Differentiation of the Bacterial Cell Division Site

INTERNATIONAL REVIEW OF CYTOLOGY, VOL. 118

Differentiation of the Bacterial Cell Division Site WILLIAM

R.

COOK, PIET A.

J. DE BOER, AND LAWRENCE I. ROTHFIELD

Department of Microbiology, University of Connecticut Health Center, Farmington, Connecticut 06032

I. Introduction

A. BACKGROUND In most bacterial species cell division occurs by the ingrowth of a division septum at the midpoint of the cell. This process is under strict topological and temporal control. We will discuss this subject as a problem in subcellular differentiation, in which a complex structure is constructed at a specific location in the cell in a process that must be coordinated with a variety of other cellular events, most notably with chromosome replication and segregation. In recent years a major change in thinking about bacterial division has come from the demonstration that early events in the differentiation process can be detected long before the initiation of septa1 ingrowth. In addition, more recent evidence suggests that the residual division site continues to carry out specific functions even after cell separation is completed. Thus, a major theme of this review will be the idea that the division site itself has a developmental history in which the stages of genesis, maturation, and localization, and ultimate fate have become accessible to study. In this context we will describe evidence obtained from several disciplines: genetics, microscopy, and, to a lesser extent, biochemistry. We will begin by briefly discussing differences between eukaryotic and prokaryotic cell division, and will describe aspects of bacterial cell envelope organization and chromosomal replication that are relevant to the later discussion of the division process itself. The main body of the review will be divided into three parts concerning (1) the formation and localization of the division apparatus, (2) the formation of the division septum, and (3) the coordination of cell division with other cellular events, such as DNA replication and chromosome segregation. The discussion will be largely limited to information derived from studies of gram-negative bac1 Copyright 0 1989 by Academic Press, Inc. All rights of reproduction in any form reserved.

3

&

WILLIAM K. COOK

ET A L .

teria, most especially of Escherichiu coli, because the combined application of genetics with other techniques to study cell division in this organism has been particularly fruitful in recent years. It should be pointed out that a substantial body of important work has also been done in other species. Although division in both bacteria and eukaryotic cells is accomplished by the circumferential invagination of the cytoplasmic membrane and cell envelope, the details differ considerably, reflecting the different organization of surface structures in the two groups. For example, contractile elements have not been described in most bacterial species. Therefore, it is not surprising that the contractile ring that appears t o play an important role in cytokinesis in animal cells (Beams and Kessel, 1976) is absent from dividing bacteria. On the other hand, the bacterial cell envelope contains a highly crosslinked peptidoglycan structure, the murein (described more fully later), that is not found in eukaryotic cells but plays a key role in the bacterial division process. The murein is the only known rigid structure in bacterial cells and, therefore, although it lies outside of the cytoplasmic membrane, may fulfill some of the roles ascribed t o the intracellular cytoskeleton of higher cells. The driving force in formation of the bacterial division septum may come from the circumferential inward growth of the rigid murein layer. In the simplest model, inner membrane is passively pushed ahead of the ingrowing murein while outer membrane is pulled behind, thereby leading to the coordinate inward movement of the three layers of the septum. This simplistic model provides a convenient starting point for thinking about the septation process.

B. ORGANIZATION OF THE BACTERIAL CELLENVELOPE As indicated in Fig. 1 , the cell envelopes of E. coli and other gramnegative bacteria contain three morphological layers: cytoplasmic (inner) membrane, murein, and outer membrane (Inouye, 1979). The compartment between the inner and outer membranes is termed the periplasmic compartment or periplasmic space. Formation of the division septum requires the ingrowth of this complex structure at the proper site within the cell. The cytoplasmic membrane acts as the major osmotic barrier of the cell, and contains a large number of specific transport proteins as well as proteins involved in energy transduction and other cellular processes. In contrast, the outer membrane contains a more limited spectrum of proteins, including several “porins” that permit the passage of various low molecular weight solutes into the periplasm (Nikaido, 1979). The murein layer is composed of an extensively crosslinked peptidogly-

DIFFERENTIATION OF BACTERIAL CELL DIVISION SITES

3

FIG. 1. Diagram of the Escherichia coli cell envelope. OM, Outer membrane; OmpA, OmpA protein; Mur, murein layer; PPS, periplasmic space; IM, inner membrane; MLP, bound form of murein-lipoprotein.

can that completely surrounds the cell. This bag-shaped molecule determines the shape of the organism and is responsible for the rigidity and the resistance of the cell to mechanical and osmotic stresses. Cell elongation and septal ingrowth both require that new peptidoglycan units be inserted into the continuous murein structure (Mirelman, 1979). This requires the controlled breaking of covalent bonds within the structure to permit the directed insertion of the new material. There is reason to believe that different biochemical mechanisms are used, at least in part, for insertion of septal murein as opposed to “elongation murein” (i.e., the murein along the length of the cell cylinder) (Mirelman, 1979). When examined by conventional electron microscopy, the murein layer of gram-negative bacteria appears as a thin, dense layer of 2-3 nm thickness that is closely apposed to the outer membrane (Fig. 1). Studies with alternative methods of preparation were interpreted as showing a less densely packed peptidoglycan domain between the dense murein layer and the inner membrane (Hobot et a / . , 1984; Leduc et al., 1985). Biochemical evidence shows that a number of intrinsic outer-membrane proteins (e.g., OmpA and murein-lipoprotein) are involved in attaching the outer membrane to the murein layer (Fig. 1). The outer-membrane proteins that mediate this attachment are sufficiently abundant (-lo5 copies per cell) to provide -400,000 contact points between murein and outer membrane (Park, 1987). The periplasmic compartment contains a number of water-soluble proteins. These proteins include degradative enzymes, proteins that inactivate toxic exogenous substances, and proteins that participate in solute transport and in chemotaxis (Brass, 1986). It is likely that other functional periplasmic proteins remain to be identified, and the possibility must be considered that some of these may play a role in the septation process.

4

WlLLlAM R. COOK

ET A L

Proteins can diffuse within the periplasmic space, although the diffusion rate is slower than in free aqueous solution (Brass, 1986; Foley rt ul., 1989). The periplasm can be operationally divided into two regions: the inner periplasm that lies between the inner membrane and the dense murein layer, and the outer periplasm located between murein and outer membrane (Fig. I). Free diffusion within the outer periplasm may be limited by the close apposition of murein and outer membrane and by the very high concentration of fixed proteins (most prominently OmpA and murein-lipoprotein) in this region. Significant movement of proteins within the periplasmic space may therefore be restricted to the inner periplasm.

C. DIFFERENCES BETWEEN CHROMOSOME SEGREGATION IN PROKARYOTIC AND EUKARYOTIC CELLS There are several differences in the process of chromosome segregation between bacteria and eukaryotic cells. Most strikingly, microtubules or analogous structures have not been shown to exist in bacteria, suggesting that the link between the chromosome and the cellular site responsible for directing daughter chromosomes to progeny cells in bacteria is likely to be less complex than in eukaryotes. Consistent with this fact, a mitotic apparatus has not yet been described in bacteria. It is widely believed-though without any direct evidence-that chromosome segregation occurs in bacteria by the attachment of daughter chromosomes to cell envelope sites that direct each of the daughter chromosomes to a different progeny cell. In the model originally proposed by Jacob et ul. (19631, the chromosomal site is located at o r near the origin of replication. In this model, still the most widely accepted one, the processes of chromosome replication and chromosome segregation are coupled and the driving force for chromosome separation is provided by insertion of new cell envelope material between the attachment sites. We will discuss in this review relevant aspects of this problem together with the question of how the cell coordinates the placement and synthesis of the division septum with the processes of DNA replication and chromosome segregation.

XI. Structure of the Division Apparatus A. THE PERISEPTAL ANNULARAPPARATUS Until the past few years, the only visible event in the cell division process was the formation of the division septum. However, earlier stages in the differentiation process have now become accessible to study. This

DIFFERENTIATION OF BACTERIAL CELL DIVISION SITES

5

was made possible by the discovery of a new organelle, the periseptal annular apparatus, which is present at the future division site before the onset of septal ingrowth (Anba et al., 1984; MacAlister et al., 1983). Studies of the structure and biogenesis of the periseptal annuli have provided a large amount of new information dealing with the process of formation and localization of the division site. The periseptal annuli are two concentric rings that surround the cell at the site of cell division (Fig. 2A). Electron microscopy has revealed that each annulus consists of a narrow zone within the cell envelope, in which the inner membrane, murein, and outer membrane lie in close apposition to each other (Fig. 2B). Three-dimensional reconstructions from serialsection electron micrographs showed that the zones of membrane-murein association are continuous structures that run completely around the cell cylinder. As discussed in more detail later, the annuli appear at the future division site long before the onset of septal invagination, thereby defining the future division site (Cook et al., 1986, 1987). The septum is later formed within the cell envelope domain (the periseptal domain) that lies between the two annuli. In bacterial cells the osmotic pressure resulting from the high intracellular concentration of impermeant solutes pushes the cytoplasmic membrane against the murein layer. Therefore, to visualize structures such as the annular attachments, cells are first plasmolyzed by brief exposure to hypertonic solutions of sucrose or other solutes that can enter the periplasm but do not readily cross the cytoplasmic membrane barrier. This results in a loss of cytoplasmic volume, causing the inner membrane to retract from the rigid murein-outer membrane layer. As shown by Bayer

FIG.2. Diagrammatic representation of periseptal annuli in surface view (A), and cross section (B) at several stages of progression through the cell cycle. PSA, Periseptal annuli; PA, polar annuli: OM, outer membrane; IM, inner membrane; M, murein. From Rothfield er al. (1986).

