Gene Editing Protocols Research Articles

IDH1 R132H and TP53 R248Q Mutations Modulate Glioma Cell Migration and Adhesion on Different ECM Components

Mutations in IDH1 and TP53 have a significant impact on glioma prognosis and progression; however, their roles in tumor cell invasion in terms of interactions with particular components of the extracellular matrix (ECM) are still unclear. Using gene editing protocol based on CRISPR-Cas 9 with cytidine deaminase, we introduced point mutations into U87MG glioblastoma cells to establish modified cell lines with heterozygous IDH1 R132H, homozygous TP53 R248Q and heterozygous IDH1 R132H, homozygous TP53 R248Q genotypes. A comparative study of cell migration on major ECM components was carried out by high-content microscopy. IDH1 R132H mutation introduced to U87MG glioblastoma cells was shown to decrease the migration speed on Matrigel and collagen IV substrates compared to the wild-type. This data were supported by cell adhesion quantification via the lateral shift assay performed by atomic force microscopy (AFM). TP53 R248Q mutation increased cell adhesion to various substrates and significantly promoted cell migration on hyaluronic acid and chondroitin sulfate but did not change the migration rates on laminin and collagens IV and I. A double-mutant genotype produced by consequently introducing IDH1 R132H and TP53 R248Q to parental glioblastoma cells was characterized by the highest migration among all the cell lines, with particularly faster motility on chondroitin sulfate. These findings underscore the complex interactions between glioma cells, with the most important driver mutations and specific ECM components regulating cancer cell migration, offering valuable insights for potential therapeutic targets in glioma treatment.

Efficient CRISPR/Cas9-mediated genome editing in the European corn borer, Ostrinia nubilalis.

The European corn borer (Ostrinia nubilalis) is an agricultural pest and burgeoning model for research on speciation, seasonal adaptation and insect resistance management. Although previous work in O. nubilalis has identified genes associated with differences in life cycle, reproduction, and resistance to Bt toxins, the general lack of a robust gene-editing protocol for O. nubilalis has been a barrier to functional validation of candidate genes. Here, we demonstrate an efficient and practical methodology for heritable gene mutagenesis in O. nubilalis using the CRISPR/Cas9 genome editing system. Precise loss-of-function (LOF) mutations were generated at two circadian clock genes, period (per) and pigment-dispersing factor receptor (pdfr), and a developmental gene, prothoracicotropic hormone (ptth). Precluding the need for a visible genetic marker, gene-editing efficiency remained high across different single guide RNAs (sgRNA) and germline transmission of mutations to F1 offspring approached 100%. When single or dual sgRNAs were injected at a high concentration, gene-specific phenotypic differences in behaviour and development were identified in F0 mutants. Specifically, F0 gene mutants demonstrated that PER, but not PDFR, is essential for normal timing of eclosion. PTTH F0 mutants were significantly heavier and exhibited a higher incidence of diapause. This work will accelerate future studies of gene function in O. nubilalis and facilitate the development of similar screens in other Lepidopteran and non-model insects.

The Evolutionary Biology of Chelicerata.

Chelicerata constitutes an ancient, biodiverse, and ecologically significant group of Arthropoda. The study of chelicerate evolution has undergone a renaissance in the past decade, resulting in major changes to our understanding of the higher-level phylogeny and internal relationships of living orders. Included among these conceptual advances are the discoveries of multiple whole-genome duplication events in a subset of chelicerate orders, such as horseshoe crabs, spiders, and scorpions. As a result, longstanding hypotheses and textbook scenarios of chelicerate evolution, such as the monophyly of Arachnida and a single colonization of land by the common ancestor of arachnids, have come into contention. The retention of ancient, duplicated genes across this lineage also offers fertile ground for investigating the role of gene duplication in chelicerate macroevolution. This new frontier of investigation is paralleled by the timely establishment of the first gene editing protocols for arachnid models, facilitating a new generation of experimental approaches.

