DNA polymerases δ and λ cooperate in repairing double-strand breaks by microhomology-mediated end-joining in Saccharomyces cerevisiae

Maintenance of genome stability is carried out by a suite of DNA repair pathways that ensure the repair of damaged DNA and faithful replication of the genome. Of particular importance are the repair pathways, which respond to DNA double-strand breaks (DSBs), and how the efficiency of repair is influenced by sequence homology. In this study, we developed a genetic assay in diploid Saccharomyces cerevisiae cells to analyze DSBs requiring microhomologies for repair, known as microhomology-mediated end-joining (MMEJ). MMEJ repair efficiency increased concomitant with microhomology length and decreased upon introduction of mismatches. The central proteins in homologous recombination (HR), Rad52 and Rad51, suppressed MMEJ in this system, suggesting a competition between HR and MMEJ for the repair of a DSB. Importantly, we found that DNA polymerase delta (Pol δ) is critical for MMEJ, independent of microhomology length and base-pairing continuity. MMEJ recombinants showed evidence that Pol δ proofreading function is active during MMEJ-mediated DSB repair. Furthermore, mutations in Pol δ and DNA polymerase 4 (Pol λ), the DNA polymerase previously implicated in MMEJ, cause a synergistic decrease in MMEJ repair. Pol λ showed faster kinetics associating with MMEJ substrates following DSB induction than Pol δ. The association of Pol δ depended on RAD1, which encodes the flap endonuclease needed to cleave MMEJ intermediates before DNA synthesis. Moreover, Pol δ recruitment was diminished in cells lacking Pol λ. These data suggest cooperative involvement of both polymerases in MMEJ.DNA double-strand breaks (DSBs) are toxic lesions that can be repaired by two major pathways in eukaryotes: nonhomologous end-joining (NHEJ) and homologous recombination (HR) (1). Although HR repairs DSBs in a template-dependent, high-fidelity manner, NHEJ functions to ligate DSB ends together using no or very short (1–4 bp) homology. Recently, a new pathway was identified in eukaryotes, which uses microhomologies (MHs) to repair a DSB and does not require the central proteins used in HR (Rad51, Rad52) or NHEJ (Ku70–Ku80) (2–5). In mammalian cells, this pathway of repair is known as alternative end-joining (Alt-EJ) and is often but not always associated with MHs, whereas in budding yeast, the commensurate pathway, MH-mediated end-joining (MMEJ), will typically use 5–25 bp of MH (6, 7). These pathways are associated with genomic rearrangements, and cancer genomes show evidence of MH-mediated rearrangements (8–12). In addition, eukaryotic genomes contain many dispersed repetitive elements that can lead to genome rearrangements when recombination occurs between them (13–16). Therefore, controlling DSB repair in the human genome, which features a variety of repeats, is especially important given the fact that recombination between repetitive elements has been implicated in genomic instability associated with disease (17–20).The original characterization of Alt-EJ in mammalian cells suggested it did not represent a significant DNA repair pathway and only operated in the absence of functional HR and NHEJ pathways. More recent analyses demonstrate a physiological role of Alt-EJ during DNA repair in the presence of active HR and NHEJ pathways (2, 12, 21, 22). Furthermore, examination of I-SceI–induced translocation junctions in mammalian cells revealed the frequent presence of MHs (23, 24). NHEJ-deficient and p53-null mice develop pro–B-cell lymphomas, and nonreciprocal translocations characterized by small MHs are found at their break point junctions (25–28). Similarly, in human cancers, many translocation break point junctions contain MHs, suggesting a role for Alt-EJ in cancer development (29–31) and resistance to chemotherapy and genetic disease (32–36). Hence, the presence of many short repetitive sequences in the human genome is likely to increase rearrangements mediated by MHs following the creation of a DSB.MMEJ is a distinct DSB repair pathway that operates in the presence of functional NHEJ and HR pathways (10, 37). The genetic requirements of MMEJ are being studied in the model eukaryote Saccharomyces cerevisiae and involve components traditionally considered specific to the NHEJ (Pol λ) and HR (Rad1–Rad10, Rad59, and Mre11–Rad50–Xrs2) pathways (4, 5, 10, 38). Although being clearly independent of the central NHEJ factor Ku70–Ku80 heterodimer (10, 37), the involvement of the key HR factor Rad52 in MMEJ remains uncertain. It has been reported that Rad52 is required for MMEJ repair (4, 10, 38), whereas in another assay system Rad52 suppresses MMEJ repair (37). More recently, it has been proposed that the replication protein A (RPA) regulates pathway choice between HR and MMEJ (37). In addition, several models have been proposed that identify specific pathways that may use MHs for the repair of DNA damage (39–41). Despite current advancements in our understanding of MMEJ, the precise involvement of DNA polymerases in supporting the repair of DSBs using MHs remains poorly understood. DNA polymerase λ (also called Pol4 in budding yeast) and its human homolog Pol λ are considered to be the primary candidates for the DNA polymerases working in NHEJ and MMEJ (4, 5, 42–46). Both genetic and biochemical evidence shows that Pol δ is recruited during HR to extend Rad51-dependent recombination intermediates (47–50). Recent analysis using pol32 mutants (5, 10) implicated the Pol32 subunit of Pol δ in MMEJ. Pol32 and Pol31 were also identified as subunits of the DNA polymerase zeta complex (Pol ζ) (51, 52), but previous analysis showed no effect of rev3 mutants in MMEJ (10). REV3 encodes the catalytic subunit of Pol ζ. However, an involvement of Pol δ had not been demonstrated directly before, and it is possible that Pol32 could act in conjunction with yet another DNA polymerase.Here, we report the development of a series of interchromosomal MMEJ assays in diploid S. cerevisiae to assess the mechanisms underlying the repair of DSBs using varying MHs. We focus on diploid cells, as they represent the natural state of budding yeast, which is a diplontic organism (53). The yeast mating-type switching system represents a mechanism to return haploid yeast as efficiently as possible to diploidy (54). Using a combination of genetic, molecular, and in vivo chromatin immunoprecipitation (ChIP) experiments, we provide compelling evidence for a direct involvement of Pol δ in coordinating with Pol λ in MMEJ in budding yeast.     

