Cas9 and Cas12 are RNA-guided DNA endonucleases derived from prokaryotic CRISPR-Cas adaptive immune systems that have been repurposed as versatile genome-engineering tools. Computational mining of genomes and metagenomes has expanded the diversity of Cas9 and Cas12 enzymes that can be used to develop versatile, orthogonal molecular toolboxes. Structural information is pivotal to uncovering the precise molecular mechanisms of newly discovered Cas enzymes and providing a foundation for their application in genome editing. In this chapter, we describe detailed protocols for the preparation of Cas9 and Cas12 enzymes for cryo-electron microscopy. These methods will enable fast and robust structural determination of newly discovered Cas9 and Cas12 enzymes, which will enhance the understanding of diverse CRISPR-Cas effectors and provide a molecular framework for expanding CRISPR-based genome-editing technologies.

The threat of emerging infectious diseases (e.g., SARS-CoV-2 the RNA virus responsible for the COVID-19 pandemic) has highlighted the importance of accurate and rapid testing for screening, patient diagnosis, and effective treatment of infectious disease. Nucleic acid diagnostic tools such as qPCR are considered the gold standard, providing a sensitive, accurate, and robust method of detection. However, these conventional diagnostic platforms are resource intensive, limited in some applications, and are almost always confined to laboratory settings. With the increasing demand for low-cost, rapid, and accurate point-of-care diagnostics, CRISPR-based systems have emerged as powerful tools to augment detection capabilities. Of note is the potent RNA detection enzyme, Leptotrichia buccalis (Lbu) Cas13a, which is capable of rapid RNA detection in complex mixtures with or without pre-amplification. To support its wide-spread use, we describe a detailed method for the expression, purification, and validation of LbuCas13a for use in molecular diagnostics.

Since their introduction, CRISPR-based DNA base editors (BEs) have become essential in the field of precision genome editing, revolutionizing the correction of pathogenic SNPs for both basic research and therapeutic applications. As this technology advances, more laboratories are implementing these tools into their workflow. The delivery of BEs as BE-guide RNA complexes (RNPs), rather than as mRNA or plasmids, has been shown to exhibit lower off-target effects, establishing it as the preferred method of delivery. However, there are no protocols describing in detail how to obtain high-purity and highly active BE RNPs. Here, we offer a comprehensive guide for the expression, purification, RNP reconstitution, and in vitro activity assessment of TadA-based BEs. The protocol includes guidance on performing activity assays using commercial denaturing gels, which is convenient and uses standard molecular biology equipment. This allows for rapid quality control testing of reconstituted BE RNPs prior to more expensive and time-consuming in vivo genome editing experiments. Overall, this protocol aims to empower more laboratories to generate tailored BE RNPs for diverse in vitro and in vivo applications.

Pathogenic microorganisms, such as viruses, have threatened human health and will continue to contribute to future epidemics and pandemics, highlighting the importance of developing effective diagnostics. To contain viral outbreaks within populations, fast and early diagnosis of infected individuals is essential. Although current standard methods are highly sensitive and specific, like RT-qPCR, some can have slow turnaround times, which can hinder the prevention of viral transmission. The discovery of CRISPR-Cas systems in bacteria and archaea initially revolutionized the world of genome editing. Intriguingly, CRISPR-Cas enzymes also have the ability to detect nucleic acids with high sensitivity and specificity, which sparked the interest of researchers to also explore their potential in diagnosis of viral pathogens. In particular, the CRISPR-Cas13 system has been used as a tool for detecting viral nucleic acids. Cas13's capability to detect both target RNA and non-specific RNAs has led to the development of detection methods that leverage these characteristics through designing specific detection read-outs. Optimization of viral sample collection, amplification steps and the detection process within the Cas13 detection workflow has resulted in assays with high sensitivity, rapid turnaround times and the capacity for large-scale implementation. This review focuses on the significant innovations of various CRISPR-Cas13-based viral nucleic acid detection methods, comparing their strengths and weaknesses while highlighting Cas13's great potential as a tool for viral diagnostics.

