Cells communicate via extracellular vesicles (EVs) containing functional RNAs, proteins, and lipids. Knowledge on the fate of internalized EVs, especially their capacity to fuse with target cell membranes and deliver luminal cargo, is limited. Currently available EV-cargo delivery assays are indirect and thus unlikely to uncover molecular players and conditions that specifically control the EV-fusion step. Here, we present a novel live-cell imaging assay for detection of EV-binding, -uptake, and -fusion in time and space. We employed the SunTag system for exceptional signal amplification. EV-donor cells were engineered to tag the luminal EV-membrane with a fluorescent label coupled to SunTag peptides. Recipient cells express fluorescent single-chain anti-SunTag antibody (STAb), which binds EV-enclosed SunTag upon its cytosolic exposure. Using SunTagged EVs carrying fusogen VSV-G, we visualize the EV-fusion process, quantify fusion kinetics and efficiency, and determine subcellular localization of fusion events. We term this methodology the Extracellular Vesicle Fusion Spatiotemporal Imaging Method (EV-FUSIM). In the future, this technology can support the identification of fusogenic EV-subsets, as well as molecular players and drugs that modulate EV-fusion, without confounding effects of post-fusion processes. This will extend knowledge on EV-biology and can aid in the engineering of EVs that efficiently deliver intraluminal therapeutic payloads.

Recent discoveries have shown the presence of ribonucleic acid (RNA) on the cell surface, defying the view that RNA only functions intracellularly. However, how RNA is presented on the cell surface and what its biological relevance is are poorly understood. We established Toll-like receptor 7 (TLR7) as a cell-surface RNA (csRNA) probe. Employing it in a genome-wide knockout screening, we identified heparan sulfate (HS) as a crucial factor for csRNA presentation. Cell-surface proximity labeling revealed that HS-associated csRNAs (hepRNAs) are in the vicinity of RNA-binding proteins (RBPs). These observations led us to a model wherein cell-surface HS, RNA, and RBP form ternary complexes, validated by our spatio-selective RNA-protein crosslinking technology in a TLR7-orthogonal manner. We further revealed the identities of hepRNA and found that they can recruit the immune receptor killer cell immunoglobulin-like receptor 2DL5 (KIR2DL5), potentially enhancing receptor-ligand interactions. Employing human cell lines, our findings lay the groundwork for investigating how cell-surface ribonucleoproteins contribute to immune modulation.

The WEE1 kinase negatively regulates CDK1/2 to control DNA replication and mitotic entry. Genetic factors that determine sensitivity to WEE1 inhibitors (WEE1i) are largely unknown. A genome-wide insertional mutagenesis screen revealed that mutation of EIF2A, a translation regulator, sensitized to WEE1i. Additionally, a genome-wide CRISPR-Cas9 screen revealed that inactivation of integrated stress response (ISR) kinase GCN2 or its co-factor GCN1 rescued WEE1i-mediated cytotoxicity. Conversely, loss of the collided ribosome sensor ZNF598 increased sensitivity to WEE1i. Mechanistically, WEE1i induced paradoxical GCN2 activation, ATF4 upregulation, and altered ribosome dynamics. ISR activation was independent of WEE1 presence, pointing at off-target GCN2 engagement by multiple chemically distinct WEE1i. ISR activation was observed in cancer cells as well as non-transformed cells, and required GCN1 and ongoing translation. Consequently, WEE1i induce multiple independent cellular effects: DNA damage, premature mitotic entry and sensitization to DNA-damaging chemotherapeutics in an ISR-independent fashion, as well as ISR activation independently of CDK1/2 activation. Importantly, low-dose WEE1 inhibition did not induce ISR activation, while it still synergized with PKMYT1 inhibition. Taken together, WEE1i trigger toxic ISR activation and translational shutdown, which can be prevented by low-dose or combination treatments, while retaining the cell cycle checkpoint-perturbing effects.

Enteroviruses dramatically remodel the cellular infrastructure for efficient replication and curtailing host antiviral responses. The roles of viral proteins in these processes have been studied mostly in vitro, by ectopic overexpression, or by surrogate infection systems, all of which have shortcomings. Here, we replace the essential 2A cleavage site at the P1-P2 junction with an internal ribosome entry site (IRES), 3CD cleavage site, or T2A sequence, allowing us to catalytically inactivate 2Apro in the virus context. Viruses with an inactive 2A^pro are hampered in replication in cell lines and are severely attenuated in a Coxsackievirus B3 (CVB3) mouse pancreatitis infection model. We show that 2A^pro is essential for disturbing nucleocytoplasmic transport, shutting down host mRNA translation, suppressing stress granule formation, suppressing the induction of the IFN response, and overcoming IFN-induced restriction factors. Moreover, using an advanced single-molecule live cell imaging approach, we reveal that 2A^pro is important for the initial round of replication of the incoming viral RNA, which is a bottleneck for efficient infection. Thus, 2A^pro plays a critical role in subverting antiviral responses and establishing a favorable environment to expedite enterovirus replication.