The cytochrome P450 subfamilies CYP392A and CYP392D are key players in acaricide metabolism in Tetranychusurticae

The spider mite Tetranychus urticae is a major agricultural pest with a global distribution, extremely diverse host range and a remarkable ability to develop resistance to a wide variety of acaricides. P450 mono‐oxygenases have been frequently associated with resistance development in this species. In particular enzymes of the CYP392A-subfamily were shown to metabolize a number of key acaricides, including abamectin, amitraz, fenpyroximate and the active metabolite of pyflubumide. However, transcriptomic studies comparing highly resistant and susceptible populations have often revealed high expression of members of the CYP392D-subfamily, but these have been only poorly studied. Here, we conducted a meta-analysis of gene expression data of 20 populations and identified two key enzymes of this family, CYP392D2 and CYP392D8, whose expression is associated with resistance. We subsequently functionally expressed these enzymes, together with CYP392A11 and CYP392A16 as known metabolizers, and compared their potential to accept a wide diversity of acaricides as substrate. This study overall confirms previous discovered substrates for CYP392A11 and CYP392A16, but also reveals unreported metabolic activity towards new acaricides. These include carbaryl, chlorpyrifos and etoxazole for CYP392A16 and carbaryl, chlorpyrifos and NNI-0711-NH pyflubumide for CYP392A11. For the newly studied CYP392D-family, we show that CYP392D2 metabolizes pyridaben, fenpyroximate, etoxazole and chlorpyrifos, while CYP392D8 metabolizes carbaryl, fenazaquin and tebufenpyrad. Last, we observed that both CYP392A- and CYP392D-subfamily enzymes activate chlorpyrifos to its corresponding oxon. Our study indicates that there is both overlap and specificity in the activity of A- and D-subfamily enzymes against acaricides and model substrates. With the recent advent of highly efficient CRISPR/Cas9 gene editing protocols in T. urticae, the way is now paved to conduct further genetic experiments revealing and quantifying the role of these enzymes in the resistance phenotype in field populations.

Tackling mosaicism in gene edited livestock

The farming of livestock has a critical role to play in global nutritional security and poverty alleviation. To meet these goals through more efficient, environmentally sustainable and animal welfare focused means, gene editing technologies could be integrated into current breeding programs. A common issue with gene editing in livestock zygotes is the high incidence of genetic mosaicism. Genetic mosaicism, characterised by a single individual carrying distinct genotypes in different cell lineages, can lead to inconsistent presentation of a desired trait phenotypically, or the absence of the intended genotype in the animal’s germline. This review explores the present status of genetic mosaicism associated with CRISPR-Cas9 gene editing in cattle, sheep, and pigs, and identifies four areas for refinement; (1) the type of CRISPR-Cas9 genome editor used; (2) the CRISPR-Cas9 formats and timing of gene editing during embryonic development; (3) the method used to deliver the genome editor and (4) the genetic screening strategies applied. We also discuss alternatives to direct zygote gene editing, including surrogate sire technology and blastomere separation, which circumvent the production of mosaic offspring. By exploring these avenues for reducing mosaicism, gene editing protocols in livestock could become more efficient and effective, which will ultimately pave the way for traits to be introduced that improve animal welfare standards and help address gaps in the security of global nutrition access.

Ex vivo gene editing and cell therapy for hereditary tyrosinemia type 1.

