Plasmolysis Bays in Escherichia coli: Are They Related toDevelopment and Positioning of Division Sites?
EGBERT MULDER AND CONRAD L. WOLDRINGH*
Section ofMolecular Cytology, Department ofMolecular Cell Biology, University ofAmsterdam,Plantage Muidergracht 14, 1018 TVAmsterdam, The Netherlands
Received 2 October 1992/Accepted 3 February 1993
Plasmolysis bays, induced in Escherichia coli by hypertonic treatment, are flanked by zones of adhesionbetween the plasma membrane and the cell wall. To test the proposition of Cook et al. (W. R. Cook, F.Joseleau-Petit, T. J. MacAlister, and L. I. Rothfield, Proc. Natl. Acad. Sci. USA 84:7144-7148, 1987) thatthese zones, called periseptal annuli, play a role in determining the division site, we analyzed the positions ofthese zones by phase-contrast and electron microscopy. In situ treatment of cells grown in agar showed that theyoungest cell pole was the most susceptible to plasmolysis, whereas the constriction site was resistant. Lateralbays occurred only at some distance from a polar bay or a resistant constriction site. Orienting cells with theirmost prominently plasmolyzed polar bay in one direction showed that the lateral bays were always displacedaway from the polar bay at about half the distance to the other cell pole. If no poles were plasmolyzed, lateralbays occurred either in the centers of nonconstricting cells or at the 1/4 or 3/4 position of cell length inconstricting cells. The asymmetric positions of lateral plasmolysis bays, caused by their abrupt displacement inthe presence of polar bays or constriction sites, does not confirm the periseptal annulus model (Cook et al.),which predicts a gradual and symmetric change in the position of lateral bays with increasing cell length. Ouranalysis indicates that plasmolysis bays have no relation to the development and positioning of the futuredivision site.
Even in a simple organism like Escherichia coli, theremust be a mechanism to ensure that cell division alwaysoccurs between the segregated daughter chromosomes. Sev-eral proteins have been implicated to be specifically involvedin the division process (5), but the positioning mechanismitself has remained elusive. Two opposing models for posi-tioning of division sites have been proposed: in one model, anewborn cell already contains a preexisting division siteformed by replication and lateral displacement of an annularenvelope structure (2, 3, 13) and in the other model, the siteis generated only after chromosome duplication and nucleoidsegregation (9, 19, 20).
In the first model, the growing envelope contains concen-tric rings of adhesion (so-called periseptal annuli) betweenthe plasma membrane and the cell wall (peptidoglycan layerand outer membrane). The periseptal annulus model as-sumes that each newborn cell contains a pair of such annuliin its center. During subsequent cellular growth, new pairs ofannuli are generated at both sides of the central structure andare laterally displaced in a gradual way to 1/4 and 3/4positions of the cell length. The central annuli are used forseptum formation in the periseptal domain between theannuli (14). After division, each newborn cell inherits anannulus at its pole and a central pair of annuli as a presep-tation structure. This model predicts preexisting divisionsites in newborn cells and a commitment to division in thecell center independent of the DNA replication cycle.
In the second model, it is the presence of the nucleoidwhich directs the division site by influencing the rate ofpeptidoglycan synthesis in two opposing ways. The nucleoidinhibits cell wall synthesis in its vicinity, as has beendemonstrated by autoradiography (10). This inhibiting effect
has been called nucleoid occlusion (1). The opposing effect isactivation of peptidoglycan synthesis in the region of termi-nation of DNA replication, which usually occurs in the cellcenter. This activation, which has been shown to be neces-sary for initiation of constriction (16, 18), is supposed tooccur at a position along the cell envelope, which has beenabandoned by the segregating nucleoids, thus alleviatingtheir occlusion effect.The first concept of envelope-directed (pre)positioning of
division sites is based on observations of plasmolysis bays,which can be induced in gram-negative bacteria by hyper-tonic treatment (2, 8). The sensitivity to plasmolysis ap-peared to vary along the cell envelope: cell poles are moresensitive than constrictions (1, 8). According to Cook et al.(2) and MacAlister et al. (8), plasmolysis bays are localizedregions of periplasm bordered by zones of adhesion betweenthe plasma membrane and the cell wall, the so-called peri-septal annuli. Electron micrographs of serial sections ofplasmolyzed cells indicated that the annuli extend to variousdegrees around the cell cylinder and have led MacAlister etal. (8) to propose that the annuli mature during lateral
displacement until finally they flank the division site allaround the cell cylinder.