6

WILLIAM R. COOK t r

AL

et al. in an important series of papers (Bayer, 1979), random electron micrographs of plasmolyzed cells reveal numerous sites where the inner membrane fails to retract from the outer layers of the cell envelope. These attachments were named zones of adhesion by Bayer, and will be referred to as “Bayer bridges” in this review to avoid confusion with the adhesion zones that comprise the periseptal annuli. It is interesting to note that zones of adhesion between inner and outer membranes of isolated chloroplasts and mitochondria can also be visualized following exposure to hypertonic solutions (Cremers et al., 1988; Hackenbrock, 1968). The circumferential membrane-murein attachments that form the periseptal annuli and their biogenetic progenitors resemble the Bayer bridges when viewed in individual thin sections (MacAlister et al., 1983). It is not known whether all of the Bayer bridges that are visible in electron micrographs of random sections are in fact components of continuous structures such as the periseptal annuli, o r whether there are several types of adhesion zones that are structurally and functionally distinct.

B. ROLE OF T H E PERISEFTAL ANNULARAPPARATUS Although there is good evidence that the periseptal annuli are associated with the division process, the physiological role of the annular apparatus has not been established. Two possible roles, which are not mutually exclusive, have been suggested: ( I ) the annuli act as gaskets to segregate the division site from the remainder of the cell envelope, and (2) essential elements of the machinery are components of the annuli themselves. The gasket hypothesis was based on the morphological appearance of the annular attachments. The circumferential membrane-murein adhesion zones that comprise the annuli appear in electron micrographs to separate the periplasmic space into separate compartments, one of which defines the periseptal domain that lies between the paired annuli at midcell. It therefore was hypothesized that the annuli might act as physical barriers to prevent the unrestricted movement of molecules into and out of the periseptal domain. This would permit the cell to accumulate at this site proteins o r other components required for septum formation, ensuring that septation was restricted to the proper location. Studies by Foley et al. (1989) have provided experimental support for the gasket hypothesis. In these studies proteins labeled with fluorescent groups were introduced into the periplasm. The ability of the labeled proteins t o diffuse into different regions of the periplasmic space was followed by measuring the recovery of local fluorescence after the irreversible photobleaching of probe molecules within localized regions. The

DIFFERENTIATION OF BACTERIAL CELL DIVISION SITES

7

results indicated that certain regions of the periplasm were not in free communication with the remainder of the periplasmic space. These sequestered regions were preferentially located at potential division sites and at cell poles. This supports the view that the periseptal and polar annuli act as functional barriers to the passage of molecules into and out of the periseptal and polar domains that previously had been identified morphologically. It therefore is reasonable to suggest that one role of the annuli is to segregate the periseptal domain from the remainder of the cell envelope. It will be of interest to extend these studies to the inner and outer membranes to determine whether the barrier function is limited to the periplasm or whether membrane components are also sequestered at division sites by the annular gaskets. It is also possible that essential division components may be part of the annular attachments themselves. This question has not been accessible to experimental study because the annuli have not yet been isolated.

c. MOLECULARORGANIZATION OF PERISEPTAL ADHESIONZONES The molecular organization of Bayer bridges and of the periseptal adhesion zones is unknown. It appears likely t o us that the adhesion zones represent sites where inner membrane is attached to murein. This is based on the observation that the inner membrane at these locations resists the strong inward pull resulting from the plasmolysis procedure. This requires that inner membrane be attached to a rigid structure that can itself resist the inward pull. Since the murein is the only rigid structure that could provide an anchor for the inner membrane, we think it likely that protein(s) at these sites interact simultaneously with inner membrane and with the peptidoglycan that comprises the murein sacculus. The following general models are compatible with this formulation (Fig. 3): 1 . The adhesion zones are sites at which proteins directly attach inner membrane to murein without any involvement of outer membrane (Fig. 3A). This would explain the fact that inner membrane remains associated with the murein-outer membrane layer at these sites despite the inward pull of the plasmolysis procedure. 2. The adhesion zones contain protein that attach both the inner and outer membranes to the murein layer (Fig. 3B). This would explain the fact that inner membrane remains associated with the murein-outer membrane layer at these sites despite the inward pull of the plasmolysis procedure and would also be compatible with the evidence that the annular adhesion zones act as diffusional barriers within the periplasm. It should be noted that an outer membrane-murein attachment

8

WILLIAM R. COOK

ET AL

FIG.3. Hypothetical models of zones of adhesion. For details see text and Fig. 1.

would not be needed to explain the barrier function if lateral diffusion of proteins did not occur at a significant rate in the outer periplasm (i.e., the region between murein and outer membrane), as discussed in Section I,B. Though less likely, in our opinion, than the models just described, it is also possible that the adhesion sites represent regions of fusion between the lipid matrices of inner and outer membranes. Two such models are illustrated in Fig. 3C and D. In Fig. 3C the adhesion zones represent sites where the fusion of inner- and outer-membrane bilayers results in a continuous bilayer that connects the two membranes. This model is untenable in its simplest form, since an aqueous pore would be formed between cytoplasm and the external medium. This problem could be avoided if the sites contained protein(s) that plugged the potential pore. In Fig. 3D the adhesion zones represent sites where the inner- and outer-membrane bilayers have fused to form a new mixed bilayer. This structure would pre-

DIFFERENTIATION OF BACTERIAL CELL DIVISION SITES

9

C

D FIG.3. (con?.)

sumably be relatively unstable because of the suboptimal packing of phospholipid molecules at the junctions. This problem might be avoided if the sites contained proteins that served both to stabilize the common bilayer phase and to anchor the membrane to murein at these sites.

D. MEMBRANE-PEPTIDOGLYCAN ATTACHMENT AT THE LEADING EDGE OF THE SEPTUM A second differentiated structure based on the attachment of murein to inner and outer membranes has also been identified in high-resolution electron micrographs of the nascent septum (Fig. 4; MacAlister et al., 1987). The structure, called the septal attachment site (SAS) or membrane attachment at the leading edge (MALE), is located at the leading edge of the ingrowing septum throughout the septation process, forming a pursestringlike zone at the leading edge of the ingrowing septum. It consists of a sharply localized zone in which inner membrane, murein, and outer membrane are tightly apposed. The attachment zone at the septal edge

10

WILLIAM R. COOK

E I - AI

FIG 4. Membrane attachment at the leading edge (MALE) of the nascent septum. ( A ) Electron micrograph of septa1 region of a dividing cell from a culture of Esclii~richirrc d i . The cells were plasmolyzed in 20% sucrose prior to fixation. X 88,320. (B) Enlargement of (A) showing the characteristic bulbous enlargement of the murein (M)-outer membrane (OM) layers that lie in close apposition to the inner membrane (IM) at the leading edge. x 320,000. (C) Schematic representation of (B) indicating the relationships of the various cell envelope layers. PPS, Periplasmic space. From MacAlister et al. (1987).

has a characteristic ultrastructural appearance that distinguishes it from the adhesion zones that form the periseptal annuli. After septal closure and cell separation, the attachment site appears to remain at the pole of the newborn cell, where it can be detected as a bacterial birth scar. It has been suggested that MALE may represent the site at which new murein units are inserted into the murein layer of the nascent septum (MacAlister et ul., 1987). If correct, this would explain the vectorial movement of the septum toward the interior of the cell, with the inner membrane being pushed inward by the ingrowing murein. This idea has gained support from the demonstration that incorporation of the murein precursor [3H]diaminopimelic acid ([3H] DAP) occurs predominantly at the leading septal edge, as determined by autoradiography of septating cells that were pulse-labeled with [3H]DAP (Wientjes and Nanninga, 1989). 111. Biogenesis and Localization of the Division Site