Structure–function analysis of ribonucleotide bypass by B family DNA replicases

Ribonucleotides are frequently incorporated into DNA during replication, they are normally removed, and failure to remove them results in replication stress. This stress correlates with DNA polymerase (Pol) stalling during bypass of ribonucleotides in DNA templates. Here we demonstrate that stalling by yeast replicative Pols δ and ε increases as the number of consecutive template ribonucleotides increases from one to four. The homologous bacteriophage RB69 Pol also stalls during ribonucleotide bypass, with a pattern most similar to that of Pol ε. Crystal structures of an exonuclease-deficient variant of RB69 Pol corresponding to multiple steps in single ribonucleotide bypass reveal that increased stalling is associated with displacement of Tyr391 and an unpreferred C2´-endo conformation for the ribose. Even less efficient bypass of two consecutive ribonucleotides in DNA correlates with similar movements of Tyr391 and displacement of one of the ribonucleotides along with the primer-strand DNA backbone. These structure–function studies have implications for cellular signaling by ribonucleotides, and they may be relevant to replication stress in cells defective in ribonucleotide excision repair, including humans suffering from autoimmune disease associated with RNase H2 defects.Replication of the eukaryotic nuclear genome initiates when RNA primase synthesizes RNA primers of about 10 nucleotides (1). Because this occurs at multiple replication origins and at ∼200-bp intervals on the lagging strand template, about 5% of the genome is initially synthesized as chains of consecutive ribonucleotides. These ribonucleotides are subsequently removed during Okazaki fragment maturation by the combined action of ribonucleases (RNases) H (2) and flap endonucleases (3). Ribonucleotides are also incorporated into DNA by DNA polymerases (Pol) α, δ, and ε, because they discriminate against ribonucleoside triphosphates (rNTPs) efficiently but imperfectly (4) and because cellular rNTP concentrations are much higher than dNTP concentrations (4). As a consequence, large numbers of ribonucleotides are incorporated during replication, and are present in the genomes of cells defective in the repair enzymes that initiate their removal, RNase H2 (5-9) and topoisomerase 1 (10).Ribonucleotides in DNA are a dual-edged sword, in that they have both beneficial and deleterious consequences. On the beneficial side, two consecutive ribonucleotides in the genome are signals for mating type switching in Schizosaccharomyces pombe (11). In addition, recent evidence suggests that RNase H2-dependent processing of ribonucleotides incorporated into the Saccharomyces cerevisiae genome by Pol ε, the primary leading strand replicase, generates a signal that can direct mismatch repair (MMR) to correct replication errors in the nascent leading strand (9, 12). Other possible beneficial signaling roles for ribonucleotides have also been considered (4, 13).On the deleterious edge of the sword, the 2′-oxygen on a ribose sugar in DNA can attack the backbone and render DNA chemically unstable. Yeast strains defective in RNase H2-dependent ribonucleotide excision repair (RER) (5, 14) exhibit several characteristics of replicative stress, including strongly elevated rates for deleting 2–5 bp from repetitive DNA sequences (5, 15), events that are initiated by topoisomerase 1 cleavage of a ribonucleotide in DNA (10, 16). Yeast strains defective in RNase H2 and RNase H1 progress slowly through S phase, accumulate ubiquitylated proliferating cell nuclear antigen (PCNA), and are sensitive to treatment with hydroxyurea (17). Moreover, their survival in the presence of hydroxyurea partly depends on MMS2-dependent template switching and on REV3, which encodes the catalytic subunit of the translesion synthesis (TLS) enzyme Pol ζ. When ribonucleotide incorporation during leading strand replication is increased by a M644G substitution in the Pol ε active site, a defect in RNase H2 results in elevated deletion mutagenesis, elevated dNTP pools, slow growth and activation of the S-phase checkpoint (5, 10, 18), and concomitant deletion of the RNH1 gene encoding RNases H1 is lethal (17). In mice, knocking out any of the genes encoding the three subunits of RNase H2 is embryonic lethal (7, 19). RNase H2 null embryos grow slowly due to reduced cell proliferation and exhibit genome instability and a p53-dependent DNA damage response. Fibroblasts from these embryos contain more than a million single and/or di-ribonucleotides in their genomes and elevated numbers of strand breaks, γ-H2A histone family, member X foci, micronuclei, and chromosomal aberrations. In humans, mutations in the genes encoding RNase H2 are associated with Aicardi-Goutières syndrome, a rare neuroinflammatory condition resembling congenital viral infection (20).These phenotypes of RNase-deficient cells are characteristic of stress that could arise from difficulty in replicating DNA templates containing unrepaired ribonucleotides. This idea is consistent with knowledge that replicases require normal DNA helix geometry to achieve efficient and accurate DNA synthesis, and with crystallographic and NMR studies (21–24) showing that ribonucleotides in DNA alter helix parameters. Recent studies have shown that Pols δ and ε have difficulty bypassing ribonucleotides in DNA templates (4), whereas Pol ζ does not (17). The probability that Pol ε will pause during single ribonucleotide bypass increases after dNTP insertion opposite the ribonucleotide and for several additional insertions opposite deoxynucleotides (5, 25).In this study, we quantify stalling by yeast replicative Pols δ and ε as the number of consecutive ribonucleotides in the DNA template increases from one to four. We show that stalling increases as the number of consecutive ribonucleotides in the DNA template increases, with Pol δ being more efficient at ribonucleotide bypass than Pol ε. We then examine the structural basis for difficulty in ribonucleotide bypass using a homologous B family replicase, bacteriophage RB69 DNA polymerase, as a surrogate that is highly amenable to structural studies (26, 27). To promote the structural analysis, we used a variant of RB69 Pol containing a phenylalanine substituted for Leu415 (28). Leu415 is adjacent to invariant Tyr416, which interacts with the sugar of the incoming dNTP and has an important role in preventing rNTP incorporation (29). An initial crystal structure of L415F RB69 Pol with correctly base-paired dTTP opposite template dA (28) revealed that the phenylalanine ring is accommodated within a cavity present in the WT polymerase without steric clash or major change in active site geometry, consistent with retention of high catalytic efficiency for correct incorporation. Moreover, L415F RB69 Pol can also bypass 8-oxo-guanine more efficiently than can WT RB69 Pol (28). These bypass results were encouraging because the yeast replicases bypass single ribonucleotides with efficiencies somewhat similar to those for bypass of 8-oxo-guanine, and L415F RB69 Pol may therefore facilitate crystallization of ternary complexes with ribonucleotides in the DNA template (25). We show that WT and L415F RB69 Pol also stall during ribonucleotide bypass, to a degree most closely resembling stalling by Pol ε. We describe seven unique crystal structures relevant to L415F RB69 Pol bypass of one or two ribonucleotides. The data are discussed in relation to the consequences of ribonucleotides in the genomes of cells defective in their removal.     

Polymerase exchange on single DNA molecules reveals processivity clamp control of translesion synthesis

Translesion synthesis (TLS) by Y-family DNA polymerases alleviates replication stalling at DNA damage. Ring-shaped processivity clamps play a critical but ill-defined role in mediating exchange between Y-family and replicative polymerases during TLS. By reconstituting TLS at the single-molecule level, we show that the Escherichia coli β clamp can simultaneously bind the replicative polymerase (Pol) III and the conserved Y-family Pol IV, enabling exchange of the two polymerases and rapid bypass of a Pol IV cognate lesion. Furthermore, we find that a secondary contact between Pol IV and β limits Pol IV synthesis under normal conditions but facilitates Pol III displacement from the primer terminus following Pol IV induction during the SOS DNA damage response. These results support a role for secondary polymerase clamp interactions in regulating exchange and establishing a polymerase hierarchy.Despite the action of several DNA repair pathways, unrepaired damage is encountered by replicative DNA polymerases, which stall at DNA-distorting lesions. Translesion synthesis (TLS), most notably by Y-family polymerases, is one pathway that alleviates such roadblocks. In TLS, a Y-family polymerase switches with a stalled replicative polymerase, synthesizes across from and past the lesion, and then, switches back to allow resumption of normal synthesis (1). The ability of Y-family polymerases to bypass damaged DNA comes at the cost of lower fidelity, requiring careful regulation of polymerase exchange (2).Processive synthesis by DNA polymerases requires their tethering to the protein-binding cleft of a ring-shaped processivity clamp by a conserved clamp-binding motif (CBM). Canonical clamps, such as the bacterial β and eukaryotic proliferating cell nuclear antigen (PCNA), are multimeric, with a binding cleft on each protomer. Biochemical experiments with bacterial (3, 4) and eukaryotic (5) proteins have suggested that clamps can simultaneously bind multiple DNA polymerases during active DNA synthesis, serving as a molecular toolbelt (6). This multivalency may facilitate rapid polymerase exchange and lesion bypass. However, it remains unclear if and when large multisubunit replicative polymerases can accommodate Y-family polymerases on the clamp. Furthermore, most organisms have multiple Y-family polymerases and many other clamp-binding proteins—at least 10 in Escherichia coli (7) and over 50 in humans (8). It is consequently an open question how the correct polymerase is selected at a DNA lesion (1).To further elucidate the role of processivity clamps in polymerase trafficking, we studied the E. coli replicative polymerase, the polymerase (Pol) III heterotrimer αεθ, and the Y-family Pol IV, which individually bind the dimeric β clamp. Pol IV, encoded by the gene dinB, is widely conserved across the three domains of life, and it is the homolog of human Pol κ (9). In addition to its function in lesion bypass (10), E. coli Pol IV is required for the mechanistically controversial phenomenon of stress-induced mutagenesis (11, 12), which is proposed to occur by its preferential synthesis at double-strand break intermediates (13, 14), and involved in reactive oxygen species-mediated antibiotic lethality by its incorporation of oxidized nucleotides into the genome (15).Interactions of Pol IV with β and their implications for the toolbelt model have generated widespread interest (4, 16). The structure of the C-terminal little finger domain of Pol IV bound to β revealed that Pol IV can simultaneously interact with the cleft and the rim, a secondary site of β near its dimer interface, which positions the Pol IV catalytic domain well away from the DNA running through the center of the clamp (17). Although this potential inactive binding mode for Pol IV has been interpreted as evidence for the toolbelt model, Pol III was shown to contain a second weak CBM in its ε exonuclease subunit (18) in addition to the CBM in its α catalytic subunit that has a strong affinity for the β clamp. A recent study proposed that Pol III would, therefore, occlude Pol IV from clamp binding during replication, only accommodating simultaneous binding after a lesion-induced stall (19).Previous efforts to reconstitute this model system for polymerase exchange have involved stalling Pol III on β at a primer terminus by nucleotide omission to synchronize a population of molecules and simulate a lesion-induced block (3, 4). These studies were not able to resolve an exchange back to Pol III after Pol IV synthesis. To bypass these limitations and elucidate the molecular mechanism of exchange between Pol III and Pol IV, we developed a single-molecule assay to observe the whole TLS reaction, quantifying polymerase exchange and bypass at site-specific DNA lesions. Here, we show that Pol III and Pol IV can simultaneously bind β during active synthesis, enabling rapid lesion bypass, and report a previously unidentified inactive binding mode for Pol III. We also observe that, at high concentrations (corresponding to up-regulated levels during the SOS DNA damage response), Pol IV occupies a secondary contact on β, promoting dissociation of Pol III. These results support a model in which secondary contacts between processivity clamps and Y-family polymerases establish a hierarchy for polymerase selection.     

