The Replicase Sliding Clamp Dynamically Accumulates behind Progressing Replication Forks in Bacillus subtilis Cells

Molecular Cell

Article The Replicase Sliding Clamp Dynamically Accumulates behind Progressing Replication Forks in Bacillus subtilis Cells Masayuki Su’etsugu1 and Jeff Errington1,* 1Centre for Bacterial Cell Biology, Institute for Cell and Molecular Biosciences, Newcastle University, Richardson Road, Newcastle-upon-Tyne NE2 4AX, UK *Correspondence: [email protected] DOI 10.1016/j.molcel.2011.02.024

SUMMARY

The sliding clamp is an essential component of the replisome required for processivity of DNA synthesis and several other aspects of chromosome metabolism. However, the in vivo dynamics of the clamp are poorly understood. We have used various biochemical and cell biological methods to study the dynamics of clamp association with the replisome in Bacillus subtilis cells. We find that clamps form large assemblies on DNA, called ‘‘clamp zones.’’ Loading depends on DnaG primase and is probably driven by Okazaki fragment initiation on the lagging strand. Unloading, which is probably regulated, only occurs after many clamps have accumulated on the DNA. On/off cycling allows chromosomal zones of about 200 accumulated clamps to follow the replisome. Since we also show that clamp zones recruit proteins bearing a clamp-binding sequence to replication foci, the results highlight the clamp as a central organizer in the structure and function of replication foci.

INTRODUCTION Duplication of genomic DNA is a highly organized process performed by a multicomponent complex, the replisome (Baker and Bell, 1998; Johnson and O’Donnell, 2005). One of the essential components needed to coordinate this process is the sliding clamp. The architecture and function of the clamp are conserved in both prokaryotes and eukaryotes (DnaN dimer and PCNA trimer, respectively) (Indiani and O’Donnell, 2006). This ringshaped protein encircles DNA and is able to slide along it freely. The clamp loader loads the clamp ring onto DNA by opening and resealing it at the primer-template DNA. The primary function of the clamp is tethering the replicative DNA polymerases to DNA as a processivity factor. Besides this function, the clamp interacts with various proteins involved in many chromosomal metabolic processes, including DNA repair, Okazaki fragment maturation, and timely regulation of replication initiation (Katayama et al., 2010; Lo´pez de Saro, 2009; Moldovan et al., 2007). 720 Molecular Cell 41, 720–732, March 18, 2011 ª2011 Elsevier Inc.

In typical prokaryotic cells, including B. subtilis, their circular chromosome is replicated from a unique origin, oriC. Replication forks proceed bidirectionally from oriC and eventually meet in the ter zone. The replication process is well understood in E. coli, and its basic mechanism is conserved in B. subtilis (Kornberg and Baker, 1992; Sanders et al., 2010; Schaeffer et al., 2005). DnaA binds to oriC to initiate replication by unwinding the DNA. The replicative helicase (DnaB in E. coli and DnaC in B. subtilis) is then loaded and expands the zone of singlestranded DNA, onto which DnaG primase synthesizes an RNA primer. SSB stabilizes the single-stranded DNA. The primertemplate junction is recognized by the clamp loader complex, which then loads the DnaN clamp. At the replicaton fork, the clamp loader connects the helicase and the leading and lagging polymerases. Fork progression provides the antiparallel templates for the replicative DNA polymerases. Therefore, while leading-strand synthesis is basically continuous, the lagging strand is synthesized discontinuously in 1–2 kb Okazaki fragments (Ogawa and Okazaki, 1980). The initiation of each Okazaki fragment requires both the primer generated by DnaG and the loading of DnaN. After Okazaki fragment synthesis, the replicative polymerase hops from the original DnaN to a newly loaded DnaN (Stukenberg et al., 1994). DnaN is stable on DNA in vitro (half-life, t1/2 =
115 min) (Leu et al., 2000). Polymerase-free DnaN thus potentially remains on the DNA behind replication forks. Indeed, DnaN molecules remaining on DNA after lagging-strand synthesis have been detected in an in vitro rolling circle replication system (Yuzhakov et al., 1996). In principle, this can explain how different machineries can utilize DnaN without interfering with replication (Lo´pez de Saro et al., 2003). On the other hand, in vivo it is likely that DnaN molecules left on DNA are actively unloaded and recycled, since the cellular content of DnaN (
300–400 dimers per E. coli cell) (Leu et al., 2000) is much less than the number of Okazaki fragments needed for one round of chromosomal replication (2000–4000 fragments per 4.6 Mb chromosome in E. coli). In both eukaryotes and prokaryotes, many replisome components, including the clamp, have been reported to form subcellular foci (replication foci) visible by fluorescence microscopy (Lemon and Grossman, 1998, 2000; Leonhardt et al., 2000; Meile et al., 2006; Onogi et al., 2002). The replication reaction is thought to take place within these foci. In B. subtilis and E. coli cells, a single replication focus arises following replication initiation during slow growth. Because of the independent

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(A) Localization of GFP-DnaN in asynchronously replicating cells. Strain MS104 (gfp-dnaN) was grown in MMglyC medium at 30 C. The DAPI staining image of nucleoid DNA and the phasecontrast image are overlaid. Scale bar: 2 mm. (B) Localization of GFP-DnaN in the absence of replication. Strain MS114 (dnaB134ts, gfp-dnaN) was grown in MMglyC medium at 30 C and then shifted to 45 C for 2 hr. Cells were then immediately subjected to microscopic analysis. The phase-contrast image shows the outline of each cell. Scale bar: 2 mm. (C–E) After the completion of ongoing rounds of replication in (B), replication initiation was released on an agar pad at room temperature (25 C), and cells were analyzed by time-lapse microscopy. Typical still frames taken at 10 s intervals are shown in (C). Scale bar: 2 mm. Fluorescence signals in an area of the focus (indicated by white arrows) were quantified. An outline of the image is given in (D). As shown in (E), the ratio of the focus signal to the whole-cell signal was plotted over time. The dotted line represents the background value in the quantified area before focus formation.

