enAsCas12a Enables CRISPR-Directed Evolution to Screen for Functional Drug Resistance Mutations in Sequences Inaccessible to SpCas9

Activating UCHL1 through the CRISPR activation system promotes cartilage differentiation mediated by HIF-1α/SOX9

Developing strategies to enhance cartilage differentiation in mesenchymal stem cells and preserve the extracellular matrix is crucial for successful cartilage tissue reconstruction. Hypoxia-inducible factor-1α (HIF-1α) plays a pivotal role in maintaining the extracellular matrix and chondrocyte phenotype, thus serving as a key regulator in chondral tissue engineering strategies. Recent studies have shown that Ubiquitin C-terminal hydrolase L1 (UCHL1) is involved in the deubiquitylation of HIF-1α. However, the regulatory role of UCHL1 in chondrogenic differentiation has not been investigated. In the present study, we initially validated the promotive effect of UCHL1 expression on chondrogenesis in adipose-derived stem cells (ADSCs). Subsequently, a hybrid baculovirus system was designed and employed to utilize three CRISPR activation (CRISPRa) systems, employing dead Cas9 (dCas9) from three distinct bacterial sources to target UCHL1. Then UCHL1 and HIF-1α inhibitor and siRNA targeting SRY-box transcription factor 9 (SOX9) were used to block UCHL1, HIF-1α and SOX9, respectively. Cartilage differentiation and chondrogenesis were measured by qRT-PCR, immunofluorescence and histological staining. We observed that the CRISPRa system derived from Staphylococcus aureus exhibited superior efficiency in activating UCHL1 compared to the commonly used the CRISPRa system derived from Streptococcus pyogenes. Furthermore, the duration of activation was extended by utilizing the Cre/loxP-based hybrid baculovirus. Moreover, our findings show that UCHL1 enhances SOX9 expression by regulating the stability and localization of HIF-1α, which promotes cartilage production in ADSCs. These findings suggest that activating UCHL1 using the CRISPRa system holds significant potential for applications in cartilage regeneration.

Enhanced single-base mutation diversity by the combination of cytidine deaminase with DNA-repairing enzymes in yeast

The occurrence of random mutations can increase the diversity of the genome and promote the evolutionary process of organisms. High efficiency mutagenesis techniques significantly accelerate the evolutionary process. In this work, we describe a targeted mutagenesis system named MutaT7trans to significantly increase mutation rate and generate mutations across all four nucleotides in yeast. We constructed different DNA-repairing enzyme-PmCDA1-T7 RNA polymerase (T7 RNAP) fusion proteins, achieved targeted mutagenesis by flanking the target gene with T7 promoters, and tuned the mutation spectra by introducing different DNA-repairing enzymes. With this mutagenesis tool, the proportion of non-C → T mutations was 10-11-fold higher than the cytidine deaminase-based evolutionary tools, and the transversion mutation frequency was also elevated. The mutation rate of the target gene was significantly increased to 5.25 × 10-3 substitutions per base (s. p. b.). We also demonstrated that MutaT7trans could be used to evolve the CrtE, CrtI, and CrtYB gene in the β-carotene biosynthesis process and generate different types of mutations.

Base editor screens for in situ mutational scanning at scale

A fundamental challenge in biology is understanding the molecular details of protein function. How mutations alter protein activity, regulation, and response to drugs is of critical importance to human health. Recent years have seen the emergence of pooled base editor screens for in situ mutational scanning: the interrogation of protein sequence-function relationships by directly perturbing endogenous proteins in live cells. These studies have revealed the effects of disease-associated mutations, discovered novel drug resistance mechanisms, and generated biochemical insights into protein function. Here, we discuss how this "base editor scanning" approach has been applied to diverse biological questions, compare it with alternative techniques, and describe the emerging challenges that must be addressed to maximize its utility. Given its broad applicability toward profiling mutations across the proteome, base editor scanning promises to revolutionize the investigation of proteins in their native contexts.

Integration of CRISPR/Cas9 with artificial intelligence for improved cancer therapeutics

Gene editing has great potential in treating diseases caused by well-characterized molecular alterations. The introduction of clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9 (Cas9)–based gene-editing tools has substantially improved the precision and efficiency of gene editing. The CRISPR/Cas9 system offers several advantages over the existing gene-editing approaches, such as its ability to target practically any genomic sequence, enabling the rapid development and deployment of novel CRISPR-mediated knock-out/knock-in methods. CRISPR/Cas9 has been widely used to develop cancer models, validate essential genes as druggable targets, study drug-resistance mechanisms, explore gene non-coding areas, and develop biomarkers. CRISPR gene editing can create more-effective chimeric antigen receptor (CAR)-T cells that are durable, cost-effective, and more readily available. However, further research is needed to define the CRISPR/Cas9 system’s pros and cons, establish best practices, and determine social and ethical implications. This review summarizes recent CRISPR/Cas9 developments, particularly in cancer research and immunotherapy, and the potential of CRISPR/Cas9-based screening in developing cancer precision medicine and engineering models for targeted cancer therapy, highlighting the existing challenges and future directions. Lastly, we highlight the role of artificial intelligence in refining the CRISPR system's on-target and off-target effects, a critical factor for the broader application in cancer therapeutics.