A. DIVISIONSITESARELOCALIZED LONGBEFORE THE ONSETOF SEFTATION The association of periseptal annuli with the division septum was originally established by serial-section electron microscopy (MacAlister et uf., 1983). The same experiments showed that structures resembling perisep-

DlFFERENTIATION OF BACTERIAL CELL DIVISION SITES

11

tal annuli were also located at other sites along the length of the cell, where septation was not occurring. Because these structures did not extend completely around the cell cylinder, it was suggested that they might be precursors of the circumferential periseptal annuli that flanked the division septum (Cook et al., 1987; MacAlister et al., 1983). As noted before, the annular attachments form the limiting borders of localized regions of plasmolysis (plasmolysis bays). Thus, although visualization of the attachments themselves requires electron microscopy, the plasmolysis bays that mark their locations can easily be visualized by light microscopy (Cook et al., 1986). The positions of the plasmolysis bays thereby have been used to identify the locations of the annular structures along the length of the cell. This made it possible to study large numbers of cells to establish the pattern of genesis and localization of the annular apparatus during the course of the division cycle. These studies led to the unexpected observation that the future division site was placed at its proper position along the length of the cell during the cell cycle that precedes the cycle in which it participates in septum formation (Cook et al., 1987). In these studies the fact that plasmolysis bays were already localized at midcell in newborn cells originally suggested that new annuli were formed very early in the life of the cell. Evidence that formation and localization of the periseptal annuli in fact occurred during the preceding cell cycle came from the demonstration that predivision cells contained annular structures at one-quarter and three-quarters cell lengths, in addition to the periseptal annuli at midcell. The annuli at the cell quarters were retained during division to become the midcell annuli of the newborn cells. Since the annuli mark the site at which the division septum is formed, the question of how the cell determines the correct location of the division site therefore becomes a question of how the cell localizes the annular apparatus at one-quarter and three-quarters cell lengths.

B. AREDIVISIONSITES GENERATED FROM PREEXISTING SITES BY A REPLICATION-DISPLACEMENT MECHANISM? Studies of annulus distribution at intermediate stages of the division cycle showed that new pairs of annuli were first detected immediately adjacent to the midcell annuli that are present in the youngest cells in the population (Cook et al., 1987). As the cells elongated, the paracentral annuli that were located on both sides of the periseptal annuli appeared to move progressively away from midcell toward the two poles until, in the longest cells, the structures were tightly clustered at one-quarter and three-quarters cell lengths (Fig. 5). These results led to the proposal that new division sites are generated

WILLIAM R. COOK

12

ET A L

I

I

. .

*.

1

.....

i.:. . ,

.

D

FIG 5. Formation and displacement of nascent periseptal annuli during the division cycle. From Cook el ul. (1987).

and localized in a three-step process by a repetitive cycle of (1) replication of the preexisting sites at midcell, generating nascent division sites on each side of the central annuli; (2) displacement of the nascent annuli toward the two poles; and (3) arrest of the lateral displacement when the annuli have arrived at their final positions at one-quarter and three-quarters cell lengths. It is likely that the nascent annuli continue to mature during the displacement stage, since the annuli at intermediate positions appear in electron micrographs not to extend completely around the cell. Although this is difficult to prove, the micrographs suggest that the annuli continue to grow circumferentially around the cylinder during the displacement period with final closure of the ring occurring sometime after their final arrival at one-quarter and three-quarters cell lengths. If, as seems likely, septum formation cannot begin until the periseptal annuli extend completely around the cylinder to seal off the future division site, the coupling of the localization process with the process of annulus maturation would help to ensure that septation is not initiated at ectopic sites. Because the two new sets of annuli are generated on opposite sides of the preexisting central annuli, each of the nascent annular pairs is com-

DIFFERENTIATION OF BACTERIAL CELL DIVISION SITES

13

mitted to one of the two daughter cells from the moment of its generation at midcell. It has been pointed out that the nascent annuli are attractive candidates to serve as the structures that anchor the daughter chromosomes to the cell envelope during the process of DNA replication and chromosomal segregation (Cook et al., 1987). This would ensure that each daughter cell received one copy of the genome. Although we find this idea attractive, it is at present a hypothesis without supporting evidence. The recent demonstration that the E. coli origin of DNA replication binds specifically to a cell envelope fraction in vitro may facilitate identification of the cell envelope proteins responsible for chromosome binding, and this in turn may make it possible to test the hypothesis that the DNA-binding sites are located within the annuli (Ogden et al., 1988). c . RESIDUAL DIVISIONSITES REMAINAT THE CELL POLES AFTER CELL DIVISIONIs COMPLETED When cells divide, each daughter cell inherits one of the two periseptal annuli that had flanked the septum prior to cell separation. The residual annulus remains at the cell pole, defining a polar domain that lies between the polar annulus and the end of the cell. Because the polar domain is derived from the periseptal domain that delimited the septation site during the preceding division cycle, it is not unreasonable to expect that elements of the division machinery might be retained in this domain after cell separation is completed. Evidence that the residual division site at the new pole has the potential to support another cycle of septum formation has come from studies of minicell mutants of E. coli. These mutants are characterized by the frequent aberrant placement of the division site at the pole of the cell, leading to formation of large numbers of very small spherical cells (“minicells”) that lack chromosomal DNA (Frazer and Curtiss, 1975). It was originally suggested by Teather et al. (1974) that the minicell mutation results in loss of an activity that is needed to inactivate residual polar division sites. Later studies showed that deletion of the minicell genetic locus (minB) results in appearance of the minicell phenotype (de Boer et al. 1989). These studies have identified two gene products of the minB locus, the products of the minC and minD genes, that act together to form an inhibitor of septation that is required to prevent minicell formation. A third gene product, the minE protein, appears to give the minCD division inhibitor specificity for polar sites as opposed to normal sites at midcell. The demonstration that a septation inhibitor is required to prevent formation of polar septa is consistent with the view that the residual sites at

the cell poles retain the capacity to undergo additional cycles of septum formation. The phenotype of minicell-forming strains also includes cells that span a broad range of cell lengths, ranging from normal-sized cells to mediumlength filaments. Analysis of the length distribution of minB cells led Teather et af. ( 1974) t o propose that in minicell-forming strains the number of septation events per unit increase in cell mass remains normal. Thus, only one septum would be formed with each cell doubling, with an equal probability that septation will occur at the “normal” site at midcell or at one of the two residual division sites at the cell poles. If correct, this implies that at least one essential division component is present in limiting amounts, reaching a sufficient level at some point to support one septation event per division cycle. The hypothetical “division potential” would be used up after triggering septation, thereby preventing extra septation events. Alternatively, one could imagine that the initiation of one septation event could provoke a cellular change that excluded subsequent events until cell growth diluted the hypothetical inhibitor. A possible candidate for the hypothetical “division potential” molecule might be the products of the ftsZ gene, since overexpression of ftsZ appears to increase the number of septation events, resulting in formation of minicells and of chromosome-containing cells that are shorter than normal (Lutkenhaus, 1988; Ward and Lutkenhaus, 1985). It has been shown that some septation events in minicell strains result in the formation of rod-shaped cells lacking chromosomes (Jaffe et al., 19881, and that nucleoid distribution patterns are disturbed in the minB filaments (E. Mulder and C. L. Woldringh, personal communication). This suggests that the products of the minB gene may also be involved in maintaining the normal DNA segregation pattern.

IV. Formation of the Septum A. COORDINATION OF INGROWTH OF OUTER MEMBRANE-MUREIN-INNER MEMBRANE Septum formation requires the invagination of the inner-membrane, murein, and outer-membrane layers of the cell envelope. In E. coli the three layers appear to invaginate coordinately, based on thin-section and freeze-etch electron micrographs of septating cells. In a contrary view, Murray and collaborators have suggested that the invagination of inner membrane and murein precedes the invagination of outer membrane during normal septation in E. cofi, based on the use of alternative fixation

DIFFERENTIATION OF BACTERIAL CELL DIVISION SITES

15

procedures to the usual glutaraldehyde fixation protocol (Burdette and Murray, 1974a,b; Gilleland and Murray, 1975). For a more complete discussion of this question the reader is referred to Rothfield et al. (1986). However, it is clear that the ingrowth of the three layers is not obligatorily coupled, since mutants strains of E. coli and Salmonella typhimurium have been identified in which the ingrowth of the inner membrane and murein is uncoupled from the ingrowth of the outer membrane (Donachie et al., 1984; Donachie and Robinson, 1987; Fung et al., 1978; Weigand et al., 1976). The affected genes are cha and IkyD, respectively. In the uncoupled mutants the inner membrane and murein appear to invaginate normally, resulting in formation of filaments that contain multiple inner membrane-murein crosswalls that divide the cytoplasm into cell-length units. The units are held together by bridges of outer membrane that failed to participate in septal ingrowth. Thus invagination of outer membrane requires one or more proteins that are not required for invagination of inner membrane and murein. These proteins could, for example be involved in attaching outer membrane to the leading edge of the invaginating septum.

