Rtt105 promotes high-fidelity DNA replication and repair by regulating the single-stranded DNA-binding factor RPA

Single-stranded DNA (ssDNA) covered with the heterotrimeric Replication Protein A (RPA) complex is a central intermediate of DNA replication and repair. How RPA is regulated to ensure the fidelity of DNA replication and repair remains poorly understood. Yeast Rtt105 is an RPA-interacting protein required for RPA nuclear import and efficient ssDNA binding. Here, we describe an important role of Rtt105 in high-fidelity DNA replication and recombination and demonstrate that these functions of Rtt105 primarily depend on its regulation of RPA. The deletion of RTT105 causes elevated spontaneous DNA mutations with large duplications or deletions mediated by microhomologies. Rtt105 is recruited to DNA double-stranded break (DSB) ends where it promotes RPA assembly and homologous recombination repair by gene conversion or break-induced replication. In contrast, Rtt105 attenuates DSB repair by the mutagenic single-strand annealing or alternative end joining pathway. Thus, Rtt105-mediated regulation of RPA promotes high-fidelity replication and recombination while suppressing repair by deleterious pathways. Finally, we show that the human RPA-interacting protein hRIP-α, a putative functional homolog of Rtt105, also stimulates RPA assembly on ssDNA, suggesting the conservation of an Rtt105-mediated mechanism.

Faithful DNA replication and repair are essential for the maintenance of genetic material (1). Even minor defects in replication or repair can cause high loads of mutations, genome instability, cancer, and other diseases (1). Deficiency in different DNA repair or replication proteins can lead to distinct mutation patterns (2–4). For example, deficiency in mismatch repair results in increased microsatellite instability, while deficiency in homologous recombination repair is often associated with tandem duplications or deletions (3–7). Sequence analysis of various cancer types has identified many distinct genome rearrangement and mutation signatures (8). However, the genetic basis for some of these signatures remains poorly understood, thus requiring further investigation in experimental models (8).In eukaryotic cells, Replication Protein A (RPA), the major single-stranded DNA (ssDNA) binding protein complex, is essential for DNA replication, repair, and recombination (9–13). It is also crucial for the suppression of mutations and genome instability (14–17). RPA acts as a key scaffold to recruit and coordinate proteins involved in different DNA metabolic processes (14, 15, 17). As the first responder of ssDNA, RPA participates in both replication initiation and elongation (10, 12, 13). During replication or under replication stresses, the exposed ssDNA must be protected and stabilized by RPA to prevent formation of secondary structures (14, 16). RPA is also essential for DNA double-stranded break (DSB) repair by the homologous recombination (HR) pathway (18–21). During HR, the 5′-terminated strands of DSBs are initially processed by the resection machinery, generating 3′-tailed ssDNA (22). The 3′-ssDNA becomes bound by the RPA complex to activate the DNA damage checkpoint (23). RPA is subsequently replaced by the Rad51 recombinase to form a Rad51 nucleoprotein filament (19, 24). This recombinase filament catalyzes invasion of the 3′-strands at the homologous sequence to form the D-loop structure, followed by repair DNA synthesis and resolution of recombination intermediates (18, 19, 24). During HR, RPA prevents the formation of DNA secondary structures and protects 3′-ssDNA from nucleolytic degradation (25). In addition, recent work implies a role of RPA in homology recognition (26).RPA is composed of three subunits, Rfa1, Rfa2, and Rfa3, and with a total of six oligonucleotide-binding (OB) motifs that mediate interactions with ssDNA or proteins (14, 17, 27). RPA can associate with ssDNA in different modes (28). It binds short DNA (8 to 10 nt) in an unstable mode and longer ssDNA (28 to 30 nt) in a high-affinity mode (28–31). Recent single-molecule studies revealed that RPA binding on ssDNA is highly dynamic (28, 32). It can rapidly diffuse within the bound DNA ligand and quickly exchange between the free and ssDNA-bound states (32–35). The cellular functions of RPA rely on its high ssDNA-binding affinity and its ability to interact with different proteins (28). Although RPA has a high affinity for ssDNA, recent studies have suggested that the binding of RPA on chromatin requires additional regulations (36). How RPA is regulated to ensure replication and repair fidelity remains poorly understood.Rtt105, a protein initially identified as a regulator of the Ty1 retrotransposon, has recently been shown to interact with RPA and acts as an RPA chaperone (36). It facilitates the nuclear localization of RPA and stimulates the loading of RPA at replication forks in unperturbed conditions or under replication stresses (36). Rtt105 exhibits synthetic genetic interactions with genes encoding replisome proteins and is required for heterochromatin silencing and telomere maintenance (37). The deletion of RTT105 results in increased gross chromosomal rearrangements and reduced resistance to DNA-damaging agents (36, 38). In vitro, Rtt105 can directly stimulate RPA binding to ssDNA, likely by changing the binding mode of RPA (36).In this study, by using a combination of genetic, biochemical, and single-molecule approaches, we demonstrate that Rtt105-dependent regulation of RPA promotes high-fidelity genome duplication and recombination while suppressing mutations and the low-fidelity repair pathways. We provide evidence that human hRIP-α, the putative functional homolog of yeast Rtt105, could regulate human RPA assembly on ssDNA in vitro. Our study unveils a layer of regulation on the maintenance of genome integrity that relies on dynamic RPA binding on ssDNA to ensure high-fidelity replication or recombination.     

SLFN11 promotes CDT1 degradation by CUL4 in response to replicative DNA damage,while its absence leads to synthetic lethality with ATR/CHK1 inhibitors

Schlafen-11 (SLFN11) inactivation in ∼50% of cancer cells confers broad chemoresistance. To identify therapeutic targets and underlying molecular mechanisms for overcoming chemoresistance, we performed an unbiased genome-wide RNAi screen in SLFN11-WT and -knockout (KO) cells. We found that inactivation of Ataxia Telangiectasia- and Rad3-related (ATR), CHK1, BRCA2, and RPA1 overcome chemoresistance to camptothecin (CPT) in SLFN11-KO cells. Accordingly, we validate that clinical inhibitors of ATR (M4344 and M6620) and CHK1 (SRA737) resensitize SLFN11-KO cells to topotecan, indotecan, etoposide, cisplatin, and talazoparib. We uncover that ATR inhibition significantly increases mitotic defects along with increased CDT1 phosphorylation, which destabilizes kinetochore-microtubule attachments in SLFN11-KO cells. We also reveal a chemoresistance mechanism by which CDT1 degradation is retarded, eventually inducing replication reactivation under DNA damage in SLFN11-KO cells. In contrast, in SLFN11-expressing cells, SLFN11 promotes the degradation of CDT1 in response to CPT by binding to DDB1 of CUL4^CDT2 E3 ubiquitin ligase associated with replication forks. We show that the C terminus and ATPase domain of SLFN11 are required for DDB1 binding and CDT1 degradation. Furthermore, we identify a therapy-relevant ATPase mutant (E669K) of the SLFN11 gene in human TCGA and show that the mutant contributes to chemoresistance and retarded CDT1 degradation. Taken together, our study reveals new chemotherapeutic insights on how targeting the ATR pathway overcomes chemoresistance of SLFN11-deficient cancers. It also demonstrates that SLFN11 irreversibly arrests replication by degrading CDT1 through the DDB1–CUL4^CDT2 ubiquitin ligase.