An ultrasensitive, rapid and portable method for screening oseltamivir-resistant virus based on CRISPR/Cas12a combined with immunochromatographic strips

Influenza is a significant public health challenge because of the emergence of antigenically shifted or highly virulent strains. The neuraminidase inhibitor oseltamivir is used as an antiviral drug in clinical treatment. However, its therapeutic effects can be greatly compromised by the emergence of drug-resistant mutant viruses. Thus, there is an urgent need to distinguish drug-resistant strains with a simple method. To address this, in the present study, we develop a rapid, sensitive and convenient molecular diagnosis method based on CRISPR/Cas12a technology and lateral flow detection (LFD). By targeting mutant sequences amplified by recombinase polymerase amplification (RPA) reaction, crRNA is designed to develop the CRISPR/Cas12a assay, and 2000 copies can be directly observed by the naked eye under blue light-emitting diode (LED) light. Combined with LFD, the limit of detection of RPA-CRISPR/Cas12a-LFD is about 20 copies of target sequence per reaction. Collectively, RPA-CRISPR/Cas12a-LFD method provides a novel alternative for the sensitive, specific and portable detection to diagnose oseltamivir-resistant mutant strains.

Target Identification of Small Molecules Using Large-Scale CRISPR-Cas Mutagenesis Scanning of Essential Genes

Target deconvolution of new bioactive agents identified from phenotypic screens remains a challenging task. The discovery of mutations that confer resistance to such agents is regarded as the gold standard proof of target identification. Here, we describe a method that exploits the error-prone repair of CRISPR-induced DNA double-strand breaks to enhance mutagenesis and increase the incidence of drug resistance mutations in essential genes. As each DNA double-strand break is introduced at a targeted genomic site predefined by the presence of a protospacer adjacent motif (PAM) and a particular CRISPR single guide RNA (sgRNA), the genetic location of drug resistance mutations can be easily uncovered through targeted sequencing of CRISPR sgRNAs. Moreover, the method allows for the identification of not only the drug target gene, but also the drug-binding domain within the target gene.

CRISPR-based therapeutics: current challenges and future applications

The discovery, only a decade ago, of the genome editing power of clustered regularly interspaced short palindromic repeats (CRISPR)-associated nucleases is already reinventing the therapeutic process, from how new drugs are discovered to novel ways to treat diseases. CRISPR-based screens can aid therapeutic development by quickly identifying a drug's mechanism of action and escape mutants. Additionally, CRISPR-Cas has advanced emerging ex vivo therapeutics, such as cell replacement therapies. However, Cas9 is limited as an in vivo therapeutic due to ineffective delivery, unwanted immune responses, off-target effects, unpredictable repair outcomes, and cellular stress. To address these limitations, controls that inhibit or degrade Cas9, biomolecule-Cas9 conjugates, and base editors have been developed. Herein, we discuss CRISPR-Cas systems that advance both conventional and emerging therapeutics.

Genome editor-directed in vivo library diversification

The generation of a library of variant genes is a prerequisite of directed evolution, a powerful tool for biomolecular engineering. As the number of all possible sequences often far exceeds the diversity of a practical library, methods that allow efficient library diversification in living cells are essential for in vivo directed evolution technologies to effectively sample the sequence space and allow hits to emerge. While traditional whole-genome mutagenesis often results in toxicity and the emergence of "cheater" mutations, recent developments that exploit the targeting and editing abilities of genome editors to facilitate in vivo library diversification have allowed for precise mutagenesis focused on specific genes of interest, higher mutational density, and reduced the occurrence of cheater mutations. This minireview summarizes recent advances in genome editor-directed in vivo library diversification and provides an outlook on their future applications in chemical biology.

CRISPR/Cas9-based directed evolution in mammalian cells

An increasingly powerful set of new CRISPR/Cas-based methods is becoming available for directed evolution of proteins in mammalian cells. Although in vitro techniques or microbial expression systems have been dominating directed evolution, there are now promising approaches to diversify proteins in mammalian cells in situ. This can be achieved by simple indel mutagenesis or more sophisticated homology repair mechanisms for cassette mutagenesis of coding sequences. Cas9 variant fusions to base editors and other effectors pose another promising way to introduce diversity into proteins. CRISPR/Cas9-based directed evolution in mammalian cells opens a new exciting era of discovery for the many classes of proteins for which a mammalian cellular context is preferable.