CMG helicase and DNA polymerase ε form a functional 15-subunit holoenzyme for eukaryotic leading-strand DNA replication

DNA replication in eukaryotes is asymmetric, with separate DNA polymerases (Pol) dedicated to bulk synthesis of the leading and lagging strands. Pol α/primase initiates primers on both strands that are extended by Pol ε on the leading strand and by Pol δ on the lagging strand. The CMG (Cdc45-MCM-GINS) helicase surrounds the leading strand and is proposed to recruit Pol ε for leading-strand synthesis, but to date a direct interaction between CMG and Pol ε has not been demonstrated. While purifying CMG helicase overexpressed in yeast, we detected a functional complex between CMG and native Pol ε. Using pure CMG and Pol ε, we reconstituted a stable 15-subunit CMG–Pol ε complex and showed that it is a functional polymerase–helicase on a model replication fork in vitro. On its own, the Pol2 catalytic subunit of Pol ε is inefficient in CMG-dependent replication, but addition of the Dpb2 protein subunit of Pol ε, known to bind the Psf1 protein subunit of CMG, allows stable synthesis with CMG. Dpb2 does not affect Pol δ function with CMG, and thus we propose that the connection between Dpb2 and CMG helps to stabilize Pol ε on the leading strand as part of a 15-subunit leading-strand holoenzyme we refer to as CMGE. Direct binding between Pol ε and CMG provides an explanation for specific targeting of Pol ε to the leading strand and provides clear mechanistic evidence for how strand asymmetry is maintained in eukaryotes.Replisomes are multisubunit protein complexes that coordinately unwind duplex DNA and duplicate both parental strands during chromosomal replication. Detailed studies of cellular and viral systems show that the basic functional units of replication—helicase, primase, and DNA polymerase (Pol)—are common to all replisomes whereas the evolutionary histories of the individual components in different kingdoms are distinctive and diverse (1). Accordingly, the sequence and structure of replisome components are unrelated, and thus connections and coordination among the different functional units can be expected to vary widely.The most well-studied cellular replisome to date, bacterial Escherichia coli, uses multiple copies of a single DNA polymerase to replicate both parental strands, and the action of these polymerases is coordinated by a multifunctional clamp loader that also connects to the replicative helicase (2). For reasons that are still unclear, the eukaryotic replisome uses three different polymerases for normal chromosome duplication, including one for the leading strand (Pol ε) and two for the lagging strand (Pol α/primase and Pol δ) (3–5). Similarly, whereas the replicative helicase in E. coli is a homohexamer of DnaB, the eukaryotic CMG (Cdc45-MCM-GINS) helicase consists of 11 distinct subunits assembled on chromatin by loading of the heterohexameric Mcm2-7 helicase core at an origin and its subsequent activation by association with Cdc45 and the heterotetrameric GINS (Sld5-Psf1-Psf2-Psf3) complex at the onset of S-phase to form the CMG complex (6–8). Among other things, the complexity of the eukaryotic system reflects the need to restrict chromosome duplication to a single round in a normal cell cycle so that proper ploidy can be maintained across multiple chromosomes after cell division.Detailed biochemical studies of the E. coli replisome show that the leading and lagging strand replicases are coupled and intimately linked to the replicative helicase, a feature also common to the well-characterized T4 and T7 bacteriophage replication systems (9–11). For this reason, it has been assumed that the same would be true of eukaryotic systems, and this notion has been strongly reinforced by the identification in yeast of replication progression complexes (RPCs), large multiprotein complexes containing, among other proteins, CMG, Mcm10, Mrc1, and Ctf4 (12, 13). The RPC also contains Pol α/primase under low-salt conditions, suggesting that it is more weakly bound, and binding of Pol α to the replisome is abolished in cells lacking Ctf4 or its metazoan counterpart, AND-1 (13–16). Ctf4 binds both the catalytic Pol1 subunit of Pol α and GINS in yeast and thus is thought to tether Pol α to CMG in the replisome (13, 16, 17).Neither Pol δ nor Pol ε is found in the most highly purified RPCs, which are defined by mass spectrometry of proteins bound after sequential affinity purification of two separate CMG components from a cell extract (12, 13). However, the noncatalytic Dpb2 protein subunit of Pol ε is known to bind to the GINS component of CMG, and recent evidence suggests that this interaction helps maintain Pol ε at the replication fork (18, 19). Pol δ was shown to bind Pol α via its nonessential Pol32 subunit (20), suggesting that Pol δ might be recruited from solution to extend primers initiated by Pol α/primase and may only associate transiently with the core replisome.To study the eukaryotic replisome in detail, we initiated a long-term project to purify the numerous components of the RPC/replisome from the model eukaryote Saccharomyces cerevisiae. Pioneering work on Drosophila and human CMG showed that an active helicase complex could be obtained by coexpression of all 11 subunits in insect cells (7, 21) so we cooverexpressed all 11 CMG subunits in yeast and purified the complex to homogeneity (22). We showed that, like its human counterpart, yeast CMG is capable of catalyzing replication of a model replication-fork substrate (21, 22). Using this system, we also showed that CMG enforces a preference for Pol ε over Pol δ in leading-strand replication whereas proliferating cell nuclear antigen (PCNA) enforces the opposite preference on the lagging strand (22). Preferential binding of Pol δ to PCNA has been clearly demonstrated and provides an explanation for the dominance of Pol δ in lagging-strand synthesis (23), but the nature of any interaction between Pol ε and CMG on the leading strand is poorly understood.While purifying CMG from yeast, we identified a direct interaction between overexpressed CMG and native Pol ε to form a multifunctional eukaryotic leading-strand holoenzyme that we refer to as CMGE. Using separately purified CMG and Pol ε, we reconstituted a stable, 15-subunit CMGE and showed that it is an active helicase–polymerase in vitro. We also show that the Dpb2 subunit of Pol ε, which binds to the Psf1 protein subunit of GINS, promotes efficient Pol ε function with CMG. Direct binding of Pol ε to the full CMG complex has not been previously demonstrated, and this interaction provides a mechanistic foundation for preferential replication of the leading strand by Pol ε as part of a stable helicase–polymerase holoenzyme (4).     

Programmable motion of DNA origami mechanisms

DNA origami enables the precise fabrication of nanoscale geometries. We demonstrate an approach to engineer complex and reversible motion of nanoscale DNA origami machine elements. We first design, fabricate, and characterize the mechanical behavior of flexible DNA origami rotational and linear joints that integrate stiff double-stranded DNA components and flexible single-stranded DNA components to constrain motion along a single degree of freedom and demonstrate the ability to tune the flexibility and range of motion. Multiple joints with simple 1D motion were then integrated into higher order mechanisms. One mechanism is a crank–slider that couples rotational and linear motion, and the other is a Bennett linkage that moves between a compacted bundle and an expanded frame configuration with a constrained 3D motion path. Finally, we demonstrate distributed actuation of the linkage using DNA input strands to achieve reversible conformational changes of the entire structure on ∼minute timescales. Our results demonstrate programmable motion of 2D and 3D DNA origami mechanisms constructed following a macroscopic machine design approach.The ability to control, manipulate, and organize matter at the nanoscale has demonstrated immense potential for advancements in industrial technology, medicine, and materials (1–3). Bottom-up self-assembly has become a particularly promising area for nanofabrication (4, 5); however, to date designing complex motion at the nanoscale remains a challenge (6–9). Amino acid polymers exhibit well-defined and complex dynamics in natural systems and have been assembled into designed structures including nanotubes, sheets, and networks (10–12), although the complexity of interactions that govern amino acid folding make designing complex geometries extremely challenging. DNA nanotechnology, on the other hand, has exploited well-understood assembly properties of DNA to create a variety of increasingly complex designed nanostructures (13–15).Scaffolded DNA origami, the process of folding a long single-stranded DNA (ssDNA) strand into a custom structure (16–18), has enabled the fabrication of nanoscale objects with unprecedented geometric complexity that have recently been implemented in applications such as containers for drug delivery (19, 20), nanopores for single-molecule sensing (21–23), and templates for nanoparticles (24, 25) or proteins (26–28). The majority of these and other applications of DNA origami have largely focused on static structures. Natural biomolecular machines, in contrast, have a rich diversity of functionalities that rely on complex but well-defined and reversible conformational changes. Currently, the scope of biomolecular nanotechnology is limited by an inability to achieve similar motion in designed nanosystems.DNA nanotechnology has enabled critical steps toward that goal starting with the work of Mao et al. (29), who developed a DNA nanostructure that took advantage of the B–Z transition of DNA to switch states. Since then, efforts to fabricate dynamic DNA systems have primarily focused on strand displacement approaches (30) mainly on systems comprising a few strands or arrays of strands undergoing ∼nm-scale motions (31–37) in some cases guided by DNA origami templates (38–40). More recently, strand displacement has been used to reconfigure DNA origami nanostructures, for example opening DNA containers (19, 41, 42), controlling molecular binding (43, 44), or reconfiguring structures (45). The largest triggerable structural change was achieved by Han et al. in a DNA origami Möbius strip (one-sided ribbon structure) that could be opened to approximately double in size (45). Constrained motion has been achieved in systems with rotational motion (19, 20, 32, 41, 44, 46, 47) in some cases to open lid-like components (19, 20, 41) or detect molecular binding (44, 48, 49). A few of these systems achieved reversible conformational changes (32, 41, 44, 46), although the motion path and flexibility were not studied. Constrained linear motion has remained largely unexplored. Linear displacements on the scale of a few nanometers have been demonstrated via conformational changes of DNA structure motifs (50–55), strand invasion to open DNA hairpins (36, 55, 56), or the reversible sliding motion of a DNA tile actuator (56); these cases also did not investigate the motion path or flexibility of motion.Building on these prior studies, this work implements concepts from macroscopic machine design to build modular parts with constrained motion. We demonstrate an ability to tune the flexibility and range of motion and then integrate these parts into prototype mechanisms with designed 2D and 3D motion. We further demonstrate reversible actuation of a mechanism with complex conformational changes on minute timescales.     

