross-bands consisted of concentric circular striations, possibly of peptidoglycan and membranes (Jones and Schmidt, 1973). Because the stalk continues to elongate as the stalked cell participates in increasing rounds of division, it was postulated that stalk length and number of cross-bands could indicate cell age. Quantitative analysis suggested that one crossband is added to the stalk toward the end of each reproductive cycle, possibly accompanying division (Poindexter and Staley, 1996; Schlimpert et al., 2012). The idea that cross-bands consist of peptidoglycan has been challenged by the discovery that crossbands are the product of the macromolecular assembly of least four proteins, StpABCD (Schlimpert et al., 2012). However, a relationship between cross-band assembly and the peptidoglycan synthesis machinery may still exist.

Like septation, lateral, or medial elongation in *C. crescentus*, stalk synthesis and elongation appear to be highly tuned to cell cycle regulation and polar positioning mechanisms. Moreover, the mechanism for stalk synthesis appears to depend on the expansion and diversification of the core machinery. Bactofilins BacA and BacB constitute a class of scaffolding proteins with proline-rich terminal regions that localize to the stalked pole of the cell during the swarmer to stalked cell transition (Kühn et al., 2010). Deletion of these genes led to a 45% reduction in stalk length, suggesting a recruitment role in stalk assembly. Indeed, the localization of the stalk-specific bifunctional peptidoglycan synthase PbpC to the stalk was shown to be dependent on BacA and BacB. PbpC potentially acts in conjunction with PbpX in stalk elongation (Yakhnina and Gitai, 2013; Strobel et al., 2014), however, it has also been shown to have a role in recruiting and modifying a stalk-specific protein. StpX is a stalk-specific membrane protein that promotes the elongation of the stalk in nutrient-limiting conditions (Hughes et al., 2010). PbpC is not only required for localizing StpX to the stalk-cell junction at onset of stalk synthesis, but appears to be directly or indirectly involved in the process of tethering StpX to the outer membrane or to an outer membrane protein (Hughes et al., 2013). That this phenomenon takes place regardless of PbpC's transglycosylase or transpeptidase activities suggests alternative roles for various PBPs and other enzyme classes in regulating the peptidoglycan synthesis machinery, opening up the diversity of mechanisms for the evolution of novel functions.

Phylogenic studies indicate that stalk positioning evolved from an ancestral, single polar stalk in the order *Caulobacteraceae*, to a single sub-polar stalk in *Asticcacaulis excentricus*, and subsequently to bilateral stalks in *Asticcacaulis biprosthecum* (Jiang et al., 2014). The stalk structure appears identical in *Caulobacter* and *Asticcacaulis* (Pate and Ordal, 1965) and all three species appear to share the same stalk synthesis mechanism of inserting peptidoglycan at the base of the stalk (Jiang et al., 2014). The natural variation and evolution of stalk positioning in these species correlates with the localization of SpmX, a developmental regulator (Radhakrishnan et al., 2008) that has been shown to be necessary for stalk synthesis in*Asticcacaulis* but not in*Caulobacter* (Radhakrishnan et al., 2008; Jiang et al., 2014). Therefore, in *Asticcacaulis* SpmX has been co-opted as a stalk-positioning factor. SpmX consists of three defined regions: a highly conserved N-terminal muramidase domain, an unstructured intermediate region that has been expanded by over 370 amino acids in the *Asticcacaulis* genus, and a two-pass transmembrane domain. Recent work has demonstrated that either the intermediate region or the transmembrane domain contain the residues responsible for SpmX's new role in sub-polar and bilateral stalk positioning (Jiang et al., 2014). The muramidase domain, while necessary for overall function and localization, is interchangeable amongst species. How a murein hydrolase participates in SpmX's function and localization will be an interesting question for future studies.

The recent discovery that SpmX coordinates stalk placement in the *Asticcacaulis* genus underscores the utility of not only studying the stalk as a model system for zonal growth, but of exploiting the natural diversity of stalked morphologies observed in Alphaproteobacteria. In marked contrast to division and elongation genes, which require depletion strains, stalk-less mutants are far easier to identify in a genetic screen. *Asticcacaulis* provides an opportunity for screening stalkless mutants with fewer chances of impacting elements critical for cell viability, since stalk synthesis no longer overlaps with polar regulation. Finally, altering the location of the stalk may prove to be a far more advantageous phenotype than losing the stalk entirely in terms of discovering localization and recruitment factors for stalk initiation and maintenance. Therefore studying stalk localization, initiation, and maintenance in *Asticcacaulis* will likely make significant inroads into understanding how bacteria can redirect the core peptidoglycan machinery to create novel morphologies.

### **Conclusion and Summary**

Critical advances in studying bacterial growth and division in model bacteria such as *E. coli*, *B. subtilis*, and *C. crescentus* have identified and characterized the fundamental components of the molecular machinery responsible for creating and shaping the sacculus. This expanding body of work reveals a general strategy for growing and shaping the bacterial cell wall. Moreover, it suggests ways in which the core can be adapted for various bacterial lifestyles as well as molecular mechanisms for the evolution of novel morphologies. The peptidoglycan synthesis machinery consists of a central core of cytoskeletal-like scaffolding proteins, inner membrane-spanning elements, and periplasmic or extracellular peptidoglycan-modifying elements. The modularity and evolvability of this system is inherent in how different elements have been retuned or repurposed in each genus for species-specific growth modes and morphologies. As different species of bacteria become genetically tractable, it is becoming clear that the diversification and expansion of any of the core elements of the peptidoglycan synthesis assembly underlie morphological diversity and alternate growth modes.

As studies in *C. crescentus* and the closely related genus *Asticcacaulis* demonstrate, much can be learned about general growth strategies using comparative approaches with related families. The great diversity of morphologies and growth modes in Alphaproteobacteria (**Figure 4**) offers an opportunity to exploit natural diversity to dissect mechanisms for stalk localization and synthesis, as well as polar growth. The recent advances in expressing fluorescent fusions in *Hyphomonas neptunium* open up yet another opportunity for discovering genes responsible for polar growth and stalk development (Jung et al., 2014). The increasing ease and affordability of genome sequencing allows further investigation into the suites of genes underlying

### **References**

uncharacterized growth modes in other genera. New reagents that fluorescently label sites of peptidoglycan synthesis are providing simple methods to determine growth modes, even in understudied species and environmental samples (Kuru et al., 2012; Pilhofer et al., 2013; Liechti et al., 2014). Determining how bacteria actively restructure their morphologies, or how they have evolved various morphologies over time, remains a major goal in bacteriology. Expanding investigations into understudied genera with novel morphologies and growth modes will complement and enrich our understanding of how bacteria grow and proliferate.

### **Acknowledgments**

Our research on morphogenesis is supported by National Institutes of Health Grants GM051986 and GM113172 to YB AR was supported by National Institutes of Health National Research Service Award F32GM112362. We thank David Kysela for the phylogenetic tree, and Kevin Young and members of the Brun laboratory for critical reading of the manuscript.

**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 Randich and Brun. 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.*

# Structural constraints and dynamics of bacterial cell wall architecture

#### *Miguel A. de Pedro1,2 and Felipe Cava2\**

*<sup>1</sup> Centro de Biología Molecular "Severo Ochoa" – Consejo Superior de Investigaciones Científicas, Universidad Autónoma de Madrid, Madrid, Spain, <sup>2</sup> Laboratory for Molecular Infection Medicine Sweden, Department of Molecular Biology, Umeå Center for Microbial Research, Umeå University, Umeå, Sweden*

The peptidoglycan wall (PG) is a unique structure which confers physical strength and defined shape to bacteria. It consists of a net-like macromolecule of peptide interlinked glycan chains overlying the cell membrane. The structure and layout of the PG dictates that the wall has to be continuously modified as bacteria go through division, morphological differentiation, and adaptive responses. The PG is poorly known in structural terms. However, to understand morphogenesis a precise knowledge of glycan strand arrangement and of local effects of the different kinds of subunits is essential. The scarcity of data led to a conception of the PG as a regular, highly ordered structure which strongly influenced growth models. Here, we review the structure of the PG to define a more realistic conceptual framework. We discuss the consequences of the plasticity of murein architecture in morphogenesis and try to define a set of minimal structural constraints that must be fulfilled by any model to be compatible with present day information.

### *Edited by:*

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

#### *Reviewed by:*

*Waldemar Vollmer, Newcastle University, UK Kevin D. Young, University of Arkansas for Medical Sciences, USA*

#### *\*Correspondence:*

*Felipe Cava, Laboratory for Molecular Infection Medicine Sweden, Department of Molecular Biology, Umeå Center for Microbial Research, Umeå University, 90187 Umeå, Sweden<EMAIL_ADDRESS>
#### *Specialty section:*

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

> *Received: 05 March 2015 Accepted: 25 April 2015 Published: 08 May 2015*

#### *Citation:*

*de Pedro MA and Cava F (2015) Structural constraints and dynamics of bacterial cell wall architecture. Front. Microbiol. 6:449. doi: 10.3389/fmicb.2015.00449* Keywords: cell wall, peptidoglycan, structure, cross-link, chain length, HPLC, muropeptides

### Introduction

The bacterial cell wall was first described as a region of specific staining behavior wrapping the cell body (Churchman, 1927). This structure was soon associated with the processes of cell division and growth (Knaysi, 1930, 1941). Introduction of electron microscopy in conjunction with the development of cell wall purification methods, demonstrated the ability of purified walls to retain cell shape (Salton and Horne, 1951a,b), and characterized cell walls as mesh bag-like macromolecules called sacculi (from the Latin for small bags, sing. sacculus; Weidel et al., 1960). These early techniques were also critical to define the glyco-peptidic nature of the cell wall, which became also known as the peptidoglycan (PG) layer, and led to the establishment of its polymeric nature (Salton, 1952, 1953, 1956, 1960; Weidel, 1960; Primosigh et al., 1961; Salton, 1961; Weidel and Pelzer, 1964). Schleifer and Kandler (1972) neatly summarized the knowledge on the structure of bacterial walls accumulated until the early 1970s and sort of wrote in stone most of the basic notions that still hold today, namely: the universality of the basic structural subunit; the long-glycan/short peptide idea; the chemical uniformity of PG and its planar, ordered, net-like organization in Gramnegative bacteria, among others. Unavoidably, some of these ideas derived into a geometric and static perception of the PG layer as depicted in so many articles and textbooks.

An aspect of PG studies important to stress at an early point is quantification. Most studies present the abundance of the individual kinds of PG subunits (muropeptides) as a percentage of the total number because this makes measurements independent of the absolute amount of sample. However, this practice makes it easy to underestimate the relevance of minor components. The amount of PG in an *Escherichia coli* cell is around 3.5 × 106 monomeric units (Wientjes et al., 1991). Therefore, values as low as 0.1% mean that each cell has about 3 × 10<sup>3</sup> molecules of that particular muropeptide, a meaningful number on biological terms.

Here, we will review the experimental evidence accumulated on PG structure and metabolism in an attempt to define a series of basic structural constraints that can help us to revisit current models of the bacterial cell wall. In this study we will center on the features of Gram-negatives because of the relative abundance of studies, and the (apparently) simpler organization of their sacculi. However, most of the information available comes from a very limited number of bacterial species and generalizations have to be taken with great precaution. Particular emphasis will be made on the influence of individual muropeptides on local structure and on how subtle changes in the subunits might influence the properties of the cell wall.

### The Structure of the PG Monomers and Linear Polymers

The primary structure of PG monomers was defined in the early 1960s and it was soon found out that most Gram-negatives shared as common basic subunit the disaccharide pentapeptide: GlcNAc-(β,1−→4)MurNAc-(L)ala-(D)glu-(γ)-(*meso*)Dap- (D)ala-(D)ala (M5; Schleifer and Kandler, 1972). The monomeric subunit might suffer a number of chemical modifications, but in most instances, the alternancy of L- and D-amino acids, as well as the presence of a terminal D-ala-D-ala dipeptide is respected (Vollmer, 2008; Vollmer et al., 2008).

The PG monomer is a molecule with rather unusual biochemical and structural properties (**Figure 1A**) that, according to molecular modeling studies, results in the peptide moiety adopting a curled, compact disposition with the terminal D-ala interacting with the L-ala at position 2 by means of hydrogen bonds (**Figure 1B**; Barnickel et al., 1979; Meroueh et al., 2006; Gumbart et al., 2014). This disposition of the amino acid residues means that the stem peptides can potentially span considerably larger distances, from *ca.* 10 Å up to *ca.* 25 Å when stretched (up to 2.5 fold), as predicted by molecular modeling (Barnikel et al., 1983).

