The circadian clock ensures successful DNA replication in cyanobacteria

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

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

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.     

Cholinergic suppression of hippocampal sharp-wave ripples impairs working memory

Learning and memory are assumed to be supported by mechanisms that involve cholinergic transmission and hippocampal theta. Using G protein–coupled receptor-activation–based acetylcholine sensor (GRAB_ACh3.0) with a fiber-photometric fluorescence readout in mice, we found that cholinergic signaling in the hippocampus increased in parallel with theta/gamma power during walking and REM sleep, while ACh3.0 signal reached a minimum during hippocampal sharp-wave ripples (SPW-R). Unexpectedly, memory performance was impaired in a hippocampus-dependent spontaneous alternation task by selective optogenetic stimulation of medial septal cholinergic neurons when the stimulation was applied in the delay area but not in the central (choice) arm of the maze. Parallel with the decreased performance, optogenetic stimulation decreased the incidence of SPW-Rs. These findings suggest that septo–hippocampal interactions play a task-phase–dependent dual role in the maintenance of memory performance, including not only theta mechanisms but also SPW-Rs.

The neurotransmitter acetylcholine is thought to be critical for hippocampus-dependent declarative memories (1, 2). Reduction in cholinergic neurotransmission, either in Alzheimer’s disease or in experiments with cholinergic antagonists, such as scopolamine, impairs memory function (3–8). Acetylcholine may bring about its beneficial effects on memory encoding by enhancing theta rhythm oscillations, decreasing recurrent excitation, and increasing synaptic plasticity (9–11). Conversely, drugs which activate cholinergic receptors enhance learning and, therefore, are a neuropharmacological target for the treatment of memory deficits in Alzheimer’s disease (5, 12, 13).The contribution of cholinergic mechanisms in the acquisition of long-term memories and the role of the hippocampal–entorhinal–cortical interactions are well supported by experimental data (5, 12, 13). In addition, working memory or “short-term” memory is also supported by the hippocampal–entorhinal–prefrontal cortex (14–16). Working memory in humans is postulated to be a conscious process to “keep things in mind” transiently (16). In rodents, matching to sample task, spontaneous alternation between reward locations, and the radial maze task have been suggested to function as a homolog of working memory [“working memory like” (17)].Cholinergic activity is a critical requirement for working memory (18, 19) and for sustaining theta oscillations (10, 20–22). In support of this contention, theta–gamma coupling and gamma power are significantly higher in the choice arm of the maze, compared with those in the side arms where working memory is no longer needed for correct performance (23–26). It has long been hypothesized that working memory is maintained by persistent firing of neurons, which keep the presented items in a transient store in the prefrontal cortex and hippocampal–entorhinal system (27–31), although the exact mechanisms are debated (32–37). An alternative hypothesis holds that items of working memory are stored in theta-nested gamma cycles (38). Common in these models of working memory is the need for an active, cholinergic system–dependent mechanism (39–41). However, in spontaneous alternation tasks, the animals are not moving continuously during the delay, and theta oscillations are not sustained either. During the immobility epochs, theta is replaced by intermittent sharp-wave ripples (SPW-R), yet memory performance does not deteriorate. On the contrary, artificial blockade of SPW-Rs can impair memory performance (42, 43), and prolongation of SPW-Rs improves performance (44). Under the cholinergic hypothesis of working memory, such a result is unexpected.To address the relationship between cholinergic/theta versus SPW-R mechanism in spontaneous alternation, we used a G protein–coupled receptor-activation–based acetylcholine sensor (GRAB_ACh3.0) (45) to monitor acetylcholine (ACh) activity during memory performance in mice. In addition, we optogenetically enhanced cholinergic tone, which suppresses SPW-Rs by a different mechanism than electrically or optogenetically induced silencing of neurons in the hippocampus (43, 44). We show that cholinergic signaling in the hippocampus increases in parallel with theta power/score during walking and rapid eye movement (REM) sleep and reaches a transient minimum during SPW-Rs. Selective optogenetic stimulation of medial septal cholinergic neurons decreased the incidence of SPW-Rs during non-REM sleep (46–48), as well as during the delay epoch of a working memory task and impaired memory performance. These findings demonstrate that memory performance is supported by complementary theta and SPW-R mechanisms.     