Okazaki fragment synthesis. We have been studying the equivalent protein in B. subtilis, DnaN, and find that its behavior is quite different. Here, clamps remain associated with DNA for a protracted period of time, forming accumulations that dynamically track with the replisome and provide a landing platform, or ‘‘clamp zone,’’ to which other replisomeassociated proteins can associate. This may allow DnaN to play a pivotal role in the spatial organization of the B. subtilis replisome in vivo.

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Gradual Accumulation of GFP-DnaN in Foci after Synchronous Initiation of DNA Replication To investigate DnaN behavior in living cells, the dnaN gene was replaced with a gfp-dnaN fusion at its endogenous locus. The fusion protein was apparently fully functional, as judged by culture growth rate and DAPI staining. Fluorescence microscopy of the cells revealed the presence of GFP-DnaN foci, as previously reported (Meile et al., 2006) (Figure 1A). To check whether the foci depend on replication, initiation was arrested using a dnaB134 mutant that encodes a temperature-sensitive variant of the replication initiation protein DnaB (Mendelson and Gross, 1967). After 2 hr at the nonpermissive temperature, to allow run-out of ongoing rounds of replication, GFP-DnaN foci disappeared and the fluorescence became dispersed throughout the cell (Figure 1B). To investigate how GFP-DnaN foci form following replication

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progression of the bidirectional forks, E. coli cells have two separated foci during replication (Reyes-Lamothe et al., 2008). In contrast, in B. subtilis, the two forks seem to mainly colocalize, so that most cells exhibit a single focus or two closely spaced foci (Berkmen and Grossman, 2006; Migocki et al., 2004). After the completion of replication, all foci disappear. Reyes-Lamothe et al. (2010) have recently investigated the stoichiometry of various replisome components in single E. coli cells. For most of the proteins studied, the stoichiometry is very low (about three copies per replication fork). That of the sliding clamp was slightly higher (3–6 copies), suggesting that unloading of the clamp is tightly coupled to reinitiation of

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Figure 2. Formation of GFP-DnaN Foci in Wild-Type Cells (A and B) Strain MS104 (gfp-dnaN) was grown in MMgly medium at 30 C and analyzed by time-lapse microscopy. Typical still frames and the signal ratio are shown as in Figure 1. Cell generating a single focus (5 s intervals) is shown in (A); in (B), cell generating two foci (foci b and c) and the disappearance of a previous focus (focus a) (10 s intervals). The phase-contrast image was taken, and cell outlines are shown by dotted lines. Scale bar: 2 mm. (C and D) Cellular content of DnaN. Strains 168 (wild-type) and MS104 (gfp-dnaN) were grown to A600 = 0.2 in the same condition as in (A) and (B). Portions of culture (50 ml) were subjected to western blot analysis using anti-DnaN antiserum (C). Amounts of wild-type DnaN and GFP-DnaN were deduced from the

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initiation, the initiation arrest was released relatively synchronously by shifting the cells back to the permissive temperature, then the generation of GFP-DnaN foci was monitored by timelapse microscopy. Initially (70 s) (Figure 1C), a single, very weak focus appeared within a cell with otherwise dispersed fluorescence. The intensity of the focus then increased gradually, accompanied by a decrease in the dispersed fluorescence outside the focus, suggesting that protein in the cytoplasmic pool was being recruited to the focus. To facilitate quantitative measurement, the focus signal was normalized to the wholecell signal, which revealed that GFP-DnaN molecules accumulate linearly over a period of
2 min after focus appearance, and then the accumulation plateaus (Figure 1E). A Similar DnaN Accumulation Behavior in Wild-Type Replication Foci To test whether the above result held in a wild-type situation, we analyzed a dnaB+ gfp-dnaN strain. Cells were grown asynchronously in minimal medium with glycerol as the only carbon source (doubling time, Td = 110 min) to prevent new rounds of replication from overlapping with the previous round. Timelapse analysis revealed that DnaN foci tend to appear at roughly the ¼ and 3/4 positions in cells when they attain a length of approximately 1.33 the average cell length. Previous cell-cycle studies have shown that these cells would have two chromosomes (Sharpe et al., 1998), so that the events at ¼ and 3/4 positions represent initiation on sister segregated chromosomes. In most cells, initiation occurs on each chromosome within a few minutes of that of its sister (examples shown in Figures 2A and 2B, foci b and c). In rare cases, only one focus appeared during the short analysis period (Figure 2A). In other cells, two new foci appeared soon after the disappearance of a mid-cell focus, which presumably represented the dissociating focus from a previous round of replication (e.g., Figure 2B, focus a). Quantitative measurements again revealed a linear accumulation of fluorescent molecules, as in the synchronous experiment. Interestingly, in the cells generating two foci, the first focus to form (focus b) showed a partial decrease in intensity when the second focus (focus c) began to appear (Figure 2B). This suggests that DnaN molecules can equilibrate between foci in the same cell. By applying image quantitation to the appearance of 20 independent foci, we obtained a value of 8.4% ± 2.7% min1 (signal ratio of the focus to whole cell) for the accumulation rate of GFPDnaN molecules and a period of 2.8 ± 0.9 min. These results include data for foci generated alone as well as for foci generated in the presence of an earlier focus, as the kinetics of accumulation were indistinguishable. To be able to relate focus intensity to the number of DnaN molecules in the cell, we quantified the cellular content of GFP-DnaN. The gfp-dnaN strain (MS104) was grown under the same conditions as in the microscopy experiment described above, and samples of culture were analyzed to measure cell numbers and DnaN protein content.