Schlafen-11 (SLFN11) is an emergent restriction factor against genomic instability acting by eliminating cells with replicative damage (1–6) and potentially acting as a tumor suppressor (6, 7). SLFN11-expressing cancer cells are consistently hypersensitive to a broad range of chemotherapeutic drugs targeting DNA replication, including topoisomerase inhibitors, alkylating agents, DNA synthesis, and poly(ADP-ribose) polymerase (PARP) inhibitors compared to SLFN11-deficient cancer cells, which are chemoresistant (1, 2, 4, 8–17). Profiling SLFN11 expression is being explored for patients to predict survival and guide therapeutic choice (8, 13, 18–24).The Cancer Genome Atlas (TCGA) and cancer cell databases demonstrate that SLFN11 mRNA expression is suppressed in a broad fraction of common cancer tissues and in ∼50% of all established cancer cell lines across multiple histologies (1, 2, 5, 8, 13, 25, 26). Silencing of the SLFN11 gene, like known tumor suppressor genes, is under epigenetic mechanisms through hypermethylation of its promoter region and activation of histone deacetylases (HDACs) (21, 23, 25, 26). A recent study in small-cell lung cancer patient-derived xenograft models also showed that SLFN11 gene silencing is caused by local chromatin condensation related to deposition of H3K27me3 in the gene body of SLFN11 by EZH2, a histone methyltransferase (11). Targeting epigenetic regulators is therefore an attractive combination strategy to overcome chemoresistance of SLFN11-deficient cancers (10, 25, 26). An alternative approach is to attack SLFN11-negative cancer cells by targeting the essential pathways that cells use to overcome replicative damage and replication stress. Along these lines, a prior study showed that inhibition of ATR (Ataxia Telangiectasia- and Rad3-related) kinase reverses the resistance of SLFN11-deficient cancer cells to PARP inhibitors (4). However, targeting the ATR pathway in SLFN11-deficient cells has not yet been fully explored.SLFN11 consists of two functional domains: A conserved nuclease motif in its N terminus and an ATPase motif (putative helicase) in its C terminus (2, 6). The N terminus nuclease has been implicated in the selective degradation of type II tRNAs (including those coding for ATR) and its nuclease structure can be derived from crystallographic analysis of SLFN13 whose N terminus domain is conserved with SLFN11 (27, 28). The C terminus is only present in the group III Schlafen family (24, 29). Its potential ATPase activity and relationship to chemosensitivity to DNA-damaging agents (3–5) imply that the ATPase/helicase of SLFN11 is involved specifically in DNA damage response (DDR) to replication stress. Indeed, inactivation of the Walker B motif of SLFN11 by the mutation E669Q suppresses SLFN11-mediated replication block (5, 30). In addition, SLFN11 contains a binding site for the single-stranded DNA binding protein RPA1 (replication protein A1) at its C terminus (3, 31) and is recruited to replication damage sites by RPA (3, 5). The putative ATPase activity of SLFN11 is not required for this recruitment (5) but is required for blocking the replication helicase complex (CMG-CDC45) and inducing chromatin accessibility at replication origins and promoter sites (5, 30). Based on these studies, our current model is that SLFN11 is recruited to “stressed” replication forks by RPA filaments formed on single-stranded DNA (ssDNA), and that the ATPase/helicase activity of SLFN11 is required for blocking replication progression and remodeling chromatin (5, 30). However, underlying mechanisms of how SLFN11 irreversibly blocks replication in DNA damage are still unclear.Increased RPA-coated ssDNA caused by DNA damage and replication fork stalling also triggers ATR kinase activation, promoting subsequent phosphorylation of CHK1, which transiently halts cell cycle progression and enables DNA repair (32). ATR inhibitors are currently in clinical development in combination with DNA replication damaging drugs (33, 34), such as topoisomerase I (TOP1) inhibitors, which are highly synergistic with ATR inhibitors in preclinical models (35). ATR inhibitors not only inhibit DNA repair, but also lead to unscheduled replication origin firing (36), which kills cancer cells (37, 38) by inducing genomic alterations due to faulty replication and mitotic catastrophe (33).The replication licensing factor CDT1 orchestrates the initiation of replication by assembling prereplication complexes (pre-RC) in G1-phase before cells enter S-phase (39). Once replication is started by loading and activation of the MCM helicase, CDT1 is degraded by the ubiquitin proteasomal pathway to prevent additional replication initiation and ensure precise genome duplication and the firing of each origin only once per cell cycle (39, 40). At the end of G2 and during mitosis, CDT1 levels rise again to control kinetochore-microtubule attachment for accurate chromosome segregation (41). Deregulated overexpression of CDT1 results in rereplication, genome instability, and tumorigenesis (42). The cellular CDT1 levels are tightly regulated by the damage-specific DNA binding protein 1 (DDB1)–CUL4^CDT2 E3 ubiquitin ligase complex in G1-phase (43) and in response to DNA damage (44, 45). How CDT1 is recognized by CUL4^CDT2 in response to DNA damage remains incompletely known.In the present study, starting with a human genome-wide RNAi screen, bioinformatics analyses, and mechanistic validations, we explored synthetic lethal interactions that overcome the chemoresistance of SLFN11-deficient cells to the TOP1 inhibitor camptothecin (CPT). The strongest synergistic interaction was between depletion of the ATR/CHK1-mediated DNA damage response pathways and DNA-damaging agents in SLFN11-deficient cells. We validated and expanded our molecular understanding of combinatorial strategies in SLFN11-deficient cells with the ATR (M4344 and M6620) and CHK1 (SRA737) inhibitors in clinical development (33, 46, 47) and found that ATR inhibition leads to CDT1 stabilization and hyperphosphorylation with mitotic catastrophe. Our study also establishes that SLFN11 promotes the degradation of CDT1 by binding to DDB1, an adaptor molecule of the CUL4^CDT2 E3 ubiquitin ligase complex, leading to an irreversible replication block in response to replicative DNA damage.     

The circadian clock ensures successful DNA replication in cyanobacteria

Disruption of circadian rhythms causes decreased health and fitness, and evidence from multiple organisms links clock disruption to dysregulation of the cell cycle. However, the function of circadian regulation for the essential process of DNA replication remains elusive. Here, we demonstrate that in the cyanobacterium Synechococcus elongatus, a model organism with the simplest known circadian oscillator, the clock generates rhythms in DNA replication to minimize the number of open replication forks near dusk that would have to complete after sunset. Metabolic rhythms generated by the clock ensure that resources are available early at night to support any remaining replication forks. Combining mathematical modeling and experiments, we show that metabolic defects caused by clock–environment misalignment result in premature replisome disassembly and replicative abortion in the dark, leaving cells with incomplete chromosomes that persist through the night. Our study thus demonstrates that a major function of this ancient clock in cyanobacteria is to ensure successful completion of genome replication in a cycling environment.

Circadian clocks, internally generated rhythms in physiology with a ∼24 h period, are found in all domains of life. These clocks allow organisms to coordinate their physiological activities in anticipation of the daily cycle in the external environmental (1–3). Disruption of clocks caused either by mutation or clock–environment mismatch leads to decreased health and reproductive fitness in multiple organisms (4–6). In mammals, risk for age-related diseases such as cancer and cardiometabolic dysfunction is enhanced by circadian disruption (7, 8).Although much is now understood about the molecular mechanisms that generate rhythms, the origin of these health defects is still incompletely understood. A common target of circadian control shared across many species is the progression of the cell cycle (9–12). In animals, disrupted circadian rhythms are often linked to aberrant cell proliferation and tumorigenesis (13). Successful duplication of the genome is essential for the production of viable progeny. Replicating a bacterial genome can take up to several hours, a timescale over which external illumination from sunlight can change substantially. We therefore speculated that initiation of DNA replication could be a key point of circadian control. The cyanobacterium Synechococcus elongatus, which has the simplest known circadian system, is a powerful model system to address these issues, both because its clock is intimately coupled to cell cycle (9, 14–16) and because clock–environment misalignment has profound effects on reproductive fitness (17). Here, we analyze whether replication is clock-regulated in S. elongatus and the consequences of clock–environment mismatch on DNA replication.     

