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Go with the actomyosin flow Cortical flows of actomyosin are central to many processes in cellular and developmental biology. In the one-cell Caenorhabditis elegans embryo, anteroposterior polarity, a prerequisite for asymmetric cell division, is established by large-scale flow of the actomyosin cortex that segregates cortical polarity proteins between the anterior and posterior domains. The underlying forces and physical principles behind long-range flow remain unclear. Mayer et al . have devised a novel method to measure cortical tension (total mechanical tension) and find that cortical flows are driven by contractility of the actomyosin network. The direction of flow depends on anisotropies in the cortical tension, and long-range cortical flow occurs only if the cortex is sufficiently viscous. In the one-cell Caenorhabditis elegans embryo, anteroposterior polarization is facilitated by large-scale flow of the actomyosin cortex, which segregates cortical polarity proteins into anterior and posterior domains. The underlying forces and physical principles behind long-range flow are unclear. Here, a new method is described by which to measure cortical tension. The results identify two prerequisites for large-scale cortical flow: a gradient in actomyosin contractility and a sufficiently large viscosity of the cortex. Asymmetric cell divisions are essential for the development of multicellular organisms. To proceed, they require an initially symmetric cell to polarize 1 . In Caenorhabditis elegans zygotes, anteroposterior polarization is facilitated by a large-scale flow of the actomyosin cortex 2 , 3 , 4 , which directs the asymmetry of the first mitotic division. Cortical flows appear in many contexts of development 5 , but their underlying forces and physical principles remain poorly understood. How actomyosin contractility and cortical tension interact to generate large-scale flow is unclear. Here we report on the subcellular distribution of cortical tension in the polarizing C. elegans zygote, which we determined using position- and direction-sensitive laser ablation. We demonstrate that cortical flow is associated with anisotropies in cortical tension and is not driven by gradients in cortical tension, which contradicts previous proposals 5 . These experiments, in conjunction with a theoretical description of active cortical mechanics, identify two prerequisites for large-scale cortical flow: a gradient in actomyosin contractility to drive flow and a sufficiently large viscosity of the cortex to allow flow to be long-ranged. We thus reveal the physical requirements of large-scale intracellular cortical flow that ensure the efficient polarization of the C. elegans zygote.

In cells, many vital processes involve myosin-driven motility that actively remodels the actin cytoskeleton and changes cell shape. Here we study how the collective action of myosin motors organizes actin filaments into contractile structures in a simplified model system devoid of biochemical regulation. We show that this self-organization occurs through an active multistage coarsening process. First, motors form dense foci by moving along the actin network structure followed by coalescence. Then the foci accumulate actin filaments in a shell around them. These actomyosin condensates eventually cluster due to motor-driven coalescence. We propose that the physical origin of this multistage aggregation is the highly asymmetric load response of actin filaments: they can support large tensions but buckle easily under piconewton compressive loads. Because the motor-generated forces well exceed this threshold, buckling is induced on the connected actin network that resists motor-driven filament sliding. We show how this buckling can give rise to the accumulation of actin shells around myosin foci and subsequent coalescence of foci into superaggregates. This new physical mechanism provides an explanation for the formation and contractile dynamics of disordered condensed actomyosin states observed in vivo.

The specificity of CRISPR/Cas9 genome editing is largely determined by the sequences of guide RNA (gRNA) and the targeted DNA, yet the sequence-dependent rules underlying off-target effects are not fully understood. To systematically explore the sequence determinants governing CRISPR/Cas9 specificity, here we describe a dual-target system to measure the relative cleavage rate between off- and on-target sequences (off-on ratios) of 1902 gRNAs on 13,314 synthetic target sequences, and reveal a set of sequence rules involving 2 factors in off-targeting: 1) a guide-intrinsic mismatch tolerance (GMT) independent of the mismatch context; 2) an “epistasis-like” combinatorial effect of multiple mismatches, which are associated with the free-energy landscape in R-loop formation and are explainable by a multi-state kinetic model. These sequence rules lead to the development of MOFF, a model-based predictor of Cas9-mediated off-target effects. Moreover, the “epistasis-like” combinatorial effect suggests a strategy of allele-specific genome editing using mismatched guides. With the aid of MOFF prediction, this strategy significantly improves the selectivity and expands the application domain of Cas9-based allele-specific editing, as tested in a high-throughput allele-editing screen on 18 cancer hotspot mutations. The sequence determinants governing CRISPR/Cas9 specificity are not fully understood. Here, the authors devise a high-throughput dual-target synthetic system to explore the sequence features associated with CRISPR/Cas9 off-target effect, reveal a set of sequence-dependent rules, and develop an off-target prediction model and a strategy for Cas9-based allele-specific editing.

