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.     

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.     

Rad51 recombinase prevents Mre11 nuclease-dependent degradation and excessive PrimPol-mediated elongation of nascent DNA after UV irradiation

After UV irradiation, DNA polymerases specialized in translesion DNA synthesis (TLS) aid DNA replication. However, it is unclear whether other mechanisms also facilitate the elongation of UV-damaged DNA. We wondered if Rad51 recombinase (Rad51), a factor that escorts replication forks, aids replication across UV lesions. We found that depletion of Rad51 impairs S-phase progression and increases cell death after UV irradiation. Interestingly, Rad51 and the TLS polymerase polη modulate the elongation of nascent DNA in different ways, suggesting that DNA elongation after UV irradiation does not exclusively rely on TLS events. In particular, Rad51 protects the DNA synthesized immediately before UV irradiation from degradation and avoids excessive elongation of nascent DNA after UV irradiation. In Rad51-depleted samples, the degradation of DNA was limited to the first minutes after UV irradiation and required the exonuclease activity of the double strand break repair nuclease (Mre11). The persistent dysregulation of nascent DNA elongation after Rad51 knockdown required Mre11, but not its exonuclease activity, and PrimPol, a DNA polymerase with primase activity. By showing a crucial contribution of Rad51 to the synthesis of nascent DNA, our results reveal an unanticipated complexity in the regulation of DNA elongation across UV-damaged templates.The DNA-binding protein Rad51 is a central component of homologous recombination repair (HRR). HRR repairs double-strand breaks (DSBs) in an error-free way and processes one-ended DSBs to reactivate collapsed replication forks (1). During HRR, DSBs are processed by the 3′-to-5′ exonuclease activity of the double strand break repair nuclease (Mre11) to generate protruding 3′ ssDNA at DSBs. The ssDNA is then coated with Rad51, a factor that catalyzes homology search and strand invasion. The loading and stabilization of Rad51/ssDNA complexes are supported by multiple mediators, such as the tumor suppressor BRCA2 (breast cancer 2) (1). Moreover, Rad51 promotes XPF1- and Exo1-mediated DSB formation after gemcitabine-induced irreversible ribonucleotide reductase inhibition, thus promoting cell death (2). The signals that redirect Rad51 into a DSB formation pathway rather than DSB repair are not yet known.The functions of Rad51 are not limited to the processing/generation of DSBs. Over the past few years, it has become evident that Rad51 escorts ongoing replication forks regardless of the presence of DSBs (3–5). Specifically, Rad51 protects persistently stalled replication forks from Mre11-mediated nucleolytic degradation and facilitates replication fork restart when the replication-halting agent hydroxyurea (HU) or aphidicolin (APH) is removed (6–19). Such novel functions of Rad51 require many HRR factors, including BCRA2, FANCD2 (Fanconi Anemia Complementation group protein D2), CtIP, BRCA1, and the WRN helicase, but are independent of HRR effectors, such as Rad54 (6, 7). Rad51-dependent fork-restart and fork-protection are distinct mechanisms, because proteins like the BLM helicase promote the former, but not the latter, process after HU treatment (6, 20). Mre11-mediated nucleolytic degradation of nascent DNA in BCRA2- and FANCD2-depleted, HU-treated cells was suggested to take place at the unprotected ends of reversed forks, which may mimic DSBs (6–8). Conversely, two recent reports suggest that Rad51 prevents pathological Mre11-dependent nucleolytic degradation of nonreversed stalled forks and promotes controlled DNA2-dependent exonucleolytic processing of reversed forks (15, 21).DSB-independent Rad51 functions were revealed by the use of agents that cause a significant degree (more than 40%) of replication fork stalling, such as HU, camptothecin (CPT), and mitomycin C (MMC) (15). Much less is known about the participation of Rad51 in the replication across DNA lesions that do not persistently halt replication forks and only cause a moderate reduction in the replication speed (e.g., UV-induced DNA lesions) (22). Multiple mechanisms aid DNA replication after UV irradiation. First, strongly distorting UV lesions are effectively removed by nucleotide excision repair (NER). Second, mildly distorting lesions, which are less efficiently removed by NER, can be used as replication templates in translesion DNA synthesis (TLS) events (23). TLS avoids fork stalling by loading specialized DNA polymerases that use damaged DNA as replication templates (24).It is currently unclear whether non-TLS events aid DNA replication across UV-damaged DNA in mammalian systems. Importantly, Rad51 is recruited to DNA after UV irradiation (4, 21, 25). HRR factors contribute to the repair of infrequent DSBs generated by high UV doses (80 J/m²) in NER-deficient backgrounds (26). Interestingly, however, cancer cells depleted from Rad51 were sensitive to much lower UV doses (1–5 J/m²) (26), thus suggesting DSB-independent functions of Rad51 in the cellular response to UV light. More recently, Rad51 was shown to maintain continuous DNA replication after treatment with methyl-methane sulfonate (MMS), a DNA-damaging agent that induces bulky lesions similar to the lesions caused by UV radiation (4). Whether Rad51 prevents nucleolytic degradation of nascent DNA in response to UV irradiation has not yet been explored. Remarkably, fork reversal takes place frequently after UV irradiation, similar to the case with HU, MMC, and CPT (21).We therefore set out to investigate the contribution of Rad51 to DNA replication across UV lesions. Using the DNA stretching technique (27), we uncovered two DSB-independent roles of Rad51 in the replication of UV-damaged DNA. First, Rad51 protected the nascent DNA from rapid and time-limited Mre11-dependent exonucleolytic degradation. Second, Rad51 prevented excessive DNA elongation after UV irradiation. Such dysregulated elongation of DNA was orchestrated by Mre11, but not by its exonuclease activity, and a DNA polymerase with primase activity, PrimPol (primase polymerase). Our results therefore suggest that Rad51 depletion increases repriming after UV irradiation. Intriguingly, both Rad51-mediated functions affected the accumulation of DNA damage response (DDR) markers at later time points, but only the excessive fork elongation in Rad51-depleted cells was associated with cell death. Finally, we demonstrate that the TLS DNA polymerase polη and Rad51 are both required to achieve optimal elongation of nascent UV-damaged DNA.     

