Stoichiometry and Architecture of Active DNA Replication Machinery in Escherichia coli

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Science  23 Apr 2010:
Vol. 328, Issue 5977, pp. 498-501
DOI: 10.1126/science.1185757

Forking Replisomes

Replisomes are multiprotein machines that replicate DNA. Significant insight into how they work comes from in vitro studies, but how replisomes are organized in living cells has remained unclear. Reyes-Lamothe et al. (p. 498) have watched the replisome in living Escherichia coli cells using single-molecule fluorescence spectroscopy with millisecond time resolution. Cells expressing fluorescent derivatives of 10 different replisome components revealed both the stoichiometry and spatial distribution of the components at active replication forks in Escherichia coli. A similar technique could be used to study other molecular machines as they function.


The multiprotein replisome complex that replicates DNA has been extensively characterized in vitro, but its composition and architecture in vivo is unknown. Using millisecond single-molecule fluorescence microscopy in living cells expressing fluorescent derivatives of replisome components, we have examined replisome stoichiometry and architecture. Active Escherichia coli replisomes contain three molecules of the replicative polymerase, rather than the historically accepted two. These are associated with three molecules of τ, a clamp loader component that trimerizes polymerase. Only two of the three sliding clamps are always associated with the core replisome. Single-strand binding protein has a broader spatial distribution than the core components, with 5 to 11 tetramers per replisome. This in vivo technique could provide single-molecule insight into other molecular machines.

Replisomes are dynamic multiprotein machines that replicate DNA by copying the leading-strand template continuously and the lagging-strand template discontinuously. In E. coli the replisome couples the activities of more than 11 proteins during genome replication (1, 2). The DnaB helicase, loaded onto the lagging-strand template, separates the two templates that are subsequently copied by Pol III polymerase (αεθ). Pol III processivity results from binding to a sliding clamp (β) encircling duplex DNA; sliding clamps are added and removed by a clamp loader [(τ/γ)3δδ′ψχ] whose τ component oligomerizes Pol III. Unwound DNA on the lagging-strand template is bound by single-strand binding protein (Ssb) tetramers that remove DNA secondary structure and protect against nucleases. Primase binds to helicase during cycles of priming and DNA synthesis on the lagging-strand template.

In vitro studies of the replisome have yielded details of replisome organization and the replication mechanism but have not revealed how replication is organized within living cells. Additional copies of known replisome components may be present at replication forks, whereas additional cellular factors, absent from in vitro assays, may modify the composition of the replisome and act in DNA processing. Furthermore, the techniques used to determine stoichiometry and architecture in vitro favor strong interactions, potentially biasing estimates on numbers and interactions, and are subject to complications if any component is proteolytically sensitive. We thus investigated active replisome architecture in living cells by means of a fluorescence microscopy protocol with single-molecule sensitivity and millisecond temporal resolution.

Using fully functional fluorescent C- or N-terminal YPet derivatives of E. coli replisome components expressed from their endogenous promoters, we showed previously that the two spatially separable sister replisomes derived from a single initiation event at the replication origin, oriC, track independently along DNA (3). Using these and additional fusions, we analyzed 10 components of individual replisomes by “slimfield” fluorescence microscopy, which uses a compact Gaussian laser excitation field (~30 μm2) that encompasses single cells with an excitation intensity ~100 times that of wide-field fluorescence (4, 5). This imaging allows quantitative detection of single fluorescent molecules at 3-ms capture rates (Fig. 1, A and B). The high laser excitation intensity does not abolish DNA replication, as judged by a factor of 2 difference in turnover on DNA of Ssb-YPet in replicating and nonreplicating cells exposed to an excitation intensity and duration similar to that of slimfield microscopy (fig. S1), consistent with the demonstration that similar exposures do not inhibit flagellar rotation (4).

