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CRISPR/Cas9-based genome editing can easily generate knockout mouse models by disrupting the gene sequence, but its efficiency for creating models that require either insertion of exogenous DNA (knock-in) or replacement of genomic segments is very poor. The majority of mouse models used in research involve knock-in (reporters or recombinases) or gene replacement (e.g., conditional knockout alleles containing exons flanked by LoxP sites). A few methods for creating such models have been reported that use double-stranded DNA as donors, but their efficiency is typically 1-10% and therefore not suitable for routine use. We recently demonstrated that long single-stranded DNAs (ssDNAs) serve as very efficient donors, both for insertion and for gene replacement. We call this method efficient additions with ssDNA inserts-CRISPR (Easi-CRISPR) because it is a highly efficient technology (efficiency is typically 30-60% and reaches as high as 100% in some cases). The protocol takes â^¼2 months to generate the founder mice.

The liver is composed of two epithelial cell types: hepatocytes and liver ductal cells. Culture conditions for expansion of human liver ductal cells in vitro as organoids were previously described in a protocol; however, primary human hepatocytes remained hard to expand, until recently. In this protocol, we provide full details of how we overcame this limitation, establishing culture conditions that facilitate long-term expansion of human fetal hepatocytes as organoids. In addition, we describe how to generate (multi) gene knockouts using CRISPR–Cas9 in both human fetal hepatocyte and adult liver ductal organoid systems. Using a CRISPR–Cas9 and homology-independent organoid transgenesis (CRISPR-HOT) approach, efficient gene knockin can be achieved in these systems. These gene knockin and knockout approaches, and their multiplexing, should be useful for a variety of applications, such as disease modeling, investigating gene functions and studying processes, such as cellular differentiation and cell division. The protocol to establish human fetal hepatocyte organoid cultures takes ~1–2 months. The protocols to genome engineer human liver ductal organoids and human fetal hepatocyte organoids take 2–3 months. Culture conditions are described for long-term expansion of human fetal hepatocytes as 3D organoids. Gene knockin and knockout approaches are also described for organoids derived from human fetal hepatocytes and human adult liver ductal cells.

Targeted insertion of transgenes at pre-determined plant genomic safe harbors provides a desirable alternative to insertions at random sites achieved through conventional methods. Most existing cases of targeted gene insertion in plants have either relied on the presence of a selectable marker gene in the insertion cassette or occurred at low frequency with relatively small DNA fragments (<1.8 kb). Here, we report the use of an optimized CRISPR-Cas9-based method to achieve the targeted insertion of a 5.2 kb carotenoid biosynthesis cassette at two genomic safe harbors in rice. We obtain marker-free rice plants with high carotenoid content in the seeds and no detectable penalty in morphology or yield. Whole-genome sequencing reveals the absence of off-target mutations by Cas9 in the engineered plants. These results demonstrate targeted gene insertion of marker-free DNA in rice using CRISPR-Cas9 genome editing, and offer a promising strategy for genetic improvement of rice and other crops. Existing examples of targeted gene insertion in plants either rely on a selectable marker gene or result in short DNA inserts. Here, the authors use an optimized CRISPR-Cas9 method to insert a 5.2 kb carotenoid biosynthesis cassette into genomic safe harbors in rice, and obtain marker-free lines with high carotenoid content.

Genome engineering using programmable nucleases enables homologous recombination (HR)-mediated gene knock-in. However, the labour used to construct targeting vectors containing homology arms and difficulties in inducing HR in some cell type and organisms represent technical hurdles for the application of HR-mediated knock-in technology. Here, we introduce an alternative strategy for gene knock-in using transcription activator-like effector nucleases (TALENs) and clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated 9 (Cas9) mediated by microhomology-mediated end-joining, termed the PITCh (Precise Integration into Target Chromosome) system. TALEN-mediated PITCh, termed TAL-PITCh, enables efficient integration of exogenous donor DNA in human cells and animals, including silkworms and frogs. We further demonstrate that CRISPR/Cas9-mediated PITCh, termed CRIS-PITCh, can be applied in human cells without carrying the plasmid backbone sequence. Thus, our PITCh-ing strategies will be useful for a variety of applications, not only in cultured cells, but also in various organisms, including invertebrates and vertebrates. One challenge facing the use of programmable nucleases in genome engineering is the requirement for homologous recombination. Here, Nakade et al. harness microhomology-mediated end-joining as a means of inserting exogenous coding sequences into the genome using both TALEN and CRISPR/Cas9 technologies.

To gain insight into how mutant huntingtin (mHtt) CAG repeat length modifies Huntington's disease pathogenesis, the authors profiled mRNA in over 600 brain and peripheral tissue samples from Huntington's disease knock-in mice with increasing CAG repeat lengths. Coexpression network analyses reveal 13 striatal and 5 cortical modules that are highly correlated with CAG length and age. To gain insight into how mutant huntingtin ( mHtt ) CAG repeat length modifies Huntington's disease (HD) pathogenesis, we profiled mRNA in over 600 brain and peripheral tissue samples from HD knock-in mice with increasing CAG repeat lengths. We found repeat length-dependent transcriptional signatures to be prominent in the striatum, less so in cortex, and minimal in the liver. Coexpression network analyses revealed 13 striatal and 5 cortical modules that correlated highly with CAG length and age, and that were preserved in HD models and sometimes in patients. Top striatal modules implicated mHtt CAG length and age in graded impairment in the expression of identity genes for striatal medium spiny neurons and in dysregulation of cyclic AMP signaling, cell death and protocadherin genes. We used proteomics to confirm 790 genes and 5 striatal modules with CAG length–dependent dysregulation at the protein level, and validated 22 striatal module genes as modifiers of mHtt toxicities in vivo .