Plasmolysis bays can also be visualized by phase-contrastmicroscopy (2). The continuous change in the position ofplasmolysis bays attracted attention and led to the conceptthat periseptal annuli, generated at midcell, migrate toward1/4 and 3/4 positions along the cell (2). Additional confirma-tion for predetermination of division sites through periseptalannuli stemmed from analyses of plasmolysis bays in divi-sion mutants. InftsA filaments, plasmolysis bays are locatedat future division sites between the segregated nucleoids. Onthe other hand, in ftsZ filaments, positioning of plasmolysisbays is random (13 and this study), which has been inter-preted to mean that the FtsZ protein is involved in the
positioning process (13). Our phase-contrast and electronmicroscope analyses of plasmolysis bays in wild-type cellsand filamentous mutants have caused us to offer an alterna-tive explanation for the positioning of plasmolysis bays inbacterial cells. We propose that the positioning of bays alongthe cell's long axis is in principle random, but cell poles aremore sensitive and constriction sites are more resistant toplasmolysis. Our analysis thus shows that plasmolysis bayshave no relation to the development and positioning of thefuture division apparatus.
(Part of this research was presented at the 47th Sympo- FIG. 1. Phase-contrast micrographssium of the Society for General Microbiology held at the grown for more than 1 mass doubling IUniversity of Edinburgh in 1991 [11].) sucrose in situ. Note that the cell poles ithe most prominently plasmolyzed. Bar,
MATERIALS AND METHODS RESULTS
Bacterial strains and growth conditions. E. coli MC4100lysA and its isogenic derivatives ftsA1882 and ftsZ84 havebeen described previously (15). Cells were cultured understeady-state growth conditions at 30 or 370C in minimalmedium (15), supplemented with glucose (0.4%) as the solecarbon source, thiamine (4 pug/ml), and the required aminoacids (50 pug/ml [each]). The cell division ts mutants werecultured in 100 mosM medium to prevent the suppression bysalt at restrictive temperatures (15). Temperature shifts from42 to 30'C were carried out by fourfold dilutions of steady-state cultures (optical density at 450 nm, 0.2 to 0.3) intoprewarmed medium. The parent strain was grown at either100 or 300 mosM, obtained by the addition of 0.1 M NaCI.Osmolality was checked using a microosmometer (model3MO; Advanced Instruments, Needham Heights, Mass.).
Plasmolysis of cells suspended in agar. Cells were grown inLuria broth (Difco) to a concentration of about 108/ml.About 4 ,ul of exponentially growing cells was suspended in50 to 100 ,ul of 2% agar made up in Luria broth and kept at46°C. Subsequently, 20 ,l of the agar suspension was appliedto an object slide and compressed with a coverslip, previ-ously rubbed with silicon grease. After solidification, thepreparation was incubated at 37°C to allow cell growth. Aftermost cells had divided, usually after about 30 min, thecoverslip was removed and 4 RI of 15% sucrose (containing10 mM NaN3 to prevent recovery of the cells from plasmol-ysis; 12) was placed on the agar surface and covered with aclean coverslip for examination with an Olympus BH2-RFCA phase-contrast microscope.