A mechanistic understanding of the different pathways involved in the repair of DSBs is a timely, yet challenging task. CRISPR-Cas9 is a powerful tool to induce DNA double-strand breaks (DSB) at defined genomic locations to study the ensuing repair response, but Cas9 studies are typically limited by i) low-throughput induction of DSB, by targeting only one or a few genomic sites, or ii) the use of genetically integrated reporter systems, which do not always reflect endogenous phenotypes. To address these limitations, we developed multi-target CRISPR, a Cas9-based tool to controllably induce DSBs in high-throughput at endogenous sites, by leveraging repetitive genomic regions. In this Chapter, we describe how to design and execute a multi-target CRISPR experiment. We also detail how to analyze next-generation sequencing data for characterization of DSB repair events at multiple cut sites. We envision that multi-target CRISPR will become a valuable tool for the study of mammalian DSB repair mechanisms.

CRISPR-associated transposons (CASTs) are potential gene editing tools because of their RNA-guided DNA insertion activity. It is essential to understand the mechanisms underlying the transposition for the application of CASTs. Here, we provide protocols for the biochemical reconstitution of a type I-B CAST for RNA-guided transposition. The procedures may be applicable to other types of CASTs and facilitate the mechanism studies of various CASTs.

CRISPR-Cas9 has transformed genome editing through its programmability and versatility. Its DNA cleavage activity involves dynamic conformational changes during gRNA binding, DNA recognition, R-loop formation, and endonuclease activation. Understanding these molecular transitions is critical for improving the specificity and efficiency of Cas9, but this remains challenging precisely due to these rapid structural rearrangements. Early structural studies provided foundational insights but were limited to static states under catalytically inactive conditions. Cryo-EM has since enabled visualization of the dynamic nature of active Cas9, by enriching for specific conformations. This chapter introduces a kinetics-informed cryo-EM approach to capture the stepwise activation of Cas9 in real time. With thorough kinetic analyses, such as stopped-flow measurements of R-loop formation, we describe how to identify optimal timepoints to visualize key conformational states with cryo-EM. Integration of kinetic and structural data enables precise mapping of the conformational landscape of Cas9 and other dynamic enzymes, advancing our understanding of their molecular mechanisms and providing a framework for engineering enhanced variants.

The CRISPR-associated (Cas) nuclease Cas12a2 from Sulfuricurvum sp. PC08-66 (SuCas12a2) binds RNA targets with a complementary guide (g)RNA. Target RNA binding causes a major conformational rearrangement in Cas12a2 that activates a RuvC nuclease domain to collaterally cleave RNA, ssDNA and dsDNA, arresting growth and providing population-level immunity. Here, we report in vivo, cell-free, and in vitro methods to characterize the collateral cleavage activity of SuCas12a2 as well as a procedure for gRNA design. As part of the in vivo methods, we describe how to capture growth arrest through plasmid interference and induction of an SOS DNA damage response in the bacterium Escherichia coli. We further apply cell-free transcription-translation to affirm collateral cleavage activity triggered by an expressed RNA target. Finally, as part of the in vitro methods, we describe how to purify active nuclease and subsequently conduct biochemical cleavage assays. In total, the outlined methods should accelerate the exploration of SuCas12a2 and other related Cas nucleases, revealing new features of CRISPR biology and helping develop new CRISPR technologies for molecular diagnostics and other applications.

Prime editing enables the generation of nearly any small genetic variant. However, the process of prime editing guide RNA (pegRNA) design is challenging and requires automated computational design tools. We developed Prime Editing Guide Generator (PEGG), a fast, flexible, and user-friendly Python package that enables the rapid generation of pegRNA and pegRNA-sensor libraries. Here, we describe the installation and use of PEGG (https://pegg.readthedocs.io) to rapidly generate custom pegRNA-sensor libraries for use in high-throughput prime editing screens.

