Sequential de novo centromere formation and inactivation on a chromosomal fragment in maize

The ability of centromeres to alternate between active and inactive states indicates significant epigenetic aspects controlling centromere assembly and function. In maize (Zea mays), misdivision of the B chromosome centromere on a translocation with the short arm of chromosome 9 (TB-9Sb) can produce many variants with varying centromere sizes and centromeric DNA sequences. In such derivatives of TB-9Sb, we found a de novo centromere on chromosome derivative 3-3, which has no canonical centromeric repeat sequences. This centromere is derived from a 288-kb region on the short arm of chromosome 9, and is 19 megabases (Mb) removed from the translocation breakpoint of chromosome 9 in TB-9Sb. The functional B centromere in progenitor telo2-2 is deleted from derivative 3-3, but some B-repeat sequences remain. The de novo centromere of derivative 3-3 becomes inactive in three further derivatives with new centromeres being formed elsewhere on each chromosome. Our results suggest that de novo centromere initiation is quite common and can persist on chromosomal fragments without a canonical centromere. However, we hypothesize that when de novo centromeres are initiated in opposition to a larger normal centromere, they are cleared from the chromosome by inactivation, thus maintaining karyotype integrity.The centromere is an important chromosomal region responsible for correct chromosome segregation during cell division. Centromeres are found in the primary constriction region on the chromosome, upon which the kinetochore complex assembles to produce a platform for spindle binding (1). Centromere function is conserved among different species, and several epigenetic markers of active centromeres have been found, including a histone H3 variant referred to as CENH3 in plants (2, 3) or CENP-A in animals (4–6) and phosphorylation of histone H2A at Thr133 in plants (7). Correct loading of CENH3 to the centromere region is a key component of kinetochore assembly (8). Centromeric DNA sequences have experienced rapid evolution (9, 10), and arrangements of DNA sequence in centromere regions differ in species and even in different chromosomes of an individual organism (11). Myriad repeat sequences exist in the centromeres of higher plants. In maize (Zea mays), there are two major types of centromere specific DNA sequences: the simple satellite repeat sequence CentC (12) and centromeric retrotransposon of maize (CRM) (3). Many epigenetic features have been identified in centromeric regions, including DNA methylation levels, histone variants, histone modifications, and RNA components (11). Both epigenetic elements and DNA sequences take part in centromere formation and maintenance, but it is still unknown how genetic and epigenetic factors work together in this process.The centromere is one of the most complex regions on the chromosome, and complete DNA sequencing through the centromeric region is difficult to obtain due to their highly repetitive nature. Centromere sizes, defined by CENP-A/CENH3 binding regions, range from 125 bp in Saccharomyces cerevisiae to 500–1,500 kb in humans and mice (11). In plants, centromere sizes can range to several megabases (Mb) with many repetitive transposable elements, which makes it difficult to study centromere structure and function. For example, the sizes of centromere 2 and 5 in maize are roughly 2 and 7 Mb, respectively.Previous work sought misdivision derivatives of the B chromosome centromere using a translocation between the supernumerary chromosome and the short arm of chromosome 9 (9S) to reduce the size of the centromere for functional studies (13–15). B chromosomes are extra chromosomes that have been found in many plants, animals, and fungi. In maize, a reciprocal translocation between a B chromosome and the short arm of chromosome 9 produced two chromosomes referred to as B-9 and 9-B (13), together referred to as TB-9Sb. Chromosome 9-B contains the long arm of the B chromosome and most of chromosome 9, including its centromere. Correspondingly, chromosome B-9 contains part of the short arm of chromosome 9 and the other part of the B chromosome with the active B centromere. The translocation breakpoint is near Wx1, which is located on 9-B (16). The B centromere of B-9 can undergo misdivision during meiosis, producing many derivatives (14). The first misdivision derivative was a pseudoisochromosome, and subsequently, many telocentric chromosomes and isochromosomes were derived by additional misdivisions (13, 17). Misdivision events can be recognized in crosses of TB-9Sb onto a tester via a fusion-breakage cycle recognized by the behavior of the C1 color marker on the B-9 chromosome. The cycle continues during endosperm development to produce a mosaic phenotype but is “healed” in the embryo, which when grown and analyzed cytologically will reveal the nature of the new chromosomes formed (13, 17). This type of screen was used to assemble a large collection of misdivisions to examine the structural features