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).     

C-terminal region of activation-induced cytidine deaminase (AID) is required for efficient class switch recombination and gene conversion

Activation-induced cytidine deaminase (AID) introduces single-strand breaks (SSBs) to initiate class switch recombination (CSR), gene conversion (GC), and somatic hypermutation (SHM). CSR is mediated by double-strand breaks (DSBs) at donor and acceptor switch (S) regions, followed by pairing of DSB ends in two S regions and their joining. Because AID mutations at its C-terminal region drastically impair CSR but retain its DNA cleavage and SHM activity, the C-terminal region of AID likely is required for the recombination step after the DNA cleavage. To test this hypothesis, we analyzed the recombination junctions generated by AID C-terminal mutants and found that 0- to 3-bp microhomology junctions are relatively less abundant, possibly reflecting the defects of the classical nonhomologous end joining (C-NHEJ). Consistently, the accumulation of C-NHEJ factors such as Ku80 and XRCC4 was decreased at the cleaved S region. In contrast, an SSB-binding protein, poly (ADP)-ribose polymerase1, was recruited more abundantly, suggesting a defect in conversion from SSB to DSB. In addition, recruitment of critical DNA synapse factors such as 53BP1, DNA PKcs, and UNG at the S region was reduced during CSR. Furthermore, the chromosome conformation capture assay revealed that DNA synapse formation is impaired drastically in the AID C-terminal mutants. Interestingly, these mutants showed relative reduction in GC compared with SHM in chicken DT40 cells. Collectively, our data indicate that the C-terminal region of AID is required for efficient generation of DSB in CSR and GC and thus for the subsequent pairing of cleaved DNA ends during recombination in CSR.Activation-induced cytidine deaminase (AID) is essential for three different genetic events: class switch recombination (CSR), gene conversion (GC), and somatic hypermutation (SHM), which contribute to Ig gene diversification (1–5). Although AID generates single-strand breaks (SSBs) in the Ig genes, subsequent repair steps for CSR and GC are similar to each other but are distinct from SHM in their mechanistic properties, i.e, in (i) generation of the double-strand breaks (DSBs), (ii) recombination, and (iii) the requirement for uracil-DNA-glycosylase (UNG) for the pairing of the DSB ends (6–10). Despite the similarities between GC and CSR, their repair mechanisms have distinct features: CSR recombination requires nonhomologous end joining (NHEJ), and GC depends on homologous recombination (HR). During CSR, DSB ends normally are joined by classical NHEJ (C-NHEJ), which requires specific repair proteins such as Ku80, XRCC4, or DNA ligase IV (11, 12). In the absence of C-NHEJ, a back-up end-joining pathway called “alternative end joining” (A-EJ), which is reported to be slower and also more error prone than C-NHEJ, joins the broken DSBs ends (13). On the other hand, HR, the most common form of homology-directed repair, requires long sequence homology between donor and acceptor DNA to complete the recombination step by recruiting a distinct set of repair proteins such as RAD54, RAD52, and RAD51 to the break sites (14, 15).Various studies on AID mutations in the N-terminal or C-terminal regions (4, 8, 9, 16–19) have shown that N-terminal AID mutants are compromised for CSR and are defective in SHM, indicating that the N-terminal region of AID is required for DNA cleavage (9, 16, 19). On the other hand, the C-terminal region of AID, which contains a nuclear-export signal and is responsible for AID’s shuttling activity between the nucleus and cytoplasm, is required for CSR-specific activity but not for DNA cleavage activity and SHM (8, 16). Among the series of AID C-terminal mutants examined, two mutants show characteristic features: P20, which has an insertion of 34 amino acids at residue 182 and normal nuclear-cytoplasmic shuttling activity, and JP8Bdel, which has a 16-amino acid truncation at residue 183, accumulates in the nucleus, and shows higher DNA break activity at the donor switch (S) region (16, 17). Although several reports suggest that the C-terminal region of AID is involved in protein stability (20, 21), C-terminal mutants of AID stabilized by fusing the hormone-binding domain of estrogen receptor (ER) also show similar CSR-defective phenotypes (8). Taken together, these data suggest that the DNA cleavage activity and CSR-specific activity depend on different regions of AID (8, 19). In addition, the C-terminal region of AID is essential for the interaction of AID with poly (A)⁺ RNA via a specific cofactor (22). Because CSR requires de novo protein synthesis, we proposed that after DNA cleavage the C-terminal region of AID may be involved in the regulation of the recombination step through generation of a new protein (8, 16, 22).DSBs induced by AID during CSR ultimately are joined by the efficient DNA repair pathway that requires C-NHEJ factors such as Ku70/80 (12, 23). However, in the absence of C-NHEJ, the A-EJ pathway that relies on microhomology can join the broken DNA ends, although this pathway is associated with chromosomal translocations (11, 24). Previously, we reported that JP8Bdel enhances aberrant c-myc/IgH translocations and that it fails to carry out the efficient recombination between donor and acceptor S regions in the IgH locus (8). Therefore, it is important to examine whether the AID C-terminal mutants affect the S–S joining in CSR.In the current work we examined whether the C-terminal region of AID is involved in DNA synapse formation and recombination during CSR in CH12F3-2 and spleen B cells. We also examined the effect of AID C-terminal mutations on GC in chicken DT40 cells, which depends on HR between pseudo V genes and the downstream IgVλ region. Using these CSR- and GC-monitoring systems, we demonstrate that efficient CSR and GC require the C-terminal region of AID for the formation of DSB from SSB and subsequent end synapse. Considering these findings together, we conclude that, in addition to DNA cleavage, AID has a unique function in the generation of DSBs, which is required for S–S synapse formation and joining in CSR and recombination in GC.     

