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.     

Identifying hydrophobic protein patches to inform protein interaction interfaces

Interactions between proteins lie at the heart of numerous biological processes and are essential for the proper functioning of the cell. Although the importance of hydrophobic residues in driving protein interactions is universally accepted, a characterization of protein hydrophobicity, which informs its interactions, has remained elusive. The challenge lies in capturing the collective response of the protein hydration waters to the nanoscale chemical and topographical protein patterns, which determine protein hydrophobicity. To address this challenge, here, we employ specialized molecular simulations wherein water molecules are systematically displaced from the protein hydration shell; by identifying protein regions that relinquish their waters more readily than others, we are then able to uncover the most hydrophobic protein patches. Surprisingly, such patches contain a large fraction of polar/charged atoms and have chemical compositions that are similar to the more hydrophilic protein patches. Importantly, we also find a striking correspondence between the most hydrophobic protein patches and regions that mediate protein interactions. Our work thus establishes a computational framework for characterizing the emergent hydrophobicity of amphiphilic solutes, such as proteins, which display nanoscale heterogeneity, and for uncovering their interaction interfaces.

Protein–protein interactions play a crucial role in numerous biological processes, ranging from signal transduction and immune response to protein aggregation and phase behavior (1–3). Consequently, being able to understand, predict, and modulate protein interactions has important implications for understanding cellular processes and mitigating the progression of disease (4, 5). A necessary first step toward this ambitious goal is uncovering the interfaces through which proteins interact (6–8). In principle, identifying hydrophobic protein regions, which interact weakly with water, should be a promising strategy for uncovering protein interaction interfaces (9, 10). Indeed, the release of weakly interacting hydration waters from hydrophobic regions can drive protein interactions, as well as other aqueous assemblies (11–13). However, even when the structure of a protein is available at atomistic resolution, it is challenging to identify its hydrophobic patches because they are not uniformly nonpolar, but display variations in polarity and charge at the nanoscale. Moreover, the emergent hydrophobicity of a protein patch stems from the collective response of protein hydration waters to the nanoscale chemical and topographical patterns displayed by the patch (14–20) and cannot be captured by simply counting the number of nonpolar groups in the patch, or even through more involved additive approaches, such as hydropathy scales or surface-area models (21–28).To address this challenge, we build upon seminal theoretical advances and molecular simulation studies, which have shown that near a hydrophobic surface, it is easier to disrupt surface–water interactions and form interfacial cavities (29–34). To uncover protein regions that have the weakest interactions with water, here, we employ specialized molecular simulations, wherein protein–water interactions are disrupted by systematically displacing water molecules from the protein hydration shell (35–37). By identifying the protein patches that nucleate cavities most readily in our simulations, we are then able to uncover the most hydrophobic protein regions. Interestingly, we find that both hydrophobic and hydrophilic protein patches are highly heterogeneous and contain comparable numbers of nonpolar and polar atoms. Our results thus highlight the nontrivial relationship between the chemical composition of protein patches and their emergent hydrophobicity (24–26), and further emphasize the importance of accounting for the collective solvent response in characterizing protein hydrophobicity (16). We then interrogate whether the most hydrophobic protein patches, which nucleate cavities readily, are also likely to mediate protein interactions. Across five proteins that participate in either homodimer or heterodimer formation, we find that roughly 60 to 70% of interfacial contacts and only about 10 to 20% of noncontacts nucleate cavities. Our work thus provides a versatile computational framework for characterizing hydrophobicity and uncovering interaction interfaces of not just proteins, but also of other complex amphiphilic solutes, such as cavitands, dendrimers, and patchy nanoparticles (38–41).     

Prespacers formed during primed adaptation associate with the Cas1–Cas2 adaptation complex and the Cas3 interference nuclease–helicase

For Type I CRISPR-Cas systems, a mode of CRISPR adaptation named priming has been described. Priming allows specific and highly efficient acquisition of new spacers from DNA recognized (primed) by the Cascade-crRNA (CRISPR RNA) effector complex. Recognition of the priming protospacer by Cascade-crRNA serves as a signal for engaging the Cas3 nuclease–helicase required for both interference and primed adaptation, suggesting the existence of a primed adaptation complex (PAC) containing the Cas1–Cas2 adaptation integrase and Cas3. To detect this complex in vivo, we here performed chromatin immunoprecipitation with Cas3-specific and Cas1-specific antibodies using cells undergoing primed adaptation. We found that prespacers are bound by both Cas1 (presumably, as part of the Cas1–Cas2 integrase) and Cas3, implying direct physical association of the interference and adaptation machineries as part of PAC.