To understand viral infection and virus–host interactions, real-time, single-cell assays to track viral infection progression are essential. Many conventional assays sample large numbers of cells for single measurements, averaging out the cell-to-cell heterogeneity that is intrinsic to viral infection. Moreover, conventional assays often require cell fixation or lysis, limiting analysis to a single timepoint and masking the temporal and spatial dynamics of infection. We have developed virus infection real-time imaging (VIRIM), a method to visualize the translation of individual RNAs of viruses in real-time. The single-molecule and live-cell nature of VIRIM allows the examination of the earliest events of viral infection, when viral protein and RNA levels are still low, and allows study into the origins and consequences of cell-to-cell heterogeneity during virus infection. Here we provide a step-by-step description of the VIRIM assay, including a detailed procedure for designing, producing and validating the viruses required for VIRIM. In addition, we provide guidelines for generating the reporter cell line, performing the time-lapse imaging and analyzing the fluorescence microscopy data. Once established, a typical VIRIM experiment requires 2–5 days to complete.

A recent ground-breaking study suggested that small RNA from mammalian cells can undergo N-glycan modifications (termed glycoRNA)¹. The discovery relied upon a metabolic glycan labeling strategy in combination with commonly used phase-separation-based RNA isolation. Following the reported procedure, here we likewise identify an N-glycosylated species in the RNA fraction. However, our results suggest that the reported RNase sensitivity of the glycosylated species depends on the specific RNA purification method. This suggests the possibility of copurifying unexpected RNase-insensitive N-glycoconjugates during glycoRNA isolation. The co-existence of two independent, yet highly similar molecular entities, complicates biochemical assays on glycoRNA and calls for more specific approaches for glycoRNA analysis. To address this, we propose a control experiment that can help distinguish genuine glycoRNA species from copurified glycoconjugates.

Respiratory syncytial virus (RSV) is the most prevalent cause of acute lower respiratory infection in young children. Currently, the first RSV vaccines are approved by the FDA. Recently, N6-methyladenosine (m⁶A) RNA methylation has been implicated in the regulation of the viral life cycle and replication of many viruses, including RSV. m⁶A methylation of RSV RNA has been demonstrated to promote replication and prevent anti-viral immune responses by the host. Whether m⁶A is also involved in viral entry and whether m⁶A can also affect RSV infection via different mechanisms than methylation of viral RNA is poorly understood. Here, we identify m⁶A reader YTH domain-containing protein 1 (YTHDC1) as a novel negative regulator of RSV infection. We demonstrate that YTHDC1 abrogates RSV infection by reducing the expression of RSV entry receptor CX3C motif chemokine receptor 1 (CX3CR1) on the cell surface of lung epithelial cells. Altogether, these data reveal a novel role for m⁶A methylation and YTHDC1 in the viral entry of RSV. These findings may contribute to the development of novel treatment options to control RSV infection.

Single-molecule fluorescence imaging experiments generally require sub-nanomolar protein concentrations to isolate single protein molecules, which makes such experiments challenging in live cells due to high intracellular protein concentrations. Here, we show that single-molecule observations can be achieved in live cells through a drastic reduction in the observation volume using overmilled zero-mode waveguides (ZMWs- subwavelength-size holes in a metal film). Overmilling of the ZMW in a palladium film creates a nanowell of tunable size in the glass layer below the aperture, which cells can penetrate. We present a thorough theoretical and experimental characterization of the optical properties of these nanowells over a wide range of ZMW diameters and overmilling depths, showing an excellent signal confinement and a 5-fold fluorescence enhancement of fluorescent molecules inside nanowells. ZMW nanowells facilitate live-cell imaging as cells form stable protrusions into the nanowells. Importantly, the nanowells greatly reduce the cytoplasmic background fluorescence, enabling the detection of individual membrane-bound fluorophores in the presence of high cytoplasmic expression levels, which could not be achieved with TIRF microscopy. Zero-mode waveguide nanowells thus provide great potential to study individual proteins in living cells.

Antiviral signalling, which can be activated in host cells upon virus infection, restricts virus replication and communicates infection status to neighbouring cells. The antiviral response is heterogeneous, both quantitatively (efficiency of response activation) and qualitatively (transcribed antiviral gene set). To investigate the basis of this heterogeneity, we combined Virus Infection Real-time IMaging (VIRIM), a live-cell single-molecule imaging method, with real-time readouts of the dsRNA sensing pathway to analyse the response of human cells to encephalomyocarditis virus (EMCV) infection. We find that cell-to-cell heterogeneity in viral replication rates early in infection affect the efficiency of antiviral response activation, with lower replication rates leading to more antiviral response activation. Furthermore, we show that qualitatively distinct antiviral responses can be linked to the strength of the antiviral signalling pathway. Our analyses identify variation in early viral replication rates as an important parameter contributing to heterogeneity in antiviral response activation.