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.     

From the Cover: PNAS Plus: Single-molecule motions and interactions in live cells reveal target search dynamics in mismatch repair

MutS is responsible for initiating the correction of DNA replication errors. To understand how MutS searches for and identifies rare base-pair mismatches, we characterized the dynamic movement of MutS and the replisome in real time using superresolution microscopy and single-molecule tracking in living cells. We report that MutS dynamics are heterogeneous in cells, with one MutS population exploring the nucleoid rapidly, while another MutS population moves to and transiently dwells at the replisome region, even in the absence of appreciable mismatch formation. Analysis of MutS motion shows that the speed of MutS is correlated with its separation distance from the replisome and that MutS motion slows when it enters the replisome region. We also show that mismatch detection increases MutS speed, supporting the model for MutS sliding clamp formation after mismatch recognition. Using variants of MutS and the replication processivity clamp to impair mismatch repair, we find that MutS dynamically moves to and from the replisome before mismatch binding to scan for errors. Furthermore, a block to DNA synthesis shows that MutS is only capable of binding mismatches near the replisome. It is well-established that MutS engages in an ATPase cycle, which is necessary for signaling downstream events. We show that a variant of MutS with a nucleotide binding defect is no longer capable of dynamic movement to and from the replisome, showing that proper nucleotide binding is critical for MutS to localize to the replisome in vivo. Our results provide mechanistic insight into the trafficking and movement of MutS in live cells as it searches for mismatches.DNA mismatch repair (MMR) is the highly conserved process responsible for correcting DNA replication errors (1). Although replication errors occur infrequently in bacteria (∼1 error per 31 million bp) (2), the consequences of MMR failure on human health are severe (3). MutS is the first protein involved in the MMR pathway, and it is responsible for detecting rare base-pairing errors. In Bacillus subtilis, MutS then recruits MutL, an endonuclease in most bacteria and eukaryotic organisms, to incise the DNA (4, 5). After MutL incision, the error-containing strand is removed, and the DNA is resynthesized to complete error correction (6).The mechanism by which MutS homologs locate a single mismatch among millions of correctly paired nucleotides has been studied extensively by bulk biochemistry, in vitro atomic force microscopy and single-molecule imaging, and visualizing MutS using in vivo cell biology approaches (7–20). In vitro single-molecule studies largely indicate that MutS operates as a searching clamp diffusing along DNA in a 1D search process (8, 10, 13). In this model, after mismatch recognition, MutS can dwell at the mismatch before exchanging ADP for ATP, converting into a stable ATP-bound sliding clamp, and subsequently, diffusing away from the mismatch at a faster rate in search of MutL and possible strand discrimination signals (8, 10, 15). Studying the mechanistic steps of the search process with DNA curtains has provided evidence that MutS may identify errors through a combination of 1D sliding and a 3D pathway (15). These results show that MutS predominantly searches DNA by 1D diffusion but is also capable of mismatch recognition through a 3D pathway, in which MutS binds the mismatch without engaging in a prior 1D search. The 3D pathway for mismatch recognition has been proposed to help MutS circumvent protein barriers that would exist in vivo. Additional analysis of MutS on DNA curtains suggests that DNA binding proteins largely prevent 1D diffusion by MutS homologs (14). These results indicate that a search mechanism limited to 1D diffusion