CRISPR-Cas systems of adaptive immunity provide prokaryotes with resistance against bacteriophages and plasmids (1–4). They consist of CRISPR DNA arrays and cas genes. Functionally, CRISPR defense can be subdivided into the interference and adaptation steps. The interference step involves specific recognition of regions in foreign nucleic acids, named protospacers, based on their complementarity to CRISPR arrays spacers followed by their destruction (5). The CRISPR adaptation step leads to integration of new spacers into the array (6, 7), forming inheritable memory that allows the entire lineage of cells derived from a founder that acquired a particular spacer to do away with genetic invaders carrying matching protospacers (8).Both interference and adaptation can be subdivided into multiple steps. For interference to occur, the CRISPR array is transcribed from a promoter located in the upstream leader region. The resulting pre-CRISPR RNA (pre-crRNA) is processed into short CRISPR RNAs (crRNAs), each containing a spacer flanked by repeat fragments (9). Individual crRNAs are bound by Cas proteins forming the effector complex, which is capable of recognizing sequences complementary to the spacer part of crRNA (10). Upon protospacer recognition, the target is destroyed either by a protein component of the effector complex or by additional recruitable Cas nucleases (3, 11–14). In a well-studied Type I-E CRISPR-Cas system of Escherichia coli, the effector comprises a multisubunit Cascade protein complex bound to a crRNA (11, 12, 15). The complementary interaction of Cascade-bound crRNA with a target protospacer leads to localized protospacer DNA melting and formation of an R-loop complex, where the crRNA spacer is annealed to the protospacer “target” strand, while the opposing “nontarget” strand is displaced and is present in a single-stranded form (16, 17). To avoid potentially suicidal recognition of CRISPR array spacers from which crRNAs originate, target recognition and R-loop complex formation require, in addition to complementarity with the crRNA spacer, the presence of a three-nucleotide long PAM (protospacer adjacent motif) preceding the protospacer (15, 18, 19). For E. coli type I-E system, the consensus PAM sequence is 5′-AAG-3′ on the nontarget strand. Some other trinucleotides also allow target recognition, though with decreased efficiency (15, 20). Below, we will refer to consensus PAM as “PAM^AAG.” The Cas3 nuclease-helicase is recruited to the R-loop complex and is responsible for target destruction (21–24). Cas3 first introduces a single-stranded break in the nontarget protospacer strand 11 to 15 nucleotides downstream of the PAM on the nontarget strand (25). Next, Cas3 unwinds and cleaves DNA in the 3′-5′ direction from the PAM (26–29). In vitro, Cas3-dependent degradation of DNA at the other side of the protospacer was also detected (16). Bidirectional Cas3-dependent degradation of DNA was also detected in vivo (30). The details of Cas3 “molecular gymnastics” required for such bidirectional destruction of DNA around the R-loop complex are not known.The main proteins of CRISPR adaptation are Cas1 and Cas2. In vitro, these proteins interact with each other, and the resulting complex is capable of inserting spacer-sized fragments in substrate DNA molecules containing at least one CRISPR repeat and a repeat-proximal leader region (31, 32). In the course of spacer integration, the Cas1–Cas2 complex first catalyzes a direct nucleophilic attack by the 3′-OH end of the incoming spacer at a phosphodiester bond between the leader and the first repeat in the top CRISPR strand (32, 33). This reaction proceeds via concurrent cleavage of the leader-repeat junction and covalent joining of one spacer strand to the 5′ end of the repeat. Subsequently, the 3′-OH on the second spacer strand attacks the phosphodiester bond at the repeat-spacer junction in the bottom CRISPR strand leading to full integration (32, 33). As a result, an intermediate with the newly incorporated spacer flanked by single-stranded repeat sequences is formed (32, 34). The gaps are filled in by a DNA polymerase, possibly DNA polymerase I (35).When overexpressed, E. coli Cas1 and Cas2 can integrate new spacers into the array in the absence of other Cas proteins (7, 36). During such “naive” adaptation, ∼50% of newly acquired spacers are selected from sequences flanked by the 5′-AAG-3′ trinucleotide, that is, consensus interference-proficient PAM^AAG. It thus follows that at least 50% of spacers acquired by Cas1 and Cas2 alone will be defensive during the interference step. The adaptation process must be tightly controlled, activated in the presence of the infecting mobile genetic elements, and directed toward foreign DNA, for otherwise, spacers acquired from host DNA will lead to suicidal self-interference. The details of the activation of CRISPR adaptation