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

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

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

Characterization of the strain-rate–dependent mechanical response of single cell–cell junctions

Cell–cell adhesions are often subjected to mechanical strains of different rates and magnitudes in normal tissue function. However, the rate-dependent mechanical behavior of individual cell–cell adhesions has not been fully characterized due to the lack of proper experimental techniques and therefore remains elusive. This is particularly true under large strain conditions, which may potentially lead to cell–cell adhesion dissociation and ultimately tissue fracture. In this study, we designed and fabricated a single-cell adhesion micro tensile tester (SCAµTT) using two-photon polymerization and performed displacement-controlled tensile tests of individual pairs of adherent epithelial cells with a mature cell–cell adhesion. Straining the cytoskeleton–cell adhesion complex system reveals a passive shear-thinning viscoelastic behavior and a rate-dependent active stress-relaxation mechanism mediated by cytoskeleton growth. Under low strain rates, stress relaxation mediated by the cytoskeleton can effectively relax junctional stress buildup and prevent adhesion bond rupture. Cadherin bond dissociation also exhibits rate-dependent strengthening, in which increased strain rate results in elevated stress levels at which cadherin bonds fail. This bond dissociation becomes a synchronized catastrophic event that leads to junction fracture at high strain rates. Even at high strain rates, a single cell–cell junction displays a remarkable tensile strength to sustain a strain as much as 200% before complete junction rupture. Collectively, the platform and the biophysical understandings in this study are expected to build a foundation for the mechanistic investigation of the adaptive viscoelasticity of the cell–cell junction.

