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).     

Structure of the Pseudomonas aeruginosa transamidosome reveals unique aspects of bacterial tRNA-dependent asparagine biosynthesis

Many prokaryotes lack a tRNA synthetase to attach asparagine to its cognate tRNA^Asn, and instead synthesize asparagine from tRNA^Asn-bound aspartate. This conversion involves two enzymes: a nondiscriminating aspartyl-tRNA synthetase (ND-AspRS) that forms Asp-tRNA^Asn, and a heterotrimeric amidotransferase GatCAB that amidates Asp-tRNA^Asn to form Asn-tRNA^Asn for use in protein synthesis. ND-AspRS, GatCAB, and tRNA^Asn may assemble in an ∼400-kDa complex, known as the Asn-transamidosome, which couples the two steps of asparagine biosynthesis in space and time to yield Asn-tRNA^Asn. We report the 3.7-Å resolution crystal structure of the Pseudomonas aeruginosa Asn-transamidosome, which represents the most common machinery for asparagine biosynthesis in bacteria. We show that, in contrast to a previously described archaeal-type transamidosome, a bacteria-specific GAD domain of ND-AspRS provokes a principally new architecture of the complex. Both tRNA^Asn molecules in the transamidosome simultaneously serve as substrates and scaffolds for the complex assembly. This architecture rationalizes an elevated dynamic and a greater turnover of ND-AspRS within bacterial-type transamidosomes, and possibly may explain a different evolutionary pathway of GatCAB in organisms with bacterial-type vs. archaeal-type Asn-transamidosomes. Importantly, because the two-step pathway for Asn-tRNA^Asn formation evolutionarily preceded the direct attachment of Asn to tRNA^Asn, our structure also may reflect the mechanism by which asparagine was initially added to the genetic code.Accurate translation of the genetic code into a protein sequence relies on a covalent attachment of amino acids to cognate tRNAs that are later used in protein synthesis (1). This attachment is catalyzed by aminoacyl-tRNA synthetases (aaRSs), each specific to one amino acid and a set of tRNA isoacceptors (2). However, the majority of prokaryotes lack several tRNA synthetases, particularly asparaginyl-tRNA synthetase (AsnRS), which ligates asparagine to tRNA^Asn (3, 4). In these organisms, asparagine is synthesized in a two-step, tRNA-dependent pathway (5). First, a nondiscriminating aspartyl-tRNA synthetase (ND-AspRS) attaches aspartate to tRNA^Asn to form Asp-tRNA^Asn (6, 7). Then the tRNA-bound aspartate is converted to asparagine by the amidotransferase (AdT) GatCAB to yield the final product, Asn-tRNA^Asn (6, 8–13). Likewise, in prokaryotes lacking glutaminyl-tRNA synthetase (GlnRS), Gln-tRNA^Gln is formed by the sequential actions of a nondiscriminating glutamyl-tRNA synthetase (ND-GluRS) (14) and an AdT (5). In bacteria, the role of AdT is played by GatCAB (15), whereas in archaea, it is played by GatDE (16, 17).More than 25 years ago (18), it was proposed that ND-aaRSs and AdTs may form a complex—now called a transamidosome—to couple the two steps of Asn-tRNA^Asn formation in space and time and allow efficient transfer of Asp-tRNA^Asn from the aaRS to the AdT. The first characterized transamidosome was the Asn-transamidosome from Thermus thermophilus (TtAsn-transamidosome) (19, 20). This complex was identified as a tRNA-dependent association of AspRS2 (TtAspRS2) and GatCAB in a 2:2:2 ratio. It was shown that transamidosome formation stabilizes interactions between subunits of GatCAB (21) and protects Asn-tRNA^Asn from hydrolysis, with product release being rate-limiting (19). In the complex, the AspRS forms a dimer with only one catalytic site active at a time (21). It was suggested that the key advantages of asparagine formation by the transamidosome compared with separate enzymes are enhanced aspartylation of tRNA^Asn and better prevention of the misacylated Asp-tRNA^Asn from use in translation, because this would compromise the fidelity of protein synthesis (19–21).Importantly, TtAspRS2 was acquired through horizontal gene transfer from archaea (10), and lacks the GAD domain typical of bacterial AspRSs (22). The TtAsn-transamidosome crystal structure suggests that a complex between bacterial ND-AspRS and GatCAB should be less stable and more structurally distinct than the T. thermophilus complex, owing to the presence of the GAD domain in bacterial ND-AspRS (21, 23). Consistent with this notion, the stable association of the Helicobacter pylori ND-AspRS (HpND-AspRS) with GatCAB requires the presence of an auxiliary factor, Hp0100 (23, 24). In the complex, the activity of the HpND-AspRS is unchanged, although the activity of GatCAB increases (23); however, Hp0100 is phylogenetically limited to ε-proteobacteria (23). Therefore, most bacteria have a structurally and, possibly, functionally distinct class of transamidosomes than those described by the T. thermophilus and H. pylori complexes.In the bacterium Pseudomonas aeruginosa, Asn-tRNA^Asn formation is catalyzed by GatCAB and bacterial ND-AspRS, and thus represents the most common type of bacterial Asn-transamidosome (25). Here we report the crystal structure of the P. aeruginosa Asn-transamidosome (PaAsn-transamidosome), which represents the transamidation state of the Asn-tRNA^Asn formation. The structure suggests that the additional GAD domain within the ND-AspRS changes the overall architecture of the complex relative to the previously described TtAsn-transamidosome. Consistent with the structure, our in vitro measurements show that PaAsn-transamidosome has unique kinetic properties and functions primarily to enhance tRNA^Asn turnover and facilitate Asp-tRNA^Asn handoff from AspRS to GatCAB.     