upon the entry of foreign DNA into the cell remain elusive. Some data indicate that active replication and/or a small size of phage or plasmid DNA may be responsible for a preferential selection of spacers from these molecules compared to selection of self-targeting spacers from host chromosomes (19). In addition, DNA repair/recombination signals present in host DNA, but lacking in foreign DNA may also increase the bias of the adaptation machinery to the latter (37).The bias of spacer acquisition machinery toward foreign DNA does not have to be significant, for acquisition of a self-targeting spacer by an infected cell will lead to the demise of such a cell in an act of altruism that would help control the spread of the infectious agent through the population. In contrast, acquisition of interference-proficient spacers from foreign DNA may allow the infected cell to survive, clear the infection, and endow its progeny with inheritable resistance—clearly an advantageous trait.To overcome CRISPR resistance, viruses and plasmids accumulate “escaper” mutations in the targeted protospacer or its PAM (36, 38). Given that the acquisition of protective spacers in infected cells is likely to be a rare event and the ease with which escaper mutations accumulate, the complex multistage CRISPR defense could become costly and ineffective (39). To increase the efficiency of CRISPR defense and counter the spread of mobile genetic elements with escaper mutations, CRISPR-Cas systems have evolved a specialized mode of spacer acquisition referred to as “primed adaptation” or “priming” (36, 40–47). Unlike the naive adaptation, in Type I CRISPR-Cas systems, priming requires, in addition to Cas1 and Cas2, a Cascade charged with crRNA recognizing the foreign target and the Cas3 nuclease–helicase. Spacers acquired during priming originate almost exclusively from DNA located in cis with the protospacer initially recognized by the effector complex (referred to hereafter as the “priming protospacer” or “PPS”). Furthermore, 90% or more of spacers acquired during priming by the I-E system of E. coli originate from protospacers with PAM^AAG and are therefore capable of efficient interference. Another hallmark of primed adaptation is the following: spacers acquired from DNA located at different sides of the PPS map to opposite DNA strands. The mapping of spacers acquired during naive adaptation shows no strand bias (48). Thus, the strand bias of spacers acquired during priming is probably related to Cas3 nuclease activity; however, exact details are lacking.The overall yield of spacers acquired during priming is increased when the PPS is imperfectly matched with a Cascade-bound crRNA spacer or when the PAM of the PPS is suboptimal (49). Thus, escaper protospacers serve as PPS, and priming initiated by inefficient recognition of such protospacers allows cells to quickly update their immunological memory by specific and efficient acquisition of additional interference-proficient spacers from mobile genetic elements that accumulated escaper mutations to earlier acquired spacers.The exact molecular mechanism of primed adaptation is not fully understood. Clearly, it should involve tight coordination between suboptimal interference against escaper targets and the spacer acquisition process. The DNA fragments produced by Cas3, a nuclease responsible for target degradation during interference, may feed primed adaptation, directly or indirectly, providing a functional link between the interference and adaptation arms of the CRISPR-Cas response. Based on results of in vitro experiments, it has been proposed that Cas3-generated degradation products may be used as substrates for the generation of prespacers (50)—DNA fragments that can be incorporated by the Cas1–Cas2 complex into arrays. However, no Cas3-generated products were detected in cells undergoing interference only, suggesting that Cas3 may degrade DNA to very short, subspacer length products (30). On the other hand, mutations abolishing the Cas3 nuclease activity lead to very little primed adaptation, indicating that priming requires the Cas3 nuclease activity (51). A possible way out from this impasse would be the existence of a “priming complex” that includes both Cas1–Cas2 and Cas3 and is responsible for the generation of prespacers by the Cas1–Cas2 complex from DNA along which Cas3 moves. Single-molecule analysis supports the existence of such a complex and even suggests that PPS-bound Cascade may be part of the priming complex (52). Here, we show that both Cas1–Cas2 and Cas3 associate with the same set of prespacers in cells undergoing primed adaptation, functionally linking CRISPR interference and adaptation machineries during priming. We also investigate the phenomenon of strand bias of spacer acquisition during priming and show that this bias does not depend on the orientation of PPS.     