of the B centromere (18). Centromere sizes of these derivatives were changed and progressively reduced. In these previous studies, molecular analysis of centromere size relied on studying the B centromere-specific DNA repeat before the maize centromere elements, CentC and CRM, were known. The B-specific repeat allows this centromere to be studied against the background of the other centromeres; it surrounds and is interspersed within the active core of the B centromere (15, 19).New functional centromeres formed at ectopic locations rather than native centromeric regions on the chromosomes are called de novo centromeres. Many de novo centromeres have been found in human patients and other organisms (20, 21). There are reports of de novo centromeres in plants, such as barley (Hordeum vulgare) (22), oat (Avena sativa)-maize addition lines (23), and maize (24, 25). The conditions for de novo centromere formation remain unclear (26). Recent research revealed that many de novo centromeres prefer to form near native centromeric regions or in the heterochromatic regions, such as the pericentromere and telomere (27, 28). There are also de novo centromeres in human formed far from native centromeres (20). We have previously described two de novo centromeres in maize: one is near the position of the native centromere (25) and the other is distal to the site of the corresponding native centromere (24). Specific chromatin environments may be required for centromere formation, but the major elements are as yet unknown.DNA sequence alone is insufficient to direct centromere formation, and dicentric chromosomes containing two centromeres are good examples. To be stable, structurally dicentric chromosomes must have one inactive and one active centromere; otherwise, two active centromeres will lead to chromosome breakage during cell division. In maize, many dicentric chromosomes have been reported from B-A translocation chromosome derivatives (29). Dicentric chromosomes can be produced through the process of the chromosome type breakage-fusion-bridge (BFB) cycle, and the inactive centromeres can be reactivated by intrachromosomal recombination (30). The DNA sequences of the active and inactive centromeres of dicentric chromosomes are essentially identical, but the centromere activity states are completely different. We screened several misdivision derivatives using FISH probes specific to maize centromere sequences, CentC and CRM, as well as probes specific to the B centromere repeat sequence (B-repeat) to gain further insight into the nature of the centromeres in this collection. We discovered that one such chromosome, derivative 3-3, lacks detectable CentC and CRM signals, but still has a functional centromere that is not associated with the B-repeat sequence. The results of chromatin immunoprecipitation sequencing (ChIP-seq) using maize CENH3 antibody revealed that a 288-kb region on 9S is involved in the de novo centromere formation. The functional B centromere of progenitor telo2-2 is deleted from derivative 3-3. Further, new derivatives of derivative 3-3 had been selected (31) but there was no change in the B-specific repeat patterns. Here, we found that the de novo centromere of 3-3 has become inactive in all of its derivatives, and in each case a shift to a new de novo centromere position occurred; one of these contains only a 200-kb CENH3 binding region within 9S. The other two are apparently in B chromosome sequences. Thus, sequential de novo centromere formation and exchange of centromere activity occurred in chromosome 3-3 and its derivatives, providing new insight into centromere formation and maintenance.These results help formulate the nature of de novo centromere formation. In all of the examples now documented in maize, the size range is within a few hundred kilobases. In contrast, normal maize centromeres, as noted above, are typically several megabases. The regular occurrence of de novo centromeres found here and previously (24, 25) indicates that they are capable of being formed regularly on chromosomal fragments that are structurally acentric; however, they do not persist in normal chromosomes. The reason might reside in the previous observation in maize (30) and wheat (32) that in functional dicentrics the smaller centromere becomes inactive in a tug of war between large and small. However, in the absence of a normal centromere, the present work illustrates that de novo centromeres can persist. Thus, in normal chromosomes, if a de novo is initiated, it will be as quickly inactivated in opposition to the much larger preexisting centromere; the chromosome will not be affected, and will seldom change structure over evolutionary time despite such a high rate of de novo formation. This hypothesis also suggests that a selective pressure will be placed on the normal centromeres to expand to a size that can regularly inactivate de novo centromeres based on their initial size at formation.     

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.     

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.     

