Molecular Microbiology (2004)

51

(2), 461–469 doi:10.1046/j.1365-2958.2003.03837.x

© 2003 Blackwell Publishing Ltd

Blackwell Science, LtdOxford, UKMMIMolecular Microbiology 1365-2958Blackwell Publishing Ltd, 200351

2461469

Original Article

Kinetics of plasmid segregationS. Gordon, J. Rech, D. Lane and A. Wright

Accepted 22 September, 2003. *For correspondence. E-maildave@!ibcg.biotoul.fr; Tel. (+33) 5 61 33 59 68; Fax (+33) 5 6133 58 86. †Present address: Whitehead Institute, MassachusettsInstitute of Technology, Nine Cambridge Center, Cambridge, MA02142–1479, USA.

Kinetics of plasmid segregation in

Escherichia coli

Scott Gordon,

1†

Jerôme Rech,

2

David Lane

2

* and Andrew Wright

1

1

Department of Molecular Biology and Microbiology, Tufts University Health Sciences Campus, Boston, Massachusetts, USA.

2

Laboratoire de Microbiologie et Génétique Moléculaire, Centre National de Recherche Scientifique, 118 route de Narbonne, 31062 Toulouse, France.

Summary

Low copy-number bacterial replicons occupy specificlocations in their host cells. Production of a GFP-Lacrepressor hybrid protein in cells carrying F or P1 plas-mids tagged with a

lac

operator array reveals that insmaller (younger) cells these plasmids are seenmainly as a single fluorescent focus at mid-cell,whereas larger cells tend to have two foci, one at eachquarter-cell position. Duplication of the central focusis presumed to represent active partition of plasmidcopies. We report here our investigation by time-lapsemicroscopy of the subsequent movement of thesecopies to the quarter positions. Following duplicationof the central focus, the new foci migrated rapidly anddirectly to their quarter-cell destinations, where theyremained until the next cell cycle. The speed of move-ment was about five times faster than poleward migra-tion of

oriC

and 50 times faster than cell elongation.Aberrant positioning of mini-F lacking its

sopC

cen-tromere demonstrated the requirement for the parti-tion system in this localization process. From themeasured number of F plasmid copies per cell itappears that each migrating focus contains two ormore plasmid molecules. The molecular basis of thisclustering, and evidence for phasing of the partitionevent in the cell cycle, are discussed.

Introduction

Partition of bacterial chromosomes and plasmids servesthe same purpose as eukaryotic mitosis, stable inherit-

ance of the genome. Segregation of chromosomes isreadily seen in eukaryotic cells, where motor proteinspropel them along microtubules during anaphase ofmitosis. In bacteria the process is more obscure. Thesmallness of the objects concerned, and the tendency ofcell cycle events to overlap rather than to occupy distinctphases as in eukaryotes, renders visualization of segre-gating bacterial replicons difficult. Molecular geneticstudies of dispensable, low copy-number plasmids, suchas F, P1 and R1 of

Escherichia coli

, have revealed thepresence in each plasmid of specific partition determi-nants – genes encoding an ATPase, a centromere-binding protein, and the centromere itself (Gordon andWright, 2000; Hiraga, 2000). The presence of functionalanalogues of these elements in eubacterial chromo-somes attests to their general importance (Ireton

et al

.,1994; Sharpe and Errington, 1996; Mohl and Gober,1997; Kim

et al

., 2000; Godfrin-Estevenon

et al

., 2002).However, we can presently see only dimly how theseelements displace and position their replicons. Recentwork showing that certain partition ATPases oscillate orform actin-like filaments offers encouraging signs (Ebers-bach and Gerdes, 2001; Moller-Jensen

et al

., 2002), butthe relationship of these properties to partition remainsfor the moment unclear. Correlating these activities withthe movements that replicons undergo in the course of acell cycle is essential to establishing the nature of thisrelationship.