Although some species as *Caulobacter crescentus* retain the terminal D-ala-D-ala in a high proportion of PG subunits (Takacs et al., 2010), most species contain mostly tetrapeptides and tripeptides following the sequential elimination of the D-ala residues to variable extents (Quintela et al., 1995). The kinetics of elimination of the terminal D-ala have been measured in *E. coli* (Glauner and Höltje, 1990). Assuming every monomer is added as a pentapeptide, nascent PG accumulated for as little as 20 s exhibits only about 25% pentapeptides, 60 s later it drops to about 5% and in a further few minutes goes well below 0.5%. As the terminal D-ala is required for the polymerization of PG the modified muropeptides are disabled to act as donors in DD-transpeptidation reactions (see below). Elimination of the terminal D-ala also affects folding of the stem peptides favoring a more extended conformation and displacements of the potential NH2 acceptor sites in the *(meso)*DAP residue (**Figure 2**).

#### FIGURE 1 | Continued

software (http://avogadro.cc/) using the MMFF94s field force, the steepest descent algorithm and a convergence of 10−7. Muropeptides were modeled in the absence of solvent. The optimized molecular structure was visualized and prepared for publication with Molsoft ICM software (http://www.molsoft.com/).

In order to weave the polymeric cell wall the subunits must first form linear polymers, a reaction performed by bifunctional DD-transpeptidase-transglycosylase enzymes, the Class A Penicillin-Binding Proteins (PBP's; Typas et al., 2012). The polymerization reaction happens in the periplasmic space and occurs by the transfer of a molecule of the activated precursor M5-P-Bactoprenol (lipid-II) to the C4-OH of the GlcNAc moiety in a second lipid-II molecule to generate a (β, 1−→4) glycosidic bond (van Heijenoort, 2001). The growing glycan is then reiteratively transferred to new molecules of lipid-II causing longitudinal extension of the PG strand. The mechanism for the termination and release of newly made glycan strands remains unclear. In most bacteria, glycan strands display a molecule of (1−→6) anhydro MurNAc (Anh-MurNAc) as the C1 terminal sugar, suggesting the participation of a transglycosylase activity in the process. However, there is no reason to discard the existence of alternative mechanisms. Indeed, *Agrobacterium tumefaciens* and *Mesorhizobium meliloti* either lack Anh-MurNAc or have amounts undetectable by HPLC and MALDI-MS techniques (Quintela et al., 1995; Brown et al., 2012).

The sugar backbone of a PG linear polymer tends to adopt a right-handed helical structure (Leps et al., 1987; Meroueh et al., 2006; Gumbart et al., 2014). Calculations derived from actual NMR analysis of a linear dimer indicate three disaccharides per turn with consecutive subunits at roughly 120◦, and a pitch of *ca.* 3 nm (**Figure 3B**; Meroueh et al., 2006; Gumbart et al., 2014) although a certain degree of variation in the helicity looks likely (Gumbart et al., 2014). This disposition implies that in a state of minimal energy, the sugar backbone is almost fully stretched, therefore when subjected to tensile forces the glycan polymer remains essentially unyielding. This fact contrasts with the relaxed disposition of the peptide moieties and makes the cell wall anisotropic respect to some properties (Gumbart et al., 2014). **Figure 3**, shows a likely conformation for a linear dodecamer as calculated by computer modeling. Other than the compact disposition of the stem peptides, is interesting to note that the critical regions, the D-ala-D-ala dipeptides and the D-NH2 groups of (*meso*)DAP molecules, involved in the all-important crosslinking reactions occupy rather accessible positions at the surface of the molecule (**Figure 3B**).

### Length of Glycan Strands

Length of the glycan strands is an important structural parameter of PG. Because the glycan backbone of PG strands is inextensible, the longer the strands the more rigid the structure of the sacculus. The PG can be compared to a composite material in that the global properties of the structure depend to

FIGURE 2 | Elimination of the terminal D-ala residue has strong influence on the 3D structure of PG subunits. PG monomeric subunits derived from the M5 by elimination of the terminal D-ala (M4) or the terminal D-ala-D-ala (M3) are compared in two orientations, transversal to (upper panel) and from above (lower panel) the surface of the sacculus. To facilitate

comparison muropeptides were rotated until GlcNAc residues overlap. Red designates the glycan moiety and blue the stem peptide. Terminal D-ala residues are highlighted in yellow. The 3D structures were calculates as in Figure 1. Hydrogen atoms were removed in the image, but considered for the calculation of the molecular surface.

a large extent on the length, orientation, and bonding of the fibers.

Because of the specific presence of Anh-MurNAc at the glycan ends (van Heijenoort, 2001), it is possible to calculate experimentally the average length (AL) of PG strands as the inverse of the molar fraction of Anh-MurNAc-containing subunits in the PG (Glauner, 1988). Available information supports a large variability in the AL of glycan strands amongst different bacteria. In *E. coli* the AL for cells actively growing in rich media is around 30 monomers/strand (M/S; Glauner et al., 1988). Most Gram-negatives analyzed so far fall in the 20–100 M/S range with a preference for the lower values (10–40 M/S). However, values lower than 10 M/S have been measured in *C. crescentus* and *Helicobacter pylori* (Costa et al., 1999; Takacs et al., 2010). AL is not constant but strongly influenced by the state of growth (Pisabarro et al., 1985; Lam et al., 2009; Cava et al., 2011), and by mutational or chemical alterations of cell wall metabolism (Romeis et al., 1991; Varma et al., 2007). In *E. coli* growth arrest causes a progressive reduction in AL (25–30%) until a constant value is reached in resting cells. Resumption of growth is accompanied by a progressive rise in AL which reaches its characteristic value after 2–3 doublings in mass. However, measurements of AL in newly made *vs.* total PG in *E. coli*, showed that AL was essentially constant for the new PG made during the transitions (Pisabarro et al., 1985). These observations support the idea that AL variations are not due to a change in the frequency of glycan chain terminating events, but rather to processing of macromolecular PG. Therefore, bacteria must have some modulatory mechanisms able to modify the AL of PG in response to environmental conditions.

An important aspect is that although the length of average PG strands (∼10–100 nm) is very small when compared to the dimensions of the cell, in most cases the strands are considerably longer than the thickness of the sacculus, which is between 2.5 and 7 nm for hydrated sacculi (Yao et al., 1999).

Although AL certainly gives important structural information, it is the distribution of glycan chain lengths that really influences the properties of the sacculus. However, experimental determination of length distributions has been achieved in a very limited number of cases. The only extensive study was performed in *E. coli* (Harz et al., 1990). Whether or not the information obtained is representative for other bacteria is uncertain. The analysis showed a very wide and positively skewed size distribution with a modal value of eight M/S (Harz et al., 1990). However, this distribution only accounts for about 70% of the total PG. The remaining 30% could not be resolved into individual length classes but presents an AL equivalent of 45 M/S (Harz et al., 1990). At present, there are no data to choose a particular distribution of lengths for the longer fraction. However, as the shorter strands have to be at least 31 disaccharides long, and the AL is 45, the existence of extremely long (>500 disaccharides or roughly 500 nm) glycans is not very likely. The determination of both the AL and length distribution indicates that the sacculus is mostly made of rather short glycan strands but covering a very wide range of lengths, an aspect often neglected in cell wall models.

There have been a limited number of attempts to study how different conditions affect the glycan strand length distribution (Romeis et al., 1991; Obermann and Holtje, 1994; Ishidate et al., 1998; Ursinus et al., 2004). Particularly interesting is the comparison of regular rods and mini-cells in a mini-cell producing *E. coli* mutant (Obermann and Holtje, 1994). The results showed moderately shorter strands in the mini-cells with a reduction in AL from 28 to 25 M/S and a drift toward shorter lengths in the size distribution. A similar study performed with *E. coli* cell division mutants is also enlightening about local variations in glycan length distributions (Ishidate et al., 1998). This study showed that PG associated with the initiation of septation is likely made of relatively long glycan strands, and interestingly, the cell division protein FtsN requires strands >25 M/S to interact with septal PG (Ursinus et al., 2004).

Most of the information above is consistent with the existence of mechanisms able to make glycan strands of adequate lengths as a function of environmental conditions and surface topology.

### Cross-Linking of Glycan Strands

Formation of the sacculus requires glycan strands to be crosslinked to each other in a net-like fashion. Cross-linking is catalyzed by specific peptidyl-transferases which are able to transfer a peptide bond from a stem peptide (donor) to the free NH2 group at the D-center of the *(meso)*DAP residue of a second stem peptide (acceptor) from a nearby glycan strand. Most bacteria are capable of cross-linking reactions mediated by DDtranspeptidase activities residing in the bifunctional class A PBPs (Typas et al., 2012). These enzymes catalyze the transfer of the peptide bond linking the two terminal D-ala residues of the donor moiety, to generate a new peptide bond with a DD configuration. DD-transpeptidation seems to be a universal mechanism to interlink glycan strands, but it is not necessarily unique. Indeed, a few alternative cross-linking reactions have been described (Vasstrand et al., 1982; Glauner et al., 1988; Boniface et al., 2009; Bui et al., 2009). The more extended "accessory" cross-linking reaction is mediated by LD-peptidyl transferases (LD-TPases) which transfer the L–D peptide bond between the L-center of *(meso)*DAP and the D-ala at position 4 in the donor stem peptide to the NH2 group of the acceptor *(meso)*DAP (the very same as for the DD-reaction), generating a L-*(meso)*DAP → D-*(meso)*DAP peptide bond (Glauner and Höltje, 1990). LD-TPases are monofunctional enzymes, structurally unrelated to the PBPs, and are involved in a number of functions such as attachment of PGbound lipoprotein (Magnet et al., 2007) and incorporation of D-amino acids (Cava et al., 2011). Although *E. coli* and most other Gram-negatives that have been analyzed have both kinds of cross-links, LD cross-linking is neither universal (Cava et al., 2011) nor essential for bacteria with both LD and DD crosslinkage (Magnet et al., 2008; Cava et al., 2011). The relative abundances of LD and DD cross-linking are very variable, but in some species can be equally abundant (Lavollay et al., 2008; Brown et al., 2012). Why some species have both kinds of crosslinks is not clear yet. The fact that LD-enzymes are, in most cases, insensitive to beta-lactams has led to the proposal of LD-crosslinks as contributors to escape antibiotic action (Gupta et al., 2010).

DD cross-linking of new strands is coupled to linear polymerization and is the reaction responsible for the incorporation of nascent strands into the sacculus. Indeed, analysis of nascent PG shows that right after incorporation, the new material already exhibits a content of DD-cross-links that is close to the average value for total PG (Glauner and Höltje, 1990). However, LD cross-links are initially absent (or below experimental detection limits) and then increase in a progressive way as PG ages (typically hours). This difference in kinetics supports the idea that LD cross-links are not involved primarily in the insertion of new precursors, but rather as part of PG maturation (Glauner and Höltje, 1990; Brown et al., 2012). However, LD-TPases involved in LD cross-linking work preferentially on those regions where incorporation of precursors occurs (Kuru et al., 2012; Siegrist et al., 2013), suggesting a close association between PG synthesis and LD-TPase activity. These considerations suggest that LD cross-links are an accessory feature of PG and may play important physiological and structural functions, particularly in those species where they are more abundant (Cameron et al., 2014).

Comparison of computer models suggests important differences between LD and DD cross-links. The absence of the D-ala residue linking the two *(meso)*DAP molecules in LD crosslinks makes the stem peptide more rigid, with a more extended conformation reaching a longer distance than the equivalent DD cross-link (1 nm *vs.* 0.7 nm distance between the MurNAc-C3 carbons of each strand, respectively; **Figure 4**). The possibility of additional hydrogen bonds in the DD cross-links affects the preferred orientations of the disaccharide moieties, and favors a more compact folding of the stem peptides. These differences imply that each kind of cross-link might fit optimally into subtly different relative orientations and distances between the glycans, and that local accumulations of one or the other crosslink should influence the local properties of PG differently. The longer range of LD cross-links might be especially well suited to connect glycans which are already under stress, as expected from the apparently post-insertional nature of this kind of cross-linkage.

