Genome editing in the fall armyworm, Spodoptera frugiperda: Multiple sgRNA/Cas9 method for identification of knockouts in one generation.

The CRISPR/Cas9 system is an efficient genome editing method that can be used in functional genomics research. The fall armyworm, Spodoptera frugiperda, is a serious agricultural pest that has spread over most of the world. However, very little information is available on functional genomics for this insect. We performed CRISPR/Cas9-mediated site-specific mutagenesis of three target genes: two marker genes [Biogenesis of lysosome-related organelles complex 1 subunit 2 (BLOS2) and tryptophan 2, 3-dioxygenase (TO)], and a developmental gene, E93 (a key ecdysone-induced transcription factor that promotes adult development). The knockouts (KO) of BLOS2, TO and E93 induced translucent mosaic integument, olive eye color, and larval-pupal intermediate phenotypes, respectively. Sequencing RNA isolated from wild-type and E93 KO insects showed that E93 promotes adult development by influencing the expression of the genes coding for transcription factor, Krüppel homolog 1, the pupal specifier, Broad-Complex, serine proteases, and heat shock proteins. Often, gene-edited insects display mosaicism in which only a fraction of the cells are edited as intended, and establishing a homozygous line is both costly and time-consuming. To overcome these limitations, a method to completely KO the target gene in S. frugiperda by injecting the Cas9 protein and multiple sgRNAs targeting one exon of the E93 gene into embryos was developed. Ten percent of the G0 larvae exhibited larval-pupal intermediates. The mutations were confirmed by T7E1 assay, and the mutation frequency was determined as >80%. Complete KO of the E93 gene was achieved in one generation using the multiple sgRNA method, demonstrating a powerful approach to improve genome editing in lepidopteran and other non-model insects.

Electroporation of CRISPR-Cas9 into Malignant B Cells for Loss-of-Function Studies of Target Gene Via Knockout.

CRISPR-Cas9 is a unique technology that enables geneticists and medical researchers to edit genomic DNA for studying biology, pathogenesis, and molecular basis of treatment in malignant B cells. Unfortunately, malignant B cells are extremely difficult to transfect by most traditional methods. In this chapter, we describe the use of the Nucleofector™ Technology-based electroporation system with optimized transfection conditions for generating a malignant B cell model, JEKO-1, with ROR1-gene knockout via CRISPR-Cas9 technology.

Structure, function and techniques of investigation of the P2X7 receptor (P2X7R) in mammalian cells.

The P2X7 receptor [P2X7R or P2RX7 in National Center for Biotechnology Information (NCBI) gene nomenclature] is a member of the P2X receptor (P2XR) subfamily of P2 receptors (P2Rs). The P2X7R is an extracellular ATP-gated ion channel with peculiar permeability properties expressed by most cell types, mainly in the immune system, where it has a leading role in cytokine release, oxygen radical generation, T lymphocyte differentiation and proliferation. A role in cancer cell growth and tumor progression has also been demonstrated. These features make the P2X7R an appealing target for drug development in inflammation and cancer. The functional P2X7R, recently (partially) crystallized and 3-D solved, is formed by the assembly of three identical subunits (homotrimer). The P2X7R is preferentially permeable to small cations (Ca2+, Na+, K+), and in most (but not all) cell types also to large positively charged molecules of molecular mass up to 900Da. Permeability to negatively charged species of comparable molecular mass (e.g., Lucifer yellow) is debated. Several highly selective P2X7R pharmacological blockers have been developed over the years, thus providing powerful tools for P2X7R studies. Biophysical properties and coupling to several different physiological responses make the P2X7R amenable to investigation by electrophysiology and cell biology techniques, which allow its identification and characterization in many different cell types and tissues. A careful description of the physiological features of the P2X7R is a prerequisite for an effective therapeutic development. Here we describe the most common techniques to asses P2X7R functions, including patch-clamp, intracellular calcium measurements, and membrane permeabilization to large fluorescent dyes in a selection of different cell types. In addition, we also describe common toxicity assays used to verify the effects of P2X7R stimulation on cell viability.

CRISPR/Cas9-based knockout pipeline for reverse genetics in mammalian cell culture.