Using purified His6-DnaN protein as a quantitative standard, the amount of GFP-DnaN was estimated to be
600 dimers/ cell (Figures 2C and 2D), similar to that for DnaN in wild-type cells (
500 dimers/cell). Assuming that the whole-cell fluorescence consists of 800 GFP-DnaN dimers during focus formation, based on the cells being 1.3-fold longer than average, the accumulation rate of DnaN was estimated as 67 ± 22 dimers/min. To test whether other replication proteins also showed a gradual accumulation, similar time-lapse analyses were performed for DnaX-YFP and PolC-GFP (Figure S1). DnaX is a subunit of the clamp loader and PolC is the replicative polymerase. Both fluorescence fusions form foci dependent on replication initiation, similar to GFP-DnaN (Lemon and Grossman, 1998). Although the signals were relatively weak, for both fusions the foci seemed to exhibit a single stepwise increase in fluorescence, quite distinct from the extended gradual increase in GFP-DnaN foci (Figure S1). Thus, gradual accumulation seems to be a special feature of DnaN. DNA Loading of DnaN Is Required for Focus Formation A simple explanation for the gradual accumulation of DnaN in replication foci would be that they arise by sequential loading on the chromosome during lagging-strand replication. To test this idea, we asked whether DNA loading of DnaN is required for focus formation. For the loading reaction, DnaN requires primer-template DNA, which is generated by DnaG primase (Yuzhakov et al., 1996). Therefore, we first tested whether focus formation depends on DnaG. The dnaG gene is located upstream of the essential sigA gene within the same operon (Wang et al., 1985). We constructed an allele of dnaG, controlled by the IPTG-inducible PspacHY promoter, and placed this at an ectopic locus. The N-terminal coding part of the endogenous dnaG was then deleted, leaving intact a promoter for the downstream genes within dnaG (Wang et al., 1985). The resultant construct showed a complete growth dependence on IPTG, as expected. As a convenient means of examining the effects of depletion of DnaG on formation of DnaN foci, we prepared spores (which contain fully replicated chromosomes) (Oishi et al., 1964) in the presence of IPTG. Spores can be germinated and outgrown in the absence of inducer to follow the effects of protein depletion on a relatively synchronous first round of DNA replication (Harry, 2001). In the presence of IPTG, the germinated spores showed normal GFP-DnaN foci, as expected (Figure 3A). In the absence of IPTG, outgrowth occurred normally, and a background fluorescence was evident, but no foci were detected (Figure 3B). This result indicates that DnaG is essential for the generation of DnaN foci, which supports the idea that DnaN focus formation depends on the DNA-loading reaction. To further assess this idea in replicating cells, we constructed a DnaN mutant deficient in DNA loading. It has previously been demonstrated that a ‘‘monomer’’ mutant of E. coli DnaN, which bears amino acid substitutions at the dimer interface, is impaired in the formation of dimer rings and is thus unable to stay on DNA

quantitative standard of purified His6-DnaN protein (His-DnaN). Asterisk (*) indicates a nonspecific band. A portion of the same culture was subjected to cell counting by plating (D). The cellular contents of DnaN and GFP-DnaN were calculated based on the calculated molecular mass of the DnaN dimer (84 kDa) and Avogadro’s number (6.02 3 1023 molecules/mol). See also Figure S1 for focus formation in cells bearing DnaX-YFP or PolC-GFP.

