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Uncoupling of Leading- and Lagging-Strand DNA Replication During Lesion Bypass in Vivo

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Science  23 May 2003:
Vol. 300, Issue 5623, pp. 1300-1303
DOI: 10.1126/science.1083964

Abstract

Numerous agents attack DNA, forming lesions that impair normal replication. Specialized DNA polymerases transiently replace the replicative polymerase and copy past lesions, thus generating mutations, the major initiating cause of cancer. We monitored, in Escherichia coli, the kinetics of replication of both strands of DNA molecules containing a single replication block in either the leading or lagging strand. Despite a block in the leading strand, lagging-strand synthesis proceeded further, implying transient uncoupling of concurrent strand synthesis. Replication through the lesion requires specialized DNA polymerases and is achieved with similar kinetics and efficiencies in both strands.

The normal replication machinery accurately copies nondamaged template DNA but is unable to bypass most DNA lesions. For example, lesions caused by ultraviolet light interfere with DNA replication and lead to the formation of daughter-strand gaps in E. coli and in human cells (1, 2). However, until now all in vivo data have relied on randomly distributed lesions, preventing the fine structure of a blocked replication fork from being analyzed. To investigate the effect on fork progression of defined replication blocks in the leading versus lagging strand, we implemented a strategy in which the kinetics of synthesis of both DNA strands can be monitored separately in vivo. The assay involves electroporation-facilitated transformation of plasmid molecules and the analysis of their early replication products. The plasmids used here replicate unidirectionally from their ColE1 replication origin and use the same elongation apparatus as E. coli chromosome (3).

As a control experiment, we introduced lesion-free plasmid DNA into bacteria and analyzed its replication products as a function of time. The synthesis of both strands can be monitored separately by the presence of a local sequence heterology, formed by a +3-nucleotide (nt) bulge, which introduces a new restriction site in one of the strands after replication (fig. S1) (4). At time points as early as 15 min, bands specific for leading- and lagging-strand replication products were observed (Fig. 1A). In agreement with models involving coupled replication of both DNA strands, leading and lagging replication products appear with similar kinetics (Fig. 1B).

Fig. 1.

Kinetics of replication of nondamaged plasmid. (A) Replication products are analyzed by electrophoresis (8% denaturing polyacrylamide gel electrophoresis) at various time points after replication in the SOS-induced strain JM103, uvrA,mutS. Mismatch repair is inactivated in this strain to prevent repair of the sequence heterology. Fractions of the total sample (one-half or one-quarter) have been extracted and loaded as indicated to prevent saturation at longer time points. An equal amount of external standard plasmid (72-nt band) is added to all samples before extraction. Leading- (87 nt) and lagging- (63 nt) strand replication products appear with kinetics similar to those expected for concurrent strand replication. Quantification of the bands is made possible by applying known amounts of control plasmid (calibration tracks). Sequence reactions (A and C) of the marker strand are also applied on the gel. Of the plasmids that became irreversibly bound to cells (i.e., resistant to washes), only a small fraction (5 to 10%) engages in replication cycles. The remainder, which is not being replicated, is revealed as a 90-nt band, the amount of which stays essentially constant over time when corrected for the amount of sample loaded in each track. (B) The amounts of leading-and lagging-strand replication products are plotted as a function of time (average of two independent experiments). The three first time points are interpolated to derive the average doubling time. Error bars are standard deviations from two to three independent determinations.

To analyze the effect of lesions on the overall replication kinetics of double-stranded DNA, we constructed plasmid molecules containing a single lesion in either the leading- or lagging-strand templates. The DNA lesion is formed by covalent linkage of the chemical carcinogen N-2-acetylaminofluorene (AAF) to the C-8 position of a guanine residue (G-AAF) (5, 6). A plasmid carrying a single G-AAF lesion in the leading-strand DNA template is introduced into SOS-induced bacteria to express the specialized DNA polymerases required for translesion synthesis (TLS) (7). Unexpectedly, the G-AAF lesion in the leading strand did not affect the kinetics of appearance of the lagging-strand replication products, which accumulate as in lesion-free control plasmid (Figs. 1 and 2). In contrast, the synthesis of replication products from the lesion-containing leading strand is strongly delayed (Fig. 2). Replication intermediates blocked in the vicinity of the lesion site (L-1 and L-2, Fig. 2A) are observed at early time points (15 min) and subsequently converted into full-length bypass products with a delay of ∼50 min (Fig. 2B). This delay reflects the average time required for TLS at the single AAF lesion in vivo. After this initial delay, leading-strand products accumulate with a doubling time similar to that observed for non-damaged DNA (i.e., ∼11 min). Similarly, when the G-AAF lesion is located in the lagging strand, plasmids derived from the leading strand accumulate with the kinetics observed for nondamaged molecules, whereas the appearance of progeny from the lagging-strand template is delayed by ∼50 min (8). In the absence of SOS induction, no TLS products are observed, which shows complete dependence of lesion bypass on SOS-controlled gene products (8).

