Nucleic-acid detection via isothermal amplification and collateral cleavage of reporter molecules by CRISPR-associated enzymes is a promising alternative to quantitative polymerase chain reaction (qPCR). Here, we report the clinical validation of the SHERLOCK (specific high-sensitivity enzymatic reporter unlocking) assay using the enzyme Cas13a from Leptotrichia wadei for the detection of SARS-CoV-2 (severe acute respiratory syndrome coronavirus 2) -the virus that causes COVID-19 (coronavirus disease 2019) -in 154 nasopharyngeal and throat swab samples collected at Siriraj Hospital, Thailand. Within a detection limit of 42 RNA copies per

Nature uses 64 codons to encode the synthesis of proteins from the genome, and chooses 1 sense codon-out of up to 6 synonyms-to encode each amino acid. Synonymous codon choice has diverse and important roles, and many synonymous substitutions are detrimental. Here we demonstrate that the number of codons used to encode the canonical amino acids can be reduced, *

Biological microscopy would benefit from smaller alternatives to green fluorescent protein for imaging specific proteins in living cells. Here we introduce PRIME (PRobe Incorporation Mediated by Enzymes), a method for fluorescent labeling of peptide-fused recombinant proteins in living cells with high specificity. PRIME uses an engineered fluorophore ligase, which is derived from the natural Escherichia coli enzyme lipoic acid ligase (LplA). Through structureguided mutagenesis, we created a mutant ligase capable of recognizing a 7-hydroxycoumarin substrate and catalyzing its covalent conjugation to a transposable 13-amino acid peptide called LAP (LplA Acceptor Peptide). We showed that this fluorophore ligation occurs in cells in 10 min and that it is highly specific for LAP fusion proteins over all endogenous mammalian proteins. By genetically targeting the PRIME ligase to specific subcellular compartments, we were able to selectively label spatially distinct subsets of proteins, such as the surface pool of neurexin and the nuclear pool of actin.fluorescence microscopy | biotechnology | enzyme engineering T echniques for posttranslational labeling of proteins in living cells address some of the shortcomings of green fluorescent protein (GFP) by expanding the repertoire of chemical probes available for protein visualization (1). However, most of these methods, such as HaloTag (2), SNAP/CLIP (3), and DHFR (4), still use large protein tags that sterically interfere with protein trafficking and function, as GFP is known to do (5, 6). FlAsH (7) is the only peptide-based posttranslational labeling method that currently works inside living cells. At 12 amino acids, the FlAsH tag is much smaller than GFP, but poor labeling specificity (7-9), cellular toxicity, and undesired palmitoylation (7) and oxidation (8) of the Cys 4 recognition motif limit its utility.Here we introduce a method for protein labeling that utilizes a peptide tag while preserving high sequence specificity inside living cells. Our method, called PRIME (PRobe Incorporation Mediated by Enzymes), is based on a "fluorophore ligase" that is engineered from the Escherichia coli enzyme lipoic acid ligase (LplA). LplA's natural function is to ligate lipoic acid onto three E. coli proteins involved in oxidative metabolism (10). We previously used LplA for labeling of cell-surface proteins by demonstrating that the wild-type enzyme can ligate an azidoalkanoic acid instead of lipoic acid (11) (Fig. 1A, Middle). Ligated azide could then be chemoselectively derivatized using cyclooctynefluorophore conjugates.In this work, we wished to extend LplA-mediated labeling to intracellular proteins but recognized the challenges associated with our two-step labeling scheme. First, labeling sensitivity is limited by the kinetics of strain-promoted ½3 þ 2 cycloaddition, which has a rate constant of 4.3 × 10 −3 M −1 sec −1 (12). Second, for intracellular applications, two washout steps would be needed: first to remove excess azide, and second to remove excess fluorophore. We s...

The efficient, site-specific introduction of unnatural amino acids into proteins in mammalian cells is an outstanding challenge in realizing the potential of genetic code expansion approaches. Addressing this challenge will allow the synthesis of modified recombinant proteins and augment emerging strategies that introduce new chemical functionalities into proteins to control and image their function with high spatial and temporal precision in cells. The efficiency of unnatural amino acid incorporation in response to the amber stop codon (UAG) in mammalian cells is commonly considered to be low. Here we demonstrate that tRNA levels can be limiting for unnatural amino acid incorporation efficiency, and we develop an optimized pyrrolysyl-tRNA synthetase/tRNACUA expression system, with optimized tRNA expression for mammalian cells. In addition, we engineer eRF1, that normally terminates translation on all three stop codons, to provide a substantial increase in unnatural amino acid incorporation in response to the UAG codon without increasing readthrough of other stop codons. By combining the optimized pyrrolysyl-tRNA synthetase/tRNACUA expression system and an engineered eRF1, we increase the yield of protein bearing unnatural amino acids at a single site 17- to 20-fold. Using the optimized system, we produce proteins containing unnatural amino acids with comparable yields to a protein produced from a gene that does not contain a UAG stop codon. Moreover, the optimized system increases the yield of protein, incorporating an unnatural amino acid at three sites, from unmeasurably low levels up to 43% of a no amber stop control. Our approach may enable the efficient production of site-specifically modified therapeutic proteins, and the quantitative replacement of targeted cellular proteins with versions bearing unnatural amino acids that allow imaging or synthetic regulation of protein function.