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(Stewart et al., 2001). As the equivalent residues are conserved in B. subtilis DnaN (Figure 3C), we constructed a mutant encoding the L280A and L281A substitutions and expressed the protein as a YFP-fusion under the control of the xylose-inducible promoter Pxyl at an ectopic locus in the presence of endogenous wild-type DnaN (Figure 3D). The YFP-DnaN monomer mutant showed no dominant effect on cell growth. The control construct ectopically expressing wild-type YFP-DnaN formed foci, as expected (Figure 3E). In contrast, overall fluorescence for the YFP-DnaN monomer mutant was mainly dispersed throughout the cell (Figure 3F). These results indicate that the DNA-loaded form of DnaN is required for focus formation. Faint foci of the monomer mutant were observed in some cells (Figure 3F), which could be due to an ability of the monomer mutant to associate with replisome components, such as the clamp loader (see below) (Stewart et al., 2001). Alterations in Primase Concentration Affect the DnaN Accumulation Rate The above results suggested that multiple DnaN molecules are loaded onto DNA and that these accumulate to form the foci. This idea is consistent with the in vitro data showing that one DnaN dimer is loaded for each Okazaki fragment initiated and that it remains on DNA after synthesis of the fragment is finished (Yuzhakov et al., 1996). The rate of synthesis of Okazaki fragments in two bidirectional forks is estimated to lie in the range of 30–60 fragments/min, given that the fragment size is
1–2 kb (Ogawa and Okazaki, 1980) and the replication rate per fork is 30 kb/min (Wang et al., 2007). This rate is comparable to the accumulation rate of DnaN (67 ± 22 dimers/min) measured in Figure 2, supporting the idea that the DnaN accumulation rate depends on the rate of initiation of Okazaki fragment synthesis. If correct, reducing the frequency of Okazaki fragment synthesis would result in a slower rate of DnaN accumulation. To test this, we used the DnaG-inducible strain described above. DnaG is required for the initiation of synthesis of each Okazaki fragment. An in vitro reconstituted system of replication has demonstrated that a decrease in DnaG concentration reduces the frequency of Okazaki fragment initiation (Sanders et al., 2010; Wu et al., 1992). We first investigated the IPTG concentration critical for the strain growth, in which DnaG concentration should be low enough to affect the frequency of Okazaki fragment synthesis. The strain grew normally when the IPTG concentration was decreased from 1 mM to 30 mM (Td = 120 min), but an effect on culture growth was detected at a concentration of 10 mM (Td = 280 min). At this IPTG concentration, some elongated cells with aberrant DnaN foci were observed, presumably due to inhibition of replication. However, in nonelongating cells, the overall intensity of the GFP-DnaN foci was similar to that in wild-type cells, and we were able to detect the focus formation process (Figure 3G). The signal ratio of focus to whole cell was then quantified over the focus formation process in ten cells (example shown in Figure 3H). The average revealed about a 3-fold decrease in the accumulation rate (2.6% ± 1.1% min1) and a concomitant increase of about 3-fold in the accumulation period (8.7 ± 3 min) in comparison to wild-type cells. These results suggest that the DnaN accumulation rate depends on the cellular activity of DnaG, probably via its action in initiating 724 Molecular Cell 41, 720–732, March 18, 2011 ª2011 Elsevier Inc.

Okazaki fragment synthesis. This then supports the idea that DnaN focus formation reflects the temporary retention of multiple DnaN molecules accumulated after sequential rounds of Okazaki fragment synthesis. Chromosomal Zones of DnaN Association Track with Replication Forks We next investigated the chromosomal distribution of DnaN during synchronous replication. To enable the application of a chromatin affinity precipitation (ChAP) assay, the dnaN gene was replaced with an allele encoding an N-terminal His tag. We also introduced the dnaB134 temperature-sensitive mutation with which to synchronize replication. To facilitate synchronization, rifampicin was added 10 min after the release of initiation arrest, which blocks further rounds of initiation (Laurent, 1973). Cells were then crosslinked at various times and collected. His-tagged DnaN was pulled down using Co2+-conjugated TALON beads, and then the amounts of DNA in the input and the pull-down fractions were analyzed by quantitative PCR with primer pairs specific for a locus near the origin (a), loci around the right and left arms of the chromosome (b and d, c and e, respectively), and close to the terminus (ter, f) (Figure 4A). Marker frequency measurements relative to the ter locus confirmed that replication forks progressed bidirectionally and relatively synchronously from the origin. The approximate times of duplication of each locus (Figures 4A and 4B) were consistent with expectation based on the speed of replication fork progression (see above) (Wang et al., 2007). Figure 4C shows the ChAP recovery normalized to input DNA. The results showed that DnaN first associates with a locus near the origin (a) and subsequently the locus moved bidirectionally (b-e) toward ter. For each locus tested, a peak of recovery was observed at about the same time as fork traverse detected in the maker frequency analysis. Although the precision of these data is relatively low, they are consistent with expectation that the chromosomal zones of DnaN association move in parallel with replication fork progression. We further assessed the localization of DnaN foci relative to replication forks by comparing their position in cells with that of the DnaC helicase using a strain containing DnaN-mCherry and GFP-DnaC. The results revealed that most DnaN foci lay close to or coincident with DnaC foci (Figures 4D–4F). Similar results were obtained when CFP-DnaN was compared with DnaX-YFP as a marker for fork position (Figures 4G–4I). These results are consistent with the idea that DnaN foci follow replication fork progression. Replication Fork Arrest Leads to Dispersion of DnaN To begin investigating the mechanism of removal of clamps from DNA, we used 6-hydroxy-phenylazo-uracil (HPUra), a specific inhibitor for DNA polymerase, PolC (Brown, 1970), to arrest replication. Immediately after the addition of HPUra (2 min), the intensity of GFP-DnaN foci was significantly reduced, and the dispersed fluorescence became evident outside of the foci (Figure 5B). Prolonged incubation in the presence of HPUra did not dissipate the weak foci (60 min) (Figure 5B). We also performed time-lapse analysis, and the signal ratio of focus to whole cell was quantified for cells with a single focus (Figures 5C and 5D).

Molecular Cell Cellular Dynamics of Sliding Clamp

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(A and B) DnaG dependence of DnaN focus formation in outgrowing spores. Spores of strain MS242 (gfp-dnaN, amyE::PspacHY-dnaG, DdnaG) were geminated and outgrown in germination medium in the presence (A) or absence (B) of 1 mM IPTG. The phase-contrast image reveals the outline of each cell. Scale bar: 2 mm. (C–F) Localization of the dimerization-deficient DnaN mutant protein. Alignments of amino acid residues flanking the mutation sites in E. coli (Ec) and B. subtilis (Bs) DnaNs are shown in (C). Schematic view of this experiment is given in (D). Strains MS320 (amyE::Pxyl -yfp-dnaN) (E) and MS328 (amyE::Pxyl -yfp-dnaNmono) (F) were grown in MMgly medium in the presence of 0.5% xylose at 30 C. The phase-contrast image reveals the outline of each cell. Scale bar: 2 mm. (G and H) A decrease in the DnaG concentration slows the rate of DnaN accumulation. Strain MS242 (gfp-dnaN, amyE::PspacHY-dnaG, DdnaG) was grown in MMgly medium in the presence of 10 mM IPTG at 30 C. Focus generation was analyzed by time-lapse microscopy at 20 s intervals, as in Figure 1. An image of 5 s exposure was taken after the time lapse (long exposure), and cell outlines are shown by dotted lines. Scale bar: 2 mm.

