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.     

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.     

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.     

A limited role of NKCC1 in telencephalic glutamatergic neurons for developing hippocampal network dynamics and behavior

NKCC1 is the primary transporter mediating chloride uptake in immature principal neurons, but its role in the development of in vivo network dynamics and cognitive abilities remains unknown. Here, we address the function of NKCC1 in developing mice using electrophysiological, optical, and behavioral approaches. We report that NKCC1 deletion from telencephalic glutamatergic neurons decreases in vitro excitatory actions of γ-aminobutyric acid (GABA) and impairs neuronal synchrony in neonatal hippocampal brain slices. In vivo, it has a minor impact on correlated spontaneous activity in the hippocampus and does not affect network activity in the intact visual cortex. Moreover, long-term effects of the developmental NKCC1 deletion on synaptic maturation, network dynamics, and behavioral performance are subtle. Our data reveal a neural network function of NKCC1 in hippocampal glutamatergic neurons in vivo, but challenge the hypothesis that NKCC1 is essential for major aspects of hippocampal development.

Intracellular chloride concentration ([Cl⁻]_i) is a major determinant of neuronal excitability, as synaptic inhibition is primarily mediated by chloride-permeable receptors (1). In the mature brain, [Cl⁻]_i is maintained at low levels by chloride extrusion, which renders γ-aminobutyric acid (GABA) hyperpolarizing (2) and counteracts activity-dependent chloride loads (3). GABAergic inhibition in the adult is crucial not only for preventing runaway excitation of glutamatergic cells (4) but also for entraining neuronal assemblies into oscillations underlying cognitive processing (5). However, the capacity of chloride extrusion is low during early brain development (6, 7). Additionally, immature neurons are equipped with chloride uptake mechanisms, particularly with the Na⁺/K⁺/2Cl⁻ cotransporter NKCC1 (8–12). NKCC1 contributes to the maintenance of high [Cl⁻]_i in the developing brain (13), favoring depolarization through GABA_A receptor (GABA_AR) activation in vivo (14, 15).When GABA acts as a depolarizing neurotransmitter, neural circuits generate burst-like spontaneous activity (16–20), which is crucial for their developmental refinement (21–24). In vitro evidence indicates that GABAergic interneurons promote neuronal synchrony in an NKCC1-dependent manner (10, 12, 25–28). However, the in vivo developmental functions of NKCC1 are far from understood (29, 30). One fundamental question is to what extent NKCC1 and GABAergic depolarization supports correlated spontaneous activity in the neonatal brain. In the neocortex, GABA imposes spatiotemporal inhibition on network activity already in the neonatal period (14, 25, 31, 32). Whether a similar situation applies to other brain regions is unknown, as two recent chemo- and optogenetic studies in the hippocampus yielded opposing results (25, 33). Manipulations of the chloride driving force are potentially suited to resolve these divergent findings, but pharmacological (34–36) or conventional knockout (10, 11, 37) strategies suffer from unspecific effects that complicate interpretations.Here, we overcome this limitation by selectively deleting Slc12a2 (encoding NKCC1) from telencephalic glutamatergic neurons. We show that chloride uptake via NKCC1 promotes synchronized activity in acute hippocampal slices, but has weak and event type-dependent effects in CA1 in vivo. Long-term loss of NKCC1 leads to subtle changes of network dynamics in the adult, leaving synaptic development unperturbed and behavioral performance intact. Our data suggest that NKCC1-dependent chloride uptake is largely dispensable for several key aspects of hippocampal development in vivo.     

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

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

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

CAP1 binds and activates adenylyl cyclase in mammalian cells

CAP1 (Cyclase-Associated Protein 1) is highly conserved in evolution. Originally identified in yeast as a bifunctional protein involved in Ras-adenylyl cyclase and F-actin dynamics regulation, the adenylyl cyclase component seems to be lost in mammalian cells. Prompted by our recent identification of the Ras-like small GTPase Rap1 as a GTP-independent but geranylgeranyl-specific partner for CAP1, we hypothesized that CAP1-Rap1, similar to CAP-Ras-cyclase in yeast, might play a critical role in cAMP dynamics in mammalian cells. In this study, we report that CAP1 binds and activates mammalian adenylyl cyclase in vitro, modulates cAMP in live cells in a Rap1-dependent manner, and affects cAMP-dependent proliferation. Utilizing deletion and mutagenesis approaches, we mapped the interaction of CAP1-cyclase with CAP’s N-terminal domain involving critical leucine residues in the conserved RLE motifs and adenylyl cyclase’s conserved catalytic loops (e.g., C1a and/or C2a). When combined with a FRET-based cAMP sensor, CAP1 overexpression–knockdown strategies, and the use of constitutively active and negative regulators of Rap1, our studies highlight a critical role for CAP1-Rap1 in adenylyl cyclase regulation in live cells. Similarly, we show that CAP1 modulation significantly affected cAMP-mediated proliferation in an RLE motif–dependent manner. The combined study indicates that CAP1-cyclase-Rap1 represents a regulatory unit in cAMP dynamics and biology. Since Rap1 is an established downstream effector of cAMP, we advance the hypothesis that CAP1-cyclase-Rap1 represents a positive feedback loop that might be involved in cAMP microdomain establishment and localized signaling.

