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换一换High-throughput single-cell transcriptomics of bacteria using combinatorial barcoding
Karl D. Gaisser, Sophie N. Skloss, Leandra M. Brettner, Luana Paleologu, Charles M. Roco, Alexander B. Rosenberg, Matthew Hirano, R. William DePaolo, Georg Seelig, Anna Kuchina
Microbial split-pool ligation transcriptomics (microSPLiT) is a high-throughput single-cell RNA sequencing method for bacteria. With four combinatorial barcoding rounds, microSPLiT can profile transcriptional states in hundreds of thousands of Gram-negative and Gram-positive bacteria in a single experiment without specialized equipment. As bacterial samples are fixed and permeabilized before barcoding, they can be collected and stored ahead of time. During the first barcoding round, the fixed and permeabilized bacteria are distributed into a 96-well plate, where their transcripts are reverse transcribed into cDNA and labeled with the first well-specific barcode inside the cells. The cells are mixed and redistributed two more times into new 96-well plates, where the second and third barcodes are appended to the cDNA via in-cell ligation reactions. Finally, the cells are mixed and divided into aliquot sub-libraries, which can be stored until future use or prepared for sequencing with the addition of a fourth barcode. It takes 4 days to generate sequencing-ready libraries, including 1 day for collection and overnight fixation of samples. The standard plate setup enables single-cell transcriptional profiling of up to 1 million bacterial cells and up to 96 samples in a single barcoding experiment, with the possibility of expansion by adding barcoding rounds. The protocol requires experience in basic molecular biology techniques, handling of bacterial samples and preparation of DNA libraries for next-generation sequencing. It can be performed by experienced undergraduate or graduate students. Data analysis requires access to computing resources, familiarity with Unix command line and basic experience with Python or R.
AI 解读Capturing acyl–enzyme intermediates with genetically encoded 2,3-diaminopropionic acid for hydrolase substrate identification
Juan Luo, Yao Yu, Ke Wang, Sizhe He, Longjie Wang, Fangfang Liang, Jason W. Chin, Shan Tang
Catalytic mechanism-based, light-activated traps have recently been developed to identify the substrates of cysteine or serine hydrolases. These traps are hydrolase mutants whose catalytic cysteine or serine are replaced with genetically encoded 2,3-diaminopropionic acid (DAP). DAP-containing hydrolases specifically capture the transient thioester- or ester-linked acyl–enzyme intermediates resulting from the first step of the proteolytic reaction as their stable amide analogs. The trapped substrate fragments allow the downstream identification of hydrolase substrates by mass spectrometry and immunoblotting. In this protocol, we provide a detailed step-by-step guide for substrate capture and identification of the peptidase domain of the large tegument protein deneddylase (UL36USP) from human herpesvirus 1, both in mammalian cell lysate and live mammalian cells. Four procedures are included: Procedure 1, DAP-mediated substrate trapping in mammalian cell lysate (~8 d); Procedure 2, DAP-mediated substrate trapping in adherent mammalian cells (~6 d); Procedure 3, DAP-mediated substrate trapping in suspension mammalian cells (~5 d); and Procedure 4, substrate identification and validation (~12–13 d). Basic skills to perform protein expression in bacteria or mammalian cells, affinity enrichment and proteomic analysis are required to implement the protocol. This protocol will be a practical guide for identifying substrates of serine or cysteine hydrolases either in a complex mixture, where genetic manipulation is challenging, or in live cells such as bacteria, yeasts and mammalian cells.
AI 解读Integrating genome-wide CRISPR screens and in silico drug profiling for targeted antidote development
Bei Wang, Yu Xu, Arabella H. Wan, Guohui Wan, Qiao-Ping Wang
Numerous toxins threaten humans, but specific antidotes are unavailable for most of them. Although CRISPR screening has aided the discovery of the mechanisms of some toxins, developing targeted antidotes remains a significant challenge. Recently, we established a systematic framework to develop antidotes by combining the identification of novel drug targets by using a genome-wide CRISPR screen with a virtual screen of drugs approved by the US Food and Drug Administration. This approach allows for a comprehensive understanding of toxin mechanisms at the whole-genome level and facilitates the identification of promising antidote drugs targeting specific molecules. Here, we present step-by-step instructions for executing genome-scale CRISPR–Cas9 knockout screens of toxins in HAP1 cells. We also provide detailed guidance for conducting an in silico drug screen and an in vivo drug validation. By using this protocol, it takes ~4 weeks to perform the genome-scale screen, 4 weeks for sequencing and data analysis, 4 weeks to validate candidate genes, 1 week for the virtual screen and 2 weeks for in vitro drug validation. This framework has the potential to accelerate the development of antidotes for a wide range of toxins and can rapidly identify promising drug candidates that are already known to be safe and effective. This could lead to the development of new antidotes much more quickly than traditional methods, protecting lives from diverse toxins and advancing human health.
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