CYK-1/Formin activation in cortical RhoA signaling centers promotes organismal left–right symmetry breaking

Proper left–right symmetry breaking is essential for animal development, and in many cases, this process is actomyosin-dependent. In Caenorhabditis elegans embryos active torque generation in the actomyosin layer promotes left–right symmetry breaking by driving chiral counterrotating cortical flows. While both Formins and Myosins have been implicated in left–right symmetry breaking and both can rotate actin filaments in vitro, it remains unclear whether active torques in the actomyosin cortex are generated by Formins, Myosins, or both. We combined the strength of C. elegans genetics with quantitative imaging and thin film, chiral active fluid theory to show that, while Non-Muscle Myosin II activity drives cortical actomyosin flows, it is permissive for chiral counterrotation and dispensable for chiral symmetry breaking of cortical flows. Instead, we find that CYK-1/Formin activation in RhoA foci is instructive for chiral counterrotation and promotes in-plane, active torque generation in the actomyosin cortex. Notably, we observe that artificially generated large active RhoA patches undergo rotations with consistent handedness in a CYK-1/Formin–dependent manner. Altogether, we conclude that CYK-1/Formin–dependent active torque generation facilitates chiral symmetry breaking of actomyosin flows and drives organismal left–right symmetry breaking in the nematode worm.

The emergence of left–right asymmetry is essential for normal animal development and, in the majority of animal species, one type of handedness is dominant (1). The actin cytoskeleton plays an instrumental role in establishing the left–right asymmetric body plan of invertebrates like fruit flies (2–6), nematodes (7–11), and pond snails (12–15). Moreover, an increasing number of studies showed that vertebrate left–right patterning also depends on a functional actomyosin cytoskeleton (13, 16–22). Actomyosin-dependent chiral behavior has even been reported in isolated cells (23–28) and such cell-intrinsic chirality has been shown to promote left–right asymmetric morphogenesis of tissues (29, 30), organs (21, 31), and entire embryonic body plans (12, 13, 32, 33). Active force generation in the actin cytoskeleton is responsible for shaping cells and tissues during embryo morphogenesis. Torques are rotational forces with a given handedness and it has been proposed that in plane, active torque generation in the actin cytoskeleton drives chiral morphogenesis (7, 8, 34, 35).What could be the molecular origin of these active torques? The actomyosin cytoskeleton consists of actin filaments, actin-binding proteins, and Myosin motors. Actin filaments are polar polymers with a right-handed helical pitch and are therefore chiral themselves (36, 37). Due to the right-handed pitch of filamentous actin, Myosin motors can rotate actin filaments along their long axis while pulling on them (33, 38–42). Similarly, when physically constrained, members of the Formin family rotate actin filaments along their long axis while elongating them (43). In both cases the handedness of this rotation is determined by the helical nature of the actin polymer. From this it follows that both Formins and Myosins are a potential source of molecular torque generation that could drive cellular and organismal chirality. Indeed, chiral processes across different length scales, and across species, are dependent on Myosins (19), Formins (13–15, 26), or both (7, 8, 21, 44). It is, however, unclear how Formins and Myosins contribute to active torque generation and the emergence chiral processes in developing embryos.In our previous work we showed that the actomyosin cortex of some Caenorhabditis elegans embryonic blastomeres undergoes chiral counterrotations with consistent handedness (7, 35). These chiral actomyosin flows can be recapitulated using active chiral fluid theory that describes the actomyosin layer as a thin-film, active gel that generates active torques (7, 45, 46). Chiral counterrotating cortical flows reorient the cell division axis, which is essential for normal left–right symmetry breaking (7, 47). Moreover, cortical counterrotations with the same handedness have been observed in Xenopus one-cell embryos (32), suggesting that chiral counterrotations are conserved among distant species. Chiral counterrotating actomyosin flow in C. elegans blastomeres is driven by RhoA signaling and is dependent on Non-Muscle Myosin II motor proteins (7). Moreover, the Formin CYK-1 has been implicated in actomyosin flow chirality during early polarization of the zygote as well as during the first cytokinesis (48, 49). Despite having identified a role for Myosins and Formins, the underlying mechanism by which active torques are generated remains elusive.Here we show that the Diaphanous-like Formin, CYK-1/Formin, is a critical determinant for the emergence of actomyosin flow chirality, while Non-Muscle Myosin II (NMY-2) plays a permissive role. Our results show that cortical CYK-1/Formin is recruited by active RhoA signaling foci and promotes active torque generation, which in turn tends to locally rotate the actomyosin cortex clockwise. In the highly connected actomyosin meshwork, a gradient of these active torques drives the emergence of chiral counterrotating cortical flows with uniform handedness, which is essential for proper left–right symmetry breaking. Together, these results provide mechanistic insight into how Formin-dependent torque generation drives cellular and organismal left–right symmetry breaking.     

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.     

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.     

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.     

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).     

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.     

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.     

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.     