Computational studies of anaplastic lymphoma kinase mutations reveal common mechanisms of oncogenic activation

Kinases play important roles in diverse cellular processes, including signaling, differentiation, proliferation, and metabolism. They are frequently mutated in cancer and are the targets of a large number of specific inhibitors. Surveys of cancer genome atlases reveal that kinase domains, which consist of 300 amino acids, can harbor numerous (150 to 200) single-point mutations across different patients in the same disease. This preponderance of mutations—some activating, some silent—in a known target protein make clinical decisions for enrolling patients in drug trials challenging since the relevance of the target and its drug sensitivity often depend on the mutational status in a given patient. We show through computational studies using molecular dynamics (MD) as well as enhanced sampling simulations that the experimentally determined activation status of a mutated kinase can be predicted effectively by identifying a hydrogen bonding fingerprint in the activation loop and the αC-helix regions, despite the fact that mutations in cancer patients occur throughout the kinase domain. In our study, we find that the predictive power of MD is superior to a purely data-driven machine learning model involving biochemical features that we implemented, even though MD utilized far fewer features (in fact, just one) in an unsupervised setting. Moreover, the MD results provide key insights into convergent mechanisms of activation, primarily involving differential stabilization of a hydrogen bond network that engages residues of the activation loop and αC-helix in the active-like conformation (in >70% of the mutations studied, regardless of the location of the mutation).

Neuroblastoma (NB) is the third most common cancer in children. Most NBs begin in sympathetic nerve ganglia in the abdomen—about half in the adrenal gland—and children with high-risk NB have a 5-y survival of only around 50%. These high-risk tumors are genomically and genetically heterogeneous, presenting with gene amplifications (mainly of the MYCN gene) and in some cases mutations in other genes—notably ALK (anaplastic lymphoma kinase), which encodes a receptor tyrosine kinase (RTK) (1, 2). Although germline ALK mutations in familial NB were reported first, somatic mutations were subsequently identified in patients, and the majority of all mutations occur in the cytoplasmic tyrosine kinase domain (TKD) of ALK (3). This discovery was important because aberrant kinase activity of the ALK TKD can be inhibited with existing drugs (3–6). Indeed, therapeutic targeting of ALK in other tumors such as non-small cell lung cancer (NSCLC), in which it is activated in an oncogenic fusion protein (7), has been successful. However, as shown for EGFR in NSCLC (8–10) and for ALK in earlier studies in NB, TKD mutations vary in the degree to which they activate the kinase—leading to oncogenesis—and in their effects on sensitivity to inhibition with small molecule inhibitors (5, 11, 12).We previously identified ALK mutations or amplifications in 14% of 1,600 patients with NB (11). Three hot spots in the ALK TKD (positions 1174, 1245, and 1275) account for 85% of kinase mutations, although mutations at numerous other sites have also been reported (10, 11, 13). These include clearly activating mutations, silent mutations (i.e., those shown not to be activating), and mutations that confer resistance to known ALK kinase inhibitors.A key challenge is to develop approaches for rapidly identifying which kinase domain mutations in such a list can be classified as cancer drivers (i.e., have an impact on cancer progression or treatment) and which are “passenger” mutations with no clinical consequence (14). Several approaches have been proposed for predicting and/or explaining the effects of mutations on kinase regulation. These include molecular dynamics (MD) simulations, structural bioinformatics methods based on evolutionary analyses, network analysis, and machine learning (ML) (15). The earliest attempts to understand how well sequence changes are tolerated were undertaken not in the context of cancer, but rather as efforts to understand evolutionary distances between sequences. These methods give probabilities of mutation based on phylogenetic trees (16) or sequence alignments (17), but were not designed to predict the effects of mutations on protein function. One of the earliest methods for predicting whether a mutation is deleterious is called Sorts Intolerant From Tolerant (SIFT), which uses sequence conservation