We previously demonstrated the successful use of in vivo CRISPR gene editing to delete 4-hydroxyphenylpyruvate dioxygenase (HPD) to rescue mice deficient in fumarylacetoacetate hydrolase (FAH), a disorder known as hereditary tyrosinemia type 1 (HT1). The aim of this study was to develop an ex vivo gene-editing protocol and apply it as a cell therapy for HT1. We isolated hepatocytes from wild-type (C57BL/6J) and Fah-/- mice and then used an optimized electroporation protocol to deliver Hpd-targeting CRISPR-Cas9 ribonucleoproteins into hepatocytes. Next, hepatocytes were transiently incubated in cytokine recovery media formulated to block apoptosis, followed by splenic injection into recipient Fah-/- mice. We observed robust engraftment and expansion of transplanted gene-edited hepatocytes from wild-type donors in the livers of recipient mice when transient incubation with our cytokine recovery media was used after electroporation and negligible engraftment without the media (mean: 46.8% and 0.83%, respectively; p=0.0025). Thus, the cytokine recovery medium was critical to our electroporation protocol. When hepatocytes from Fah-/- mice were used as donors for transplantation, we observed 35% and 28% engraftment for Hpd-Cas9 ribonucleoproteins and Cas9 mRNA, respectively. Tyrosine, phenylalanine, and biochemical markers of liver injury normalized in both Hpd-targeting Cas9 ribonucleoprotein and mRNA groups independent of induced inhibition of Hpd through nitisinone, indicating correction of disease indicators in Fah-/- mice. The successful liver cell therapy for HT1 validates our protocol and, despite the known growth advantage of HT1, showcases ex vivo gene editing using electroporation in combination with liver cell therapy to cure a disease model. These advancements underscore the potential impacts of electroporation combined with transplantation as a cell therapy.

Gene Editing in the Chagas Disease Vector Rhodnius prolixus by Cas9-Mediated ReMOT Control.

Rhodnius prolixus is currently the model vector of choice for studying Chagas disease transmission, a debilitating disease caused by Trypanosoma cruzi parasites. However, transgenesis and gene editing protocols to advance the field are still lacking. Here, we tested protocols for the maternal delivery of CRISPR-Cas9 (clustered regularly spaced palindromic repeats/Cas-9 associated) elements to developing R. prolixus oocytes and strategies for the identification of insertions and deletions (indels) in target loci of resulting gene-edited generation zero (G0) nymphs. We demonstrate successful gene editing of the eye color markers Rp-scarlet and Rp-white, and the cuticle color marker Rp-yellow, with highest effectiveness obtained using Receptor-Mediated Ovary Transduction of Cargo (ReMOT Control) with the ovary-targeting BtKV ligand. These results provide proof of concepts for generating somatic mutations in R. prolixus and potentially for generating germ line-edited lines in triatomines, laying the foundation for gene editing protocols that could lead to the development of novel control strategies for vectors of Chagas disease.

Hematopoietic stem cell gene editing rescues B-cell development in X-linked agammaglobulinemia

X-linked agammaglobulinemia (XLA) is an inborn error of immunity that renders boys susceptible to life-threatening infections due to loss of mature B cells and circulating immunoglobulins. It is caused by defects in the gene encoding the Bruton tyrosine kinase (BTK) that mediates the maturation of B cells in the bone marrow and their activation in the periphery. This paper reports on a gene editing protocol to achieve "knock-in" of a therapeutic BTK cassette in hematopoietic stem and progenitor cells (HSPCs) as a treatment for XLA. To rescue BTK expression, this study employed a clustered regularly interspaced short palindromic repeats/CRISPR-associated protein 9 system that creates a DNA double-strand break in an early exon of the BTK locus and an adeno-associated virus 6 virus that carries the donor template for homology-directed repair. The investigators evaluated the efficacy of the gene editing approach in HSPCs from patients with XLA that were cultured invitro under B-cell differentiation conditions or that were transplanted in immunodeficient mice to study B-cell output invivo. A (feeder-free) B-cell differentiation protocol was successfully applied to blood-mobilized HSPCs to reproduce invitro the defects in B-cell maturation observed in patients with XLA. Using this system, the investigators could show the rescue of B-cell maturation by gene editing. Transplantation of edited XLA HSPCs into immunodeficient mice led to restoration of the human B-cell lineage compartment in the bone marrow and immunoglobulin production in the periphery. Gene editing efficiencies above 30% could be consistently achieved in human HSPCs. Given the potential selective advantage of corrected cells, as suggested by skewed X-linked inactivation in carrier females and by competitive repopulating experiments in mouse models, this work demonstrates the potential of this strategy as a future definitive therapy for XLA.

