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CRISPR-Cas nucleases are powerful tools for manipulating nucleic acids; however, targeted insertion of DNA remains a challenge, as it requires host cell repair machinery. Here we characterize a CRISPR-associated transposase from cyanobacteria Scytonema hofmanni (ShCAST) that consists of Tn7-like transposase subunits and the type V-K CRISPR effector (Cas12k). ShCAST catalyzes RNA-guided DNA transposition by unidirectionally inserting segments of DNA 60 to 66 base pairs downstream of the protospacer. ShCAST integrates DNA into targeted sites in the Escherichia coli genome with frequencies of up to 80% without positive selection. This work expands our understanding of the functional diversity of CRISPR-Cas systems and establishes a paradigm for precision DNA insertion.

Many chromatin features play critical roles in regulating gene expression. A complete understanding of gene regulation will require the mapping of specific chromatin features in small samples of cells at high resolution. Here we describe Cleavage Under Targets and Tagmentation (CUT&Tag), an enzyme-tethering strategy that provides efficient high-resolution sequencing libraries for profiling diverse chromatin components. In CUT&Tag, a chromatin protein is bound in situ by a specific antibody, which then tethers a protein A-Tn5 transposase fusion protein. Activation of the transposase efficiently generates fragment libraries with high resolution and exceptionally low background. All steps from live cells to sequencing-ready libraries can be performed in a single tube on the benchtop or a microwell in a high-throughput pipeline, and the entire procedure can be performed in one day. We demonstrate the utility of CUT&Tag by profiling histone modifications, RNA Polymerase II and transcription factors on low cell numbers and single cells. Understanding gene regulation will require mapping specific chromain features in a small number of cells at high resolution. Here the authors describe CUT&Tag, which uses antibody-mediated tethering of Tn5 transposase to a chromatin protein to generate high resolution libraries.

The assay for transposase-accessible chromatin using sequencing (ATAC-seq) provides a simple and scalable way to detect the unique chromatin landscape associated with a cell type and how it may be altered by perturbation or disease. ATAC-seq requires a relatively small number of input cells and does not require a priori knowledge of the epigenetic marks or transcription factors governing the dynamics of the system. Here we describe an updated and optimized protocol for ATAC-seq, called Omni-ATAC, that is applicable across a broad range of cell and tissue types. The ATAC-seq workflow has five main steps: sample preparation, transposition, library preparation, sequencing and data analysis. This protocol details the steps to generate and sequence ATAC-seq libraries, with recommendations for sample preparation and downstream bioinformatic analysis. ATAC-seq libraries for roughly 12 samples can be generated in 10 h by someone familiar with basic molecular biology, and downstream sequencing analysis can be implemented using benchmarked pipelines by someone with basic bioinformatics skills and with access to a high-performance computing environment.A protocol for generating chromatin accessibility profiles from a broad variety of cell and tissue types, including a step-by-step workflow for library preparation and guidelines for data processing and downstream analysis.

We recently introduced Cleavage Under Targets & Tagmentation (CUT&Tag), an epigenomic profiling strategy in which antibodies are bound to chromatin proteins in situ in permeabilized nuclei. These antibodies are then used to tether the cut-and-paste transposase Tn5. Activation of the transposase simultaneously cleaves DNA and adds adapters (‘tagmentation’) for paired-end DNA sequencing. Here, we introduce a streamlined CUT&Tag protocol that suppresses DNA accessibility artefacts to ensure high-fidelity mapping of the antibody-targeted protein and improves the signal-to-noise ratio over current chromatin profiling methods. Streamlined CUT&Tag can be performed in a single PCR tube, from cells to amplified libraries, providing low-cost genome-wide chromatin maps. By simplifying library preparation CUT&Tag requires less than a day at the bench, from live cells to sequencing-ready barcoded libraries. As a result of low background levels, barcoded and pooled CUT&Tag libraries can be sequenced for as little as $25 per sample. This enables routine genome-wide profiling of chromatin proteins and modifications and requires no special skills or equipment. The authors describe a streamlined epigenomic profiling protocol based on cut-and-paste tagmentation by the Tn5 transposase targeted to a chromatin protein of interest.

The current technologies to place new DNA into specific locations in plant genomes are low frequency and error-prone, and this inefficiency hampers genome-editing approaches to develop improved crops 1 , 2 . Often considered to be genome ‘parasites’, transposable elements (TEs) evolved to insert their DNA seamlessly into genomes 3 – 5 . Eukaryotic TEs select their site of insertion based on preferences for chromatin contexts, which differ for each TE type 6 – 9 . Here we developed a genome engineering tool that controls the TE insertion site and cargo delivered, taking advantage of the natural ability of the TE to precisely excise and insert into the genome. Inspired by CRISPR-associated transposases that target transposition in a programmable manner in bacteria 10 – 12 , we fused the rice Pong transposase protein to the Cas9 or Cas12a programmable nucleases. We demonstrated sequence-specific targeted insertion (guided by the CRISPR gRNA) of enhancer elements, an open reading frame and a gene expression cassette into the genome of the model plant Arabidopsis . We then translated this system into soybean—a major global crop in need of targeted insertion technology. We have engineered a TE ‘parasite’ into a usable and accessible toolkit that enables the sequence-specific targeting of custom DNA into plant genomes. Fusion of rice Pong transposase to the Cas9 or Cas12a programmable nucleases provides sequence-specific targeted insertion of enhancer elements, an open reading frame and gene expression cassette into the genome of the model plant Arabidopsis and crop soybean .