to determine “deleteriousness” (18, 19) and remains a benchmark in the field of mutation classification. Several other algorithms have been developed that use sequence conservation or homology to predict the effects of single-nucleotide polymorphisms (SNPs) (20–24). In particular, PolyPhen-2 utilizes several sequence-based and structure-based features for the classification of driver versus passenger mutations arising from SNPs. Another approach (25) uses the mutation rate of noncoding genomic regions as a baseline and tries to identify genes in which there is a statistically significant deviation from this baseline. Several groups have also developed ML techniques to separate driver from passenger mutations. Methods used include random forest (26, 27), entropic methods (28), support vector machines (SVM) (15, 29, 30), graph/network analysis (31), and convolutional neural networks (32). A systematic assessment of the balanced accuracy of these methods is difficult to obtain as the published reports are applied across different datasets. However, a recent review of the predictive power of a subset of the methods outlined here concluded that MD-based and ML-based methods performed better in terms of balanced accuracies (15). MD methods in particular have the additional advantage over other predictive algorithms as they also provide a mechanistic (rather than only correlative) explanation for the results.Numerous groups have used MD simulations to assess the effects of mutations. MD simulations probe motions on the order of nanoseconds to microseconds, whereas catalysis by protein kinases takes place on the scale of milliseconds to seconds (11, 33–36). Careful analysis of simulation trajectories is therefore needed to gain insight into how mutations affect activity. These analyses can generally be fit into three broad categories (37): 1) analysis of alteration in structure- or energy-based functions, 2) analysis of collective motions, and 3) computation of free energy landscapes. The first category includes methods such as analysis of hydrogen bonds and salt bridges, changes in solvent accessible surface area (SASA), or of hydration dynamics. The second category includes measurements such as root mean squared deviation (RMSD) or fluctuation and calculations based on interatomic covariance matrices such as protein structure networks or principal component analyses. The third category includes a large and growing number of methods for understanding the energetic relationship between different conformational states of a protein. These methods generally rely on some prior knowledge of different conformational states of a protein (e.g., “active” and “inactive” conformations of a kinase) and apply some energetic potential to help the system explore desired states (38–42).To overcome the limitation of timescales accessible by MD, enhanced sampling methods that allow more rapid exploration of conformational space and determination of energy landscapes have been used on EGFR (43), ABL (44), ALK (45), B-RAF (46), CDK5 (47), insulin receptor kinase (39), c-KIT (48), HCK (49), RET and MET (44), and SRC (50, 51). Changes in hydrogen bonding networks, salt bridges, and hydrophobic interactions, which are easy to compute in MD simulations, have been used as proxies for comparing the stabilities of active and inactive conformations for a given mutated variant (11, 35). We therefore hypothesize that MD simulations can be utilized to classify activating and nonactivating mutations—based on which conformation they favor—balancing both accuracy and interpretability in computational analysis of cancer mutations.A key limitation of most previous studies is that they have either considered conformational changes only in the wild-type protein or have assessed only a handful of mutated proteins. Where MD has been applied as a predictive tool to classify mutations (11, 15), a key limitation is that any test set of mutations derived from a cancer study is imbalanced in terms of activating mutations (which dominate) and nonactivating mutations. Although upsampling techniques (15) can partially mitigate this issue, the optimal solution is to incorporate a balance between activating and nonactivating mutations in the study design. Here, we investigate the predictive power of MD and ML methods by carefully curating a list of 42 mutations in ALK from clinical data, the Catalogue of Somatic Mutations in Cancer (COSMIC) database (13), and additional “synthetic” test nonactivating mutations that we introduced to improve database balance and/or to address specific mechanistic questions.     