Oocyte electroporation prior to in vitro fertilization is an efficient method to generate single, double, and multiple knockout porcine embryos of interest in biomedicine and animal production

Genetically modified pigs play a critical role in mimicking human diseases, xenotransplantation, and the development of pigs resistant to viral diseases. The use of programmable endonucleases, including the CRISPR/Cas9 system, has revolutionized the generation of genetically modified pigs. This study evaluates the efficiency of electroporation of oocytes prior to fertilization in generating edited gene embryos for different models. For single gene editing, phospholipase C zeta (PLC ζ) and fused in sarcoma (FUS) genes were used, and the concentration of sgRNA and Cas9 complexes was optimized. The results showed that increasing the concentration resulted in higher mutation rates without affecting the blastocyst rate. Electroporation produced double knockouts for the TPC1/TPC2 genes with high efficiency (79 %). In addition, resistance to viral diseases such as PRRS and swine influenza was achieved by electroporation, allowing the generation of double knockout embryo pigs (63 %). The study also demonstrated the potential for multiple gene editing in a single step using electroporation, which is relevant for xenotransplantation. The technique resulted in the simultaneous mutation of 5 genes (GGTA1, B4GALNT2, pseudo B4GALNT2, CMAH and GHR). Overall, electroporation proved to be an efficient and versatile method to generate genetically modified embryonic pigs, offering significant advances in biomedical and agricultural research, xenotransplantation, and disease resistance. Electroporation led to the processing of numerous oocytes in a single session using less expensive equipment. We confirmed the generation of gene-edited porcine embryos for single, double, or quintuple genes simultaneously without altering embryo development to the blastocyst stage. The results provide valuable insights into the optimization of gene editing protocols for different models, opening new avenues for research and applications in this field.

Efficient Gene-Editing in Human Pluripotent Stem Cells Through Simplified Assembly of Adeno-Associated Viral (AAV) Donor Templates.

Gene-edited human pluripotent stem cells provide attractive model systems to functionally interrogate the role of specific genetic variants in relevant cell types. However, the need to isolate and screen edited clones often remains a bottleneck, in particular when recombination rates are sub-optimal. Here, we present a protocol for flexible gene editing combining Cas9 ribonucleoprotein with donor templates delivered by adeno-associated virus (AAV) vectors to yield high rates of homologous recombination. To streamline the workflow, we designed a modular system for one-step assembly of targeting vectors based on Golden Gate cloning and developed a rapid protocol for small-scale isolation of AAV virions of serotype DJ. High homology-directed repair (HDR) rates in human pluripotent stem cells (hPSCs), ~70% in ACTB and ~30% in LMNB1, were achieved using this approach, also with short (300 bp) homology arms. The modular design of donor templates is flexible and allows for the generation of conditional and/or complex alleles. This protocol thus provides a flexible and efficient strategy workflow to rapidly generate gene-edited hPSC lines. Key features • Versatile approach combining AAV-DJ donors and CRISPR ribonucleoproteins, providing an efficient method for long and short edits, insertions, and deletions in human pluripotent stem cells. • One-step cloning method for rapid generation of customized AAV donor plasmids. • Simplified AAV purification pipeline for ready-to-infect virion preparations.

Autologous MPO Knock-out Hematopoietic Stem and Progenitor Cells for Pulmonary Arterial Hypertension