would present a significant challenge for identifying replication errors in vivo. Therefore, it is still not clear how MutS searches for errors within the supercoiled, compacted, and protein-laden nucleoid in the crowded 3D environment of a living bacterial cell.Each of these in vitro single-molecule studies has provided important insight into the target search dynamics of MutS, and these studies and other bulk studies have elucidated many of the mechanistic steps leading to mismatch identification on naked DNA in vitro. Of course, most of these studies have been limited to analysis of MutS in isolation on protein-free DNA; they often do not incorporate replication proteins or other in vivo obstacles that are likely to impact the in vivo search process. The search dynamics of MutS have not been investigated at single-molecule resolution for any MutS homolog in live cells.The need to understand how the biochemistry of MutS homologs translates into mismatch identification in live cells has led to several studies across many organisms using a variety of approaches, including fluorescent protein fusions and bulk fluorescence imaging. These studies have shown that MutS homologs form foci that colocalize with the DNA replication machinery (replisome) and that interaction of MutS with replication processivity clamps [proliferating cell nuclear antigen (PCNA) or β-clamp] is important for focus formation (16, 17, 19–22). In Saccharomyces cerevisiae, MMR is also temporally coupled to DNA replication (23). However, the colocalization between bacterial MutS and the replisome is far from absolute. For example, in the absence of an exogenous mutagen, bulk microscopy detects fluorescent MutS foci in only ∼9% of B. subtilis cells. Furthermore, only about one-half of these 9% of cells with a MutS focus show colocalization of MutS to the replisome (20). Therefore, only about 4.5% of cells under normal growth conditions in B. subtilis show colocalization between MutS and the replisome (20). Obviously, the behavior of MutS in the remaining 91% of cells without a MutS focus has been impossible to determine using ensemble fluorescence techniques.Most of the MutS pool cannot be visualized by bulk microscopy, because the large fluorescent foci that are visible by ensemble fluorescence imaging account for only 10% of cellular MutS (that is, ∼8 of 80 dimers present in a cell) (24). Therefore, prior studies have been unable to image and quantify the behavior of MutS outside of large static foci visible during ensemble fluorescence microscopy. Therefore, to understand MutS location and dynamics in vivo, only a more sensitive method with higher spatial and temporal resolutions can unambiguously determine the extent to which MutS is enriched near the replisome and answer two open questions. Is MutS enrichment at the replisome constitutive in bacteria? Does MutS search for mismatches away from the replisome in vivo?To further understand the process of MMR inside cells and gain insight into the movement and location of MutS molecules in vivo throughout MMR, we probed the spatial distribution, dynamics, and genomic distribution of MutS in live bacteria with superresolution fluorescence imaging (25–27), single-molecule tracking (28–30), and ChIP-sequencing (ChIP-seq) approaches. We investigated the effect of replisome association, active DNA synthesis, mismatch binding, nucleotide binding, and presence of MutL on the dynamics of MutS movement in vivo. This study applies single-molecule imaging to a dedicated DNA repair pathway and captures the trafficking behavior of MutS, a protein that is conserved from bacteria to humans. A high degree of spatial and temporal resolution has allowed us to view the dynamic movement and search process of single MutS molecules during MMR in vivo.     

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.     

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.     