Fig. 1

Slimfield microscopy and photobleach analysis. (A) Slimfield schematic. A laser underfills the back aperture of an objective lens, generating an intense Gaussian field at the sample large enough to image single E. coli. (B and C) Overlaid bright-field (gray) and 90-ms frame-averaged fluorescence images (yellow) of ε-YPet strain; arrows indicate spots with a stoichiometry of ~3 [cyan, (B)] and ~6 [red, (C)] ε-YPet molecules, with corresponding single 3-ms frames taken after 45 ms, showing that stochastic photobleaching generates different brightnesses. (D) Raw intensity (blue) and filtered data (red) for a putative single (left panel) and double (right panel) replisome spot with surface-immobilized YPet in vitro; arrows indicate the 45-ms point. (E) Fourier spectral analysis for a photobleach trace of the ε-YPet strain with mean ± SD peak indicated for brightness of a single YPet.

For estimation of stoichiometry, image frames were averaged over 90 ms to define “regions of interest” that are hotspots for localization of a given YPet fusion protein. The position, size, shape, and intensity of the spots were measured automatically for each individual image frame, generating step-like intensity traces as photobleaching occurred (Fig. 1, D and E). We measured the step spacing with the use of an edge-preserving filter (6, 7) combined with Fourier spectral analysis and compared these with intensity traces from purified YPet in vitro (810); the in vivo steps were approximately integer multiples of the in vitro intensity of a single YPet molecule (4, 5, 813). Thus, intensities prior to photobleaching enabled single-molecule stoichiometry determination (5).

For most replisome components, we observed bimodal 1:2 distributions of stoichiometries, reflecting the observation that ~75% of cells contain two spatially separated replisomes, each associated with independent forks, while ~25% have sister replisomes separated by a distance smaller than the diffraction limit of our system (~250 nm) and are thus observed as a single spot (3). Lower stoichiometries were always associated with spots from cells containing two spots; cells with a single spot displayed doubled stoichiometries (Fig. 2). With the ε-YPet strain we observed a bimodal distribution with two Gaussians, peaks centered on 2.9 ± 1.0 (mean ± SD, SEM ~ 0.2) and 6.0 ± 1.3 molecules, consistent with a model in which a single fork contains three copies of ε, the proofreading exonuclease of the core polymerase. The Pol III catalytic subunit, α, had a bimodal distribution with peaks centered on 3.1 ± 1.1 and 5.8 ± 1.1 molecules. Similarly, the clamp loader component, τ, which also oligomerizes the polymerase, had peaks at 3.1 ± 0.8 and 5.6 ± 0.8 molecules. An essential component of the clamp loader, δ, expected to be present in one copy, had peaks at 1.0 ± 0.4 and 2.0 ± 0.4 molecules. Furthermore, we observed a stoichiometry of ~6 and ~12 molecules for DnaB (fig. S2), as expected for the hexameric helicase.

Fig. 2

Stoichiometries of replisome components and spatial distributions. Upper panels: Stoichiometry distributions per spot, using unbiased kernel density estimation for different E. coli strains (N = 27 to 51 cells in each data set); shown are two-Gaussian fits (black) with contributing single-Gaussian curves (red and blue) and mean ± SD of Gaussian peaks. Insets show examples of overlaid bright-field (gray) and single 3-ms fluorescence images (yellow) for each; arrows indicate foci in cells containing two (cyan) and one (red) replisome. Lower panels: False-color contour plots for 2D averaged spatial distributions for each strain (N = 42 to 151 spots in each data set). Estimates for mean FWHM 〈σ〉 of a symmetrical 2D Gaussian fit and the ratio σxy of the FWHM for the 1D Gaussian fits through the mean spot parallel to the x and y axes are indicated; SD errors are in parentheses.