Preparation of plasmolyzed cells for electron microscopy.Cells were plasmolyzed, fixed with glutaraldehyde, andwashed by centrifugation as described previously by Cook etal. (3), except that sucrose was dissolved in water. Concen-trations of sucrose used for plasmolysis ranged from 10 to35% in steps of 1 to 5%. Samples of plasmolyzed cells, withan optimal number of plasmolysis bays (see Results), wereselected by phase-contrast microscopy for agar filtration (17)and electron microscope examination (for a discussion of thevisualization of plasmolysis bays in agar filter preparations,see reference 12). Electron micrographs of agar filters (17)containing plasmolyzed cells were projected on a digitizingscreen (Summagraphics Co.) and measured and analyzed aspreviously described (10). Cell dimensions and positions ofbays and constrictions were measured by two observersusing a cross-shaped cursor to mark the respective points onthe cell's long axis (see the points indicated in Fig. 7). Foranalysis of separate categories of plasmolyzed cells, thepopulation was remeasured with a specially developed pro-gram (6).
; of cells mixed in agar,time, and plasmolyzed withfrom the recent division are, 5 jum.
In situ plasmolysis observed by phase-contrast microscopy.Previous observations showed that at relatively low sucroseconcentrations, most E. coli cells are only plasmolyzed atone pole (3, 12; see also Fig. 2A). To determine whether thesusceptibility of this pole is dependent on its age, cells weregrown in agar for 1 to 2 mass doublings before plasmolysis insitu. Cells suspended in agar appeared to elongate and todivide normally. After cell separation, however, the tips ofthe daughter cells slide past each other, as if pushed back bythe elastic agar. The two tips lying side by side, represent theyoungest cell poles. Scoring of 153 of such divided cell pairsafter in situ plasmolysis revealed that 89% contained bays attheir youngest poles, as illustrated in Fig. 1. It could alsoclearly be seen that lateral bays appeared only in longer cellsand that no bays occurred at constriction sites (see below).
Analysis of plasmolysis bays by electron microscopy. Elec-tron microscopy was used to analyze the positions of bays ata higher magnification. Most nondividing cells grown at 100mosM and plasmolyzed with 12% sucrose (Fig. 2A) showedplasmolysis at only one cell pole. At a sucrose concentrationof 18%, additional plasmolysis bays in the lateral part of thecell or at the other pole appeared (Fig. 2B). No plasmolysisbays could be observed at constriction sites. With sucroseconcentrations higher than 18%, the bays became bigger butless distinct because the entire cell collapsed (results notshown; see also reference 3). To analyze plasmolysis baypositions, we selected a sample of cells that were plasmo-lyzed with 18% sucrose.
Cell lengths, plasmolysis bay positions, and constrictionsites of about 1,500 cells were measured and plotted (Fig. 3)by a method similar to that previously performed by Cook etal. (see Fig. 2 and 4 of reference 2), but with the mostprominently plasmolyzed polar bay oriented to position 1 ofnormalized cell length and with unconstricted and con-stricted cells separated (Fig. 3A and B, respectively). At thetop of each panel in Fig. 3, the dot plot shows the normalizedpositions of individual bays as a function of cell length. Atthe bottom of each panel, the distribution of plasmolysisbays as a function of their position along the cell length isplotted.
In agreement with the results of Cook et al. (2), very fewlateral plasmolysis bays occur in the regions directly adja-cent to the polar bays, suggesting the presence of a zone ofresistance about 0.5 ,um wide. However, in contrast to Cooket al. (2), who observed central and symmetrical distribu-tions of lateral bays, it is evident (Fig. 3A) that the averageposition of the lateral bay is shifted away from the mostprominent polar bay, even in the shortest cells. On average,the lateral bays were found at position 0.43 of normalized
POSITIONING OF PLASMOLYSIS BAYS IN ESCHERICHIA COLI 2243
FIG. 2. Electron micrographs of E. coli grown at 370C in minimal medium of 100 mosM, plasmolyzed for 3 min with sucrose concentrationsof 12 (A) and 18% (B). The cells in panel A have polar bays only, whereas the cells in panel B also have lateral bays. Bar, 1 pum.