Pulmonary Arterial Hypertension (PAH) is a progressive condition for which there is no cure. Even with substantial pharmacologic advances in the modern treatment era, existing treatments provide only symptomatic relief and survival remains unacceptably poor. Patients who fail to respond to medications usually have the worst prognosis and patients with persistently elevated pulmonary pressure and right heart failure usually die within 5 years. PAH is a severe complication of Systemic sclerosis (SSc) affecting approximately 8%-12% of patients. SSc affects 13.5-44.3 per 100,000 individuals and is complex and diverse, with limited treatment options. SSc-associated PAH is the second largest group of patients with PAH. Once diagnosed, it becomes the leading cause of mortality among this patient population, with a 50% mortality rate within 3 years of PAH diagnosis. Inflammation and endothelial injury being the main components of the syndrome, investigation regarding the underlying mechanisms of this immune response may lead to the development of a new efficient therapy. There is extensive evidence for a pathogenic role for neutrophil myeloperoxidase (MPO) in PAH disease. PAH patients with higher levels of MPO have a poorer outcome. Studies in murine models of PAH demonstrate significant disease amelioration by elimination of neutrophil MPO by hematopoietic stem cell (HSC) transplantation with HSC lacking MPO from germline MPO gene knock-out. We developed a gene therapy approach to PAH in which autologous HSCs will undergo gene editing using CRISPR/Cas9 to achieve bi-allelic disruption of the MPO gene. These MPO-knock-out HSCs will then be autologous transplanted. That autologous transplantation of HSPC in which the MPO gene is disrupted is expected to prevent progression of PAH due to the essential role of neutrophil MPO activity on disease development. We identified a sgRNA that leads to high frequency disruption of MPO in primary human CD34+ hematopoietic stem and progenitor cells (HSPC), leading to efficient elimination of MPO protein in the resultant neutrophils. That sequence was assessed for genome-wide off-target and showed no detectable locus, validating it as a lead candidate for safe gene editing strategy. We validated in an in vitro neutrophil differentiation model the functional efficiency of the knock-out. MPO KO differentiated cells exhibit reduced neutrophil extracellular traps formation (NETosis) and absence of MPO protein. An optimized gene editing protocol for HSC was optimized and showed consistent allelic disruption over 80%. The edited cells retain hematopoietic potential and showed no skewing in the generated lineages in an in vitro colony forming assay. We developed a surrogate gene editing protocol for murine cells and optimized electroporation to achieve high allelic disruption of the murine MPO gene in lineage negative murine hematopoietic cells. The protocol will be tested in a hypoxia-induced PAH murine model. The presented work can represent a major improvement for PAH patient. The limited therapeutic options and severity of the disease urges to develop new approaches. The gene therapy protocol developed showed promising result in vitro and rely on clinical evidences for the implication of MPO in the disease. Results will be tested in a pre-clinical murine model

CRISPR-Cas9 editing of TLR4 to improve the outcome of cardiac cell therapy

Inflammation and fibrosis limit the reparative properties of human mesenchymal stromal cells (hMSCs). We hypothesized that disrupting the toll-like receptor 4 (TLR4) gene would switch hMSCs toward a reparative phenotype and improve the outcome of cell therapy for infarct repair. We developed and optimized an improved electroporation protocol for CRISPR-Cas9 gene editing. This protocol achieved a 68% success rate when applied to isolated hMSCs from the heart and epicardial fat of patients with ischemic heart disease. While cell editing lowered TLR4 expression in hMSCs, it did not affect classical markers of hMSCs, proliferation, and migration rate. Protein mass spectrometry analysis revealed that edited cells secreted fewer proteins involved in inflammation. Analysis of biological processes revealed that TLR4 editing reduced processes linked to inflammation and extracellular organization. Furthermore, edited cells expressed less NF-ƙB and secreted lower amounts of extracellular vesicles and pro-inflammatory and pro-fibrotic cytokines than unedited hMSCs. Cell therapy with both edited and unedited hMSCs improved survival, left ventricular remodeling, and cardiac function after myocardial infarction (MI) in mice. Postmortem histologic analysis revealed clusters of edited cells that survived in the scar tissue 28 days after MI. Morphometric analysis showed that implantation of edited cells increased the area of myocardial islands in the scar tissue, reduced the occurrence of transmural scar, increased scar thickness, and decreased expansion index. We show, for the first time, that CRISPR-Cas9-based disruption of the TLR4-gene reduces pro-inflammatory polarization of hMSCs and improves infarct healing and remodeling in mice. Our results provide a new approach to improving the outcomes of cell therapy for cardiovascular diseases.

