RNA-guided endonucleases (RGENs), derived from the prokaryotic adaptive immune system known as CRISPR/Cas, enable targeted genome engineering in cells and organisms. RGENs are ribonucleoproteins that consist of guide RNA and Cas9, a protein component originated from Streptococcus pyogenes. These enzymes cleave chromosomal DNA, whose sequence is complementary, to guide RNA in a targeted manner, producing site-specific DNA double-strand breaks (DSBs), the repair of which gives rise to targeted genome modifications. Despite broad interest in RGEN-mediated genome editing, these nucleases are limited by off-target mutations and unwanted chromosomal translocations associated with off-target DNA cleavages. Here, we show that off-target effects of RGENs can be reduced below the detection limits of deep sequencing by choosing unique target sequences in the genome and modifying both guide RNA and Cas9. We found that both the composition and structure of guide RNA can affect RGEN activities in cells to reduce off-target effects. RGENs efficiently discriminated on-target sites from off-target sites that differ by two bases. Furthermore, exome sequencing analysis showed that no off-target mutations were induced by two RGENs in four clonal populations of mutant cells. In addition, paired Cas9 nickases, composed of D10A Cas9 and guide RNA, which generate two single-strand breaks (SSBs) or nicks on different DNA strands, were highly specific in human cells, avoiding off-target mutations without sacrificing genome-editing efficiency. Interestingly, paired nickases induced chromosomal deletions in a targeted manner without causing unwanted translocations. Our results highlight the importance of choosing unique target sequences and optimizing guide RNA and Cas9 to avoid or reduce RGEN-induced off-target mutations.

Transcription activator-like (TAL) effector nucleases (TALENs) can be readily engineered to bind specific genomic loci, enabling the introduction of precise genetic modifications such as gene knockouts and additions. Here we present a genome-scale collection of TALENs for efficient and scalable gene targeting in human cells. We chose target sites that did not have highly similar sequences elsewhere in the genome to avoid off-target mutations and assembled TALEN plasmids for 18,740 protein-coding genes using a high-throughput Golden-Gate cloning system. A pilot test involving 124 genes showed that all TALENs were active and disrupted their target genes at high frequencies, although two of these TALENs became active only after their target sites were partially demethylated using an inhibitor of DNA methyltransferase. We used our TALEN library to generate single- and double-gene-knockout cells in which NF-κB signaling pathways were disrupted. Compared with cells treated with short interfering RNAs, these cells showed unambiguous suppression of signal transduction.

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Senescence, defined as irreversible cell-cycle arrest, is the main driving force of aging and age-related diseases. Here, we performed high-throughput screening to identify compounds that alleviate senescence and identified the ataxia telangiectasia mutated (ATM) inhibitor KU-60019 as an effective agent. To elucidate the mechanism underlying ATM's role in senescence, we performed a yeast two-hybrid screen and found that ATM interacted with the vacuolar ATPase V subunits ATP6V1E1 and ATP6V1G1. Specifically, ATM decreased E-G dimerization through direct phosphorylation of ATP6V1G1. Attenuation of ATM activity restored the dimerization, thus consequently facilitating assembly of the V and V domains with concomitant reacidification of the lysosome. In turn, this reacidification induced the functional recovery of the lysosome/autophagy system and was coupled with mitochondrial functional recovery and metabolic reprogramming. Together, our data reveal a new mechanism through which senescence is controlled by the lysosomal-mitochondrial axis, whose function is modulated by the fine-tuning of ATM activity.

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Rapid and highly sensitive detection of biomolecules is greatly needed for pathogen diagnosis in clinical samples, but the method needs to be significantly improved in terms of sensitivity and specificity for actual use in clinical settings. Here, we report the development of an improved molecular diagnostics tool that utilizes CRISPR/dCas9mediated biosensor that couples a nuclease inactivated Cas9 (dCas9) and single microring resonator biosensor, enables label-free and real-time detection of pathogenic DNA and RNA. We addressed the clinical utility of this CRISPR/dCas9-mediated biosensor in tick-borne illnesses including scrub typhus (ST) and severe fever with thrombocytopenia syndrome (SFTS), whose clinical presentations are too similar to be easily differentiated. By using CRISPR/dCas9-mediated biosensor, we achieved single molecule sensitivity for the detection of ST (0.54 aM) and SFTS (0.63 aM); this detection sensitivity is 100 times more sensitive than that of RT-PCR assay. Finally, CRISPR/dCas9-mediated biosensor was able to clearly distinguish between ST and SFTS in serum samples within 20 min. We believe that CRISPR/dCas9-mediated biosensor will be useful for rapid and accurate molecular diagnostic tool that is suitable for immediate clinical applications.

Genetic mutations in BRCA1, which is crucial for the process of DNA repair and maintenance of genomic integrity, are known to increase markedly the risk of breast and ovarian cancers. Clinical genetic testing has been used to identify new BRCA1 variants; however, functional assessment and determination of their pathogenicity still poses challenges for clinical management. Here, we describe that CRISPR-mediated cytosine base editor, known as BE3, can be used for the functional analysis of BRCA1 variants. We performed CRISPR-mediated base-editing screening using 745 gRNAs targeting all exons in BRCA1 to identify loss-of-function variants and identified variants whose function has heretofore remained unknown, such as c.-97C>T, c.154C>T, c.3847C>T, c.5056C>T, and c.4986+5G>A. Our results show that CRISPR-mediated base editor is a powerful tool for the reclassification of variants of uncertain significance (VUSs) in BRCA1.

Although prime editors are a powerful tool for genome editing, which can generate various types of mutations such as nucleotide substitutions, insertions, and deletions in the genome without double-strand breaks or donor DNA, the conventional prime editors are still limited to their target scopes because of the PAM preference of the Streptococcus pyogenes Cas9 (spCas9) protein. Here, we describe the engineered prime editors to expand the range of their target sites using various PAM-flexible Cas9 variants. Using the engineered prime editors, we could successfully generate more than 50 types of mutations with up to 51.7% prime-editing activity in HEK293T cells. In addition, we successfully introduced the BRAF V600E mutation, which could not be induced by conventional prime editors. These variants of prime editors will broaden the applicability of CRISPR-based prime editing technologies in biological research.