Photosynthesis tunes quantum-mechanical mixing of electronic and vibrational states to steer exciton energy transfer

Photosynthetic species evolved to protect their light-harvesting apparatus from photoxidative damage driven by intracellular redox conditions or environmental conditions. The Fenna–Matthews–Olson (FMO) pigment–protein complex from green sulfur bacteria exhibits redox-dependent quenching behavior partially due to two internal cysteine residues. Here, we show evidence that a photosynthetic complex exploits the quantum mechanics of vibronic mixing to activate an oxidative photoprotective mechanism. We use two-dimensional electronic spectroscopy (2DES) to capture energy transfer dynamics in wild-type and cysteine-deficient FMO mutant proteins under both reducing and oxidizing conditions. Under reducing conditions, we find equal energy transfer through the exciton 4–1 and 4–2-1 pathways because the exciton 4–1 energy gap is vibronically coupled with a bacteriochlorophyll-a vibrational mode. Under oxidizing conditions, however, the resonance of the exciton 4–1 energy gap is detuned from the vibrational mode, causing excitons to preferentially steer through the indirect 4–2-1 pathway to increase the likelihood of exciton quenching. We use a Redfield model to show that the complex achieves this effect by tuning the site III energy via the redox state of its internal cysteine residues. This result shows how pigment–protein complexes exploit the quantum mechanics of vibronic coupling to steer energy transfer.

Photosynthetic organisms convert solar photons into chemical energy by taking advantage of the quantum mechanical nature of their molecular systems and the chemistry of their environment (1–4). Antenna complexes, composed of one or more pigment–protein complexes, facilitate the first steps in the photosynthesis process: They absorb photons and determine which proportion of excitations to move to reaction centers, where charge separation occurs (4). In oxic environments, excitations can generate highly reactive singlet oxygen species. These pigment–protein complexes can quench excess excitations in these environments with molecular moieties such as quinones and cysteine residues (1, 5–7).The Fenna–Matthews–Olson (FMO) complex, a trimer of pigment–protein complexes found in the green sulfur bacterium Chlorobaculum tepidum (8), has emerged as a model system to study the photophysical properties of photosynthetic antenna complexes (9–19). Each subunit in the FMO complex contains eight bacteriochlorophyll-a site molecules (Protein Data Bank, ID code: 3ENI) that are coupled to form a basis of eight partially delocalized excited states called excitons (Fig. 1) (20–23). Previous experiments on FMO have observed the presence of long-lived coherences in nonlinear spectroscopic signals at both cryogenic and physiological temperatures (11, 13). The coherent signals are thought to arise from some combination of electronic (24–26), vibrational (16–18), and vibronic (27) coherences in the system (28–30). One previous study reported that the coherent signals in FMO remain unchanged upon mutagenesis of the protein, suggesting that the signals are ground state vibrational coherences (17). Others discuss the role of vibronic coupling, where electronic and nuclear degrees of freedom become coupled (29). Other dimeric model systems have demonstrated the regimes in which these vibronically coupled states produce coherent or incoherent transport and vibronic coherences (31–33). Recent spectroscopic data has suggested that vibronic coupling plays a role in driving efficient energy transfer through photosynthetic complexes (27, 31, 33, 34), but to date there is no direct experimental evidence suggesting that biological systems use vibronic coupling as part of their biological function.Open in a separate windowFig. 1.(Left) Numbered sites and sidechains of cysteines C353 and C49 in the FMO pigment–protein complex (PDB ID code: 3ENI) (20). (Right) Site densities for excitons 4, 2, and 1 in reducing conditions with the energy transfer branching ratios for the WT oxidized and reduced protein. The saturation of pigments in each exciton denotes the relative contribution number to the exciton. The C353 residue is located near excitons 4 and 2, which have most electron density along one side of the complex, and other redox-active residues such as the Trp/Tyr chain. C353 and C49 surround site III, which contains the majority of exciton 1 density. Excitons 2 and 4 are generally delocalized over sites IV, V, and VII.It has been shown that redox conditions affect excited state properties in pigment-protein complexes, yet little is known about the underlying microscopic mechanisms for these effects (1, 9). Many commonly studied light-harvesting complexes—including the FMO complex (20), light-harvesting complex 2 (LH2) (35), the PC645 phycobiliprotein (36), and the cyanobacterial antenna complex isiA (37)—contain redox-active cysteine residues in close proximity to their chromophores. As the natural low light environment of C. tepidum does not necessitate photoprotective responses to light quantity and quality, its primary photoprotective mechanism concerns its response to oxidative stress. C. tepidum is an obligate anaerobe, but the presence of many active anoxygenic genes such as sodB for superoxide dismutase and roo for rubredoxin oxygen oxidoreductase (38) suggests that it is frequently exposed to molecular oxygen (7, 39). Using time-resolved fluorescence measurements, Orf et al. demonstrated that two cysteine residues in the FMO complex, C49 and C353, quench excitons under oxidizing conditions (1), which could protect the excitation from generating reactive oxygen species (7, 40–42). In two-dimensional electronic spectroscopy (2DES) experiments, Allodi et al. showed that redox conditions in both the wild-type and C49A/C353A double-mutant proteins affect the ultrafast dynamics through the FMO complex (9, 43). The recent discovery that many proteins across the evolutionary landscape possess chains of tryptophan and tyrosine residues provides evidence that these redox-active residues may link the internal protein behavior with the chemistry of the surrounding environment (41, 43).In this paper, we present data showing that pigment–protein complexes tune the vibronic coupling of their chromophores and that the absence of this vibronic coupling activates an oxidative photoprotective mechanism. We use 2DES to show that a pair of cysteine residues in FMO, C49 and C353, can steer excitations toward quenching sites in oxic environments. The measured reaction rate constants demonstrate unusual nonmonotonic behavior. We then use a Redfield model to determine how the exciton energy transfer (EET) time constants arise from changing chlorophyll site energies and their system-bath couplings (44, 45). The analysis reveals that the cysteine residues tune the resonance between exciton 4–1 energy gap and an intramolecular chlorophyll vibration in reducing conditions to induce vibronic coupling and detune the resonance in oxidizing conditions. This redox-dependent modulation of the vibronic coupling steers excitations through different pathways in the complex to change the likelihood that they interact with exciton quenchers.     