Nonequilibrium dynamics and ultraslow relaxation of confined DNA during viral packaging

Many viruses use molecular motors that generate large forces to package DNA to near-crystalline densities inside preformed viral proheads. Besides being a key step in viral assembly, this process is of interest as a model for understanding the physics of charged polymers under tight 3D confinement. A large number of theoretical studies have modeled DNA packaging, and the nature of the molecular dynamics and the forces resisting the tight confinement is a subject of wide debate. Here, we directly measure the packaging of single DNA molecules in bacteriophage phi29 with optical tweezers. Using a new technique in which we stall the motor and restart it after increasing waiting periods, we show that the DNA undergoes nonequilibrium conformational dynamics during packaging. We show that the relaxation time of the confined DNA is >10 min, which is longer than the time to package the viral genome and 60,000 times longer than that of the unconfined DNA in solution. Thus, the confined DNA molecule becomes kinetically constrained on the timescale of packaging, exhibiting glassy dynamics, which slows the motor, causes significant heterogeneity in packaging rates of individual viruses, and explains the frequent pausing observed in DNA translocation. These results support several recent hypotheses proposed based on polymer dynamics simulations and show that packaging cannot be fully understood by quasistatic thermodynamic models.DNA packaging is both a critical step in viral assembly and a unique model for understanding the physics of polymers under strong confinement. Before packaging, the DNA (∼6–60 µm long) forms a loose random coil of diameter ∼1–3 µm. After translocation into the viral prohead (∼50–100 nm in diameter), a ∼10,000-fold volume compaction is achieved. Packaging is driven by a powerful molecular motor that must work against the large forces resisting confinement arising from DNA bending, repulsion between DNA segments, and entropy loss (1–8).DNA packaging in bacteriophages phi29, lambda, and T4 has been directly measured via single-molecule manipulation with optical tweezers and the packaging motors have been shown to generate forces of >60 pN, among the highest known for biomotors, while translocating DNA at rates ranging from ∼100 bp (for phage phi29, which packages a 19.3-kbp genome into a 42 × 54-nm prohead shell) up to as high as ∼2,000 bp/s (for phage T4, which packages a 171-kbp genome into a 120 × 86-nm prohead) (9–15). The force resisting packaging rises steeply with prohead filling and has been proposed to play an important role in driving viral DNA ejection (16).Recently, a variety of theoretical models for viral DNA packaging have been proposed (3–5, 17–21). The simplest treat DNA as an elastic rod with repulsive self-interactions and assume that packaging is a quasistatic thermodynamic process, i.e., that the DNA is able to continuously relax to a free-energy minimum state (3–5, 19–21). The DNA arrangement is generally assumed to be an inverse spool with local hexagonal close packing between DNA segments, as suggested by electron microscopy and X-ray scattering studies (22, 23). Such models yield exact analytical predictions that reproduce many of the experimental trends, including the sharp rise in resistance during the latter stages of packaging (3–5, 20).Dynamic simulations, however, predict differing results. Depending on model and simulation protocol, some predict rapid equilibration into ordered spool or folded toroid conformations, whereas others predict nonequilibrium dynamics and disordered conformations (3, 6, 24–31). The packaged DNA conformation also depends on ionic conditions, capsid size and shape, and shape of the internal core structure found in some phages (6, 30). Notably, some electron microscopy studies have also been interpreted as suggesting ordered spooled conformations (22), whereas others have been interpreted as suggesting partly disordered conformations (29). Although some simulations predict nonequilibrium dynamics, several potential caveats are that (i) the DNA has been represented by coarse-grained polymer models with various approximations for physical interactions (6), (ii) the packaging rate used in the simulations is >10⁵ times higher than the measured packaging rate due to computational constraints (3, 26–28), and (iii) it has been pointed out by some authors that simulation timescale cannot be directly related to experimental timescale because of the use of coarse-grained models for DNA (25, 28). As noted in early modeling studies, the calculations based on quasistatic models may represent a lower bound on the required packaging forces due to dissipative dynamic losses (4). Whether nonequilibrium dynamics play a significant role in real systems has thus remained an important open question.     

Enhanced expression of DNA polymerase eta contributes to cisplatin resistance of ovarian cancer stem cells

Cancer stem cells (CSCs) with enhanced tumorigenicity and chemoresistance are believed to be responsible for treatment failure and tumor relapse in ovarian cancer patients. However, it is still unclear how CSCs survive DNA-damaging agent treatment. Here, we report an elevated expression of DNA polymerase η (Pol η) in ovarian CSCs isolated from both ovarian cancer cell lines and primary tumors, indicating that CSCs may have intrinsically enhanced translesion DNA synthesis (TLS). Down-regulation of Pol η blocked cisplatin-induced CSC enrichment both in vitro and in vivo through the enhancement of cisplatin-induced apoptosis in CSCs, indicating that Pol η-mediated TLS contributes to the survival of CSCs upon cisplatin treatment. Furthermore, our data demonstrated a depletion of miR-93 in ovarian CSCs. Enforced expression of miR-93 in ovarian CSCs reduced Pol η expression and increased their sensitivity to cisplatin. Taken together, our data suggest that ovarian CSCs have intrinsically enhanced Pol η-mediated TLS, allowing CSCs to survive cisplatin treatment, leading to tumor relapse. Targeting Pol η, probably through enhancement of miR-93 expression, might be exploited as a strategy to increase the efficacy of cisplatin treatment.Ovarian cancer is the most lethal malignancy of the female reproductive tract with a 5-y survival rate of only 27% in advanced stages (1). The American Cancer Society estimates that in 2014, about 21,980 new cases of ovarian cancer will be diagnosed and 14,270 women will die of ovarian cancer in the United States (1). The mainline treatment of ovarian cancer is cytoreductive surgery followed by platinum (Pt)-based chemotherapy (2). Chemotherapy with Pt is initially effective for most patients. However, the majority eventually becomes refractory to Pt treatment, and around 70% of patients have tumor relapses (3). Poor understanding of the underlying mechanisms of this acquired drug resistance and tumor relapse poses a critical cancer research challenge.cis-diamminedichloroplatinum(II) (cisplatin), the first member of Pt-based chemotherapeutic agents, has been widely used to treat various malignant tumors, including ovarian cancer (4). Mechanistically, cisplatin reacts with DNA bases to cross-link adjacent purines. These cross-links block DNA replication and invoke apoptosis in rapidly dividing cells (5). Thus, the preferential activation of the DNA damage responses, especially the efficient removal of these DNA lesions, or prompt rescue of the replication, will prevent replication fork collapse and promote survival of the cells upon cisplatin treatment, eventually leading to cisplatin resistance. The cisplatin-induced DNA cross-links are primarily removed by the nucleotide excision repair (NER) pathway (6) or bypassed during replication through translesion DNA synthesis (TLS) (7–10). TLS is mediated by specialized DNA polymerases (Pols), which are characterized by low fidelity and an ability to replicate across certain types of damaged sites in template DNA with the assistance of monoubiquitylated proliferating cell nuclear antigen (ub-PCNA) (11). TLS rescues cells from the collapse of the replication fork and thus is believed to contribute to the development of cisplatin resistance (8, 12–17).It has been increasingly evident that heterogeneous ovarian cancers contain a subpopulation of cancer stem cells (CSCs) with enhanced tumorigenicity and chemoresistance. These CSCs are believed to be responsible for treatment failure and tumor relapse. Ovarian CSCs have been successfully isolated, based on the expression of distinctive cell surface markers CD44 and CD117 (18, 19), their ability to efflux the Hoechst 33342 fluorescent dye (Side population, SP) (20), the activity of ALDH (21), and their ability to grow as floating spheres in serum-free medium (19). The CD44⁺CD117⁺ cells, SP cells, ALDH⁺ cells, and spheroid cells isolated from both ovarian cancer cell lines and primary human ovarian tumors fulfill all currently accepted criteria for the existence of a subpopulation of tumor-initiating cells (19, 22, 23). Most importantly, these CSCs also demonstrate increased cisplatin resistance. However, it is still unclear how CSCs survive cisplatin treatment. In this study, we demonstrated that the expression level of TLS Pol η is higher in ovarian CSCs isolated from both cancer cell lines and primary tumors than the bulk cancer cells. Down-regulation of Pol η expression blocked cisplatin-induced enrichment of the CSC population, through facilitating the killing of CSCs by cisplatin. Mechanistic investigation demonstrated that decreased expression of miR-93 in ovarian CSCs contributes, at least partially, to the enhanced expression of Pol η. Taken together, our study suggests that Pol η-mediated TLS could be a target to facilitate the eradication of ovarian CSCs by cisplatin.     