B. BIOCHEMISTRY OF SEPTUMFORMATION 1. Role of the ftsl Protein (PBP3) The only division-related protein for which a biochemical function has been identified is the product of theftsl gene (Spratt, 1983). Theftsl gene encodes a 60-kDa inner-membrane protein that has been identified as penicillin-binding protein 3 (PBP3). The specific interaction of the f ts l gene product with p-lactams implies that the protein plays a role in murein biosynthesis or metabolism. It has also been reported that purified preparations of PBP3 show two enzymatic activities presumably involved in murein biosynthesis, a transglycosylase and a transpeptidase, the transpeptidase being the most sensitive to inhibition by p-lactam antibiotics (Ishino and Matsuhashi, 1981). Evidence that PBP3 is required for septum formation has come from studies of thermosensitiveftsl mutants, in which growth at elevated temperature results in formation of nonseptate filaments (Nishimura et al., 1977; Spratt, 1977; Suzuki et al., 1978). Additional support for this view comes from the observation that filament formation also is induced by treatment of normal bacteria with antibiotics, such as cephalexin, that have a high affinity for PBP3 (Botta and Park, 1981; Spratt, 1975). Taken together, the aforementioned evidence strongly suggests that PBP3 plays a direct role in synthesis of septal murein. This implies that PBP3 either

I6

WILLIAM R. COOK

ET Al.

is preferentially located at division sites or is more randomly distributed and can be functionally activated at division sites at the appropriate time in the cell cycle. These possibilities should be accessible to study by immunoelectron microscopy using antibody directed against PBP3 or by autoradiography of cells labeled with isotopically labeled PBP3-specific compounds. 2 . Other Proteins Implicated in Cell Division A number of other gene products have been implicated in the division process based on the isolation of conditional mutants that form nonseptate filaments when grown under nonpermissive conditions. It is not known whether any of the gene products participate directly in the differentiation process. They have been reviewed elsewhere (Donachie and Robinson, 1987; Lutkenhaus. 1988). V. Regulation of the Division Process A. THEE. coli CELLCYCLE 1. Description of the Division Cycle

In E. coli the cell division cycle is a repeating series of events, punctuated by chromosome replication and septum formation (Helmstetter, 1987). For E. coli B/rA grown at a generation time of 60 minutes, chromosome replication (defined as the C phase) requires -40 minutes. Septa1 ingrowth begins 8-10 minutes after completion of replication. Approximately 10-12 minutes are then required for completion of septation and physical separation of daughter cells, although the time required for septum ingrowth increases when doubling times are >60-70 minutes. An additional interdivision period is present before the initiation of the next round of chromosome replication (Helmstetter and Pierucci, 1976; Kubitschek et al., 1967; Skarstad et a / . , 1983, 1985). This may be equivalent to the G , phase of the eukaryotic cell cycle (Cooper, 1984). Similar values have been obtained in both B/r and K-12 strains (Helmstetter, 1987). The components required for formation of the division septum are generated prior to the termination of chromosome replication. This is shown by the observation that in cells that have just completed chromosome replication, septation is not prevented by inhibitors of DNA, RNA, and protein synthesis(BrehmerandChuang, 198lb;Clark, 1968; Dix and Helmstetter, 1973; Helmstetter and Pierucci, 1968; Jones and Donachie, 1973; Kubitshek, 1974; Pierucci and Helmstetter, 1969; Woldringh et al., 1977). The durations of the C phase and septation periods are generally invariant

DIFFERENTIATION OF BACTERIAL CELL DIVISION SITES

17

when generation times fall between 20 and 60 minutes, but they gradually slow down when generation times are prolonged beyond 60 minutes.

2. Initiation of Chromosomal Replication Slowly growing cells (generation time >50-60 minutes) contain approximately two completely replicated chromosomes per cell at the time of septum formation. On the other hand, the number of partially or completely replicated chromosomes per cell increases by a factor of 2-4 in rapidly growing cells (Helmstetter, 1987). This reflects the fact that initiation of new replication forks at the chromosomal start site (oriC in E. cofi K-12) can begin prior to completion of ongoing rounds of replication. As a consequence, rapidly growing cells contain multiple replication forks at the time of septation. Therefore, replication events can be initiated in one division cycle and completed in the next. This demonstrates that division cycle events can be overlapped. The number of replication forks (and therefore initiation events) per cell (Cooper and Helmstetter, 1968; Helmstetter and Cooper, 1968) and the average mass per cell are both dependent on the growth rate. Donachie (1968) deduced that the ratio of initiation events to cell mass was invariant over a broad range of growth rates, and concluded that initiation began when the ratio of chromosomal origins to cell mass fell below a critical value. This led to suggestions that a positive-acting substance required for initiation of chromosome replication is synthesized at a rate proportional to the overall increase in cell mass (Donachie, 1968; Jacob et al., 1963; Lark, 1979; Margalit et al., 1984; Sompayrac and Maaloe, 1973). Following the initiation event, levels of the initiator would be depleted, thereby preventing further initiations until the critical concentration was reestablished. In the alternative model, initiation of replication would be prevented by an inhibitor molecule that is synthesized once during each replication cycle, after replication of the appropriate gene locus (Pritchard et al., 1969; Pritchard, 1984). The intracellular concentration of the inhibitor would be diluted with increasing cell mass, until the level was too low to prevent initiation. At this point a new round of initiation would take place, followed by a new burst of inhibitor synthesis. 3. Termination of Chromosome Replication It has been suggested that formation of the division septum may be coupled to the termination of chromosome replication. This suggestion was originally based on studies of cells in which DNA replication was permitted to resume after 2 hours of thymine starvation. Both RNA and

18

WILLIAM R. COOK ET

AL

protein synthesis were required to restore septum-forming ability to these cells. By adding rifampicin o r chloramphenicol at various times after thymine supplementation, Jones and Donachie (1973) showed that septation was dependent on a 5- to I0-minute period of RNA and protein synthesis occurring 35-40 minutes after resumption of DNA synthesis. Septation occurred shortly thereafter. The initial round of chromosome replication also terminated at this time, leading to the proposal that “termination protein(s1” formed shortly after termination of chromosome replication are required for the initiation of septation (Jones and Donachie, 1973). The possibility also exists that the period of required protein synthesis reflected the time required for recovery from the SOS response (see later) and that the apparent relationship to termination was coincidental. A correlation between termination of chromosome replication and septum formation was also observed by Grossman et al. (1989). In these experiments, the presence of methionine during amino acid starvation was used to slow chromosome replication, resulting in a delay in the time of termination. There was a corresponding delay in the first burst of division that was seen when protein synthesis was allowed to resume, consistent with the hypothesis that termination is required for septum formation. It is also possible that the delay in division was not causally related to the delay in termination but rather reflected other secondary effects of the rnethionine treatment. It should be noted that septation can occur in cells blocked in DNA elongation and termination (see later), demonstrating that termination of chromosome replication is not absolutely required for septum formation. B.

COUPLING OF SEPTATION TO CHROMOSOME

SEGREGATION

REPLICATION AND

Under normal conditions the formation of anucleate cells is extremely rare. This implies that septation is both temporally and topologically coupled to the processes of DNA replication and segregation. Temporal coupling requires that chromosome replication and the decatenation and separation of the daughter chromosomes must be completed prior to completion of the septal crosswall. Topological coupling requires that the new septum must be located between the daughter chromosomes. If temporal and topological coupling did not occur, one can imagine two possible results: ( 1) a mixture of anucleate cells and cells containing more than one chromosome would be produced, and/or (2) septal closure would have a guillotine effect on the unreplicated o r unseparated DNA. Under normal circumstances neither of these effects occurs with significant fre-

DIFFERENTIATION OF BACTERIAL CELL DIVISION SITES

19

quency, implying that an effective cellular mechanism exists to coordinate septation with DNA replication and/or segregation. Septa1 ingrowth can begin prior to the completion of DNA replication and segregation (Fig. 6; Woldringh et al., 1977). Thus, if direct coupling does exist between DNA segregation and the timing of septum formation it appears unlikely to involve a control on the initiation of septal invagination. On the other hand, a mechanism to prevent closure of the crosswall until DNA replication and separation are completed remains possible. 1. Prevention of Formation of Cells Lacking Chromosomes When DNA

Synthesis Is Blocked

a. Negative Regulation of Septation by the SOS System. The best understood of the pathways that prevent formation of anucleate cells is a part of the SOS response, a complex cascade of reactions in which the presence of damaged DNA or the inhibition of DNA replication results in the induction of a number of genes required for DNA repair and other processes (D’Ari, 1985; Holland, 1987; Little and Mount, 1982; Walker, 1984; Witkin, 1976). One of the consequences of SOS induction is that septation is reversibly inhibited, preventing formation of daughter cells until the defective DNA has been repaired. The mechanism by which septation is halted after induction of the SOS response begins with the activation of the product of the recA gene to an active protease in response to damaged DNA (Little and Mount, 1982;

FIG.6. Thin-section electron micrograph of Escherichiu coli B/rK showing initiation of septal constriction prior to complete separation of nucleoids. X 66,000. From Woldringh et al. (1977).