Self-assembly of alkynylplatinum(II) terpyridine amphiphiles into nanostructures via steric control and metal–metal interactions

A series of mono- and dinuclear alkynylplatinum(II) terpyridine complexes containing the hydrophilic oligo(para-phenylene ethynylene) with two 3,6,9-trioxadec-1-yloxy chains was designed and synthesized. The mononuclear alkynylplatinum(II) terpyridine complex was found to display a very strong tendency toward the formation of supramolecular structures. Interestingly, additional end-capping with another platinum(II) terpyridine moiety of various steric bulk at the terminal alkyne would lead to the formation of nanotubes or helical ribbons. These desirable nanostructures were found to be governed by the steric bulk on the platinum(II) terpyridine moieties, which modulates the directional metal−metal interactions and controls the formation of nanotubes or helical ribbons. Detailed analysis of temperature-dependent UV-visible absorption spectra of the nanostructured tubular aggregates also provided insights into the assembly mechanism and showed the role of metal−metal interactions in the cooperative supramolecular polymerization of the amphiphilic platinum(II) complexes.Square-planar d⁸ platinum(II) polypyridine complexes have long been known to exhibit intriguing spectroscopic and luminescence properties (1–54) as well as interesting solid-state polymorphism associated with metal−metal and π−π stacking interactions (1–14, 25). Earlier work by our group showed the first example, to our knowledge, of an alkynylplatinum(II) terpyridine system [Pt(tpy)(C ≡ CR)]⁺ that incorporates σ-donating and solubilizing alkynyl ligands together with the formation of Pt···Pt interactions to exhibit notable color changes and luminescence enhancements on solvent composition change (25) and polyelectrolyte addition (26). This approach has provided access to the alkynylplatinum(II) terpyridine and other related cyclometalated platinum(II) complexes, with functionalities that can self-assemble into metallogels (27–31), liquid crystals (32, 33), and other different molecular architectures, such as hairpin conformation (34), helices (35–38), nanostructures (39–45), and molecular tweezers (46, 47), as well as having a wide range of applications in molecular recognition (48–52), biomolecular labeling (48–52), and materials science (53, 54). Recently, metal-containing amphiphiles have also emerged as a building block for supramolecular architectures (42–44, 55–59). Their self-assembly has always been found to yield different molecular architectures with unprecedented complexity through the multiple noncovalent interactions on the introduction of external stimuli (42–44, 55–59).Helical architecture is one of the most exciting self-assembled morphologies because of the uniqueness for the functional and topological properties (60–69). Helical ribbons composed of amphiphiles, such as diacetylenic lipids, glutamates, and peptide-based amphiphiles, are often precursors for the growth of tubular structures on an increase in the width or the merging of the edges of ribbons (64, 65). Recently, the optimization of nanotube formation vs. helical nanostructures has aroused considerable interests and can be achieved through a fine interplay of the influence on the amphiphilic property of molecules (66), choice of counteranions (67, 68), or pH values of the media (69), which would govern the self-assembly of molecules into desirable aggregates of helical ribbons or nanotube scaffolds. However, a precise control of supramolecular morphology between helical ribbons and nanotubes remains challenging, particularly for the polycyclic aromatics in the field of molecular assembly (64–69). Oligo(para-phenylene ethynylene)s (OPEs) with solely π−π stacking interactions are well-recognized to self-assemble into supramolecular system of various nanostructures but rarely result in the formation of tubular scaffolds (70–73). In view of the rich photophysical properties of square-planar d⁸ platinum(II) systems and their propensity toward formation of directional Pt···Pt interactions in distinctive morphologies (27–31, 39–45), it is anticipated that such directional and noncovalent metal−metal interactions might be capable of directing or dictating molecular ordering and alignment to give desirable nanostructures of helical ribbons or nanotubes in a precise and controllable manner.Herein, we report the design and synthesis of mono- and dinuclear alkynylplatinum(II) terpyridine complexes containing hydrophilic OPEs with two 3,6,9-trioxadec-1-yloxy chains. The mononuclear alkynylplatinum(II) terpyridine complex with amphiphilic property is found to show a strong tendency toward the formation of supramolecular structures on diffusion of diethyl ether in dichloromethane or dimethyl sulfoxide (DMSO) solution. Interestingly, additional end-capping with another platinum(II) terpyridine moiety of various steric bulk at the terminal alkyne would result in nanotubes or helical ribbons in the self-assembly process. To the best of our knowledge, this finding represents the first example of the utilization of the steric bulk of the moieties, which modulates the formation of directional metal−metal interactions to precisely control the formation of nanotubes or helical ribbons in the self-assembly process. Application of the nucleation–elongation model into this assembly process by UV-visible (UV-vis) absorption spectroscopic studies has elucidated the nature of the molecular self-assembly, and more importantly, it has revealed the role of metal−metal interactions in the formation of these two types of nanostructures.     