The S. pyogenes (Sp) Cas9 endonuclease is an important gene-editing tool. Sp Cas9 is directed to target sites based on complementarity to a complexed single-guide RNA (sgRNA). However, Sp Cas9-sgRNA also binds and cleaves genomic off-targets with only partial complementarity. To date, we lack the ability to predict cleavage and binding activity quantitatively, and rely on binary classification schemes to identify strong off-targets. We report a quantitative kinetic model that captures the Sp Cas9-mediated strand-replacement reaction in free-energy terms. The model predicts binding and cleavage activity as a function of time, target, and experimental conditions. Trained and validated on high-throughput bulk-biochemical data, our model predicts the intermediate R-loop state recently observed in single-molecule experiments, as well as the associated conversion rates. Finally, we show that our quantitative activity predictor can be reduced to a binary off-target classifier that outperforms the established state-of-the-art. Our approach is extensible, and can characterize any CRISPR-Cas nuclease – benchmarking natural and future high-fidelity variants against Sp Cas9; elucidating determinants of CRISPR fidelity; and revealing pathways to increased specificity and efficiency in engineered systems. Cas9 off-target sites can be predicted by many bioinformatics tools. Here the authors present low complexity mechanistic model that characterizes SpCas9 kinetics in free-energy terms, allowing quantitative prediction of off-target activity in bulk-biochemistry, single molecule, and whole-genome profiling experiments.

The absence of ‘shovel-ready’ anti-coronavirus drugs during vaccine development has exceedingly worsened the SARS-CoV-2 pandemic. Furthermore, new vaccine-resistant variants and coronavirus outbreaks may occur in the near future, and we must be ready to face this possibility. However, efficient antiviral drugs are still lacking to this day, due to our poor understanding of the mode of incorporation and mechanism of action of nucleotides analogs that target the coronavirus polymerase to impair its essential activity. Here, we characterize the impact of remdesivir (RDV, the only FDA-approved anti-coronavirus drug) and other nucleotide analogs (NAs) on RNA synthesis by the coronavirus polymerase using a high-throughput, single-molecule, magnetic-tweezers platform. We reveal that the location of the modification in the ribose or in the base dictates the catalytic pathway(s) used for its incorporation. We show that RDV incorporation does not terminate viral RNA synthesis, but leads the polymerase into backtrack as far as 30 nt, which may appear as termination in traditional ensemble assays. SARS-CoV-2 is able to evade the endogenously synthesized product of the viperin antiviral protein, ddhCTP, though the polymerase incorporates this NA well. This experimental paradigm is essential to the discovery and development of therapeutics targeting viral polymerases. To multiply and spread from cell to cell, the virus responsible for COVID-19 (also known as SARS-CoV-2) must first replicate its genetic information. This process involves a ‘polymerase’ protein complex making a faithful copy by assembling a precise sequence of building blocks, or nucleotides. The only drug approved against SARS-CoV-2 by the US Food and Drug Administration (FDA), remdesivir, consists of a nucleotide analog, a molecule whose structure is similar to the actual building blocks needed for replication. If the polymerase recognizes and integrates these analogs into the growing genetic sequence, the replication mechanism is disrupted, and the virus cannot multiply. Most approaches to study this process seem to indicate that remdesivir works by stopping the polymerase and terminating replication altogether. Yet, exactly how remdesivir and other analogs impair the synthesis of new copies of the virus remains uncertain. To explore this question, Seifert, Bera et al. employed an approach called magnetic tweezers which uses a magnetic field to manipulate micro-particles with great precision. Unlike other methods, this technique allows analogs to be integrated under conditions similar to those found in cells, and to be examined at the level of a single molecule. The results show that contrary to previous assumptions, remdesivir does not terminate replication; instead, it causes the polymerase to pause and backtrack (which may appear as termination in other techniques). The same approach was then applied to other nucleotide analogs, some of which were also found to target the SARS-CoV-2 polymerase. However, these analogs are incorporated differently to remdesivir and with less efficiency. They also obstruct the polymerase in distinct ways. Taken together, the results by Seifert, Bera et al. suggest that magnetic tweezers can be a powerful approach to reveal how analogs interfere with replication. This information could be used to improve currently available analogs as well as develop new antiviral drugs that are more effective against SARS-CoV-2. This knowledge will be key at a time when treatments against COVID-19 are still lacking, and may be needed to protect against new variants and future outbreaks.

Transcription in bacteria is controlled by multiple molecular mechanisms that precisely regulate gene expression. It has been recently shown that initial RNA synthesis by the bacterial RNA polymerase (RNAP) is interrupted by pauses; however, the pausing determinants and the relationship of pausing with productive and abortive RNA synthesis remain poorly understood. Using single-molecule FRET and biochemical analysis, here we show that the pause encountered by RNAP after the synthesis of a 6-nt RNA (ITC6) renders the promoter escape strongly dependent on the NTP concentration. Mechanistically, the paused ITC6 acts as a checkpoint that directs RNAP to one of three competing pathways: productive transcription, abortive RNA release, or a new unscrunching/scrunching pathway. The cyclic unscrunching/scrunching of the promoter generates a long-lived, RNA-bound paused state; the abortive RNA release and DNA unscrunching are thus not as tightly linked as previously thought. Finally, our new model couples the pausing with the abortive and productive outcomes of initial transcription. RNA synthesis by bacterial RNA polymerase is interrupted by pauses but their role in RNA synthesis is poorly understood. Here the authors use single-molecule FRET and biochemical analysis to show that pausing regulates branching between the abortive and productive outcomes of initial transcription.