20

WILLIAM R. COOK Er A L

Walker, 1984). The RecA protease cleaves LexA, a repressor of the SOS regulon (Little and Mount, 1982; Walker, 1984). One of the derepressed genes, the sfiA gene (also known as sufA),codes for an inhibitor of septation (Huisman and D’Ari, 1981; Huisman et a f . , 1984; Mizusawa et a f . , 1983). After removal of the inducing stimulus, LexA is no longer cleaved by RecA protease, and normal division is restored (Maguin et a f . , 1986a; Mizusawa et al., 1983; Mizusawa and Gottesman, 1983; Shoemaker et al., 1984). For further information on the SOS response the reader is referred to Walker (1987). Genetic evidence indicates that the target of SfiA action is the essential cell division gene, f t s Z , or its gene product (Gayda et a f . , 1976; George et a f . , 1975; Huisman et a f . , 1984; Jones and Holland, 1984, 1985; Lutkenhaus, 1983). It has been suggested that inhibition of septation involves a direct interaction between the SfiA and FtsZ proteins. This is based on the observation that the rate of degradation of the SfiA protein by Lon protease is much slower in cells containing wild-type FtsZ protein than in certainftsZ mutants [i.e., sfiB (also known as sufB)mutants] in which the SfiA-mediated division inhibition is lost (Jones and Holland, 1984; Mizusawa and Gottesman, 1983; Mizusawa et al., 1983; Shoemaker et ul., 1984). Consistent with this interpretation is the observation that overproduction of FtsZ can suppress the division-inhibitory effect of SfiA (Lutkenhaus et a f . , 1986; Ward and Lutkenhaus, 1985). Alternative explanations are possible, and a final judgment on the idea that SfiA acts directly on the FtsZ protein must await more direct biochemical experiments. The SOS response appears to be a damage control system rather than a mechanism responsible for coupling DNA replication and septation under normal conditions, since sfiA(sufA)and sfiB(sufB)mutations that inactivate the SOS division inhibition response have no apparent effect on the normal division pattern. A second RecA-dependent coupling between chromosome replication and septation is mediated by the sfiC gene, which is present in some but not all strains (D’Ari and Huisman, 1983). As with SfiA, SfiC is derepressed after RecA activation. However, sfiC derepression is not dependent on LexA cleavage, indicating that an alternative repressor, also subject to RecA cleavage must exist. sfiC has been identified as a component of excisible element €14 (Maguin et a f . , 1986b). Genetic evidence indicates that the target of SfiC action, like SfiA, isftsZ (D’Ari and Huisman, 1983). The action of SfiC is irreversible, and cells do not recover after removing the original inducer (Maguin et a f . , 1986a), making the physiological significance of SfiC action questionable.

DIFFERENTIATION OF BACTERIAL CELL DIVISION SITES

21

b. Regulation of Septation by SOS-Zndependent Mechanisms. Filamentation also occurs when DNA replication is blocked in strains containing mutations that result in an inoperative SOS pathway. This indicates the presence of a non-SOS mechanism that couples chromosome replication and septum formation (Burton and Holland, 1983, Huisman et al., 1980, 1983; Jaffe et al., 1986; Jaffe and D’Ari, 1985). Such a mechanism could play a role in coordinating chromosome replication and septation in unperturbed cultures (Huisman et al., 1983; Jaffe et al., 1986; Jones and Donachie, 1973), but this will remain conjectural until the responsible gene or genes have been identified and it has been shown that inactivation of these genes perturbs the normal division pattern. Although nonseptate filaments are formed when DNA synthesis is blocked in strains that lack the SOS system, a number of chromosomefree cells are also produced. This indicates that septation inhibition in the non-SOS system is not as effective as that provided by the action of sfiA in the SOS system (Jaffe et al., 1986; Jaffe and D’Ari, 1985). It is possible to envision two types of mechanism by which the SOSindependent coupling of cell division and DNA replication might be mediated. In the first mechanism, coupling could be mediated by a positive control element similar to the “termination protein” proposed by Jones and Donachie (1973), as discussed earlier. In this view the effector would be required for initiation of septation during each division cycle. It has been suggested that the product of thefrsA gene may be the coupling effector (Tormo et al., 1980, 1985a,b, 1986; Tormo and Vicente, 1984). Conversely, the non-SOS coupling could be mediated by a negativecontrol element. In this case the partial inhibition of septum formation that occurs when DNA synthesis is inhibited in SOS-defective cells could reflect the induction of a division inhibitor analogous to the SfiA septation inhibitor of the SOS system.

2. Coordination of Chromosome Replication and Cell Division As noted before, the evidence for direct coupling between chromosome replication and cell division-in the sense that a signal is passed from the replicating or postreplication chromosome to the septation machinery to trigger septation-is not compelling. In an alternative model, DNA replication and septum formation are not directly coupled but both processes are independently triggered by factors related to progression through the cell cycle. This would result in the temporal coupling of the two processes without requiring direct signaling between the chromosome replication process and the septation machinery. This is consistent with the observation that septation occurs in the absence of chromosome replication pro-

22

WILLIAM R. COOK

ET A1

vided that the SOS-induced division inhibition mechanism is not present (see later). The idea that chromosome replication and septation are separate, parallel pathways is also consistent with the fact that the timing of initiation of chromosome replication is much more precisely regulated than the initiation of septation (Brehmer and Chuang, 1981a; Koppes et al., 1978; Kubitschek, 1%2; Newman and Kubitschek, 1978; Schaechter et al., 1962; Skarstad et al., 1986).

c. RELATIONOF CHROMOSOME SEGREGATION TO LOCATIONOF THE DIVISIONSITE

1. Mutations that Affect Nucleoid Segregation

A number of genes have been identified that affect the fidelity of nucleoid segregation into daughter cells. These fall into three general groups: genes coding for products that are known to affect DNA synthesis, genes that affect DNA supercoiling (DNA gyrase mutants), and genes the biochemical function of which is still unknown (par mutants). a. DNA Synthesis Mutants. Inhibition of DNA synthesis induced by temperature shift in thermosensitive mutants blocked in the initiation or elongation steps of DNA replication, or by thymine starvation, results in formation of filaments. Nucleoids are restricted primarily to the center of the filament, presumably representing the original unreplicated chromosome. This was shown most clearly by the studies of Jaffe and collaborators, who examined the effects on the division pattern of blocking DNA synthesis in a number of different ways in strains that d o not express the SOS response (Jaffe et al., 1986; Jaffe and D’Ari, 1985). A two-phase response was observed. During the first few hours, filaments accumulated in the culture, ascribed to the non-SOS-mediated coupling of DNA replication and septum formation. Later, however, substantial numbers of anucleate cells appeared (620% after 4 hours), reflecting the resumption of septation after the initial period of blocked division. Thus the septation block that is mediated by the non-SOS-mediated DNA-division coupling system was transient (Jaffe et al., 1986). The formation of anucleate cells was abolished by cya or crp mutations and therefore is dependent on the presence of of cAMP and the CAMP-binding protein. The mechanism that underlines the cAMP dependence of this septation activity has not yet been defined (D’Ari et ul., 1988; Jaffe et ul., 1986).