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2009). This idea is supported by the observation that the weak foci of the YFP-DnaN monomer mutant seen in Figure 3F were not dispersed by HPUra (Figure 5E). We also confirmed that the dispersion effect is specific for DnaN using other replisome components, YFP-SSB and GFP-DnaC, the foci of which did not disperse (Figures 5F and 5G). These experiments show that most DnaN molecules can dissociate from replication foci in the absence of elongation, suggesting an unloading mechanism whereby DnaN foci are maintained at an equilibrium level while fork progression continuously generates new loading sites for DnaN (Figure 5A).

Dynamic Turnover of DnaN Molecules within Replication Foci If DnaN molecules are continually loaded 1200 1400 and unloaded within foci during replication, this dynamic behavior ought to be detectable by fluorescence recovery after photobleaching (FRAP) analysis. We tested this using a strain harboring a yfp-dnaN fusion under the control of the Pxyl promoter at an ectopic locus and the endogenous dnaN under the control of the IPTG-inducible Pspac promoter. The strain was grown in a minimal medium with a long doubling time (Td = 110), in the presence of xylose but absence of IPTG, Molecular Cell 41, 720–732, March 18, 2011 ª2011 Elsevier Inc. 725

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Time after +Rif (min) Figure 4. Localization of DnaN Foci Relative to the Replication Fork (A) B. subtilis chromosomal loci chosen for analysis. The distance (kb) from oriC and the estimated time (min) of fork traverse are shown. (B and C) Strain MS137 (dnaB134ts, his12-dnaN) was grown at 30 C and then shifted to 45 C for 1.5 hr. After initiation release at 30 C for 10 min, 250 mg/ml rifampicin was added (time = 0). Cells were collected at the indicated times and subjected to marker frequency analysis to monitor the fork progression (B) and ChAP analysis (C). The experiment was repeated twice with different time points for different loci, and a representative data set is shown. (D–I) Strains MS157 (dnaN-mCherry, gfp-dnaC) (D–F) and MS110 (cfp-dnaN, dnaX-yfp) were grown in MMgly medium at 30 C. The images shown are: mCherry (D), GFP (E), overlay of mCherry (red) and GFP (green) (F), CFP (G), YFP (H), and overlay of CFP (blue) and YFP (yellow) (I). Scale bar: 2 mm.

to express only the YFP-fusion version of DnaN. The cells were then analyzed in the absence of xylose to minimize the effect of de novo synthesis of YFP-DnaN. Figures 6A–6D illustrate representative experiments of many repeats. Figure 6A shows a cell with a single YFP-DnaN focus 726 Molecular Cell 41, 720–732, March 18, 2011 ª2011 Elsevier Inc.

(the focus temporarily became two closely spaced foci, presumably due to separation of the two forks; note that in B. subtilis, the left and right forks mostly remain close together, in contrast to E. coli) (Berkmen and Grossman, 2006; Migocki et al., 2004; Reyes-Lamothe et al., 2008). After bleaching the focus area,

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Figure 5. Replication Arrest Disperses DnaN Foci (A) Model for translocation of the DnaN accumulation zone. (B–D) Strain MS104 (gfp-dnaN) was grown in MMgly medium at 30 C. Cells were taken before (HPUra) and 2, 20, and 60 min after the addition of 47 mg/ml HPUra and subjected to microscopic analysis using 10 s exposure times (B). Two minutes after the addition of HPUra, time-lapse analysis using 200 ms exposure times was started (+HPUra) (C and D). A control analysis was done with no HPUra (HPUra). Typical still frames, with phase-contrast image showing the outlines of each cell, are shown (C). Scale bar: 2 mm. The signal ratio of focus to whole cell was quantified for cells with a single focus (n = 10) (D), and the average was plotted with error bars representing standard deviations. To deduce a dispersion rate, the background ratio was subtracted (dashed line), and the data were fitted to an exponential curve (gray line). (E–G) Strains MS328 (amyE::Pxyl-yfp-dnaNmono) (E), MS151 (yfp-ssb) (F), and MS150 (gfp-dnaC) (G) were grown in MMgly medium (supplemented with 1% xylose for MS328) at 30 C and analyzed before (HPUra) and 20 min after (+HPUra) the addition of 47 mg/ml HPUra. The phase-contrast image shows the outline of each cell. Scale bar: 2 mm.

the focus intensity fell precipitously, as expected, and then recovered, but only partially (Figures 6A and 6B). As a control, the focus in a nonbleached cell in the same field showed only moderate fading (0.25% per image acquisition), and the data corrected using this rate are shown in Figure 6B (blue circles) and the following panels. In cells with a single DnaN focus, each focus represented
40% of the cellular DnaN (see Figure 2A), and therefore more than half (56%) of the total cellular fluorescence was extinguished when an area covering the focus and some background were bleached in this experiment (see ‘‘total cellular fluorescence’’ under panel A). Thus, the limited recovery would be due to the extensive loss of total cellular fluorescence.