We present a straightforward protocol for reverse genetics in cultured mammalian cells, using CRISPR/Cas9-mediated homology-dependent repair (HDR) based insertion of a protein trap cassette, resulting in a termination of the endogenous gene expression. Complete loss of function can be achieved with monoallelic trap cassette insertion, as the second allele is frequently disrupted by an error-prone non-homologous end joining (NHEJ) mechanism. The method should be applicable to any expressed gene in most cell lines, including those with low HDR efficiency, as the knockout alleles can be directly selected for.

Use of Organoids to Characterize Signaling Pathways in Cancer Initiation.

The development of intestinal organoid technology has greatly accelerated research in the field of colorectal cancer. Contrary to traditional cancer cell lines, organoids are composed of multiple cell types arranged in 3D structures highly reminiscent of their native tissues. Thus, organoids provide a near-physiological and readily accessible model to study tissue morphogenesis, adult stem cell behavior and tumorigenesis. Here, we provide protocols for establishing intestinal organoid cultures from genetically modified mouse lines and describe methods to overexpress and knockout genes of interest using lentiviral-based approaches.

Pyviko: an automated Python tool to design gene knockouts in complex viruses with overlapping genes.

BACKGROUND: Gene knockouts are a common tool used to study gene function in various organisms. However, designing gene knockouts is complicated in viruses, which frequently contain sequences that code for multiple overlapping genes. Designing mutants that can be traced by the creation of new or elimination of existing restriction sites further compounds the difficulty in experimental design of knockouts of overlapping genes. While software is available to rapidly identify restriction sites in a given nucleotide sequence, no existing software addresses experimental design of mutations involving multiple overlapping amino acid sequences in generating gene knockouts. RESULTS: Pyviko performed well on a test set of over 240,000 gene pairs collected from viral genomes deposited in the National Center for Biotechnology Information Nucleotide database, identifying a point mutation which added a premature stop codon within the first 20 codons of the target gene in 93.2% of all tested gene-overprinted gene pairs. This shows that Pyviko can be used successfully in a wide variety of contexts to facilitate the molecular cloning and study of viral overprinted genes. CONCLUSIONS: Pyviko is an extensible and intuitive Python tool for designing knockouts of overlapping genes. Freely available as both a Python package and a web-based interface ( http://louiejtaylor.github.io/pyViKO/ ), Pyviko simplifies the experimental design of gene knockouts in complex viruses with overlapping genes.

CRISPR/Cas9-mediated gene knockout in the mouse brain using in utero electroporation.

The CRISPR/Cas9 system has recently been adapted for generating knockout mice to investigate physiological functions and pathological mechanisms. Here, we report a highly efficient procedure for brain-specific disruption of genes of interest in vivo. We constructed pX330 plasmids expressing humanized Cas9 and single-guide RNAs (sgRNAs) against the Satb2 gene, which encodes an AT-rich DNA-binding transcription factor and is responsible for callosal axon projections in the developing mouse brain. We first confirmed that these constructs efficiently induced double-strand breaks (DSBs) in target sites of exogenous plasmids both in vitro and in vivo. We then found that the introduction of pX330-Satb2 into the developing mouse brain using in utero electroporation led to a dramatic reduction of Satb2 expression in the transfected cerebral cortex, suggesting DSBs had occurred in the Satb2 gene with high efficiency. Furthermore, we found that Cas9-mediated targeting of the Satb2 gene induced abnormalities in axonal projection patterns, which is consistent with the phenotypes previously observed in Satb2 mutant mice. Introduction of pX330-NeuN using our procedure also resulted in the efficient disruption of the NeuN gene. Thus, our procedure combining the CRISPR/Cas9 system and in utero electroporation is an effective and rapid approach to achieve brain-specific gene knockout in vivo.

Recombineering: using drug cassettes to knock out genes in vivo.

