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Ulva prolifera is a fast-growing green seaweed that has garnered considerable interest in both fundamental and applied research. Here, we established a molecular tool by employing a selectable marker gene that allowed the isolation of U. prolifera cells integrating exogenous DNA. We developed a modular plasmid for expressing exogenous genes in U. prolifera based on the bacterial antibiotic-resistance marker, aminoglycoside phosphotransferase gene ( aph7” ). Integration of aph7” in macroalgae can generate transformants resistant to hygromycin B. In addition, we characterized the promoter region of the small subunit of ribulose-1,5-bisphosphate carboxylase/oxygenase gene (pUpRbcS) to drive the expression of aph7” . The transcripts were consistently confirmed from antibiotic-selected transformants, stably retaining the exogenous gene in the succeeding generations. Subsequently, a CRISPR-based knock-in system was established, facilitating the integration of aph7” cassette in the endogenous selection gene encoding for adenine phosphoribosyltransferase ( UpAPT ). APT gene can serve as an endogenous marker in algae that exhibits a lethal phenotype under cultivation with 2-fluoroadenine. The resulting knock-in mutants could resist the co-selection of the antibiotic hygromycin B and 2-fluoroadenine. Our results advance U. prolifera as a genetic platform, enabling functional research to elucidate Ulva biology, and to bring forth biotechnological utilization of algal resources.

Efficient precision genome engineering requires high frequency and specificity of integration at the genomic target site. Here, we describe a set of resources to streamline reporter gene knock-ins in zebrafish and demonstrate the broader utility of the method in mammalian cells. Our approach uses short homology of 24–48 bp to drive targeted integration of DNA reporter cassettes by homology-mediated end joining (HMEJ) at high frequency at a double strand break in the targeted gene. Our vector series, pGTag (plasmids for Gene Tagging), contains reporters flanked by a universal CRISPR sgRNA sequence which enables in vivo liberation of the homology arms. We observed high rates of germline transmission (22–100%) for targeted knock-ins at eight zebrafish loci and efficient integration at safe harbor loci in porcine and human cells. Our system provides a straightforward and cost-effective approach for high efficiency gene targeting applications in CRISPR and TALEN compatible systems.

Background Conditional knockout mice and transgenic mice expressing recombinases, reporters, and inducible transcriptional activators are key for many genetic studies and comprise over 90% of mouse models created. Conditional knockout mice are generated using labor-intensive methods of homologous recombination in embryonic stem cells and are available for only ~25% of all mouse genes. Transgenic mice generated by random genomic insertion approaches pose problems of unreliable expression, and thus there is a need for targeted-insertion models. Although CRISPR-based strategies were reported to create conditional and targeted-insertion alleles via one-step delivery of targeting components directly to zygotes, these strategies are quite inefficient. Results Here we describe Easi- CRISPR ( E fficient a dditions with s sDNA i nserts-CRISPR), a targeting strategy in which long single-stranded DNA donors are injected with pre-assembled crRNA + tracrRNA + Cas9 ribonucleoprotein (ctRNP) complexes into mouse zygotes. We show for over a dozen loci that Easi -CRISPR generates correctly targeted conditional and insertion alleles in 8.5–100% of the resulting live offspring. Conclusions Easi- CRISPR solves the major problem of animal genome engineering, namely the inefficiency of targeted DNA cassette insertion. The approach is robust, succeeding for all tested loci. It is versatile, generating both conditional and targeted insertion alleles. Finally, it is highly efficient, as treating an average of only 50 zygotes is sufficient to produce a correctly targeted allele in up to 100% of live offspring. Thus, Easi- CRISPR offers a comprehensive means of building large-scale Cre- LoxP animal resources.

Targeted integration of transgenes can be achieved by strategies based on homologous recombination （HR）, mi- crohomology-mediated end joining （MMEJ） or non-homologous end joining （NHEJ）. The more generally used HR is inefficient for achieving gene integration in animal embryos and tissues, because it occurs only during cell division, although MMEJ and NHEJ can elevate the efficiency in some systems. Here we devise a homology-mediated end joining （HMEJ）-based strategy, using CRISPR/Cas9-mediated cleavage of both transgene donor vector that contains guide RNA target sites and -800 bp of homology arms, and the targeted genome. We found no significant improve- ment of the targeting efficiency by the HMEJ-based method in either mouse embryonic stem cells or the neuroblas- toma cell line, N2a, compared to the HR-based method. However, the HMEJ-based method yielded a higher knock- in efficiency in HEK293T cells, primary astrocytes and neurons. More importantly, this approach achieved transgene integration in mouse and monkey embryos, as well as in hepatocytes and neurons in vivo, with an efficiency much greater than HR-, NHEJ- and MMEJ-based strategies. Thus, the HMEJ-based strategy may be useful for a variety of applications, including gene editing to generate animal models and for targeted gene therapies.