CAP/srv2 was originally identified in yeast biochemically as an adenylyl cyclase–associated protein (1) and genetically as a suppressor of the hyperactive Ras2-V19 allele (2). CAP/srv2-deficient yeast cells are unresponsive to active Ras2, and adenylyl cyclase activity is no longer regulated by Ras2 in these cells (1, 2), indicating the involvement of CAP/srv2 in the Ras/cyclase pathway. However, some mutant CAP/srv2 alleles presented phenotypes not observed in strains with impaired Ras/cyclase pathway (1–3), indicating the existence of Ras/cyclase-independent functions downstream of CAP/srv2. These two phenotype groups, that is, Ras/cyclase-linked and Ras/cyclase-independent, could be suppressed by expression of an N-terminal half and a C-terminal half of CAP/srv2, respectively (4). Subsequent studies showed that the C-terminal half of CAP/srv2 was able to bind monomeric G-actin (5–8) and other actin regulators establishing a role in F-actin dynamics (9–16). Thus, CAP/srv2 is a bifunctional protein with an N-terminal domain involved in Ras/cyclase regulation and a C-terminal domain involved with F-actin dynamics regulation (16–18).CAP1 is structurally conserved in all eukaryotes (18–22); however, their functions are not. Expression of the closely related Schizosaccharomyces pombe cap or mammalian CAP1 in yeast can only suppress the phenotypes associated with deletion of CAP/srv2’s C-terminal but not its N-terminal domain (19, 20, 22), suggesting that only the F-actin dynamics function was conserved while the Ras/cyclase regulation diverged early on in evolution (16–18). CAP/srv2’s N-terminal 1 to 36 domain was sufficient for cyclase binding in yeast involving a conserved RLE motif with predicted coiled-coil folding (23). Interestingly, this domain is also involved in CAP1 oligomerization both in yeast and mammalian cells (24–26), where it purifies as a high-molecular complex of ∼600 kDa consistent with a 1:1 stoichiometric CAP1-actin hexameric organization (12, 25, 27, 28). Importantly, removal of this domain disrupted CAP1 oligomerization, reduced F-actin turnover in vitro and caused defects in cell growth, cell morphology, and F-actin organization in vivo (24, 29). However, whether the conserved RLE motif in mammalian CAP1 interacts with other coiled-coil–containing proteins is for the moment unknown.Ras2-mediated cyclase regulation in yeast requires its farnesylation (30–32). However, the lipid target involved was not identified in the original studies. We have recently shown that mammalian CAP1 interacts with the small GTPase Rap1. The interaction involves Rap1’s C-terminal hypervariable region (HVR) and its lipid moiety in a geranylgeranyl-specific manner; that is, neither the closely related Ras1 nor engineered farnesylated Rap1 interacted with CAP1 (33). Thus, we raised the question whether CAP1-Rap1, similar to CAP/srv2-Ras2 in yeast, plays a role in cAMP dynamics in mammalian cells.In this study, we report that CAP1 binds to and activates mammalian adenylyl cyclase in vitro. The interaction involves CAP1’s conserved RLE motifs and cyclase’s conserved catalytic subdomains (e.g., C1a and/or C2a). Most importantly, we show that both CAP1 and Rap1 modulate cAMP dynamics in live cells and are critical players in cAMP-dependent proliferation.     

Structural insights into α-synuclein monomer–fibril interactions

Protein aggregation into amyloid fibrils is associated with multiple neurodegenerative diseases, including Parkinson’s disease. Kinetic data and biophysical characterization have shown that the secondary nucleation pathway highly accelerates aggregation via the absorption of monomeric protein on the surface of amyloid fibrils. Here, we used NMR and electron paramagnetic resonance spectroscopy to investigate the interaction of monomeric α-synuclein (α-Syn) with its fibrillar form. We demonstrate that α-Syn monomers interact transiently via their positively charged N terminus with the negatively charged flexible C-terminal ends of the fibrils. These intermolecular interactions reduce intramolecular contacts in monomeric α-Syn, yielding further unfolding of the partially collapsed intrinsically disordered states of α-Syn along with a possible increase in the local concentration of soluble α-Syn and alignment of individual monomers on the fibril surface. Our data indicate that intramolecular unfolding critically contributes to the aggregation kinetics of α-Syn during secondary nucleation.

Synucleinopathies, including Parkinson’s disease (PD), are associated with the accumulation of intracellular neuronal aggregates termed as Lewy bodies and Lewy neuritis, which contain high concentration of the protein α-synuclein (α-Syn) in an aggregated state (1, 2). The disease-relevant role of α-Syn is further highlighted by mutations in the α-Syn gene (SNCA) causing familial PD [i.e., A30P (3), E46K (4), H50Q (5), G51D (6), A53E (7), and A53T (8)] and the duplication or triplication of the SNCA leading to early-onset PD in affected families (9, 10). α-Syn is a 140-residue intrinsically disordered protein (IDP) in solution (11) but adopts a helical structure in the presence of acidic lipid surfaces (12, 13). The positively charged N terminus (residues 1 to 60) is rich in lysine residues and contains KTKEGV binding repeats associated with vesicle binding (14). Moreover, the N-terminal domain includes all known SNCA familial PD mutations. The central region (residues 61 to 95) defines the non-amyloid-β component (NAC) (15), which is essential for α-Syn aggregation (16), while the C terminus (residues 96 to 140) is highly negatively charged.In vitro, α-Syn forms polymorphic amyloid fibrils (17–19) with unique arrangements of cross-β-sheet motifs (20–22). When injected into model animals, these fibrils induce a PD-like pathology (23) where the aggregation pathway of α-Syn plays a key role in the development of the disease (24). A detailed analysis of the aggregation kinetics of α-Syn into amyloids is therefore important toward understanding the toxic mechanisms relevant for synucleinopathies.Amyloid formation of α-Syn is very sensitive to solution conditions, including pH (25), temperature (26), and salt concentration (27). It further requires the presence of an air–water interface (28) or negatively charged lipid membranes (29) for which α-Syn has a high affinity. Previous studies suggest that amyloid fibril growth of α-Syn occurs via a nucleation-dependent polymerization reaction (30). Following a fairly slow primary nucleus formation, α-Syn fibrils are elongated by addition of single monomers. In a next step, the amyloid fibrils multiply by fragmentation or can catalyze the formation of new amyloids from monomers on their surface—a process known as secondary nucleation that was first described for sickle cell anemia 40 y ago (31). Fragmentation and secondary nucleation critically depend on the fibril mass and accelerate the aggregation kinetics (30). In the case of α-Syn aggregation under quiescent condition fragmentation does not exist and only the described secondary nucleation process occurs. While detailed kinetic experiments showed no significant secondary nucleation at pH 7, it strongly contributes at pH values lower than 6 (25, 30). However, mechanistic or structural information of the secondary nucleation process in α-Syn aggregation has been lacking so far.In this study we investigated the structural properties of α-Syn monomer–fibril interactions by NMR and electron paramagnetic resonance (EPR) spectroscopy. Our results provide insights into how monomeric α-Syn transiently interacts in vitro via its positively charged N terminus with the negatively charged C-terminal residues of the α-Syn fibrils, giving detailed insights into the mechanism of the secondary nucleation process.     

Tropospheric carbonyl sulfide mass balance based on direct measurements of sulfur isotopes

Robust estimates for the rates and trends in terrestrial gross primary production (GPP; plant CO₂ uptake) are needed. Carbonyl sulfide (COS) is the major long-lived sulfur-bearing gas in the atmosphere and a promising proxy for GPP. Large uncertainties in estimating the relative magnitude of the COS sources and sinks limit this approach. Sulfur isotope measurements (³⁴S/³²S; δ³⁴S) have been suggested as a useful tool to constrain COS sources. Yet such measurements are currently scarce for the atmosphere and absent for the marine source and the plant sink, which are two main fluxes. Here we present sulfur isotopes measurements of marine and atmospheric COS, and of plant-uptake fractionation experiments. These measurements resulted in a complete data-based tropospheric COS isotopic mass balance, which allows improved partition of the sources. We found an isotopic (δ³⁴S ± SE) value of 13.9 ± 0.1‰ for the troposphere, with an isotopic seasonal cycle driven by plant uptake. This seasonality agrees with a fractionation of −1.9 ± 0.3‰ which we measured in plant-chamber experiments. Air samples with strong anthropogenic influence indicated an anthropogenic COS isotopic value of 8 ± 1‰. Samples of seawater-equilibrated-air indicate that the marine COS source has an isotopic value of 14.7 ± 1‰. Using our data-based mass balance, we constrained the relative contribution of the two main tropospheric COS sources resulting in 40 ± 17% for the anthropogenic source and 60 ± 20% for the oceanic source. This constraint is important for a better understanding of the global COS budget and its improved use for GPP determination.