Therapeutic Gene Editing of HSCs Ex Vivo without in Vitro Culture Avoids Genotoxicity, Simplifies Procedures, and Preserves Efficiency and Stemness

Therapeutic gene editing of HSCs promises to cure inherited blood disorders. However, current ex vivo gene editing protocols may be associated with genotoxic potential and complex in vitro cell manufacturing procedures. Base and prime editing reduce double strand breaks (DSBs) but are generally less efficient and might have unique genotoxic potential (e.g. untargeted deaminase activity). Here we test the hypothesis that therapeutic gene editing with targeted endonucleases disrupting HbF-repressing elements for b-hemoglobinopathies may be made safer, simpler, and at least as effective by focusing the editing on nondividing cells. Genotoxic risks of DSBs appear to be concentrated in cycling cells. For example, Cas9-induced DSBs can result in long deletions at the on-target site that appear to be mainly the result of unidirectional extensive resection selectively active in S/G2 cell cycle phases1. In addition, DSBs must be resolved prior to mitosis to prevent chromosome mis-segregation leading to copy number variation and cytoplasmic micronuclei2,3. We measured the cell cycle state of mobilized PB CD34+ HSPCs immediately after collection, or after 24 or 48 hours of culture with cytokines (SCF, TPO, and FLT3L each at 100 ng/ml). 100% of cells were in G0/G1 at 0 hours, 94.6%±1.5% at 24 hours, and 76.7%±2.6% at 48 hours. We electroporated CD34+ HSPCs with 0, 24, or 48 hours of cytokine culture before performing 3xNLS-HiFi-SpCas9:sgRNA ribonucleoprotein (RNP) electroporation targeting the BCL11A erythroid enhancer. We employed a flow cytometry assay to quantify micronuclei in edited HSPCs. The fraction of micronuclei was increased in cells electroporated with Cas9:sgRNA RNP following 24 or 48 hours of cytokine stimulation but not in unstimulated cells. We performed ddPCR on sorted nuclei and micronuclei using assays specific to distal chromosome 2p (telomeric to the BCL11A cleavage site) and to chr1 as a reference autosome. We found ~9-fold enrichment of chr2p in micronuclei after 24 or 48 hours of pre-stimulation cytokine culture prior to gene editing, but strikingly did not observe any enrichment of chr2p in micronuclei in cells edited without pre-stimulation culture, suggesting that chromosome mis-segregation and unwanted downstream consequences could be avoided by editing quiescent HSCs (Fig. 1A). In addition, we developed ddPCR assays to measure genome editing outcomes after editing the BCL11A enhancer with paired cleavages (at the +58 and +55 enhancers) separated by 3-kb. These assays measure: 1) programmed 3-kb deletions, 2) programmed 3-kb inversions, or 3) short indels or unedited alleles. Any missing signals (genomic copies fewer than found on a reference nontargeted autosome) may represent long deletions or rearrangements. Editing cells without pre-stimulation culture was associated with undetectable missing signals (inferred long deletions and rearrangements) as compared to editing after 24 or 48 hours of pre-stimulation culture. We compared our standard protocol of 24 hours of pre-stimulation cytokine culture and 24 hours of post-electroporation cytokine culture to a protocol in which CD34+ HSPCs were electroporated immediately after thawing and then cryopreserved 15 minutes after electroporation, without any cytokine culture, before infusion to NBSGW mice. After 16 weeks, HbF levels were potently induced to similar levels after gene editing in recipients of HSPCs without and with cytokine culture (Fig. 1B). Overall human engraftment was similar for all groups, as was multilineage repopulation. Gene edit frequencies were similar for recipients of HSPCs without and with cytokine culture. We observed comparable editing and engraftment potency of HSPCs without as compared to with cytokine culture across mobilization type (G-CSF or plerixafor), donor disease status (healthy and SCD), and sgRNA target sequence. Together, these results suggest that editing HSPCs ex vivo without in vitro culture: avoids unintended on-target genotoxicity potential, including production of micronuclei, long deletions, and rearrangements; simplifies protocols, eliminating time, cost, and complexity; and preserves gene editing efficiency and HbF induction and HSC repopulation potency. Editing nondividing cells selectively could be a straightforward modification to improve ex vivo editing as well as to advance future efforts for safe and potent in vivo editing of HSCs. Figure 1View largeDownload PPTFigure 1View largeDownload PPT Close modal

HideRNAs protect against CRISPR-Cas9 re-cutting after successful single base-pair gene editing

