Explore the vast range of titles available.

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.

Although single-component Class 2 CRISPR systems, such as type II Cas9 or type V Cas12a (Cpf1), are widely used for genome editing in eukaryotic cells, the application of multi-component Class 1 CRISPR has been less developed. Here we demonstrate that type I-E CRISPR mediates distinct DNA cleavage activity in human cells. Notably, Cas3, which possesses helicase and nuclease activity, predominantly triggered several thousand base pair deletions upstream of the 5′-ARG protospacer adjacent motif (PAM), without prominent off-target activity. This Cas3-mediated directional and broad DNA degradation can be used to introduce functional gene knockouts and knock-ins. As an example of potential therapeutic applications, we show Cas3-mediated exon-skipping of the Duchenne muscular dystrophy ( DMD ) gene in patient-induced pluripotent stem cells (iPSCs). These findings broaden our understanding of the Class 1 CRISPR system, which may serve as a unique genome editing tool in eukaryotic cells distinct from the Class 2 CRISPR system. Class 1 CRISPR systems are not as developed for genome editing as Class 2 systems are. Here the authors show that Cas3 can be used to generate functional knockouts and knock-ins, as well as Cas3-mediated exon-skipping in DMD cells.

Enzyme turnover numbers (k cats) are essential for a quantitative understanding of cells. Because k cats are traditionally measured in low-throughput assays, they can be inconsistent, labor-intensive to obtain, and can miss in vivo effects. We use a data-driven approach to estimate in vivo k cats using metabolic specialist Escherichia coli strains that resulted from gene knockouts in central metabolism followed by metabolic optimization via laboratory evolution. By combining absolute proteomics with fluxomics data, we find that in vivo k cats are robust against genetic perturbations, suggesting that metabolic adaptation to gene loss is mostly achieved through other mechanisms, like gene-regulatory changes. Combining machine learning and genome-scale metabolic models, we show that the obtained in vivo k cats predict unseen proteomics data with much higher precision than in vitro k cats. The results demonstrate that in vivo k cats can solve the problem of inconsistent and low-coverage parameterizations of genome-scale cellular models.

Summary Plants offer fast, flexible and easily scalable alternative platforms for the production of pharmaceutical proteins, but differences between plant and mammalian N‐linked glycans, including the presence of β‐1,2‐xylose and core α‐1,3‐fucose residues in plants, can affect the activity, potency and immunogenicity of plant‐derived proteins. Nicotiana benthamiana is widely used for the transient expression of recombinant proteins so it is desirable to modify the endogenous N‐glycosylation machinery to allow the synthesis of complex N‐glycans lacking β‐1,2‐xylose and core α‐1,3‐fucose. Here, we used multiplex CRISPR/Cas9 genome editing to generate N. benthamiana production lines deficient in plant‐specific α‐1,3‐fucosyltransferase and β‐1,2‐xylosyltransferase activity, reflecting the mutation of six different genes. We confirmed the functional gene knockouts by Sanger sequencing and mass spectrometry‐based N‐glycan analysis of endogenous proteins and the recombinant monoclonal antibody 2G12. Furthermore, we compared the CD64‐binding affinity of 2G12 glycovariants produced in wild‐type N. benthamiana, the newly generated FX‐KO line, and Chinese hamster ovary (CHO) cells, confirming that the glyco‐engineered antibody performed as well as its CHO‐produced counterpart.

With the groundbreaking advancements in genome editing technologies, particularly CRISPR-Cas9, creating knockout mutants has become highly efficient. However, the CRISPR-Cas9 system introduces DNA double-strand breaks, increasing the risk of chromosomal rearrangements and posing a major obstacle to simultaneous multiple gene knockout. Base-editing systems, such as Target-AID, are safe alternatives for precise base modifications without requiring DNA double-strand breaks, serving as promising solutions for existing challenges. Nevertheless, the absence of adequate tools to support Target-AID-based gene knockout highlights the need for a comprehensive system to design guide RNAs (gRNAs) for the simultaneous knockout of multiple genes. Here, we aimed to develop KOnezumi-AID, a command-line tool for gRNA design for Target-AID-mediated genome editing. KOnezumi-AID facilitates gene knockout by inducing the premature termination codons or promoting exon skipping, thereby generating experiment-ready gRNA designs for mouse and human genomes. Additionally, KOnezumi-AID exhibits batch processing capacity, enabling rapid and precise gRNA design for large-scale genome editing, including CRISPR screening. In summary, KOnezumi-AID is an efficient and user-friendly tool for gRNA design, streamlining genome editing workflows and advancing gene knockout research.