Oncogenic KRAS engages an RSK1/NF1 pathway to inhibit wild-type RAS signaling in pancreatic cancer

Pancreatic ductal adenocarcinoma (PDAC) is a lethal malignancy with limited treatment options. Although activating mutations of the KRAS GTPase are the predominant dependency present in >90% of PDAC patients, targeting KRAS mutants directly has been challenging in PDAC. Similarly, strategies targeting known KRAS downstream effectors have had limited clinical success due to feedback mechanisms, alternate pathways, and dose-limiting toxicities in normal tissues. Therefore, identifying additional functionally relevant KRAS interactions in PDAC may allow for a better understanding of feedback mechanisms and unveil potential therapeutic targets. Here, we used proximity labeling to identify protein interactors of active KRAS in PDAC cells. We expressed fusions of wild-type (WT) (BirA-KRAS4B), mutant (BirA-KRAS4B^G12D), and nontransforming cytosolic double mutant (BirA-KRAS4B^G12D/C185S) KRAS with the BirA biotin ligase in murine PDAC cells. Mass spectrometry analysis revealed that RSK1 selectively interacts with membrane-bound KRAS^G12D, and we demonstrate that this interaction requires NF1 and SPRED2. We find that membrane RSK1 mediates negative feedback on WT RAS signaling and impedes the proliferation of pancreatic cancer cells upon the ablation of mutant KRAS. Our findings link NF1 to the membrane-localized functions of RSK1 and highlight a role for WT RAS signaling in promoting adaptive resistance to mutant KRAS-specific inhibitors in PDAC.