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.     

Real-time observation of Cas9 postcatalytic domain motions

CRISPR-Cas9 from Streptococcus pyogenes is an RNA-guided DNA endonuclease, which has become the most popular genome editing tool. Coordinated domain motions of Cas9 prior to DNA cleavage have been extensively characterized but our understanding of Cas9 conformations postcatalysis is limited. Because Cas9 can remain stably bound to the cleaved DNA for hours, its postcatalytic conformation may influence genome editing mechanisms. Here, we use single-molecule fluorescence resonance energy transfer to characterize the HNH domain motions of Cas9 that are coupled with cleavage activity of the target strand (TS) or nontarget strand (NTS) of DNA substrate. We reveal an NTS-cleavage-competent conformation following the HNH domain conformational activation. The 3′ flap generated by NTS cleavage can be rapidly digested by a 3′ to 5′ single-stranded DNA-specific exonuclease, indicating Cas9 exposes the 3′ flap for potential interaction with the DNA repair machinery. We find evidence that the HNH domain is highly flexible post-TS cleavage, explaining a recent observation that the HNH domain was not visible in a postcatalytic cryo-EM structure. Our results illuminate previously unappreciated regulatory roles of DNA cleavage activity on Cas9’s conformation and suggest possible biotechnological applications.

The CRISPR (clustered regularly interspaced short palindromic repeats)-Cas (CRISPR-associated) system targets foreign nucleic acids for destruction in bacteria and archaea (1). Among different types of the system, CRISPR-Cas9 from Streptococcus pyogenes has been widely used for genome editing in plant and animal cells (2–5). DNA cleavage by the CRISPR-Cas9 system involves multiple steps (6). The Cas9 enzyme associates with a guide RNA, consisting of a programmable CRISPR RNA (crRNA) and a transactivating RNA (tracrRNA), to form a Cas9 ribonucleoprotein complex (RNP) (7). The DNA substrate of Cas9 RNP contains a protospacer region complementary to the spacer sequence of crRNA, and a protospacer adjacent motif (PAM) (NGG for S. pyogenes Cas9) flanking the protospacer (7). After Cas9 RNP binding, the DNA is directionally unwound from the PAM-proximal region to the PAM-distal region (8–10), and the unwound target strand (TS) is hybridized to the spacer sequence of crRNA. The TS and nontarget strand (NTS) of the DNA are cleaved by the HNH domain (residue 780 to 906) and the RuvC domain (residues 1 to 56, 718 to 765, and 926 to 1,099), respectively (7, 11). Engineering of the active site of the HNH domain or RuvC domain creates NTS nickase (Cas9_dHNH, Cas9 with H840A mutation which cleaves NTS only) or TS nickase (Cas9_dRuvC, Cas9 with D10A mutation which cleaves TS only) (7). The TS and NTS Cas9 nickases have also been used in many genome editing applications to achieve higher editing specificity or avoid generating double-stranded breaks (12–14). Cas9 RNP remains stably bound to the cleaved DNA for hours in vitro (8, 9, 15, 16), likely hindering DNA repair processes in cells (15). Therefore, a characterization of the postcatalysis state of Cas9 and its nickase variants has the potential to provide insights into genome editing mechanisms.Fluorescence resonance energy transfer (FRET) and structural studies have demonstrated the key roles of the HNH domain conformation in Cas9-mediated DNA cleavage (17–24). The HNH domain undergoes large conformational changes from the “undocked” inactive conformations to the “docked” active conformation with respect to its TS substrate upon on-target DNA binding, which represents a conformational activation of the HNH domain preceding DNA cleavage (18, 19). The HNH activation also involves conformational changes of the REC2 domain (residue 167 to 307) and REC3 domain (residue 497 to 713) (25). The HNH domain was not visible in a recent cryo-EM structure of the “product” state Cas9-RNA-DNA complex, suggesting the HNH domain is flexible after DNA cleavage (26). However, the previous FRET study showed that the docked conformation persists after DNA cleavage and the product state HNH domain motions were not observed (except for a special case in which the 3′ flap of the NTS was completely removed after DNA cleavage) (18), possibly because the labeling positions for the FRET donor and acceptor pair were not sensitive to the conformational differences between the docked state and the product state of the HNH domain. In this study, we created new Cas9 FRET constructs with increased sensitivity to small distance changes between the FRET pair and observed postcatalytic HNH domain motions using single-molecule FRET.     

Spine dynamics of PSD-95-deficient neurons in the visual cortex link silent synapses to structural cortical plasticity

Critical periods (CPs) are time windows of heightened brain plasticity during which experience refines synaptic connections to achieve mature functionality. At glutamatergic synapses on dendritic spines of principal cortical neurons, the maturation is largely governed by postsynaptic density protein-95 (PSD-95)-dependent synaptic incorporation of α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) receptors into nascent AMPA-receptor silent synapses. Consequently, in mouse primary visual cortex (V1), impaired silent synapse maturation in PSD-95-deficient neurons prevents the closure of the CP for juvenile ocular dominance plasticity (jODP). A structural hallmark of jODP is increased spine elimination, induced by brief monocular deprivation (MD). However, it is unknown whether impaired silent synapse maturation facilitates spine elimination and also preserves juvenile structural plasticity. Using two-photon microscopy, we assessed spine dynamics in apical dendrites of layer 2/3 pyramidal neurons (PNs) in binocular V1 during ODP in awake adult mice. Under basal conditions, spine formation and elimination ratios were similar between PSD-95 knockout (KO) and wild-type (WT) mice. However, a brief MD affected spine dynamics only in KO mice, where MD doubled spine elimination, primarily affecting newly formed spines, and caused a net reduction in spine density similar to what has been observed during jODP in WT mice. A similar increase in spine elimination after MD occurred if PSD-95 was knocked down in single PNs of layer 2/3. Thus, structural plasticity is dictated cell autonomously by PSD-95 in vivo in awake mice. Loss of PSD-95 preserves hallmark features of spine dynamics in jODP into adulthood, revealing a functional link of PSD-95 for experience-dependent synapse maturation and stabilization during CPs.