The development of methods for visualizing plasmidsbased on green fluorescent protein (GFP) (Robinett

et al

.,1996; Gordon

et al

., 1997) and fluorescence

in situ

hybrid-ization (Niki and Hiraga, 1997) has made it possible todetermine how plasmids are positioned in the cell. Wehave previously shown that F and P1 plasmids carryingan array of tandemly repeated

lac

operators could betagged in living

Escherichia coli

cells by a GFP-LacI hybridprotein, resulting in fluorescent spots which mark the plas-mids’ location (Gordon

et al

., 1997). These studies haveshown where plasmids are located at different stages ofthe cell cycle, but they have not told us how fast or bywhat path they get there. We report here the use of time-lapse microscopy of GFP-tagged plasmids to determinethe speed and trajectory of the F and P1 plasmids under-going segregation, and thus to reveal directly the kineticsof their displacement in the cell and how this differs fromthat of the chromosomal origin.

462

S. Gordon, J. Rech, D. Lane and A. Wright

© 2003 Blackwell Publishing Ltd,

Molecular Microbiology

,

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, 461–469

Results

Mode of plasmid movement

Logarithmically growing cells of strain SG101 carryingthe GFP::LacI-producing plasmid, pSG20, together withF

¢

lacO

+

or P1

¢

lacO

+

were treated with arabinose to induceGFP::LacI synthesis and applied to a nutrient agar-coatedmicroscope slide at room temperature (

~

20

∞

C) for obser-vation. Under these conditions, the cell doubling time wasabout 140 min and more than 90% of the cells containedeither one or two fluorescent foci. Fields of cells werephotographed at 15 second intervals and the photographssubsequently scanned for foci in the process of duplica-tion

.

Duplication of foci occurred in about 0.5–1% of thecells during a 30-minute period of observation, lower thanexpected for the

~

20% of the cell cycle traversed duringthis period. This deficit probably results from growth arrestof a significant fraction of the population, as is often seenin fluorescence-based time-lapse experiments (data notshown), as we also observed that a lower than expectedfraction of the cells (5.3% of 300 counted) divided duringthe 30-minute period, and that duplications were observedonly within the first 10 minutes of an experiment. Theduplications took place within a narrow range of cell sizes,2.6–3.3

m

m long in the case of F, 3.5–4.3

m

m long for P1:the cell length range for the total population was 2.2–6.3

m

m. The newly duplicated foci were seen to moverapidly from the centre towards the quarter positions overa period of 1.0–1.5 min, and then to remain immobile ordrift slightly polewards during the following few minutes(Fig. 1).

The displacement of individual foci during successive15-second intervals was plotted relative to their initial posi-tions: examples are shown in Fig. 2. These plots revealthat the rate of movement was not constant. Periods ofrelative immobility and bursts of rapid motion were seen,sometimes simultaneously for a given pair of foci, as inthe case of the P1 sample A pair at 165–195 s. Butregardless of these irregularities in velocity, movementwas consistently directed away from the centre. Slightreversals in the plots of Fig. 2 are within the error ofmeasurement. Fluorescent foci in the great majority ofcells remained relatively stationary even at the earliesttime-points in the time-lapse series, suggesting that theplasmids are tethered rather than being free to movearound within the cell.

Rates of plasmid movement

The calculated average rates of movement for F and P1are similar (Table 1), 0.43 and 0.40

m

m minute

-

1

respec-tively. To compare these migration rates with that of thechromosomal origin, we used origin displacement datapreviously obtained with a derivative of SG101 carryingthe array of tandem

lac

operators near

oriC

(Gordon

et al

.,1997). Plotting movement of origin foci in the same man-ner as used for the F and P1 plasmids yielded an averagemigration rate of about 0.1

m

m per minute. The corre-sponding rate of cell elongation, for cells on the agaroselayer under the microscope, was about 0.01

m

m perminute. Thus, plasmids F and P1 migrate about six timesfaster than the chromosomal origin and 40–50 times faster

Fig. 1.

Time-lapse images of

E. coli

cells pro-ducing GFP-LacI. The cells contained either F (pOX38) or P1 tagged with the

lacO

cassette. Two cells (A and B) of each are shown. Images were acquired every 15 s from the moment when focus duplication took place, as shown on the top line. Positions of foci are indicated by superimposed pointers. The constancy of the distance between these after about 45 s shows that most of the plasmid movement occurred beforehand. In the first panel of P1 B the contrast has been enhanced to reveal cell dimensions, whereas subsequent panels are untouched to preserve focus definition. Scale bars = 1 micron.