A total of 60,430 new cases of pancreatic cancer were estimated for 2021, and the 5-y relative survival rate has consistently remained below 11% (1). About 85% of these pancreatic cancer tumors are pancreatic ductal adenocarcinoma (PDAC) (2). Poor outcomes of PDAC cases result from late diagnoses leading to unresectable and heterogeneous tumors as well as ineffective therapies, which only prolong survival on the order of months (3–5). Mutations in the KRAS proto-oncogene are present in over 90% of PDAC cases and are associated with a poor prognosis (6). Furthermore, mice expressing mutant KRAS in the pancreas develop precursor lesions, which sporadically progress into frank PDAC. This progression is accelerated when combined with other mutations or deletion of tumor suppressor genes (7–11). Additionally, independent studies have shown that the maintenance of murine PDAC cells require KRAS (12–14).As a RAS GTPase, KRAS acts as a molecular switch at the plasma membrane that relays growth factor signaling from receptor tyrosine kinases to downstream pathways such as RAF/MEK and PI3K/AKT (15). GTP binding alters the conformation of the KRAS G domain, thereby creating binding sites for downstream effectors to trigger enzymatic cascades that promote cell transformation (16–19). Intrinsically, KRAS slowly hydrolyzes GTP into GDP to halt signaling; however, GTPase activating proteins (GAPs) such as neurofibromin 1(NF1) catalyze this process (20). In contrast, guanine nucleotide exchange factors, such as son of sevenless homolog 1 (SOS1), catalyze the exchange of GTP for bound GDP. In most PDAC cases, KRAS is mutated at the 12th residue located in the G domain from glycine to either a valine (G12V), or more commonly, aspartate (G12D). These mutations sterically prevent the “arginine finger domain” of GAPs from entering the GTPase site, thereby blocking extrinsic allosteric GTPase activation and stabilizing RAS-GTP (21, 22). Activating mutations in KRAS constitutively trigger RAF/MEK and PI3K/AKT pathways leading to increased cell proliferation as well as other prooncogenic behaviors (15). KRAS signaling not only relies on the G domain but also the C-terminal hypervariable domain (HVR), which is required to stabilize KRAS on membranes where signaling is most efficient (23–26). Independent studies suggest that specific biochemical and cellular consequences of KRAS activation are attributed to the unique properties of the HVR of the predominant splice form KRAS4B, namely the polybasic domain and the lipid anchor (27–30). Localization of RAS proteins to the plasma membrane requires the prenylation of the CAAX motif (23). Additionally, for KRAS4B, the hypervariable region contains a highly polybasic domain consisting of several consecutive lysines, which can interact with the negative charges on the polar heads of phospholipids and stabilize protein interactions (31). Structural and biochemical characterization of the HVR and G domain has contributed to a better understanding of the signaling outputs of KRAS and led to KRAS-targeting strategies.Various approaches to inhibit KRAS include direct inhibition, expression interference, mislocalization, and targeting of downstream effectors (32). Thus far, direct inhibitors against KRAS have only successfully targeted the G12C mutant, which comprises 2.9% of KRAS mutant PDAC (21, 33). For other KRAS mutants, targeting downstream effectors of KRAS in pancreatic cancer remains an alternative approach. Unfortunately, dual inhibition of MEK and AKT pathways was ineffective in PDAC patients (34). Difficulty in targeting KRAS due to adaptive resistance and feedback regulation motivates a better understanding of KRAS biology (35). For example, although PDAC typically features a mutant KRAS, there may be a role for its wild-type (WT) counterpart as well as WT RAS paralogs (HRAS and NRAS), which are GAP sensitive and subject to signaling feedback. While oncogenic KRAS has been shown to activate WT HRAS and NRAS via allosteric stimulation of SOS1 (36), WT KRAS has been proposed to be a tumor suppressor in some KRAS mutant cancers based on the commonly observed mutant-specific allele imbalance that occurs throughout tumor progression (37). Additionally, the reintroduction of WT KRAS abolished tumor T cell acute lymphoblastic leukemia development and impaired tumor growth in KRAS mutant lung cancer cells in vivo (37–39). The discovery of novel KRAS protein interactors involved in downstream signaling or feedback and compensatory pathways may elucidate why inhibition of downstream pathways have had limited clinical impact in PDAC. Here, we perform proximity labeling experiments by expressing a fusion of BirA^R118G biotin ligase and KRAS in PDAC cells, which, in the presence of high concentrations of biotin, generates reactive biotinoyl-AMP that labels lysines of nearby proteins, such as interactors of its fusion partner KRAS (40–42). The biotinylated interactor proteins can be isolated by streptavidin pulldown and analyzed by proteomics to identify novel protein interactors (43–45). Because covalent labeling occurs in living cells, enzymatic labeling may potentially identify transient interactors and protein complexes.Two recent studies used proximity-dependent biotin identification (BioID) labeling methods to identify KRAS interactors in 293T and colon cancer cells (46, 47). These studies uncovered and validated the functional relevance of PIP5KA1 and mTORC2 in PDAC cells. However, BirA-KRAS screens in PDAC models have not yet been performed. Since the tumor context may determine protein expression and relevant interactions, we sought to perform a BirA-KRAS screen in PDAC cells. We hypothesize that proximity labeling with BioID presents a means for identifying new mutant KRAS-specific interactions in PDAC, which may unveil new insights into therapeutic design for this malignancy.     

ICAM-1 induced rearrangements of capsid and genome prime rhinovirus 14 for activation and uncoating

Most rhinoviruses, which are the leading cause of the common cold, utilize intercellular adhesion molecule-1 (ICAM-1) as a receptor to infect cells. To release their genomes, rhinoviruses convert to activated particles that contain pores in the capsid, lack minor capsid protein VP4, and have an altered genome organization. The binding of rhinoviruses to ICAM-1 promotes virus activation; however, the molecular details of the process remain unknown. Here, we present the structures of virion of rhinovirus 14 and its complex with ICAM-1 determined to resolutions of 2.6 and 2.4 Å, respectively. The cryo-electron microscopy reconstruction of rhinovirus 14 virions contains the resolved density of octanucleotide segments from the RNA genome that interact with VP2 subunits. We show that the binding of ICAM-1 to rhinovirus 14 is required to prime the virus for activation and genome release at acidic pH. Formation of the rhinovirus 14–ICAM-1 complex induces conformational changes to the rhinovirus 14 capsid, including translocation of the C termini of VP4 subunits, which become poised for release through pores that open in the capsids of activated particles. VP4 subunits with altered conformation block the RNA–VP2 interactions and expose patches of positively charged residues. The conformational changes to the capsid induce the redistribution of the virus genome by altering the capsid–RNA interactions. The restructuring of the rhinovirus 14 capsid and genome prepares the virions for conversion to activated particles. The high-resolution structure of rhinovirus 14 in complex with ICAM-1 explains how the binding of uncoating receptors enables enterovirus genome release.