Promiscuous activity of the Streptococcus pyogenes DNA nuclease CRISPR-Cas9 can result in destruction of a successfully modified sequence obtained by templated repair of a Cas9-induced DNA double-strand break. To avoid re-cutting, additional target-site-disruptions (TSDs) are often introduced on top of the desired base-pair alteration in order to suppress target recognition. These TSDs may lower the efficiency of introducing the intended mutation and can cause unexpected phenotypes. Alternatively, successfully edited sites can be protected against Cas9 re-cutting activity. This method exploits the finding that Cas9 complexed to trimmed guideRNAs can still tightly bind specific genomic sequences but lacks nuclease activity. We show here that the presence of a guideRNA plus a trimmed guideRNA that matches the successfully mutated sequence, which we call hideRNA, can enhance the recovery of precise single base-pair substitution events tenfold. The benefit of hideRNAs in generating a single point mutation was demonstrated in cell lines using plasmid-based delivery of CRISPR-Cas9 components and in mouse zygotes injected with Cas9/guideRNA plus Cas9/hideRNA ribonucleoprotein complexes. However, hRNA protection sometimes failed, which likely reflects an unfavorable affinity of hRNA/Cas9 versus gRNA/Cas9 for the DNA target site. HideRNAs can easily be implemented into current gene editing protocols and facilitate the recovery of single base-pair substitution. As such, hideRNAs are of great value in gene editing experiments demanding high accuracy.

Rapid and efficient generation of antigen-specific isogenic T cells from cryopreserved blood samples.

Clustered regularly interspaced short palindromic repeats/CRISPR‐associated protein 9 (CRISPR/Cas9)‐mediated gene editing has been leveraged for the modification of human and mouse T cells. However, limited experience is available on the application of CRISPR/Cas9 electroporation in cryopreserved T cells collected during clinical trials. To address this, we aimed to optimize a CRISPR/Cas9‐mediated gene editing protocol compatible with peripheral blood mononuclear cells (PBMCs) samples routinely produced during clinical trials. PBMCs from healthy donors were used to generate knockout T‐cell models for interferon‐γ, Cbl proto‐oncogene B (CBLB), Fas cell surface death receptor (Fas) and T‐cell receptor (TCRαβ) genes. The effect of CRISPR/Cas9‐mediated gene editing on T cells was evaluated using apoptosis assays, cytokine bead arrays and ex vivo and in vitro stimulation assays. Our results demonstrate that CRISPR/Cas9‐mediated gene editing of ex vivo T cells is efficient and does not overtly affect T‐cell viability. Cytokine release and T‐cell proliferation were not affected in gene‐edited T cells. Interestingly, memory T cells were more susceptible to CRISPR/Cas9 gene editing than naïve T cells. Ex vivo and in vitro stimulation with antigens resulted in equivalent antigen‐specific T‐cell responses in gene‐edited and untouched control cells, making CRISPR/Cas9‐mediated gene editing compatible with clinical antigen‐specific T‐cell activation and expansion assays. Here, we report an optimized protocol for rapid, viable and highly efficient genetic modification in ex vivo human antigen‐specific T cells, for subsequent functional evaluation and/or expansion. Our platform extends CRISPR/Cas9‐mediated gene editing for use in gold‐standard clinically used immune‐monitoring pipelines and serves as a starting point for development of analogous approaches, such as those including transcriptional activators and/or epigenetic modifiers.

Gene Editing in Green Alga Chlamydomonas reinhardtii via CRISPR-Cas9 Ribonucleoproteins.

With the establishment of the CRISPR-Cas9 molecular tool as a DNA editing system in 2012, the handling of gene editing experiments was strongly facilitated pushing reverse genetics approaches forward in many organisms. These new gene editing technologies also drastically increased the possibilities for design-driven synthetic biology. Here, we describe a protocol for gene editing in the green algae Chlamydomonas reinhardtii using preassembled CRISPR-Cas9 ribonucleoproteins.The three sections of the protocol guide through a complete gene editing experiment, starting with the experimental design and the choice of suitable CRISPR target sites and how to perform a Cas9 in vitro test digestion. The second part covers the transformation of algal cells with Cas9 RNPs using electroporation. In the last part, the PCR-based screening for mutants and isolation of clones is explained.