Cross-linking most frequently takes place between the stem peptides of muropeptides belonging to two glycan strands. However, in a variable but often considerable number of cases, three or, more rarely, four glycan strands become interconnected by their stem peptides generating cross-linked trimers and tetramers, respectively (**Figure 5**). Cross-linked trimers seem to be rather universal and have been detected in most Gram-negatives studied in relatively high proportions. Crosslinked tetramers are rarer than trimers. In most cases tetramers are not detected, but in a few instances sizable amounts have been measured, as in the case of *C. crescentus* (Takacs et al., 2010). Most abundant trimers are fully DD cross-linked. However, smaller amounts of hybrid (DD plus LD) and full LD cross-linked trimers are present in PG from bacteria able to LD cross-link PG (Glauner and Höltje, 1990; Brown et al., 2012).

According to Glauner and Höltje (1990), in *E. coli* DD trimers are readily detected in nascent PG with an age of 1 min. As the proportion of trimers in PG is rather low (about

transversal view, and lower images to a view from above. Each pair was oriented according to the GlcNAc residues indicated by the black arrows. Red, green, and blue designate the disaccharide for the donor moiety, the disaccharide for the acceptor moiety and the stem peptide, respectively. Yellow highlights the position of the cross-link. Note the inverted orientation predicted for the donor and acceptor moieties.

3–4% of total muropeptides), this supports the idea of DDtrimers being synthesized concomitantly with the incorporation of new glycans into the pre-existing PG fabric. This is further supported by the kinetics of D-ala-D-ala elimination from nascent PG. Formation of DD-trimers should depend on the presence of subunits with a pentapeptide stem peptide in the donor moiety. The terminal D-ala is quickly removed from nascent PG. Therefore, DD-trimers should be made concomitantly or very shortly (<2 min) after incorporation of new subunits into a growing strand. With respect to how, when and where DD/LD hybrids or full LD trimers are made, there are no experimental data reported.

The presence of sizable proportions of cross-linked trimers and tetramers in the PG has deep structural consequences. As illustrated in **Figure 5A**, in the case of trimers the geometric constraints imposed by the nature of the molecule mean that the three glycan strands cross each other in a non-planar configuration. Indeed, at least one of the glycans has to follow a trajectory on a markedly different plane to that defined by the other two strands. A similar, even more complex situation is generated by the presence of tetramers (**Figure 5B**) which force a disposition of the strands in different planes. The normal abundance of trimers is rather variable, but taking *E. coli* as a reference (4–5% of total muropeptides) a sacculus would contain roughly 6 <sup>×</sup> <sup>10</sup><sup>4</sup> such glycan-strand knots. As the number of strands per sacculus is *ca*. 1.1 <sup>×</sup> <sup>10</sup><sup>5</sup> (3.5 <sup>×</sup> <sup>10</sup><sup>6</sup> monomers per sacculus with an AL of *ca*. 30 M/S) (Harz et al., 1990; Wientjes et al., 1991) on average, each glycan chain participates in 1–2 of these knots. The contribution of tetramers is more difficult to estimate, as their presence in many species is doubtful. In *E. coli* their amount is close to the detection limit (0.2% of total muropeptides). This apparently low abundance still means about 2 × 10<sup>3</sup> tetramers/sacculus, or that roughly one out of 16 strands is cross-linked by one tetramer. In those species were they are present in larger amounts as in *C. crescentus* (>3%; Takacs et al., 2010), their effect should lead to a very disordered orientation of the strands. Such a situation indicates that most strands weave over and under other strands and therefore, cannot form the kind of ordered, planar net often presented in representations and models of the cell wall.

## Knitting the Net, Cross-Links, and Glycan Strands

*Escherichia coli* PG has a considerable proportion (about 30%) of strands shorter than eight disaccharide subunits (DSs; Harz et al., 1990). The degree of cross-linkage is about 30% for *E. coli*. How this value is calculated means that there are 30 peptide bridges for each 100 muropeptides. Therefore, most strands equal or shorter than eight DS would statistically be cross-linked at two or fewer points. Such a structure would behave like a single long crosslink when subjected to stress (Costa et al., 1999), unless one of the cross-linking points were a trimer or tetramer connecting the short strand to two or three other strands, respectively. Quite interestingly, the terminal muropeptides of PG strands are hyper cross-linked (Glauner and Höltje, 1990; Lam et al., 2009; Takacs et al., 2010). Most strands terminate in a cross-linked dimer, but many in trimers and tetramers. The preference for a particular kind of terminal muropeptide could work as a compensating mechanism to ensure the continuity of the net-like structure under such circumstances. For similar reasons strands shorter than four DS (still about 10% of total PG) would be cross-linked at only one point and could not contribute to the strength of the stress bearing structure. These extremely short "strands" could represent material being recycled on its way out of the sacculus. Nevertheless, the above arguments assume a homogeneous distribution of cross-links, but this assumption could be wrong. An alternative would be that short chains were proportionally more cross-linked than long ones. Under these circumstances the short strands could play the role of staples joining longer chains at certain locations, whereas the long, under-cross linked, strands could deform more easily.

In some cases low AL values are associated with elevated proportions of cross-linked muropeptides, in particular trimers and tetramers (Takacs et al., 2010). The high proportions of trimers and tetramers might compensate for the shortness of the strands increasing the probability that they will cross-link to more than two other strands and therefore help create a relatively tight net made up of short strands. *H. pylori* represents an extreme case, with an AL of about 7 M/S in actively growing cells, but with a PG that lacks trimers and has a low degree of cross-linking (27% of dimers). In cases like this the NAc-muraminyl end of a short strand could be linked to the NAc-glucosaminyl end of another

by a peptide cross-link, generating a longer linear polymer with alternating glycan and peptidic regions that could interconnect relatively distant strands and contribute to the strength of the sacculus (Costa et al., 1999).

### Organization and Orientation of PG Strands

Two important parameters define the structure of the sacculus: the total amount of PG subunits available, and the orientation of the glycan strands. Indeed, how PG is stretched to cover the area of the cell and how many PG layers exist in Gram-negatives have been matters of debate since the earliest works on cell walls (Wientjes et al., 1991).

Small-angle neutron diffraction experiments on purified *E. coli* sacculi (Labischinski et al., 1991) indicated that the thickness of PG in *E. coli* is non-homogeneous, with about one third of the surface thick enough (7 nm) to accommodate a triple layer of PG, whilst the rest has a thickness (2.5 nm) compatible with a single layered structure. However, the method only estimates the proportion of areas with each thickness, but gives no data about their distribution. These data are compatible with the distortions expected from the presence of cross-linked trimers and tetramers, as discussed above (Labischinski et al., 1991). Further support for a (partially) multilayered organization was obtained from cryo-EM (Hobot et al., 1984) and AFM (Yao et al., 1999). The later methodology provided the first measurements of PG thickness in the hydrated state and showed that hydration strongly influences this parameter. Direct determination of the amount of PG per cell (3.5 <sup>±</sup> 0.6 <sup>×</sup> <sup>10</sup><sup>6</sup> monomers/cell) and per unit of cell surface area (PG density) was also compatible with the kind of partially multilayered ordering suggested by other techniques (Wientjes et al., 1991). However, the calculations were based on a rather primitive model for the PG network (Oldmixon et al., 1974), and ignored the size distribution of strands and its possible relaxation effects on the PG fabric (Huang et al., 2012).

A second very important, and often neglected, observation from the AFM work was the demonstration that purified sacculi from Gram-negatives could have different thicknesses. Indeed, sacculi from *Pseudomonas aeruginosa* were about half the thickness of those from *E. coli* (Yao et al., 1999). This result clearly indicates that PG thickness might be a variable parameter. Indeed, studies on the effects of specific mutations (Lam et al., 2009), partial starvation for PG precursors (Prats and de Pedro, 1989) and the effects of D-amino acids on the structure of PG (Caparros et al., 1994; Cava et al., 2011) indicate that, at least some bacteria can accommodate large variations in the amount of PG per unit of cell surface, without apparent consequences for morphology, growth, and division. A relevant observation in *E. coli* is that the only significant variation detected in the composition of PG from cells with low PG content was a shortening in the AL of glycan strands (from 28 down to 19 M/S), a change that might change the elasticity of the sacculus to favor stretching (Huang et al., 2008). The surface density of PG seems to suffer large changes not only in response to harsh treatments, but also during the regular life cycle of bacteria (leaving aside those having specialized resistance forms), with resting cells accumulating considerably more PG per unit area than their actively growing counterparts (Caparros et al., 1994; Cava et al., 2011). Therefore, in at least some bacteria, the organization of the sacculus has to be compatible with drastic alterations in the PG surface density without causing macroscopic alteration in the shape and growth of the cell.

The relative orientation of the glycan strands with respect to the axes of the cell is another "hot topic" in cell wall biology. Because PG is not isotropic with respect to some characteristics, particularly elasticity (Gumbart et al., 2014), the disposition of strands in the sacculus has an important influence on cell wall properties. Measurements of the mechanical characteristics of sacculi show that they behave as a perfectly elastic material but with a high anisotropy (Yao et al., 1999). In fact, the elastic modulus for hydrated PG sacculi from either *P. aeruginosa* or *E. coli*,

### References

Barnickel, G., Labischinski, H., Bradaczek, H., and Giesbrecht, P. (1979). Conformational energy calculation on the peptide part of murein. *Eur. J. Biochem.* 95, 157–165. doi: 10.1111/j.1432-1033.1979.tb12950.x

was 2–3 times larger in the longitudinal dimension (the long axis of the cell) than in the transverse direction, meaning that under pressure the sacculus would become proportionally longer. Such a behavior is a strong indication that the glycan strands transverse the long axis of the cell. In contrast, a perfectly (or close to perfect) perpendicular orientation of the glycan chains would be difficult to reconcile with arguments against a planar organization of these strands. Instead, analysis of purified sacculi by cryo-electron tomography supports a disordered circumferential arrangement of the glycan strands, which show a preferential, but not strict, transverse alignment relative to the cell axis (Gan et al., 2008). This disposition is also supported by advanced computational approaches, which find that such a layout best fits the mechanical properties of sacculi (Huang et al., 2008; Gumbart et al., 2014).

### Concluding Remarks

The arguments discussed above indicate that there is not such a thing as a "Gram-negative sacculus" and if there were, it would be quite far from a regular or highly ordered structure. Nonetheless, the most important point we wish to make is how little real structural information is available. Many readers will be surprised by the age of the citations in this review. The reason is simply that many important lines of work have been abandoned or not pursued with the intensity that is required to acquire more comprehensive information on cell wall biology. The biological diversity of the cell wall seems to be much larger than initially expected in terms of structure and composition, questioning the universality of the models developed on the basis of *E. coli* data. To explore the real range of cell wall structural variability an "omics" approach looks like the best direction to go. Recent methodological advances make fast analysis of large numbers of PG samples feasible, and open the way to a methodical, large scale survey of bacterial cell walls. New techniques in electron, atomic force, and optical (fluorescence) microscopy are also demonstrating enough resolving power to contribute substantially to the unraveling of the fine structure of cell sacculi. Further development and systematic application of these new capabilities should provide the data required for a realistic understanding of bacterial cell wall structure and biology.

### Acknowledgments

This work was supported by the Laboratory for Infection Medicine Sweden (MIMS), the Knut and Alice Wallenberg Foundation (KAW), and the Swedish Research Council to FC, and UCMR Linneus professorship to MAP.

Barnikel, G., Naumann, D., Bradaczek, H., Labischinski, H., and Giesbrecht, P. (1983). "Computer-aided molecular modelling of the three dimensional structure of bacterial peptidoglycan," in *The Target of Penicillin: The Murein Sacculus of Bacterial Cell Walls, Architecture and Growth*, eds J.-V. H. R. Hakenbeck and H. Labischinski (Berlin: de Gruyter), 61–66.