Adhesive organelles between neighboring epithelial cells form an integrated network as the foundation of complex tissues (1). As part of normal physiology, this integrated network is constantly exposed to mechanical stress and strain, which is essential to normal cellular activities, such as proliferation (2–4), migration (5, 6), differentiation (7), and gene regulation (7, 8) associated with a diverse set of functions in tissue morphogenesis (9–11) and wound healing (9). A host of developmental defects or clinical pathologies in the form of compromised cell–cell associations will arise when cells fail to withstand external mechanical stress due to genetic mutations or pathological perturbations (12, 13). Indeed, since the mechanical stresses are mainly sustained by the intercellular junctions, which may represent the weakest link and limit the stress tolerance within the cytoskeleton network of a cell sheet, mutations or disease-induced changes in junction molecules and components in adherens junctions and desmosomes lead to cell layer fracture and tissue fragility, which exacerbate the pathological conditions (14–17). This clinical relevance gives rise to the importance of understanding biophysical transformations of the cell–cell adhesion interface when cells are subjected to mechanical loads.As part of their normal functions, cells often experience strains of tens to a few hundred percent at strain rates of 10⁻⁴ to 1 s⁻¹ (18–21). For instance, embryonic epithelia are subjected to strain rates in the range of 10⁻⁴ to 10⁻³ s⁻¹ during normal embryogenesis (22). Strain rates higher than 0.1 s⁻¹ are often experienced by adult epithelia during various normal physiological functions (21, 23, 24), such as breathing motions in the lung (1 to 10 s⁻¹) (25), cardiac pulses in the heart (1 to 6.5 s⁻¹) (20), peristaltic movements in the gut (0.4 to 1.5 s⁻¹), and normal stretching of the skin (0.1 to 5 s⁻¹). Cells have different mechanisms to dissipate the internal stress produced by external strain to avoid fracture, often via cytoskeleton remodeling and cell–cell adhesion enhancement (26, 27). These coping mechanisms may have different characteristic timescales. Cytoskeleton remodeling can dissipate mechanical stress promptly due to its viscoelastic nature and the actomyosin-mediated cell contractility (17, 28–32). Adhesion enhancement at the cell–cell contact is more complex in terms of timescale. Load-induced cell–cell adhesion strengthening has been shown via the increase in the number of adhesion complexes (33–35) or by the clustering of adhesion complexes (36–39), which occurs on a timescale ranging from a few minutes up to a few hours after cells experience an initial load (28). External load on the cell–cell contact also results in a prolonged cell–cell adhesion dissociation time (40, 41), suggesting cadherin bonds may transition to catch bonds under certain loading conditions (42, 43), which can occur within seconds (44). With the increase in cellular tension, failure to dissipate the stress within the cell layer at a rate faster than the accumulation rate will inevitably lead to the fracture of the cell layer (45). Indeed, epithelial fracture often aggravates the pathological outcomes in several diseases, such as acute lung injuries (46), skin disorders (47), and development defects (48). It is generally accepted that stress accumulation in the cytoskeleton network (49, 50) and potentially in the cytoplasm is strain-rate–dependent (51). However, to date, there is a lack of understanding about the rate-dependent behavior of cell–cell adhesions, particularly about which of the stress-relaxation mechanisms are at play across the spectrum of strain rates. In addition, it remains unclear how the stress relaxation interplays with adhesion enhancement under large strains, especially at high strain rates which may lead to fracture, that is, a complete separation of mature cell–cell adhesions under a tensile load (45, 52, 53). Yet, currently, there is a lack of quantitative technology that enables the investigation of these mechanobiological processes in a precisely controlled manner. This is especially true at high strain rates.To delineate this mechanical behavior, the cleanest characterization method is to directly measure stress dynamics at a single mature cell–cell adhesion interface. Specifically, just as a monolayer cell sheet is a reduction from three-dimensional (3D) tissue, a single cell–cell adhesion interface, as a reduction from a monolayer system, represents the smallest unit to study the rheological behavior of cellular junctions. The mechanistic understanding uncovered with this single unit will inform cellular adaptations to a more complex stress microenvironment in vivo and in vitro, in healthy and diseased conditions. To this end, we developed a single-cell adhesion micro tensile tester (SCAµTT) platform based on nanofabricated polymeric structures using two-photon polymerization (TPP). This platform allows in situ investigation of stress–strain characteristics of a mature cell–cell junction through defined strains and strain rates. With SCAµTT, we reveal some interesting biophysical phenomena at the single cell–cell junction that were previously not possible to observe using existing techniques. We show that cytoskeleton growth can effectively relax intercellular stress between an adherent cell pair in a strain-rate–dependent manner. Along with cadherin-clustering–induced bond strengthening, it prevents failure to occur at low strain rates. At high strain rates, insufficient relaxation leads to stress accumulation, which results in cell–cell junction rupture. We show that a remarkably large strain can be sustained before junction rupture (>200%), even at a strain rate as high as 0.5 s⁻¹. Collectively, the rate-dependent mechanical characterization of the cell–cell junction builds the foundation for an improved mechanistic understanding of junction adaptation to an external load and potentially the spatiotemporal coordination of participating molecules at the cell–cell junction.     

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.     

Violet light suppresses lens-induced myopia via neuropsin (OPN5) in mice

Myopia has become a major public health concern, particularly across much of Asia. It has been shown in multiple studies that outdoor activity has a protective effect on myopia. Recent reports have shown that short-wavelength visible violet light is the component of sunlight that appears to play an important role in preventing myopia progression in mice, chicks, and humans. The mechanism underlying this effect has not been understood. Here, we show that violet light prevents lens defocus–induced myopia in mice. This violet light effect was dependent on both time of day and retinal expression of the violet light sensitive atypical opsin, neuropsin (OPN5). These findings identify Opn5-expressing retinal ganglion cells as crucial for emmetropization in mice and suggest a strategy for myopia prevention in humans.