CRISPR–Cas9 technology has revolutionized genome editing and is applicable to the organoid field. However, precise integration of exogenous DNA sequences into human organoids is lacking robust knock-in approaches. Here, we describe CRISPR–Cas9-mediated homology-independent organoid transgenesis (CRISPR–HOT), which enables efficient generation of knock-in human organoids representing different tissues. CRISPR–HOT avoids extensive cloning and outperforms homology directed repair (HDR) in achieving precise integration of exogenous DNA sequences into desired loci, without the necessity to inactivate TP53 in untransformed cells, which was previously used to increase HDR-mediated knock-in. CRISPR–HOT was used to fluorescently tag and visualize subcellular structural molecules and to generate reporter lines for rare intestinal cell types. A double reporter—in which the mitotic spindle was labelled by endogenously tagged tubulin and the cell membrane by endogenously tagged E-cadherin—uncovered modes of human hepatocyte division. Combining tubulin tagging with TP53 knock-out revealed that TP53 is involved in controlling hepatocyte ploidy and mitotic spindle fidelity. CRISPR–HOT simplifies genome editing in human organoids.Artegiani, Hendriks et al. describe a CRISPR–Cas9-based method to efficiently generate human knock-in organoids using non-homologous end joining to study rare intestinal cell types and human hepatocyte division.

Homologous recombination-based gene targeting is a powerful tool for precise genome modification and has been widely used in organisms ranging from yeast to higher organisms such as Drosophila and mouse. However, gene targeting in higher plants, including the most widely used model plant Arabidopsis thaliana , remains challenging. Here we report a sequential transformation method for gene targeting in Arabidopsis . We find that parental lines expressing the bacterial endonuclease Cas9 from the egg cell- and early embryo-specific DD45 gene promoter can improve the frequency of single-guide RNA-targeted gene knock-ins and sequence replacements via homologous recombination at several endogenous sites in the Arabidopsis genome. These heritable gene targeting can be identified by regular PCR. Our approach enables routine and fine manipulation of the Arabidopsis genome. Efficient gene targeting in higher plants remains challenging. Here, the authors develop a sequential transformation method for CRISPR/Cas9-mediated gene targeting in Arabidopsis and demonstrate its functionality at five genomic sites in two endogenous loci.

Inherited retinal degenerations are caused by mutations in >250 genes that affect photoreceptor cells or the retinal pigment epithelium and result in vision loss. For autosomal recessive and X-linked retinal degenerations, significant progress has been achieved in the field of gene therapy as evidenced by the growing number of clinical trials and the recent commercialization of the first gene therapy for a form of congenital blindness. However, despite significant efforts to develop a treatment for the most common form of autosomal dominant retinitis pigmentosa (adRP) caused by >150 mutations in the rhodopsin (RHO) gene, translation to the clinic has stalled. Here, we identified a highly efficient shRNA that targets human (and canine) RHO in a mutation-independent manner. In a single adeno-associated viral (AAV) vector we combined this shRNA with a human RHO replacement cDNA made resistant to RNA interference and tested this construct in a naturally occurring canine model of RHO-adRP. Subretinal vector injections led to nearly complete suppression of endogenous canine RHO RNA, while the human RHO replacement cDNA resulted in up to 30% of normal RHO protein levels. Noninvasive retinal imaging showed photoreceptors in treated areas were completely protected from retinal degeneration. Histopathology confirmed retention of normal photoreceptor structure and RHO expression in rod outer segments. Long-term (>8 mo) follow-up by retinal imaging and electroretinography indicated stable structural and functional preservation. The efficacy of this gene therapy in a clinically relevant large-animal model paves the way for treating patients with RHO-adRP.

Recent use of the CRISPR/Cas9 system has dramatically reduced the time required to produce mutant mice, but the involvement of a time-consuming microinjection step still hampers its application for high-throughput genetic analysis. Here we developed a simple, highly efficient and large-scale genome editing method, in which the RNAs for the CRISPR/Cas9 system are electroporated into zygotes rather than microinjected. We used this method to perform single-stranded oligodeoxynucleotide (ssODN)-mediated knock-in in mouse embryos. This method facilitates large-scale genetic analysis in the mouse.

CRISPR–Cas9 paired with adeno-associated virus serotype 6 (AAV6) is among the most efficient tools for producing targeted gene knockins. Here, we report that this system can lead to frequent concatemeric insertions of the viral vector genome at the target site that are difficult to detect. Such errors can cause adverse and unreliable phenotypes that are antithetical to the goal of precision genome engineering. The concatemeric knockins occurred regardless of locus, vector concentration, cell line or cell type, including human pluripotent and hematopoietic stem cells. Although these highly abundant errors were found in more than half of the edited cells, they could not be readily detected by common analytical methods. We describe strategies to detect and thoroughly characterize the concatemeric viral vector insertions, and we highlight analytical pitfalls that mask their prevalence. We then describe strategies to prevent the concatemeric inserts by cutting the vector genome after transduction. This approach is compatible with established gene editing pipelines, enabling robust genetic knockins that are safer, more reliable and more reproducible. AAV vectors form difficult-to-detect concatemers at Cas9 target sites.