**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 de Pedro and Cava. 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 membrane: transertion as an organizing principle in membrane heterogeneity

*Kouji Matsumoto1\*, Hiroshi Hara1, Itzhak Fishov2, Eugenia Mileykovskaya3 and Vic Norris4*

*<sup>1</sup> Department of Biochemistry and Molecular Biology, Graduate School of Science and Engineering, Saitama University, Saitama, Japan, <sup>2</sup> Department of Life Sciences, Ben-Gurion University of the Negev, Beer-Sheva, Israel, <sup>3</sup> Department of Biochemistry and Molecular Biology, University of Texas Medical School at Houston, Houston, TX, USA, <sup>4</sup> Laboratory of Microbiology Signals and Microenvironment EA 4312, Department of Science, University of Rouen, Mont-Saint-Aignan, France*

#### *Edited by:*

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

#### *Reviewed by:*

*Marc Bramkamp, Ludwig Maximilians University of Munich, Germany Ding J. Jin, National Institutes of Health, USA Germán Rivas, Consejo Superior de Investigaciones Científicas, Spain*

#### *\*Correspondence:*

*Kouji Matsumoto, Department of Biochemistry and Molecular Biology, Graduate School of Science and Engineering, Saitama University, 255 Shimo-ohkubo, Sakura-ku, Saitama-shi, Saitama 338-8570, Japan<EMAIL_ADDRESS>
#### *Specialty section:*

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

> *Received: 26 March 2015 Accepted: 25 May 2015 Published: 12 June 2015*

#### *Citation:*

*Matsumoto K, Hara H, Fishov I, Mileykovskaya E and Norris V (2015) The membrane: transertion as an organizing principle in membrane heterogeneity. Front. Microbiol. 6:572. doi: 10.3389/fmicb.2015.00572* The bacterial membrane exhibits a significantly heterogeneous distribution of lipids and proteins. This heterogeneity results mainly from lipid–lipid, protein–protein, and lipid– protein associations which are orchestrated by the coupled transcription, translation and insertion of nascent proteins into and through membrane (transertion). Transertion is central not only to the individual assembly and disassembly of large physically linked groups of macromolecules (*alias* hyperstructures) but also to the interactions between these hyperstructures. We review here these interactions in the context of the processes in *Bacillus subtilis* and *Escherichia coli* of nutrient sensing, membrane synthesis, cytoskeletal dynamics, DNA replication, chromosome segregation, and cell division.

Keywords: membrane, transertion, hyperstructures, lipid domain, *Escherichia coli, Bacillus subtilis*

### Introduction

The laterally heterogeneous distribution of the lipid and the protein components of bacterial cell membranes is the current paradigm, having replaced the homogeneous distribution of these components that is assumed in the fluid mosaic membrane model. This heterogeneity is involved in producing the specific environments needed for membrane proteins to participate in the many important processes that are associated with cell membranes. This raises the question of the nature of the mechanisms responsible for membrane heterogeneity.

One of the main causes of large-scale heterogeneity in bacterial membranes is the coupled transcription, translation and insertion of nascent proteins into and through membrane, *alias* transertion (Norris and Madsen, 1995; Binenbaum et al., 1999; Bakshi et al., 2014; **Figure 1**). The tethering of nascent proteins orchestrates interactions that result in the formation of membrane domains. These interactions include protein–protein interactions and lipid–protein interactions – and therefore also include lipid–lipid interactions (Vereb et al., 2003). The multitude of processes in which transertion is implicated has led to it being proposed as a powerful force in maintaining the nucleoid in expanded state, in the integrative sensing of metabolic activity and in coupling chromosome segregation to cell division (Norris and Amar, 2012; Fishov and Norris, 2012b).

cells by fluorescent microscopy. Left: image of an *E. coli* cell division mutant *pbpB*(ts) transferred to the non-permissive temperature (time zero, bacterium at the top), double-stained with DAPI (top half of image) and FM4-64 (bottom half of image) for visualization of nucleoids and membrane, respectively; adapted from Fishov and Woldringh (1999), © John Wiley and (DAPI, blue line; FM4-64, red). (B) The relationship between proteolipid domains, transertion, and clusters of genes either "looped out" from or buried within the nucleoid. n1, n2, nascent proteins; m, mRNA; r, ribosomes; p, RNA polymerase; G, gene clusters; red arrow indicates division site; adapted from Norris and Madsen (1995), © Elsevier.

It has long been evident that bacteria are highly structured (for references see Norris et al., 1994). It is now emerging that this structuring is in the form of *hyperstructures* which are large, physically linked, assemblies of macromolecules that serve specific functions (Norris et al., 2007a). Transertion plays a central role in the formation of one class of hyperstructures and also in the dialog between these hyperstructures that determines the behavior of the cell itself. Here, we review membrane heterogeneity focussing on transertion hyperstructures and their relationships with the processes of nutrient sensing, membrane synthesis, cytoskeletal dynamics, DNA replication, chromosome segregation, and cell division.

## Lipid Domains

### Cardiolipin and Other Anionic Phospholipid Domains

Although the lipids present in the inner membrane of *Escherichia coli* will spontaneously separate into domains *in vitro* (Zerrouk et al., 2008), until fairly recently, the lipid molecules in the membranes of bacterial cells were assumed to be homogeneously distributed, since the fluidity of biological membranes had become generally accepted following the publication of the fluid mosaic model (Singer and Nicolson, 1972). However, cell membranes must be laterally polarized to produce specific environments for certain membrane proteins, in particular, chemoreceptor proteins and proteins that promote polymerization of actin in eukaryotic hosts, as well as proteins involved in cell division at mid-cell and at asymmetric positions (Shapiro et al., 2002). Microscopic visualization of membrane lipids in cells has reinforced the view that bacterial membranes do possess structural heterogeneity. Uneven distribution of fluorescent lipophilic dyes and selective staining of septal regions has been observed in mycobacteria and an uneven distribution of fluorescence has been observed in lipophilic dye (FM4- 64)-stained *E. coli* cells (Christensen et al., 1999; Fishov and Woldringh, 1999; **Figure 1**).

Unequivocal visualization of cardiolipin (CL) domains in the septal region and in the poles in the membranes of *E. coli* cells has been accomplished by means of the CL-specific fluorescent dye 10-*N*-nonyl-3,6-bis(dimethylamino)acridine (10- *N*-nonyl-acridine orange: NAO; Mileykovskaya and Dowhan, 2000, 2009; Mileykovskaya, 2007; **Figures 2A–C**). Time-lapse microscopy of *E. coli* cells showed positioning of NAO stained domains at nascent division sites and their gradual development into septal domains (Mileykovskaya and Dowhan, 2009; Mileykovskaya and Margolin, 2012; **Figure 2D**). NAO only binds to anionic phospholipids owing to an interaction between its quaternary amine and the phosphate residue of phospholipids and an intercalation of the hydrophobic acridine moiety into the membrane bilayer (Petit et al., 1992). The stoichiometry between NAO and monoacidic phospholipids is 1:1 and the emitted fluorescence is green whilst with CL, which contains two phosphate groups per molecule, NAO forms a dimer and the emitted fluorescence of the CL complex shifts to red due to the metachromatic effect of acridine molecules (Petit et al., 1994; Mileykovskaya et al., 2001; **Figure 2G**). CL domains have also been observed in *Bacillus subtilis* cells, both during exponential growth in the septal region and the poles and during sporulation in the engulfment and the forespore membranes (Kawai et al., 2004, 2006; **Figures 2E,F**). Note that in *B. subtilis* cells PE is also localized in polar and septal membranes (Nishibori et al., 2005). The CL localization to the polar membranes in *E. coli* is consistent with the increase in the content of CL in the membranes of minicells, which are rich in polar membranes (Koppelman et al., 2001).

An *E. colipgsA* mutant lacks phosphatidylglycerophosphate synthase, which catalyzes the committed step of biosynthesis of the major acidic phospholipids, and thus lacks phosphatidylglycerol (PG) and CL (both < 0.01% of total phospholipids); this mutant, which is viable if it also has an *lpp* mutation, accumulates the anionic biosynthetic precursors, phosphatidic acid (PA) and CDP-diacylglycerol (4.0 and 3.2%, respectively; Kikuchi et al., 2000; Shiba et al., 2004). PG is required for modification of prolipoprotein (the precursor of Braun's lipoprotein, the product of *lpp*); cells lacking PG in an *lpp*+ background are not viable because the accumulation of unmodified prolipoprotein in the inner membrane causes a tight membrane fusion (Suzuki et al., 2002). The viability of a *pgsA lpp*− mutant suggests that the anionic phospholipids can substitute for one another in essential biological functions that include (1) the initiation of DNA replication, which depends on the rejuvenation of the DnaA protein by CL, and (2) the selection of the division site, which depends on the inhibition of inappropriate FtsZ polymerization in the polar regions by a complex, MinCD, with a high affinity for anionic phospholipids (Matsumoto, 2001; Szeto et al., 2002; Mileykovskaya et al., 2003; Norris et al., 2004; Mileykovskaya and Dowhan, 2005; Mazor et al., 2008; Vecchiarelli et al., 2014). In the *pgsA* mutant cells, *N*-acyl-phosphatidylethanolamine (*N*-acyl-PE) and PA have been localized to polar and septal membrane domains (Mileykovskaya et al., 2009), indicating that these normally minor anionic phospholipids are segregated, like CL, into similar anionic lipid domains. Thus, *E. coli* has a mechanism for preferential segregation of anionic phospholipids to the polar and septal regions. One possible segregation mechanism depends on the shape of lipid molecules. 'Cone-shaped' lipid molecules, in which the cross-sectional area of the polar head group is less than the cross-sectional area of the hydrophobic domain, prefer regions of membranes that have a negative curvature; hence CL and PA are concentrated in negatively curved regions of the inner leaflet of the bacterial membranes (Cullis et al., 1983; Gennis, 1989; Seddon, 1990; Renner and Weibel, 2011; Jouhet, 2013; Renner et al., 2013). Using a mathematical model, this sensing of membrane curvature – and consequent positioning in the poles – has been attributed to stable, conical clusters of CL molecules forming due to the membrane being pinned to the cell wall, which is itself the result of the balance between the osmotic-pressure difference across the membrane and the inward pressure of the cell wall (Huang et al., 2006; Huang and Ramamurthi, 2010; **Figure 2F**).

A recent report from Weibel's lab has claimed that there is no clear preference of NAO for binding to CL compared to PG and other anionic phospholipids, and that NAO produces an intense red-shifted fluorescence emission with PG, PA, and phosphatidylserine (PS) that is comparable to that with CL (Oliver et al., 2014). They suggested that not only CL but also PG is concentrated in the polar regions of *E. coli* cell membranes. However, they used much higher concentrations of NAO than previous authors used for bacterial staining (Mileykovskaya and Dowhan, 2000; Kawai et al., 2004; Romantsov et al., 2007, 2008; Rosch et al., 2007; Lobasso et al., 2009, 2013; Maloney et al., 2011; Muchová et al., 2011; Kicia et al., 2012; Tran et al., 2013). Moreover, PG microdomains, in contrast to CL domains, cannot sense membrane curvature, since the cross-sectional area of PG molecule is cylindrical and lamellar structures are preferred (Cullis et al., 1983; Gennis, 1989; Seddon, 1990). As Weibel's lab had already demonstrated that the location of NAO-stained domains depends directly on membrane curvature (Renner and Weibel, 2011; Renner et al., 2013), this group did not propose a specific physical mechanism responsible for localizing anionic phospholipid domains stained by NAO (Oliver et al., 2014). In *B. subtilis*, phospholipid-specific dyes including FM4-64 are localized in a helical pattern extending along the long axis (Barák et al., 2008). Helical FM4-64 domains are missing in cells depleted of MurG, an enzyme involved in peptidoglycan synthesis, indicating a link between the helical

FIGURE 2 | Anionic phospholipid domains in *E. coli* and *B. subtilis* cells revealed by NAO staining. (A) Fluorescence microscopy of cardiolipin (CL)-containing liposomes stained with NAO. Liposomes composed of CL and phosphatidylcholine (60 mol%/40 mol%). Excitation: 490 nm, emission: 617 nm (a) and 528 (b), DIC (c); modified from Mileykovskaya (2007), © John Wiley and Sons. (B) Deconvolved images of an optical section of *E. coli* W3899 stained with NAO. Cells were grown in LB media in the presence of 200 nM NAO to OD600 1.5, and then immobilized on a microscope slide cover glass with poly-L-lysine; modified from Mileykovskaya and Dowhan (2009), © Elsevier. Excitation is 490 nm and emission is 617 nm. (C) *E. coli* cells stained with NAO (red) for CL (a) and DAPI (blue) for the nucleoid (b). Overlay of images (c) demonstrates localization of CL at the cell poles and at the division site; scale bar, 2.5 μM; modified from Mileykovskaya and Dowhan (2000), © American Society for Microbiology. (D) The dynamics of CL domain formation during the cell cycle of growing *E. coli*. Arrows of the same color indicate the progression of the corresponding NAO-stained structures with time; adapted from Mileykovskaya and Dowhan (2009). (E) Staining of wild type *B. subtilis* cells with NAO. Wild type *B. subtilis* 168 cells were harvested in exponential growth and sporulation phase (at T2 and T4) and stained with 100 nM NAO for 20 min. Fluorescence images of exponential growth (a) and sporulation phase cells at T2 (b) and T4 (c). An arrow indicates a sharp fluorescence band in the center of the cell. Two fluorescence dot

structures in the cell center are indicated by a pair of arrowheads. Regions of NAO stained nascent poles in the cells that are separating are indicated with a pair of arrows. Panels are adapted from Kawai et al. (2004), © American Society for Microbiology. (F) Osmotic pressure and turgor-induced localization of lipid clusters during cell division and sporulation. During exponential growth, high-intrinsic-curvature clusters of CL localize to the poles of the inner leaflet [green curve (a) or blue clusters in (b)], driven by differences in membrane curvature. A lower osmotic-pressure differential across the septal/forespore-engulfing membrane [green area, (c)/(d), respectively] induces relocalization of the lipid clusters to the septal membrane; (d) CL clusters migrate along the continuous leaflet consisting of the inner leaflet of the mother cell and the outer leaflet of the forespore-engulfing membrane to localize around the spore due to low osmotic-pressure differential (green circle); modified from Huang et al. (2006) and Huang and Ramamurthi (2010), © John Wiley and Sons with permission. (G) The proposed arrangement of CL in the presence of NAO. A top view of the bilayer in which the hexagonal array of large circles represents the fatty acid chains is shown. The small internal circles containing P represent the phosphate groups, hydrogen-bonded tightly by the hydroxyl of the connecting glycerol, above the two central circles of the four fatty acid chains of CL (red). This tight array provides room for the NAO molecules (green) to stack in between the rows of CL head groups; adapted from Mileykovskaya et al. (2001), © Elsevier.