Colon cancer-associated mutator DNA polymerase δ variant causes expansion of dNTP pools increasing its own infidelity

Defects in DNA polymerases δ (Polδ) and ε (Polε) cause hereditary colorectal cancer and have been implicated in the etiology of some sporadic colorectal and endometrial tumors. We previously reported that the yeast pol3-R696W allele mimicking a human cancer-associated variant, POLD1-R689W, causes a catastrophic increase in spontaneous mutagenesis. Here, we describe the mechanism of this extraordinary mutator effect. We found that the mutation rate increased synergistically when the R696W mutation was combined with defects in Polδ proofreading or mismatch repair, indicating that pathways correcting DNA replication errors are not compromised in pol3-R696W mutants. DNA synthesis by purified Polδ-R696W was error-prone, but not to the extent that could account for the unprecedented mutator phenotype of pol3-R696W strains. In a search for cellular factors that augment the mutagenic potential of Polδ-R696W, we discovered that pol3-R696W causes S-phase checkpoint-dependent elevation of dNTP pools. Abrogating this elevation by strategic mutations in dNTP metabolism genes eliminated the mutator effect of pol3-R696W, whereas restoration of high intracellular dNTP levels restored the mutator phenotype. Further, the use of dNTP concentrations present in pol3-R696W cells for in vitro DNA synthesis greatly decreased the fidelity of Polδ-R696W and produced a mutation spectrum strikingly similar to the spectrum observed in vivo. The results support a model in which (i) faulty synthesis by Polδ-R696W leads to a checkpoint-dependent increase in dNTP levels and (ii) this increase mediates the hypermutator effect of Polδ-R696W by facilitating the extension of mismatched primer termini it creates and by promoting further errors that continue to fuel the mutagenic pathway.When functioning properly, DNA replication is phenomenally accurate, making ∼10⁻¹⁰ mutations per base pair during each replication cycle (1). In respect to human biology, it means that, on average, less than one mutation occurs each time the human genome is replicated. This amazing exactitude is contingent upon the serial action of DNA polymerase selectivity, exonucleolytic proofreading, and DNA mismatch repair (MMR). MMR defects increase spontaneous mutagenesis in numerous model systems (2) and give rise to cancer in mice (3). In humans, inherited mutations in MMR genes predispose to colorectal cancer (CRC) in Lynch syndrome (4, 5). Additionally, MMR genes are inactivated via hypermethylation in ∼15% of sporadic CRC, endometrial cancer (EC), and gastric cancer (6).Like defects in MMR, mutations that decrease the base selectivity or proofreading activity of replicative DNA polymerases elevate spontaneous mutagenesis in eukaryotic cells (7–14) and cancer incidence in mice (15–18). Germline mutations affecting the exonuclease domains of DNA polymerases δ (Polδ) and ε (Polε) cause hereditary CRC (19), and somatic changes in the exonuclease domain of Polε were found in sporadic hypermutated CRC and EC (20–23). Two of these exonuclease domain mutations were modeled in yeast and found to increase mutation rate, supporting their role in the development of hypermutated tumors (19, 24). Because of these discoveries, recent publications emphasized proofreading deficiency as the initiating cause of some human tumors (25–28). The potential role of base selectivity defects in human cancers received much less attention. At the same time, evidence is accumulating that base selectivity defects do occur in tumors and can have dramatic consequences for genome stability. Modeling in yeast of the Polε exonuclease domain mutation most prevalent in CRC and EC produced an increase in the mutation rate far exceeding the increase expected from loss of proofreading, suggesting additional fidelity defects (24). Several studies of colon cancer cell lines and primary tumors reported amino acid changes in Polδ and Polε outside of the exonuclease domain, including some in the conserved DNA polymerase motifs (20, 23, 29). We have previously shown that the yeast analog of one such variant, Polδ-R696W, is an extremely mutagenic DNA polymerase that, notably, retains full exonuclease activity (30). Expression of the pol3-R696W allele was estimated to result in a more than 10,000-fold increase in the mutation rate, which is incompatible with life in both haploid and diploid cells and represents the strongest mutator effect described to date in eukaryotic cells. The corresponding human mutation, POLD1-R689W, was found in two colon cancer cell lines, DLD-1 (29) and HCT15 (31), which were independently derived from the same primary tumor (32), indicating that the R689W change existed in the tumor before the establishment of these cell lines.To understand how a single amino acid change in Polδ could cause such a phenomenal increase in mutagenesis, we first set out to characterize the fidelity of purified Polδ-R696W during in vitro DNA synthesis. Surprisingly, Polδ-R696W was not dramatically less accurate than previously studied Polδ variants that are only moderately mutagenic in vivo. We found, however, that yeast cells producing Polδ-R696W undergo S-phase checkpoint-dependent expansion of dNTP pools. Simulating this dNTP increase in vitro reduced the fidelity of Polδ-R696W to a level that explains its extraordinary mutator effect in vivo. We present evidence that the mutagenesis in pol3-R696W strains results from a combination of a Polδ nucleotide selectivity defect with activation of the signaling pathway that elevates dNTP pools and further increases the rate of Polδ errors. This vicious circle represents a new mechanism through which DNA polymerase alterations can promote the genome instability and which should be taken into consideration when trying to understand the development of hypermutated human cancers.     