From the Cover: Human Pol ζ purified with accessory subunits is active in translesion DNA synthesis and complements Pol η in cisplatin bypass

DNA polymerase ζ (Pol ζ) is a eukaryotic B-family DNA polymerase that specializes in translesion synthesis and is essential for normal embryogenesis. At a minimum, Pol ζ consists of a catalytic subunit Rev3 and an accessory subunit Rev7. Mammalian Rev3 contains >3,000 residues and is twice as large as the yeast homolog. To date, no vertebrate Pol ζ has been purified for biochemical characterization. Here we report purification of a series of human Rev3 deletion constructs expressed in HEK293 cells and identification of a minimally catalytically active human Pol ζ variant. With a tagged form of an active Pol ζ variant, we isolated two additional accessory subunits of human Pol ζ, PolD2 and PolD3. The purified four-subunit Pol ζ4 (Rev3–Rev7–PolD2–PolD3) is much more efficient and more processive at bypassing a 1,2-intrastrand d(GpG)-cisplatin cross-link than the two-subunit Pol ζ2 (Rev3–Rev7). We show that complete bypass of cisplatin lesions requires Pol η to insert dCTP opposite the 3′ guanine and Pol ζ4 to extend the primers.DNA polymerase ζ (Pol ζ), composed of the catalytic Rev3 and accessary Rev7 subunits, is an error-prone DNA translesion polymerase that causes both spontaneous and DNA damage-induced mutagenesis (1, 2). More than two-thirds of the 1,504 residues in yeast Rev3 share sequence homology with all B-family DNA polymerases, including Pols α, δ, and ε, which are responsible for the bulk of high-fidelity genomic replication in eukaryotes (3). Unlike the typical B-family polymerases, Pol ζ lacks an intrinsic 3′–5′ exonuclease activity and thus has no proofreading function (2). Human homologs of REV3 (REV3L) and REV7 (MAD2L2; hereafter referred to as REV7) genes were identified shortly after yeast Pol ζ was characterized. Human Rev3 contains 3,130 residues and is twice as large as the yeast counterpart (4). Human and yeast Rev7 are homologous (5) and bear sequence similarity to the mitotic checkpoint proteins Mad2 (6). Unlike Saccharomyces cerevisiae REV3 and REV7 genes, which are nonessential and whose knockout leads only to a decreased rate of damage-induced mutagenesis (7, 8), Rev3l knockout in mice is embryonic-lethal (9), and mouse Rev3l^−/− embryonic stem cells are not viable (10, 11). Human and mouse cell cultures obtained from conditional Rev3l knockout show genome instability and growth defects without an external challenge of DNA damage (12–14). DNA pol ζ is apparently essential for normal cell proliferation and embryogenesis in mammals.Translesion synthesis (TLS) and DNA-damage-induced mutagenesis are the best-characterized functions of Pol ζ. Absence of the yeast REV3 gene leads to sensitivity to UV light and intrastrand and interstrand cross-linking agents (2, 15). DNA Pol ζ has been shown to induce multiple base substitutions as well as more complex mutations in yeast (7, 16, 17) and may contribute to hypermutation in Ig genes in mammals (18, 19). The TLS function of DNA Pol ζ has been implicated in its role of mediating resistance to platinum-based chemotherapies (20–22). Owing to the conservation of B-family DNA polymerases, a distorted DNA template base is unlikely to be accommodated in the active site of DNA Pol ζ. In fact, yeast DNA Pol ζ is unable to insert a nucleotide opposite either a cis–syn thymine or a 6-4 photoproduct (23). Genetic data indicate that a complete lesion bypass event may require two TLS DNA polymerases (24)—one for nucleotide incorporation opposite a lesion (insertion step) and the other for the subsequent primer extension (extension step). The insertion step of TLS is often accomplished by a Y-family polymerase, whose active site is uncommonly large, solvent-exposed, and flexible (25). Studies of another B-family TLS DNA polymerase from Escherichia coli (Pol II) show that it efficiently extends a DNA primer after a lesion by looping out the damaged DNA template strand, leading to frameshift and mixed-type mutations (26).In budding yeast, REV3 has been shown to be epistatic with POL32, a subunit of DNA Pol δ. Inactivating either REV3 or POL32 leads to reduced spontaneous mutagenesis (27–29). As with all eukaryotic B-family DNA polymerases, Rev3 contains a Cys-rich C-terminal domain (CTD) (30–33), which forms a zinc-finger domain followed by a [4Fe–4S] cluster (34). In Pol α, δ, and ε, each CTD interacts with its specific accessary subunits (32, 35). Recently, three groups have independently shown that the [4Fe–4S] cluster of yeast Rev3 interacts with Pol31 and Pol32 subunit (36), thus forming an stoichiometric four-subunit Pol ζ (Pol ζ4; Rev3–Rev7–Pol31–Pol32) (23, 37, 38). Baranovskiy et al. further showed that the CTDs of human Pol ζ and δ share the same accessary subunits p50 and p66, homologs of yeast Pol31 and Pol32, respectively (37). The interaction between yeast Rev3 and Pol31 is reported to be direct, and Pol32 is essential to stabilize Pol31 and, furthermore, via its interactions with proliferating cell nuclear antigen (PCNA), recruits and activates Pol ζ to carry out TLS (38). The catalytic activity of yeast Pol ζ is improved by the presence of Pol31 and Pol32 (23, 38).Purification and characterization of Pol ζ has so far been limited to the yeast protein. Perhaps because of its large size, mammalian Pol ζ has not been purified for biochemical characterization. To overcome this roadblock, we coexpressed human REV3L and REV7 in mammalian cells in culture. Initially, very low expression level and heterogeneity was encountered, but these problems were solved by targeted deletion of various internal segments of human REV3L. We succeeded in purifying an active two-subunit form of human Pol ζ (Pol ζ2). By differential pull-down experiments using Pol ζ2 variants with and without the CTD of Rev3, we isolated two CTD-dependent Pol ζ accessary subunits, PolD2 and PolD3. We report here purification of an active form of human four-subunit Pol ζ4 and the collaboration of two TLS polymerases, Pol η and Pol ζ, in lesion bypass.     

Structure-based cleavage mechanism of Thermus thermophilus Argonaute DNA guide strand-mediated DNA target cleavage

We report on crystal structures of ternary Thermus thermophilus Argonaute (TtAgo) complexes with 5′-phosphorylated guide DNA and a series of DNA targets. These ternary complex structures of cleavage-incompatible, cleavage-compatible, and postcleavage states solved at improved resolution up to 2.2 Å have provided molecular insights into the orchestrated positioning of catalytic residues, a pair of Mg²⁺ cations, and the putative water nucleophile positioned for in-line attack on the cleavable phosphate for TtAgo-mediated target cleavage by a RNase H-type mechanism. In addition, these ternary complex structures have provided insights into protein and DNA conformational changes that facilitate transition between cleavage-incompatible and cleavage-compatible states, including the role of a Glu finger in generating a cleavage-competent catalytic Asp-Glu-Asp-Asp tetrad. Following cleavage, the seed segment forms a stable duplex with the complementary segment of the target strand.Argonaute (Ago) proteins, critical components of the RNA-induced silencing complex, play a key role in guide strand-mediated target RNA recognition, cleavage, and product release (reviewed in refs. 1–3). Ago proteins adopt a bilobal scaffold composed of an amino terminal PAZ-containing lobe (N and PAZ domains), a carboxyl-terminal PIWI-containing lobe (Mid and PIWI domains), and connecting linkers L1 and L2. Ago proteins bind guide strands whose 5′-phosphorylated and 3′-hydroxyl ends are anchored within Mid and PAZ pockets, respectively (4–7), with the anchored guide strand then serving as a template for pairing with the target strand (8, 9). The cleavage activity of Ago resides in the RNase H fold adopted by the PIWI domain (10, 11), whereby the enzyme’s Asp-Asp-Asp/His catalytic triad (12–15) initially processes loaded double-stranded siRNAs by cleaving the passenger strand and subsequently processes guide-target RNA duplexes by cleaving the target strand (reviewed in refs. 16–18). Such Mg²⁺ cation-mediated endonucleolytic cleavage of the target RNA strand (19, 20) resulting in 3′-OH and 5′-phosphate ends (21) requires Watson–Crick pairing of the guide and target strands spanning the seed segment (positions 2–2′ to 8–8′) and the cleavage site (10′–11′ step on the target strand) (9). Insights into target RNA recognition and cleavage have emerged from structural (9), chemical (22), and biophysical (23) experiments.Notably, bacterial and archaeal Ago proteins have recently been shown to preferentially bind 5′-phosphoryated guide DNA (14, 15) and use an activated water molecule as the nucleophile (reviewed in ref. 24) to cleave both RNA and DNA target strands (9). Structural studies have been undertaken on bacterial and archaeal Ago proteins in the free state (10, 15) and bound to a 5′-phosphorylated guide DNA strand (4) and added target RNA strand (8, 9). The structural studies of Thermus thermophilus Ago (TtAgo) ternary complexes have provided insights into the nucleation, propagation, and cleavage steps of target RNA silencing in a bacterial system (9). These studies have highlighted the conformational transitions on proceeding from Ago in the free state to the binary complex (4) to the ternary complexes (8, 9) and have emphasized the requirement for a precisely aligned Asp-Asp-Asp triad and a pair of Mg²⁺ cations for cleavage chemistry (9), typical of RNase H fold-mediated enzymes (24, 25). Structural studies have also been extended to binary complexes of both human (5, 6) and yeast (7) Agos bound to 5′-phosphorylated guide RNA strands.Despite these singular advances in the structural biology of RNA silencing, further progress was hampered by the modest resolution (2.8- to 3.0-Å resolution) of TtAgo ternary complexes with guide DNA (4) and added target RNAs (8, 9). This precluded identification of water molecules coordinated with the pair of Mg²⁺ cations, including the key water that acts as a nucleophile and targets the cleavable phosphate between positions 10′-11′ on the target strand. We have now extended our research to TtAgo ternary complexes with guide DNA and target DNA strands, which has permitted us to grow crystals of ternary complexes that diffract to higher (2.2–2.3 Å) resolution in the cleavage-incompatible, cleavage-compatible, and postcleavage steps. These high-resolution structures of TtAgo ternary complexes provide snapshots of distinct key steps in the catalytic cleavage pathway, opening opportunities for experimental probing into DNA target cleavage as a defense mechanism against plasmids and possibly other mobile elements (26, 27).     

