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Nonsense-mediated mRNA decay (NMD) is an mRNA degradation pathway that eliminates transcripts containing premature termination codons (PTCs). Half-lives of the mRNAs containing PTCs demonstrate that a small percent escape surveillance and do not degrade. It is not known whether this escape represents variable mRNA degradation within cells or, alternatively cells within the population are resistant. Here we demonstrate a single-cell approach with a bi-directional reporter, which expresses two β-globin genes with or without a PTC in the same cell, to characterize the efficiency of NMD in individual cells. We found a broad range of NMD efficiency in the population; some cells degraded essentially all of the mRNAs, while others escaped NMD almost completely. Characterization of NMD efficiency together with NMD regulators in single cells showed cell-to-cell variability of NMD reflects the differential level of surveillance factors, SMG1 and phosphorylated UPF1. A single-cell fluorescent reporter system that enabled detection of NMD using flow cytometry revealed that this escape occurred either by translational readthrough at the PTC or by a failure of mRNA degradation after successful translation termination at the PTC. Here the author developed a single-cell reporter system to identify cell-to-cell variability of nonsense-mediated mRNA decay (NMD). This approach provides a sensitive tool to investigate cellular heterogeneity of NMD in various physiological conditions.

Translation is the fundamental biological process converting mRNA information into proteins. Single-molecule imaging in live cells has illuminated the dynamics of RNA transcription; however, it is not yet applicable to translation. Here, we report single-molecule imaging of nascent peptides (SINAPS) to assess translation in live cells. The approach provides direct readout of initiation, elongation, and location of translation. We show that mRNAs coding for endoplasmic reticulum (ER) proteins are translated when they encounter the ER membrane. Single-molecule fluorescence recovery after photobleaching provides direct measurement of elongation speed (5 amino acids per second). In primary neurons, mRNAs are translated in proximal dendrites but repressed in distal dendrites and display \"bursting\" translation. This technology provides a tool with which to address the spatiotemporal translation mechanism of single mRNAs in living cells.

MicroRNAs (miRNAs) are small non-coding RNAs, which regulate the expression of thousands of genes; miRNAs silence gene expression from complementary mRNAs through translational repression and mRNA decay. For decades, the function of miRNAs has been studied primarily by ensemble methods, where a bulk collection of molecules is measured outside cells. Thus, the behavior of individual molecules during miRNA-mediated gene silencing, as well as their spatiotemporal regulation inside cells, remains mostly unknown. Here we report single-molecule methods to visualize each step of miRNA-mediated gene silencing in situ inside cells. Simultaneous visualization of single mRNAs, translation, and miRNA-binding revealed that miRNAs preferentially bind to translated mRNAs rather than untranslated mRNAs. Spatiotemporal analysis based on our methods uncovered that miRNAs bind to mRNAs immediately after nuclear export. Subsequently, miRNAs induced translational repression and mRNA decay within 30 and 60 min, respectively, after the binding to mRNAs. This methodology provides a framework for studying miRNA function at the single-molecule level with spatiotemporal information inside cells. For decades, miRNAs have been studied primarily by ensemble methods, where a bulk collection of molecules is measured outside cells. Here, Kobayashi and Singer report methods to image miRNA function at the single-molecule level inside cells.

Key Points The asymmetrical distribution of mRNAs in cells is used by various organisms to spatially control gene expression. RNA localization has a role in diverse biological processes, such as development, cell motility, neuron connectivity and mating type switching in yeast. Recent technical advents and the development of new methods for mRNA detection in live and fixed cells allow the tracking and quantification of single mRNAs in a variety of cell types. Single-molecule imaging of mRNA in fixed and live cells revealed a complex cooperativity between RNA-binding proteins (RBPs) and motor proteins to regulate active transport of mRNAs. The composition of mRNA–protein (mRNP) complexes is intricate, and future research will reveal how they assemble into RNA granules with unique localization and functions. Neurons and unicellular organisms, such as yeast and bacteria, use both convergent and disparate mechanisms of targeting mRNAs to different regions. The subcellular localization of mRNAs enables the spatial regulation of protein translation and generates functional and structural asymmetries in cells. New imaging (and other) techniques for tracking single-mRNA dynamics have unravelled mechanisms of mRNA movements and localization patterns in various cell types. The spatial regulation of protein translation is an efficient way to create functional and structural asymmetries in cells. Recent research has furthered our understanding of how individual cells spatially organize protein synthesis, by applying innovative technology to characterize the relationship between mRNAs and their regulatory proteins, single-mRNA trafficking dynamics, physiological effects of abrogating mRNA localization in vivo and for endogenous mRNA labelling. The implementation of new imaging technologies has yielded valuable information on mRNA localization, for example, by observing single molecules in tissues. The emerging movements and localization patterns of mRNAs in morphologically distinct unicellular organisms and in neurons have illuminated shared and specialized mechanisms of mRNA localization, and this information is complemented by transgenic and biochemical techniques that reveal the biological consequences of mRNA mislocalization.