The Earth system is going through rapid changes as the climate warms and CO₂ level rises. This rise in CO₂ is mitigated by plant uptake; hence, it is important to estimate global and regional photosynthesis rates and trends (1). Yet, robust tools for investigating these processes at a large scale are scarce (2). Recent studies suggest that carbonyl sulfide (COS) could provide an improved constraint on terrestrial photosynthesis (gross primary production, GPP) (2–12). COS is the major long-lived sulfur-bearing gas in the atmosphere and the main supplier of sulfur to the stratospheric sulfate aerosol layer (13), which exerts a cooling effect on the Earth’s surface and regulates stratospheric ozone chemistry (14).During terrestrial photosynthesis, COS diffuses into leaf stomata and is consumed by photosynthetic enzymes in a similar manner to CO₂ (3–5). Contrary to CO₂, COS undergoes rapid and irreversible hydrolysis mainly by the enzyme carbonic-anhydrase (6, 7). Thus, COS can be used as a proxy for the one-way flux of CO₂ removal from the atmosphere by terrestrial photosynthesis (2, 8–11). However, the large uncertainties in estimating the COS sources weaken this approach (10–12, 15). Tropospheric COS has two main sources: the oceans and anthropogenic emissions, and one main sink–terrestrial plant uptake (8, 10–13). Smaller sources include biomass burning, soil emissions, wetlands, volcanoes, and smaller sinks include OH destruction, stratospheric destruction, and soil uptake (12). The largest source of COS to the atmosphere is the ocean, both as direct COS emission, and as indirect carbon disulfide (CS₂) and dimethylsulfide (DMS) emissions that are rapidly oxidized to COS (10, 16–20). Recent studies suggest oceanic COS emissions are in the range of 200–4,000 GgS/y (19–22). The second major COS source is the anthropogenic source, which is dominated by indirect emissions derived from CS₂ oxidation, mainly from the use of CS₂ as an industrial solvent. Direct emissions of COS are mainly derived from coal and fuel combustion (17, 23, 24). Recent studies suggest that anthropogenic emissions are in the range of 150–585 GgS/y (23, 24). The terrestrial plant uptake is estimated to be in the range of 400–1,360 GgS/y (11). Measurements of sulfur isotope ratios (δ³⁴S) in COS may be used to track COS sources and thus reduce the uncertainties in their flux estimations (15, 25–27). However, the isotopic mass balance approach works best if the COS end members are directly measured and have a significantly different isotopic signature. Previous δ³⁴S measurements of atmospheric COS are scarce and there have been no direct measurements of two important components: the δ³⁴S of oceanic COS emissions, and the isotopic fractionation of COS during plant uptake (15, 25–27). In contrast to previous studies that used assessments for these isotopic values, our aim was to directly measure the isotopic values of these missing components, and to determine the tropospheric COS δ³⁴S variability over space and time.     

Neuromodulator release in neurons requires two functionally redundant calcium sensors

Neuropeptides and neurotrophic factors secreted from dense core vesicles (DCVs) control many brain functions, but the calcium sensors that trigger their secretion remain unknown. Here, we show that in mouse hippocampal neurons, DCV fusion is strongly and equally reduced in synaptotagmin-1 (Syt1)- or Syt7-deficient neurons, but combined Syt1/Syt7 deficiency did not reduce fusion further. Cross-rescue, expression of Syt1 in Syt7-deficient neurons, or vice versa, completely restored fusion. Hence, both sensors are rate limiting, operating in a single pathway. Overexpression of either sensor in wild-type neurons confirmed this and increased fusion. Syt1 traveled with DCVs and was present on fusing DCVs, but Syt7 supported fusion largely from other locations. Finally, the duration of single DCV fusion events was reduced in Syt1-deficient but not Syt7-deficient neurons. In conclusion, two functionally redundant calcium sensors drive neuromodulator secretion in an expression-dependent manner. In addition, Syt1 has a unique role in regulating fusion pore duration.

To date, over 100 genes encoding neuropeptides and neurotrophic factors, together referred to as neuromodulators, are identified, and most neurons express neuromodulators and neuromodulator receptors (1). Neuromodulators travel through neurons in dense core vesicles (DCVs) and, upon secretion, regulate neuronal excitability, synaptic plasticity, and neurite outgrowth (2–4). Dysregulation of DCV secretion is linked to many brain disorders (5–7). However, the molecular mechanisms that regulate neuromodulator secretion remain largely elusive.Neuromodulator secretion, like neurotransmitter secretion from synaptic vesicles (SVs), is tightly controlled by Ca²⁺. The Ca²⁺ sensors that regulate secretion have been described for other secretory pathways but not for DCV exocytosis in neurons. Synaptotagmin (Syt) and Doc2a/b are good candidate sensors due to their interaction with SNARE complexes, phospholipids, and Ca²⁺ (8–11). The Syt family consists of 17 paralogs (12, 13). Eight show Ca²⁺-dependent lipid binding: Syt1 to 3, Syt5 to 7, and Syt9 and 10 (14, 15). Syt1 mediates synchronous SV fusion (8), consistent with its low Ca²⁺-dependent lipid affinity (15, 16) and fast Ca²⁺/membrane dissociation kinetics (16, 17). Syt1 is also required for the fast fusion in chromaffin cells (18) and fast striatal dopamine release (19). Synaptotagmin-7 (Syt7), in contrast, drives asynchronous SV fusion (20), in line with its a higher Ca²⁺ affinity (15) and slower dissociation kinetics (16). Syt7 is also a major calcium sensor for neuroendocrine secretion (21) and secretion in pancreatic cells (22–24). Other sensors include Syt4, which negatively regulates brain-derived neurothropic factor (25) and oxytocin release (26), in line with its Ca²⁺ independency. Syt9 regulates hormone secretion in the anterior pituitary (27) and, together with Syt1, secretion from PC12 cells (28, 29). Syt10 controls growth factor secretion (30). However, Syt9 and Syt10 expression is highly restricted in the brain (31–33). Hence, the calcium sensors for neuronal DCV fusion remain largely elusive. Because DCVs are generally not located close to Ca²⁺ channels (34), we hypothesized that DCV fusion is triggered by high-affinity Ca²⁺ sensors. Because of their important roles in vesicle secretion, their Ca²⁺ binding ability, and their high expression levels in the brain (20, 31, 35–38), we addressed the roles of Doc2a/b, Syt1, and Syt7 in neuronal DCV fusion.In this study, we used primary Doc2a/b-, Syt1-, and Syt7-null (knockout, KO) neurons expressing DCV fusion reporters (34, 39–41) with single-vesicle resolution. We show that both Syt1 and Syt7, but not Doc2a/b, are required for ∼60 to 90% of DCV fusion events. Deficiency of both Syt1 and Syt7 did not produce an additive effect, suggesting they function in the same pathway. Syt1 overexpression (Syt1-OE) rescued DCV fusion in Syt7-null neurons, and vice versa, indicating that the two proteins compensate for each other in DCV secretion. Moreover, overexpression of Syt1 or Syt7 in wild-type (WT) neurons increased DCV fusion, suggesting they are both rate limiting for DCV secretion. We conclude that DCV fusion requires two calcium sensors, Syt1 and Syt7, that act in a single/serial pathway and that both sensors regulate fusion in a rate-limiting and dose-dependent manner.     