The retromer is co-opted to deliver lipid enzymes for the biogenesis of lipid-enriched tombusviral replication organelles

Biogenesis of viral replication organelles (VROs) is critical for replication of positive-strand RNA viruses. In this work, we demonstrate that tomato bushy stunt virus (TBSV) and the closely related carnation Italian ringspot virus (CIRV) hijack the retromer to facilitate building VROs in the surrogate host yeast and in plants. Depletion of retromer proteins, which are needed for biogenesis of endosomal tubular transport carriers, strongly inhibits the peroxisome-associated TBSV and the mitochondria-associated CIRV replication in yeast and in planta. In vitro reconstitution revealed the need for the retromer for the full activity of the viral replicase. The viral p33 replication protein interacts with the retromer complex, including Vps26, Vps29, and Vps35. We demonstrate that TBSV p33-driven retargeting of the retromer into VROs results in delivery of critical retromer cargoes, such as 1) Psd2 phosphatidylserine decarboxylase, 2) Vps34 phosphatidylinositol 3-kinase (PI3K), and 3) phosphatidylinositol 4-kinase (PI4Kα-like). The recruitment of these cellular enzymes by the co-opted retromer is critical for de novo production and enrichment of phosphatidylethanolamine phospholipid, phosphatidylinositol-3-phosphate [PI(3)P], and phosphatidylinositol-4-phosphate [PI(4)P] phosphoinositides within the VROs. Co-opting cellular enzymes required for lipid biosynthesis and lipid modifications suggest that tombusviruses could create an optimized lipid/membrane microenvironment for efficient VRO assembly and protection of the viral RNAs during virus replication. We propose that compartmentalization of these lipid enzymes within VROs helps tombusviruses replicate in an efficient milieu. In summary, tombusviruses target a major crossroad in the secretory and recycling pathways via coopting the retromer complex and the tubular endosomal network to build VROs in infected cells.

Viruses are intracellular parasites which co-opt cellular resources to produce abundant viral progeny. Positive-strand (+)RNA viruses replicate on subcellular membranes by forming viral replication organelles (VROs) (1–5). VROs sequester the viral proteins and viral RNAs together with co-opted host factors to provide an optimal subcellular environment for the assembly of numerous viral replicase complexes (VRCs), which are then responsible for robust viral RNA replication. VROs also spatially and temporally organize viral replication. Importantly, the VROs hide the viral RNAs from cellular defense mechanisms as well (5, 6). VROs consist of extensively remodeled membranes with unique lipid composition. How viruses achieve these membrane remodeling and lipid modifications and lipid enrichment is incompletely understood. Therefore, currently, there is a major ongoing effort to dissect the VRC assembly process and to understand the roles of viral and host factors in driving the biogenesis of VROs (1, 3, 7).Tomato bushy stunt virus (TBSV), a plant-infecting tombusvirus, has been shown to induce complex rearrangements of cellular membranes and alterations in lipid and other metabolic processes during infections (8–10). The VROs formed during TBSV infections include extensive membrane contact sites (vMCSs) and harbor numerous spherules (containing VRCs), which are vesicle-like invaginations in the peroxisomal membranes (8, 11–13). A major gap in our understanding of the biogenesis of VROs, including vMCSs and VRCs, is how the cellular lipid-modifying enzymes are recruited to the sites of viral replication.Tombusviruses belong to the large Flavivirus-like supergroup that includes important human, animal, and plant pathogens. Tombusviruses have a small single-component (+)RNA genome of ∼4.8 kb that codes for five proteins. Among those, there are two essential replication proteins, namely p33 and p92^pol, the latter of which is the RdRp protein and it is translated from the genomic RNA via readthrough of the translational stop codon in p33 open reading frame (14). The smaller p33 replication protein is an RNA chaperone, which mediates the selection of the viral (+)RNA for replication (14–16). Altogether, p33 is the master regulator of VRO biogenesis (3). We utilize a TBSV replicon (rep)RNA, which contains four noncontiguous segments from the genomic RNA, and it can efficiently replicate in yeast and plant cells expressing p33 and p92^pol (14, 17).Tombusviruses hijack various cellular compartments and pathways for VRO biogenesis (18). These include peroxisomes by TBSV or mitochondria (in the case of the closely related carnation Italian ringspot virus [CIRV]), the endoplasmic reticulum (ER) network, Rab1-positive COPII vesicles, and the Rab5-positive endosomes (8, 19–23). Tombusviruses also induce membrane proliferation, new lipid synthesis, and enrichment of lipids, most importantly phosphatidylethanolamine (PE), sterols, phosphatidylinositol-4-phosphate [PI(4)P], and phosphatidylinositol-3-phosphate [PI(3)P] phosphoinositides in peroxisomal or mitochondrial membranes for different tombusviruses (13, 24–27). This raised the question that how TBSV could hijack lipid synthesis enzymes from other subcellular locations that leads to enrichment of critical lipids in the large VROs in model yeast and plant hosts.The endosomal network (i.e., early, late, and recycling endosomes) is a collection of pleomorphic organelles which sort membrane-bound proteins and lipids either for vacuolar/lysosomal degradation or recycling to other organelles. With the help of the so-called retromer complex, tubular transport carriers formed from the endosomes recycle cargoes to the Golgi and ER or to the plasma membrane (28–31). The core retromer complex consists of three conserved proteins, Vps26, Vps29, and Vps35, which are involved in cargo sorting and selection. The retromer complex affects several cellular processes, including autophagy through the maturation of lysosomes (32), neurodegenerative diseases (33), plant root hair growth (34), and plant immunity (35).The cellular retromer is important for several pathogen–host interactions. For example, the retromer is targeted by Brucella, Salmonella, and Legionella bacteria (36–39) and the rice blast fungus (40). The retromer is also involved in the intracellular transport of the Shigella and Cholera toxins and the plant ricin toxin. The NS5A replication protein of hepatitis C virus (HCV) interacts with Vps35 and this interaction is important for HCV replication in human cells (41). The cytoplasmic tail of the Env protein of HIV-1 binds to the retromer components Vps35 and Vps26, which is required for Env trafficking and infectious HIV-1 morphogenesis (42). Moreover, the retromer complex affects the morphogenesis of vaccinia virus (43) and HPV16 human papillomavirus entry and delivery to the trans-Golgi network (44). Despite the importance of the retromer in pathogen–host interactions, the mechanistic insights are far from complete.In the case of tombusviruses, enrichment of PE and PI(3)P within VROs is facilitated by co-opting the endosomal Rab5 small GTPase and Vps34 PI3K (20, 24), suggesting that the endosome-mediated trafficking pathway might be involved in viral replication in host cells. However, the actual mechanism of how tombusviruses exploit the endosomal/endocytic pathway and induce lipid enrichment within VROs is not yet dissected. Therefore, in this work, we targeted the retromer complex, based on previous genome-wide screens using yeast gene-deletion libraries, which led to the identification of VPS29 and VPS35 as host genes affecting TBSV replication and recombination, respectively (45, 46). These proteins are components of the retromer complex (28–31). We found TBSV and the closely related CIRV co-opt the retromer complex for the biogenesis of VROs in yeast and plants. We observed that depletion of retromer proteins strongly inhibited TBSV and CIRV replication. The recruitment of the retromer is driven by the viral p33 replication protein, which interacts with Vps26, Vps29, and Vps35 retromer proteins. We show that the retromer helps delivering critical cargo proteins, such as Psd2 phosphatidylserine decarboxylase, Vps34 phosphatidylinositol 3-kinase (PI3K), and Stt4 phosphatidylinositol 4-kinase (PI4Kα-like). These co-opted cellular enzymes are then involved in de novo production and enrichment of PE phospholipid, PI(3)P, and PI(4)P phosphoinositides within the VROs. Altogether, these virus-driven activities create an optimized membrane microenvironment within VROs to support efficient tombusvirus replication.     