cell length. Considering separate length classes, the positionof lateral bays is gradually shifted from 0.45 in newborn cells(cell length, 1.3 pum) to 0.28 in cells with a length of 2.2 pum(Fig. 3A). These latter cells are about to initiate cell con-striction. As has been observed previously (8, 12), Fig. 3Bdemonstrates again that constriction sites are resistant toplasmolysis and that the distribution of bays is asymmetric.Because Olijhoek et al. (12) had found that plasmolysis
occurred more readily in long cells, we wondered whetherplasmolysis bays in longer cells would also be larger. Anincrease in polar bay size with cell length would displace thelateral bays if the bays were formed, on average, in themiddle, between the polar bay and the other cell pole. Toexamine this possibility, the sizes of polar bays were deter-mined and different categories of plasmolyzed cells (Table 1)were plotted separately (Fig. 4). The categories and theirproportions in the population are shown in Table 1. Eightypercent of the cells displayed a plasmolysis bay, and in 70%,a polar bay was present. The cells plotted in Fig. 4A and Cwere oriented with the most prominent polar bay downward.
In the unipolar cells, the size of the polar bay, measured asthe distance from cell pole to its border, is plotted as afunction of cell length (Fig. 4A). In unconstricted cells withcell lengths of less than 2.1 pum, the size of an average polarbay increased from 0.25 at the length of a newborn cell (1.3pum) to 0.5 pim at a cell length of 2.1 ,um. Figure 4B shows the
TABLE 1. Different categories of cells containingplasmolysis baysa
Category Type(s) of plasmolysis % of total % Constricted cellsbay population in categoryb
a A total of 1,520 cells were scored; the cells represented the same
population analyzed in Fig. 3 and 4.The percentage of constricted cells in the total population was 26%.
c See plots of the same categories in Fig. 4A to C. See also Fig. 7.d Nonplasmolyzed cells may either have become permeable during the
fixation and drying process or may have collapsed as a whole.
relatively few cells (9%; Table 1) that did not contain a polarbay. In the unconstricted cells, the lateral bays clusteredaround the cell center, with some lateral deviation in thelonger unconstricted cells (at 2 pum). In constricting cells, thebays cluster at 25 and 75% cell lengths (Fig. 4B). In cellscontaining both a polar bay and a lateral bay (Fig. 4C), theaverage polar bay was smaller than in unipolar cells (Fig.4A), but their size likewise increased with cell length (fromabout 0.2 to 0.4 pum). In bipolar cells (Table 1, category D),one polar bay was always larger (0.45 pum) than the other(0.25 pum) and both increased in size with increasing celllength (result not shown). This increase in the size of polarbays in longer cells confirms the results of Olijhoek et al.(12).The lateral bays in Fig. 4C do not cluster around the cell
center, as in Fig. 4B, but are abruptly displaced to, onaverage, the middle of the cell, between the polar bay andother pole (see the line indicated by the arrow in Fig. 4C). Inthe constricting cells, the lateral bays are again abruptlydisplaced to about 3/4 of the cell length, just as in the cells ofFig. 4B. Neither in Fig. 4B nor in Fig. 4C is a gradual changeof lateral bays from the cell center to the 3/4 cell positionapparent. Only when the two categories (B and C in Fig. 4)are combined in one plot is the impression of a progressivedisplacement with cell length, as seen in Fig. 3A, obtained.
Plasmolysis bay distributions in E. coli fts4 and ftsZ fila-ments. As previously reported (15), theftsA(Ts) cell divisionmutant, but not ftsZ(Ts), forms filaments at restrictivetemperature that show abortive constrictions (bottlenecks)at regular intervals. Cook and Rothfield (4) showed byphase-contrast microscopy that in ftsA but not in ftsZfilaments, plasmolysis bays are clustered at sites along thelength of the filament corresponding to potential divisionsites. In Fig. 5, we confirm this observation by electronmicroscopy, using filaments grown for 2 mass doublings at420C and treated with 15% sucrose. Only the ftsA(Ts)filaments (Fig. SB) showed clear positioning at 1/2 celllength, and more diffuse positioning at 1/4 and 3/4 celllengths. In longer ftsA(Ts) filaments, some additional posi-tioning at 1/8, 3/8, and 5/8 cell length is found. As in normallydividing cells (Fig. 3 and 4), one pole appeared moresusceptible to plasmolysis than the other, and additionalplasmolysis bays were similarly displaced, away from thelarger polar bay (oriented at the diagonal in Fig. 5 and 6). We
normalized cell lengthFIG. 3. Positioning of plasmolysis bays in E. coli plasmolyzed
with 18% sucrose. (A) Nonconstricting cells (n = 741) and (B)constricting cells (n = 280). The cells were measured from electronmicrographs, with the more prominently plasmolyzed pole placed atposition 1 of normalized cell length. In the dot plots, the normalizedpositions of plasmolysis bays are depicted as a function of celllength. Note that the plasmolysis bays are displaced away from themore prominent polar bay at position 1. The average length ofnewborn cells is 1.3 pum. The first constrictions appear at a celllength of 1.8 pum.