The intrinsic instability of the hydrolase domain of lipoprotein lipase facilitates its inactivation by ANGPTL4-catalyzed unfolding

The complex between lipoprotein lipase (LPL) and its endothelial receptor (GPIHBP1) is responsible for the lipolytic processing of triglyceride-rich lipoproteins (TRLs) along the capillary lumen, a physiologic process that releases lipid nutrients for vital organs such as heart and skeletal muscle. LPL activity is regulated in a tissue-specific manner by endogenous inhibitors (angiopoietin-like [ANGPTL] proteins 3, 4, and 8), but the molecular mechanisms are incompletely understood. ANGPTL4 catalyzes the inactivation of LPL monomers by triggering the irreversible unfolding of LPL’s α/β-hydrolase domain. Here, we show that this unfolding is initiated by the binding of ANGPTL4 to sequences near LPL’s catalytic site, including β2, β3–α3, and the lid. Using pulse-labeling hydrogen‒deuterium exchange mass spectrometry, we found that ANGPTL4 binding initiates conformational changes that are nucleated on β3–α3 and progress to β5 and β4–α4, ultimately leading to the irreversible unfolding of regions that form LPL’s catalytic pocket. LPL unfolding is context dependent and varies with the thermal stability of LPL’s α/β-hydrolase domain (T_m of 34.8 °C). GPIHBP1 binding dramatically increases LPL stability (T_m of 57.6 °C), while ANGPTL4 lowers the onset of LPL unfolding by ∼20 °C, both for LPL and LPL•GPIHBP1 complexes. These observations explain why the binding of GPIHBP1 to LPL retards the kinetics of ANGPTL4-mediated LPL inactivation at 37 °C but does not fully suppress inactivation. The allosteric mechanism by which ANGPTL4 catalyzes the irreversible unfolding and inactivation of LPL is an unprecedented pathway for regulating intravascular lipid metabolism.

The lipolytic processing of triglyceride-rich lipoproteins (TRLs) along the luminal surface of capillaries plays an important role in the delivery of lipid nutrients to vital tissues (e.g., heart, skeletal muscle, adipose tissue). A complex of lipoprotein lipase (LPL) and its endothelial cell transporter, glycosylphosphatidylinositol-anchored high-density lipoprotein-binding protein 1 (GPIHBP1), is responsible for the margination of TRLs and their lipolytic processing (1–3). The importance of the LPL•GPIHBP1 complex for TRL processing is underscored by the development of severe hypertriglyceridemia (chylomicronemia) with loss-of-function mutations in LPL or GPIHBP1 or with GPIHBP1 autoantibodies that disrupt GPIHBP1•LPL interactions (4–7). Chylomicronemia is associated with a high risk for acute pancreatitis, which is debilitating and often life threatening (8, 9). Interestingly, increased efficiency of plasma triglyceride processing appears to be beneficial, reducing both plasma triglyceride levels and the risk for coronary heart disease (CHD). For example, genome-wide population studies have revealed that single-nucleotide polymorphisms that limit the ability of angiopoietin-like proteins 3 or 4 (ANGPTLs) to inhibit LPL are associated with lower plasma triglyceride levels and a reduced risk of CHD (10–14).GPIHBP1 is an atypical member of the LU domain superfamily because it contains a long intrinsically disordered and highly acidic N-terminal extension in addition to a canonical disulfide-rich three-fingered LU domain (15). At the abluminal surface of capillaries, GPIHBP1 is responsible for capturing LPL from heparan sulfate proteoglycans (HSPGs) in the subendothelial spaces and shuttling it to its site of action in the capillary lumen (3, 16). The capture of LPL from subendothelial HSPGs depends on electrostatic interactions with GPIHBP1’s intrinsically disordered acidic domain and stable hydrophobic interactions with GPIHBP1’s LU domain (15, 17, 18). In the setting of GPIHBP1 deficiency, LPL never reaches the capillary lumen and remains mislocalized, bound to HSPGs, in the subendothelial spaces. Aside from promoting the formation of GPIHBP1•LPL complexes, the acidic domain plays an important role in preserving LPL activity. The acidic domain is positioned to form a fuzzy complex with a large basic patch on the surface of LPL, which is formed by the confluence of several heparin-binding motifs. This electrostatic interaction stabilizes LPL structure and activity, even in the face of physiologic inhibitors of LPL (e.g., ANGPTL4) (19–22).Distinct expression profiles for ANGPTL-3, -4, and -8 underlie the tissue-specific regulation of LPL and serve to match the supply of lipoprotein-derived lipid nutrients to the metabolic demands of nearby tissues (21, 23–29). In the fasted state, ANGPTL4 inhibits LPL activity in adipose tissue, resulting in increased delivery of lipid nutrients to oxidative tissues. In the fed state, ANGPTL3•ANGPTL8 complexes inhibit LPL activity in oxidative tissues and thereby channel lipid delivery to adipocytes. While the physiologic relevance of tissue-specific LPL regulation is clear, the mechanisms by which ANGPTL proteins inhibit LPL activity remain both incompletely understood and controversial. One view holds that ANGPTL4 inhibits LPL activity by a reversible mechanism (30, 31). An opposing view, formulated early on by the laboratory of Gunilla Olivecrona, is that ANGPTL4 irreversibly inhibits LPL by a “molecular unfolding chaperone-like mechanism” (32). Hydrogen–deuterium exchange mass spectrometry (HDX-MS) studies have supported the latter view. Recent studies by our group revealed that ANGPTL4 catalyzes the irreversible unfolding (and inactivation) of LPL’s α/β-hydrolase domain and that the unfolding is substantially mitigated by the binding of GPIHBP1 to LPL (17, 19, 22). We further showed that ANGPTL4 functions by unfolding catalytically active LPL monomers rather than by promoting the dissociation of catalytically active LPL homodimers (33, 34). Despite these newer findings, a host of issues remains unresolved. For example, the binding site for ANGPTL4 on LPL has been controversial (31, 35); the initial conformational changes induced by ANGPTL4 binding have not been delineated, and how the early conformational changes in LPL progress to irreversible inactivation is unknown. In the current studies, we show, using time-resolved HDX-MS, that ANGPTL4 binds to LPL sequences proximal to the entrance of LPL’s catalytic pocket. That binding event triggers alterations in the dynamics of LPL secondary structure elements that are central to the architecture of the catalytic triad. Progression of those conformational changes leads to irreversible unfolding and collapse of LPL’s catalytic pocket. The binding of GPIHBP1 to LPL limits the progression of these allosteric changes, explaining why GPIHBP1 protects LPL from ANGPTL4-induced inhibition.     