Mechanism of staphylococcal multiresistance plasmid replication origin assembly by the RepA protein

The staphylococcal multiresistance plasmids are key contributors to the alarming rise in bacterial multidrug resistance. A conserved replication initiator, RepA, encoded on these plasmids is essential for their propagation. RepA proteins consist of flexibly linked N-terminal (NTD) and C-terminal (CTD) domains. Despite their essential role in replication, the molecular basis for RepA function is unknown. Here we describe a complete structural and functional dissection of RepA proteins. Unexpectedly, both the RepA NTD and CTD show similarity to the corresponding domains of the bacterial primosome protein, DnaD. Although the RepA and DnaD NTD both contain winged helix-turn-helices, the DnaD NTD self-assembles into large scaffolds whereas the tetrameric RepA NTD binds DNA iterons using a newly described DNA binding mode. Strikingly, structural and atomic force microscopy data reveal that the NTD tetramer mediates DNA bridging, suggesting a molecular mechanism for origin handcuffing. Finally, data show that the RepA CTD interacts with the host DnaG primase, which binds the replicative helicase. Thus, these combined data reveal the molecular mechanism by which RepA mediates the specific replicon assembly of staphylococcal multiresistant plasmids.The emergence of multidrug-resistant bacteria is a mounting global health crisis. In particular, multidrug-resistant Staphylococcus aureus is a major cause of nosocomial and community-acquired infections and is resistant to most antibiotics commonly used for patient treatment (1). Hospital intensive care units in many countries, including the United States, now report methicillin-resistant S. aureus infection rates exceeding 50% (2, 3). Antibiotic resistance in contemporary infectious S. aureus strains, such as in hospitals, is often encoded by plasmids that can be transmitted between strains via horizontal DNA transfer mechanisms. These plasmids are typically classified as small (<5 kb) multicopy plasmids, which usually encode only a single resistance gene; medium-sized (8–40 kb) multirestance plasmids that confer resistance to multiple antibiotics, disinfectants, and/or heavy metals; and large (>40 kb) conjugative multiresistance plasmids that additionally encode a conjugative DNA transfer mechanism (4–6). Importantly, sequence analyses have shown that most staphylococcal conjugative and nonconjugative multiresistance plasmids encode a highly conserved replication initiation protein, denoted RepA_N (5–15). RepA_N proteins are also encoded by plasmids from other Gram-positive bacteria as well as by some phage, underscoring their ubiquitous nature (10). These RepA proteins are essential for replication of multiresistance plasmids, and hence plasmid carriage and dissemination, yet the mechanisms by which these proteins function in replication are currently unknown.The DNA replication cycle can be divided into three stages: initiation, elongation, and termination. Replication initiation proteins (RepA) mediate the crucial first step of initiation. Bacterial chromosome replication is initiated by the chromosomal replication initiator protein, DnaA, which binds the origin and recruits the replication components known as the primosome (16). In Gram-negative bacteria the primosome includes DnaG primase, the replicative helicase (DnaB), and DnaC (17). Replication initiation in Gram-positive bacteria involves DnaG primase and helicase (DnaC) and the proteins DnaD, DnaI, and DnaB (18–22). DnaD binds first to DnaA at the origin. This is followed by binding of DnaI/DnaB and DnaG, which together recruit the replicative helicase (23, 24). Instead of DnaA, plasmids encode and use their own specific replication initiator binding protein. Structures are only available for RepA proteins (F, R6K, and pPS10 Rep) harbored in Gram-negative bacteria. These proteins contain winged helix-turn-helix (winged HTH) domains and bind iteron DNA as monomers to, in some still unclear manner, drive replicon assembly (25–27).Replication mechanisms used by plasmids harbored in Gram-positive bacteria are less well understood and are distinct from their Gram-negative counterparts. Indeed, most plasmid RepA proteins in Gram-negative and Gram-positive bacteria show no sequence homology and seem to be unrelated. The multiresistance RepA proteins are arguably among the most abundant of plasmid Rep proteins, yet how they function is not known. Data suggest that these proteins are composed of three main regions: an N-terminal domain (NTD) consisting of ∼120 aa, a long and variable linker region (∼30–50 residues), and a C-terminal domain (CTD) of ∼120 residues (28–31). The NTD and CTD are both essential for replication. The NTD exhibits the highest level of sequence conservation, which has resulted in the designation of plasmids that encode these proteins as the RepA_N replicon family (10). Although not as well conserved as the NTD, RepA CTD regions show homology between plasmids found in genus-specific clusters, suggesting that this domain may perform a host-specific role (28–32). Although the function of the RepA CTD remains enigmatic, recent studies have indicated that the NTD mediates DNA binding and interacts with iterons that reside within the plasmid origin (30). The essential roles played by RepA proteins in multiresistance plasmid retention marks them as attractive targets for the development of specific chemotherapeutics. However, the successful design of such compounds necessitates structural and mechanistic insight. Here, we describe a detailed dissection of the RepA proteins from the multiresistance plasmids pSK41 and pTZ2126. The combined data reveal the molecular underpinnings of a minimalist replication assembly mediated by multiresistance RepA proteins.     