domains and peptidoglycan synthesis (Muchová et al., 2011). As the helical structures are absent from the cells with repressed *pgsA* expression (in which the levels of PG and CL are much reduced; Hashimoto et al., 2009), the dyes are considered to be associated with the anionic phospholipids (Barák et al., 2008). As the CL-specific NAO dye is located in the septal regions and the poles of *B. subtilis* cells (Kawai et al., 2004), the principal component of the FM4-64 helical structures is likely to be PG (Barák et al., 2008). It has, however, been suggested that experiments showing changes in FM 4-64 staining of *B. subtilis* cells with repressed *pgsA* expression cannot serve as unambiguous evidence of FM 4-64 specificity for PG since, first, changes in the distribution of FM 4-64 resulted from the dissipation of membrane potential by the uncoupler CCCP and, second, the membrane potential was much lower in *B. subtilis* cells with repressed *pgsA* expression (Strahl et al., 2014). In *E. coli* no discernible helical structures were observed with FM4-64 staining (Barák et al., 2008) as reported previously (Fishov and Woldringh, 1999).

Microdomains that are functionally similar to the lipid rafts of eukaryotic cells have recently been found in *B. subtilis* membranes (Lopez and Kolter, 2010). These microdomains contain homologues of eukaryotic flotillins, YuaG and YqfA, referred to as FloT and FloA, respectively (see Heterogeneous Distribution of Envelope and Envelope-Associated Proteins). The distribution of these microdomains was in a punctuated pattern along the cytoplasmic membrane (Donovan and Bramkamp, 2009; Lopez and Kolter, 2010), which differs from the patterns of FM4-64-stained helices and of CL-domains in the septal region and the poles. YisP is involved in biofilm formation in *B. subtilis* and has been predicted to produce C30 isoprenoids; the enzyme acts as a phosphatase, catalyzing formation of farnesol from farnesyl diphosphate (Feng et al., 2014). Inhibition of formation of these microdomains using zaragozic acid impaired biofilm formation and protein secretion but not cell viability and it has been suggested that polyisoprenoids are constituents of the microdomains (Lopez and Kolter, 2010; Feng et al., 2014; Bramkamp and Lopez, 2015).

### Lipid Synthases and Membrane Domains

The septal and polar locations of specific phospholipids may be related to the subcellular location of the enzymes involved in their synthesis. In *E. coli*, a relationship between the location of CL (Mileykovskaya and Dowhan, 2009; Mileykovskaya and Margolin, 2012; see Cardiolipin and Other Anionic Phospholipid Domains) and that of CL synthase (ClsA) and phosphatidylglycerolphosphate synthase (PgsA; Tan et al., 2012; Dowhan, 2013) has yet to be reported. In *B. subtilis*, green fluorescent protein (GFP) fusions to ClsA and PssA, a PS synthase that catalyzes the committed step of biosynthesis of PE, are located in the septal membranes even when the corresponding genes are expressed at a low level from their natural promoters (Nishibori et al., 2005). Thus, ClsA is probably concentrated in the septal membranes under natural conditions thereby playing an important role in the septal localization of CL in *B. subtilis* (**Figure 3**). Localizing the enzymes involved in the synthesis of other lipids has yielded unexpected, interesting results. GFP fusions to several phospholipid synthases were localized to the septum in a thick, bright fluorescence band. These synthases include PgsA, PssA, Psd, which converts PS into PE, CdsA, which produces CDP-diacylglycerol, MprF, which transfers lysine to PG to produce lysylPG, and UgtP, which is responsible for glucolipid synthesis. The dot-pair distribution of UgtP corresponds to its role in regulating FtsZ assembly and hence differs from the distribution of the phospholipid synthases

#### FIGURE 3 | Septal localization of lipid synthases in *B. subtilis* cells. Typical florescence images from cells harboring *gfp* fusions in the *amyE* locus cultivated in sporulation medium (DSM) up to late logarithmic growth phase are shown. Green fluorescence from the GFP-fusions was detected by using a standard GFP(R)-BP filter unit. For the name of the enzymes and their

catalytic activities refer to the figure of the biosynthetic pathway (Figure 5B). Panels are from Figures 4 and 5 of Nishibori et al. (2005), © American Society for Microbiology. GFP-fusions of ClsA and PssA are located in the septal membranes even when the corresponding genes are expressed from their natural promoters (see Figure 4 of Nishibori et al., 2005).

(see Sensing). The locations of these enzymes thus differ from the cytoplasmic location of GpsA, which catalyzes the production of G3P (Nishibori et al., 2005; **Figure 3**), and the uniformly distributed locations of the membrane proteins AtpC, a subunit of ATP synthase, and SecY (Matsumoto et al., 2012). It has also been shown that the enzymes for lipoteichoic acid (LTA) synthesis, LtaS and YfnI, are septally localized (Schirner et al., 2009; Matsuoka et al., 2011b), but DG kinase (DgkB), which converts DG that is produced in LTA synthesis into PA, is not localized (Matsuoka et al., 2011b).

How are the lipid synthases targeted to the septal membranes? These enzymes presumably have specific regions responsible either directly for their localization or indirectly via interaction with certain cell division proteins. Selective inactivation of putative targeting regions has shown that ClsA has regions of amphipathic α-helices at its COOH-terminus that are responsible for its septal localization (Kusaka et al., manuscript in preparation). PgsA has no such amphipathic α-helices at its COOH-terminus, suggesting that the system for its septal localization is different from that of ClsA. Many other proteins with amphipathic α-helices, including MinD (see Lipid–Protein Interactions), have membrane binding properties that depend on electrostatic interactions, *via* their net positive charges, and on hydrophobicity. In addition, the catalytic domains (or consensus sequences) of *B. subtilis* PssA (Saha et al., 1996b) and *E. coli* and *B. subtilis* PgsA (Usui et al., 1994; Matsumoto, 1997) have an amphipathic α-helix structure which allows them to access their hydrophilic substrates, serine and G3P, in the cytoplasm (Matsumoto et al., 2012). The septal location of the phospholipid synthases in *B. subtilis* cells implies that most phospholipids are synthesized at the septal membranes, and, given the septal location of PE and CL, it is likely that these newly synthesized lipids are somehow prevented from diffusing into the lateral membranes (Matsumoto et al., 2006).

### Chemical Basis for Generation of Lipid Domains

What then is the chemical basis for the generation of lipid domains? How do the lipid molecules form domains in membranes that are fluid? Both lipid–lipid interactions and lipid–membrane protein interactions are suspected to induce the formation of microdomains, which comprise at most some tens of molecules of a specific lipid (for a review, see Edidin, 1997). The following properties of phospholipid molecules may account for the formation of microdomains through lipid–lipid interactions.

Cardiolipin, which is a diphosphatidylglycerol, has a unique head group, with a tightly locked, surprisingly small configuration and only one negative charge (Haines and Dencher, 2002); this result contradicts the previous view reported in textbooks. The two phosphates in the head group trap a proton and are locked in a bicyclic array held together by the hydroxyl residue of the glycerol which connects the two halves. This small polar head group makes for a tighter packing of hydrophobic acyl chains between CL molecules by van der Waals force interactions than is found in lipids with larger polar head groups. It has also been suggested (Haines and Dencher, 2002) that interaction between adjacent head-groups creates a compact array of CL molecules, which becomes manifest in the presence of associated NAO arrays (Mileykovskaya et al., 2001).

Recently, a microtechnology for manipulating bacterial membrane curvature and quantitatively measuring its effect on the localization of CL in spheroplasts has been developed (Renner and Weibel, 2011). CL domains were localized to regions of high intrinsic negative curvature and MinD was found to be associated to the region of large negative curvature (see below). This localization of CL to membranes of highly negative curvature is ascribable to its cone-like molecular configuration (Gennis, 1989; Jouhet, 2013; see Cardiolipin and Other Anionic Phospholipid Domains). Polar localization of PA domains found in *E. colipgsA* cells lacking both CL and PG can also be ascribable to the cone-like configuration of PA molecule (Norris et al., 2002; Mileykovskaya et al., 2009).

Polar head groups of PG molecules interact by an extensive network of hydrogen bonds, ionic bonds, and coordination bonds between glycerol hydroxyls and the unesterified phosphate oxygen both in the anhydrous crystal and the hydrated gel state (Boggs, 1987; Pascher et al., 1987; Seddon, 1990). In fact, PG is probably segregated into distinct domains, in both *B. subtilis* and *E. coli* membranes, according to studies using pyrene-labeled phospholipids (Vanounou et al., 2003). This extensive and tight network of PG molecules may cause exclusion of CL molecules to produce patches of CL in the membranes (Mileykovskaya et al., 2001), since the small and tightly locked head group of CL cannot interact with PG.

The head group of the PE molecule has both a cationic amine residue and an anionic phosphate residue. Each amine and unesterified phosphate oxygen can participate in short distance intermolecular hydrogen bonds. The ethanolamine groups thus form a linkage between phosphorus groups of adjacent PE molecules and the tendency to be hydrated must be less than that of phosphatidylcholine, so producing a very compact, rigid head-group network of PE molecules (Elder et al., 1977; Hauser et al., 1981; Boggs, 1987; Seddon, 1990), and giving PE substantially higher *T*<sup>m</sup> values than phosphatidylcholine which has an identical acyl chain structure. This compact head-group network of PE may well suffice to explain the formation of PE microdomains. Interactions with certain membrane proteins might then modify the lipid organization and further stabilize the head-group network, as suggested (Edidin, 1997). Polar and septal localization of PE in *B. subtilis* cells (Nishibori et al., 2005) is likely ascribable to its cone-like molecular shape (see Cardiolipin and Other Anionic Phospholipid Domains). For details refer to the previous review (Matsumoto et al., 2006).

The potential role for polyamines in lipid domain formation is largely unexplored although it has been suggested that polyamine binding to the acidic groups of phospholipids in membranes could lead to clusters forming (Schuber, 1989). Ionic interactions between polyamines and acidic phospholipids are strongest between the polyamine with the highest positive charges and the phospholipid with the highest content of negative groups (Yung and Green, 1986). Interaction with polyamines is believed to reduce the repulsive forces between negatively charged membrane components by bridging proteins and lipids and by shielding the surface charges (Schuber, 1989). The binding of ions such as calcium to anionic phospholipids can also result in domain formation (Haverstick and Glaser, 1987) and for references see Cannon et al. (2003), but, again, this is relatively unexplored.