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.     

Ccp1-Ndc80 switch at the N terminus of CENP-T regulates kinetochore assembly

Kinetochores, a protein complex assembled on centromeres, mediate chromosome segregation. In most eukaryotes, centromeres are epigenetically specified by the histone H3 variant CENP-A. CENP-T, an inner kinetochore protein, serves as a platform for the assembly of the outer kinetochore Ndc80 complex during mitosis. How CENP-T is regulated through the cell cycle remains unclear. Ccp1 (counteracter of CENP-A loading protein 1) associates with centromeres during interphase but delocalizes from centromeres during mitosis. Here, we demonstrated that Ccp1 directly interacts with CENP-T. CENP-T is important for the association of Ccp1 with centromeres, whereas CENP-T centromeric localization depends on Mis16, a homolog of human RbAp48/46. We identified a Ccp1-interaction motif (CIM) at the N terminus of CENP-T, which is adjacent to the Ndc80 receptor motif. The CIM domain is required for Ccp1 centromeric localization, and the CIM domain–deleted mutant phenocopies ccp1Δ. The CIM domain can be phosphorylated by CDK1 (cyclin-dependent kinase 1). Phosphorylation of CIM weakens its interaction with Ccp1. Consistent with this, Ccp1 dissociates from centromeres through all stages of the cell cycle in the phosphomimetic mutant of the CIM domain, whereas in the phospho-null mutant of the domain, Ccp1 associates with centromeres during mitosis. We further show that the phospho-null mutant disrupts the positioning of the Ndc80 complex during mitosis, resulting in chromosome missegregation. This work suggests that competitive exclusion between Ccp1 and Ndc80 at the N terminus of CENP-T via phosphorylation ensures precise kinetochore assembly during mitosis and uncovers a previously unrecognized mechanism underlying kinetochore assembly through the cell cycle.