Early life of an animal is characterized by time windows of functionally and structurally enhanced brain plasticity known as critical periods (CPs), which have been described initially in the primary visual cortex (V1) of kittens (1). During CPs, experience refines the connectivity of principal excitatory neurons to establish the mature functionality of neural networks. This refinement is governed by the constant generation and elimination of nascent synapses on dendritic spines that sample favorable connections to be consolidated and unfavorable ones to be eliminated (2–5). A fraction of nascent synapses is or becomes α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA)-receptor silent, expressing N-methyl-D-aspartate (NMDA) receptors only (6–8). At eye opening, silent synapses are abundant in the primary visual cortex (V1) (9, 10) and mature during CPs by stable AMPA receptor incorporation (11–14). The pace of silent synapse maturation is governed by the opposing yet cooperative function of postsynaptic density protein of 95 kDa (PSD-95) and its paralog PSD-93, two signaling scaffolds of the postsynaptic density of excitatory synapses (12, 13). However, whether silent synapses are preferential substrates for spine elimination during CPs remains to be investigated.In juvenile mice (postnatal days [P] 20 to 35), a brief monocular deprivation (MD) of the dominant contralateral eye results in a shift of the ocular dominance (OD) of binocular neurons in V1 toward the open eye, mediated by a reduction of responses to visual stimulation of the deprived eye (15–17). Structurally, MD induces an increase in spine elimination in apical dendrites of layer (L) 2/3 and L5 pyramidal neurons (PNs) which is only observed during the CP and constitutes a hallmark of juvenile OD plasticity (jODP) (18–20). After CP closure, cortical plasticity declines progressively, and in standard cage-raised mice beyond P40, a 4-d MD no longer induces the functional nor anatomical changes associated with jODP (21–24).At least three different mechanisms involved in experience-dependent maturation of cortical neural networks have been described, but the molecular and cellular mechanisms that cause CP closure remain highly debated (18, 25, 26). First, plasticity of local inhibitory neurons, such as increased inhibitory tone or a reduction of release probability by experience-dependent endocannabinoid receptor 1 (CB1R) activation was reported to close the critical period in rodent V1 (27–29). Second, the expression of so-called “plasticity brakes,” such as extracellular matrix (ECM), Nogo receptor 1 (NgR1), paired immunoglobulin-like receptor B (PirB), and Lynx1 were correlated with the end of critical periods (30–33). Experimentally decreasing the inhibitory tone or absence of plasticity brakes enhanced ODP expression in various knockout (KO) mouse models (32, 34, 35), among which only Lynx1 KO mice were shown to exhibit functional hallmarks of jODP, such as selective deprived eye depression after a short MD (36). Structurally, Lynx1 KO mice exhibited elevated spine dynamics at baseline; however, MD induced a reduction in spine elimination in apical dendrites of L5 PNs, whereas in L2/3 PNs there was no change (37). Thus, the effects of removing plasticity brakes on structural plasticity are variable, and it remains unclear to what extend manipulating the plasticity brakes can reinstate cellular signatures of CP plasticity in the adult wild-type (WT) brain (38). Third, the progressive maturation of AMPAR-silent synapses was correlated with the closure of the CP for jODP (12, 13). Consequently, in PSD-95 KO mice, the maturation of silent synapses is impaired; their fraction remains at the eye opening level, and jODP is preserved lifelong (13). Furthermore, visual cortex-specific knockdown (KD) of PSD-95 in the adult brain reinstated jODP. In contrast, in PSD-93 KO mice, silent synapses mature precociously and the CP for jODP closes precociously (12), correlating the presence of silent synapses with functional plasticity during CPs.While these three mechanisms of CP closure are not mutually exclusive in regulating cortical plasticity (26), it remains elusive whether CP-like structural plasticity can be expressed in the adult brain and whether silent synapses might be substrates for it. Here, we performed chronic two-photon imaging of dendrites of L2/3 pyramidal neurons in binocular V1 of PSD-95 KO (and KD) and WT mice, tracking the same dendritic spines longitudinally before, during, and after a 4-d period of MD. As previous studies have reported anesthesia effects on spine dynamics (39–41), we performed our experiments in awake mice, thoroughly trained for head fixation under the two-photon microscope. Our chronic spine imaging experiments revealed that in adult PSD-95 KO and KD mice, a brief MD indeed increased spine elimination about twofold, while adult WT mice did not display experience-dependent changes in spine elimination or spine formation. Thus, the loss of PSD-95 led to a high number of AMPAR-silent synapses which were correlated with jODP after MD, and with juvenile-like structural plasticity even in the adult brain, underscoring the importance of silent synapses for CP-timing and network maturation and stabilization.     

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.     