Exchange protein directly activated by cAMP plays a critical role in bacterial invasion during fatal rickettsioses

Rickettsiae are responsible for some of the most devastating human infections. A high infectivity and severe illness after inhalation make some rickettsiae bioterrorism threats. We report that deletion of the exchange protein directly activated by cAMP (Epac) gene, Epac1, in mice protects them from an ordinarily lethal dose of rickettsiae. Inhibition of Epac1 suppresses bacterial adhesion and invasion. Most importantly, pharmacological inhibition of Epac1 in vivo using an Epac-specific small-molecule inhibitor, ESI-09, completely recapitulates the Epac1 knockout phenotype. ESI-09 treatment dramatically decreases the morbidity and mortality associated with fatal spotted fever rickettsiosis. Our results demonstrate that Epac1-mediated signaling represents a mechanism for host–pathogen interactions and that Epac1 is a potential target for the prevention and treatment of fatal rickettsioses.Rickettsiae are responsible for some of the most devastating human infections (1–4). It has been forecasted that temperature increases attributable to global climate change will lead to more widespread distribution of rickettsioses (5). These tick-borne diseases are caused by obligately intracellular bacteria of the genus Rickettsia, including Rickettsia rickettsii, the causative agent of Rocky Mountain spotted fever (RMSF) in the United States and Latin America (2, 3), and Rickettsia conorii, the causative agent of Mediterranean spotted fever endemic to southern Europe, North Africa, and India (6). A high infectivity and severe illness after inhalation make some rickettsiae (including Rickettsia prowazekii, R. rickettsii, Rickettsia typhi, and R. conorii) bioterrorism threats (7). Although the majority of rickettsial infections can be controlled by appropriate broad-spectrum antibiotic therapy if diagnosed early, up to 20% of misdiagnosed or untreated (1, 3) and 5% of treated RMSF cases (8) result in a fatal outcome caused by acute disseminated vascular endothelial infection and damage (9). Fatality rates as high as 32% have been reported in hospitalized patients diagnosed with Mediterranean spotted fever (10). In addition, strains of R. prowazekii resistant to tetracycline and chloramphenicol have been developed in laboratories (11). Disseminated endothelial infection and endothelial barrier disruption with increased microvascular permeability are the central features of SFG rickettsioses (1, 2, 9). The molecular mechanisms involved in rickettsial infection remain incompletely elucidated (9, 12). A comprehensive understanding of rickettsial pathogenesis and the development of novel mechanism-based treatment are urgently needed.Living organisms use intricate signaling networks for sensing and responding to changes in the external environment. cAMP, a ubiquitous second messenger, is an important molecular switch that translates environmental signals into regulatory effects in cells (13). As such, a number of microbial pathogens have evolved a set of diverse virulence-enhancing strategies that exploit the cAMP-signaling pathways of their hosts (14). The intracellular functions of cAMP are predominantly mediated by the classic cAMP receptor, protein kinase A (PKA), and the more recently discovered exchange protein directly activated by cAMP (Epac) (15). Thus, far, two isoforms, Epac1 and Epac2, have been identified in humans (16, 17). Epac proteins function by responding to increased intracellular cAMP levels and activating the Ras superfamily small GTPases Ras-proximate 1 and 2 (Rap1 and Rap2). Accumulating evidence demonstrates that the cAMP/Epac1 signaling axis plays key regulatory roles in controlling various cellular functions in endothelial cells in vitro, including cell adhesion (18–21), exocytosis (22), tissue plasminogen activator expression (23), suppressor of cytokine signaling 3 (SOCS-3) induction (24–27), microtubule dynamics (28, 29), cell–cell junctions, and permeability and barrier functions (30–37). Considering the critical importance of endothelial cells in rickettsioses, we examined the functional roles of Epac1 in rickettsial pathogenesis in vivo, taking advantage of the recently generated Epac1 knockout mouse (38) and Epac-specific inhibitors (39, 40) generated from our laboratory. Our studies demonstrate that Epac1 plays a key role in rickettsial infection and represents a therapeutic target for fatal rickettsioses.     