DNA polymerase ι compensates for Fanconi anemia pathway deficiency by countering DNA replication stress

Fanconi anemia (FA) is caused by defects in cellular responses to DNA crosslinking damage and replication stress. Given the constant occurrence of endogenous DNA damage and replication fork stress, it is unclear why complete deletion of FA genes does not have a major impact on cell proliferation and germ-line FA patients are able to progress through development well into their adulthood. To identify potential cellular mechanisms that compensate for the FA deficiency, we performed dropout screens in FA mutant cells with a whole genome guide RNA library. This uncovered a comprehensive genome-wide profile of FA pathway synthetic lethality, including POLI and CDK4. As little is known of the cellular function of DNA polymerase iota (Pol ι), we focused on its role in the loss-of-function FA knockout mutants. Loss of both FA pathway function and Pol ι leads to synthetic defects in cell proliferation and cell survival, and an increase in DNA damage accumulation. Furthermore, FA-deficient cells depend on the function of Pol ι to resume replication upon replication fork stalling. Our results reveal a critical role for Pol ι in DNA repair and replication fork restart and suggest Pol ι as a target for therapeutic intervention in malignancies carrying an FA gene mutation.

Fanconi anemia (FA) is a genomic instability disorder caused by biallelic or x-linked mutations in any of 22 genes. FA patients are characterized by multiple developmental abnormalities, progressive bone marrow failure, and profound cancer susceptibility (1–3). Germ-line FA mutations predispose an individual to breast, ovarian, pancreatic, and hematological malignancies. Somatic FA mutations have been identified in sporadic acute leukemia and breast cancer (4–6).The FA pathway is the major cellular mechanism responding to DNA crosslinking damage and replication stress. The 22 FA gene products fall into several functional groups. In response to DNA damage, the FANCD2/FANCI complex is monoubiquitinated, signifying the activation of the canonical FA pathway (7, 8). The monoubiquitinated FANCD2/FANCI complex most likely orchestrates the recruitment of nucleolytic factors for the processing of crosslinking DNA damage (9, 10). The FA core complex, consisting of FANCA, -B, -C, -E, -F, -G, and -M, FAAP20, FAAP24, FAAP100, and the RING domain protein FANCL, provides the E3 ligase activity for the damage-induced monoubiquitination of FANCD2/FANCI (11–16). FANCP/XPF and FANCQ/SLX4, the third group of FA gene products, are nucleases or part of the nuclease scaffold, taking part in DNA cleavage for the removal of the crosslinking lesions (8, 17–21). DNA double-strand breaks, as an intermediate structure of ICL (Interstrand CrossLink) repair, depend on the fourth group of FA proteins, required in homologous recombination (FANCD1/BRCA2, FANCO/RAD51C, FANCJ/BARD1, and FANCR/RAD51) (22–26).In addition to the direct role in crosslinking damage repair, FA pathway components are linked to the protection of replication fork integrity during replication interruption that is not directly caused by damage to the DNA. BRCA1/2 are important in stabilizing stalled forks in an MRE11-dependent manner (27, 28). Similarly, FANCD2 and FANCI have been shown to prevent collapse of stalled replication forks (29, 30). Defects in the FA and recombination mechanisms lead to severe fork erosion and endogenous DNA damage accumulation upon reversible replication block, suggesting that the FA pathway plays a crucial role in DNA replication under both normal and perturbed growth conditions (8, 23, 31–34).Given the important role of the FA pathway in replication stress, it is perplexing that cells with a completely impaired FA mechanism are capable of sustained proliferation (34, 35). Overt abnormalities are absent in mice with knockout of several key FA genes (36–39). Moreover, individuals can survive without a functional FA pathway for decades (median life expectancy of 30 y for FA patients) (40). More recently, a genome-scale CRISPR-Cas9 guide RNA (gRNA) library screen has defined gene sets essential for proliferation of common model cell lines (41). None of the classic FA genes which participate in the monoubiquitination process appear to be essential in these screens. Cells deficient in classic FA genes can sustain growth despite the accumulation of endogenous DNA damage. Thus, it seems likely that compensatory mechanisms exist in FA mutant cells to support long-term viability.In this study, we sought to identify cellular mechanisms that are important for the survival of cells deficient in the FA pathway. Comparative genome-scale CRISPR/Cas9 screens were carried out in isogenic FA pathway-proficient and -deficient cells. Genes that exhibit synthetic lethality in FA mutant cells are candidates which compensate for the loss of the FA pathway function. Among the top candidates, we validated and investigated DNA polymerase (Pol) ι as a critical factor for the survival of FA mutant cells. We found that, in FA-deficient cells, Pol ι is crucial in the resumption of stressed replication forks and in suppressing the accumulation of endogenous DNA damage. This reveals a function for Pol ι in relieving DNA damage stress.     

DDX11 loss causes replication stress and pharmacologically exploitable DNA repair defects

DDX11 encodes an iron–sulfur cluster DNA helicase required for development, mutated, and overexpressed in cancers. Here, we show that loss of DDX11 causes replication stress and sensitizes cancer cells to DNA damaging agents, including poly ADP ribose polymerase (PARP) inhibitors and platinum drugs. We find that DDX11 helicase activity prevents chemotherapy drug hypersensitivity and accumulation of DNA damage. Mechanistically, DDX11 acts downstream of 53BP1 to mediate homology-directed repair and RAD51 focus formation in manners nonredundant with BRCA1 and BRCA2. As a result, DDX11 down-regulation aggravates the chemotherapeutic sensitivity of BRCA1/2-mutated cancers and resensitizes chemotherapy drug–resistant BRCA1/2-mutated cancer cells that regained homologous recombination proficiency. The results further indicate that DDX11 facilitates recombination repair by assisting double strand break resection and the loading of both RPA and RAD51 on single-stranded DNA substrates. We propose DDX11 as a potential target in cancers by creating pharmacologically exploitable DNA repair vulnerabilities.

Faithful DNA replication and DNA repair processes are essential for genome integrity. Inherited mutations in BRCA1 or BRCA2 genes predispose to breast and ovarian cancer, among other types of malignancies such as pancreatic cancers and brain tumors (1). Mechanistically, BRCA1 and BRCA2 are critical for double strand break (DSB) repair by homologous recombination (HR) and for the protection of stalled replication forks by facilitating RAD51 filament formation (2).Tumors with mutations in HR factors, the most widespread being those harboring mutations in BRCA1 and BRCA2, are sensitive to chemotherapeutic drugs that block replication and cause DSBs (3). Platinum drugs, such as cisplatin, create intra- and interstrand adducts that require HR activities for DNA repair during replication and therefore are effective in killing HR-defective cancers. Analysis of the plateau of the survival curve of different cancers revealed that patients often develop resistance, and thus, alternative strategies are needed. The advent of PARP (poly ADP ribose polymerase) inhibitors (PARPi), including olaparib, which exhibit synthetic lethal effects when applied to cells and tumors defective in HR (4, 5), holds significant promise. PARP1, 2, and 3 are required to repair numerous DNA single-strand breaks (SSBs) resulting from oxidative damage and during base excision repair. When PARP enzymes are locally trapped at SSBs, they prevent fork progression and generate DSBs (6), which need to be repaired by BRCA1/2 and other HR factors (4, 5). While the synthetic lethality of PARPi and HR deficiency is being exploited clinically, many BRCA-mutated carcinomas acquire resistance to PARPi (2). Identifying key factors that are functionally linked with BRCA1/2 and/or PARP during replication stress response may indicate useful alternative or combinatorial chemotherapeutic strategies.DDX11 is a conserved iron–sulfur (Fe–S) cluster 5′ to 3′ DNA helicase facilitating chromatin structure and DNA repair in manners that are not fully understood. Biallelic DDX11 mutations in humans cause the developmental disorder Warsaw breakage syndrome (WBS), which presents overlaps with Fanconi anemia in terms of chromosomal instability induced by intra- and interstrand crosslinking (ICL) agents and with cohesinopathies in terms of sister chromatid cohesion defects (7, 8). DDX11 has also strong ties to cancer. Specifically, DDX11 is highly up-regulated or amplified in diverse cancers, such as breast and ovarian cancers, including one-fifth of high-grade serous ovarian cancers (cBioPortal and The Cancer Genome Atlas [TCGA]). Moreover, DDX11 is required for the survival of advanced melanomas (9), lung adenocarcinomas (10), and hepatocellular carcinomas (11). In terms of molecular functions, DDX11 interacts physically with the replication fork component Timeless to assist replisome progression and to facilitate epigenetic stability at G-quadruplex (G4) structures and sister chromatid cohesion (12–16). Notably, DDX11 also contributes along 9–1-1, Fanconi anemia factors, and SMC5/6 to prevent cytotoxicity of PARPi and ICLs (17–20). However, if the DNA damage tolerance functions of DDX11 are relevant for tumorigenesis or cancer therapies remains currently unknown.Here, we find that targeting DDX11 sensitizes ovarian and other cancer cell lines to drug therapies involving cisplatin and the PARP inhibitor olaparib. We established DDX11 knockout (KO) in HeLa uterine and U2OS osteosarcoma cancer cell lines and uncovered via chemical drug screens and immunofluorescence of DNA damage markers that they show typical hallmarks of increased replication stress. DDX11 helicase activity and the Fe–S domain are critical to prevent cellular sensitization to olaparib and ICLs and to avert accumulation of DSB markers. Mechanistically, we uncover that DDX11 facilitates homology-directed repair of DSBs and RAD51 focus formation downstream of 53BP1. Importantly, DDX11 is required for viability in BRCA1-depleted cells that are resistant to chemotherapy by concomitant depletion of 53BP1, REV7, and other shieldin components (21, 22), indicating roles for DDX11 in the activated BRCA2-dependent HR pathway, often accounting for the resistance of BRCA1-mutated tumors (2). DDX11 DNA repair function is nonredundant with BRCA1 and BRCA2 pathways, facilitating resection and loading of both RPA and RAD51 on single-stranded DNA substrates. Altogether, our results define a DDX11-mediated DNA repair pathway that creates pharmaceutically targetable vulnerabilities in cancers.     