Nonessential χ and ψ, which heterodimerize, have been reported to be present at one copy per replisome because of ψ binding to γ/τ in the clamp loader (1, 14). Instead, we found a mean of ~4 copies per single replisome spot (Fig. 2); single-spot cells had ~8 molecules per spot, with a broader distribution of stoichiometries than for other low-copy components. We propose that this results from one χψ heterodimer being tethered to the clamp loader while the other χψ dimers bind available C-terminal tails of the same or different Ssb tetramers by a characterized interaction through χ (14). The intensity of ψ-YPet foci is greatly reduced when χ is absent, but χ focus intensity is unchanged when ψ is absent, supporting this hypothesis (fig. S3).

dnaX expresses comparable amounts of τ and a truncated form, γ, formed by a programmed frameshift (15); τ and γ can interchangeably be clamp loader constituents, but only τ oligomerizes Pol III (1). Our demonstration that three copies of τ are associated with the replisome suggests that γ is not associated with the single clamp loader in most replisomes. To test this, we constructed two strains: dnaX), which did not express γ because the frameshift was abrogated, and dnaX(γ-YPet), which expressed γ-YPet. The strain that failed to express γ grew well, confirming that γ is nonessential (16), while the strain expressing γ-YPet showed fluorescent replisome-associated foci (fig. S4A). Because γ can interact with χψ, we considered whether γ might be Ssb-associated via a linking interaction with the χψ heterodimers not associated with the clamp loader (Fig. 3). Τargeted proteolysis of degron-tagged ψ, or deletion of χ or ψ, led to a loss of replisome-associated γ-YPet foci, but not ε-YPet or τ-YPet foci; the remaining γ-YPet foci were of reduced intensity (fig. S4B). Thus, the single replicative clamp loader in each replisome contains τ but not γ, and the clamp loader–independent copies of χψ associated with Ssb are likely to recruit γ to Ssb in addition to acting in primase release (17). We propose that γ may replace τ as a clamp loader component in postreplication repair-associated events at the replication fork. Comparison of τ stoichiometry in dnaX and dnaX) strains showed a ~30% increase when γ was absent (fig. S5), indicating that γ and τ can compete for binding to Ssb.

Fig. 3

Schematic model for replisome components. (A) Two engaged polymerases and one of the three β clamps at a distance from the core replisome (circle of diameter 50 nm shown in gray). The data indicate that ~75% of replisomes have this organization, whereas ~25% have all three β clamps associated with the core replisome and potentially associated with active Pol III. (B) Expanded view of clamp loader (τ3δδ′ψχ) and three additional molecules of χψ interacting with Ssb tails. The χψ heterodimer bound to the clamp loader may also contact Ssb (14); γ (shown as a trimer, but the stoichiometry is unknown) then interacts with Ssb-associated χψ.

In contrast to the structural skeleton components of the replisome, Ssb showed a broad distribution of stoichiometries (Fig. 2) with a periodicity of ~4 molecules (fig. S6) and a mean of 31.8 ± 11.1 molecules per spot for cells containing two spots per cell, consistent with 8 ± 3 Ssb tetramers per replication fork. Single-spot cells had a stoichiometry larger by a factor of ~2, at ~70 molecules per spot. The number of Ssb molecules bound at the replication fork is expected to be proportional to the length of single-stranded DNA (ssDNA). The average stoichiometry of Ssb within replisomes approximately doubled in cells treated with hydroxyurea (5), as expected, because ssDNA accumulates at the fork when replication is stalled (18). The ssDNA associated with each Ssb tetramer is either 35 or 65 nucleotides (nt) in vitro; (Ssb)35 forms compact filaments, whereas (Ssb)65 forms dimers of tetramers covering 170 nt (19). The contribution of each binding mode in vivo is unknown. Assuming that the stretch of ssDNA at the replication fork equals the Okazaki fragment length [~650 nt at 22°C (20)], then the presence of (Ssb)65 would give an occupancy of ~8 tetramers, close to our mean stoichiometry estimate for single replisome spots (Fig. 2).