Summary Genome editing based on the homology‐directed repair (HDR) pathway enables scar‐free and precise genetic manipulations. However, the low frequency of HDR hinders its application in plant genome editing. In this study, we engineered the fusion of Cas9 and a viral replication protein (Rep) as a molecular bridge to tether donor DNA in vivo, which enhances the efficiency of targeted gene insertion via the HDR pathway. This Rep‐bridged knock‐in (RBKI) method combines the advantages of rolling cycle replication of viral replicons and in vivo enrichment of donor DNA at the target site for HDR. Chromatin immunoprecipitation indicated that the Cas9‐Rep fusion protein bound up to 66‐fold more donor DNA than Cas9 did. We exemplified the RBKI method by inserting small‐ to middle‐sized tags (33–519 bp) into 3 rice genes. Compared to Cas9, Cas9‐Rep fusion increased the KI frequencies by 4–7.6‐fold, and up to 72.2% of stable rice transformants carried in‐frame knock‐in events in the T0 generation. Whole‐genome sequencing of 6 plants segregated from heterozygous KI lines indicated that the knock‐in events were faithfully inherited by the progenies with neither off‐target editing nor random insertions of the donor DNA fragment. Further analysis suggested that the RBKI method reduced the number of byproducts from nonhomologous end joining; however, HDR‐mediated knock‐in tended to accompany microhomology‐mediated end joining events. Together, these findings show that the in vivo tethering of donor DNAs with Cas9‐Rep is an effective strategy to increase the frequency of HDR‐mediated genome editing.

We present a robust method called improved-Genome editing via Oviductal Nucleic Acids Delivery ( i -GONAD) that delivers CRISPR ribonucleoproteins to E0.7 embryos via in situ electroporation. The method generates mouse models containing single-base changes, kilobase-sized deletions, and knock-ins. The efficiency of i -GONAD is comparable to that of traditional microinjection methods, which rely on ex vivo handling of zygotes and require recipient animals for embryo transfer. In contrast, i- GONAD avoids these technically difficult steps, and it can be performed at any laboratory with simple equipment and technical expertise. Further, i -GONAD-treated females retain reproductive function, suggesting future use of the method for germline gene therapy.

Genetically engineered mouse models (GEMMs) are crucial for investigating disease mechanisms, developing therapeutic strategies, and advancing fundamental biological research. While CRISPR gene editing has greatly facilitated the creation of these models, existing techniques still present technical challenges and efficiency limitations. Here, we establish a CRISPR-VLP-induced targeted mutagenesis (CRISPR-VIM) strategy, enabling precise genome editing by co-culturing zygotes with virus-like particle (VLP)-delivered gene editing ribonucleoproteins (RNPs) without requiring physical manipulation or causing cellular damage. We generate Plin1- and Tyr -knockout mice through VLP-based SpCas9 or adenine base editor (ABE)/sgRNA RNPs and characterize their phenotype and germline transmission. Additionally, we demonstrate cytosine base editor (CBE)/sgRNA-based C-to-T substitution or SpCas9/sgRNA-based knock-in using VLPs. This method further simplifies and accelerates GEMM generation without specialized techniques or equipment. Consequently, the CRISPR-VIM method can facilitate mouse modeling and be applied in various research fields. Gene-edited mouse models are crucial for disease research but remain challenging to create. Here, authors introduce the CRISPR-VIM, using virus-like particles to efficiently deliver CRISPR tools into zygotes without physical damage, streamlining the creation of genetically engineered mouse models.

Quantifying the genetic heterogeneity of a cell population is essential to understanding of biological systems. We develop a universal method to label individual DNA molecules for single-base-resolution haplotype-resolved quantitative characterization of diverse types of rare variants, with frequency as low as 4 × 10 −5 , using both short- or long-read sequencing platforms. It provides the first quantitative evidence of persistent nonrandom large structural variants and an increase in single-nucleotide variants at the on-target locus following repair of double-strand breaks induced by CRISPR-Cas9 in human embryonic stem cells.

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.

Background The CRISPR-Cas9 system is a widely utilized platform for transgenic animal production in various species, although its off-target effects should be addressed. Several applications of this tool have been proposed in model animals but remain insufficient for transgenic livestock production. Results Here, we report the first application of single Cas9 nickase (Cas9n) to induce gene insertion at a selected locus in cattle. We identify the main binding sites of a catalytically inactive Cas9 (dCas9) protein in bovine fetal fibroblast cells (BFFs) with chromatin immunoprecipitation sequencing (ChIP-seq). Subsequently, we demonstrate that a single Cas9n-induced single-strand break can stimulate the insertion of the natural resistance-associated macrophage protein-1 ( NRAMP1 ) gene with reduced, but still considerable, off-target effects. Through somatic cell nuclear transfer, we finally obtain transgenic cattle with increased resistance to tuberculosis. Conclusions Our results contribute to the development of CRISPR-Cas9 system for agriculture applications.