To lessen the cellular loss of fluorescence, we analyzed a cell in which initiation was ongoing on two chromosomes and thus with two well-separated foci (Figures 6C–6E). In this case, each focus had a lower relative content of DnaN molecules (20%–30% of total cellular molecules) (Figure 2B) than for cells with single foci. After bleaching of one of the two foci (focus a), the loss of total cellular fluorescence was moderate (28%) (Figure 6C), and the subsequent recovery of focus intensity was higher than for the single-focus cell, as expected (Figure 6D, blue circles). Interestingly, the nonbleached focus (focus b) in the same cell showed a decrease in intensity, approximately in parallel with the recovery of the bleached focus (Figure 6D, red circles). The values of both foci then stabilized after
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Figure 6. DnaN within Foci Undergoes a Continuous Unloading-Reloading Cycle (A–E) FRAP/FLIP behaviors of DnaN. Strain MS356 (amyE::Pxyl-yfp-dnaN, dnaN::Pspac-dnaN) was grown in MMgly medium in the presence of 0.01% xylose at 30 C and mounted on agar pads with no xylose. Cells with a single DnaN focus (A and B) or with two foci (C–E) were analyzed. Dotted circles indicate the bleached areas, and recovery was monitored at 20 s intervals. Typical still frames are shown (A and C). Total cellular fluorescence before (20 s) or just after (0 s) bleaching was quantified, and the relative values are shown. Long exposure images (5 s) were taken at the end of each time-lapse series, and the outline of each cell is indicated. Scale bar: 2 mm. Relative fluorescence intensity of the bleached focus (B and D, blue circles) or the adjacent nonbleached focus in the same cell (red circles) were plotted over time. For cells with two foci, the average of 15 experiments is also shown (E), with error bars representing standard deviations. Five foci in nonbleached cells were also analyzed to obtain their fading rate during image acquisition (0.25% per acquisition as an average). The data were corrected using this rate. (F) Computer models for FRAP and FLIP behavior in cells with a single focus or two foci. DnaN molecules on (fON and nfON) or off (fOFF and nfOFF) DNA are modeled in the focus (circle) or cell (ellipse) areas, respectively. OFF molecules are allowed to diffuse. The equations used for the fluorescent molecules in the single focus model are shown. Laser treatment was modeled by changing fluorescent molecules (fON and fOFF) in the dotted circle area to nonfluorescent molecules (nfON and nfOFF). The simulation was performed using parameter values of Koff = 0.0028 s1 for the single-focus model or 0.0057 s1 for the two-foci

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which time the two foci had almost equivalent intensities. Similar results were obtained in a series of experiments, and an average of 15 experiments is shown in Figure 6E. This FLIP (fluorescence loss in photobleaching) behavior shows that two foci in the same cell can exchange their DnaN molecules. Taken together, these observations support the idea that DnaN molecules undergo dynamic turnover in the foci. Mathematical Modeling of Cellular Dynamics of DnaN To test whether the FRAP data kinetically support the continuous loading-unloading action of DnaN, a two-dimensional computer simulation was performed (Figure 6F). The DNA-free (OFF) and loaded (ON) molecules were modeled in cell (ellipse) and focus (circle) areas, respectively. Given that DnaN linearly accumulates into foci with a rate limited by primer synthesis (Kpri,
1 molecule s1), the loading reaction rate (JON) can be described by the equation JON = Kpri. For the unloading reaction rate (JOFF), we assumed a mass action reaction described by the equation JOFF = Koff $[ON]. The value for [ON] at equilibrium (JON = JOFF) was estimated from the intensity of prebleached foci in the experiment, from which a value for Koff was obtained using the equation Koff = Kpri / [ON]. This estimation gave a reasonable Koff value (0.0028–0.0057 s1, t1/2 = 2–4 min) that explains the kinetics of focus dispersion in the replication arrest experiment (see Figure 5D). We also assumed that bleached molecules compete with fluorescent molecules for the loading site. The mathematical model simulated well the limited recovery of focus intensity observed in the cell with a single focus (Figure 6B, cyan line). Also, in the case of cells with two foci, the simulated behaviors of FRAP and FLIP both fitted reasonably well to the experimental data (Figure 6E, cyan and pink lines). When the DnaN accumulation process was simulated, the mathematical model gave a nonlinear pattern (Figure 6G, cyan line), in contrast to the experimental data (Figure 2A: data overlaid as gray circles in Figure 6G). This implies that the unloading reaction is regulated such that DNA-loaded clamps accumulate until a certain level on DNA is reached, or the free pool of clamps falls below a threshold level. DnaN Recruits GFP Fusions Bearing a Clamp-Binding Sequence to Replication Foci Recent work on DNA replication proteins of E. coli in vivo showed that replication foci contain small numbers of replication protein subunits and, in particular, only 3–6 copies of the sliding clamp (Reyes-Lamothe et al., 2010). The situation is clearly different in B. subtilis. What might be the reason for the apparently excessive accumulation of DnaN at the replisome? One possibility was that the patch of DnaN molecules tracking with the replication fork could provide a molecular landing pad with which to recruit proteins involved in transactions with newly replicated DNA to relevant sites on the chromosome. In both E. coli and B. subtilis, many DnaN-binding partners share a five amino acid sequence (consensus: QL[S/D]LF), which is termed the clamp-binding