### Transertion and Membrane Heterogeneity

### Heterogeneous Distribution of Envelope and Envelope-Associated Proteins

A heterogeneous distribution of proteins in the cell envelope of bacteria, despite their size, has been widely demonstrated (see reviews, Shapiro et al., 2002; Fishov and Norris, 2012b; Govindarajan et al., 2012). During the course of investigation of specific biological functions and processes, this heterogeneity has been revealed mainly by visualization with GFP-fusions and immunofluorescence to take the form of polar, patchy and helix-like distributions in the membranes. Examples of proteins localized to the septal regions include lipid synthases in *B. subtilis* (Nishibori et al., 2005; Matsumoto et al., 2012; Takada et al., 2014; described in Lipid Synthases and Membrane Domains). Reciprocally, ATP synthase and succinate dehydrogenase are at low levels in (or absent from) the mid-cell region at the onset of cell division in *B. subtilis*, which may reflect an association with lipid domains elsewhere that are rich in PG or other lipids rather than with lipid domains in the mid-cell region (i.e., the division site) where these lipids may be at low levels (Meredith et al., 2008). Recently, FloT and FloA, homologues of eukaryotic flotillin proteins found exclusively in lipid rafts along with proteins involved in signaling and transport have been localized to discrete microdomains in the membrane of *B. subtilis* (see Cardiolipin and Other Anionic Phospholipid Domains); significantly, these microdomains, which are likely to be present in many other bacteria, also contain other proteins involved in signal transduction and cell–cell communication such as the sensor kinase, KinC, and protein secretion such as SecY in *B. subtilis* (Donovan and Bramkamp, 2009; Lopez and Kolter, 2010; Bach and Bramkamp, 2013; Bramkamp and Lopez, 2015). Flotillins are believed to play a large part in maintaining the overall physical heterogeneity of the membrane since, in their absence, lipid-ordered domains coalesce (Bach and Bramkamp, 2013; Bramkamp and Lopez, 2015). Cytoplasmic membrane proteins located in the polar regions of *E. coli* cells include ProP, LacY, and MscS (Romantsov et al., 2010) and the MCPs (methyl-accepting chemotaxis proteins; Alley et al., 1992; Sourjik and Armitage, 2010) whilst proteins located at the sites of cell constriction in *E. coli* include the components of *trans-*envelope Tol-Pal complex, TolA, TolQ, and TolR (in the cytoplasmic membrane), the peptidoglycanassociated lipoprotein Pal, (anchored to the outer membrane), and TolB (a soluble periplasmic protein; Gerding et al., 2007).

Two systematic approaches have been taken to find proteins that are preferentially located in the poles. Using twodimensional gel electrophoresis, Lai et al. (2004) identified proteins preferentially located in *E. coli* minicells and created a catalog of polar proteins; these included the inner membrane proteins MCPs, AtpA, AtpB, YiaF, and AcrA, the membraneassociated protein FtsX, and the outer membrane proteins YbhC, OmpW, Tsx, Pal, FadL, OmpT, and BtuB. In a complementary, cataloging approach, Li and Young (2012) used FLAG peptidetagged Tar, a known polar MCP, to isolate by affinity capture those inner membrane vesicles that originated from the poles. These vesicles were enriched in over 30 proteins. Most were in or associated with the inner membrane and included Aer, PBP1B, TnaA, DcuA, PutP, TrxA, FliP, AccD, CpxA, FliC, RpsD, YcbC, GlnP, GroEL, MlaF, NarZ, YqjD, YniB, MCPs, CheA, YijP, PBP1A, SppA, LepB, NupC, and YiaF, although a few exceptions, TolC, Pal, and BamA, were in the outer membrane. In following up the latter catalog by fusing proteins to GFP and expressing the gene from their own promoters, Aer, TnaA, GroES (rather than GroEL), YqjD, and YiaF were found to form polar foci that were distinct from inclusion bodies, thereby suggesting that these proteins are located in polar membranes *in vivo*. Moreover, the MCPs and Pal figure in the polar membrane class in both cataloges, thus lending some confidence to the results. Extensions of approaches like these should identify the remaining proteins with polar preference using the full list of *E. coli* proteins, which includes as many as 1,133 predicted integral inner membrane proteins (Bernsel and Daley, 2009) and 503 peripheral, inner membrane-associated ones (Papanastasiou et al., 2013).

GFP-fusions and immunofluorescence have provided many examples of specific proteins in polar and helix-like distributions in the membranes of *E. coli* and *B. subtilis* cells; these proteins include MreB (as well as homologues MreBH and Mbl) and the Sec translocon (for the list and references see Table 1 in Fishov and Norris, 2012a). These results have, however, been questioned due to recent findings showing that artefactual distributions can be generated by high levels of expression. The translocon in *E. coli* includes the SecYEG translocon which can associate with SecDF–YajC–YidC and SecA to enable protein transport in a reaction that is stimulated by CL (Gold et al., 2010; Schulze et al., 2014). The GFP-fusions of SecA and SecY over-expressed using 1% xylose are helically distributed in *B. subtilis* (Campo et al., 2004) whilst expression from the native promoter Psec*secA* gives a peripherally homogeneous and septal distribution of the fusion products in conditions in which careful high-level expression controls show helices (Carballido-Lopez et al., 2006- Supplementary Figure S6; Halbedel et al., 2014). This peripherally homogeneous distribution of SecA in *B. subtilis* is different from that of *E. coli* GFP-SecE which has a patchy subcellular distribution (Shiomi et al., 2006). Localization of FlAsH-tagged SecY and SecE in *E. coli* also shows quite different patterns from that of *B. subtilis* SecA, with helix-like (patchy), polar dots and a diffuse distribution in cells under conditions of no induction (Gold et al., 2010); that said, the cells still have higher levels of tagged products from the high copy number plasmid pBAD22, even in the absence of the inducer, compared with those from their native promoter on the chromosome, i.e., single copy. The translocon distributions in *B. subtilis* and *E. coli* also differ from that of the *Streptococcus pyogenes* translocon, which forms a single microdomain (Rosch and Caparon, 2005; Rosch et al., 2007) and that of *S. pneumonia* D39, expressed from the native chromosomal loci, which localizes dynamically to different places (Tsui et al., 2011). GFP-fusions of MreB have been shown to move, as fragmented discrete patches, perpendicular to the long axis of *B. subtilis* cells using highly sensitive time-lapse imaging (Dominguez-Escobar et al., 2011; Garner et al., 2011). This movement of MreB did not, as one would have expected, follow a helical pattern. Recent reports and reviews suggest that the distribution pattern reported for *E. coli* MreB mainly resembles that reported for *B. subtilis* MreB: a punctuated pattern with pairs of dots or small bands generally described as helical (van Teeffelen et al., 2011; Chastanet and Carballido-Lopez, 2012; Margolin, 2012; Errington, 2015). Indeed, MreB proteins form elongated filamentous structures when over-produced or when observed in late phases of growth. In virtually all older reports describing MreB localization*,* inducible, mostly overexpressed, GFP fusions were used, and observations were often made during the late exponential stage, when the structures were easier to visualize. This could be the case for other fusion proteins showing polar and helix-like distributions under highly expressed conditions. Thus, it is difficult at present to decide how many of the proteins do indeed form long-range, helical structures at the membrane, as noted by Margolin (2012).

#### Lipid–Protein Interactions

The last stage of transertion (see below) involves interaction of nascent proteins with the polar head groups of lipids followed by insertion into non-polar acyl-chain layer of the membrane. Proteins with a preference for a specific polar head group interact to form a specific domain in the membrane. Integral inner membrane proteins with specific lipid preferences could therefore be located in specific membrane domains (for the list of proteins see Table 2 in the review of Fishov and Norris, 2012a). In addition, peripheral membrane proteins that associate weakly with specific lipids could help form specific domains and, if these proteins bind other proteins, could bring them into the domain too. Among the peripheral membrane proteins, there are amphitropic proteins that have two apparent locations: one form of the protein is in the aqueous compartment of the cytosol whilst the other form is weakly associated with the cell membrane. Amphitropic proteins can be classified into three major categories (Johnson and Cornell, 1999), based on the way they associate with membranes. The first class (A) contains motifs with binding pockets for a lipid monomer, called lipid clamps. The second class (B) contains lipid covalent anchors embedded in the lipid bilayer; this class includes lipoproteins such as the major outer membrane lipoprotein (Braun's lipoprotein), LolB, components of Bam complexes and RcsF with an *N*acyl chain and a diacylglyceryl moiety at the NH2-terminus (Sankaran and Wu, 1994; Matsumoto, 2001; Suzuki et al., 2002; Okuda and Tokuda, 2011; Shiba et al., 2012; Konovalova et al., 2014). *E. coli* has 96 lipoproteins, 58% of which have completely unknown functions (Brokx et al., 2004). The third class (C) of amphitropic proteins contains amphipathic α-helices, which bind to a membrane by partitioning into the membrane bilayer such that the hydrophobic face of the protein is sequestered away from water, yet its polar face can contact the aqueous phase (Johnson and Cornell, 1999). The helix axis lies parallel to the membrane surface. The weak (reversible) interaction of the proteins with amphipathic α-helices plays an important role in regulation of their functions. Representatives of this class include a family of prokaryotic cytoskeletal proteins such as MreB, MinD, and FtsA, having an amphipathic α-helix at their NH2− or COOH-terminus (Szeto et al., 2003; Pichoff and Lutkenhaus, 2005; Salje et al., 2011; Szwedziak et al., 2012; **Figure 4**). Membrane anchoring with amphipathic helices has yet to be reported for any eukaryotic filament (Salje et al., 2011). It has been demonstrated recently that the nucleoid occlusion protein of *B. subtilis*, Noc, also associates with the cell membrane via an NH2-terminal amphipathic α-helix. It occludes assembly of the division machinery by simultaneous binding to DNA and the membrane (Adams et al., 2015). PgsA, *B. subtilis* PssA (Matsumoto, 1997), and the well-understood mammalian CTP: phosphocholine cytidylyltransferase (CCT) have an amphipathic α-helical structure in the middle of the protein (Cornell and Taneva, 2006). The amphipathic α-helices that were found in DnaA (Garner and Crook, 1996; Makise et al., 2001) are not involved in membrane association; instead, the results of membrane retention experiment obtained with various fragments of the protein indicated that the association is through a concerted interaction of distant residues forming a surface (Regev et al., 2012).

MreB is one of the key components of the bacterial cytoskeleton that lies just underneath the membrane and organizes the cell wall synthesis machinery. In *E. coli*, MreB has an amphipathic α-helix (an 11 amino acid residue sequence which includes two methionine, two phenylalanine, one leucine on one face of the helical wheel and three basic residues on the other face) at its NH2-terminus that is responsible for its location underneath the membrane. In gram-positive bacteria and the thermotrophic archaeum *Thermotoga maritima*, the counterpart of MreB has no such helix (Salje et al., 2011) and the membrane binding of *T. maritima* MreB is mediated by a small insertion loop that contains leucine and phenylalanine (residue 93 and 94), two other hydrophobic residues, the start methionine and a leucine (residues 1 and 2; Salje et al., 2011). MinD, plays a key role in division site selection by protecting the poles from aberrant positioning of FtsZ ring, and has a small amphipathic α-helix of 8–12 residues, termed the membrane targeting sequence or MTS, at its COOH-terminus that is conserved across eubacteria, archaea, and chloroplasts; this MTS is essential for association with anionic phospholipid-enriched membrane from which, in the case of *E. coli*, it can be easily detached by MinE at the appropriate time in the Min oscillation cycle (Raskin and de Boer, 1999; Szeto et al., 2002, 2003; Mileykovskaya et al., 2003; Vecchiarelli et al., 2014). The actin-like protein FtsA has an MTS that comprises a conserved COOH-terminal amphipathic α-helix of 13–14 residues (Pichoff and Lutkenhaus, 2005); FtsA polymerizes to form an "FtsA ring." The tubulin-like protein FtsZ is tethered to FtsA through a COOH-terminal tail of 16 residues to polymerize to form an FtsZ ring underneath the A-ring (Szwedziak et al., 2012; **Figure 4**). Work on liposome division has shown the importance of the MTS in division. ATP

is needed not only for the polymerization of FtsA and also for its attachment via its MTS to lipid monolayers and to vesicle membranes; this polymerization of FtsA caused vesicle shrinkage, consistent with the protein facilitating division both indirectly by interacting with FtsZ and directly by altering the membrane (Krupka et al., 2014). A fusion of FtsZ-YFP-mts was constructed with an MTS at its COOH-terminus to tether it directly to the membrane; this allowed Z-rings to assemble in multi-lamellar tubular liposomes and to generate a constriction force in the presence of GTP without the need for any other protein (Osawa et al., 2008). Uni-lamellar liposomes incorporating FtsA and FtsZ-YFP produced more natural Z-rings, which constricted liposomes and in some cases appeared to complete the division (Osawa and Erickson, 2013). The co-polymerization of FtsZ and FtsA is proposed to lead to bending, curvature and membrane constriction because the subunit repeat lengths of FtsZ and FtsA are different, being roughly 4 and 5 nm, respectively (Szwedziak et al., 2014; **Figure 4**).