Myopia (nearsightedness) in school-age children is generally axial myopia, which is the consequence of elongation of the eyeball along the visual axis. This shape change results in blurred vision but can also lead to severe complications including cataract, retinal detachment, myopic choroidal neovascularization, glaucoma, and even blindness (1–3). Despite the current worldwide pandemic of myopia, the mechanism of myopia onset is still not understood (4–8). One hypothesis that has earned a current consensus is the suggestion that a change in the lighting environment of modern society is the cause of myopia (9, 10). Consistent with this, outdoor activity has a protective effect on myopia development (9, 11, 12), though the main reason for this effect is still under debate (7, 12, 13). One explanation is that bright outdoor light can promote the synthesis and release of dopamine in the eye, a myopia-protective neuromodulator (14–16). Another suggestion is that the distinct wavelength composition of sunlight compared with fluorescent or LED (light-emitting diode) artificial lighting may influence myopia progression (9, 10). Animal studies have shown that different wavelengths of light can affect the development of myopia independent of intensity (17, 18). The effects appear to be distinct in different species: for chicks and guinea pigs, blue light showed a protective effect on experimentally induced myopia, while red light had the opposite effect (18–22). For tree shrews and rhesus monkeys, red light is protective, and blue light causes dysregulation of eye growth (23–25).It has been shown that visible violet light (VL) has a protective effect on myopia development in mice, in chick, and in human (10, 26, 27). According to Commission Internationale de l’Eclairage (International Commission on Illumination), VL has the shortest wavelength of visible light (360 to 400 nm). These wavelengths are abundant in outside sunlight but can only rarely be detected inside buildings. This is because the ultraviolet (UV)-protective coating on windows blocks all light below 400 nm and because almost no VL is emitted by artificial light sources (10). Thus, we hypothesized that the lack of VL in modern society is one reason for the myopia boom (9, 10, 26).In this study, we combine a newly developed lens-induced myopia (LIM) model with genetic manipulations to investigate myopia pathways in mice (28, 29). Our data confirm (10, 26) that visible VL is protective but further show that delivery of VL only in the evening is sufficient for the protective effect. In addition, we show that the protective effect of VL on myopia induction requires OPN5 (neuropsin) within the retina. The absence of retinal Opn5 prevents lens-induced, VL-dependent thickening of the choroid, a response thought to play a key role in adjusting the size of the eyeball in both human and animal myopia models (30–33). This report thus identifies a cell type, the Opn5 retinal ganglion cell (RGC), as playing a key role in emmetropization. The requirement for OPN5 also explains why VL has a protective effect on myopia development.     

DDX11 loss causes replication stress and pharmacologically exploitable DNA repair defects

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

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

Structured sequences emerge from random pool when replicated by templated ligation

The central question in the origin of life is to understand how structure can emerge from randomness. The Eigen theory of replication states, for sequences that are copied one base at a time, that the replication fidelity has to surpass an error threshold to avoid that replicated specific sequences become random because of the incorporated replication errors [M. Eigen, Naturwissenschaften 58 (10), 465–523 (1971)]. Here, we showed that linking short oligomers from a random sequence pool in a templated ligation reaction reduced the sequence space of product strands. We started from 12-mer oligonucleotides with two bases in all possible combinations and triggered enzymatic ligation under temperature cycles. Surprisingly, we found the robust creation of long, highly structured sequences with low entropy. At the ligation site, complementary and alternating sequence patterns developed. However, between the ligation sites, we found either an A-rich or a T-rich sequence within a single oligonucleotide. Our modeling suggests that avoidance of hairpins was the likely cause for these two complementary sequence pools. What emerged was a network of complementary sequences that acted both as templates and substrates of the reaction. This self-selecting ligation reaction could be restarted by only a few majority sequences. The findings showed that replication by random templated ligation from a random sequence input will lead to a highly structured, long, and nonrandom sequence pool. This is a favorable starting point for a subsequent Darwinian evolution searching for higher catalytic functions in an RNA world scenario.