The development of new CRISPR–Cas genome editing tools continues to drive major advances in the life sciences. Four classes of CRISPR–Cas-derived genome editing agents—nucleases, base editors, transposases/recombinases and prime editors—are currently available for modifying genomes in experimental systems. Some of these agents have also moved rapidly into the clinic. Each tool comes with its own capabilities and limitations, and major efforts have broadened their editing capabilities, expanded their targeting scope and improved editing specificity. We analyze key considerations when choosing genome editing agents and identify opportunities for future improvements and applications in basic research and therapeutics.A growing arsenal of CRISPR-based tools enables increasingly sophisticated genome editing applications.

Up to half of children with severe developmental disorders of probable genetic origin remain without a genetic diagnosis; here, in a systematic and nationwide study of 1,133 children with severe, undiagnosed developmental disorders, and their parents, exome sequencing and array-based detection of chromosomal rearrangements reveals novel genes causing developmental disorders, increasing the proportion of children that can now be diagnosed to 31%. Gene linkage to developmental disorders Until recently, the discovery of the genetic causes of monogenic disorders has been predominantly phenotype-driven. Up to half of all children with severe developmental disorders of probable genetic origin remain without a genetic diagnosis. This publication from The Deciphering Developmental Disorders Study presents a UK-wide systematic genetic analysis of 1,133 children with severe, undiagnosed developmental disorders, and their parents. Exome sequencing and array-based detection of chromosomal rearrangements revealed 12 previously unknown developmental disorder genes and increased the proportion of children that could be diagnosed by 10%. Despite three decades of successful, predominantly phenotype-driven discovery of the genetic causes of monogenic disorders 1 , up to half of children with severe developmental disorders of probable genetic origin remain without a genetic diagnosis. Particularly challenging are those disorders rare enough to have eluded recognition as a discrete clinical entity, those with highly variable clinical manifestations, and those that are difficult to distinguish from other, very similar, disorders. Here we demonstrate the power of using an unbiased genotype-driven approach 2 to identify subsets of patients with similar disorders. By studying 1,133 children with severe, undiagnosed developmental disorders, and their parents, using a combination of exome sequencing 3 , 4 , 5 , 6 , 7 , 8 , 9 , 10 , 11 and array-based detection of chromosomal rearrangements, we discovered 12 novel genes associated with developmental disorders. These newly implicated genes increase by 10% (from 28% to 31%) the proportion of children that could be diagnosed. Clustering of missense mutations in six of these newly implicated genes suggests that normal development is being perturbed by an activating or dominant-negative mechanism. Our findings demonstrate the value of adopting a comprehensive strategy, both genome-wide and nationwide, to elucidate the underlying causes of rare genetic disorders.

Targeting membrane proteins could improve the efficacy of T cell–based immunotherapies. To facilitate the identification of T cell targets, we developed a hybrid genetic screening system where the Sleeping Beauty (SB) transposon and single guide RNA cassette are nested in an adeno-associated virus (AAV). SB-mediated genomic integration of the single guide RNA cassette enables efficient gene editing in primary murine T cells as well as a screen readout. We performed in vivo AAV–SB-CRISPR screens for membrane protein targets in CD8+ T cells in mouse models of glioblastoma (GBM). We validated screen hits by demonstrating that adoptive transfer of CD8+ T cells with Pdia3, Mgat5, Emp1 or Lag3 gene editing enhances the survival of GBM-bearing mice in both syngeneic and T-cell receptor transgenic models. Transcriptome profiling, single cell sequencing, cytokine assays and T cell signaling analysis showed that Pdia3 editing in T cells enhances effector functions. Engineered PDIA3 mutant EGFRvIII chimeric antigen T cells are more potent in antigen-specific killing of human GBM cells.

Joint profiling of chromatin accessibility and gene expression from the same single cell provides critical information about cell types in a tissue and cell states during a dynamic process. Here, we develop in situ sequencing hetero RNA–DNA-hybrid after assay for transposase-accessible chromatin-sequencing (ISSAAC-seq), a highly sensitive and flexible single-cell multi-omics method to interrogate chromatin accessibility and gene expression from the same single nucleus. We demonstrated that ISSAAC-seq is sensitive and provides high quality data with orders of magnitude more features than existing methods. Using the joint profiles from over 10,000 nuclei from the mouse cerebral cortex, we uncovered major and rare cell types and cell-type specific regulatory elements and identified heterogeneity at the chromatin level within established cell types defined by gene expression. Finally, we revealed distinct dynamics and relationships of gene expression and chromatin accessibility during an oligodendrocyte maturation trajectory. ISSAAC-seq offers a sensitive and modular tool for joint profiling gene expression and chromatin accessibility in single cells.

CRISPR-associated transposases (CASTs) enable recombination-independent, multi-kilobase DNA insertions at RNA-programmed genomic locations. However, the utility of type V-K CASTs is hindered by high off-target integration and a transposition mechanism that results in a mixture of desired simple cargo insertions and undesired plasmid cointegrate products. Here we overcome both limitations by engineering new CASTs with improved integration product purity and genome-wide specificity. To do so, we engineered a nicking homing endonuclease fusion to TnsB (named HELIX) to restore the 5′ nicking capability needed for cargo excision on the DNA donor. HELIX enables cut-and-paste DNA insertion with up to 99.4% simple insertion product purity, while retaining robust integration efficiencies on genomic targets. HELIX has substantially higher on-target specificity than canonical CASTs, and we identify several novel factors that further regulate targeted and genome-wide integration. Finally, we extend HELIX to other type V-K orthologs and demonstrate the feasibility of HELIX-mediated integration in human cell contexts. Engineered CRISPR-associated transposases improve the purity and specificity of DNA insertions.