Zika virus employs the host antiviral RNase L protein to support replication factory assembly

Infection with the flavivirus Zika virus (ZIKV) can result in tissue tropism, disease outcome, and route of transmission distinct from those of other flaviviruses; therefore, we aimed to identify host machinery that exclusively promotes the ZIKV replication cycle, which can inform on differences at the organismal level. We previously reported that deletion of the host antiviral ribonuclease L (RNase L) protein decreases ZIKV production. Canonical RNase L catalytic activity typically restricts viral infection, including that of the flavivirus dengue virus (DENV), suggesting an unconventional, proviral RNase L function during ZIKV infection. In this study, we reveal that an inactive form of RNase L supports assembly of ZIKV replication factories (RFs) to enhance infectious virus production. Compared with the densely concentrated ZIKV RFs generated with RNase L present, deletion of RNase L induced broader subcellular distribution of ZIKV replication intermediate double-stranded RNA (dsRNA) and NS3 protease, two constituents of ZIKV RFs. An inactive form of RNase L was sufficient to contain ZIKV genome and dsRNA within a smaller RF area, which subsequently increased infectious ZIKV release from the cell. Inactive RNase L can interact with cytoskeleton, and flaviviruses remodel cytoskeleton to construct RFs. Thus, we used the microtubule-stabilization drug paclitaxel to demonstrate that ZIKV repurposes RNase L to facilitate the cytoskeleton rearrangements required for proper generation of RFs. During infection with flaviviruses DENV or West Nile Kunjin virus, inactive RNase L did not improve virus production, suggesting that a proviral RNase L role is not a general feature of all flavivirus infections.

The flavivirus genus contains arthropod-transmitted viruses with a positive-sense single-stranded RNA (ssRNA) genome, including Zika virus (ZIKV), dengue virus (DENV), and West Nile virus (WNV). These viruses are transmitted by mosquitos and globally distributed, with high associated morbidity and mortality in humans (1–3). More recently, ZIKV sexual and vertical transmission has been recognized, the latter involving transplacental migration of the virus, potentially resulting in fetal microcephaly (4–9). Due to diversity in ZIKV tissue tropism, disease, and route of transmission as compared with other flaviviruses, it is possible that variances in ZIKV infection at the molecular level confer the observed shifts in clinical outcome at the organismal level. For this reason, we are interested in host machinery that supports the ZIKV replication cycle but not that of other flaviviruses, as this may improve our understanding of the molecular determinants of ZIKV pathogenesis.After entry into the host cell, the flavivirus genome, which also serves as the messenger RNA (mRNA), is directly translated at the endoplasmic reticulum (ER). Proximal to sites of translation, flaviviruses create replication factories (RFs) through extensive cytoskeletal rearrangements that generate invaginations in the folds of the ER membrane, within which new genome synthesis occurs (10–15). These RFs contain ZIKV replication complex proteins, including the NS3 protease, the replication intermediate double-stranded RNA (dsRNA), as well as template genomic ssRNA. New genome is packaged into compartments at opposite ER folds, and new virions traffic through the transgolgi network and eventually bud from the plasma membrane (16). RFs therefore enable efficient throughput of key viral processes as centers of new genome synthesis linked with viral protein translation as well as new virus assembly. In addition, RFs serve as a protective barrier to impede cytosolic innate immune sensing, as flavivirus RNA predominantly resides within RFs during the bulk of the intracellular replication cycle.Innate immune sensors within the cytoplasm of the infected cell detect viral RNA to activate antiviral responses, including the type I interferon (IFN) and oligoadenylate synthetase/ribonuclease L (OAS/RNase L) pathways. Extensive research has demonstrated that flaviviruses, including ZIKV, have evolved strategies for counteracting the type I IFN response (17–22). Since OAS genes are IFN-stimulated genes and therefore up-regulated by type I IFN signaling, the OAS/RNase L pathway can also be potentiated by type I IFN production. However, activation of RNase L can occur in the absence of type I IFN responses when basal OAS expression is sufficient (23, 24). In either event, OAS sensors detect viral dsRNA and generate 2''-5''-oligoadenylates (2-5A). RNase L, which is constitutively expressed in an inactive form, homodimerizes upon 2-5A binding to become catalytically active (25). Active RNase L cleaves both host and viral ssRNA within the cell (Fig. 1). While there are three OAS isoforms, we have shown that the OAS3 isoform is the predominant activator of RNase L during infection with a variety of viruses including ZIKV (23, 26). Activated RNase L cleavage of host ribosomal RNA (rRNA) and mRNA as well as viral ssRNA ultimately inhibits virus infection (26–31).Open in a separate windowFig. 1.Noncanonical RNase L function promotes infectious ZIKV production. (Left) Canonical RNase L antiviral activity. Viral dsRNA is detected by OAS3, which produces the small molecule 2-5A that binds inactive RNase L, inducing its homodimerization and catalytic activation, resulting in cleavage of host and viral ssRNA, leading to inhibition of viral infection. (Right) RNase L activity during ZIKV infection. ZIKV dsRNA is recognized by OAS3, which activates RNase L resulting in ssRNA cleavage; however, ZIKV production is improved with RNase L expression.Once activated, RNase L can restrict infection of a diverse range of DNA and RNA viruses, including flaviviruses DENV and WNV (30, 32–34). Many viruses have subsequently developed mechanisms for evading RNase L antiviral effects, most of which target this pathway upstream of RNase L activation through sequestration of dsRNA, which prevents OAS activation, or by degradation of 2-5A (32, 35–40). We recently showed that ZIKV avoids antiviral effects of activated RNase L and that this evasion strategy requires assembly of RFs to protect genome from RNase L cleavage (23). Despite substantial RNase L–mediated cleavage of intracellular ZIKV genome, a portion of uncleaved genome was shielded from activated RNase L within RFs. This genome was sufficient to produce high levels of infectious virus particles, as infectious ZIKV released from wild-type (WT) cells was significantly higher than from RNase L knockout (KO) cells. Unlike ZIKV, infectious DENV production was decreased by canonical RNase L antiviral activity (23, 33). These results indicated that RNase L expression was ultimately proviral during ZIKV infection (Fig. 1). As this was the initial report of viral resistance to catalytically active RNase L during infection, we sought to isolate the differences between ZIKV RFs and those constructed by other flaviviruses, to identify factors that enable this ZIKV evasion mechanism.In this study, we focused on elucidating how RNase L increases ZIKV production. An earlier study reported that an inactive form of RNase L interacts with the actin cytoskeleton to reorganize cellular framework during viral infection (41). Since flaviviruses reorganize the cellular cytoskeletal and organellar network during infection (11), we investigated the possibility that RNase L was exploited by ZIKV to assemble protective RFs that dually serve as a barrier against host sensors in addition to providing sites of replication.     