Kinetics of nucleotide-dependent structural transitions in the kinesin-1 hydrolysis cycle

To dissect the kinetics of structural transitions underlying the stepping cycle of kinesin-1 at physiological ATP, we used interferometric scattering microscopy to track the position of gold nanoparticles attached to individual motor domains in processively stepping dimers. Labeled heads resided stably at positions 16.4 nm apart, corresponding to a microtubule-bound state, and at a previously unseen intermediate position, corresponding to a tethered state. The chemical transitions underlying these structural transitions were identified by varying nucleotide conditions and carrying out parallel stopped-flow kinetics assays. At saturating ATP, kinesin-1 spends half of each stepping cycle with one head bound, specifying a structural state for each of two rate-limiting transitions. Analysis of stepping kinetics in varying nucleotides shows that ATP binding is required to properly enter the one-head–bound state, and hydrolysis is necessary to exit it at a physiological rate. These transitions differ from the standard model in which ATP binding drives full docking of the flexible neck linker domain of the motor. Thus, this work defines a consensus sequence of mechanochemical transitions that can be used to understand functional diversity across the kinesin superfamily.Kinesin-1 is a motor protein that steps processively toward microtubule plus-ends, tracking single protofilaments and hydrolyzing one ATP molecule per step (1–6). Step sizes corresponding to the tubulin dimer spacing of 8.2 nm are observed when the molecule is labeled by its C-terminal tail (7–10) and to a two-dimer spacing of 16.4 nm when a single motor domain is labeled (4, 11, 12), consistent with the motor walking in a hand-over-hand fashion. Kinesin has served as an important model system for advancing single-molecule techniques (7–10) and is clinically relevant for its role in neurodegenerative diseases (13), making dissection of its step a popular ongoing target of study.Despite decades of work, many essential components of the mechanochemical cycle remain disputed, including (i) how much time kinesin-1 spends in a one-head–bound (1HB) state when stepping at physiological ATP concentrations, (ii) whether the motor waits for ATP in a 1HB or two-heads–bound (2HB) state, and (iii) whether ATP hydrolysis occurs before or after tethered head attachment (4, 11, 14–20). These questions are important because they are fundamental to the mechanism by which kinesins harness nucleotide-dependent structural changes to generate mechanical force in a manner optimized for their specific cellular tasks. Addressing these questions requires characterizing a transient 1HB state in the stepping cycle in which the unattached head is located between successive binding sites on the microtubule. This 1HB intermediate is associated with the force-generating powerstroke of the motor and underlies the detachment pathway that limits motor processivity. Optical trapping (7, 19, 21, 22) and single-molecule tracking studies (4, 8–11) have failed to detect this 1HB state during stepping. Single-molecule fluorescence approaches have detected a 1HB intermediate at limiting ATP concentrations (11, 12, 14, 15), but apart from one study that used autocorrelation analysis to detect a 3-ms intermediate (17), the 1HB state has been undetectable at physiological ATP concentrations.Single-molecule microscopy is a powerful tool for studying the kinetics of structural changes in macromolecules (23). Tracking steps and potential substeps for kinesin-1 at saturating ATP has until now been hampered by the high stepping rates of the motor (up to 100 s⁻¹), which necessitates high frame rates, and the small step size (8.2 nm), which necessitates high spatial precision (7). Here, we apply interferometric scattering microscopy (iSCAT), a recently established single-molecule tool with high spatiotemporal resolution (24–27) to directly visualize the structural changes underlying kinesin stepping. By labeling one motor domain in a dimeric motor, we detect a 1HB intermediate state in which the tethered head resides over the bound head for half the duration of the stepping cycle at saturating ATP. We further show that at physiological stepping rates, ATP binding is required to enter this 1HB state and that ATP hydrolysis is required to exit it. This work leads to a significant revision of the sequence and kinetics of mechanochemical transitions that make up the kinesin-1 stepping cycle and provides a framework for understanding functional diversity across the kinesin superfamily.     