Dynamic competition between SARS-CoV-2 NSP1 and mRNA on the human ribosome inhibits translation initiation

Severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) is a beta-CoV that recently emerged as a human pathogen and is the causative agent of the COVID-19 pandemic. A molecular framework of how the virus manipulates host cellular machinery to facilitate infection remains unclear. Here, we focus on SARS-CoV-2 NSP1, which is proposed to be a virulence factor that inhibits protein synthesis by directly binding the human ribosome. We demonstrate biochemically that NSP1 inhibits translation of model human and SARS-CoV-2 messenger RNAs (mRNAs). NSP1 specifically binds to the small (40S) ribosomal subunit, which is required for translation inhibition. Using single-molecule fluorescence assays to monitor NSP1–40S subunit binding in real time, we determine that eukaryotic translation initiation factors (eIFs) allosterically modulate the interaction of NSP1 with ribosomal preinitiation complexes in the absence of mRNA. We further elucidate that NSP1 competes with RNA segments downstream of the start codon to bind the 40S subunit and that the protein is unable to associate rapidly with 80S ribosomes assembled on an mRNA. Collectively, our findings support a model where NSP1 proteins from viruses in at least two subgenera of beta-CoVs associate with the open head conformation of the 40S subunit to inhibit an early step of translation, by preventing accommodation of mRNA within the entry channel.

Beta-coronaviruses (CoVs) are a family of RNA viruses that include human pathogens (1). In the last two decades, two beta-CoVs have emerged from animal hosts to cause epidemic diseases of the human respiratory tract: severe acute respiratory syndrome (SARS-CoV, in 2002) (2, 3) and Middle East respiratory syndrome (MERS-CoV, in 2012) (4). A third beta-CoV emerged in late 2019—SARS-CoV-2—that is responsible for the ongoing COVID-19 pandemic (5). Given the lack of effective therapies against SARS-CoV-2, there is an urgent need for a molecular understanding of how the virus manipulates the machineries present in human cells.SARS-CoV-2 and the closely related SARS-CoV have single-stranded, positive-sense RNA genomes nearly 30 kb in length (6, 7). Upon entry of a virion into human cells, the genomic RNA is released into the cytoplasm where it must hijack human translation machinery to synthesize viral proteins (8). As the genomic RNA has a 7-methylguanosine (m⁷G) cap on the 5′ terminus, viral protein synthesis likely proceeds via a process reminiscent of that which occurs on typical human messenger RNAs (mRNAs) (9). However, as viral proteins accumulate, human translation is inhibited and host mRNAs are destabilized, which facilitates suppression of the host immune response (10–13).Studies on SARS-CoV have implicated nonstructural protein 1 (NSP1), the first encoded viral protein, as a virulence factor with a key role in the shutdown of host translation (10, 11, 14). In infected cells or upon its ectopic expression, NSP1 inhibits human translation, which is dependent on its association with the small (40S) subunit of the human ribosome (12–17). In a linked but separable activity, NSP1 destabilizes at least a subset of human mRNAs, likely via recruitment of an unidentified human endonuclease (12, 13, 15, 16, 18). NSP1 from SARS-CoV-2 is expected to employ similar mechanisms, given its ∼85% sequence identity with the SARS-CoV protein. Indeed, SARS-CoV-2 NSP1 inhibits translation by binding to the 40S subunit (19–21). Thus, NSP1 has a near-singular ability to disrupt host gene expression dramatically; yet, the mechanism by which this inhibition occurs is not clear.The 40S subunit is the nexus of translation initiation, recruiting an m⁷G-capped mRNA through a multistep, eukaryotic initiation factor (eIF)-mediated process. Prior to recruitment of an mRNA, the 40S subunit is bound by numerous eIFs, which include eIF1, eIF1A, eIF3, eIF5, and the ternary complex (TC) of eIF2–GTP–methionine initiator transfer RNA (tRNA_i^Met) (22). The eIFs make extensive contacts with the 40S subunit, including the ribosomal A and P sites (23, 24). They also manipulate the dynamics of the 40S head region to facilitate mRNA recruitment, which has structural consequences at both the mRNA entry (3′ side of mRNA) and exit (5′ end of mRNA) channels. Following mRNA recruitment and directional scanning of the 5′ untranslated region (UTR) to a start codon, a series of compositional and conformational changes occur (25, 26). This ultimately repositions the 40S subunit head into the closed conformation and accommodates the anticodon stem loop of the initiator tRNA at the start codon (23–26), enabling recruitment of the 60S subunit and entry into the elongation phase.Recently, structures of NSP1 bound to human ribosomes have been reported (19–21), including 40S preinitiation and 80S ribosomal complexes. In all instances, the N-terminal globular domain of NSP1 is flexibly localized to the solvent-exposed surface of the 40S subunit, near the entrance to the mRNA entry channel (SI Appendix, Fig. S1A). This domain is anchored by the two most C-terminal α-helices of NSP1, which were dynamic and unstructured in the free SARS-CoV NSP1 structure solved by NMR (27); in the NSP1–40S subunit complex, these helices were well resolved and docked within the mRNA entry channel, where they contact ribosomal proteins uS3 and uS5, and helix 18 of the 18S ribosomal RNA (rRNA). As noted above, this location on the ribosome is structurally flexible, adopting open and closed states upon swiveling of the 40S subunit head (23–25). The position of NSP1 in the mRNA channel may also conflict with the position of fully accommodated mRNA. Thus, the intrinsic dynamics of translation initiation present many opportunities and obstacles for NSP1 association with the ribosome, and its subsequent inhibition of translation.Here, we merge biochemical and single-molecule approaches to probe the molecular function of SARS-CoV-2 NSP1 and its interaction with the human ribosome. We showed that NSP1 potently inhibited translation of human and SARS-CoV-2 model mRNAs, and determined how the NSP1–40S subunit interaction was modulated by eIFs and mRNA. Our results reveal allosteric control of NSP1 association by key eIFs and identify a conformation of the ribosomal subunit compatible with rapid NSP1 association. They also define the dynamic competition between NSP1 and mRNA to bind the ribosome. When synthesized with recent structures, our study suggests a mechanism for how NSP1 inhibits translation initiation.     