sequence (CBS) (Dalrymple et al., 2001). We therefore tested whether fusion of a CBS peptide (QLSLF) would enable GFP to localize at DnaN foci. Genes encoding either N- or C-terminal fusions of the CBS to GFP (CBS-GFP or GFP-CBS, respectively) were placed at an ectopic locus under the control of the Pxyl promoter. For both constructs, expression of the GFP derivative (presence of xylose) had no detectable affect on culture growth. Microscopic analysis revealed weak foci for both CBS-GFP and GFP-CBS (Figures 7A and 7B). In both cases, background fluorescence in the cells was evident, presumably due to their low affinity for DnaN, and its level was not reduced by a decrease in the xylose concentration. Examination of a strain containing DnaN-mCherry and GFP-CBS revealed almost complete colocalization of their respective foci (Figures 7D–7F). To test the specificity of the putative DnaN-CBS interaction, two residues in the CBS were substituted with alanine (ALSAF) in the CBSGFP construct. As expected, the resultant CBSmut-GFP was impaired for focus formation (Figure 7C). We next tested whether the CBS-GFP foci depend on DnaN foci. Replacement of the endogenous promoter of dnaA-dnaN operon by the Pspac promoter allows cells to be depleted for DnaA and DnaN, resulting in the disappearance of DnaN foci (Figures 7G and 7H) (Meile et al., 2006). Removal of IPTG also resulted in disappearance of CBS-GFP foci (Figures 7I and 7J), supporting the idea that the CBS provides a means of recruiting proteins to replication foci via interaction with accumulated DNA-loaded molecules of DnaN. Conclusion Our results suggest the following activity cycle for the DnaN clamp in vivo. After replication initiation, fork progression constantly generates primed sites for Okazaki fragment synthesis, onto which clamps are loaded. Since each clamp temporarily remains on DNA after fragment synthesis, they accumulate linearly behind replication forks with a rate comparable to that of Okazaki fragment initiation (
1 clamp/s). Because the diffusion of clamps on DNA is restricted by other proteins bound to DNA (Stukenberg et al., 1991) and cellular DNA is highly condensed, the clusters of molecules generate discrete subcellular foci (clamp zones). The accumulation then plateaus as a result of the clamp unloading reaction coming into equilibrium with the constant loading driven by fork progression. At this point, chromosomal zones of clamp accumulation follow replication forks, dynamically maintaining a constant level of accumulation. How does clamp unloading occur? In E. coli, given the relatively tight stoichiometry (Reyes-Lamothe et al., 2010), clamp unloading is presumably coupled to reinitiation of Okazaki fragment synthesis. However, our results show that in B. subtilis, clamps remain associated with DNA during an extended period, and that there may be an active unloading mechanism in vivo. The replication arrest experiment and the in silico simulation of the FRAP data show that clamps dissociate with a rate of

model and Kpri = 0.008 mM/s for both models. Cyan and pink lines in (B) and (E) show the behaviors of fON in the simulations compared with the corresponding experimental data. (G) Computer simulation of the DnaN accumulation behavior. The Koff value was determined from the plateau level of the experimental data (0.0042 s1). After the simulation, the signal ratio of focus to whole cell was plotted (cyan line) and was compared with the experimental data of Figure 2A (gray circles).

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Figure 7. Formation of DnaN-Dependent Foci by GFP Fusions Bearing a CBS Peptide (A–C) Strains MS303 (amyE::Pxyl-cbs-gfp) (A), MS307 (amyE::Pxyl-gfp-cbs) (B), and MS304 (amyE::Pxyl-cbsmut-gfp) (C) were grown in MMglyC medium at 30 C in the presence of 1% xylose. The ratio of cells with foci (n = 250–300) is shown in each panel. Scale bar: 2 mm. (D–F) Colocalization of GFP-CBS with DnaN-mCherry. MS354 (amyE::PspacHY-gfp-cbs, dnaN-mCherry) was grown in MMglyC medium at 30 C in the presence of 1 mM IPTG. Shown are: the GFP image (D), the mCherry image (E), and overlay of the GFP (green) and the mCherry (red) images (F). Scale bar: 2 mm. (G–J) The DnaN dependence of focus formation by CBS-GFP. DnaA and DnaN were depleted as described previously (Meile et al., 2006). Stains MS111 (gfp-dnaN, dnaA::Pspac-dnaA) (G and H) and MS305 (amyE::Pxyl-cbs-gfp, dnaA::Pspac-dnaA) (I and J) were grown in LB medium at 30 C in the presence of 0.5 mM IPTG (G and I). After the removal of IPTG, incubation was continued for another 3 hr (H and J). For CBS-GFP expression, 1% xylose was added. The GFP image is overlaid with the phase contrast image (G and H). Scale bar: 2 mm.


0.004 s1 (t1/2 =
3 min), which contrasts with the in vitro stability of the clamp on DNA (t1/2 =
115 min) (Leu et al., 2000). The mathematical modeling implies that the unloading reaction during clamp accumulation is not a simple mass action mechanism but regulated. One possibility is that DNA-free clamps serve to limit unloading by titrating unloader molecules until the free pool of clamps falls. The clamp loader and its d subunit, which can unload the clamp in vitro, have affinity for DNA-free clamps (Leu et al., 2000). Alternatively, one or more clamp-binding proteins might support this regulation by protecting clamps from attack by the unloader, which would allow a certain level of clamps to accumulate readily on DNA. In any case, regulation of unloading could ensure that multiple replication forks in the same cell share out the pool of available clamps relatively evenly. Indeed, cells initiating replication of a second sister chromosome show a reduction in clamp accumulation at the first-formed focus (Figure 2B). Finally, we demonstrated that the clamp recruits clampbinding proteins to the discrete cellular locations where DNA replication takes place. Hence, the moving window of clamp accumulation, or clamp zone, might serve as a molecular landing pad with which to organize various protein tools into a factorylike higher-order structure at the replication site. In E. coli, the Dam methylation system serves to differentiate newly synthesized DNA for DNA processing reactions and to help regulate 730 Molecular Cell 41, 720–732, March 18, 2011 ª2011 Elsevier Inc.