The precise inheritance of genetic information relies on the accurate segregation of chromosomes in mitosis and meiosis. Kinetochores are large protein complexes assembled on centromeres and play a crucial role in chromosome segregation. The kinetochore links the chromosome to microtubule polymers, drives the movement of chromosomes, and ensures correct microtubule–kinetochores attachment (1–3). The kinetochore assembly is thus tightly regulated. Yet, the mechanism by which kinetochores are precisely assembled through the cell cycle remains poorly understood.The kinetochore comprises an outer region and an inner region. The outer kinetochore interacts with microtubules and is assembled on the platform of the inner kinetochore. The inner kinetochore consists of a complex of 14 to 16 subunits known as the constitutive centromere–associated network (CCAN) that is directly built on centromeric chromatin (4–6). In centromeres, the histone H3 variant, CENP-A, replaces the canonical histone H3 to form CENP-A–containing nucleosomes (7–9). Most eukaryotes contain large complex regional centromeres where CENP-A–containing nucleosomes are interspersed with canonical H3–containing nucleosomes (10–12). Regional centromeres are epigenetically specified by CENP-A (12–14). But how CENP-A– and histone H3–containing nucleosomes are balanced in centromeres remains unclear.CENP-T, an integral component of CCAN, is also a histone fold–containing protein. CENP-T provides a platform for the assembly of the Ndc80 complex (Ndc80C), an essential outer kinetochore component, during mitosis (5, 15–18). Ndc80C acts as the interface between microtubules and kinetochores and mediates the microtubule attachments (19, 20). The long N terminus of CENP-T contains a conserved Ndc80 receptor motif. The motif forms an alpha-helix that directly interacts with the Spc24-Spc25 heterodimer in Ndc80C (15, 16). The motif can be phosphorylated by cyclin-dependent kinase 1 (CDK1) to stabilize the interaction between CENP-T and Ndc80C (16, 21–24). However, how CENP-T is regulated through the cell cycle to mediate the assembly of Ndc80C is still not well understood.CENP-T has been shown to interact with three other histone fold–containing proteins, CENP-W, CENP-S, and CENP-X, to form the heterotetrameric nucleosome-like structure in vitro (25, 26). The CENP-T-W-S-X complex directly associates with centromeric DNA. The DNA binding activity of the complex is important for kinetochore formation (5, 25). Interestingly, the complex also directly associates with histone H3, not with CENP-A (5, 27), suggesting that CENP-T particles and the CENP-A nucleosome occupy different positions in centromeres. How the spatial relationship between the CENP-A nucleosome and CENP-T particles in centromeres is regulated remains unclear.The fission yeast Schizosaccharomyces pombe contains large regional centromeres and is considered to be a model system for centromere study. The CENP-A homolog, Cnp1, is enriched in centromere cores, which are surrounded by pericentromeric heterochromatin (28–30). CENP-A^Cnp1 nucleosomes nucleate kinetochore assembly. Mislocalization of CENP-A^Cnp1 results in severe chromosome segregation defects in fission yeast (28, 31–34). Fission yeast also contains the CENP-T homolog, Cnp20, which associates with centromeres throughout the cell cycle. The same as in higher eukaryotes, CENP-T^Cnp20 in S. pombe is essential for viability (35).Recently, Ccp1, a nucleosome assembly protein (NAP) family protein, has been shown to play an important role in antagonizing the loading of CENP-A in fission yeast (36). Ccp1 forms a homodimer and is enriched at centromeres. Ccp1 acts as a key player in balancing CENP-A and histone H3 levels in the region (36). How Ccp1 regulates the CENP-A level in centromeres remains elusive. Interestingly, its centromere localization is cell cycle regulated. Ccp1 is dissociated from centromeres at the onset of mitosis and reassociates with centromeres at the end of mitosis (36, 37). The biological importance of the cell cycle–dependent interaction between Ccp1 and centromeres is unknown.Here using mass spectrometry, we found that Ccp1 interacts directly with CENP-T^Cnp20 in fission yeast. We further identified a conserved Ccp1-interaction motif (CIM) at the N terminus of CENP-T^Cnp20, which is adjacent to the Ndc80 receptor motif. We demonstrated that CIM is important for Ccp1 localization. Furthermore, our data suggested that CDK1-mediated phosphorylation of the CIM motif at the onset of mitosis dissociates Ccp1 from CENP-T^Cnp20, allowing proper positioning of Ndc80C. Ccp1 associates with centromeres during mitosis in the phospho-null mutant of the CIM domain, leading to mislocalization of Ndc80C and severe chromosome segregation defects. Our study uncovers a previously unrecognized mechanism regulating kinetochore organization in regional centromeres.     

Polymerase δ promotes chromosomal rearrangements and imprecise double-strand break repair

Recent studies have implicated DNA polymerases θ (Pol θ) and β (Pol β) as mediators of alternative nonhomologous end-joining (Alt-NHEJ) events, including chromosomal translocations. Here we identify subunits of the replicative DNA polymerase δ (Pol δ) as promoters of Alt-NHEJ that results in more extensive intrachromosomal mutations at a single double-strand break (DSB) and more frequent translocations between two DSBs. Depletion of the Pol δ accessory subunit POLD2 destabilizes the complex, resulting in degradation of both POLD1 and POLD3 in human cells. POLD2 depletion markedly reduces the frequency of translocations with sequence modifications but does not affect the frequency of translocations with exact joins. Using separation-of-function mutants, we show that both the DNA synthesis and exonuclease activities of the POLD1 subunit contribute to translocations. As described in yeast and unlike Pol θ, Pol δ also promotes homology-directed repair. Codepletion of POLD2 with 53BP1 nearly eliminates translocations. POLD1 and POLD2 each colocalize with phosphorylated H2AX at ionizing radiation-induced DSBs but not with 53BP1. Codepletion of POLD2 with either ligase 3 (LIG3) or ligase 4 (LIG4) does not further reduce translocation frequency compared to POLD2 depletion alone. Together, these data support a model in which Pol δ promotes Alt-NHEJ in human cells at DSBs, including translocations.