Molecular basis of the dual role of the Mlh1-Mlh3 endonuclease in MMR and in meiotic crossover formation

In budding yeast, the MutL homolog heterodimer Mlh1-Mlh3 (MutLγ) plays a central role in the formation of meiotic crossovers. It is also involved in the repair of a subset of mismatches besides the main mismatch repair (MMR) endonuclease Mlh1-Pms1 (MutLα). The heterodimer interface and endonuclease sites of MutLγ and MutLα are located in their C-terminal domain (CTD). The molecular basis of MutLγ’s dual roles in MMR and meiosis is not known. To better understand the specificity of MutLγ, we characterized the crystal structure of Saccharomyces cerevisiae MutLγ(CTD). Although MutLγ(CTD) presents overall similarities with MutLα(CTD), it harbors some rearrangement of the surface surrounding the active site, which indicates altered substrate preference. The last amino acids of Mlh1 participate in the Mlh3 endonuclease site as previously reported for Pms1. We characterized mlh1 alleles and showed a critical role of this Mlh1 extreme C terminus both in MMR and in meiotic recombination. We showed that the MutLγ(CTD) preferentially binds Holliday junctions, contrary to MutLα(CTD). We characterized Mlh3 positions on the N-terminal domain (NTD) and CTD that could contribute to the positioning of the NTD close to the CTD in the context of the full-length MutLγ. Finally, crystal packing revealed an assembly of MutLγ(CTD) molecules in filament structures. Mutation at the corresponding interfaces reduced crossover formation, suggesting that these superstructures may contribute to the oligomer formation proposed for MutLγ. This study defines clear divergent features between the MutL homologs and identifies, at the molecular level, their specialization toward MMR or meiotic recombination functions.

During the first meiotic division, in most organisms, each pair of homologous chromosomes (homologs) needs to experience at least one crossover to ensure their accurate segregation and increase genetic diversity of the progeny (1, 2). Crossovers are generated after programmed DNA double-strand break (DSB) formation, and their subsequent repair by homologous recombination (3). Failure to achieve at least one crossover per homolog pair results in aneuploid gametes. These dysfunctions are frequent causes of spontaneous miscarriages and birth defects in humans (1). DSBs are generated by the meiosis-specific Spo11 protein and resected to form 3′ single-stranded tails that are directed to invade and pair with an unbroken homologous template, preferentially on the homolog (3). Invasion intermediates are a substrate for DNA synthesis. After the capture of the second DSB end, a subset of the intermediates is converted into double Holliday junctions (dHJs), which in meiosis are primarily resolved into crossovers (4, 5). The remaining recombination intermediates are repaired as noncrossovers.MutLγ (Mlh1-Mlh3 in yeast and MLH1-MLH3 in human) is essential for the formation of meiotic crossovers in many organisms. The MutLγ heterodimer possesses, similarly to MutLα (Mlh1-Pms1 in yeast, MLH1-PMS2 in human), a latent endonuclease activity (6–10). It has been proposed that the MutLγ endonuclease activity catalyzes the resolution of the dHJ intermediates and promotes the formation of crossovers (8, 11). In agreement with this, Mlh1 and Mlh3 form foci on pachytene chromosomes in different organisms at future crossover sites (12–14). In yeast mlh3 mutants, the crossover rates are reduced to 50 to 70% of the wild-type level (8, 15–17). These mutants exhibit failure in chromosome disjunction and consequently a decrease of spore viability. In Mlh3^-/- mice, males and females present a crossover defect that leads to aneuploidy (18). In agreement with the proposed resolvase role of MutLγ, a mutant of the active site within the conserved DQHAX₂EX₄E motif of Saccharomyces cerevisiae Mlh3, D523N, results in a loss of activity and confers a phenotype similar to the mlh3∆ mutant with decreased crossover frequencies (8). A mutation in the endonuclease domain of mouse MLH3 leads to infertile males and strongly reduced crossover numbers, despite a correct loading of factors essential for crossover resolution (19). In budding yeast and mammals, MLH1 deletion is also associated with severe dysfunctions in meiosis due to crossover defects, combined with high genetic instability due to its additional central role with Pms1 (as part of the MutLα complex) in mutation avoidance (20).In budding yeast, the majority of meiotic crossovers are formed by MutLγ, requiring in addition other proteins including the ZMM proteins (Zip1-4, Spo16, Mer3, and Msh4-Msh5), Exo1, and the Sgs1-Top3-Rmi1 complex (11, 21). The ZMM factors are proposed to stabilize the recombination intermediates and protect them from the action of helicases (reviewed in ref. 22). The 5′-3′ exonuclease, Exo1, is important for crossover formation independently of its nuclease activity and likely acts as scaffold factor in particular through its interaction motif with Mlh1 (17). We recently reported that MutLγ-Exo1 associates with recombination intermediates, followed by direct Cdc5 recruitment that triggers MutLγ crossover activity (23). Despite its central role in the formation of crossovers, to date, the MutLγ endonuclease does not show any specific activity on either a single HJ or dHJ DNA substrate in vitro, in contrast to other resolvases or structure-specific endonucleases (24). Recently, it was shown that human MutLγ is part of an ensemble with MutSγ, EXO1, RFC, and PCNA that preferentially cleaves plasmid DNA containing a HJ, although it does not cleave symmetrically across the junction, as would be expected for a canonical resolvase (25, 26).MutLγ is also an accessory factor of the postreplicative mismatch repair (MMR). In S. cerevisiae, MutLα is the major MutL homolog involved in MMR. It is targeted to DNA mismatches by the MutS homologs and introduces DNA nicks to initiate the excision of the strand containing the mismatch. MutLα contains an endonuclease site formed by three conserved motifs in Pms1 and the last conserved amino acids of Mlh1 (6, 7, 27). Mutation of this active site (e.g., pms1E707K in yeast) is associated with high mutations rates. A minor role of MutLγ has been reported in the repair of a subset of DNA mismatches recognized by Msh2-Msh3 (28, 29). The third MutL heterodimer, Mlh1-Mlh2 (MLH1-PMS1 in mammals) or MutLβ, has no endonuclease motif or activity. In yeast, mlh2Δ cells present only a slight defect in the repair of a subset of frameshift mutations (30–32). Apart from this function in MMR, we recently identified an interaction of MutLβ with Mer3 helicase that limits gene conversion tract lengths (31). A major challenge is to further characterize the molecular basis of the specific interactions and regulations of the three MutL homologs with their DNA substrates and protein partners in MMR and in meiotic recombination.MutLγ and MutLα present an overall common organization with a N-terminal domain (NTD) bearing an ATPase function, connected through a long linker to a C-terminal domain (CTD). The CTD mediates the dimerization of the complexes (33, 34) and possesses the endonuclease site (6–8). Upon ADP and ATP binding, the NTD undergoes large asymmetric conformational changes that can position it into a close proximity to the CTD (35, 36). The heterodimers MutLα and MutLγ can slide on double-stranded DNA (dsDNA) with linear diffusion modes (37, 38) and control strand excision during MMR (39). However, both heterodimers differ in their DNA-binding properties. MutLα has very low affinity for short dsDNA in physiological salt concentrations and can bind cooperatively to long dsDNA (>200 bp), forming long continuous protein tracts (40). In contrast, MutLγ binds short-branched DNA substrates with a higher affinity (in the nanomolar range) and with a marked preference for HJs (9). MutLγ also binds, albeit more weakly, short and long dsDNA and is proposed to form oligomers on long dsDNA (9, 41). The Pms1 and Mlh3 subunits present a moderate sequence identity precluding correct modeling of MutLγ from the MutLα crystal structures and thus limiting our understanding of the molecular basis of the specificity of MutLα and MutLγ heterodimers in MMR and meiosis, respectively.Here, we present the crystal structure of the S. cerevisiae MutLγ(CTD), and we compare it with the three-dimensional structure of the S. cerevisiae MutLα(CTD) that we previously reported (27). We reveal differences between the two complexes with regard to the size of the heterodimerization interface, the regulatory domain position, and the shape of the cavity surrounding the endonuclease site. We characterize the role of the last amino acids of Mlh1 using mutant alleles and identify a central role of the last three conserved residues of Mlh1 in vivo for both MMR and meiotic recombination. We analyze the DNA-binding specificities of the MutLα(CTD) and Mut-Lγ(CTD) and compare to the properties of full-length proteins. We then identify positions in Mlh3(NTD) and Mlh3(CTD) that can participate to the positioning of the NTD close to the CTD and characterize the corresponding mutants in meiosis. Finally, we report a filament arrangement of the MutLγ(CTD) in the crystal in agreement with oligomers proposed in previous studies (25, 41), and we characterize an allele mutated in this oligomerization region.     