3
75%
2
75%
02~~~~~~~~~~~~~0°. . , 50%
0
0O0%% 2. 3''' '' ' 3 'En{3-.._ #' 75%
O 2 ... . @....-t~-.~~~~~~~~~~~~~X X 5-%
2 *.. -~~~~~~~~g 25
1 2 3
cell length (eim)FIG. 4. Different categories of plasmolyzed cells (see Table 1)
obtained after remeasurement of 1,520 cells from the same preparationused in Fig. 3. Cells were oriented with their more prominently polarbay downward. The diagonal line indicates the cell center. The arrowslabeled 25, 50, and 75%, indicate the 1/4, 1/2, and 3/4 positions of celllengths, respectively. (A) Cells with only one polar bay. The distancebetween the border of the bay to the nearest cell pole is measured. Thedistance between the border of the bay to the nearest cell pole ismeasured. The regression line, calculated for unconstricted cells,indicates that the bay size increases with cell length. (B) Cells withlateral bays and no polar bays. The location of the center of the bay isindicated. (C) Cells containing both a polar bay and a lateral bay. Aregression line as in panel A is shown. The line indicated by the arrowwas calculated as half the distance between the polar bay and secondcell pole. In panel C, some overlap exists between cells with a centralplasmolysis bay and those with a central constriction. Symbols: 0,polar bays; *, lateral bays; x, constriction sites.
thus confirm that in ftsZ(Ts) filaments, plasmolysis baypositioning is essentially random, whereas in ftsA(Ts) fila-ments, plasmolysis bays are preferentially positioned at thesites of abortive constrictions.
POSITIONING OF PLASMOLYSIS BAYS IN ESCHERICHM COLI
20
1 5
1 0
0
ca;IA
5
0
20
1 5
1 0
5
00 5 10 15 20
cell length (pi)FIG. 5. Plasmolysis bay positions in filaments of ftsZ(Ts) cells
(A) and ftsA(Ts) cells grown for 2 mass doublings at 420C (B). Thelines indicate 100 and 50% of cell length. The filaments are plotted,with the largest polar bay placed at the diagonal.
In the experiment of Fig. 6, we plasmolyzed ftsZ fila-ments, which had resumed division after the shift back to thepermissive temperature, to see whether a repositioning ofbays wpuld occur, as required by the periseptal annulusmodel (13). As previously described (19), such recoveringfilaments preferentially pinch off cells with a size between 1and 2 times that of the newborn cell at the permissivetemperature. This phenomenon can be seen in Fig. 6A,where the positions of constrictions were measured 30 minafter the shift back to the permissive temperature. Thepositioning of plasmolysis bays is shown in Fig. 6B anddemonstrates that although the filaments have several poten-tial division sites, only the sites of active constriction (Fig.6A) exclude the formation of a plasmolysis bay. No posi-tioning of plasmolysis bays is observed along the remainderof the filament. We conclude that in the absence of eitherabortive (sensitive) or active (resistant) constrictions, plas-molysis bays occur randomly along the length of the cell.