A 'gene knockout' or 'knockout' is a mutation that inactivates a gene function. These mutations are very useful for classical genetic studies as well as for modern techniques including functional genomics. In the past, knockouts of bacterial genes were often made by transposon mutagenesis. In this case, laborious screens are required to find a knockout in the gene of interest. Knockouts of other organisms have traditionally been made by first using in vitro genetic engineering to modify genes contained on plasmids or bacterial artificial chromosomes (BACs) and later moving these modified constructs to the organism of interest by cell culture techniques. Other methods utilizing a combination of genetic engineering and in vivo homologous recombination were inefficient at best. Recombineering provides a new way to generate knockout mutations directly on the bacterial chromosome or to modify any plasmid or BAC in vivo as a prelude to making knockouts in other organisms. The constructs are designed to the base pair and are not dependent on suitable restriction sites. A drug cassette can be placed anywhere within a gene or the open reading frame of the gene can be replaced with the drug cassette. Either way, the desired construct is selected for.

Gene knockouts, in vivo site-directed mutagenesis and other modifications using the delitto perfetto system in Saccharomyces cerevisiae.

Gene manipulation serves the purpose of providing a better understanding of the function of specific genes as well as for developing novel variants of the genes of interest. The generation of knockout genes, the alteration, depletion, or enhancement of a particular gene function through the generation of specific gene mutations, or the generation of random mutations in a gene are all essential processes for gene manipulation. The genome of the yeast Saccharomyces cerevisiae is relatively easy to modify, owing to its efficient homologous recombination (HR) system. Gene knockout can be a very simple, one-step approach to eliminate a gene by substituting its DNA sequence with that of a genetic marker. Differently, desired mutations can be introduced into a gene by replacing the sequence of the normal gene with that of the mutated gene. Recombinant DNA can be created in vitro and then introduced into cells, most often exploiting the endogenous recombination system of the cells. However, unless the desired mutation gives a particular phenotype, a bottleneck of 'recombineering' is the requirement of a selection system to identify the recombinant clones among those unmodified. Even in an organism like yeast where the level of HR is highly above the incidence of random integration, the frequency of homologous targeting is in the range of 10(-4)-10(-6) depending on the length of the homology used (Wach et al., 1994). Thus, a selection system is always required to identify the targeted clones. Counterselectable markers, such as URA3, LYS2, LYS5, MET15, and TRP1 (Bach and LaCroute, 1972; Chattoo et al., 1979; Singh and Sherman, 1974; Toyn et al., 2000), are widely utilized in yeast and can be recycled for additional usage in the same yeast strain. If the marker is not eliminated or it is popped out via site-specific recombination between direct repeats, such as in the Flp/FRT or Cre/Lox systems, a heterologous sequence is left as a scar at the site of the modified DNA (Storici et al., 1999; Sauer, 1987). The presence of such scars can threaten the genomic stability of the strain and/or limit the number of successive genetic manipulations for that strain. Here, we describe the delitto perfetto approach for in vivo mutagenesis that combines the practicality of a general selection system with the versatility of synthetic oligonucleotides for targeting (Storici et al., 2001). It provides for generation of gene knockouts and almost any sort of mutation and genome rearrangement via HR. The delitto perfetto in vivo mutagenesis technique is designed for efficient and precise manipulation of yeast strains in a two-step process spanning ~2 weeks. Here, we present the theory and procedures of the delitto perfetto technique.

Application of the FLP/FRT system for conditional gene deletion in yeast Saccharomyces cerevisiae.

The yeast Saccharomyces cerevisiae has proved to be an excellent model organism to study the function of proteins. One of the many advantages of yeast is the many genetic tools available to manipulate gene expression, but there are still limitations. To complement the many methods used to control gene expression in yeast, we have established a conditional gene deletion system by using the FLP/FRT system on yeast vectors to conditionally delete specific yeast genes. Expression of Flp recombinase, which is under the control of the GAL1 promoter, was induced by galactose, which in turn excised FRT sites flanked genes. The efficacy of this system was examined using the FRT site-flanked genes HSP104, URA3 and GFP. The pre-excision frequency of this system, which might be caused by the basal activity of the GAL1 promoter or by spontaneous recombination between FRT sites, was detected ca. 2% under the non-selecting condition. After inducing expression of Flp recombinase, the deletion efficiency achieved ca. 96% of cells in a population within 9 h. After conditional deletion of the specific gene, protein degradation and cell division then diluted out protein that was expressed from this gene prior to its excision. Most importantly, the specific protein to be deleted could be expressed under its own promoter, so that endogenous levels of protein expression were maintained prior to excision by the Flp recombinase. Therefore, this system provides a useful tool for the conditional deletion of genes in yeast.