Creative template-dependent synthesis by human polymerase mu

Among the many proteins used to repair DNA double-strand breaks by nonhomologous end joining (NHEJ) are two related family X DNA polymerases, Pol λ and Pol µ. Which of these two polymerases is preferentially used for filling DNA gaps during NHEJ partly depends on sequence complementarity at the break, with Pol λ and Pol µ repairing complementary and noncomplementary ends, respectively. To better understand these substrate preferences, we present crystal structures of Pol µ on a 2-nt gapped DNA substrate, representing three steps of the catalytic cycle. In striking contrast to Pol λ, Pol µ “skips” the first available template nucleotide, instead using the template base at the 5′ end of the gap to direct nucleotide binding and incorporation. This remarkable divergence from canonical 3′-end gap filling is consistent with data on end-joining substrate specificity in cells, and provides insights into polymerase substrate choices during NHEJ.DNA double-strand breaks (DSBs) result from exposure to endogenous and exogenous factors including reactive oxygen species, physical or mechanical stress, ionizing radiation, or activities of nuclear enzymes on DNA (1). Another source of DSBs is programmed DNA breakage during meiotic or mitotic recombination, immunoglobin gene rearrangement in V(D)J, and class-switch recombination (1, 2). Because of the largely random nature of accidental double-strand breakage, the broken ends can have widely varying sequences, structures, or modifications. Nonhomologous end joining (NHEJ) is the predominant form of DSB repair in higher eukaryotes, and requires a certain degree of flexibility from the many factors involved in this complex repair process.DSB substrates lacking microhomology at the break site are unsuitable for immediate rejoining by ligase IV, and a polymerase is usually required to fill the gaps to generate ends that can be efficiently ligated (3). In higher eukaryotes, this role is performed by family X polymerase λ (Pol λ) and polymerase µ (Pol μ) (4–6). Pol λ has a strong preference for substrates with complementary template-strand pairing opposite the primer terminus. Pol µ can also use such complementary DSB substrates, but is uniquely active in template-dependent synthesis on DSBs entirely lacking complementarity, where the primer terminus is unpaired (7).Given partial overlap in substrate use in vitro by Pol µ and Pol λ, how does the NHEJ machinery select which enzyme to repair specific substrates with the highest fidelity? Recent work by Pryor et al. (8) (companion article in this issue) has demonstrated a strong preference for Pol λ over Pol µ in repairing the majority of complementary DSBs. This is advantageous, because Pol λ displays higher fidelity of synthesis (9, 10). However, Pol λ cannot use noncomplementary DSB substrates with unpaired primer termini. Pol µ is the only known polymerase that can use noncomplementary substrates, such that in Pol µ knockout cells, these ends are largely unrepaired (figures 1B and 3A in ref. 8, the companion article). To better understand the substrate preferences of Pol λ and Pol µ, here we present crystal structures of the human Pol µ (hPol µ) catalytic domain in complex with a 2-nt gapped DNA substrate, from DNA binding through nucleotide binding and incorporation. These structures are consistent with the in vivo NHEJ assays from Pryor et al. (8) and support a “1-nt gap” spacing rule for Pol µ in which Pol µ prefers to engage all substrates in a similar fashion, using the 5ʹ unpaired template base in the gap as though it were a single-nucleotide (1-nt) gap. Additionally, this study provides insights into Pol µ substrate preference during in vitro polymerization reactions and how this enzyme might function during NHEJ in vivo.     

Single-molecule imaging reveals the mechanism of Exo1 regulation by single-stranded DNA binding proteins

Exonuclease 1 (Exo1) is a 5′→3′ exonuclease and 5′-flap endonuclease that plays a critical role in multiple eukaryotic DNA repair pathways. Exo1 processing at DNA nicks and double-strand breaks creates long stretches of single-stranded DNA, which are rapidly bound by replication protein A (RPA) and other single-stranded DNA binding proteins (SSBs). Here, we use single-molecule fluorescence imaging and quantitative cell biology approaches to reveal the interplay between Exo1 and SSBs. Both human and yeast Exo1 are processive nucleases on their own. RPA rapidly strips Exo1 from DNA, and this activity is dependent on at least three RPA-encoded single-stranded DNA binding domains. Furthermore, we show that ablation of RPA in human cells increases Exo1 recruitment to damage sites. In contrast, the sensor of single-stranded DNA complex 1—a recently identified human SSB that promotes DNA resection during homologous recombination—supports processive resection by Exo1. Although RPA rapidly turns over Exo1, multiple cycles of nuclease rebinding at the same DNA site can still support limited DNA processing. These results reveal the role of single-stranded DNA binding proteins in controlling Exo1-catalyzed resection with implications for how Exo1 is regulated during DNA repair in eukaryotic cells.All DNA maintenance processes require nucleases, which enzymatically cleave the phosphodiester bonds in nucleic acids. Exo1, a member of the Rad2 family of nucleases, participates in DNA mismatch repair (MMR), double-strand break (DSB) repair, nucleotide excision repair (NER), and telomere maintenance (1–3). Exo1 is the only nuclease implicated in MMR, where its 5ʹ to 3ʹ exonuclease activity is used to remove long tracts of mismatch-containing single-stranded DNA (ssDNA) (2, 4–7). In addition, functionally deficient Exo1 variants have been identified in familial colorectal cancers, and Exo1-null mice exhibit a significant increase in tumor development, decreased lifespan, and sterility (8, 9). Exo1 also promotes DSB repair via homologous recombination (HR) by processing the free DNA ends to generate kilobase-length ssDNA resection products (1, 10–12). The resulting ssDNA is paired with a homologous DNA sequence located on a sister chromatid, and the missing genetic information is then restored via DNA synthesis. The central role of Exo1 in DNA repair is highlighted by the large set of genetic interactions between Exo1 and nearly all other DNA maintenance and metabolism pathways (13).Exo1 generates long tracts of ssDNA in both MMR and DSB repair (3). This ssDNA is rapidly bound by replication protein A (RPA), a ubiquitous heterotrimeric protein that participates in all DNA transactions that generate ssDNA intermediates (14). RPA protects the ssDNA from degradation, participates in DNA damage response signaling, and acts as a loading platform for downstream DSB repair proteins (15–17). RPA also coordinates DNA resection by removing secondary ssDNA structures and by modulating the Bloom syndrome, RecQ helicase-like (BLM)/DNA2- and Exo1-dependent DNA resection pathways (18–21). Reconstitution of both the yeast and human BLM (Sgs1 in yeast)/DNA2-dependent resection reactions established that RPA stimulates DNA unwinding by BLM/Sgs1 and enforces a 5′-endonuclease polarity on DNA2 (20, 22). However, the effect of RPA on Exo1 remains unresolved. Independent studies using reconstituted yeast proteins reported that RPA could both inhibit (23) and stimulate yeast Exo1 (yExo1) (18). Similarly, human RPA has variously been reported to stimulate (19) or inhibit human Exo1 (hExo1) (4, 5, 21).In addition to RPA, human cells also encode SOSS1, a heterotrimeric ssDNA-binding complex that is essential for HR (24). SOSS1 consists of INTS3 (SOSSA), hSSB1 (SOSSB1), and C9orf80 (SOSSC) (24–26). SOSSB1 encodes a single ssDNA-binding domain that bears structural homology to Escherichia coli ssDNA-binding protein (SSB) (24). SOSS1 foci form rapidly after induction of DNA breaks, and ablation of SOSS1 severely reduces DNA resection, γH2AX foci formation, and HR at both ionizing radiation- and restriction endonuclease-induced DSBs (12, 24, 25, 27). In vitro, SOSS1 stimulates hExo1-mediated DNA resection and may help to load hExo1 at ss/dsDNA junctions (21). However, the functional relationship between SOSS1 and RPA during hExo1 resection remains unresolved.Here, we use high-throughput single-molecule DNA curtains and quantitative cell biology to reveal the interplay between human and yeast Exo1 and SSBs during DNA resection. We show that both human and yeast Exo1s are processive nucleases, but are rapidly stripped from DNA by RPA. RPA inhibition is dependent on its multiple DNA binding domains. Remarkably, SOSS1 and other SSBs with fewer than three DNA binding domains support long-range resection by hExo1. In human cells, depletion of RPA increases the rate of hExo1 recruitment to laser-induced DNA damage but reduces the extent of resection. In the presence of RPA, both human and yeast Exo1 can resect DNA using a distributive, multiple-turnover mechanism, potentially reconciling prior conflicting in vitro observations. Together, our work reveals the mechanistic basis for how RPA and SOSS1 differentially modulate hExo1 activity and highlights an additional, unexpected role for these SSBs in DNA resection. We anticipate that these findings will shed light on how Exo1 is regulated in multiple genome maintenance pathways.     