Replisome bypass of a protein-based R-loop block by Pif1

Efficient and faithful replication of the genome is essential to maintain genome stability. Replication is carried out by a multiprotein complex called the replisome, which encounters numerous obstacles to its progression. Failure to bypass these obstacles results in genome instability and may facilitate errors leading to disease. Cells use accessory helicases that help the replisome bypass difficult barriers. All eukaryotes contain the accessory helicase Pif1, which tracks in a 5′–3′ direction on single-stranded DNA and plays a role in genome maintenance processes. Here, we reveal a previously unknown role for Pif1 in replication barrier bypass. We use an in vitro reconstituted Saccharomyces cerevisiae replisome to demonstrate that Pif1 enables the replisome to bypass an inactive (i.e., dead) Cas9 (dCas9) R-loop barrier. Interestingly, dCas9 R-loops targeted to either strand are bypassed with similar efficiency. Furthermore, we employed a single-molecule fluorescence visualization technique to show that Pif1 facilitates this bypass by enabling the simultaneous removal of the dCas9 protein and the R-loop. We propose that Pif1 is a general displacement helicase for replication bypass of both R-loops and protein blocks.

Efficient and faithful replication of the genome is essential to maintain genome stability and is carried out by a multiprotein complex called the replisome (1–4). There are numerous obstacles to progression of the replisome during the process of chromosome duplication. These obstacles include RNA-DNA hybrids (R-loops), DNA secondary structures, transcribing RNA polymerases, and other tightly bound proteins (5–9). Failure to bypass these barriers may result in genome instability, which can lead to cellular abnormalities and genetic disease. Cells contain various accessory helicases that help the replisome bypass these difficult barriers (10–20). A subset of these helicases act on the opposite strand of the replicative helicase (1, 2, 14, 19).All eukaryotes contain an accessory helicase, Pif1, which tracks in a 5′–3′ direction on single-stranded DNA (ssDNA) (11–16). Pif1 is important in pathways such as Okazaki-fragment processing and break-induced repair that require the removal of DNA-binding proteins as well as potential displacement of R-loops (11–13, 21, 15–18, 22–25). Genetic studies and immunoprecipitation pull-down assays indicate that Pif1 interacts with PCNA (the DNA sliding clamp), Pol ε (the leading-strand polymerase), the MCMs (the motor subunits of the replicative helicase CMG), and RPA (the single-stranded DNA-binding protein) (15, 26, 27). Pif1 activity in break-induced repair strongly depends on its interaction with PCNA (26). These interactions with replisomal components suggest that Pif1 could interact with the replisome during replication. In Escherichia coli, the replicative helicase is the DnaB homohexamer that encircles the lagging strand and moves in a 5′–3′ direction (20). E. coli accessory helicases include the monomeric UvrD (helicase II) and Rep, which move in the 3′–5′ direction and operate on the opposite strand from the DnaB hexamer. It is known that these monomeric helicases promote the bypass of barriers during replication such as stalled RNA polymerases (5). The eukaryotic replicative helicase is the 11-subunit CMG (Cdc45, Mcm2–7, GINS) and tracks in the 3′–5′ direction, opposite to the direction of Pif1 (25, 28). Once activated by Mcm10, the MCM motor domains of CMG encircle the leading strand (29–32). We hypothesized that, similar to UvrD and Rep in E. coli, Pif1 interacts with the replisome tracking in the opposite direction to enable bypass of replication obstacles.In this report, we use an in vitro reconstituted Saccharomyces cerevisiae replisome to study the role of Pif1 in bypass of a “dead” Cas9 (dCas9), which is a Cas9 protein that is deactivated in DNA cleavage but otherwise fully functional in DNA binding. As with Cas9, dCas9 is a single-turnover enzyme that can be programmed with a guide RNA (gRNA) to target either strand. The dCas9–gRNA complex forms a roadblock consisting of an R-loop and a tightly bound protein (dCas9), a construct that is similar to a stalled RNA polymerase. This roadblock (hereafter dCas9 R-loop) arrests replisomes independent of whether the dCas9 R-loop is targeted to the leading or lagging strand (30). Besides its utility due to its programmable nature (33), the use of the dCas9 R-loop allows us to answer several mechanistic questions. For example, the ability to program the dCas9 R-loop block to any specific sequence enables us to observe whether block removal is different depending on whether the block is on the leading or lagging strand. Furthermore, the inner diameter of CMG can accommodate double-stranded DNA (dsDNA) and possibly an R-loop, but not a dCas9 protein. Using the dCas9 R-loop block allows us to determine the fate of each of its components.Here, we report that Pif1 enables the bypass of the dCas9 R-loop by the replisome. Interestingly, dCas9 R-loops targeted to either the leading or lagging strand are bypassed with similar efficiency. In addition, the PCNA clamp is not required for bypass of the block, indicating that Pif1 does not need to interact with PCNA during bypass of the block. We used a single-molecule fluorescence imaging to show that both the dCas9 and the R-loop are displaced as an intact nucleoprotein complex. We propose that Pif1 is a general displacement helicase for replication bypass of both R-loops and protein blocks.     

Conditions and extent of volatile loss from the Moon during formation of the Procellarum basin

Rocks from the lunar interior are depleted in moderately volatile elements (MVEs) compared to terrestrial rocks. Most MVEs are also enriched in their heavier isotopes compared to those in terrestrial rocks. Such elemental depletion and heavy isotope enrichments have been attributed to liquid–vapor exchange and vapor loss from the protolunar disk, incomplete accretion of MVEs during condensation of the Moon, and degassing of MVEs during lunar magma ocean crystallization. New Monte Carlo simulation results suggest that the lunar MVE depletion is consistent with evaporative loss at 1,670 ± 129 K and an oxygen fugacity +2.3 ± 2.1 log units above the fayalite-magnetite-quartz buffer. Here, we propose that these chemical and isotopic features could have resulted from the formation of the putative Procellarum basin early in the Moon’s history, during which nearside magma ocean melts would have been exposed at the surface, allowing equilibration with any primitive atmosphere together with MVE loss and isotopic fractionation.