DISCUSSION
Preferred positions of plasmolysis bays. The present mea-surements indicate that in E. coli cells plasmolysis baysoccur preferentially at the (youngest) poles (Fig. 1), whereaslateral bays are in principle randomly positioned around themiddle of the cell or displaced to an asymmetric position inthe presence of a polar bay. The random location is evidentin relatively long, unconstricted cells (Fig. 4B) and in recov-eringftsZ filaments, which are dividing after the shift back tothe permissive temperature (Fig. 6B). As indicated by our
30
25
20
15
1 0
a,-I
0"4O;
I:
5
0
30
25
20
1 5
o~~~~~~~~~~~~~~
1 0
5
00 5 10 15 20 25 30
cell length (gm)FIG. 6. Positions of constriction sites (A) and plasmolysis bays
(B) in recoveringftsZ filaments, plasmolyzed 30 min after the shiftback to the permissive temperature. The arrows in both panelsindicate identical positions.
observations and by the measurements of Cook et al. (2),two different phenomena influence the positioning of lateralbays: the bays develop only at some distance from a polarbay and do not occur at a constriction site.The first phenomenon may be explained by assuming that
upon hypertonic treatment, the negative osmotic pressure isrelieved in a relatively large area around the site where themembrane is being pulled away from the wall. Such a reliefof pressure in the surrounding cytoplasm may prevent thedevelopment of a new plasmolysis bay in the immediatevicinity. The generation of such refractory zones may alsoexplain the seemingly regular positioning of bays in filamentswhich contain no abortive constrictions (Fig. SA).The second phenomenon, the exclusion of bays from an
area around active constrictions (Fig. 3B), may be explainedby a strong interaction between the plasma membrane andthe cell wall. Such a tight association was already empha-sized by MacAlister et al. (7, 8), after visualization ofplasmolyzed constriction sites in thin sections. They sug-gested the existence of a specific attachment at the leadingedge of the constriction which persisted even after cellseparation (1). Our autoradiographic observations of anincreased activity of peptidoglycan synthesis at the constric-tion site (16, 20) also suggest that the coherence between thewall and membrane might be stronger here. On the basis ofthese considerations, we originally proposed that the newestcell pole might still be more refractory to plasmolysis thanthe oldest cell pole (11). Our present observations (Fig. 1),however, clearly contradict this suggestion: it is in fact the
youngest pole that is the most sensitive to plasmolysis. Theinteraction between the wall and membrane is apparentlyabolished as soon as cell separation has taken place andpeptidoglycan is no longer synthesized. This abolishmentmay even involve an active process of modification of thepolar site, as suggested by Rothfield et al. (13). Suchinactivation could be necessary to prevent an additionalconstriction cycle, which would lead to minicell formation.The above interpretation is also in agreement with thesituation in ftsA filaments (Fig. 5B), where the sites ofabortive constrictions appear to have become the mostsensitive sites.
Displacement of plasmolysis bays. The existence of zoneswhich are refractory to plasmolysis limits the space fordevelopment of lateral bays and displaces them toward theother cell half in the presence of a polar bay. Such zones ofresistance do not seem to be supported by electron micro-scopic observations (7). Thin sections usually show smallgaps between the plasma membrane and cell wall that are toosmall to be seen with the light microscope. It is our conten-tion, however, that these gaps have been introduced duringthe dehydration procedure, which causes a dramatic shrink-age of the cell (17).
Orienting the cells with the most prominently plasmolyzedpole to one side clearly shows the asymmetric position of thelateral bays (Fig. 3A and 4C). Nevertheless, the lateral baysstill show, in Fig. 3A, a gradual displacement with increasingcell length if all cells are plotted as a function of normalizedlength (see also Fig. 4a in reference 2). This displacement ledCook et al. (2) to propose their hypothesis of periseptalannuli, which would occur at the centers of newborn cellsand would diverge symmetrically during cell elongation tothe 1/4 and 3/4 positions in dividing cells. The gradualdisplacement, however, largely disappears when separatecategories of plasmolyzed cells (Table 1) are plotted (Fig. 4).A schematic representation is given in Fig. 7 for the samethree categories as shown in Fig. 4. Although the lateral baysoccur, on average, at midcell in unconstricted cells withoutpolar bays (Fig. 7, B-i), these bays appear abruptly dis-placed in the presence of a polar bay (Fig. 7, C-1; arrow inFig. 4C) and are displaced even further when constrictionstarts (Fig. 7, C-2).