Translocations are genetic rearrangements involving the fusion of heterologous chromosomes (1) and can be initiated by two or more DNA double-strand breaks (DSBs) (2, 3). DSBs in human cells are repaired by multiple pathways with distinct genetic requirements. The first pathway, homology-directed repair (HDR), is active during the S/G2 phase of the cell cycle, when sister chromatids are present to template DNA synthesis. In the first step of HDR, the DSB ends undergo 5′-to-3′ single-strand resection that involves the Mre11/Rad50/Nbs1 (MRN) complex and the endonuclease CtIP (4). The RPA complex binds the exposed single-stranded DNA and is then exchanged for RAD51 by BRCA2 (5). The RAD51 nucleoprotein filament then facilitates strand invasion into a homologous duplex that serves as a repair template. Multiple DNA polymerases, including the replicative polymerases (Pol δ and Pol ε) and translesion DNA polymerases can participate in DNA synthesis during HDR in mammalian cells (6, 7), which is followed by ligation of the ends.The second pathway, classic nonhomologous end-joining (C-NHEJ), is active throughout the cell cycle and therefore responsible for the repair of most DSBs in somatic cells (5). In C-NHEJ, the DSB is bound by the Ku70/Ku80 (Ku) heterodimer, resulting in recruitment of DNA-dependent protein kinase and the end-bridging factors XLF and PAXX (8). End-processing factors, including Artemis and DNA polymerases μ and λ, can also be recruited for end-resection and gap-filling (9, 10). The XRCC4/LIGIV complex is recruited and ligates both strands (11).The third type of repair, alternative NHEJ (Alt-NHEJ), is often described as a back-up end-joining process, as it resolves a greater fraction of DSBs when C-NHEJ is compromised (12). Alt-NHEJ is also more error-prone than C-NHEJ and appears to contribute to mutagenesis in many types of cancer cells (13, 14). A large number of potentially redundant factors may participate in Alt-NHEJ, including CtIP, MRN, Pol θ (encoded by POLQ), poly(ADP-ribose) polymerase 1 (PARP1), and Ligases 1 and 3 (Lig1/3) (4, 12, 15–21). The large number of factors suggests that the “pathway” is actually a combination of potential mediators that compete at the break. As a result, context-specific, lineage-driven, and stochastic effects may each influence which factors ultimately mediate Alt-NHEJ at a given DSB.Similar to HDR, the first step of Alt-NHEJ can involve resection of the DNA ends to single-strands by MRN and CtIP. During HDR, single-strand resection typically extends for kilobases but the extent of resection is limited in G0/G1 phases of the cell cycle by 53BP1. Thus, Alt-NHEJ occurring during G0/G1 involves short 3′ overhangs that may anneal at sites of microhomology (4). Previous studies with mammalian cells have described varying lengths of microhomology characteristic of Alt-NHEJ junctions [e.g., ≥2 bp (22), ≥3 bp (23), ≥5 bp (24), and 2 to 6 bp (25)], suggesting that there is no absolute requirement for a given length across contexts.After annealing, DNA polymerases and ligases are needed to fill the gaps before end ligation. The role of polymerases in Alt-NHEJ (and translocation formation) remains poorly understood. Pol θ is thought to play a role in Alt-NHEJ by facilitating DNA synthesis from annealed microhomologies, and loss of Pol θ led to reduced frequency of Cas9-induced translocations (17) and increased frequency of spontaneous IgH/Myc translocations in mouse cells (26). A recent study showed that loss of Pol β can also reduce translocation frequency between endonuclease-induced breaks in human cells (27).Translocation junctions in mammalian cells tend to have longer deletions and increased use of microhomology compared to repair at single DSBs, suggesting that Alt-NHEJ is involved. Initial studies on the mechanisms of translocation formation were performed in mouse embryonic stem cells (mESCs) containing a translocation reporter that reconstitutes a neomycin resistance gene after cleavage of chromosomes 14 and 17 by the I-SceI meganuclease (22, 28, 29). mESCs lacking either Ku or Xrcc4/Lig4 had increased translocation frequencies, suggesting that these factors suppress translocations rather than promoting them (28). Furthermore, in the absence of C-NHEJ factors, translocation junctions contained longer deletions and an increased usage of microhomology in the final repair products, consistent with Alt-NHEJ mediating translocations in these cells. Additional studies in mESCs demonstrated that loss of CtIP, Parp1, Lig3, and Lig1 can each result in reduced translocation frequency, further implicating these factors in the Alt-NHEJ that mediates translocation formation (22, 29).A subsequent study in human cells painted a more complex picture. Human cell lines depleted of LIG4 had reduced translocation frequency and depletion of LIG3 only decreased translocations in LIG4-depleted cells (23). The mechanisms and cell type-specificity behind these differential end-joining requirements for translocation formation in mouse and human cells remain poorly defined (23).We previously reported a screen of short hairpin RNA (shRNA) against 169 DNA repair-related genes in human cells to identify factors that modulate translocation frequency (30). We used stringent criteria to define “hits” and validated each in multiple human cell lines. Knockdown of the SUMO E2 enzyme UBC9 or RAD50 increased translocation frequency, so we categorized these factors as translocation suppressors. Conversely, knockdown of 53BP1, DNA damage-binding protein 1 (DDB1), or PARP3 decreased translocation frequency, so we categorized these as translocation promoters (30).We noted that POLD2, an accessory subunit of the replicative polymerase Pol δ, nearly scored in the screen as a promoter of chromosomal translocations. In budding yeast, Pol δ promotes both Alt-NHEJ and microhomology-mediated chromosomal translocations (31) but this has not been assessed in mammalian cells. Here, we demonstrate that Pol δ plays a role during Alt-NHEJ in human cells and that Pol δ subunits promote translocations. Based on these findings, we propose a model in which Pol δ exonuclease and polymerase activity promote Alt-NHEJ after annealing of sequences with microhomology.     