N terminus of ASPP2 binds to Ras and enhances Ras/Raf/MEK/ERK activation to promote oncogene-induced senescence

The ASPP2 (also known as 53BP2L) tumor suppressor is a proapoptotic member of a family of p53 binding proteins that functions in part by enhancing p53-dependent apoptosis via its C-terminal p53-binding domain. Mounting evidence also suggests that ASPP2 harbors important nonapoptotic p53-independent functions. Structural studies identify a small G protein Ras-association domain in the ASPP2 N terminus. Because Ras-induced senescence is a barrier to tumor formation in normal cells, we investigated whether ASPP2 could bind Ras and stimulate the protein kinase Raf/MEK/ERK signaling cascade. We now show that ASPP2 binds to Ras–GTP at the plasma membrane and stimulates Ras-induced signaling and pERK1/2 levels via promoting Ras–GTP loading, B-Raf/C-Raf dimerization, and C-Raf phosphorylation. These functions require the ASPP2 N terminus because BBP (also known as 53BP2S), an alternatively spliced ASPP2 isoform lacking the N terminus, was defective in binding Ras–GTP and stimulating Raf/MEK/ERK signaling. Decreased ASPP2 levels attenuated H-RasV12–induced senescence in normal human fibroblasts and neonatal human epidermal keratinocytes. Together, our results reveal a mechanism for ASPP2 tumor suppressor function via direct interaction with Ras–GTP to stimulate Ras-induced senescence in nontransformed human cells.ASPP2, also known as 53BP2L, is a tumor suppressor whose expression is altered in human cancers (1). Importantly, targeting of the ASPP2 allele in two different mouse models reveals that ASPP2 heterozygous mice are prone to spontaneous and γ-irradiation–induced tumors, which rigorously demonstrates the role of ASPP2 as a tumor suppressor (2, 3). ASPP2 binds p53 via the C-terminal ankyrin-repeat and SH3 domain (4–6), is damage-inducible, and can enhance damage-induced apoptosis in part through a p53-mediated pathway (1, 2, 7–10). However, it remains unclear what biologic pathways and mechanisms mediate ASPP2 tumor suppressor function (1). Indeed, accumulating evidence demonstrates that ASPP2 also mediates nonapoptotic p53-independent pathways (1, 3, 11–15).The induction of cellular senescence forms an important barrier to tumorigenesis in vivo (16–21). It is well known that oncogenic Ras signaling induces senescence in normal nontransformed cells to prevent tumor initiation and maintain complex growth arrest pathways (16, 18, 21–24). The level of oncogenic Ras activation influences its capacity to activate senescence; high levels of oncogenic H-RasV12 signaling leads to low grade tumors with senescence markers, which progress to invasive cancers upon senescence inactivation (25). Thus, tight control of Ras signaling is critical to ensure the proper biologic outcome in the correct cellular context (26–28).The ASPP2 C terminus is important for promoting p53-dependent apoptosis (7). The ASPP2 N terminus may also suppress cell growth (1, 7, 29–33). Alternative splicing can generate the ASPP2 N-terminal truncated protein BBP (also known as 53BP2S) that is less potent in suppressing cell growth (7, 34, 35). Although the ASPP2 C terminus mediates nuclear localization, full-length ASPP2 also localizes to the cytoplasm and plasma membrane to mediate extranuclear functions (7, 11, 12, 36). Structural studies of the ASPP2 N terminus reveal a β–Grasp ubiquitin-like fold as well as a potential Ras-binding (RB)/Ras-association (RA) domain (32). Moreover, ASPP2 can promote H-RasV12–induced senescence (13, 15). However, the molecular mechanism(s) of how ASPP2 directly promotes Ras signaling are complex and remain to be completely elucidated.Here, we explore the molecular mechanisms of how Ras-signaling is enhanced by ASPP2. We demonstrate that ASPP2: (i) binds Ras-GTP and stimulates Ras-induced ERK signaling via its N-terminal domain at the plasma membrane; (ii) enhances Ras-GTP loading and B-Raf/C-Raf dimerization and forms a ASPP2/Raf complex; (iii) stimulates Ras-induced C-Raf phosphorylation and activation; and (iv) potentiates H-RasV12–induced senescence in both primary human fibroblasts and neonatal human epidermal keratinocytes. These data provide mechanistic insight into ASPP2 function(s) and opens important avenues for investigation into its role as a tumor suppressor in human cancer.     

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.