DIFFERENTIATION OF BACTERIAL CELL DIVISION SITES

23

b. DNA Gyrase Mutants. i. gyrA(parD). Hussain et al. (1987a,b) have described a strain containing a mutation (parD) at 88.4 units on the E.coli genetic map, which resulted in the apparent failure to segregate nucleoids, leading to formation of filaments and to the production of anucleate cells. Subsequent experiments showed that the abnormal phenotype was due to an amber mutation in gyrA, coding for the a subunit of DNA gyrase (Hussain et al., 1987b); a second uncharacterized mutation in the same strain also may play a role in the abnormal phenotype. No effects on chromosome replication were noted. The aberrant pattern of septation in this strain has been described in detail, and the results show the presence of chromosome-free cells that span a broad range of cell sizes. The mechanism(s) that couple initiation of septation to cell mass appeared not to be altered, as shown by the fact that the total number of septa formed per mass doubling were similar to the numbers of septa formed in a wild-type strain. ii. gyrB. Mutations in the gene coding for the p subunit of DNA gyrase (gyrB) (Fairweather et al., 1980) fail to decatenate newly replicated chromosomes (Steck and Drlica, 1984). Presumably as a result of this, the cultures contain filaments with centrally localized nucleoids. The cultures also contain small anucleate cells (Orr et a / . , 1979), reflecting the uncoupling of septum formation from chromosome segregation. c. par Mutants. Two other mutations [parA, at 95 units (Hirota et al., 1968a; Norris et al., 1986) and pa&, at 65 units (Kato et a / . , 1988)]affect chromosome partition without any apparent effect on DNA synthesis. In both cases filaments are formed that contain a large centrally located nucleoid. DNA-less cells are also present in the cultures. The fact that chromosome segregation was affected in these mutants whereas DNA synthesis appeared to be unperturbed suggests that the primary defect may be in the process of chromosome segregation. It should be noted that a third par mutant (parB) was originally thought also to fill these criteria (aberrant chromosome segregation in the presence of normal DNA synthesis). The parB mutation was later found to be located in the dnaG gene and was shown to affect the initiation of DNA replication (Filutowicz and Jonczyk, 1983; Norris et al., 1986).

2. Does Nucleoid Position Determine Septa1 Position? a. Nucleoid-Occlusion Model. All of the mutationsjust described result in the formation of filaments with centrally positioned nucleoids, as

24

WlLLlAM R. COOK

ET AL.

well as substantial numbers of smaller, nonfilamentous cells that lack nucleoids. Thus, the cells are capable of septation but appear not to form septa at the normal midcell position. Studies of the pattern of division in these strains have led to the suggestion that the nucleoid(s) play an essential role in establishing the location of the division septum by exerting a local inhibitory effect on septum formation (Donachie et d . , 1984; Woldringh et d . , 1985; Taschner et al., 1987). We will call this the nuclear-occlusion model. We will now examine the implications of this interesting model and the evidence on which it is based. The model states that the location of the nucleoids is the sole determinant of septal positioning (Woldringh et al., 1985; Hussain et al., 1987a,b). In this view the entire cell envelope is competent to initiate septal ingrowth. Following chromosome replication in wild-type cells, the two daughter nucleoids are positioned at regular intervals along the length of the cell by an active nucleoid segregation mechanism, or perhaps by the passive diffusional redistribution of daughter chromosomes. The nucleoids then would act as local inhibitors of septum formation, thereby restricting formation of the septum to a region near the middle of the cell. Three possible mechanisms can be considered for a nucleoid-occlusion effect, if it exists. 1. The nucleoid provides a simple physical block to the inward move-

ment of the septum. 2. The proximity of the nucleoid perturbs the organization of the overlying cell envelope by local physicochemical effects. 3. The nucleoid elaborates a diffusible septation inhibitor that prevents the initiation of septal ingrowth within a limited distance from the nucleoid itself.

b. Evidence for the Model. The idea that nucleoid placement is the sole determinant of septal placement was first suggested by Woldringh et al. (1983, based on studies of dnaZTs cells in which division was permitted to resume after a period of inhibition of DNA synthesis. This resulted in the production of anucleate cells and minicells. It is of interest that rninicell production has not been described when DNA synthesis has been blocked by inactivation of other dna gene products. The authors noted that the coefficient of variation in cell length was much larger for the temperature-shifted culture because both minicells and long chromosome-free cells were formed. On the basis of the increased coefficient of variation, it was suggested that there were no predetermined sites for the execution of constriction in the downshifted cells. E. Mulder and C. L.

DIFFERENTIATION OF BACTERIAL CELL DIVISION SITES

25

Woldringh (personal communication) have also noted similarly broad coefficients of variation of placement of constrictions in filaments of other DNA mutants in which anucleate cells were produced. The use of coefficients of variation as a indication of the randomness of placement of septa is based on the assumption that the distance from septal constriction to cell pole represents a single unimodal length distribution population. It should be noted that this assumption need not be correct. For example, if preexisting sites were distributed at regular intervals between cell pole and nucleoid, septation at any one of these sites would be possible. Measurement of the distance from constriction to cell pole in a large number of cells would then yield a mixture of discrete size classes, each class representing a multiple of the unit cell length. In this hypothetical case, the coefficient of variation of segment length for the entire data set would be high despite the fact that septal placement was nonrandom. It will be of interest to see whether it is possible to resolve the length distribution pattern into more than one Gaussian population before making a decision on the randomness of septal placement in these experiments. Studies of mutants with DNA gyrase defects have also provided evidence that has been interpreted to support a nucleoid-occlusion model. Growth of gyrA(parD) cells under nonpermissive conditions (see earlier) led to formation of filaments with large centrally located nucleoids, leaving long regions of the filaments devoid of visible nucleoid material. This presumably was due to failure to decatenate daughter chromosomes after completion of replication. Septation within the anucleate regions of the filaments subsequently led to formation of small chromosome-free cells (Hussain et al., 1987a). The chromosome-free cells were said to consist of short rods of all lengths between minicells and normal-length cells, although this is difficult to evaluate based on the resolution of the published results. It was pointed out that these observations are compatible with the idea that the absence of nucleoids near the cell poles led to the random placement of septa in these nucleoid-free regions (Hussain et al., 1987a). It should be noted that inactivation of DNA gyrase changes the expression of a number of chromosomal genes in which the state of chromosomal supercoiling affects promoter activity (Schmid, 1988; Wang, 1985). This raises the possibility that any septal localization defect could be secondary to altered expression of genes that affect septal placement by a mechanism other than nucleoid occlusion. c. Evaluation o f t h e Model. The idea that the entire cell envelope is competent to initiate septation during the latter portion of the division cycle raises certain problems. It is not compatible, for example, with the

26

WILLIAM R. COOK

ET A L .

observation that in slowly growing cells periseptal annuli are localized at the future division site during the previous cell cycle, long before the onset of the C phase of the division cycle in which septation occurs at this site (see Section 11). It is therefore unlikely, in our view, that the potential to initiate septum formation is randomly distributed along the entire length of the cell at the time of formation of the new septum. The most direct way t o test the nucleoid-occlusion model is to determine the size distribution of newborn cells in which nucleoids are absent from the greater part of the cell body secondary to inhibition of DNA replication. This avoids the possible complications of other secondary effects of gyrase defects or of par mutations. If future division sites were localized prior t o chromosome segregation, septa would be expected to form at regular intervals along the filaments, with the exception of the occluded site at midcell. On the other hand, if nucleoid positioning were the sole determinant of septa1 positioning as stated by the nuclear-occlusion model, then septa would be randomly distributed and the anucleate cells that are formed would show a continuum of cell lengths, down to the size of minicells. In the most extensive published study of this type, Jaffe et al. (1986) have studied the division patterns of cultures in which DNA synthesis was blocked in the absence of the SOS system. The inhibition of DNA synthesis was accomplished in a variety of ways, including thymine starvation and temperature shift in dnaA, dnaB, and dnaC mutants. This led to the formation of filaments that contained only one or two nucleoids, primarily located near the midpoint of the filament, plus significant numbers of anucleate cells. The chromosome-free cells were of normal length and were relatively uniform in size, suggesting that the positioning of septation sites was not affected by the absence of nucleoid in the vicinity of the septation event. Similar results were reported by Hirota et a / . (1968b) in studies of a thermosensitive dnaA mutant. In our view, these studies provide the most cogent argument against the proposition that division sites are randomly located along the length of the cylinder and that selection of the site depends only on the absence of nucleoid. One can imagine a more permissive variant of the model in which the nucleoid would affect the placement of the septum by vetoing septation at nearby division sites, while permitting septation to occur at other potential division sites that are located at regular intervals along the length of the cylinder. If this were correct, newly born chromosome-free cells would fall into discrete size classes equivalent to multiples of the presumed unit cell length. The possibility can also be considered that aberrant nucleoid placement affects an earlier stage in differentiation of the

DIFFERENTIATION OF BACTERIAL CELL DIVISION SITES

27

division site, such as the formation or localization of periseptal annuli, rather than affecting the septation event itself. Further studies will be needed to determine whether nucleoid position indeed plays a role in the selection of the septation site. VI. Conclusions