Human rhinoviruses are the cause of more than half of common colds (1). Medical visits and missed days of school and work cost tens of billions of US dollars annually (2, 3). There is currently no cure for rhinovirus infections, and the available treatments are only symptomatic. Rhinoviruses belong to the family Picornaviridae, genus Enterovirus, and are classified into species A, B, and C (4). Rhinoviruses A and B can belong to either “major” or “minor” groups, based on their utilization of intercellular adhesion molecule-1 (ICAM-1) or low-density lipoprotein receptor for cell entry (5–7). Type C rhinoviruses use CDHR3 as a receptor (8). Rhinovirus 14 belongs to the species rhinovirus B and uses ICAM-1 as a receptor. Receptors recognized by rhinoviruses and other enteroviruses can be divided into two groups based on their function in the infection process (9). Attachment receptors such as DAF, PSGL1, KREMEN1, CDHR3, and sialic acid enable the binding and endocytosis of virus particles into cells (10–13). In contrast, uncoating receptors including ICAM-1, CD155, CAR, and SCARB2 enable virus cell entry but also promote genome release from virus particles (5, 14–16).Virions of rhinoviruses are nonenveloped and have icosahedral capsids (17). Genomes of rhinoviruses are 7,000 to 9,000 nucleotide-long single-stranded positive-sense RNA molecules (1, 17). The rhinovirus genome encodes a single polyprotein that is co- and posttranslationally cleaved into functional protein subunits. Capsid proteins VP1, VP3, and VP0, originating from one polyprotein, form a protomer, 60 of which assemble into a pseudo-T = 3 icosahedral capsid. To render the virions mature and infectious, VP0 subunits are cleaved into VP2 and VP4 (18, 19). VP1 subunits form pentamers around fivefold symmetry axes, whereas subunits VP2 and VP3 form heterohexamers centered on threefold symmetry axes. The major capsid proteins VP1 through 3 have a jelly roll β-sandwich fold formed by two β-sheets, each containing four antiparallel β-strands, which are conventionally named B to I (20–22). The two β-sheets contain the strands BIDG and CHEF, respectively. The C termini of the capsid proteins are located at the virion surface, whereas the N termini mediate interactions between the capsid proteins and the RNA genome on the inner surface of the capsid. VP4 subunits are attached to the inner face of the capsid formed by the major capsid proteins. The surfaces of rhinovirus virions are characterized by circular depressions called canyons, which are centered around fivefold symmetry axes of the capsids (21).The VP1 subunits of most rhinoviruses, but not those of rhinovirus 14, contain hydrophobic pockets, which are filled by molecules called pocket factors (17, 21, 23, 24). It has been speculated that pocket factors are fatty acids or lipids (25). The pockets are positioned immediately below the canyons. The exposure of rhinoviruses to acidic pH induces expulsion of the pocket factors, which leads to the formation of activated particles and genome release (17, 26–32). The activated particles are characterized by capsid expansion, a reduction in interpentamer contacts, the release of VP4 subunits, externalization of N termini of VP1 subunits, and changes in the distribution of RNA genomes (17, 26–29, 33, 34). Artificial hydrophobic compounds that bind to VP1 pockets with high affinity inhibit infection by rhinoviruses (35, 36).ICAM-1 is an endothelial- and leukocyte-associated protein that stabilizes cell–cell interactions and facilitates the movement of leukocytes through endothelia (37). ICAM-1 can be divided into an extracellular amino-terminal part composed of five immunoglobulin domains, a single transmembrane helix, and a 29-residue–long carboxyl-terminal cytoplasmic domain. The immunoglobulin domains are characterized by a specific fold that consists of seven to eight β-strands, which form two antiparallel β-sheets in a sandwich arrangement (38–40). The immunoglobulin domains of ICAM-1 are stabilized by disulfide bonds and glycosylation (38–41). The connections between the immunoglobulin domains are formed by flexible linkers that enable bending of the extracellular part of ICAM-1. For example, the angle between domains 1 and 2 differs by 8° between molecules in distinct crystal forms (38, 42). As a virus receptor, ICAM-1 enables the virus particles to sequester at the cell surface and mediates their endocytosis (5). The structures of complexes of rhinoviruses 3, 14, and 16, and CVA21 with ICAM-1 have been determined to resolutions of 9 to 28 Å (42–46). It was shown that ICAM-1 molecules bind into the canyons at the rhinovirus surface, approximately between fivefold and twofold symmetry axes (42–46). ICAM-1 interacts with residues from all three major capsid proteins. It has been speculated that the binding of ICAM-1 triggers the transition of virions of rhinovirus 14 to activated particles and initiates genome release (45, 47). However, the limited resolution of the previous studies prevented characterization of the corresponding molecular mechanism.Here, we present the cryo-electron microscopy (cryo-EM) reconstruction of the rhinovirus 14 virion, which contains resolved density of octanucleotide segments of the RNA genome that interact with VP2 subunits. Furthermore, we show that the binding of ICAM-1 to rhinovirus 14 induces changes in its capsid and genome, which are required for subsequent virus activation and genome release at acidic pH.     