The high-affinity immunoglobulin receptor FcγRI potentiates HIV-1 neutralization via antibodies against the gp41 N-heptad repeat

The HIV-1 gp41 N-heptad repeat (NHR) region of the prehairpin intermediate, which is transiently exposed during HIV-1 viral membrane fusion, is a validated clinical target in humans and is inhibited by the Food and Drug Administration (FDA)-approved drug enfuvirtide. However, vaccine candidates targeting the NHR have yielded only modest neutralization activities in animals; this inhibition has been largely restricted to tier-1 viruses, which are most sensitive to neutralization by sera from HIV-1–infected individuals. Here, we show that the neutralization activity of the well-characterized NHR-targeting antibody D5 is potentiated >5,000-fold in TZM-bl cells expressing FcγRI compared with those without, resulting in neutralization of many tier-2 viruses (which are less susceptible to neutralization by sera from HIV-1–infected individuals and are the target of current antibody-based vaccine efforts). Further, antisera from guinea pigs immunized with the NHR-based vaccine candidate (ccIZN36)₃ neutralized tier-2 viruses from multiple clades in an FcγRI-dependent manner. As FcγRI is expressed on macrophages and dendritic cells, which are present at mucosal surfaces and are implicated in the early establishment of HIV-1 infection following sexual transmission, these results may be important in the development of a prophylactic HIV-1 vaccine.

Membrane fusion between HIV-1 and host cells is mediated by the viral envelope glycoprotein (Env), a trimer consisting of the gp120 and gp41 subunits. Upon interaction with cellular receptors, Env undergoes a dramatic conformational change and forms the prehairpin intermediate (PHI) (1–3), in which the fusion peptide region at the amino terminus of gp41 inserts into the cell membrane. In the PHI, the N-heptad repeat (NHR) region of gp41 is exposed and forms a stable, three-stranded α-helical coiled coil. Subsequently, the PHI resolves when the NHR and the C-heptad repeat (CHR) regions of gp41 associate to form a trimer-of-hairpins structure that brings the viral and cell membranes into proximity, facilitating membrane fusion (Fig. 1).Open in a separate windowFig. 1.HIV-1 membrane fusion. The surface protein of the HIV-1 envelope is composed of the gp120 and gp41 subunits. After Env binds to cell-surface receptors, gp41 inserts into the host cell membrane and undergoes a conformational change to form the prehairpin intermediate. The N-heptad repeat (orange) region of gp41 is exposed in the PHI and forms a three-stranded coiled coil. To complete viral fusion, the PHI resolves to a trimer-of-hairpins structure in which the C-heptad repeat (blue) adopts a helical conformation and binds the NHR region. Fusion inhibitors such as enfuvirtide bind the NHR, preventing viral fusion by inhibiting formation of the trimer of hairpins (1–3). The membrane-proximal external region (red) is located adjacent to the transmembrane (TM) region of gp41.The NHR region of the PHI is a validated therapeutic target in humans: the Food and Drug Administration (FDA)-approved drug enfuvirtide binds the NHR and inhibits viral entry into cells (4, 5). Various versions of the three-stranded coiled coil formed by the NHR have been created and used as vaccine candidates in animals (6–10). The neutralization potencies of these antisera, as well as those of anti-NHR monoclonal antibodies (mAbs) (11–15), are modest and mostly limited to HIV-1 isolates that are highly sensitive to antibody-mediated neutralization [commonly referred to as tier-1 viruses (16)]. These results have led to skepticism about the PHI as a vaccine target.Earlier studies showed that the neutralization activities of mAbs that bound another region of gp41, the membrane-proximal external region (MPER) (Fig. 1), were enhanced as much as 5,000-fold in cells expressing FcγRI (CD64) (17, 18), an integral membrane protein that binds the Fc portion of immunoglobulin G (IgG) molecules with high (nanomolar) affinity (19, 20). This effect was not attributed to phagocytosis and occurred when the cells were preincubated with antibody and washed before adding virus (17, 18). Since the MPER is a partially cryptic epitope that is not fully exposed until after Env engages with cellular receptors (21, 22), these results suggest that by binding the Fc region, FcγRI provides a local concentration advantage for MPER mAbs at the cell surface that enhances viral neutralization (17, 18). While not expressed on T cells, FcγRI is expressed on macrophages and dendritic cells (23), which are present at mucosal surfaces and are implicated in sexual HIV-1 transmission and the early establishment of HIV-1 infection (22–34).Here we investigated whether FcγRI expression also potentiates the neutralizing activity of antibodies targeting the NHR, since that region, like the MPER, is preferentially exposed during viral fusion. We found that D5, a well-characterized anti-NHR mAb (11, 12), inhibits HIV-1 infection ∼5,000-fold more potently in TZM-bl cells expressing FcγRI (TZM-bl/FcγRI cells) than in TZM-bl cells that do not. Further, while antisera from guinea pigs immunized with (ccIZN36)₃, an NHR-based vaccine candidate (7), displayed weak neutralizing activity in TZM-bl cells, they exhibited enhanced neutralization in TZM-bl/FcγRI cells, including against some tier-2 HIV-1 isolates that are more resistant to antibody-mediated neutralization (16) and that serve as benchmarks for antibody-based vaccine efforts. These results indicate that FcγRI can play an important role in neutralization by antibodies that target the PHI. Since these receptors are expressed on cells prevalent at mucosal surfaces thought to be important for sexual HIV-1 transmission, our results motivate vaccine strategies that harness this potentiating effect.     