It is therefore our contention that the positioning ofplasmolysis bays in elongating cells does not represent aprogressive and symmetrical change from midcell toward the1/4 and 3/4 positions. If plasmolysis bays are not distributedrandomly, their positioning is influenced by the local activ-ities of peptidoglycan synthesis, which may cause an adher-ence of the plasma membrane to the cell wall and a strongerresistance to plasmolysis. Consequently, at the sites of thenucleoids, where the synthetic activity of the peptidoglycanlayer has been shown to be repressed (10), the membranewould retract more easily, whereas at constriction sites, withtheir high synthetic activity (16, 18), plasmolysis would bemore difficult.
Concluding remarks. The plasmolysis bays have beenproposed (2) to reflect the location of specific structures, theperiseptal annuli, which generate new division sites in athree-step process of replication, displacement, and arrest(1, 13), and thus represent predetermined division sites. Inaddition, the migrating annuli could serve as structures thatanchor the daughter chromosomes during DNA replicationand segregation (2, 13). In principle, however, the periseptalannulus model assumes no coupling between the DNAreplication process and the generation of a division site. Wehave proposed an alternative model in which division sites
A
B-1
B-21
C-1
C-2EX.A......I
F`
C
-2 -1 0 1 2FIG. 7. Schematic representation of three of the six categories
distinguished in Table 1. Category A is the largest category of cellsplasmolyzed at only one pole. In category B, the lateral bay occurs,on average, in the centers of unconstricted cells (B-i) and at 25 and75% positions in constricted cells (B-2) (compare with Fig. 4B).Category C also consists of unconstricted (C-1) and constricted(C-2) cells. Note the asymmetric displacement of the lateral bay,first because of the polar bay (C-1), and second because of thecombined effects of polar bay and constriction (C-2). Black dotsindicate how the measurement cursor was placed along the long axisof the cell to measure the size of the plasmolysis bays. The scale atthe bottom of the figure shows the cell lengths in micrometers.
are generated only at the site of termination of DNA repli-cation after the nucleoids have segregated (9, 19, 20). In thisnucleoid occlusion model, the position of the division site isnot predetermined but is coupled to the DNA replication andsegregation process. There is thus no requirement for spe-cific envelope structures which replicate and migrate alongthe cell envelope.
ACKNOWLEDGMENTS
We thank N. Nanninga for stimulating support in this work andcritical reading of the manuscript, L. I. Rothfield and W. R. Cookfor valuable discussions, A. Zaritsky for advice and critical remarks,N. 0. E. Vischer for help with the computer programs and plots,and J. H. D. Leutscher for preparation of the micrographs.
REFERENCES1. Cook, W. R., P. A. J. De Boer, and L. I. Rothfield. 1989.
Differentiation of the bacterial cell division site. Int. Rev. Cytol.118:1-31.
2. Cook, W. R., F. Joseleau-Petit, T. J. MacAlister, and L. I.Rothfield. 1987. Proposed mechanism for generation and local-ization of new division sites during the division cycle of Es-cherichia coli. Proc. Natl. Acad. Sci. USA 84:7144-7148.
3. Cook, W. R., T. J. MacAlister, and L. I. Rothfield. 1986.
POSITIONING OF PLASMOLYSIS BAYS IN ESCHERICHIA COLI 2247
Compartmentalization of the periplasmic space at division sitesin gram-negative bacteria. J. Bacteriol. 168:1430-1438.
4. Cook, W. R., and L. I. Rothfield. 1991. Biogenesis of celldivision sites in ftsA and ftsZ filaments. Res. Microbiol. 142:321-324.
5. Donachie, W. D., and A. C. Robinson. 1987. Cell division:parameter values and the process, p. 1578-1593. In F. C.Neidhardt, J. L. Ingraham, K. B. Low, B. Magasanik, M.Schaechter, and H. E. Umbarger (ed.), Escherichia coli andSalmonella typhimunium: cellular and molecular biology. Amer-ican Society for Microbiology, Washington, D.C.