timely initiation of replication (Løbner-Olesen et al., 2005). Although this system is not conserved in B. subtilis, our results raise the possibility that the clamp zones also serve as temporary molecular markers for newly synthesized DNA. EXPERIMENTAL PROCEDURES Media and Strains Routine selection and maintenance of E. coli and B. subtilis strains was done on nutrient agar plates (Oxoid) or in Luria-Bertani (LB) medium. The B. subtilis strains used and their construction are listed in the Supplemental Experimental Procedures. B. subtilis cells were grown in Spizizen minimal medium supplemented with 6 mM MgSO4, 1.1 mg/ml FeCl3, 5 mM MnSO4, 0.5% glycerol (MMgly), or MMgly containing 0.1% glutamate (MMglyC). Supplements were further added as required: 20 mg/ml tryptophan, 40 mg/ml phenylalanine, 20 mg/ml thymine, 100 mg/ml ampicillin, 5 mg/ml chloramphenicol, 50 mg/ml spectinomycin, 15 mg/ml tetracycline, and 1 mg/ml erythromycin. Epifluorescence Microscopy To visualize cells from exponentially growing cultures, overnight cultures were diluted 100-fold in fresh medium and grown for at least three generations at the indicated temperature. Cells were mounted onto agar pads and analyzed as described previously (Murray and Errington, 2008). To visualize nucleoids, the DNA was stained with 2 mg/ml 40 -6-diamino-2-phenylindole (DAPI) (Sigma). For time-lapse microscopy, cells were mounted onto 1.2% agar pads (containing Spizizen minimal medium supplemented with 6 mM MgSO4, 1.1 mg/ml FeCl3, 0.5% glucose, and 0.02% casamino acids), immobilized within a Gene Frame (ABgene), and immediately analyzed in a 25 C–27 C room. Images

Molecular Cell Cellular Dynamics of Sliding Clamp

were analyzed with ImageJ (http://rsb.info.nih.gov/ij). Total fluorescence signals in the focus area and in the whole-cell area were quantified. After subtraction of the background signal in a blank area, the ratio of focus to whole cell was calculated.

ChAP and Marker Frequency Analyses ChAP assays were performed essentially as described by (Ishikawa et al., 2007) and are described in detail in the Supplemental Experimental Procedures. In brief, after crosslinking, cells and chromosomal DNA were disrupted by sonication. His12-DnaN and its bound materials in cleared extracts were collected using Dynabeads TALON (Invitrogen). After dissociation of crosslinks, input and elute DNAs were purified and subjected to quantitative PCR analysis. The quantitative PCR values of input DNAs were also used for marker frequency analysis.

FRAP/FLIP Analysis Exponentially growing cells were mounted on agar pads as described for timelapse microscopy. The experiments were performed in a 25 C–27 C room using a Nikon Eclipse Ti inverted microscope system coupled to a laser source, equipped with a Yokogawa spinning disc (CSU22) and Coolsnap HQ2 camera (Photometrics). Fluorescence images were taken through a Plan Apochromat objective lens (1003/1.40 numerical aperture) using a 491 nm laser, and the focus was maintained using the Perfect Focus System (Nikon). DnaN foci were bleached using a 200 ms pulse from a 405 nm laser. Pre- and postbleach images were acquired using 500 ms exposure times. The fluorescence intensity in the focus area was normalized to that before bleaching after subtraction of the background intensity.

Computer Simulation Computer simulations were created using Virtual Cell software (http://www. vcell.org). A cell was modeled as an ellipse (3.6 mm height and 0.8 mm width). Foci were modeled as a single circle or two circles (0.25 mm radius) at the center or quarter positions of the ellipse, respectively. The cellular concentration of DnaN (0.5 mM) was estimated from the western blot measurement in Figure 2D. To estimate the initial concentrations of the ON and OFF molecules, the signal ratio of focus (ON) to whole cell (ON + OFF) was quantified from a fluorescence image of the prebleaching cell. ON molecules were spatially associated with the foci area. OFF molecules were allowed to diffuse throughout the cell with a rate constant of 2.5 mm2/s, which was assumed from that of MBP-GFP protein in E. coli cells (Elowitz et al., 1999). A laser covering the focus area was modeled as a circle (0.4 mm radius), which bleaches fluorescent molecules in the area within 200 ms with magnitude similar to that in the experiments. Simulations were performed with 100 ms time steps and 0.072 mm spatial meshes.

SUPPLEMENTAL INFORMATION Supplemental Information includes Supplemental Experimental Procedures, Supplemental References, and one figure and can be found with this article online at doi:10.1016/j.molcel.2011.02.024.

ACKNOWLEDGMENTS We thank Jan-Willem Veening for strains. We thank Heath Murray and Lin Juan Wu for critical reading of the manuscript. Virtual Cell software is supported by the National Resource for Cell Analysis and Modeling. This work was supported in part by a grant from the European Research Council (250363) to J.E. M.S. was supported by a Postdoctoral Fellowship for Research Abroad from the Japan Society for the Promotion of Science (JSPS) and a Research fellowship from the Uehara Memorial Foundation. Received: December 17, 2010 Revised: January 25, 2011 Accepted: February 23, 2011 Published: March 17, 2011

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