An engineered 4-1BBL fusion protein with “activity on demand”

Engineered cytokines are gaining importance in cancer therapy, but these products are often limited by toxicity, especially at early time points after intravenous administration. 4-1BB is a member of the tumor necrosis factor receptor superfamily, which has been considered as a target for therapeutic strategies with agonistic antibodies or using its cognate cytokine ligand, 4-1BBL. Here we describe the engineering of an antibody fusion protein, termed F8-4-1BBL, that does not exhibit cytokine activity in solution but regains biological activity on antigen binding. F8-4-1BBL bound specifically to its cognate antigen, the alternatively spliced EDA domain of fibronectin, and selectively localized to tumors in vivo, as evidenced by quantitative biodistribution experiments. The product promoted a potent antitumor activity in various mouse models of cancer without apparent toxicity at the doses used. F8-4-1BBL represents a prototype for antibody-cytokine fusion proteins, which conditionally display “activity on demand” properties at the site of disease on antigen binding and reduce toxicity to normal tissues.

Cytokines are immunomodulatory proteins that have been considered for pharmaceutical applications in the treatment of cancer patients (1–3) and other types of disease (2). There is a growing interest in the use of engineered cytokine products as anticancer drugs, capable of boosting the action of T cells and natural killer (NK) cells against tumors (3, 4), alone or in combination with immune checkpoint inhibitors (3, 5–7).Recombinant cytokine products on the market include interleukin-2 (IL-2) (Proleukin) (8, 9), IL-11 (Neumega) (10, 11), tumor necrosis factor (TNF; Beromun) (12), interferon (IFN)-α (Roferon A, Intron A) (13, 14), IFN-β (Avonex, Rebif, Betaseron) (15, 16), IFN-γ (Actimmune) (17), granulocyte colony-stimulating factor (Neupogen) (18), and granulocyte macrophage colony-stimulating factor (Leukine) (19, 20). The recommended dose is typically very low (often <1 mg/d) (21–23), as cytokines may exert biological activity in the subnanomolar concentration range (24). Various strategies have been proposed to develop cytokine products with improved therapeutic index. Protein PEGylation or Fc fusions may lead to prolonged circulation time in the bloodstream, allowing the administration of low doses of active payload (25, 26). In some implementations, cleavable polyethylene glycol polymers may be considered, yielding prodrugs that regain activity at later time points (27). Alternatively, tumor-homing antibody fusions have been developed, since the preferential concentration of cytokine payloads at the tumor site has been shown in preclinical models to potentiate therapeutic activity, helping spare normal tissues (28–34). Various antibody-cytokine fusions are currently being investigated in clinical trials for the treatment of cancer and of chronic inflammatory conditions (reviewed in refs. 2, 33, 35–37).Antibody-cytokine fusions display biological activity immediately after injection into patients, which may lead to unwanted toxicity and prevent escalation to therapeutically active dosage regimens (9, 22, 38). In the case of proinflammatory payloads (e.g., IL-2, IL-12, TNF-α), common side effects include hypotension, nausea, and vomiting, as well as flu-like symptoms (24, 39–42). These side effects typically disappear when the cytokine concentration drops below a critical threshold, thus providing a rationale for slow-infusion administration procedures (43). It would be highly desirable to generate antibody-cytokine fusion proteins with excellent tumor-targeting properties and with “activity on demand”— biological activity that is conditionally gained on antigen binding at the site of disease, helping spare normal tissues.Here we describe a fusion protein consisting of the F8 antibody specific to the alternatively spliced extra domain A (EDA) of fibronectin (44, 45) and of murine 4-1BBL, which did not exhibit cytokine activity in solution but could regain potent biological activity on antigen binding. The antigen (EDA+ fibronectin) is conserved from mouse to man (46), is virtually undetectable in normal adult tissues (with the exception of the placenta, endometrium, and some vessels in the ovaries), but is expressed in the majority of human malignancies (44, 45, 47, 48). 4-1BBL, a member of the TNF superfamily (49), is expressed on antigen-presenting cells (50, 51) and binds to its receptor, 4-1BB, which is up-regulated on activated cytotoxic T cells (52), activated dendritic cells (52), activated NK and NKT cells (53), and regulatory T cells (54). Signaling through 4-1BB on cytotoxic T cells protects them from activation-induced cell death and skews the cells toward a more memory-like phenotype (55, 56).We engineered nine formats of the F8-4-1BBL fusion protein, one of which exhibited superior performance in quantitative biodistribution studies and conditional gain of cytokine activity on antigen binding. The antigen-dependent reconstitution of the biological activity of the immunostimulatory payload represents an example of an antibody fusion protein with “activity on demand.” The fusion protein was potently active against different types of cancer without apparent toxicity at the doses used. The EDA of fibronectin is a particularly attractive antigen for cancer therapy in view of its high selectivity, stability, and abundant expression in most tumor types (44, 45, 47, 48).