Kinetics of plasmid segregation

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than elongation of the cell. This rate is also somewhatfaster than the 0.3

m

m min

-

1

average measured for RK2(Pogliano

et al

., 2001), when account is taken of thehigher incubation temperature (30

∞

C) used in that study.Nevertheless an RK2 duplication event seen by Poglianoand co-workers followed similar kinetics to those shownin Fig. 2.

Role of the partition system in plasmid positioning

If the pattern of plasmid positioning discerned hererepresents operation of the partition system, partition-defective derivatives of the tagged plasmids should notdisplay it. A direct test of the dependence of F

¢

lacO

move-ment on Sop function proved not to be feasible, becausewe were unable to combine the

lacO

array with any ofseveral

sop

locus deletions in either an F

¢

or mini-F. How-ever, an analogous detection system, consisting of anarray of

tet

operator sequences to which a Y(ellow)FP-TetR protein binds (Lau

et al

., 2003), was readily insertedinto a

D

sopC

mini-F plasmid. The rate of loss of the

tetO

-array derivative (pDAG480) during growth at 20

∞

C in LBmedium, 5.7

±

1.0% per generation (standard error ofthree measurements), was similar to that of the original

D

sopC

mini-F (pDAG115), 4.8

±

0.6%.The localization of pDAG480 and of its stable

sop

+

equivalent, pDAG479, were examined (Fig. 3).Figure 3A–C show that positioning of mini-F

sop

+

foci wasvery similar to that of F

¢

lacO

foci previously analysed(Gordon

et al

., 1997). In contrast, the

D

sopC

plasmid

Fig. 2.

Kinetics of F and P1 displacement following focus duplication. Each plot shows the movement of the paired plasmid foci shown and labelled as in Fig. 1, measured as the distance from the point where focus duplication was first observed. Tick marks at the right of each panel show the positions of and cell lengths. Cell length (

m

m) at the first and final time points were: A

-

2.9 and 3.5, B

-

3.1 and 3.6, C

-

3.9 and 4.4, D

-

4.1 and 4.45.

14

34

Table 1.

Rate of focus movement after duplication.

Rate (

m

m minute

-

1

)

a

Minimum Maximum Average

F

¢

lacO

0.08 1.89 0.43P1

¢

lacO

0.08 1.62 0.40

oriC

0.01 0.18 0.07cell length 0.007 0.012 0.009

a.

F and P1 data are each based on five focus duplications. Ratesare calculated from movements measured within each of the 15-second intervals from focus splitting till arrival within 0.1

m

m of thecell quarter position. The minimum and maximum figures (other thanfor cell length) are the lowest and highest non-zero values obtainedfor each DNA type (or cell length).

464

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showed no tendency to follow the wild-type pattern(Fig. 3D–H). Rather, its foci were located off the main axisof the cell, with a strong preference for the poles and thecell margins, as previously described for FISH localizationof a partition-defective mini-F (Niki and Hiraga, 1997).Even without a time-lapse analysis of

D

sopC

mini-Fmovement, it is obvious from these observations thatpositioning of mini-F, and by extension of F

¢

plasmids,near mid-cell and the cell quarter positions requires afunctioning partition system. Perturbed localization of apartition-defective mini-P1 has been reported recently (Liand Austin, 2002).

Plasmid content of fluorescent foci

In the time-lapse experiments the cells were never seento contain more than two foci. This was a surprising obser-

vation, as the number of F and P1 copies per average cellof

E. coli

growing in rich medium at 37

∞

C had been mea-sured at 4–5 (Prentki

et al

., 1977; Bremer and Dennis,1987; Austin and Eichorn, 1992; Biek and Strings, 1995).It suggested either that each focus represents two or moreplasmid molecules, or that the copy number drops to aminimum of one molecule per newborn cell owing to fail-ure of plasmid replication to keep pace with cell growthand division at the lower temperature, 20

∞

C, used in theseexperiments. We therefore determined the intracellularconcentrations of F and chromosomal DNA under thegrowth conditions used for the kinetic analysis.

The data presented in Fig. 4 show that certain cellparameters do indeed change upon shifting the tempera-ture of growth in LB medium from 37

∞

C to 20

∞

C. Wemeasured the relative concentrations of three chromo-somal markers and of mini-F by hybridizing probes of

Fig. 3.

