While Pol α clearly initiates DNA synthesis at origins and of Okazaki fragments, the roles of Pols δ and ε in leading and lagging strand replication in vivo have been difficult to discern, in part because mutations in the yeast genes encoding these polymerases (POL3 for Pol δ and POL2 for Pol ε) are lethal or nearly so [9,10]. The situation changed when structure-function studies identified novel variants of yeast Pol α [11], Pol δ [12–14] and Pol ε [15] that were used for tracking where each polymerase synthesizes DNA in a cell. Each variant contains a replacement for a conserved leucine (α/δ) or methionine (ε) at the polymerase active site. These variants have high replication capacity allowing normal cell growth. They also have low fidelity that elevates spontaneous mutation rates in vivo, thus identifying the polymerase responsible for making a replication error. Most importantly, they have error signatures that distinguish whether the error was made during copying the leading or lagging strand template. For example, between two mismatches that could give rise to T-A to C-G transition substitutions, L612M Pol δ generates T-dGTP mismatches at a ≥ 28-fold higher rate than A-dCTP mismatches (Fig. 1A, top) during DNA synthesis in vitro [14]. In a MMR-defective pol3-L612M msh2Δ strain that monitors uncorrected replication errors made by L612M Pol δ in vivo, T to C substitutions at base pair 97 in the URA3 gene were created at a high rate (58 × 10⁻⁷) when URA3 was located close to a well characterized replication origin (ARS306), and distant from the next closest origin [16]. Given the direction of fork movement as base pair 97 is being replicated, and inferring that the T to C substitution results from a T-dGTP mismatch, the error would be generated by Pol δ during lagging strand replication (Fig. 1A, OR1). Importantly, T to C substitutions at base pair 97 occurred at a much lower rate (3.1 × 10⁻⁷) when the URA3 orientation (OR2) was reversed relative to ARS306. In this orientation, the T-dGTP mismatch would be a leading strand error, so the much lower rate implies that Pol δ has at most a minor role in replicating the leading strand template. When a similar logic was applied to other orientation-dependent mutations across the URA3 open reading frame (e.g., deletion of a T-A base pair (Fig. 1A) and see other events in [16]), the results supported the idea that Pol δ primarily copies the lagging strand template.

rNTPs in DNA present a potential risk to genome stability because the reactive 2´ hydroxyl would sensitize the DNA backbone to strand cleavage. This is interesting because rNTP concentrations in eukaryotes are much higher than dNTP concentrations [23,28]. Although most DNA polymerases discriminate well against rNTP insertion into DNA [29], a recent study showed that in reactions containing cellular rNTP and dNTP concentrations, yeast Pols α, δ and ε incorporate substantial numbers of ribonucleotides into DNA during synthesis in vitro [28]. Soon thereafter [30], rNTPs were shown to be incorporated during replication by yeast Pol ε in vivo, and also that they are normally removed by RNase H2-dependent repair. However, failure to repair these ribonucleotides in RNase H2-defective strains slightly increased replicative stress and strongly increased the rate of 2–5 base pair deletions in repetitive sequences. In some cases (e.g., deletion of a CA dinucleotide, Fig. 2B), the deletion rate was higher for one orientation of URA3 as compared to the other. These data suggest strand-specific rNTP incorporation by Pol ε, once again underscoring the asymmetry of replication. Interestingly, the rate of 2–5 base pair deletions in the pol2-M644G rnh201 strain was not strongly increased by loss of mismatch repair [31]. This suggests a model wherein, when rNMPs incorporated by M644G Pol ε during leading strand replication are not repaired by RNase H2, they are processed outside the context of replication in a manner that gives rise to misaligned intermediates that result in short deletions. In support of this hypothesis, a recent study [32] showed that topoisomerase 1 is required for formation of 2–5 base pair deletions in rnh201Δ strains. Topoisomerase 1 normally relieves DNA supercoils formed during transcription, and it can also cleave the backbone of DNA containing a single ribonucleotide. This cleavage creates DNA ends that must be processed to allow ligation, and it is this processing that is proposed to provide the opportunity for strand misalignments in repetitive sequences that ultimately yield the deletions.

Early genetic studies in E. coli (e.g., see [35] and references therein) revealed that mismatch repair most efficiently corrects mismatches that are most commonly made by its major replicase, DNA polymerase III. This type of ‘reciprocity’ is also seen for replication errors generated by yeast Pol δ. Pol δ most frequently creates mismatches that could result in transitions and single base deletions in mononucleotide runs, and it is these errors that are corrected most efficiently by MMR [16]. Does reciprocity extend even further? Might MMR correct lagging strand replication errors made by less accurate Pol α more efficiently than those made by more accurate Pol δ? One reason to consider this possibility is that errors made by Pol α are closer to the 5´ ends of Okazaki fragments than are errors made by Pol δ. This proximity may be relevant because the 5´ ends of DNA can serve as signals for MMR in vitro [36], and because when working in conjunction with the asymmetric PCNA sliding clamp [37], DNA 5´ ends have been suggested to be strand discrimination signals in vivo [38]. When this type of reciprocity was tested in yeast using the Leu to Met mutator alleles of Pol δ and Pol α, the results (Fig. 3A and [39]) suggest that apparent MMR efficiencies may indeed be higher for base-base mismatches made by Pol α as compared to the same mismatches made by Pol δ. This supports a model (Fig. 3B) wherein the 5′ ends of Okazaki fragments serve as strand discrimination signals. The proximity of a Pol α error to a 5´ DNA end may result in higher efficiency for the general MMR machinery, just as the efficiency of E. coli MMR is proportional to the distance between the mismatch and the strand discrimination signal [36]. It is also possible that mismatches near the 5′ ends of Okazaki fragments are repaired by a MMR pathway specialized to protect the genome against particularly abundant replication errors made by Pol α (Fig. 3B). This could involve mismatch removal via strand displacement, as recently observed in vitro [40], or MMR involving nucleases such as Exo1 or the exonuclease activity of yeast FEN1, a nuclease involved in Okazaki fragment maturation that has also been suggested to participate in Msh2-Msh6-dependent MMR [41].

The studies reviewed here illustrate a growing appreciation of the intricate enzymology required to asymmetrically replicate large and complex eukaryotic nuclear genomes with the accuracy needed to preserve the information content of both strands. The ultimate outcome reflects not just the polymerases discussed here, but also their highly differentiated accessory proteins [5] and the coordination of replication with repair, recombination, transcription [47], nucleosome loading, and chromatin status. This complexity makes it improbable that genome stability can be generally increased [48]. This complexity also provides many opportunities for destabilizing the genome, some undoubtedly yet to be discovered. Achieving a detailed understanding of these opportunities is motivated by the beneficial consequences of mutations to organisms, like evolution, phase variation at contingency loci in microbes and somatic hypermutation of immunoglobulin genes, as well as from the many associations between mutations and human diseases [49,50]. As just one example that may be related to asymmetric relication of the leading and lagging strands, defects in the 3´ exonuclease activities of Pols δ and ε both confer cancer susceptibility in mice, but do so at different ages of onset and with different tissue specificity [51]. Could this be related to differences in strand-specific mutagenesis and/or tissue-specific differences in replication origin firing during development?