Significance The mechanism of RAD51-recombinase filament formation is visualized and quantified with single-molecule resolution using a combination of dual optical tweezers, fluorescence microscopy, and microfluidics. With this method, short-lived transient intermediates formed during nascent RAD51 filament assembly were observed directly. It is observed that RAD51 nuclei consisting of a variable number of monomers bind from solution to DNA, with an interaction time that increases with nucleus size. Nuclei that remain bound to DNA long enough can grow by the incorporation of additional RAD51 monomers, stabilizing the RAD51 filament. During recombinational repair of double-stranded DNA breaks, RAD51 recombinase assembles as a nucleoprotein filament around single-stranded DNA to form a catalytically proficient structure able to promote homology recognition and strand exchange. Mediators and accessory factors guide the action and control the dynamics of RAD51 filaments. Elucidation of these control mechanisms necessitates development of approaches to quantitatively probe transient aspects of RAD51 filament dynamics. Here, we combine fluorescence microscopy, optical tweezers, and microfluidics to visualize the assembly of RAD51 filaments on bare single-stranded DNA and quantify the process with single-monomer sensitivity. We show that filaments are seeded from RAD51 nuclei that are heterogeneous in size. This heterogeneity appears to arise from the energetic balance between RAD51 self-assembly in solution and the size-dependent interaction time of the nuclei with DNA. We show that nucleation intrinsically is substrate selective, strongly favoring filament formation on bare single-stranded DNA. Furthermore, we devised a single-molecule fluorescence recovery after photobleaching assay to independently observe filament nucleation and growth, permitting direct measurement of their contributions to filament formation. Our findings yield a comprehensive, quantitative understanding of RAD51 filament formation on bare single-stranded DNA that will serve as a basis to elucidate how mediators help RAD51 filament assembly and accessory factors control filament dynamics.

Argonaute (Ago) proteins are key players in both gene regulation (eukaryotes) and host defense (prokaryotes). Acting on single-stranded nucleic-acid substrates, Ago relies on base pairing between a small nucleic-acid guide and its complementary target sequences for specificity. To efficiently scan nucleic-acid chains for targets, Ago diffuses laterally along the substrate and must bypass secondary structures as well as protein barriers. Using single-molecule FRET in conjunction with kinetic modelling, we reveal that target scanning is mediated through loose protein-nucleic acid interactions, allowing Ago to slide short distances over secondary structures, as well as to bypass protein barriers via intersegmental transfer. Our combined single-molecule experiment and kinetic modelling approach may serve as a platform to dissect search processes and study the effect of sequence on search kinetics for other nucleic acid-guided proteins. Argonaute protein is loaded with small RNA and scans long stretch of sequences to find complementary target sites. Here, using single-molecule FRET and kinetic modelling, the authors showed that prokaryotic Argonaute protein binds target DNA loosely and slides along the DNA during target search.

The ubiquitous sliding clamp facilitates processivity of the replicative polymerase and acts as a platform to recruit proteins involved in replication, recombination and repair. While the dynamics of the E. coli β 2 -sliding clamp have been characterized in vitro , its in vivo stoichiometry and dynamics remain unclear. To probe both β 2 -clamp dynamics and stoichiometry in live E. coli cells, we use custom-built microfluidics in combination with single-molecule fluorescence microscopy and photoactivated fluorescence microscopy. We quantify the recruitment, binding and turnover of β 2 -sliding clamps on DNA during replication. These quantitative in vivo results demonstrate that numerous β 2 -clamps in E. coli remain on the DNA behind the replication fork for a protracted period of time, allowing them to form a docking platform for other enzymes involved in DNA metabolism. DNA replication is accomplished by the replisome, a multi-protein complex that comprises the sliding clamp. Here, Moolman et al . present quantitative and dynamic measurements of the number of β 2 -sliding clamps at the single-cell level in live E. coli cells to shed light on key aspects of DNA replication.

Tus protein bound to Ter sites on circular bacterial chromosomes provides a way to avoid random crashes of opposing replication forks. DNA-unzipping experiments show that the Tus– Ter –induced lock during unzipping at the nonpermissive face requires only DNA-strand separation. The bidirectional replication of a circular chromosome by many bacteria necessitates proper termination to avoid the head-on collision of the opposing replisomes. In Escherichia coli , replisome progression beyond the termination site is prevented by Tus proteins bound to asymmetric Ter sites. Structural evidence indicates that strand separation on the blocking (nonpermissive) side of Tus– Ter triggers roadblock formation, but biochemical evidence also suggests roles for protein-protein interactions. Here DNA unzipping experiments demonstrate that nonpermissively oriented Tus– Ter forms a tight lock in the absence of replicative proteins, whereas permissively oriented Tus– Ter allows nearly unhindered strand separation. Quantifying the lock strength reveals the existence of several intermediate lock states that are impacted by mutations in the lock domain but not by mutations in the DNA-binding domain. Lock formation is highly specific and exceeds reported in vivo efficiencies. We postulate that protein-protein interactions may actually hinder, rather than promote, proper lock formation.