Returned samples of basaltic rocks from the Moon provided evidence decades ago that the Moon is depleted in volatile elements compared to the Earth (1), with lunar basalt abundances of moderately volatile elements (MVEs) being ∼1/5 that of terrestrial basalt abundances for alkali elements and ∼1/40 for other MVE, such as Zn, Ag, In, and Cd (2). The theme of lunar volatiles thus seemed settled. Yet, the unambiguous detection in 2008 of lunar indigenous hydrogen and other volatile elements, such as F, Cl, and S in pyroclastic glasses (3), heralded a new era of research into lunar volatiles, overturning the decades-old paradigm of a bone-dry Moon (e.g., refs. 4 and 5). Here, we define volatile elements as those with 50% condensation temperatures below these of the major rock-forming elements Fe, Mg, and Si (6). This paradigm shift was accompanied by new measurements of volatile stable isotope compositions (e.g., H, C, N, Cl, K, Cr, Cu, Zn, Ga, Rb, and Sn) in a wealth of bulk lunar samples (7–18) and in the mineral phases and melt inclusions they host (19–28). These studies have shown that the stable isotope compositions of most MVEs (e.g., K, Zn, Ga, and Rb) are enriched in their heavier isotopes with respect to the bulk silicate Earth (BSE) (9, 11, 13–15, 17). Such heavy isotope enrichment is associated with elemental depletion, which has been variously attributed to liquid–vapor exchange and vapor loss from the protolunar disk (17, 18), incomplete accretion of MVEs during condensation of the Moon (13, 29, 30), and degassing of these elements during lunar magma ocean crystallization (9, 11, 14, 15, 25, 31). Almost all these hypotheses have typically assumed that the MVE depletions and associated MVE isotope fractionations are relevant to the whole Moon. However, our lunar sample collections are biased, as all Apollo and Luna returned samples come from the lunar nearside from within or around the anomalous Procellarum KREEP Terrane (PKT) region (e.g., ref. 32), where KREEP stands for enriched in K, REEs, and P. Barnes et al. (26) proposed that the heavy Cl isotope signature measured in KREEP-rich Apollo samples resulted from metal-chloride degassing from late-stage lunar magma ocean melts in response to a large crust-breaching impact event, spatially associated with the PKT region, which facilitated exposure of these late-stage melts to the lunar surface. Here, we further investigate whether a localized impact event could have been responsible for the general MVE depletion and heavy MVE isotope enrichment measured in lunar samples.     

Stimulator of interferon genes (STING) is an essential proviral host factor for human rhinovirus species A and C

Human rhinoviruses (RVs) are positive-strand RNA viruses that cause respiratory tract disease in children and adults. Here we show that the innate immune signaling protein STING is required for efficient replication of members of two distinct RV species, RV-A and RV-C. The host factor activity of STING was identified in a genome-wide RNA interference (RNAi) screen and confirmed in primary human small airway epithelial cells. Replication of RV-A serotypes was strictly dependent on STING, whereas RV-B serotypes were notably less dependent. Subgenomic RV-A and RV-C RNA replicons failed to amplify in the absence of STING, revealing it to be required for a step in RNA replication. STING was expressed on phosphatidylinositol 4-phosphate (PI4P)-enriched membranes and was enriched in RV-A16 compared with RV-B14 replication organelles isolated in isopycnic gradients. The host factor activity of STING was species-specific, as murine STING (mSTING) did not rescue RV-A16 replication in STING-deficient cells. This species specificity mapped primarily to the cytoplasmic, ligand-binding domain of STING. Mouse-adaptive mutations in the RV-A16 2C protein allowed for robust replication in cells expressing mSTING, suggesting a role for 2C in recruiting STING to RV-A replication organelles. Palmitoylation of STING was not required for RV-A16 replication, nor was the C-terminal tail of STING that mediates IRF3 signaling. Despite co-opting STING to promote its replication, interferon signaling in response to STING agonists remained intact in RV-A16 infected cells. These data demonstrate a surprising requirement for a key host mediator of innate immunity to DNA viruses in the life cycle of a small pathogenic RNA virus.

Human rhinoviruses (RVs) are ubiquitous respiratory pathogens composing a large group of antigenically diverse, positive-strand RNA viruses classified within the Enterovirus genus of the Picornaviridae family (1, 2). The most frequent cause of the common cold, RV infections among the young are associated with the development of asthma (3, 4). In older individuals, RV infections may also lead to acute exacerbations of asthma and chronic obstructive pulmonary disease and are a significant cause of lower respiratory tract disease (5, 6). RVs are grouped phylogenetically into three species, each containing multiple serotypes (2, 7). Unlike RV-A and RV-B, which have been recognized for decades and readily propagated in conventional cell cultures, RV-C was identified more recently and replicates in vitro only in cells engineered to express a critical entry factor, cadherin-related family member 3 (CDHR3) (8). RV-C is strongly associated with severe respiratory tract infections in young children and is more closely related to RV-A than to RV-B (2, 7, 9, 10). Nearly all hospital visits related to RV-triggered asthma are due to infections with RV-A or RV-C viruses, with RV-C associated with more severe symptoms (10–12).The molecular mechanisms underlying replication of these RNA viruses are only partially understood. Enteroviral RNAs are synthesized on the cytosolic surface of membranous cytoplasmic tubulovesicular structures (13–15). These replication organelles are derived from remodeled endoplasmic reticulum (ER) or Golgi membranes and contain multiple viral nonstructural proteins, including 2B, 2C, and an RNA-dependent RNA polymerase, 3D^pol (16). The formation of replication organelles is associated with a striking reordering of cellular lipid metabolism, with phosphatidylinositol 4-kinase-IIIβ (PI4Kβ) playing a key role. PI4Kβ is recruited to membranes at the site of replication by the viral 3A protein acting in concert with host acyl-CoA binding domain-containing 3 (ACBD3) (13, 17, 18). PI4Kβ mediates the enrichment of these membranes with phosphatidylinositol 4-phosphate (PI4P), leading to subsequent recruitment of oxysterol-binding protein 1 (OSBP1), which enhances cholesterol flux into the membranes (18). Thus, ACBD3, PI4Kβ, and OSBP1 are all crucial host factors for RV replication.The intracellular replication of poliovirus, a closely-related enterovirus, is also dependent on components of host autophagic signaling, including LC3 protein that associates with the membranes of replication organelles in a nonlipidated form (19, 20). Whether this is also true for rhinoviruses is uncertain. Unlike poliovirus, RV-A1a replication is not influenced by chemical compounds that promote or inhibit autophagy, rapamycin, and 3-methyadenine (3-MA) respectively, while similar studies of RV-A2 produced conflicting results (21, 22). These latter data show that even among closely related viruses in the same picornaviral genus, host factors involved in remodeling membranes and generating replication organelles may vary substantially. Here we describe a surprising requirement for the Stimulator of Interferon Genes (STING) protein in intracellular replication of RV-A and RV-C viruses. STING (also known as MITA, ERIS, or MPYS) is an essential adaptor protein downstream of cGMP-AMP synthase (cGAS) in the innate immune cytosolic DNA-sensing pathway, and thus is typically associated with antiviral rather than proviral effects (23–27). We show that RV-A16 replication organelles are enriched in STING, and that transfected subgenomic RV-A16 and RV-C15 RNA replicons fail to amplify in the absence of STING. Genetic evidence links STING to the nonstructural 2C protein of RV-A, which is known to play a crucial role in the formation of replication organelles.     

NAD(H)-mediated tetramerization controls the activity of Legionella pneumophila phospholipase PlaB

The virulence factor PlaB promotes lung colonization, tissue destruction, and intracellular replication of Legionella pneumophila, the causative agent of Legionnaires’ disease. It is a highly active phospholipase exposed at the bacterial surface and shows an extraordinary activation mechanism by tetramer deoligomerization. To unravel the molecular basis for enzyme activation and localization, we determined the crystal structure of PlaB in its tetrameric form. We found that the tetramer is a dimer of identical dimers, and a monomer consists of an N-terminal α/β-hydrolase domain expanded by two noncanonical two-stranded β-sheets, β-6/β-7 and β-9/β-10. The C-terminal domain reveals a fold displaying a bilobed β-sandwich with a hook structure required for dimer formation and structural complementation of the enzymatic domain in the neighboring monomer. This highlights the dimer as the active form. Δβ-9/β-10 mutants showed a decrease in the tetrameric fraction and altered activity profiles. The variant also revealed restricted binding to membranes resulting in mislocalization and bacterial lysis. Unexpectedly, we observed eight NAD(H) molecules at the dimer/dimer interface, suggesting that these molecules stabilize the tetramer and hence lead to enzyme inactivation. Indeed, addition of NAD(H) increased the fraction of the tetramer and concomitantly reduced activity. Together, these data reveal structural elements and an unprecedented NAD(H)-mediated tetramerization mechanism required for spatial and enzymatic control of a phospholipase virulence factor. The allosteric regulatory process identified here is suited to fine tune PlaB in a way that protects Legionella pneumophila from self-inflicted lysis while ensuring its activity at the pathogen–host interface.