Reconstitution of long and short patch mismatch repair reactions using Saccharomyces cerevisiae proteins

A problem in understanding eukaryotic DNA mismatch repair (MMR) mechanisms is linking insights into MMR mechanisms from genetics and cell-biology studies with those from biochemical studies of MMR proteins and reconstituted MMR reactions. This type of analysis has proven difficult because reconstitution approaches have been most successful for human MMR whereas analysis of MMR in vivo has been most advanced in the yeast Saccharomyces cerevisiae. Here, we describe the reconstitution of MMR reactions using purified S. cerevisiae proteins and mispair-containing DNA substrates. A mixture of MutS homolog 2 (Msh2)–MutS homolog 6, Exonuclease 1, replication protein A, replication factor C-Δ1N, proliferating cell nuclear antigen and DNA polymerase δ was found to repair substrates containing TG, CC, +1 (+T), +2 (+GC), and +4 (+ACGA) mispairs and either a 5′ or 3′ strand interruption with different efficiencies. The Msh2–MutS homolog 3 mispair recognition protein could substitute for the Msh2–Msh6 mispair recognition protein and showed a different specificity of repair of the different mispairs whereas addition of MutL homolog 1–postmeiotic segregation 1 had no affect on MMR. Repair was catalytic, with as many as 11 substrates repaired per molecule of Exo1. Repair of the substrates containing either a 5′ or 3′ strand interruption occurred by mispair binding-dependent 5′ excision and subsequent resynthesis with excision tracts of up to ∼2.9 kb occurring during the repair of the substrate with a 3′ strand interruption. The availability of this reconstituted MMR reaction now makes possible detailed biochemical studies of the wealth of mutations identified that affect S. cerevisiae MMR.DNA mismatch repair (MMR) is a critical DNA repair pathway that is coupled to DNA replication in eukaryotes where it corrects misincorporation errors made during DNA replication (1–9). This pathway prevents mutations and acts to prevent the development of cancer (10, 11). MMR also contributes to gene conversion by repairing mispaired bases that occur during the formation of recombination intermediates (3, 4, 12). Finally, MMR acts to suppress recombination between divergent but homologous DNA sequences, thereby preventing the formation of genome rearrangements that can result from nonallelic homologous recombination (4, 13–15).Our knowledge of the mechanism of eukaryotic MMR comes from several general lines of investigation (3–9). Studies of bacterial MMR have provided a basic mechanistic framework for comparative studies (5). Genetic and cell-biology studies, primarily in Saccharomyces cerevisiae, have identified eukaryotic MMR genes, provided models for how their gene products define MMR pathways, and elucidated some of the details of how MMR pathways interact with replication (1–4). Reconstitution studies, primarily in human systems, have identified some of the catalytic features of eukaryotic MMR (7–9, 16, 17). Biochemical and structural studies of S. cerevisiae and human MMR proteins have provided information about the function of individual MMR proteins (6–9).In eukaryotic MMR, mispairs are bound by MutS homolog 2 (Msh2)–MutS homolog 6 (Msh6) and Msh2–MutS homolog 3 (Msh3), two partially redundant complexes of MutS-related proteins (3, 4, 18, 19). These complexes recruit a MutL-related complex, called MutL homoloh 1 (Mlh1)–postmeiotic segregation 1 (Pms1) in S. cerevisiae and Mlh1–postmeiotic segregation 2 (Pms2) in human and mouse (3, 4, 20–23). The Mlh1–Pms1/Pms2 complex has an endonuclease activity suggested to play a role in the initiation of the excision step of MMR (24, 25). Downstream of mismatch recognition is a mispair excision step that can be catalyzed by Exonuclease 1 (Exo1) (26–28); however, defects in both S. cerevisiae and mouse Exo1 result in only a partial MMR deficiency, suggesting the existence of additional excision mechanisms (26, 27, 29). DNA polymerase δ, the single-strand DNA binding protein replication protein A (RPA), the sliding clamp proliferating cell nuclear antigen (PCNA), and the clamp loader replication factor C (RFC) are also required for MMR at different steps, including activation of Mlh1–Pms1/Pms2, stimulation of Exo1, potentially in Exo1-independent mispair excision, and in the gap-filling resynthesis steps of MMR (3, 16, 17, 24, 27, 30–36). Although much is known about these core MMR proteins, it is not well understood how eukaryotic MMR is coupled to DNA replication (1, 2), how excision is targeted to the newly replicated strand (1, 25, 37–39), or how different MMR mechanisms such as Exo1-dependent and -independent subpathways are selected or how many such subpathways exist (1, 24, 27, 29).S. cerevisiae has provided a number of tools for studying MMR, including forward genetic screens for mutations affecting MMR, including dominant and separation-of-function mutations, the ability to evaluate structure-based mutations in vivo, cell biological tools for visualizing and analyzing MMR proteins in vivo, and overproduction of individual MMR proteins for biochemical analysis. However, linking these tools with biochemical systems that catalyze MMR reactions in vitro for mechanistic studies has not yet been possible. Here, we describe the development of MMR reactions reconstituted using purified proteins for the analysis of MMR mechanisms.     

RecQ helicase and RecJ nuclease provide complementary functions to resect DNA for homologous recombination