Direct visualization of genomic loci in the 3D nucleus is important for understanding the spatial organization of the genome and its association with gene expression. Various DNA FISH methods have been developed in the past decades, all involving denaturing dsDNA and hybridizing fluorescent nucleic acid probes. Here we report a novel approach that uses in vitro constituted nuclease-deficient clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated caspase 9 (Cas9) complexes as probes to label sequence-specific genomic loci fluorescently without global DNA denaturation (Cas9-mediated fluorescence in situ hybridization, CASFISH). Using fluorescently labeled nuclease-deficient Cas9 (dCas9) protein assembled with various single-guide RNA (sgRNA), we demonstrated rapid and robust labeling of repetitive DNA elements in pericentromere, centromere, G-rich telomere, and coding gene loci. Assembling dCas9 with an array of sgRNAs tiling arbitrary target loci, we were able to visualize nonrepetitive genomic sequences. The dCas9/sgRNA binary complex is stable and binds its target DNA with high affinity, allowing sequential or simultaneous probing of multiple targets. CASFISH assays using differently colored dCas9/sgRNA complexes allow multicolor labeling of target loci in cells. In addition, the CASFISH assay is remarkably rapid under optimal conditions and is applicable for detection in primary tissue sections. This rapid, robust, less disruptive, and cost-effective technology adds a valuable tool for basic research and genetic diagnosis.

A simple and general chemical structure change to a panel of cell-permeable small-molecule fluorophores increases their brightness and photostability, which will enable improved single-molecule studies and super-resolution imaging. Specific labeling of biomolecules with bright fluorophores is the keystone of fluorescence microscopy. Genetically encoded self-labeling tag proteins can be coupled to synthetic dyes inside living cells, resulting in brighter reporters than fluorescent proteins. Intracellular labeling using these techniques requires cell-permeable fluorescent ligands, however, limiting utility to a small number of classic fluorophores. Here we describe a simple structural modification that improves the brightness and photostability of dyes while preserving spectral properties and cell permeability. Inspired by molecular modeling, we replaced the N , N -dimethylamino substituents in tetramethylrhodamine with four-membered azetidine rings. This addition of two carbon atoms doubles the quantum efficiency and improves the photon yield of the dye in applications ranging from in vitro single-molecule measurements to super-resolution imaging. The novel substitution is generalizable, yielding a palette of chemical dyes with improved quantum efficiencies that spans the UV and visible range.

Cellular messenger RNA levels are achieved by the combinatorial complexity of factors controlling transcription, yet the small number of molecules involved in these pathways fluctuates stochastically. It has not yet been experimentally possible to observe the activity of single polymerases on an endogenous gene to elucidate how these events occur in vivo. Here, we describe a method of fluctuation analysis of fluorescently labeled RNA to measure dynamics of nascent RNA—including initiation, elongation, and termination—at an active yeast locus. We find no transcriptional memory between initiation events, and elongation speed can vary by threefold throughout the cell cycle. By measuring the abundance and intranuclear mobility of an upstream transcription factor, we observe that the gene firing rate is directly determined by trans-activating factor search times.

RNA–protein interactions are essential for proper gene expression regulation, particularly in neurons with unique spatial constraints. Currently, these interactions are defined biochemically, but a method is needed to evaluate them quantitatively within morphological context. Colocalization of two-color labels using wide-field microscopy is a method to infer these interactions. However, because of chromatic aberrations in the objective lens, this approach lacks the resolution to determine whether two molecules are physically in contact or simply nearby by chance. Here, we developed a robust super registration methodology that corrected the chromatic aberration across the entire image field to within 10 nm, which is capable of determining whether two molecules are physically interacting or simply in proximity by random chance. We applied this approach to image single-molecule FISH in combination with immunofluorescence (smFISH-IF) and determined whether the association between an mRNA and binding protein(s) within a neuron was significant or accidental. We evaluated several mRNA-binding proteins identified from RNA pulldown assays to determine which of these exhibit bona fide interactions. Surprisingly, many known mRNA-binding proteins did not bind the mRNA in situ, indicating that adventitious interactions are significant using existing technology. This method provides an ability to evaluate two-color registration compatible with the scale of molecular interactions.

A graphical processing unit implementation of an efficient Bayesian-based multiview deconvolution method brings the resolution and contrast advantages of multiview deconvolution to more users of light-sheet fluorescence microscopy. Light-sheet fluorescence microscopy is able to image large specimens with high resolution by capturing the samples from multiple angles. Multiview deconvolution can substantially improve the resolution and contrast of the images, but its application has been limited owing to the large size of the data sets. Here we present a Bayesian-based derivation of multiview deconvolution that drastically improves the convergence time, and we provide a fast implementation using graphics hardware.

Analysis of single molecules in living cells has provided quantitative insights into the kinetics of fundamental biological processes; however, the dynamics of messenger RNA (mRNA) translation have yet to be addressed. We have developed a fluorescence microscopy technique that reports on the first translation events of individual mRNA molecules. This allowed us to examine the spatiotemporal regulation of translation during normal growth and stress and during Drosophila oocyte development. We have shown that mRNAs are not translated in the nucleus but translate within minutes after export, that sequestration within P-bodies regulates translation, and that oskar mRNA is not translated until it reaches the posterior pole of the oocyte. This methodology provides a framework for studying initiation of protein synthesis on single mRNAs in living cells.