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.     

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.     

Sequence of the supernumerary B chromosome of maize provides insight into its drive mechanism and evolution

B chromosomes are enigmatic elements in thousands of plant and animal genomes that persist in populations despite being nonessential. They circumvent the laws of Mendelian inheritance but the molecular mechanisms underlying this behavior remain unknown. Here we present the sequence, annotation, and analysis of the maize B chromosome providing insight into its drive mechanism. The sequence assembly reveals detailed locations of the elements involved with the cis and trans functions of its drive mechanism, consisting of nondisjunction at the second pollen mitosis and preferential fertilization of the egg by the B-containing sperm. We identified 758 protein-coding genes in 125.9 Mb of B chromosome sequence, of which at least 88 are expressed. Our results demonstrate that transposable elements in the B chromosome are shared with the standard A chromosome set but multiple lines of evidence fail to detect a syntenic genic region in the A chromosomes, suggesting a distant origin. The current gene content is a result of continuous transfer from the A chromosomal complement over an extended evolutionary time with subsequent degradation but with selection for maintenance of this nonvital chromosome.

Supernumerary chromosomes were first discovered in the leaf-footed plant bug Metapodius more than a century ago (1). Since then, they have been reported in numerous plant, animal, and fungal species (2). A common feature of these so-called B chromosomes is that they are nonessential and are present only in some individuals in the population of a particular species. Through their evolution, they have developed various modes of behavior, e.g., tissue-specific elimination in Aegilops (3), preferential fertilization in Zea (4), or sex manipulation in Nasonia (5). In many plant species, they undergo controlled nondisjunction—unequal allocation to daughter nuclei during postmeiotic divisions (6). Their effect on frequency and distribution of meiotic crossovers along the standard A chromosomes has also been described (7, 8). Despite the peculiar behavior and unclear origins, no high-quality B chromosome reference sequence has been previously obtained in any organism.The B chromosome of maize is one of the most thoroughly studied supernumerary chromosomes (9–11) (Fig. 1). It can be found in numerous landraces and also in populations of Mexican teosinte, the maize wild relative (12). Despite being dispensable, it is maintained in populations by two properties: nondisjunction at the second pollen mitosis giving rise to unequal sperm and then preferential fertilization of the egg by the B chromosome-containing sperm (4, 13) (Fig. 1). This acrocentric chromosome is smaller than standard A chromosomes. Its long arm comprises proximal (PE) and distal euchromatin (DE), proximal heterochromatin (PH), and four large distal blocks of heterochromatin (DH1-4) (Fig. 1). Its short arm is minute. In a majority of genetic backgrounds, the presence of B chromosomes is not detrimental unless at high copy number (10), but in some others will cause breakage at the second pollen mitosis of some A chromosomes that contain heterochromatic knobs (14). This effect is thought to be an extension of the B chromosome drive mechanism involved with nondisjunction at this particular mitosis. The B chromosome has also evolved the property of increasing recombination in heterochromatic regions in general, probably to ensure its own chiasmata for proper distribution in meiosis (7, 15–17). Further, it has acquired a mechanism for its faithful transmission as a univalent (18, 19), which would also enhance its transmission, given that it can sometimes be present in odd numbers. Thus, classical cytogenetic studies established multiple properties of this unusual chromosome that act to maintain it in populations, but the molecular basis of its non-Mendelian inheritance remained obscure. To date, the efforts to generate B-specific sequences in maize have been limited to DNA marker development, general characterization of repetitive sequences, and low-pass sequencing (20–23). While genomes of several maize lines were sequenced to reference quality (24–26), comparable information for the supernumerary chromosome has not yet been available.Open in a separate windowFig. 1.The maize B chromosome. (A) Root tip metaphase spread of a line possessing nine B chromosomes (red signal). The red signal identifies the ZmBs B chromosome repeat in and around the centromere with a minor representative at the distal tip of the B long arm. Green signal identifies several chromosomal features, namely, the CentC centromeric satellite, 45S ribosomal DNA repeats, and TAG microsatellite clusters. DAPI stains the chromosomes (blue). (B) Schematic view of the acrocentric maize B chromosome at pachynema of meiosis. The chromosome is divided into the B short arm (BS), B centromere (BC), proximal heterochromatin (PH), proximal euchromatin (PE), four blocks of distal heterochromatin (DH1-4), and the distal euchromatin (DE). The B-specific repeat ZmBs, CentC satellite, CRM2 retrotransposon, knob heterochromatin, and TAG microsatellite cluster are color coded along the length of the chromosome. (C) Depiction of nondisjunction of the B chromosome. The B chromosome (blue with a red centromere) is shown in the generative nucleus (G) after the first microspore division. After replication, the two chromatids proceed to the same pole at the second microspore mitosis in the vast majority of divisions. Thus, most mature pollen grains contain two sperm (S) with only one containing the B chromosomes. V: vegetative cell. (D) Depiction of preferential fertilization. For most lines of maize, the sperm with the two B chromosomes will preferentially fertilize the egg (E) as compared with the central cell (C) in the process of double fertilization. The fertilized egg develops into the next generation embryo and the fertilized central cell develops into the endosperm. The combination of nondisjunction at the second pollen mitosis and preferential fertilization comprise the drive mechanism of the B chromosome.     

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.     

ATM controls the extent of DNA end resection by eliciting sequential posttranslational modifications of CtIP

DNA end resection is a critical step in the repair of DNA double-strand breaks (DSBs) via homologous recombination (HR). However, the mechanisms governing the extent of resection at DSB sites undergoing homology-directed repair remain unclear. Here, we show that, upon DSB induction, the key resection factor CtIP is modified by the ubiquitin-like protein SUMO at lysine 578 in a PIAS4-dependent manner. CtIP SUMOylation occurs on damaged chromatin and requires prior hyperphosphorylation by the ATM protein kinase. SUMO-modified hyperphosphorylated CtIP is targeted by the SUMO-dependent E3 ubiquitin ligase RNF4 for polyubiquitination and subsequent degradation. Consequently, disruption of CtIP SUMOylation results in aberrant accumulation of CtIP at DSBs, which, in turn, causes uncontrolled excessive resection, defective HR, and increased cellular sensitivity to DSB-inducing agents. These findings reveal a previously unidentified regulatory mechanism that regulates CtIP activity at DSBs and thus the extent of end resection via ATM-dependent sequential posttranslational modification of CtIP.