### Membrane Curvature and Protein Location

In rod-shaped bacterial cells, negatively curved membranes characterize the poles and parts of the developing septum. In *B. subtilis*, it has been suggested that this curvature is sensed by DivIVA (Lenarcic et al., 2009; Ramamurthi and Losick, 2009), which is located at the septum and at mature poles and which is responsible for the polar location of the division inhibitor MinC/MinD via MinJ, a system different from the Min system of *E. coli* (Marston et al., 1998; Bramkamp et al., 2008; Patrick and Kearns, 2008). In cells with engineered curvature, DivIVA is indeed located preferentially in regions of high negative curvature (Renner et al., 2013; **Figure 2F**). The location of DivIVA depends on SecA although DivIVA does not contain a signal sequence (Halbedel et al., 2014). Instead, SecA is considered to act as a chaperone in the folding of DivIVA, which has essentially an α-helical structure with several coiled-coil helices that contribute to the curvature of the DivIVA tetramer (Halbedel et al., 2014). The crystal structure of this tetramer resembles the crescent shape of eukaryotic BAR domains which bind to curved membranes and also introduce curvature (Lemmon, 2008; Oliva et al., 2010; Mim and Unger, 2012). Enzyme I (EI), which is a part of the phosphoenolpyruvate-phosphotransferase system (PTS) responsible for the sensing and uptake of many extracellular sugars, is located in the poles of *E. coli* and *B. subtilis* due, it is proposed, to its affinity for negatively curved membrane, an affinity shared by DivIVA (Govindarajan et al., 2013). Conversely, SpoVM, a small peripheral membrane protein with an amphipathic α-helix, associates with the positively curved (convex) membrane surfaces of the forespore to form the spore coat complex with SpoVA, in the mother cell during sporulation (Ramamurthi, 2010; **Figure 2F**). Recently, it has been suggested that the actual amphipathic α-helix region of SpoVM (26 aa) is atypical; this region is short with only 13 residues (from 11 to 23**)**, contains only one positively charged residue (Arg) and three Gly residues, and inserts deeply into the membrane, unlike many amphipathic α-helix molecules that only insert shallowly into the membrane (Gill et al., 2015). The authors hypothesize that deep insertion of SpoVM into membrane, which involves extensive interactions with acyl chains to sense packing differences in differently curved membranes and which may involve cooperative interactions with other SpoVM proteins, drives its preferential localization onto slightly convex membranes surface such as the outer surface of the forespore.

### Transertion

In the transertion hypothesis, it is proposed that the coupled transcription-translation of genes encoding membrane proteins (1) structures the membranes, both physically and chemically, and (2) positions these genes close to the membrane (Norris and Madsen, 1995; Woldringh, 2002; **Figure 1**). Despite early evidence in its favor (Binenbaum et al., 1999), this hypothesis lacked further evidence to support it for long time. Recently, Libby et al. (2012) have shown that the synthesis of membrane proteins expression affects the position of chromosome loci in *E. coli* cells. They observed that two genetic loci*, lacY* and *tetA*, which encode membrane proteins, were rapidly shifted toward the membrane upon induction whilst the chromosomal locus for a cytoplasmic protein was not shifted; in addition, antibiotics that block transcription and translation prevented these shifts toward the membrane. More recently still, a radial contraction of the *E. coli* nucleoids was observed immediately after the addition of inhibitors of either transcription or translation, again consistent with transertion normally exerting an expanding force on the nucleoid (Bakshi et al., 2014). In the case of inhibition of transcription and hence transertion by Rifampicin, the eventual expansion of the nucleoid has recently been attributed to the penetration of the nucleoid by the ribosomal subunits (Bakshi et al., 2014; Sanamrad et al., 2014). That said, it should be noted that transcription is the essential source of supercoiling (particularly at high growth rates when 50–80% of active RNA polymerases are transcribing *rrn* genes) which is a major cause of nucleoid compaction; hence, inhibition of transcription initiation with Rifampicin results in RNA polymerase run-off, less supercoiling, and nucleoid expansion; this expansion can happen even after nucleoid compaction by chloramphenicol when transertion is absent (for a review, see Jin et al., 2013).

Over 1,100 genes distributed throughout the *E. coli* genome are predicted to encode integral inner membrane proteins (Bernsel and Daley, 2009) and therefore their expression could maintain the chromosome in an expanded and dynamic state, consistent with the transertion hypothesis. Significant numbers of ribosome and RNA polymerase copies have been shown by superresolution imaging to extend from the nucleoid to the cytoplasmic membrane (Bakshi et al., 2012), indicating that transertion exerts a direct radially expanding force on the nucleoid (Woldringh, 2002; Woldringh and Nanninga, 2006). In this study, however, few copies of RNA polymerase were found near the polar caps, indicating that transertion could not occur in the regions away from the nucleoid (polar caps) and therefore it might seem that the radially expanding force on the nucleoid of transertion could not be a direct axially expanding force. Such axial expansion might, however, result from the penetration of the nucleoid by the 30S and 50S subunits released from the 70S ribosomes following treatment with inhibitors of translation (Bakshi et al., 2014; Sanamrad et al., 2014). It should be noted that although less than 15% of *E. coli*'s ribosomes are reported to be available to participate in transertion (Bakshi et al., 2012; Gahlmann and Moerner, 2014), this figure does not necessarily reflect the percentage of the ribosomes that are actually engaged in translation. A higher percentage is likely to be available in *C. crescentus* where genes and mRNA have been found to be close to one another (Llopis et al., 2010).

There are plenty of polar membrane proteins as noted in the previous section (see Heterogeneous Distribution of Envelope and Envelope-Associated Proteins). Transertion is unlikely to be an important factor in directly controlling their location since, firstly, RNA polymerase is relatively scarce near the polar caps and, secondly, the Sec machinery responsible for inserting integral membrane proteins into membranes is not specifically located in the poles in either *B. subtilis* (Carballido-Lopez et al., 2006-Supplementary Figure S6) or *E. coli* (Shiomi et al., 2006; Gold et al., 2010; see Heterogeneous Distribution of Envelope and Envelope-Associated Proteins). Thus, some integral membrane proteins (e.g., the MCPs) are inserted into the cytoplasmic membrane around the nucleoid via the Sec machinery and then freely migrate in the membrane to the polar regions (Shiomi et al., 2006). One possibility is that integral membrane proteins such as the MCPs are excluded from the transertion domains around the chromosome because they have no affinity for the lipids in which those domains are enriched. The reason for this is that transertion may create a problem if it leads to the formation of an array of MCPs of the wrong size in the wrong place. Another, recently proposed, possibility is that polar localization of the diffused MCPs occurs independently of the phospholipid composition of the cytoplasmic membrane and is not be dictated by the curvature of the cell poles, instead, MCPs interact with components of the trans-envelope Tol-Pal complex which restricts the diffusion of MCP arrays (Santos et al., 2014; see Transertion Problems).

In the original transertion hypothesis, the structuring of the membrane and the tethering of genes was restricted to those genes encoding proteins inserted into or through membranes. Although the role of the former class of proteins (i.e., integral membrane proteins) is easy to imagine, the latter class (i.e., secreted and exported proteins) is also of interest. For example, there could be a role in transertion for Braun's lipoprotein if the lipid modification at the NH2-terminus of the lipoprotein were to occur before the synthesis of the rest of the protein. Moreover, the transertion hypothesis could be extended to genes encoding peripheral membrane proteins such as SeqA (Slater et al., 1995; Shakibai et al., 1998) and DnaA in *E. coli* (Regev et al., 2012) and Noc in *B. subtilis* (Adams et al., 2015).

Finally, it has been proposed recently on entropic grounds that transertion enables ribosomal subunits to penetrate the nucleoids and to initiate the more general process of co-transcriptional translation (Bakshi et al., 2014). Co-transcriptional translation would then protect nascent mRNA (Deana and Belasco, 2005; Deneke et al., 2013) and prevent the backtracking of RNA polymerases (McGary and Nudler, 2013), thereby giving transertion a key role in the optimization of transcription and translation (Bakshi et al., 2014). In the context in which two chemically identical chromosomes are in competition for the transcriptional and translational apparatuses, such optimization constitutes a positive feedback that could underpin differentiation (Norris and Madsen, 1995; Norris and Amar, 2012).

### Hyperstructures

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 accumulating evidence for their existence allows them to be classed in several ways (Norris et al., 2007b). In the case of the membrane, the best known include those involved in motility (such as the MCPs and flagella), lactose metabolism, energy generation, and cell division. It has been further proposed that a dialog between hyperstructures determines the phenotype of the cell (Norris et al., 2014a). Here we focus on some of the hyperstructures involved with the membrane.

### Transertion Hyperstructures

The coupled transcription-translation-insertion of membrane proteins (transertion) has been proposed to tether networks of nascent proteins, mRNAs and genes to the cytoplasmic membrane thereby attaching DNA dynamically to the bacterial cell envelope (Norris, 1995; Norris and Madsen, 1995; Woldringh, 2002; **Figure 1**). This means that the high level expression of a gene encoding a membrane protein could lead to the formation of a domain in the membrane enriched by the nascent proteins, possibly specific lipids, and translocon complexes, along with an adjacent, highly structured cytoplasm to give a transertion hyperstructure (Norris, 1995; Norris and Madsen, 1995; Binenbaum et al., 1999; Woldringh, 2002).

The *lac* operon is an obvious candidate for giving rise to a transertion hyperstructure. In *E. coli*, full induction of this operon has been calculated to give 23 RNA polymerases on *lacZ* and 5 or 6 on *lacY* and *lacA* with the ensemble of the nascent, full length and decaying mRNAs being translated by several hundred ribosomes (Kennell and Riezman, 1977). *lacY* encodes the membrane-bound permease with an average of four ribosomes calculated as translating the corresponding full length mRNA (Kennell and Riezman, 1977). LacZ-encoding mRNA has been shown to remain close to the *lacZYA* locus, consistent with both transcription and translation occurring in the same place (Llopis et al., 2010). Expression of the *lac* operon brings the locus closer to the membrane and this movement depends on the presence in the operon of *lacY*, consistent with transertion (Libby et al., 2012). Hence, calculations and experiments suggest that a transertion hyperstructure exists that comprises the *lac* genes dynamically attached to tens of nascent mRNAs and to 100s of ribosomes and the nascent enzymes some of which would be inserted into the membrane. This does not, of course, mean that the Lac proteins once synthesized remain within the transertion hyperstructure. Indeed, a tagged version of LacY – albeit expressed from a plasmid – was localized to the poles (Romantsov et al., 2010).

RNA degradation may also play a part in the dynamics of transertion hyperstructures. Using epifluorescence microscopy and molecular dynamics simulation, RNase E, the backbone of the RNA degradosome, was shown to diffuse over the entire inner membrane of *E. coli* to form, along with other proteins, short-lived hyperstructures (Strahl et al., 2015). The existence of these hyperstructures depends on the presence of the RNA substrate (Strahl et al., 2015) so they may be considered as *functioning-dependent* structures, that is, structures that form due to their activity (Norris et al., 2007a). It should be noted that such separation of the degradosomes from the sites of transcription would favor nascent transcripts being translated by polyribosomes, that is, would favor the coupling of transcription and translation over the separation of these processes (Strahl et al., 2015).

### Transertion Problems

Transertion is not only a likely solution to many of the problems that confront cells but also a source of potential problems. One problem would occur if a transertion hyperstructure were to determine, inappropriately, the size, position or composition of another class of hyperstructure. Consider, for example, the importance in signal transduction of the distribution of the MCPs and related proteins into many small clusters or into one giant cluster (Bray et al., 1998). These distribution confer different sensitivities but this relationship could be lost if transertion were to dominate by creating a large chemosignaling hyperstructure. In *E. coli*, part of the solution may be that the genes that encode chemotaxis proteins are located on the chromosome so that their transertion associates them with another major hyperstructure, that of the flagellum; it is then conceivable that differences in the affinities of the chemotaxis and flagellar proteins for the chemical or physical properties of lipids are responsible for the chemotaxis proteins relocating from the flagellar hyperstructure(s) to the poles (Cabin-Flaman et al., 2005). Consistent with this, Tar-GFP does diffuse away from the sites where this MCP is synthesized to the poles (Shiomi et al., 2006). It has been recently found that this polar localization of the diffused MCPs does not depend on either the phospholipid composition of the cytoplasmic membrane or the curvature of the cell poles; instead, MCPs interact with components of the trans-envelope Tol-Pal complex which restricts the diffusion of MCP arrays (Santos et al., 2014). The Tol-Pal complex is also part of the cell division hyperstructure, to which it is recruited by FtsN so as to play a role in the invagination of the outer membrane during division (Gerding et al., 2007).