6. Huls, P. G., N. Nanninga, E. A. van Spronsen, J. A. C.Valkenburg, N. 0. E. Vischer, and C. L. Woldringh. 1992. Acomputer-aided measuring system for the characterization ofyeast populations combining 2D-image analysis, electronic par-ticle counter, and flow cytometry. Biotechnol. Bioeng. 39:343-350.
7. MacAlister, T. J., W. R. Cook, R. Weigand, and L. I. Rothfield.1987. Membrane-murein attachment at the leading edge of thedivision septum: a second membrane-murein structure associ-ated with morphogenesis of the gram-negative bacterial divisionseptum. J. Bacteriol. 169:3945-3951.
8. MacAlister, T. J., B. MacDonald, and L. I. Rothfield. 1983. Theperiseptal annulus: an organelle associated with cell division inGram-negative bacteria. Proc. Natl. Acad. Sci. USA 80:1372-1376.
9. Mulder, E., and C. L. Woldringh. 1989. Actively replicatingnucleoids influence positioning of division sites in Eschenchiacoli filaments forming cells lacking DNA. J. Bacteriol. 171:4303-4314.
10. Mulder, E., and C. L. Woldringh. 1991. Autoradiographicanalysis of diaminopimelic acid incorporation in filamentouscells of Escherichia coli: repression of peptidoglycan synthesisaround the nucleoid. J. Bacteriol. 173:4751-4756.
11. Nanninga, N., F. B. Wientjes, E. Mulder, and C. L. Woldringh.1991. Envelope growth in Escherichia coli-spatial and tempo-ral organization: a new perspective, p. 185-223. In S. Mohan, C.
Dow, and J. A. Coles (ed.), Prokaryotic structure and function:47th Symposium of the Society for General Microbiology.Cambridge University Press, Cambridge.
12. OlUhoek, A. J. M., C. G. van Eden, F. J. Trueba, E. Pas, and N.Nanninga. 1982. Plasmolysis during the division cycle of Es-cherichia coli. J. Bacteriol. 152:479-484.
13. Rothfield, L. I., W. R. Cook, and P. A. De Boer. 1991. Biogen-esis of the Escherichia coli cell division system. Cold SpringHarb. Symp. Quant. Biol. 56:751-756.
14. Rothfield, L. I., T. J. MacAlister, and W. Cook. 1986. Murein-membrane interactions in cell division, p. 247-275. In M. Inouye(ed.), Bacterial outer membranes as model systems. Wiley andSons, New York.
15. Taschner, P. E. M., P. Huls, E. Pas, and C. L. Woldringh. 1988.Division behavior and shape changes in isogenic ftsZ, ftsQ,ftsA, pbpB, and ftsE cell division mutants of Escherichia coliduring temperature shift experiments. J. Bacteriol. 170:1533-1540.
16. Wientjes, F. B., and N. Nanninga. 1989. Rate and topography ofpeptidoglycan synthesis during cell division in Escherichia coli:concept of a leading edge. J. Bacteriol. 171:3412-3419.
17. Woldringh, C. L., M. A. De Jong, W. Van der Berg, and L. J. H.Koppes. 1977. Morphological analysis of the cell division cycleof two Escherichia coli substrains during slow growth. J.Bacteriol. 131:270-279.
18. Woldringh, C. L., P. G. Huls, E. Pas, G. J. Brakenhoff, and N.Nanninga. 1987. Topography of peptidoglycan synthesis duringelongation and polar cap formation in a cell division mutant ofEscherichia coli MC4100. J. Gen. Microbiol. 133:575-586.
19. Woldringh, C. L., E. Mulder, P. G. Huls, and N. 0. E. Vischer.1991. Toporegulation of bacterial division according to thenucleoid occlusion model. Res. Microbiol. 142:309-320.
20. Woldringh, C. L., E. Mulder, J. A. C. Valkenburg, F. B.Wientjes, A. Zaritsky, and N. Nanninga. 1990. Role of thenucleoid in toporegulation of division. Res. Microbiol. 141:39-49.