DNA double-strand breaks (DSBs) constitute one of the most severe forms of DNA damage and can result in a wide variety of genetic alterations including mutations, deletions, translocations, and chromosome loss (1, 2). Extensive studies have shown that DSBs can be repaired primarily via two pathways, classical nonhomologous end joining and homologous recombination (HR), both of which are highly conserved among all eukaryotes (3–5). Classical nonhomologous end joining, which directly rejoins the two broken ends of a DSB, occurs throughout interphase (3–5). In contrast, DSB repair by HR requires the presence of a sister chromatid and is therefore restricted to the late S and G2 phases of the cell cycle (3–5). HR is initiated by resection of the 5′ strand of the DSB ends, yielding 3′ single-stranded DNA (ssDNA) tails that are initially coated with the replication protein A (RPA) complex (3–5). The resulting RPA-coated ssDNA is an essential intermediate not only in HR repair but also in ATR-CHK1 pathway activation (3–5). Studies conducted in yeast and mammalian cells have established that resection of DSB ends is a two-step process (3–5). First, the conserved MRE11/RAD50/NBS1 complex (MRE11/RAD50/XRS2 in Saccharomyces cerevisiae) cooperates with the key resection factor CtIP (Sae2 in S. cerevisiae, Ctp1 in Schizosaccharomyces pombe) to catalyze limited resection of broken DNA ends (3–6). Second, the resulting short 3′ overhangs are further processed through the action of either the 5′–3′ exonuclease EXO1 or the nuclease–helicase protein complex DNA2–BLM (DNA2–Sgs1 in S. cerevisiae) (3–5). Whereas extensive end resection is required for HR initiation and full checkpoint activation, uncontrolled and excessive processing of DSB ends can have deleterious consequences such as large deletions at DSB sites, persistent checkpoint signaling, and cell death (7–11). However, the mechanisms by which cells precisely control the extent of end resection at DSB sites undergoing homology-directed repair remain obscure.It has been well established that posttranslational modifications of DNA repair proteins play crucial roles in the cellular response to genotoxic stress (12, 13). For example, phosphorylation of CtIP at threonine 847 (serine 267 in Sae2) by CDK1/2 restricts its activity to the S and G2 phases of the cell cycle (14–17), and promotes its capacity to stimulate the MRE11 endonuclease activity (18) as well as the annealing of broken DNA ends (19). In addition, phosphorylation of CtIP at serine 327 by CDK2 and/or Aurora A is a prerequisite for its interactions with BRCA1 and PLK1 (20–23). Furthermore, CtIP undergoes phosphorylation at threonine 315 by CDK2, and this phosphorylation event regulates CtIP protein stability by facilitating its interaction with the phosphorylation-specific prolyl isomerase PIN1 (24). In addition to acting as a CDK substrate, CtIP can also be hyperphosphorylated by ATM (or ATR in Xenopus) at multiple serine/threonine–glutamine sites in response to DSBs, which is manifested by the appearance of a slow-migrating form of CtIP (25, 26). ATM-dependent hyperphosphorylation of CtIP not only facilitates its association with damaged DNA (27) but also promotes the recruitment of BLM and EXO1 to DSB sites (25). Moreover, a previous study showed that the putative ATM-targeted residues serine 231, serine 664, and serine 745 as well as the CDK-targeted residues serine 276, threonine 315, and serine 347 within CtIP are critical for its endonuclease activity, although the relative contributions of the individual modifications have not been fully characterized (28). In addition to phosphorylation, CtIP is also subject to other posttranslational modifications, such as ubiquitination and acetylation (21, 29–36). Strict regulation of CtIP activity via various posttranslational modifications is crucial for accurate processing and repair of DSBs; however, precisely how these modifications are regulated in a coordinated manner remains unclear.In this study, we provide evidence that CtIP becomes SUMOylated primarily at lysine 578 upon exposure to DSB-inducing agents, and that this modification controls the activated CtIP level at DSBs and thereby the extent of DSB end resection. CtIP SUMOylation at lysine 578 is dependent on its prior hyperphosphorylation by the protein kinase ATM. SUMO-modified hyperphosphorylated CtIP can be targeted by the SUMO-dependent E3 ubiquitin ligase RNF4 for polyubiquitination and subsequent degradation. As a consequence, cells expressing non-SUMOylatable CtIP mutants exhibit aberrant accumulation of CtIP at DSB sites, uncontrolled excessive end resection, and defective HR. Our results suggest that active CtIP triggers its own SUMOylation and degradation, establishing a negative feedback loop that restricts CtIP activity at DSBs and thereby prevents excessive end resection and genome instability.     

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.