Chemokine CCL5 promotes robust optic nerve regeneration and mediates many of the effects of CNTF gene therapy

Ciliary neurotrophic factor (CNTF) is a leading therapeutic candidate for several ocular diseases and induces optic nerve regeneration in animal models. Paradoxically, however, although CNTF gene therapy promotes extensive regeneration, recombinant CNTF (rCNTF) has little effect. Because intraocular viral vectors induce inflammation, and because CNTF is an immune modulator, we investigated whether CNTF gene therapy acts indirectly through other immune mediators. The beneficial effects of CNTF gene therapy remained unchanged after deleting CNTF receptor alpha (CNTFRα) in retinal ganglion cells (RGCs), the projection neurons of the retina, but were diminished by depleting neutrophils or by genetically suppressing monocyte infiltration. CNTF gene therapy increased expression of C-C motif chemokine ligand 5 (CCL5) in immune cells and retinal glia, and recombinant CCL5 induced extensive axon regeneration. Conversely, CRISPR-mediated knockdown of the cognate receptor (CCR5) in RGCs or treating wild-type mice with a CCR5 antagonist repressed the effects of CNTF gene therapy. Thus, CCL5 is a previously unrecognized, potent activator of optic nerve regeneration and mediates many of the effects of CNTF gene therapy.

Like most pathways in the mature central nervous system (CNS), the optic nerve cannot regenerate once damaged due in part to cell-extrinsic suppressors of axon growth (1, 2) and the low intrinsic growth capacity of adult retinal ganglion cells (RGCs), the projection neurons of the eye (3–5). Consequently, traumatic or ischemic optic nerve injury or degenerative diseases such as glaucoma lead to irreversible visual losses. Experimentally, some degree of regeneration can be induced by intraocular inflammation or growth factors expressed by inflammatory cells (6–10), altering the cell-intrinsic growth potential of RGCs (3–5), enhancing physiological activity (11, 12), chelating free zinc (13, 14), and other manipulations (15–19). However, the extent of regeneration achieved to date remains modest, underlining the need for more effective therapies.Ciliary neurotrophic factor (CNTF) is a leading therapeutic candidate for glaucoma and other ocular diseases (20–23). Activation of the downstream signal transduction cascade requires CNTF to bind to CNTF receptor-α (CNTFRα) (24), which leads to recruitment of glycoprotein 130 (gp130) and leukemia inhibitory factor receptor-β (LIFRβ) to form a tripartite receptor complex (25). CNTFRα anchors to the plasma membrane through a glycosylphosphatidylinositol linkage (26) and can be released and become soluble through phospholipase C-mediated cleavage (27). CNTF has been reported to activate STAT3 phosphorylation in retinal neurons, including RGCs, and to promote survival, but it is unknown whether these effects are mediated by direct action of CNTF on RGCs via CNTFRα (28). Our previous studies showed that CNTF promotes axon outgrowth from neonate RGCs in culture (29) but fails to do so in cultured mature RGCs (8) or in vivo (6). Although some studies report that recombinant CNTF (rCNTF) can promote optic nerve regeneration (20, 30, 31), others find little or no effect unless SOCS3 (suppressor of cytokine signaling-3), an inhibitor of the Jak-STAT pathway, is deleted in RGCs (5, 6, 32). In contrast, multiple studies show that adeno-associated virus (AAV)-mediated expression of CNTF in RGCs induces strong regeneration (33–40). The basis for the discrepant effects of rCNTF and CNTF gene therapy is unknown but is of considerable interest in view of the many promising clinical and preclinical outcomes obtained with CNTF to date.Because intravitreal virus injections induce inflammation (41), we investigated the possibility that CNTF, a known immune modulator (42–44), might act by elevating expression of other immune-derived factors. We report here that the beneficial effects of CNTF gene therapy in fact require immune system activation and elevation of C-C motif chemokine ligand 5 (CCL5). Depletion of neutrophils, global knockout (KO) or RGC-selective deletion of the CCL5 receptor CCR5, or a CCR5 antagonist all suppress the effects of CNTF gene therapy, whereas recombinant CCL5 (rCCL5) promotes axon regeneration and increases RGC survival. These studies point to CCL5 as a potent monotherapy for optic nerve regeneration and to the possibility that other applications of CNTF and other forms of gene therapy might similarly act indirectly through other factors.     