Fig. 2.

Kinetics of replication of a plasmid with a single G-AAF adduct in the leading-strand template in SOS-induced strain JM103, uvrA,mutS. Both excision and mismatch repair pathways are inactivated in this strain to prevent repair of the lesion-containing heteroduplex. (A) Analysis is performed as detailed in Fig. 1. Replication blocks one or two nucleotides before the lesion (L-1, L-2) appear at time points as early as 15 min (lower inset with high contrast). Error-free (TLS0) and frame-shift (TLS-2) bypass products are visible at 30 min and accumulate at later time points (upper inset with high contrast). The nondamaged lagging strand accumulates (63-nt band) with the kinetics observed for nondamaged DNA (Fig. 1), showing uncoupling of the replication of the two strands. (B) The amounts of leading- (bypass products) and lagging-strand replication products are plotted as a function of time (average of two independent experiments). The lag time between the two curves represents the average time required for TLS of a single G-AAF adduct in vivo (∼50 min). Error bars are standard deviations from two to three independent determinations.

Two distinct TLS products are observed (Fig. 2A), in agreement with previous genetic studies, showing that within the specific sequence context used here (5′-GGCGAAFCC-) the G-AAF lesion can be bypassed using two distinct pathways (7, 9). An error-free pathway involves DNA polymerase Pol V (umuDC), whereas Pol II (polB) mediates a mutagenic, –2 frameshift pathway (87-and 85-nt bands, respectively) (Fig. 3A). We investigated the genetic requirement of the two TLS pathways directly using umuDC or polB deletion strains. In agreement with previous genetic data (7), no error-free bypass is observed in the ΔumuDC strain, whereas no frameshift bypass product is seen in the ΔpolB strain (Fig. 3B).

Fig. 3.

Bypass of a single G-AAF adduct: distinct error-free and frameshift pathways (7, 22). (A) Two pathways mediate TLS of a single G-AAF adduct located within the NarI sequence: an error-free pathway and a –2 frameshift pathway that require DNA Pol V and Pol II, respectively. (B) Replication of a plasmid with a G-AAF lagging-strand template lesion in SOS-induced, wild-type polB (Pol II) or umuDC (Pol V) deletion strains. Replication products are analyzed at a fixed 60-min time point. All three strains exhibit similar replication efficiencies of both strands of the nondamaged plasmid. With lesion-containing plasmid, both error-free (TLS0) and frameshift (TLS-2) bypass products are observed in the wild-type strain, whereas no TLS-2 and noTLS0 products are seen in the ΔpolB and ΔumuDC strains, respectively.

The present data show uncoupling of leading- and lagging-strand synthesis, implying the continuation of fork opening despite a block in one strand. Uncoupling of simultaneous strand synthesis may occur without disruption of the dimeric Pol III core assembly. When the lesion resides in the lagging strand, a new priming event may enable lagging-strand synthesis to continue, generating a gapped plasmid and a complete double-stranded plasmid molecule from lagging and leading strands, respectively. When the block resides in the leading strand, the lagging-strand Pol III core gets ahead of the leading-strand core, generating a complete double-stranded plasmid from the lagging strand. A partially double-stranded molecule with a single-stranded region (∼1 kb) extending from the lesion site to the end of the plasmid is formed from the leading strand. Plasmids of larger size will be required to determine how far DnaB helicase can travel before the whole fork stops. In both orientations, TLS can repair the partially replicated molecule with similar efficiency and a 50-min delay. This delay strongly depends on the chemical nature of the blocking lesion (8). Alternatively, the partially replicated intermediates may be processed by regressed fork formation (1013). In E. coli and in yeast, genetic data have indicated the bypass of specific lesions to require multiple polymerase switches and specific combinations of TLS polymerases (7, 1421). The strategy implemented here will be useful to unravel the complex biochemistry of various TLS pathways in vivo, thus providing a powerful complement to in vitro approaches.

Supporting Online Material

www.sciencemag.org/cgi/content/full/300/5623/1300/DC1

Materials and Methods

Fig. S1

References

References and Notes

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