Phospholipases are important enzymes in infectious disease pathogenesis involved in host modulation and damage. For example, some bacterial phospholipases, such as Pseudomonas aeruginosa ExoU, massively damage cells and contribute to host inflammatory response (1–4). Others, such as Legionella pneumophila VipD, facilitate bacterial intracellular replication by inhibiting phagosomal maturation, or as in the case of Listeria monocytogenes, phospholipases PlcA and PlcB are required for bacterial escape from the enclosing vacuole and for cell-to-cell spread (1, 5–8). Phospholipases have been assigned to different groups depending on the preferred cleavage site within their substrates. Phospholipases A (PLAs) and lysophospholipases A (LPLA) hydrolyze carboxyl ester bonds at the sn-1 or sn-2 position in phospholipids or lysophospholipids, respectively, and release fatty acids (1, 9). In L. pneumophila, a Gram-negative bacterium that causes Legionnaires’ disease, at least 15 genes encoding PLAs/LPLAs belonging to three families are found. Many of these are secreted to modulate the host cell but only one, PlaB, is uniquely presented at the bacterial surface (1, 10–13).In this study, we focused on PlaB, a hemolysin and virulence factor that promotes intracellular replication in macrophages by its PLA and LPLA activities (14–16). PlaB is also crucial for lung colonization and tissue destruction in guinea pig infections (13). It is the only characterized member of a recently discovered PLA family, and homologs are found in a wide array of mostly water-associated bacteria including the opportunistic pathogen P. aeruginosa (14, 15). Previous work suggested that PlaB is organized into two domains, namely an N-terminal phospholipase (amino acids 1 to ∼300) connected to a C-terminal domain (CTD) (amino acids ∼301 to 474) that is also essential for activity (15). The catalytic triad S85/D203/H251 of the N-terminal domain (NTD) of PlaB and its homologs is embedded in uncommon consensus motifs which are unique among lipases (15). The CTD is not related to formerly characterized proteins, but we have earlier shown that the last 15 amino acids of PlaB are necessary for activity, although their exact role is not understood (15, 16). Subcellular fractionation and proteinase K digests revealed that PlaB is associated with the outer membrane (OM) and exposed on the surface. However, due to the apparent lack of export signal sequences, lipid anchors, or transmembrane helices, determinants for export and membrane association have not yet been characterized and therefore still remain elusive (13, 14).PlaB represents a highly active PLA/LPLA of L. pneumophila and hydrolyzes lipids, such as phosphatidylcholine (PC) and phosphatidylglycerol (PG) found in the lung of the human host and in Legionella (14, 17–19). Hence, we reasoned that a control mechanism of enzyme activity may be crucial to prevent damage to the pathogen itself. Indeed, PlaB shows an extraordinary activation mechanism that requires protein deoligomerization. At higher PlaB concentrations, the enzyme occurs in an inactive tetrameric form, whereas it deoligomerizes in the lower nanomolar concentration range where it possesses its highest specific activity (16). However, the mechanism behind enzyme regulation and whether the resulting dimer or monomer represents the active form is not understood.To decipher the molecular basis for PlaB’s unusual activation and to gain insight on how it associates with the OM, we have determined its crystal structure. This allowed us to identify important structural features and, interestingly, revealed an NAD(H)-mediated tetramerization mechanism that controls activity.     

Wild-type α-synuclein inherits the structure and exacerbated neuropathology of E46K mutant fibril strain by cross-seeding

Heterozygous point mutations of α-synuclein (α-syn) have been linked to the early onset and rapid progression of familial Parkinson’s diseases (fPD). However, the interplay between hereditary mutant and wild-type (WT) α-syn and its role in the exacerbated pathology of α-syn in fPD progression are poorly understood. Here, we find that WT mice inoculated with the human E46K mutant α-syn fibril (hE46K) strain develop early-onset motor deficit and morphologically different α-syn aggregation compared with those inoculated with the human WT fibril (hWT) strain. By using cryo-electron microscopy, we reveal at the near-atomic level that the hE46K strain induces both human and mouse WT α-syn monomers to form the fibril structure of the hE46K strain. Moreover, the induced hWT strain inherits most of the pathological traits of the hE46K strain as well. Our work suggests that the structural and pathological features of mutant strains could be propagated by the WT α-syn in such a way that the mutant pathology would be amplified in fPD.

α-Synuclein (α-Syn) is the main component of Lewy bodies, which serve as the common histological hallmark of Parkinson’s disease (PD) and other synucleinopathies (1, 2). α-Syn fibrillation and cell-to-cell transmission in the brain play essential roles in disease progression (3–5). Interestingly, WT α-syn could form fibrils with distinct polymorphs, which exhibit disparate seeding capability in vitro and induce distinct neuropathologies in mouse models (6–10). Therefore, it is proposed that α-syn fibril polymorphism may underlie clinicopathological variability of synucleinopathies (6, 9). In fPD, several single-point mutations of SNCA have been identified, which are linked to early-onset, severe, and highly heterogeneous clinical symptoms (11–13). These mutations have been reported to influence either the physiological or pathological function of α-syn (14). For instance, A30P weakens while E46K strengthens α-syn membrane binding affinity that may affect its function in synaptic vesicle trafficking (14, 15). E46K, A53T, G51D, and H50Q have been found to alter the aggregation kinetics of α-syn in different manners (15–17). Recently, several cryogenic electron microscopy (cryo-EM) studies revealed that α-syn with these mutations forms diverse fibril structures that are distinct from the WT α-syn fibrils (18–26). Whether and how hereditary mutations induced fibril polymorphism contributes to the early-onset and exacerbated pathology in fPD remains to be elucidated. More importantly, most fPD patients are heterozygous for SNCA mutations (12, 13, 27, 28), which leads to another critical question: could mutant fibrils cross-seed WT α-syn to orchestrate neuropathology in fPD patients?E46K mutation is one of the eight disease-causing mutations on SNCA originally identified from a Spanish family with autosomal-dominant PD (11). E46K-associated fPD features early-onset motor symptoms and rapid progression of dementia with Lewy bodies (11). Studies have shown that E46K mutant has higher neurotoxicity than WT α-syn in neurons and mouse models overexpressing α-syn (29–32). The underlying mechanism is debatable. Some reported that E46K promotes the formation of soluble species of α-syn without affecting the insoluble fraction (29, 30), while others suggested that E46K mutation may destabilize α-syn tetramer and induce aggregation (31, 32). Our previous study showed that E46K mutation disrupts the salt bridge between E46 and K80 in the WT fibril strain and rearranges α-syn into a different polymorph (33). Compared with the WT strain, the E46K fibril strain is prone to be fragmented due to its smaller and less stable fibril core (33). Intriguingly, the E46K strain exhibits higher seeding ability in vitro, suggesting that it might induce neuropathology different from the WT strain in vivo (33).In this study, we found that human E46K and WT fibril strains (referred to as hE46K and hWT strains) induced α-syn aggregates with distinct morphologies in mice. Mice injected with the hE46K strain developed more α-syn aggregation and early-onset motor deficits compared with the mice injected with the hWT strain. Notably, the hE46K strain was capable of cross-seeding both human and mouse WT (mWT) α-syn to form fibrils (named as hWT_cs and mWT_cs). The cross-seeded fibrils replicated the structure and seeding capability of the hE46K template both in vitro and in vivo. Our results suggest that the hE46K strain could propagate its structure as well as the seeding properties to the WT monomer so as to amplify the α-syn pathology in fPD.