One of the dominant hypotheses to explain the origin of life (1–3) is the concept of the RNA world. It is built on the fact that catalytically active RNA molecules can enzymatically promote their own replication (4–6) via active sites in their three-dimensional structures (7–9). These so-called ribozymes have a minimal length of 30 to 41 base pairs (9, 10) and, thus, a sequence space of more than 4³⁰ ∼ 10¹⁸. The subset of functional, catalytically active sequences in this vast sequence space is vanishingly small (11), making spontaneous assembly of ribozymes from monomers or oligomers all but impossible. Therefore, prebiotic evolution has likely provided some form of selection guiding single nucleotides to form functional sequences and thereby lowering the sequence entropy of this system.The problem of nonenzymatic formation of single base nucleotides and short oligomers in settings reminiscent of the primordial soup has been studied before (12–17). However, the continuation of this evolutionary path toward early replication networks would require a preselection mechanism of oligonucleotides (see Fig. 1A), lowering the information entropy of the resulting sequence pool (18–22). In principle, such selection modes include optimization for information storage, local oligomer enrichment (e.g., in hydrogels or in catalytically functional sites).Open in a separate windowFig. 1.Templated ligation of random sequence DNA 12-mers. (A) Before cells evolved, the first ribozymes were thought to perform basic cell functions. In the exponentially vast sequence space, spontaneous emergence of a functional ribozyme is highly unlikely, therefore preselection mechanisms were likely necessary. (B) In our experiment, DNA strands hybridize at low temperatures to form three-dimensional complexes that can be ligated and preserved in the high temperature dissociation steps. The system self-selects for sequences with specific ligation site motifs as well as for strands that continue acting as templates. Hairpin sequences are therefore suppressed. (C) Concentration analysis shows progressively longer strands emerging after multiple temperature cycles. The inset (A-red, T-blue) shows that, although 12-mers (88,009 strands) have essentially random sequences (white), various sequence patterns emerge in longer strands (60-mers, 235,913 strands analyzed). (D) Samples subjected to different number (0 to 1,000) of temperature cycles between 75 °C and 33 °C. Concentration quantification is done on PAGE with SYBR poststained DNA.An important aspect of a selection mechanism is its nonequilibrium driving force. Today’s highly evolved cells function through multistep and multicomponent metabolic pathways like glycolysis in the Warburg effect (23) or by specialized enzymes like adenosine triphosphate (ATP) synthase which provide energy-rich ATP (24). In contrast, it is widely assumed (3, 4, 25–28) that selection mechanisms for molecular evolution at the dawn of life must have been much simpler (e.g., mediated by random binding between biomolecules subject to nonequilibrium driving forces such as fluid flow and cyclic changes in temperature).Here, we explored the possibility of a significant reduction of sequence entropy driven by templated ligation (19) and mediated by Watson–Crick base pairing (29). Starting from a random pool of oligonucleotides, we observed a gradual formation of longer chains showing reproducible sequence landscape inhibiting self-folding and promoting templated ligation. Here, we argue that base pairing combined with ligation chemistry can trigger processes that have many features of the Darwinian evolution.As a model oligomer, we decided to use DNA instead of RNA since the focus of our study is on base pairing, which is very similar for both (30). We started our experiments with a random pool of 12-mers formed of bases A (adenine) and T (thymine). This binary code facilitates binding between molecules and allows us to sample the whole sequence space in microliter volumes (2¹² << 10 µM × 20 µL × N_A = 10¹⁴).Formation of progressively longer oligomers from shorter ones requires ligation reactions, a method commonly employed in hairpin-mediated RNA and DNA replication (31, 32). At the origin of life, this might have been achieved by activated oligomers (33, 34) or activation agents (35–37), whereas later on the formation of simple ribozyme ligases seemed possible (38). Our study is focused on inherent properties of self-assembly by base pairing in random pools of oligomers and not on chemical mechanisms of ligation. Hence, we decided to use TAQ DNA ligase—an evolved enzyme for templated ligation of DNA (21) that is known for its ligation site sequence specificity (39, 40) and lack of sequence-dependent ligation rate (compare SI Appendix, section 21). This allowed for fast turnovers of ligation and enabled the observation of sequence dynamics.     

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.     

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).