G protein-independent, arrestin-dependent signaling is a paradigm that broadens the signaling scope of G protein-coupled receptors (GPCRs) beyond G proteins for numerous biological processes. However, arrestin signaling in the collective absence of functional G proteins has never been demonstrated. Here we achieve a state of “zero functional G” at the cellular level using HEK293 cells depleted by CRISPR/Cas9 technology of the Gs/q/12 families of Gα proteins, along with pertussis toxin-mediated inactivation of Gi/o. Together with HEK293 cells lacking β-arrestins (“zero arrestin”), we systematically dissect G protein- from arrestin-driven signaling outcomes for a broad set of GPCRs. We use biochemical, biophysical, label-free whole-cell biosensing and ERK phosphorylation to identify four salient features for all receptors at “zero functional G”: arrestin recruitment and internalization, but—unexpectedly—complete failure to activate ERK and whole-cell responses. These findings change our understanding of how GPCRs function and in particular of how they activate ERK1/2. Arrestins terminate signaling from GPCRs, but several lines of evidence suggest that they are also able to transduce signals independently of G proteins. Here, the authors systematically ablate G proteins in cell lines, and show that arrestins are unable to act as genuine signal initiators.

• Plant cellulose is synthesized by a large plasma membrane-localized cellulose synthase (CesA) complex. However, an overall functional determination of secondary cell wall (SCW) CesAs is still lacking in trees, especially one based on gene knockouts. • Here, the Cas9/gRNA-induced knockouts of PtrCesA4, 7A, 7B, 8A and 8B genes were produced in Populus trichocarpa. Based on anatomical, immunohistochemical and wood composition evidence, we gained a comprehensive understanding of five SCW PtrCesAs at the genetic level. • Complete loss of PtrCesA4, 7A/B or 8A/B led to similar morphological abnormalities, indicating similar and nonredundant genetic functions. The absence of the gelatinous (G) layer, one-layer-walled fibres and a 90% decrease in cellulose in these mutant woods revealed that the three classes of SCW PtrCesAs are essential for multilayered SCW structure and wood G-fibre. In addition, the mutant primary and secondary phloem fibres lost the n(G + L)- and G-layers and retained the thicker S-layers (L, lignified; S, secondary). Together with polysaccharide immunolocalization data, these findings suggest differences in the role of SCW PtrCesAs-synthesized cellulose in wood and phloem fibre wall structures. • Overall, this functional understanding of the SCW PtrCesAs provides further insights into the impact of lacking cellulose biosynthesis on growth, SCW, wood G-fibre and phloem fibre wall structures in the tree.

Due to an unexpected cell-penetrating property, zinc-finger nucleases (ZFNs) can be delivered to several mammalian cell types as proteins. Dose-dependent disruption of an endogenous gene was achieved with reduced activity at known off-target sites. Zinc-finger nucleases (ZFNs) are versatile reagents that have redefined genome engineering. Realizing the full potential of this technology requires the development of safe and effective methods for delivering ZFNs into cells. We demonstrate the intrinsic cell-penetrating capabilities of the standard ZFN architecture and show that direct delivery of ZFNs as proteins leads to efficient endogenous gene disruption in various mammalian cell types with minimal off-target effects.

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 .