A second problem would arise if the lipid preferences of the constituents of the transertion hyperstructure were to menace the planar, bilayer structure of the cytoplasmic membrane. The reality of this danger is evident from the proteo-lipid structures that result from the overproduction of peripheral and integral membrane proteins (Weiner et al., 1984; Arechaga et al., 2000; van den Brink-van der Laan et al., 2003; Eriksson et al., 2009). In mitochondria, the bioenergetic supercomplexes are indeed in specific membrane structures whilst in bacteria such as *E. coli*, fluorescent fusions suggest that the complexes are spatially dispersed in mobile 100–200 nm domains containing 10s–100s of complexes (Erhardt et al., 2014; Llorente-Garcia et al., 2014). A possible solution would be for an abundant structure such as the ATP synthase to have subunits with complementary lipid preferences (Arechaga et al., 2000) since even overproduction of all eight subunits results in morphological changes (von Meyenburg et al., 1984). Extending this argument, one might expect different proteins to have preferences for different lipids, which may help explain why cells have so many different lipids. An alternative or complementary solution would be the creation by a large, dynamic hyperstructure of many mobile domains with an affinity for ATP synthase.

Finally, problems due to the formation of an inappropriate transertion hyperstructure might be avoided by reducing the time during its synthesis in which the nascent protein interacts with the membrane; this might be achieved if, for example, evolution were to have selected for the membrane-interacting sequences (such as amphipathic helices) to be located at the COOH-terminus rather than at the NH2−terminus.

### Sensing

The membrane is the final frontier between the cell and its environment so membrane-based hyperstructures are in the right place to sense and to respond to environmental changes. This task is not trivial. Not only do new mRNAs and proteins have to be made but many existing mRNAs and proteins must be degraded. There are, for example, probably over a 1000 membrane proteins in *E. coli* (Bernsel and Daley, 2009) and a similar number in *B. subtilis* (Hahne et al., 2008; Otto et al., 2010; van Dijl et al., 2012) and the functioning of these proteins must be coordinated; this task is particularly difficult when major changes in the protein composition of the membrane must occur as during changes in growth conditions. On the entry of *B. subtilis* to stationary phase, the levels of the many proteins that increase include those involved in the uptake of glycerol, ribose, lactate, nucleoside, succinate, fumarate and zinc whilst the levels of those that decrease include transporters of malate, Fe3<sup>+</sup> citrate, Fe3<sup>+</sup> hydroxamate, and hydroxymethylthiamine (HMP)/thiamine transport (Otto et al., 2010). In *E. coli* cells supplemented with glucose, which is the preferred carbon and energy source, the transcriptional levels of the many genes that increase include those involved in the import of polyamines, inorganic phosphate and Mg2<sup>+</sup> whilst the levels of those that are repressed (in other words, upregulated upon glucose–starvation), are of transporters and periplasmic receptor proteins related to the import of alternative carbon and carbon–nitrogen sources, which include amino acids, carbohydrates, lactate, glycerol, peptides, dipeptides, and nucleotides (Gutierrez-Ríos et al., 2007). This transcriptome pattern is consistent with the consequences of carbon catabolite repression exerted by glucose (Saier et al., 1996). The phosphoenolpyruvate-dependent PTS controls the uptake of a large number of energetically preferred sugars. It comprises proteins, EI and HPr, which are common to all substrates, as well as sugar-specific permeases, enzymes II (EIIs). Interestingly, EI and HPr are mainly located near the poles of *E. coli* cell (Patel et al., 2004; Lopian et al., 2010; Govindarajan et al., 2012). Upon addition of the sugar to the growth medium, HPr is phosphorylated by EI; HPr-P produced is released from the polar membranes and distributes in the cell, though EI remains located near the poles of negatively curved sites (Govindarajan et al., 2013; see Membrane Curvature and Protein Location). A fraction of HPr-P gets near the membrane to phosphorylate the permeases, allowing them to transport the sugars into the cell and to phosphorylate them (Govindarajan et al., 2012).

Common affinities for molecules and ions within and between hyperstructures may help in this task of coordination. These molecules include polyamines, poly-(R)-3-hydroxybutyrate, polyphosphate and, of course, lipids (Norris et al., 2014b). Polyamines, for example, bind to nucleic acids, ribosomes and porins, and stimulate the synthesis of around 300 proteins in *E. coli* that include the sigma factors RpoS, FecI, RpoN, and related RNA polymerase omega subunit RpoZ, Cya, Cra, Fis, H-NS and SpoT (Igarashi and Kashiwagi, 2006). These multiple actions may allow polyamines to play an important, coordinating, role in the survival of *E. coli* in the transition to stationary phase (Terui et al., 2012; Norris et al., 2014b) and during osmotic changes (Munro et al., 1972).

Similar global regulatory roles can be ascribed to CL and an increase in CL levels has been proposed as a general physiological response that protects microorganisms from lysis due to osmotic stress (Catucci et al., 2004) and, in line with this, an increase in osmolality leads to transcriptional activation of the *cls* gene in *E. coli* (Romantsov et al., 2007). This increase in CL and its distribution to the poles was correlated with the polar distribution of the osmosensory transporter ProP which actively transports osmo-protectants into the cell (Mileykovskaya, 2007; Romantsov et al., 2007). In such coordination of lipid metabolism with the environment, the formation of a hyperstructure containing membrane components may help. Acyl carrier protein (ACP), a small protein of 9 kDa, interacts with diverse proteins associated with many biosynthetic pathways, including enzymes involved in synthesis of fatty acid in the cytoplasm as well as enzymes on or in the membrane involved in phospholipid or LPS synthesis; the latter assocations could account for the partial localization of ACP with *E. coli* membranes. Two enzymes involved in phospholipid synthesis, PlsB, a *sn*-glycerol-3-phosphate acyltransferase and PssA, a PS synthase, have been shown to interact with ACP and YbgC, an acyl-CoA thioesterase involved in fatty acid synthesis, using a tandem affinity purification method (Gully and Bouveret, 2006). *E. coli* PssA, which has a preference for acidic phospholipids, exists as both a membrane-associated active form and a cytoplasmic latent form and is at the heart of a cross-feedback regulation of the synthesis of zwitterionic (PE) and acidic phospholipids (Shibuya, 1992; Okada et al., 1994; Saha et al., 1996a; Matsumoto, 1997; Rilfors et al., 1999; **Figure 5A**). Bacterial two-hybrid analysis has shown that the enzymes in the phospholipid synthetic pathway, PlsB-PssA and ACP, form a complex in/on the inner membrane and further that YbgC, a fatty acid synthesis enzyme, and PlsB, form a complex in association with ACP (Gully and Bouveret, 2006). Thus, physical interactions between many of the enzymes responsible for fatty acid synthesis and phospholipid synthesis could form a hyperstructure to help coordinate lipid metabolism.

Sensing changes in nutrient availability entails bacteria transmitting this information to flagella hyperstructures to perform chemotaxis (see Transertion Problems) and to the division apparatus to regulate their size, grow faster and become larger when they are grown in nutrient-rich media. UgtP, a glucosyltransferase responsible for the synthesis of glucosylated diacylglycerols, one of which is the anchor of LTA synthesis, is localized, in a dot-pair structure similar to that of the open ring of FtsZ, to the division site in *B. subtilis* (Nishibori et al., 2005). UgtP is a division inhibitor, which prevents assembly of FtsZ and which results in an increase in the length of the cells under nutrientrich conditions (Shiomi and Margolin, 2007; Weart et al., 2007; **Figure 5**). It has been proposed that UDP-glucose acts as a proxy for nutrient availability and modulates the equilibrium between the UgtP–UgtP oligomer and the UgtP-FtsZ complex thereby serving as a molecular rheostat to help ensure that cell size is precisely co-ordinated with growth rate and nutrient availability (Chien et al., 2012). OpgH, a glucosyltransferase, is an integral membrane protein in the inner membrane of *E. coli* that is involved in the synthesis of osmoregulated periplasmic glucans, OPG (MDO); recently, OpgH has been localized to the division ring and shown to inhibit FtsZ assembly in the presence of UDP-glucose so as to delay the timing of cell division (Hill et al., 2013; **Figure 5A**). These two, very different, bacteria employ unrelated glucosyltransferases – a peripheral membrane enzyme for LTA synthesis in *B. subtilis* and an integral inner membrane enzyme for OPG synthesis in *E. coli* – that both have the second, "moonlighting," function of coupling nutrient availability to cell division by regulating FtsZ assembly (Hill et al., 2013). Disruptions of the moonlighting function of UgtP not only affect cell length: *ugtP* null mutants, which lack glucolipid products of the UgtP-catalyzed reaction, membrane glucolipids, are rounder (Price et al., 1997) and abnormally bent and distended (Lazarevic et al., 2005; Matsuoka et al., 2011a). Moreover, the extracytoplasmic function (ECF) σ factors, σM, σV, and σ*<sup>X</sup>* are constitutively activated (Salzberg and Helmann, 2008; Matsuoka et al., 2011a; Hashimoto et al., 2013), consistent with glucolipids directly influencing anti-σ<sup>M</sup> and anti-σ<sup>V</sup> factors by stabilizing conformations that sequester the respective ECF σ factors (Seki et al., 2015).

A two-hybrid-based investigation of interactions between the components of cell division machinery (27 proteins) and the enzymes involved in lipid synthesis (12 enzymes) in *B. subtilis* has revealed that FtsA interacts with PlsX (Takada et al., 2014). PlsX is an acyl-ACP:phosphate acyltransferase that synthesizes acylphosphate, an essential substrate for lysophosphatidic acid (LPA) production by PlsY, an acyltransferase that uses acyl-phosphate for the first acylation of *sn*-glycerol-3-phosphate (G3P); these are novel enzymes in a unique LPA synthesis pathway found in prokaryotic but not in eukaryotic cells (Lu et al., 2006;

#### FIGURE 5 | Lipid biosynthetic pathways and interactions of phospholipid synthases and proteins involved in cell division.

(A) Phospholipid biosynthetic pathway in *E. coli.* The gene product catalyzing each step is indicated. Abbreviations used are: G3P, glycerol-3-phosphate; LPA, lysophosphatidic acid; PA, phosphatidic acid; acyl-PO4, acylphosphate; acyl-ACP, acyl acyl-carrier protein; CDP-DG, (d)CDP-diacylglycerol; PS, phosphatidylserine; PE, phosphatidylethanolamine; PGP, phosphatidylglycerophosphate; PG, phosphatidylglycerol; CL, cardiolipin; OPG, osmoregulated periplasmic glucans (or MDO); UDP-gluc, UDP-glucose; DG, diacylglycerol; Lpp, major outer membrane lipoprotein; proLpp, prolipoprotein. Trace activities of PlsX-PlsY for LPA synthesis in *E. coli* (Lu et al., 2006/Hara et al., 2008) were indicated with thin letters and dotted arrows, different from those of the major pathway in *B. subtilis*. For Cls homologues, ClsA and ClsB, refer to Tan et al. (2012). Interaction of OpgH (a glucosyltransferase) and FtsZ indicated with thick arrow is from Hill et al. (2013); see Sensing). Activation of phosphatidylserine synthase, PssA, with acidic phospholipids indicated with thick dotted arrow is adapted from Shibuya (1992) and Matsumoto (1997). Interaction of PlsB and PssA is from Gully and Bouveret (2006), but to avoid apparent complexity the interactions between PlsB, ACP and YbgC, an Acyl-CoA thioesterase, shown by the authors, are not depicted. (B) Biosynthetic pathways for phospholipids, glucolipids and LTA in *B. subtilis*. Abbreviations used are: lysylPG, lysylphosphatidylglycerol; MGDG, monoglucosyldiacylglycerol; DGDG, diglucosyldiacylglycerol; TGDG, triglucosyldiacylglycerol; LTA, lipoteichoic acid; DHAP, dihydroxyacetone phosphate. For other abbreviations refer to (A) for *E. coli*. The pathway is adapted and compiled from Jerga et al. (2007) and Matsuoka et al. (2011b). For LtaS homologues (YflE, YfnI, YqgS and YvgJ) refer to Gründling and Schneewind (2007), Matsuoka et al. (2011b) and *(Continued)*

#### FIGURE 5 | Continued

Hashimoto et al. (2013). Interactions of FtsA and PlsX, and PlsY, both indicated with thick arrows, are from Hara et al. (2008) and Takada et al. (2014), respectively, but to avoid apparent complexity the interaction of PlsX with EzrA, DivIA and others, shown by the authors, are not depicted. Interaction of UgtP (a glucosyltransferase) and FtsZ indicated with thick arrow is from Weart et al. (2007) and Chien et al. (2012); see Sensing).