Structural investigations of replisome components suggest that their cumulative volume is contained in a sphere of maximum diameter ~50 nm (2124). To investigate this, we studied the size and shape of images from each strain (Fig. 2, lower panels). Strains expressing fluorescent α, ε, τ, δ, χ, and ψ all produced spots with a circularly symmetrical shape and a mean full width at half maximum (FWHM) across the cell strains of 305 ± 30 nm (5). There was no significant difference from the individual FWHM measurements for each strain (P = 0.05, Student t test), but all were significantly larger than the FWHM of surface-immobilized YPet in vitro of ~250 nm (fig. S7). In contrast, β and Ssb produced spots spread further over the long axis of the cell, with Ssb foci extending up to ~200 nm; ~55% of the tetramers were present within a 50-nm diameter. Although the ~4 χψ heterodimers may interact with Ssb tails, we note that the spatial distribution of χψ is that of the core replisome–clamp loader rather than that of Ssb. Furthermore, the stoichiometry of χψ is much lower than that of Ssb. Taken together, these results suggest that χψ is associated preferentially with one or a few Ssb molecules in the vicinity of the core replisome.

The stoichiometry of the dimeric β sliding clamp was expected to be partly the result of its interaction with Pol III and the clamp loader, and partly due to association with DNA at the 3′ end of Okazaki fragments. Nonetheless, its bimodal distribution peaked at ~3 and ~6 dimers (Fig. 2), which raised the possibility that it might be continually associated with Pol III. Analysis of β-YPet spots indicated two subpopulations with different spatial intensity distributions (fig. S8). The first subgroup (~27% of spots) had circular distributions that support a scheme in which three β dimers are associated with active Pol III or with the clamp loader. In the second subgroup, the spots were more extended (circularity > 1.2), consistent with at least one of the sliding clamps being localized at a distance >50 nm from the replisome core.

Historically, the replisome has been considered to have two opposing yet coordinated polymerases connected to the rest of the replisome (25). Instead, we provide strong evidence for an in vivo core replisome containing three Pol IIIs associated with a clamp loader whose three copies of τ also trimerize Pol III. In a minority of replisomes, all three Pol IIIs may be associated with sliding clamps, with two being potentially simultaneously active on the lagging strand, supporting suggestions from in vitro studies of E. coli and phage T4 replisomes (2, 26). However, the majority of replisomes appear to have only two of the polymerases associated with a sliding clamp, which suggests that the third polymerase is waiting to be loaded onto the next lagging-strand primer (Fig. 3).

We measured the number of nonreplisome Ssb tetramers to be 330 ± 105 per cell (table S1), in agreement with estimates using quantitative Western blots (5). Similarly, 0.5 to 5% of other replisome molecules in a cell were associated with each replisome (table S1), hence the measured stoichiometries reflect biologically relevant complexes and are not the consequence of the replisome’s component being rate-limiting. The level of replisome molecules is sufficient to support more than 10 cellular replication forks associated with the chromosome, plasmids, or phage.

Slimfield microscopy provides a powerful noninvasive in vivo analytical tool that extends previous analyses of the assembly and action of molecular machines (10, 27, 28). In combination with degron-targeted proteolysis of specific proteins, it has provided unanticipated insight into replisome architecture. In combination with partial prebleaching and stochastic photoactivation and photoswitching techniques, the methodology may provide new insight into biological systems that contain substantially higher numbers of freely diffusing fluorescent proteins than were investigated here.

Supporting Online Material

Materials and Methods

Figs. S1 to S11

Table S1


References and Notes

  1. See supporting material on Science Online.
  2. Supported by the Wellcome Trust and the Royal Society. M.C.L. is a Royal Society University Fellow and research fellow of Hertford College Oxford. R.R.-L. is a research fellow of New College Oxford and was supported by Conacyt and a Clarendon postgraduate award. We thank C. Possoz and C. Lesterlin for stimulating discussions, and we acknowledge National BioResource Project (NIG, Japan): E. coli for the provision of the holC and holD deletion strains.
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