In recent years, significant advances have been made in defining the E. coli cell division process. At the morphological level, the discovery of periseptal annuli has permitted the developmental history of the division site to be followed throughout the cell cycle. At the same time, genetic studies have identified a number of proteins that are implicated in formation and localization of the division apparatus, and in the process of septa1 morphogenesis. Much useful information remains to be obtained in these areas, most notably an understanding of the coordination of division site development and localization with other events of the division cycle. In this regard, the use of immunoelectron microscopy to study the cellular localization of division-related gene products has hardly been exploited. This promises to provide new information that will be essential to any detailed understanding of the mechanism of the division process. Nevertheless, if this rich body of information is to be extended to the molecular level, it will be necessary to develop methods for isolation of the division site itself. Until this is achieved, the picture that emerges will necessarily be incomplete. A second major area of interest concerns the relation between cell division and chromosome replication. It is clear that the two processes are temporally coupled under normal conditions. A considerable body of evidence has permitted the formulation of specific hypotheses regarding the basis of the coupling. However, it is still not known whether or not there is direct coupling between the two processes or, alternatively, whether both respond to a common division clock. Does the initiation or termination of chromosome replication generate a signal that normally regulates the timing of septum formation? Does the differentiating division site send a signal that affects chromosome replication? Do both processes respond to signals generated by other events of the division cycle? It has proved difficult to obtain unequivocal answers to these significant questions, and this general area remains an important field for future study. A third key question is, how does E. coli determine where to place the

28

WILLIAM R. COOK

ET A L

division septum? There was little experimental information dealing with this question until the past few years, when early stages in the localization process have been described at a morphological level. These studies have suggested that the localization of the division site takes place long before initiation of septum formation. At the same time, studies of cells in which chromosome partition is defective have led to the proposal that the position of the nucleoid plays a role in determining the placement of the division septum. The identification of gene products of the minicell locus that affect the site selection process suggests that considerable additional information about the localization process remains to be obtained from genetic approaches. Exploitation of these relatively recent advances is likely to lead to a more complete understanding of this important but poorly understood aspect of the division process.

REFERENCES Anba, J.. Bernadac, A,, Pages, J.. and Lazdunski. C. (1984). Bio/. Cell. 50, 273-278. Bayer, M. E. (1979). In “Bacterial Outer Membranes: Biogenesis and Functions” (M. Inouye. ed.). pp. 167-202. Wiley. New York. Beams. H. W., and Kessel, R. G. (1976). Am. Sci. 64, 279-290. Botta. G. A,. and Park, J. T. (1981). J. Bucteriol. 145, 333-340. Brass, J. M. (1986). Curr. Top. Microbiol. Irnmimol. 129, 1-92. Brehmer, H.. and Chuang. L. (1981a). J. Theor. B i d . 88, 47-81. Brehmer, H., and Chuang, L. (1981b). J . Theor. B i d . 93, 909-926. Burdette. 1. D. J.. and Murray. R. G. E. (1974a). J. Bucteriol. 119, 303-324. Burdette. I. D. J., and Murray, R. G . E. (1974b). J . Bucteriol. 119, 1039-1056. Burton. P.. and Holland. 1. B. (1983). Mol. Gen. Genet. 190, 309-314. Clark, D. J . (l%8). Cold Spring Harbor Svrnp. Quunt. B i d . 33, 823-838. Cook, W. R., MacAlister. T. J . , and Rothfield, L. 1. (1986). J. Bacreriol. 168, 14301438. Cook, W. R.. Kepes. F.. Joseleau-Petit. D.. MacAlister. T. J., and Rothfield, L. I. (1987). Proc. Nut/. Acod. Sci. U.S.A. 84, 7144-7148. Cooper. S. (1984). In “The Microbial Cell Cycle” (P. Nurse and E. Streiblova, eds.), pp. 7-18. CRC Press. Boca Ratan. Florida. Cooper. S.. and Helmstetter. C. E. (1968). J . Mol. Biol. 31, 519-540. Cremers, F. F. M.. Voorhout. W. F.. van der Krift, T. P., Leunissen-Bijvelt, J. J . M., and Verkleij, A. J. (1988). Biochirn. BiC>pPh?.S. Acru 933, 334-340. D’Ari, R. (1985). Biochirnir 67. 343-347. D’Ari, R., and Huisman. 0. (1983). J . Bocferiol. 156, 243-250. D’Ari, R.. Jaffe. A.. Bouloc, P., and Robin, A. (1988). J. Bacteriol. 170, 65-70. de Boer, P. A. J.. Crossley. R. E., and Rothfield, L. I. (1989). Cell (Camhridp, Mass.) 56, 641-649. Dix, D. E.. and Helmstetter, C. E. (1973). J. Bucirriol. 115, 785-795. Narrrre (London) 219, 1077-1079 Donachie. W. D. (1%). Donachie, W. D., and Robinson, A. (1987). In “Escherichia coliand Sulmonellu tvphitnrrrirrrn: Cellular and Molecular Biology” (J. L. Ingraham. K. Brooks Low. B. Magasanik,

DIFFERENTIATION O F BACTERIAL CELL DIVISION SITES

29

M. Schaechter, and H. E. Umbarger, eds.), Vol. 2, pp. 1578-1593. Am. SOC.Microbiol.. Washington, D. C. Donachie, W. D., Begg, K. J., and Sullivan, N. F. (1984). In “Microbial Development” (R. Losick and L. Shapiro, eds.), pp. 27-62. Cold Spring Harbor Lab., Cold Spring Harbor, New York. Fairweather, N. F., Om. E., and Holland, I. B. (1980). J. Eacteriol. 142, 153-161. Filutowicz, M., and Jonczyk, P. (1983). Mol. Gen. Genet. 191, 282-287. Foley, M.,Brass, J. M., Birmingham, J., Cook, W. R., Garland, P., Higgins, C. H., and Rothfield. L. I. (1989). Mol. Microbiol. (in press). Frazer, A. C., and Curtiss, R. (1975). Curr. Top. Immunol. Microbiol. 69, 1-84. Fung, J., MacAlister, T. J., and Rothfield, L. I. (1978). J . Bacteriol. 133, 1467-1471. Gayda, R. C., Yamamoto, L. T., and Markovitz, A. (1976). J . Bacteriol. 127, 1208-1216. George, J., Castellazzi, M., and Buttin, G. (1975). Mol. Gen. Genet. 140, 309-332. Gilleland, H. E., and Murray, R. G. E. (1975). J . Bacteriol. 121, 721-725. Grossman, N.,Rosner, E., and Ron, E. Z. (1989). J. Bacteriol. 171, 74-79. Hackenbrock, C. R. (1968). Proc. Natl. Acad. Sci. U.S.A. 61. 598-605. Helmstetter, C. E. (1987). In “Escherichia coli and Salmonella typhimurium: Cellular and Molecular Biology” (J. L. Ingraham, K. Brooks Low, B. Magasanik, M. Schaechter, and H. E . Umbarger, eds.), Vol. 2, pp. 1594-1605. Am. SOC.Microbiol., Washington, D. C. Helmstetter, C. E., and Cooper, S. (1968). J . Mol. B i d . 31, 507-518. Helmstetter, C. E., and Pierucci, 0. (1968). J . Bacteriol. 95, 1627-1633. Helmstetter, C. E., and Pierucci, 0. (1976). J. Mol. Biol. 102, 477-486. Hirota, Y., Jacob, F., Ryter, A., Buttin, G., and Naki, T. (1968a).J . Mol. Biol.35, 175-192. Hirota, Y., Ryter, A., and Jacob, F. (1968b). Cold Spring Harbor Symp. Quant. Biol. 33, 677-693. Hobot, J. A., Carlemalm, E., Villiger, W., and Kellenberger, E. (1984). J . Bacteriol. 160, 143-152. Holland, I. B. (1987). Cell (Cambridge, Muss.) 48, 361-362. Huisman, 0..and D’Ari, R. (1981). Nature (London) 290, 797-799. Huisman, 0.. D’Ari, R., and George, J. (1980). Mol. Gen. Genet. 177, 629-636. Huisman, O.,Jacques, M., D’Ari, R., and Caro, L. (1983). J . Bacteriol. 153, 1072-1074. Huisman, O.,D’Ari, R., and Gottesman, S. (1984). Proc. Natl. Acad. Sci. U . S . A .81,44904494. Hussain, K., Begg, K. J., Salmond, G. P. C., and Donachie, W. D. (1987a). Mol. Microbiol. 1, 73-81. Hussain, K., Elliot, E. J., and Salmond, G. P. C. (1987b). Mol. Microbiol. 1, 259-273. Inouye, M. (1979). In “Bacterial Outer Membranes: Biogenesis and Functions” (M. Inouye, ed.), pp. 1-12. Wiley, New York. Ishino, F., and Matsuhashi, M., (1981). Biochem. Biophys. Res. Commun. 101, 905-911. Jacob, F., Brenner, S., and Cuzin, F. (1963). Cold Spring Harbor Symp. Quant. Biol. 28, 329-348. Jaffe, A., and D’Ari, R. (1985). Ann. Microbiol. (Paris) 136A, 159-164. Jaffe, A., D’Ari, R., and Norris, V. (1986). J . Bacteriol. 165, 66-71. Jaffe, A,, D’Ari, R., and Hiraga, S. (1988). J. Bacteriol. 170, 3094-3101. Jones, C. A,, and Holland, I. B. (1984). EMBO J . 3, 1181-1186. Jones, C. A,, and Holland, I. B. (1985). Proc. Narl. Acad. Sci. U.S.A. 82, 6045-6049. Jones, N. C., and Donachie, W. D. (1973). Nature (London), New B i d . 243, 100-103. I(ato, J. I., Nishimura, Y., Yamada, M., Suzuki, H., and Hirota, Y. (1988). J. Bacteriol. 170, 3967-3977.