Positioning of

sop

+

and

D

sop

mini-F plasmids. Cells carrying pFX234 (

tetR::yfp

) and the

tetO

array mini-F plasmids, pDAG479 (

sop

+

A

-

C) or pDAG480 (

D

sopC

; D

-

H), were grown at 23

∞

C and induced with arabinose for 30 min before sampling and observation by fluores-cence microscopy.

Kinetics of plasmid segregation

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identical specific activity to a restriction digest of totalcellular DNA that had been resolved by electrophoresisand transferred to membrane (Fig. 4B). These datarevealed that at 20∞C the gradient of chromosomal markerfrequencies is shallower than at 37∞C (Fig. 4C), meaningthat the ratio of the rate of replication fork initiation to thatof fork progression is reduced at the lower temperature.The lower average number of forks per chromosome at20∞C was also demonstrated by adding rifampicin to

inhibit initiation and measuring the amount of DNA madeas forks complete replication in progress (Fig. 4D).Because the measured amount of DNA per cell grown at20∞C is about half that for cells grown at 37∞C (Fig. 4E,fourth column), the reduced number of forks must reflectthe rate of chromosome initiation falling behind the growthrate, while fork speed keeps step with it, as expected. Thehybridization analysis (Figs 4B and E) showed, however,that F plasmid replication is less affected, such that at

Fig. 4. Determination of the number of copies of F¢lacO+ per cell.A. Location of sequences used to measure relative frequencies of chromosomal markers. The strain SG101 is a derivative of W3110, known to have a large inversion that displaces oriC anticlockwise from its normal position in the sequenced reference strain, MG1655. To compensate for this displacement in the determination of oriC and terC gene dosage, and to avoid possible complications arising from DNA transactions in the dif (terminus) region, sequences nearly one half chromosome distant on the same replication arm were used as surrogates. The substitutes for oriC and dif are near malS and feaR, respectively; ldc is an intermediate marker.B. Southern hybridization analysis of marker frequencies, showing representative results for DNA from the F¢lacO+ strain. DNA from a culture grown into stationary phase was also analysed (stat), to verify the uniformity of probe labelling; the relatively strong hybridization signal for repE reflects limited continuation of F replication in stationary phase.C. Gradients of marker frequency derived from the Southern hybridization analysis; each point is normalized to the radioactivity binding to terminus (feaR) DNA (open square), and represents the average of two (stationary phase) or four (log phase) experiments. Marker frequencies determined for cells maintained in balanced exponential growth by sequential dilution, or for cells grown in the presence of glucose or arabinose, were not significantly different from those shown.D. Residual DNA synthesis after inhibition of initiation, shown at the right of the curves as the fractional increase in DNA.E. F¢lacO+ plasmid molecules per cell. Errors show the range of measured values about the mean. Generation time (t) was estimated from measurements of culture optical density (600 nm). Unit chromosomes per cell were calculated from chemical measurements of DNA and electronic counting of cells, using 4.64Mbp as the length of one E. coli chromosome. The number of origins per unit chromosome is numerically equal to the factor by which DNA is increased after run-out replication in the absence of initiation, and is given by (C/t).ln2/1–2–C/t where C = chromosome replication time (Bremer and Dennis, 1987); values shown are taken from (D). The origin/terminus ratio is also a function of C and t (ori/ter =2C/t), so that the measured ratios (malS/feaR) provide an independent estimate of oriC/unit chromosome (parentheses); the lower values, from run-out synthesis, were used to calculate oriC/cell. RepE/cell was calculated from oriC/cell and the repE/oriC ratio obtained from Southern hybridization (B); it was divided by ln2 to calculate the number of F¢lacO+ copies per cell just before division.

466 S. Gordon, J. Rech, D. Lane and A. Wright

© 2003 Blackwell Publishing Ltd, Molecular Microbiology, 51, 461–469

20∞C the number of F plasmids per average cell, 4.0, isonly moderately lower than the number at 37∞C, 5.2.Indeed it is high enough to indicate that the averagenumber of plasmids per fluorescent focus is >2 in cellsgrown in LB medium at 20∞C. As the number of fociper cell never exceeds two and the time of replicationof F is random with respect to the cell cycle(Helmstetter et al., 1997), the average number of F plas-mid molecules can be estimated at 5.8 just before division(Fig. 4E), or an average of about three per focus in eachnew-born cell.