According to the Clinvar database, modeling the diseases associated with pathogenic mutations requires the installation of base substitutions, small insertions or deletions. Prime editor (PE) was recently developed to precisely install any base substitutions and/or small insertions/deletions (indels) in mammalian cells and animals without requiring DSBs or donor DNA templates. PE also offers greater editing and targeting flexibility compared to other precision CRISPR editing methods because the versatile editing information is encoded in the reverse-transcription template of its prime editing guide RNA. However, optimal PE system selection and experimental design can be complex, and there are various factors that can affect PE efficiency. This chapter serves as a rapid entry-level guideline for the application of PE, providing an experimental framework for using PE at a specific genomic locus. RUNX1 was selected as a representative target site to illustrate the detailed methodology for constructing PE plasmids and the process of transfecting these plasmids into 293FT cells. We further examined the efficiency of PE-mediated genome editing in mammalian cells by using next-generation sequencing.

Molecular recording technologies record and store information about cellular history. Lineage tracing is one form of molecular recording and produces information describing cellular trajectories during mammalian development, differentiation and maintenance of adult stem cell niches, and tumor evolution. Our molecular recorder technology utilizes CRISPR-Cas9 barcode editing to generate mutations in genomically integrated, engineered DNA cassettes, which are read out by single-cell RNA sequencing and used to produce high-resolution lineage trees. Here, we describe optimized cloning and validation procedures to construct the molecular recorder lineage tracing system. We include information on considerations of technology design, cloning procedures, the generation of lineage tracing cell lines, and time course experiments to assess their performance.

Genome editing technologies have revolutionized biomedical sciences and biotechnology. However, their delivery in vivo remains one of the major obstacles for clinical translation. Here, we introduce various emerging genome editing systems and review different delivery systems have been developed to realize the promise of in vivo gene editing therapies. In particular, we focus on virus-like particles (VLPs), an emerging delivery platform and provide in depth analysis on recent advancements to improve VLPs delivery potential and highlight opportunities for future improvements. To this end, we also provide detail workflows for engineered VLP (eVLP) selection, production, and purification, along with methods for characterization and validation.

The advent of CRISPR-based technologies has enabled the rapid advancement of programmable gene manipulation in cells, tissues, and whole organisms. An emerging platform for targeted gene perturbation is epigenetic editing, the direct editing of chemical modifications on DNA and histones that ultimately results in repression or activation of the targeted gene. In contrast to CRISPR nucleases, epigenetic editors modulate gene expression without inducing DNA breaks or altering the genomic sequence of host cells. Recently, we developed the CRISPRoff epigenetic editing technology that simultaneously establishes DNA methylation and repressive histone modifications at targeted gene promoters. Transient expression of CRISPRoff and the accompanying single guide RNAs in mammalian cells results in transcriptional repression of targeted genes that is memorized heritably by cells through cell division and differentiation. Here, we describe our protocol for the delivery of CRISPRoff through plasmid DNA transfection, as well as the delivery of CRISPRoff mRNA, into transformed human cell lines and primary immune cells. We also provide guidance on evaluating target gene silencing and highlight key considerations when utilizing CRISPRoff for gene perturbations. Our protocols are broadly applicable to other CRISPR-based epigenetic editing technologies, as programmable genome manipulation tools continue to evolve rapidly.

Group II introns are transposable elements that can propagate in host genomes through the "copy and paste" mechanism. They usually comprise RNA and protein components for effective propagation. Recently, we found that some bacterial GII-C introns without protein components had multiple copies in their resident genomes, implicating their potential transposition activity. We demonstrated that some of these systems are active for hydrolytic DNA cleavage and proved their DNA manipulation capability in bacterial or mammalian cells. These introns are therefore named HYdrolytic Endonucleolytic Ribozymes (HYERs). Here, we provide a detailed protocol for the systematic identification and characterization of HYERs and present our perspectives on its potential application in nucleic acid manipulation.