Koppes. L. J . H . , Woldringh. C. L., and Nanninga. N . (1978). J. Bucteriol. 134, 423-433. Kubitschek. H. E. (1962). Exp. Cell Res. 26, 439-450. Kubitschek. H. E. (1974). Mol. Gen. Genet. 135, 123-130. Kubitschek, H. E., Bendigkeit. H. E.. and Loken. M. R. (1967). Proc. Natl. Acud. Sci. U.S.A. 57, 1611-1617. Lark, K. G. (1979). In “Gene Expression” (R. F. Goldberger ed.), Vol. I , pp. 201-207. Plenum, New York. Leduc, M., Frehel. C., and van Heijenoort, J. (1985). J. Bacteriol. 161, 627-635. Little, J. W., and Mount. D. W . (1982). Cell (Cambridge. Mass.) 29, 11-22. Lutkenhaus, J . F. (1983).J . Bocreriol. 154, 1339-1346. Lutkenhaus, J. F. (1988). Microbiol. Sci. 5, 88-91. Lutkenhaus, J. F.. Sanjanwala, B., and Lowe, M. (1986). J. Bacteriol. 166, 756-762. MacAlister, T. J., MacDonald. B., and Rothfield, L. I. (1983). Proc. Natl. Acad. Sci. U.S.A. 80, 1372-1376. MacAlister. T. J., Cook. W. R., Weigand, R., and Rothfield, L. 1. (1987). J. Bacteriol, 169, 3945-395 1. Maguin. E., Lutkenhaus, J., and D’Ari, R. (1986a). J. Bacteriol. 166, 733-738. Maguin, E., Brody, H., Hill, C. W., and D’Ari, R. (l986b). J. Bacteriol. 168, 464-466. Margalit, H.. Rosenberger, R. F., and Grover, N. B. (1984). J. Theor. B i d . 111, 183-199. Mirelman. D. (1979). In “Bacterial Outer Membranes: Biogenesis and Functions” (M. Inouye, ed.), pp. 115-166. Wiley. New York. Mizusawa, S., and Gottesman, S. (1983). Proc. Natl. Acud. Sci. U.S.A. SO, 358-362. Mizusawa, S.. Court, D., and Gottesman, S. (1983). J. Mol. B i d . 171, 337-343. Newman. C. N., and Kubitschek, H. E. (1978). J. Mol Biol. I Z I , 461-471. Nikaido. H. (1979). In “Bacterial Outer Membranes: Biogenesis and Functions” (M. Inouye. ed.). pp. 3 6 1 4 7 . Wiley. New York. Nishimura, Y.. Takeda, Y .. Nishimura, A.. Suzuki, H., Inouye, M., and Hirota, Y. (1977). Plasmid I, 67-77. Norris, V.. Alliote. T.. Jaffe, A , , and D’Ari. R. (1986).J. Bucteriol. 168, 494-504. Ogden. G. B.. Pratt. M. J., and Schaechter, M. (1988). Cell (Cambridge. Mass.) 54, 127135. Om, E., Fairweather, N. F., Holland. I . B., and Pritchard. R. H. (1979). Mol. Gen. Genet. 177, 103-1 12. Park, J . T. (1987).In “Escherichiu coli and Sulrnonellu ryphimirrirtrn: Cellular and Molecular Biology” (J. L. lngraham, K. Brooks Low, B. Magasanik. M. Schaechter. and H. E. Umbarger, eds.), Vol. I . pp. 23-30. Am. SOC.Microbiol.. Washington, D. C. Pierucci. 0..and Helmstetter. C. E. (1%9). Fed. Proc., Fed. A m . Soc. Exp. Biol. 28, 17551760. Pritchard, R. H. (1984). In “The Microbial Cell Cycle’’ (P. Nurse and E. Streiblova, eds.) pp. 19-27. CRC Press. Boca Ratan, Florida. Pritchard, R. H.. Barth, P. T.. and Collins, J . (1969). Svmp. Soc. Gen. Microhid. 19,263297. Rothfield. L. I.,MacAlister, T. J., and Cook, W. R. (1986).In “Bacterial Outer Membranes as Model Systems” (M. Inouye. ed.), pp. 247-275. Wiley. New York. Schaechter. M., Williamson. J. P.. Hood, J. R.. Jr.. and Koch, A. L. (1962).J. Gen. Microhiol. 29, 421443. Schmid, M. B. (1988). Trends Biochern. Sci. 13, 131-135. Shoemaker, J. M.. Gayda, R. G. and Markovitz, A. (1984). J. Bacteriol. 158, 551-561. Skarstad, K., Steen. H. B., and Boye, E. (1983).J . Bucreriol. 154, 656-662. Skarstad, K., Steen, H. B., and Boye, E. (1985). J. Bucteriol. 163, 661-668.

DIFFERENTIATION O F BACTERIAL CELL DIVISION SITES

31

Skarstad, K., Boye, E., and Steen, H. B. (1986). EMBO J. 5, 171 1-1717. Sompayrac, L., and Maaloe, 0. (1973). Nature (London), New Biol. 241, 133-135. Spratt, B. G. (1975). Proc. Nail. Acud. Sci. U . S . A . 72, 2999-3003. Spratt. B. G. (1977). J. Bacteriol. 131, 293-305. Spratt, B. G. (1983). J . Gen. Microbiol. 129, 1247-1260. Steck, T. R., and Drlica, K. (1984). Cell (Cambridge, Muss.) 36, 1081-1088. Suzuki, H., Nishimura, Y., and Hirota, Y. (1978). Proc. Nut/. Acud. Sci. U.S.A. 75, 664668. Taschner, P. E. M., Verest, J. G. J., and Woldringh, C. L. (1987). J . Bucteriol. 169, 19-25. Teather, R. M., Collins, J. F., and Donachie, W. D. (1974). J. Bacteriol. 118, 407-413. Tormo, A., and Vicente, M. (1984). J. Bucteriol. 157, 779-784. Tormo, A., Martinez-Salas, E., and Vicente, M. (1980). J . Bacteriol. 141, 806-813. Tormo, A., Depazo, A., de la Campa, A. G., Aldea, M., and Vicente, M. (l985a).J. Bucteriol. 164, 950-953. Tormo, A., Fernandez-Salas, C., and Vicente, M. (1985b).J. Gen. Microbiol. 131,239-244. Tormo, A., Ayala, J. A., de Pedro, M. A., Aldea, M., and Vicente, M. (1986). J. Bacteriol. 166, 985-992. Walker, G . C. (1984). Microbiol. Rev. 48, 60-93. Walker, G. C. (1987). I n “Escherichia coli and Salmonella typhimurium: Cellular and Molecular Biology” (J. L. Ingraham, K. Brooks Low, B. Magasanik, M. Schaechter, and H. E. Umbarger, eds.), Vol. 2, pp. 1346-1357. Am. SOC.Microbiol., Washington, D. C. Wang, J. C. (1985). Annu. Rev. Biochem. 54, 665-697. Ward, J. E., and Lutkenhaus, J. F. (1985). Cell (Cambridge, Mass.)42, 941-949. Weigand, R. A., Vinci, K. D., and Rothfield, L. I. (1976). Proc. Narl. Acad. Sci. U . S . A . 73, 1882-1886. Wientjes, F. B., and Nanninga, N. (1989). J. Bacteriol. 171, 3412-3419. Witkin, E. M. (1976). Bacteriol. Rev. 40,669-707. Woldringh, C. L., De Jong, M. A., van den Berg, W., and Koppes, L. (1977). J. Bacteriol. 131, 270-279. Woldringh, C. L., Valkenburgh, J. A. C., Pas, E., Taschner, P. E. M., Huls, P., and Wientjes, F. B. (1985). Ann. Microbiol. (Paris) 136A, 165-171.