Discussion

In this work we have shown that sister copies of plasmidsP1 and F, visualized as fluorescent foci in E. coli cells,move rapidly from a fixed position at the cell centre to newsites at the one quarter and three quarter positions wherethey remain until cell division. Separation was abrupt, withrates of movement up to about 1 micron per minute, fivetimes faster than the rate of separation of chromosomalorigins and 50 times faster than the rate of cell elongation.Our observations allow one to picture plasmid partition asa process that at some time before cell division dissoci-ates plasmid copies from each other and then activelydrives them apart to a fixed location in each new cell halfwhere they remain until the following cell cycle. This viewof partition is unexceptionable, in that it is straightforwardand consistent with the mechanism proposed for R1 par-tition, the system for which the greatest amount of molec-ular detail is available (Møller-Jensen et al., 2002), as wellas with previous localization studies of low copy-numberplasmids (Gordon et al., 1997; Niki and Hiraga, 1997;Weitao et al., 2000; Ho et al., 2002). Nevertheless,inspection of our results in the light of other studies sug-gests that this model should continue to be regarded withcaution. Moreover, indications that partition ATPases ofthe Par/Sop family move en masse from one end of thenucleoid to the other (Ebersbach and Gerdes, 2001; J.Leung and A. Wright, unpublished data) are not easilyincorporated in this simple view.

We observed that the duplication event which immedi-ately precedes migration of foci occurred within a narrowcell size-range, corresponding in the case of plasmid F toa restricted phase of the cell cycle around cell age 0.2(data not shown). Previous data from this laboratory alsoshowed this phasing (Gordon et al., 1997). In contrast, Fplasmid replication has been shown to occur at all stagesof the cycle (Helmstetter et al., 1997). These observationscould be reconciled by proposing that partition does notalways follow replication immediately but waits for the‘partition phase’, implying that the cell regulates, ratherthan simply executes, partition. However, this suggestionis contradicted by the recent report of Onogi et al. (2002),

who observed FISH foci of mini-F to duplicate shortly afterthe restart of replication synchronized by temperature-shifting of a repts mutant. The experimental conditionsemployed are very different from ours, and it remains tobe seen which of these situations is more representativeof F partition behaviour. It is possible that timing of parti-tion can alter in response to changes in physiologicalconditions.

A recent study more comparable to ours, becausebased on time-lapse experiments, indicated a mode of P1plasmid segregation notably different from that describedhere (Li and Austin, 2002). Whereas we observed newfoci to appear well before cell division and to proceedquickly to their destinations without significant deviation,Li and Austin concluded that sister molecules are fre-quently captured (paired) at the cell centre at the time ofcell division then ejected bi-directionally into the twodaughter cells before following an erratic course over arelatively lengthy period to arrive eventually at the new cellcentres. This pattern of movement was particularly prev-alent in cells grown in minimal medium, but was also seenduring time-lapse analysis of cells shifted from minimal torich medium at the start of microscopical observation. Onthe other hand, measurements of focus position sug-gested that delayed segregation was relatively rare in theirbroth-grown cells. Thus, some aspect of the physiology ofminimal-grown cells may account for the differencebetween the timing of segregation seen in our experi-ments and that observed by Li and Austin. For example,a cellular factor involved in plasmid partitioning might belocalized early in the cycle in rich medium but at the timeof cell division in cells grown in defined medium. However,the shift-up experiment alluded to above suggests thaterrant movement of P1 foci might characterize cells grownin both rich and minimal media, in sharp contrast to ourobservations of rapid displacement in broth-grown cells.It is possible that the tagging systems used for detectingplasmid foci, Li and Austin’s non-functional Gfp::ParB-parS or our extended Gfp::LacI-lacO, can alter someaspects of the partition system’s performance. This issueremains to be settled.