Crystal structure of a far-red–sensing cyanobacteriochrome reveals an atypical bilin conformation and spectral tuning mechanism

Cyanobacteriochromes (CBCRs) are small, linear tetrapyrrole (bilin)-binding photoreceptors in the phytochrome superfamily that regulate diverse light-mediated adaptive processes in cyanobacteria. More spectrally diverse than canonical red/far-red–sensing phytochromes, CBCRs were thought to be restricted to sensing visible and near UV light until recently when several subfamilies with far-red–sensing representatives (frCBCRs) were discovered. Two of these frCBCRs subfamilies have been shown to incorporate bilin precursors with larger pi-conjugated chromophores, while the third frCBCR subfamily uses the same phycocyanobilin precursor found in the bulk of the known CBCRs. To elucidate the molecular basis of far-red light perception by this third frCBCR subfamily, we determined the crystal structure of the far-red–absorbing dark state of one such frCBCR Anacy_2551g3 from Anabaena cylindrica PCC 7122 which exhibits a reversible far-red/orange photocycle. Determined by room temperature serial crystallography and cryocrystallography, the refined 2.7-Å structure reveals an unusual all-Z,syn configuration of the phycocyanobilin (PCB) chromophore that is considerably less extended than those of previously characterized red-light sensors in the phytochrome superfamily. Based on structural and spectroscopic comparisons with other bilin-binding proteins together with site-directed mutagenesis data, our studies reveal protein–chromophore interactions that are critical for the atypical bathochromic shift. Based on these analyses, we propose that far-red absorption in Anacy_2551g3 is the result of the additive effect of two distinct red-shift mechanisms involving cationic bilin lactim tautomers stabilized by a constrained all-Z,syn conformation and specific interactions with a highly conserved anionic residue.

Cyanobacteria have developed elaborate, spectrally tuned photoreceptors and light-harvesting systems for adaptation and survival in a wide range of ecological niches (1–5). Many photoreceptor systems are modular components of much larger signaling proteins that integrate different sensor and effector modules into a single protein molecule to interface with diverse signal transduction pathways. Photoreceptors in the phytochrome superfamily utilize a specific lineage of GAF (cGMP phosphodiesterase, adenylyl cyclase and FhlA) domain that binds a thioether-linked linear tetrapyrrole (bilin) chromophore for light perception (6–11). Bilin-based photoreceptors play critical roles in plant development as well as in regulating cyanobacterial phototaxis, development, and light harvesting (2, 3, 12–17). Protein structural changes following the primary photochemical event then alter the downstream enzymatic activities and/or protein–protein interactions via an interdomain allosteric mechanism (18).Phytochromes possess a tripartite photosensory region consisting of three N-terminal domains (PAS, GAF, and PHY), known as the photosensory core module, in which the PAS and GAF domains are tethered via a “figure-eight knot” (14, 19, 20). In prototypical phytochromes, the bilin chromophore embedded in the GAF domain adopts a protonated 5-Z,syn, 10-Z,syn, 15-Z,anti configuration in the dark-adapted state. Light absorption triggers photoisomerization of the 15,16 double bond to generate a 15E,anti photoproduct, which typically absorbs far-red light (9, 14, 21). A long extension from the adjacent PHY domain is responsible for stabilizing the far-red–absorbing Pfr state (14, 20). In cyanobacteria, the phytochrome superfamily has diversified to yield a large family of more streamlined sensors, designated cyanobacteriochromes (CBCRs) (2, 4, 22–26). Unlike canonical phytochromes, CBCR photosensory modules consist of one or more GAF domains that are sufficient for covalent attachment of bilin and photoconversion. These small CBCR domains have also been used as light-sensing modules in a variety of synthetic biology applications (27–32). In contrast to canonical red/far-red phytochromes, CBCRs are able to sense light from near UV to far-red, utilizing a common phycocyanobilin (PCB) chromophore precursor (22–24, 26).The remarkable spectral diversity of CBCRs (SI Appendix, Fig. S1A) arises from extensive molecular evolution of the GAF domain scaffold. Many CBCRs leverage two thioether linkages to sense blue, violet, or near-UV light (8, 22, 23, 25, 33–35). Such “two-Cys” CBCRs possess an additional thioether linkage to the C10 methine bridge of the bilin that splits the chromophore in half, significantly shortening the conjugated π-system. Rupture of this covalent bond can occur upon 15Z/15E photoisomerization, which restores bilin conjugation across C10 to generate a photostate absorbing at wavelengths from teal to red (8, 33, 36, 37). Dual cysteine CBCRs have evolved multiple times, yielding a wide range of photocycles with (ultra)violet, blue, teal, green, orange, and red states (22).Red/green CBCRs such as AnPixJg2 and NpR6012g4 have red-absorbing dark states similar to phytochromes that photoconvert to green-absorbing lit states. In this CBCR subfamily, the molecular mechanism responsible for photoproduct tuning relies on trapping the 15E bilin in a twisted geometry that results in blue-shifted absorption (10, 11). In contrast, green/red CBCRs exhibit a reversed photocycle: the green-absorbing 15Z dark state photoconverts to yield a red-absorbing 15E photoproduct. This subfamily uses a protochromic mechanism first reported for the light-regulated histidine kinase RcaE (SI Appendix, Fig. S1B) in which photoconversion triggers a proton transfer to an uncharged chromophore inducing a spectral red shift (2, 38).Until recently, the light-sensing range of CBCRs appeared limited to the visible spectrum, thereby implicating phytochromes to be exclusively responsible for far-red sensing in cyanobacteria. Indeed, far-red–dependent remodeling of the photosynthetic apparatus in multiple cyanobacterial species is mediated by the red/far-red phytochrome RfpA (3, 39). The discovery of two lineages of CBCRs with far-red-absorbing dark states (frCBCRs) was thus surprising (40). Upon far-red light absorption, these frCBCRs convert to either an orange- or red-absorbing photoproduct state. These frCBCRs evolved from green/red CBCRs as part of a greater green/red (GGR) lineage and independent from evolution of other frCBCRs within the XRG (extended red/green) lineage (35, 40, 41). Owing to their small size and spectral overlap with the therapeutic window of optimum tissue penetrance (700 to 800 nm) (42–46), frCBCRs represent tantalizing scaffolds for development of FR-responsive optogenetic reagents for biomedical research and imaging applications (45, 47–50).To understand the molecular basis of far-red spectral tuning of the frCBCR family that evolved within GGR lineage, we determined the crystal structures of the FR-absorbing dark state of the representative FR/O CBCR Anacy_2551g3 from Anabaena cylindrica PCC 7122 at both ambient and cryogenic temperatures. These structures revealed an all-Z,syn configuration of its PCB chromophore that differs from those found in all known CBCRs and phytochromes. Based upon these crystallographic results, spectra of site-directed mutants of Anacy_2551g3 and related frCBCRs in the GGR lineage, and comparisons with other bilin-binding proteins, we identify key protein–chromophore interactions that support two tuning mechanisms simultaneously at work for far-red light detection in this family of frCBCRs.     