The KRAS and other prenylated polybasic domain membrane anchors recognize phosphatidylserine acyl chain structure

KRAS interacts with the inner leaflet of the plasma membrane (PM) using a hybrid anchor that comprises a lysine-rich polybasic domain (PBD) and a C-terminal farnesyl chain. Electrostatic interactions have been envisaged as the primary determinant of interactions between KRAS and membranes. Here, we integrated molecular dynamics (MD) simulations and superresolution spatial analysis in mammalian cells and systematically compared four equally charged KRAS anchors: the wild-type farnesyl hexa-lysine and engineered mutants comprising farnesyl hexa-arginine, geranylgeranyl hexa-lysine, and geranylgeranyl hexa-arginine. MD simulations show that these equally charged KRAS mutant anchors exhibit distinct interactions and packing patterns with different phosphatidylserine (PtdSer) species, indicating that prenylated PBD–bilayer interactions extend beyond electrostatics. Similar observations were apparent in intact cells, where each anchor exhibited binding specificities for PtdSer species with distinct acyl chain compositions. Acyl chain composition determined responsiveness of the spatial organization of different PtdSer species to diverse PM perturbations, including transmembrane potential, cholesterol depletion, and PM curvature. In consequence, the spatial organization and PM binding of each KRAS anchor precisely reflected the behavior of its preferred PtdSer ligand to these same PM perturbations. Taken together these results show that small GTPase PBD-prenyl anchors, such as that of KRAS, have the capacity to encode binding specificity for specific acyl chains as well as lipid headgroups, which allow differential responses to biophysical perturbations that may have biological and signaling consequences for the anchored GTPase.

KRAS4B (hereafter KRAS) is a lipid-anchored small GTPase that regulates multiple signaling pathways to control cell proliferation, survival, and migration (1, 2). KRAS is one of the most frequently mutated proteins in cancer, with mutations found in 98% of pancreatic tumors, 45% of colorectal tumors, and 31% of lung tumors (1, 2). KRAS signaling is mostly compartmentalized to the plasma membrane (PM) (3), where KRAS interacts with a specific set of lipids and undergoes spatial segregation to form nanometer-sized domains, termed nanoclusters (4, 5). Nanoclusters operate as transient platforms for KRAS signal transmission such that the extent of nanoclustering directly correlates with the efficiency of effector recruitment and MAPK signal output (6–8). KRAS effectors require synergistic binding with activated KRAS and specific lipids for efficient PM recruitment and activation (9), therefore concentrating a specific set of lipids within nanoclusters is essential to KRAS function (10, 11). In this context the KRAS C-terminal membrane anchor, which comprises a hexa-lysine polybasic domain (PBD) and a farnesylated, methylesterified cysteine residue (4, 5, 12, 13), selectively sorts the monovalent anionic phospholipid phosphatidylserine (PtdSer) into nanoclusters (7, 11, 14). In consequence, PtdSer levels in the PM modulate the extent of KRAS localization to, and nanoclustering on, the PM and hence regulate KRAS-dependent effector recruitment and signaling (7, 11, 14). Depletion of PtdSer compromises the proliferation of KRAS-driven cancer cell lines and KRAS oncogenicity in mouse xenograft models (15–20). Thus, PtdSer is a key structural component of KRAS signaling nanoclusters on the PM and plays important roles in KRAS function and pathology.Electrostatics have long been considered to be the primary determinant of interactions between anionic lipids and the PBD of KRAS; however, in biological membranes these interactions are more complex. For example, KRAS nanoclusters are enriched with monovalent PtdSer, but not multivalent phosphoinositol 4,5-bisphosphate (PIP₂) or phosphoinositol 3,4,5-trisphosphate (PIP₃) (7, 11, 14, 21). Moreover, the KRAS anchor selectively binds and sorts mixed-chain PtdSer species comprising one saturated and one unsaturated acyl chain (16:0/18:1 PtdSer and 18:0/18:1 PtdSer), but not symmetric PtdSer species comprising two identical saturated or unsaturated acyl chains (di18:0 PtdSer, di18:1 PtdSer or di18:2 PtdSer) (11, 21). Molecular dynamics (MD) simulations reveal that the KRAS PBD samples diverse conformational states on bilayers (11), including a pseudohelical hairpin with only its center portion inserted into the bilayer core (11, 22). These findings suggest that the KRAS PBD anchor interacts with membranes in complex manners that extend beyond electrostatics. Here, we formally examine this hypothesis by integrating atomistic MD simulations with quantitative electron microscopy (EM)-spatial analysis of intact PM. We show that the KRAS prenyl group and PBD sequence synergistically contribute to the structure and lipid-binding pattern of the anchor on membranes. Different combinations of PBD sequence and prenyl chain can be engineered to preferentially interact with PtdSer species that have different acyl chain structures. When grafted onto KRAS the PtdSer acyl chain binding preferences of these anchors result in fundamentally different responses to multiple biophysical perturbations of PM properties, including transmembrane potential (ΔV_m), cholesterol content, and membrane curvature. Together these results show that PBD prenyl anchors by recognizing phospholipid acyl chain structure link PM biophysics to small GTPase spatiotemporal organization and potentially biological function.     

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.