The apparent ability of unsegregated P1 plasmids todelay cell division, as observed by Li and Austin (2002),seems unlikely to reflect a direct interaction with the grow-ing septum in view of the demonstration that P1 plasmidssegregate normally in FtsZ-depleted filamentous cells andthat F plasmids do so in cephalexin induced filaments,conditions under which no septal rings are formed. Suchcells do appear to have normally located replication fac-tories, as judged from their even chromosome distribution,implying that a likely candidate for the cellular structurewith which plasmid foci associate and to which they pro-ceed upon partitioning would be a component of the rep-lication factory, which, like the plasmids, is normally

Kinetics of plasmid segregation 467

© 2003 Blackwell Publishing Ltd, Molecular Microbiology, 51, 461–469

localized at mid- and quarter-cell quarter positions. Suchan association might not, however, be linked directly toreplication, as aberrantly localized, partition-defectiveplasmids are maintained at normal copy number (Fig. 3).

Our results revealed that there are multiple copies of Fand P1 per fluorescent focus, indicating that these plas-mids are segregating in groups or clusters, rather than asindividuals. This observation joins other reports of FISHor GFP-based plasmid localization that reveal numbers ofvisible foci well below the copy numbers of the plasmidsconcerned (Weitao et al., 2000; Pogliano et al., 2001; Liand Austin, 2002). It has not been ruled out that in thesecases the plasmids are co-localized, via independentattachment to a host structure for example, rather thandirectly linked to each other. However, such an attachmentwould appear to be specific to each plasmid, as differentplasmids do not co-localize (Ho et al., 2002). Provision bythe cell of a specific attachment site for each of the manyplasmids it can accommodate is an unlikely prospect(Austin and Nordstrom, 1990), and direct interaction ofplasmid molecules thus seems a more likely basis forclustering.

On the other hand, the nature of any specific links thatmaintain these clusters is unclear. One candidate is theintermolecular ‘handcuffing’ between iteron-bound RepE(F) or RepA (P1) monomers, which is thought to play animportant role in limiting replication frequency (Pal andChattoraj, 1988). An interesting consequence of, and pos-sible raison d’être for, plasmid clustering by this meanscould be the protection of replication control from thepartition event in rapidly growing cells. If plasmids weremerely paired, partition might sever the handcuffs andprecipitate untimely replication. In slowly growing cellswhere plasmid copy number reaches only two per cell,partition may indeed play such a role in replication control,as proposed by Abeles and Austin (1991), with intramo-lecular handcuffing remaining as a back-up regulatorydevice. On the other hand, a recent study of plasmidsegregation and incompatibility indicated that in rapidlygrowing cells this type of intermolecular interaction makesonly a limited contribution to clustering (J.-Y. Bouet, J.Rech, S. Egloff, D. Biek, and D. Lane, submitted).

The other likely basis of clustering is intermolecularadhesion of partition complexes. An immediate difficultyarises here however, because in the cases studied,removal of partition functions had little if any effect on thenumber of foci observed (Weitao et al., 2000; Li and Aus-tin, 2002; our results, Fig. 3). What makes this observationstill more puzzling is that, at least in the case of the DsopCmini-F examined here, the kinetics of plasmid loss fromdividing cells corresponds to an average of 4.3 segregat-ing units at division, closer to that expected for randomdistribution of 5.8 individual plasmid molecules (Fig. 4E)than the much higher rate expected for two plasmid clus-

ters. Application of time-lapse analysis to the migration ofvarious kinds of partition-defective mini-F and mini-P1plasmids should help to throw light on the nature of clus-tered state and the mechanism of plasmid movement.

Experimental procedures

Bacterial strains and plasmids

The background strain used for studies of F-lacO kinetics wasSG101 containing the GFP-LacI fusion under the control ofthe arabinose promoter on plasmid pSG20 (Gordon et al.,1997). Kinetic studies of P1-lacO were carried out in strainMC1000 containing the GFP-LacI plasmid with the additionof the P1 cre recombinase gene on plasmid pSG20-cre(Gordon et al., 1997). Plasmids F-lacO and P1-lacO aredescribed in (Gordon et al., 1997). Strain SG101 carryingpFX234 (Lau et al., 2003), a plasmid similar to pSG20 thatcarries a yfp::tetR gene instead of gfp::lacI, was used forlocalization of mini-F plasmids carrying a tetO array. Theseplasmids were constructed by substituting ~10.2kb tetO arrayfragments excised from pFX240 (Lau et al., 2003) with Nhe1(blunt-ended with Klenow polymerase and dNTPs) and eitherSalI or HindIII for the 1564 bp Bsu36 (plus Klenow) – SalIfragment between the cat gene and sopC of pDAG114 (Ravinand Lane, 1999) to form pDAG479 or the 2047 bp Bsu36(plus Klenow) – HindIII fragment of pDAG115 (Lemonnieret al., 2000) to form pDAG480.