Polysorbate (PS) is routinely used in biopharmaceutical formulation to stabilize the active pharmaceutical ingredient.Trace amounts of hydrolytic host cell proteins (HCPs) persist in the downstream purification process and can degrade PS in the final drug product over time, compromising its stability.Genomic removal of PS-degrading enzymes in the Chinese hamster ovary (CHO) host cell line allowed their traceless removal from the bioprocess.The combined knockout (KO) of the genes encoding nine previously confirmed PS-degrading HCPs resulted in a viable CHO host cell line with strongly reduced PS degradation potential.This is the first report of successful excision of a gene cluster >1 Mb in size from the CHO genome.The final multi-hydrolase KO cell line yielded competitive monoclonal antibody titers with high product quality and significantly reduced hydrolytic activity. Enzymatic degradation of polysorbates (PS) in biologic drug formulations is often caused by hydrolytic host cell proteins (HCPs) and can lead to particle formation and reduced shelf-life. Here, we present a host cell line-engineering approach by removing nine Chinese hamster ovary (CHO) host cell hydrolases, which have previously been confirmed to degrade PS. Strikingly, the sequential genomic knockout (KO) of these hydrolases, including the genetic removal of two entire gene clusters of unprecedented size, yielded viable CHO host cell line variants. This novel host cell line was further optimized using the additional KO of the two key proapoptotic genes, Bax and Bak1. Thus, we generated a competitive multi-hydrolase KO CHO host cell line that was further shown to be highly suitable for the generation of recombinant therapeutic glycoproteins. Most importantly, PS degradation and hydrolytic activity were drastically reduced, providing an avenue toward future PS degradation-free biologics-manufacturing processes. Enzymatic degradation of polysorbates (PS) in biologic drug formulations is often caused by hydrolytic host cell proteins (HCPs) and can lead to particle formation and reduced shelf-life. Here, we present a host cell line-engineering approach by removing nine Chinese hamster ovary (CHO) host cell hydrolases, which have previously been confirmed to degrade PS. Strikingly, the sequential genomic knockout (KO) of these hydrolases, including the genetic removal of two entire gene clusters of unprecedented size, yielded viable CHO host cell line variants. This novel host cell line was further optimized using the additional KO of the two key proapoptotic genes, Bax and Bak1. Thus, we generated a competitive multi-hydrolase KO CHO host cell line that was further shown to be highly suitable for the generation of recombinant therapeutic glycoproteins. Most importantly, PS degradation and hydrolytic activity were drastically reduced, providing an avenue toward future PS degradation-free biologics-manufacturing processes. [Display omitted] In the opinion of the authors, the presented technology of knocking out multiple host cell hydrolases from the Chinese hamster ovary (CHO) host cell genome to reduce polysorbate (PS) degradation is ready for implementation. We proved key considerable aspects when generating a novel CHO host cell line, such as transfectability, metabolic selection, ability to endure a conventional fed-batch process without compromising productivity, and consistent product quality attributes compared with the parental host cell line. Given that we performed cultivations in small-scale bioreactors mimicking the real-world process, we suggest a technology readiness level (TRL) of 5 out of 9 (based on NASA’s TRL system). In terms of the recently published Biomanufacturing Readiness Level (BRL) developed by the National Institute for Innovation in Manufacturing Biopharmaceuticals (NIIMBL), we would classify our technology at BLR 4 out of 9, as we have performed initial tests with industry-relevant feedstocks. Nevertheless, full-scale implementation might harbor further implications. In contrast to the parental CHO host cell line, detailed characterization data on the multi-hydrolase KO host cell line are not available. To ensure its seamless application in an industrial bioprocess, further characterization experiments should test media fit, long-term stability of clonal production cell lines, suitability to express a variety of biologics (including complex antibody formats), and capability for large-scale bioreactor cultivations. In addition, we can only predict that the presented reduction of hydrolytic activity translates all the way down to the final drug product. A complete bioprocess including the final formulation steps followed by stability studies will ultimately demonstrate whether our technology has the ability to solve the PS degradation challenge entirely. Enzymatic degradation of polysorbates (PS) in biologic formulations is often caused by hydrolytic host cell proteins (HCPs) and can lead to particle formation and reduced shelf-life. Combining hydrolase knockouts in a single Chinese hamster ovary (CHO) host cell line enabled traceless enzyme removal and represented a big step toward the production of PS degradation-free biologics.