Programmable genome editing technologies have revolutionized the ability of researchers to alter the genomes of microorganisms in a straightforward and efficient manner, significantly advancing the field of microbiology. To date, several CRISPR-Cas-based genome-editing systems have been developed for use in E. coli, including CRISPR/Cas9, base editing, and prime editing technologies. In this chapter, we describe the design and experimental application of BacPE, a variant of prime editing technology optimized for E. coli. BacPE facilitates the introduction of point mutations, insertions, and deletions without the need for double-strand DNA breaks. We demonstrate that BacPE is a powerful tool for genome editing in E. coli and highlight its potential applicability to other bacterial species.

TnpB proteins encoded in IS200/IS605 and IS607 mobile genetic elements are among the most widespread proteins in the microbial world. They function as RNA-guided DNA nucleases that play a critical role in transposon proliferation and are the predecessors of CRISPR-Cas12 effector proteins of the type V CRISPR-Cas family. Small size of TnpB nucleases makes them an attractive alternative for larger Cas9 and Cas12 proteins in genome editing applications. However, only a small fraction of TnpB nucleases characterized to date are active in human cells, highlighting the need to identify new TnpB variants that can function as genome editors. Here, we present an experimental pipeline for the characterization of TnpB proteins by combining in silico analysis with in vitro assays. To validate it we determined guide RNA and identified TAM for a set of TnpB orthologs. The proposed workflow can be employed for rapid screening and characterization of the huge TnpB protein family to identify novel TnpB variants that might expand the genome editing toolbox.

Genome editing has garnered significant attention over the last decade, resulting in a massive expansion of the genome engineering toolbox. Base editors encompass a class of tools that enable installing single-nucleotide changes in genomic DNA without the use of double-strand breaks. With the ever-increasing development of new and/or improved base editor systems, it is easy to be overwhelmed by the abundance of options. Here, we provide clear guidance to facilitate the selection of a base editor and to design guide RNAs (gRNAs) to suit various needs. Additionally, we describe in detail how to generate gRNA plasmids, transfect various mammalian cell types, and evaluate editing efficiencies. Finally, we give alternative methods and troubleshooting tips for some common pitfalls encountered during base editing.

CRISPR-Cas systems use RNA-guided CRISPR-associated (Cas) effectors to neutralize infections in bacteria and archaea. In class 2 CRISPR-Cas systems, Cas9 and Cas12 are single-protein Cas effectors that target double-stranded DNA based on complementarity to the guide RNA before cleaving the target DNA using metal-dependent endonuclease domains. Cas9 and Cas12 proteins can be readily programmed to target any DNA of interest by changing the guiding RNA sequence and have been co-opted for genome editing and other biotechnology purposes. The effect of metal ion concentration is an essential consideration in the physiological role of Cas immunity effectors as well as the biotechnological applications of Cas endonucleases. In this chapter, we describe methods for studying the effect of variable divalent metal ion conditions on the DNA binding and cleavage activities of well-studied Cas9 and Cas12a proteins.

Base editing and other precision editing agents have transformed the utility and therapeutic potential of CRISPR-based genome editing. While some native enzymes edit efficiently with their nature-derived function, many enzymes require rational engineering or directed evolution to enhance the compatibility with mammalian cell genome editing. While many methods of engineering and directed evolution exist, plate-based discrete evolution offers an ideal balance between ease of use and engineering power. Here, we describe a detailed method for the bacterial directed evolution of CRISPR base editors that compounds technical ease with flexibility of application.

Type IV CRISPR systems are phylogenetically diverse and poorly understood. However, recently, major strides have been made toward understanding type IV-A systems. In type IV-A systems, a multi-subunit ribonucleoprotein complex, called the Csf complex, uses a CRISPR-derived guide to bind double-stranded DNA, forming an R-loop to which a helicase called CRISPR-associated DinG (CasDinG) is recruited. It is proposed that the ATP-dependent helicase activity of CasDinG then unwinds duplex DNA near the targeting site, impairing RNA transcription, and gene expression. Here we describe methods used to investigate the type IV-A system from Pseudomonas aeruginosa strain 83 including a plasmid clearance assay, expression and purification of type IV ribonucleoprotein complexes and proteins, nucleic acid binding assays, and CasDinG helicase assays. These methods provide a foundation for future work aimed at understanding these enigmatic systems.