Circular swimming motility and disordered hyperuniform state in an algae system

Active matter comprises individually driven units that convert locally stored energy into mechanical motion. Interactions between driven units lead to a variety of nonequilibrium collective phenomena in active matter. One of such phenomena is anomalously large density fluctuations, which have been observed in both experiments and theories. Here we show that, on the contrary, density fluctuations in active matter can also be greatly suppressed. Our experiments are carried out with marine algae (Effrenium voratum), which swim in circles at the air–liquid interfaces with two different eukaryotic flagella. Cell swimming generates fluid flow that leads to effective repulsions between cells in the far field. The long-range nature of such repulsive interactions suppresses density fluctuations and generates disordered hyperuniform states under a wide range of density conditions. Emergence of hyperuniformity and associated scaling exponent are quantitatively reproduced in a numerical model whose main ingredients are effective hydrodynamic interactions and uncorrelated random cell motion. Our results demonstrate the existence of disordered hyperuniform states in active matter and suggest the possibility of using hydrodynamic flow for self-assembly in active matter.

Active matter exists over a wide range of spatial and temporal scales (1–6) from animal groups (7, 8) to robot swarms (9–11), to cell colonies and tissues (12–16), to cytoskeletal extracts (17–20), to man-made microswimmers (21–25). Constituent particles in active matter systems are driven out of thermal equilibrium at the individual level; they interact to develop a wealth of intriguing collective phenomena, including clustering (13, 22, 24), flocking (11, 26), swarming (12, 13), spontaneous flow (14, 20), and giant density fluctuations (10, 11). Many of these observed phenomena have been successfully described by particle-based or continuum models (1–6), which highlight the important roles of both individual motility and interparticle interactions in determining system dynamics.Current active matter research focuses primarily on linearly swimming particles which have a symmetric body and self-propel along one of the symmetry axes. However, a perfect alignment between the propulsion direction and body axis is rarely found in reality. Deviation from such a perfect alignment leads to a persistent curvature in the microswimmer trajectories; examples of such circle microswimmers include anisotropic artificial micromotors (27, 28), self-propelled nematic droplets (29, 30), magnetotactic bacteria and Janus particles in rotating external fields (31, 32), Janus particle in viscoelastic medium (33), and sperm and bacteria near interfaces (34, 35). Chiral motility of circle microswimmers, as predicted by theoretical and numerical investigations, can lead to a range of interesting collective phenomena in circular microswimmers, including vortex structures (36, 37), localization in traps (38), enhanced flocking (39), and hyperuniform states (40). However, experimental verifications of these predictions are limited (32, 35), a situation mainly due to the scarcity of suitable experimental systems.Here we address this challenge by investigating marine algae Effrenium voratum (41, 42). At air–liquid interfaces, E. voratum cells swim in circles via two eukaryotic flagella: a transverse flagellum encircling the cellular anteroposterior axis and a longitudinal one running posteriorly. Over a wide range of densities, circling E. voratum cells self-organize into disordered hyperuniform states with suppressed density fluctuations at large length scales. Hyperuniformity (43, 44) has been considered as a new form of material order which leads to novel functionalities (45–49); it has been observed in many systems, including avian photoreceptor patterns (50), amorphous ices (51), amorphous silica (52), ultracold atoms (53), soft matter systems (54–61), and stochastic models (62–64). Our work demonstrates the existence of hyperuniformity in active matter and shows that hydrodynamic interactions can be used to construct hyperuniform states.