Formation of interference-sensitive meiotic cross-overs requires sufficient DNA leading-strand elongation

Meiosis halves diploid genomes to haploid and is essential for sexual reproduction in eukaryotes. Meiotic recombination ensures physical association of homologs and their subsequent accurate segregation and results in the redistribution of genetic variations among progeny. Most organisms have two classes of cross-overs (COs): interference-sensitive (type I) and -insensitive (type II) COs. DNA synthesis is essential for meiotic recombination, but whether DNA synthesis has a role in differentiating meiotic CO pathways is unknown. Here, we show that Arabidopsis POL2A, the homolog of the yeast DNA polymerase-ε (a leading-strand DNA polymerase), is required for plant fertility and meiosis. Mutations in POL2A cause reduced fertility and meiotic defects, including abnormal chromosome association, improper chromosome segregation, and fragmentation. Observation of prophase I cell distribution suggests that pol2a mutants likely delay progression of meiotic recombination. In addition, the residual COs in pol2a have reduced CO interference, and the double mutant of pol2a with mus81, which affects type II COs, displayed more severe defects than either single mutant, indicating that POL2A functions in the type I pathway. We hypothesize that sufficient leading-strand DNA elongation promotes formation of some type I COs. Given that meiotic recombination and DNA synthesis are conserved in divergent eukaryotes, this study and our previous study suggest a novel role for DNA synthesis in the differentiation of meiotic recombination pathways.Eukaryotic sexual reproduction requires meiosis to produce haploid gametes from parents. Meiotic recombination not only ensures physical association of homologs to form bivalents but also, promotes their accurate segregation, and it is initiated by the formation of SPO11-programmed DNA double-strand breaks (DSBs) (1). Repair of meiotic DSBs is evolutionarily important, because it redistributes genetic variants in regions flanking the site of recombination [cross-overs (COs)] or gene conversion without flanking exchange [non-COs (NCOs)], resulting in offspring that are genetically distinct from parents and siblings (2, 3). Although DNA synthesis is thought to be essential for DSB repair (DSBR) (4), molecular genetic studies are quite limited. Recent genome-wide studies provide evidence that meiotic COs and NCOs may require different amounts of DNA synthesis (2, 3, 5–7), but the underlying mechanism remains unknown.Premeiotic chromosome replication requires the same DNA replication machinery as that for mitotic DNA replication (8), which is carried out by coordinated activities of DNA polymerase-α (Pol-α), -δ, and -ε (9) as well as three ancillary complexes: proliferating cell nuclear antigen, replication factor C (RFC), and the minichromosome maintenance complex. Pol-α and -δ are primarily required for lagging-strand DNA synthesis (9, 10), whereas Pol-ε carries out leading-strand DNA synthesis (11). Given that DNA synthesis is essential for S-phase replication during the mitotic cell cycle, complete disruption of these genes causes lethality, which was observed in yeast, plants, and humans (11–14), making it difficult to study their functions in meiosis. In Saccharomyces cerevisiae, a partially defective mutation of Pol-δ causes no DNA replication defect but exhibits reduction in both the number of COs and the length of meiotic gene conversion tracts (15). In Arabidopsis, a partial loss of function of DNA replication factor C1 (RFC1) results in reduction of type I COs (16). These findings suggest that DNA synthesis likely affects the length of strand exchange intermediates and influences their resolution toward different CO pathways. Moreover, genome-wide analyses in both yeast and plants showed that meiotic gene conversion tracts vary in length, ranging from several hundreds to thousands of nucleotides, and that CO-associated tracts are usually longer than those of NCOs (2, 3, 5–7). This difference raises a question of whether there is any difference in DNA synthesis among CO-associated tracts with different lengths; however, this hypothesis has not been tested experimentally. Our previous research with RFC1 supports the idea that lagging-strand synthesis promotes type I COs (16). Here, we test the hypothesis that sufficient leading-strand synthesis is important for type I COs.To investigate the role of leading-strand DNA elongation in meiotic recombination, we analyzed two partial loss-of-function alleles of the largest subunit of DNA Pol-ε (hereafter designated as POL2A). They have normal vegetative development but show defects in fertility and meiosis. Additional analyses show that POL2A has a role in the formation of normal bivalents and promoting formation of meiotic type I COs. Similar mutations in the yeast homolog showed reduced DNA replication efficiency in mitotic division (17–19), leading us to hypothesize that sufficient leading-strand DNA elongation could be a critical step for differentiation of types I and II COs. These observations provide a novel insight that meiotic types I and II COs are differentially dependent on leading-strand synthesis activity.