Recombinational DNA repair by the RecF pathway of Escherichia coli requires the coordinated activities of RecA, RecFOR, RecQ, RecJ, and single-strand DNA binding (SSB) proteins. These proteins facilitate formation of homologously paired joint molecules between linear double-stranded (dsDNA) and supercoiled DNA. Repair starts with resection of the broken dsDNA by RecQ, a 3′→5′ helicase, RecJ, a 5′→3′ exonuclease, and SSB protein. The ends of a dsDNA break can be blunt-ended, or they may possess either 5′- or 3′-single-stranded DNA (ssDNA) overhangs of undefined length. Here we show that RecJ nuclease alone can initiate nucleolytic resection of DNA with 5′-ssDNA overhangs, and that RecQ helicase can initiate resection of DNA with blunt-ends or 3′-ssDNA overhangs by DNA unwinding. We establish that in addition to its well-known ssDNA exonuclease activity, RecJ can display dsDNA exonuclease activity, degrading 100–200 nucleotides of the strand terminating with a 5′-ssDNA overhang. The dsDNA product, with a 3′-ssDNA overhang, is an optimal substrate for RecQ, which unwinds this intermediate to reveal the complementary DNA strand with a 5′-end that is degraded iteratively by RecJ. On the other hand, RecJ cannot resect duplex DNA that is either blunt-ended or terminated with 3′-ssDNA; however, such DNA is unwound by RecQ to create ssDNA for RecJ exonuclease. RecJ requires interaction with SSB for exonucleolytic degradation of ssDNA but not dsDNA. Thus, complementary action by RecJ and RecQ permits initiation of recombinational repair from all dsDNA ends: 5′-overhangs, blunt, or 3′-overhangs. Such helicase–nuclease coordination is a common mechanism underlying resection in all organisms.Homologous recombination is a relatively error-free mechanism to repair double-stranded DNA (dsDNA) breaks (DSBs) and single-stranded DNA (ssDNA) gaps, which are produced by UV light, γ-irradiation, and chemical mutagens (1). In wild-type Escherichia coli, the labor of recombinational repair is divided between the RecBCD and RecF pathways of recombination, which are responsible for the repair of DSBs and ssDNA gaps, respectively (2–5). However, the proteins of the RecF pathway are capable of DSB repair, as well as ssDNA gap repair: in recBC mutant cells containing the suppressor mutations, sbcB and sbcC (suppressors of recBC), the proteins of the RecF pathways provide the needed recombinational DNA repair functions (2, 6).The RecF pathway in E. coli involves the functions of RecA, RecF, RecG, RecJ, RecN, RecO, RecQ, RecR, RuvA, RuvB, RuvC, and single-strand DNA binding (SSB) proteins (1, 7). The RecF pathway of recombination is evolutionarily conserved across Bacteria, with most of components present in all bacteria (8). In addition, orthologs of RecF pathway proteins are found in Eukarya. RecA promotes DNA strand invasion and exchange (9–11), as does eukaryotic Rad51 (12, 13). RecO can both anneal SSB–ssDNA complexes (14, 15) and, in conjunction with RecR (and RecF), mediate loading of RecA onto SSB–ssDNA complexes (16–18). Saccharomyces cerevisiae Rad52 is a functional homolog of RecO in that it also displays both DNA-annealing and Rad51-loading activities (19–22). The RecFOR complex promotes the loading of RecA onto SSB-coated gapped DNA at ssDNA–dsDNA junctions (17, 18) and, when mutated, is suppressed by hyperactive alleles of recA (23), a property that is shared with the yeast Rad55/57 proteins (24). Furthermore, human BRCA2 protein and a fungal analog, Brh2, are partial functional analogs of the RecFOR proteins (25–27).RecQ helicase plays several roles in both early and late steps of recombination (28, 29), as do the RecQ-family helicases in Eukarya [e.g., Sgs1 and Bloom Syndrome helicase (BLM)] (30–32). In addition, eukaryotic Exonuclease 1 (Exo1) and Dna2 helicase/nuclease function somewhat analogously, although not identically, to RecJ nuclease (33–36). The in vitro reconstitution of DSB repair in E. coli, yeast, and human have shown that resection involves specific pairs of a helicase and nuclease for DNA end resection: RecQ/RecJ, Sgs1/Dna2, BLM/DNA2, and BLM/EXO1 (28, 37–39).A comparison of DSB repair by the RecBCD and RecF pathways shows that repair starts with the processing a DSB into resected dsDNA with a 3′-ssDNA overhang (7). RecJ has a 5′ to 3′ exonuclease activity on ssDNA and the action of RecJ is facilitated by RecQ, which has a 3′ to 5′ helicase activity (40, 41). The resulting processed DNA has a 3′-ssDNA overhang. The RecFOR complex binds to the 5′-end at the junction between ssDNA and dsDNA, and loads RecA protein onto the adjacent ssDNA (17, 18). Finally, the RecA nucleoprotein filament promotes pairing with homologous dsDNA (9). These steps have been reconstituted in vitro in a coordinated reaction using RecAFORQJ and SSB proteins (28).Despite progress, most studies have used DNA substrates with simple blunt-ends. However, in vivo, there are many potential structures at the end of a DSB. When the DSB is created by a replication fork encountering nicked DNA, the break can be blunt-ended (5). However, related mechanisms can produce dsDNA with either 5′- or 3′-ssDNA overhangs. Similarly, the actual intermediates of DNA processing may result in dsDNA with either 5′- or 3′-ssDNA ends. Clearly, a DNA repair pathway must be capable of dealing with such a variety of DNA end structures. In this study, we investigated the processing of DSBs by RecJ and RecQ, both individually and together. We found that a DNA with a 5′-ssDNA overhang end was degraded by RecJ nuclease and converted into an intermediate with a 3′-ssDNA overhang. Although this intermediate was no longer a substrate for RecJ, RecQ could bind to this intermediate and initiate unwinding, thereby supplying 5′-tailed ssDNA for further resection by RecJ. In addition, we established that RecQ allows RecJ to initiate nucleolytic resection on otherwise poor substrates (e.g., blunt-end DNA or DNA with 3′-ssDNA overhangs). Thus, RecQ and RecJ cooperate biochemically to create DNA intermediates for one another that enable resection of all types of broken DNA molecules.     

From the Cover: PNAS Plus: Single-molecule visualization of RecQ helicase reveals DNA melting,nucleation, and assembly are required for processive DNA unwinding

DNA helicases are motor proteins that unwind double-stranded DNA (dsDNA) to reveal single-stranded DNA (ssDNA) needed for many biological processes. The RecQ helicase is involved in repairing damage caused by DNA breaks and stalled replication forks via homologous recombination. Here, the helicase activity of RecQ was visualized on single molecules of DNA using a fluorescent sensor that directly detects ssDNA. By monitoring the formation and progression of individual unwinding forks, we observed that both the frequency of initiation and the rate of unwinding are highly dependent on RecQ concentration. We establish that unwinding forks can initiate internally by melting dsDNA and can proceed in both directions at up to 40–60 bp/s. The findings suggest that initiation requires a RecQ dimer, and that continued processive unwinding of several kilobases involves multiple monomers at the DNA unwinding fork. We propose a distinctive model wherein RecQ melts dsDNA internally to initiate unwinding and subsequently assembles at the fork into a distribution of multimeric species, each encompassing a broad distribution of rates, to unwind DNA. These studies define the species that promote resection of DNA, proofreading of homologous pairing, and migration of Holliday junctions, and they suggest that various functional forms of RecQ can be assembled that unwind at rates tailored to the diverse biological functions of RecQ helicase.DNA helicases are ubiquitous enzymes involved in many aspects of DNA metabolism, including DNA replication, repair, and recombination. These enzymes work by coupling the hydrolysis of nucleoside triphosphates (NTPs) to unwinding of double-stranded DNA (dsDNA) to produce single-stranded DNA (ssDNA) (1). This activity allows the cell’s machinery to access the information stored within the bases of the double helix. RecQ helicase from Escherichia coli is the founding member of the RecQ family of helicases (2). These enzymes belong to the superfamily 2 (SF2) group of helicases, yet share greater sequence homology with their own family members, and they play important roles in the maintenance of genomic integrity by DNA recombination and repair (1, 3). Mutations in the human RecQ-like helicases, Bloom (BLM), Werner (WRN), and RecQ4 proteins, lead to Bloom’s, Werner’s, and Rothmund–Thomson syndromes, respectively. These genetic disorders are characterized by genomic instability and an increased incidence of cancers (4).E. coli RecQ is a 3′ → 5′ helicase that functions in DNA-break repair by homologous recombination (2, 5, 6). RecQ and RecJ, a 5′ → 3′ exonuclease, process ssDNA gaps or dsDNA breaks into ssDNA for recombinational repair by RecA (7–10). In addition, RecQ ensures recombination fidelity in vivo by removing inappropriately paired joint molecules to prevent illegitimate recombination and also by disrupting joint molecule intermediates to facilitate repair by synthesis-dependent strand annealing, preventing chromosomal crossing over (7, 9, 11, 12). RecQ also functions with topoisomerase III (Topo III), a type I topoisomerase, to catenate and decatenate DNA molecules and to separate converged replication forks (13, 14). RecQ and Topo III provide an alternative to the RuvABC pathway for disengaging double Holliday junctions and do so without producing chromosomal crossovers (15).In vitro, RecQ can unwind a multitude of DNA substrates and does not require a ssDNA tail, or even a dsDNA end, to initiate unwinding; consequently, it is distinctive in being able to unwind covalently closed circular plasmid DNA (16, 17). The winged-helix domain of RecQ is important for the recognition of this broad array of DNA substrates (18). This domain binds to dsDNA yet it adopts a flexible conformation that allows it to adapt to many DNA structures. A curious feature of RecQ is that maximal unwinding requires nearly stoichiometric amounts of protein relative to the DNA (one protein per ∼10 bp), which can be partially mitigated (one protein per ∼30 bp) by including the ssDNA binding protein, SSB (5, 6, 16, 19). This behavior is compatible with the low processivity of ssDNA translocation (∼30–100 nucleotides) by RecQ (20–22) and other RecQ members (23). Paradoxically, at limiting concentrations, most RecQ helicases nonetheless efficiently unwind several kilobases of dsDNA in the course of their normal functions (9, 16, 24–26), suggesting a dynamic unwinding process. Although RecQ can unwind DNA as a monomer, it also shows a functional cooperativity when unwinding DNA with an ssDNA tail (27–29). Thus, multiple monomers can bind to ssDNA to unwind long dsDNA regions.Fluorescence techniques coupled with single-molecule microscopy have emerged as a powerful method for studying the unwinding mechanism of helicases (30–35). These assays measure individual enzymes directly or indirectly through their actions on individual DNA molecules, thus alleviating many of the drawbacks of ensemble experiments. In particular, total internal reflection fluorescence (TIRF) microscopy permits the detection of individual fluorophores with high sensitivity in the presence of a high background (30). To visualize the activity of DNA binding proteins and helicases, TIRF microscopy can be coupled with microfluidic techniques to facilitate visualization of molecules and exchange of solution components (36–39).Ensemble assays have elucidated many features of DNA unwinding by the RecQ helicase family, but the need to average over a heterogeneous and unsynchronized population of enzymes has precluded a thorough understanding of this diverse and universally important helicase family. To permit a more insightful analysis, we directly visualized unwinding of individual molecules of DNA by RecQ helicase. Unwinding was monitored using fluorescent SSB to visualize generation of ssDNA. On single DNA molecules, we could see tracks of the fluorescent SSB binding to ssDNA produced by the helicase activity of RecQ. We show that RecQ initiates DNA unwinding via melting of duplex DNA at internal sites. Once initiated, DNA unwinding propagates either uni- or bidirectionally via the cooperative action of multiple RecQ molecules at the junction of ssDNA with dsDNA. Collectively, these observations define a stable oligomeric complex of subunits involved in processive helicase action, which is concordant with both biochemical and biological function of RecQ helicases and other helicases.