Growth conditions

Cells were grown in L broth containing, when necessary,ampicillin (50 mg ml-1), kanamycin (20 mg ml-1), tetracycline(20 mg ml-1) and/or chloramphenicol (20 mg ml-1). For rea-sons discussed in Gordon et al. (1997), cells were grown at20∞C for visualization of fluorescent foci. For microscopy,cells were grown overnight at room temperature using initialdilutions which assured an OD600 between 0.3 and 0.6. Induc-tion of the GFP-LacI fusion is described in Gordon et al.(1997) except that for time-lapse imaging, the induction timewas increased to at least 45 min.

Run-out synthesis

Run-out synthesis was quantified by adding rifampicin(150 mg ml-1) to cultures growing at 20∞C and 37∞C(OD600 @ 0.3), and sampling in duplicate for chemical estima-tion of DNA as described below.

Microscopy, imaging and measurements

Microscopy was performed essentially as described (Gordonet al., 1997), with the following exceptions. Images wereacquired using a highly light-sensitive Hammamatsu C4742-95 digital CCD camera, and an automatic light shutter(MAC2000, Ludl Electronics Products). OpenLab 2.0 imagingsoftware (v.2.02, Improvision) was used for image acquisitionand control of the light shutter. During time-lapse microscopy,phase-contrast imaging of cells was omitted in the interest of

468 S. Gordon, J. Rech, D. Lane and A. Wright

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speed, and cell lengths were determined from backgroundwhole-cell fluorescence; control experiments showed thatthis procedure underestimated cell length by about 0.1 mm.Measurements of the positions of GFP foci versus celllength were carried out using the Object-Image softwarepackage (which can be obtained at http://simon.bio.uva.nl/object-image.html). YFP foci were revealed using a GFP filterand captured using a Photometrics Coolsnap camera and itsassociated software.

Copy number measurement

Derivatives of strain SG101 carrying the GFP-LacI productionplasmid, pSG20, with or without F¢lacO+, were grown in L-broth as described by Gordon et al. (1997) at 20∞C and 37∞C.At OD600 @ 0.3, when sampling for fluorescence microscopynormally occurred, samples were taken for estimation of theF:oriC DNA ratio, chemical estimation of DNA and assay ofcell number. Total DNA was extracted and cleaved with PflMI,and the fragments were transferred to nylon membrane andhybridized with radioactively labelled probe DNAs asdescribed by Ravin and Lane (1999). The probe consisted ofa mixture of four PCR fragments, each containing 418 ATbase-pairs labelled with 32P to the same specific activity anddesigned to detect PflMI fragments of 4863 bp (E. coli chro-mosome 3735843–3740706; co-ordinates from GenBankaccession number U00096) near malS, 2938 bp (209434–212373) in ldc, 1287 bp (1442816–1444103) near feaR, and3253 bp (F plasmid 44108–47361; GenBank accession num-ber NC_002483) in the F replication region (repE); a PflMIsite at F 45019 is not cut in F DNA isolated from this strain.Bound radioactivity was quantitated by phosphorimaging andused to determine the F:oriC ratio and the gradient of chro-mosomal marker frequency.

Cellular DNA content was estimated using the dipheny-lamine colorimetric assay (Burton, 1956), with herring spermDNA as the standard. The method was essentially thatdescribed for L broth-grown cells by (Bipatnath et al., 1998),except that 7.5 ml culture samples were taken, and the finalreaction mixtures clarified by centrifugation and filtrationthrough used Qiagen kit miniprep columns. Cell numberswere estimated by electronic particle counting with a Coultercounter, Model ZM, using a 30-mm orifice.

Acknowledgements

We thank François-Xavier Barre for the Tet repressor/opera-tor plasmids and members of the group Dynamiques desreplicons bactériens for useful discussions. This work wassupported by USPHS grant GM055604 (A.W.) and by thesoutien de base du LMGM from the CNRS (D.L.).

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