Chemical proteomics for identifying short-chain fatty acid modified proteins in Salmonella.

Microbiota-metabolized small molecules play important roles to regulate host immunity and pathogen virulence. Specifically, microbiota generates millimolar concentration of short-chain fatty acid (SCFA) that can directly inhibit Salmonella virulence. Here, we describe chemical proteomic methods to identify SCFA-modified proteins in Salmonella using free fatty acids as well as their salicylic acid derivatives. In addition, we include CRISPR-Cas9 gene editing protocols for epitope-tagging of specific proteins to validate SCFA-modification in Salmonella. These protocols should facilitate the discovery and functional analysis of SCFA-modified proteins in Salmonella microbiology and pathogenesis.

CRISPR/Cas9-mediated generation of biallelic F0 anemonefish (Amphiprion ocellaris) mutants.

Genomic manipulation is a useful approach for elucidating the molecular pathways underlying aspects of development, physiology, and behaviour. However, a lack of gene-editing tools appropriated for use in reef fishes has meant the genetic underpinnings for many of their unique traits remain to be investigated. One iconic group of reef fishes ideal for applying this technique are anemonefishes (Amphiprioninae) as they are widely studied for their symbiosis with anemones, sequential hermaphroditism, complex social hierarchies, skin pattern development, and vision, and are raised relatively easily in aquaria. In this study, we developed a gene-editing protocol for applying the CRISPR/Cas9 system in the false clown anemonefish, Amphiprion ocellaris. Microinjection of zygotes was used to demonstrate the successful use of our CRISPR/Cas9 approach at two separate target sites: the rhodopsin-like 2B opsin encoding gene (RH2B) involved in vision, and Tyrosinase-producing gene (tyr) involved in the production of melanin. Analysis of the sequenced target gene regions in A. ocellaris embryos showed that uptake was as high as 73.3% of injected embryos. Further analysis of the subcloned mutant gene sequences combined with amplicon shotgun sequencing revealed that our approach had a 75% to 100% efficiency in producing biallelic mutations in F0 A. ocellaris embryos. Moreover, we clearly show a loss-of-function in tyr mutant embryos which exhibited typical hypomelanistic phenotypes. This protocol is intended as a useful starting point to further explore the potential application of CRISPR/Cas9 in A. ocellaris, as a platform for studying gene function in anemonefishes and other reef fishes.

A Robust Protocol for CRISPR-Cas9 Gene Editing in Human Suspension Cell Lines.

The implementation of clustered regularly interspaced short palindromic repeats (CRISPR)-Cas9 systems in mammalian cells has sparked an exciting new era in targeted gene editing. CRISPR-Cas9 technologies allow gene function to be interrogated by gene deletions, mutations, and truncations, and by epitope tagging and promoter activity modulation. Many robust protocols have been published to date on CRISPR-Cas9 techniques, however, most of these focus on adherent cell lines. Suspension cell lines, typically of hematolymphoid origin, such as Jurkat, Daudi, and TOLEDO, pose unique challenges to the setup of CRISPR experiments. Here, using B cell lymphoma cells as a primary model, we describe a comprehensive protocol for targeted gene manipulations using the CRISPR-Cas9 system in suspension cells. We also highlight necessary optimization steps to make this approach universal to other suspension cell lines. We first describe a detailed protocol for transient expression of the Cas9 nuclease and guide RNAs. We then suggest workflows for obtaining single-cell clones and for screening for successful homozygous knockout (KO) clones in suspension lines. This protocol aims to serve as a comprehensive resource to facilitate gene editing experiments for users starting CRISPR-Cas9 gene editing protocols on suspension cell lines or those looking to optimize their current workflows. © 2021 Wiley Periodicals LLC. Basic Protocol 1: Transient CRISPR Cas9-gRNA delivery for gene knockout by NeonTM electroporation Support Protocol 1: Designing and preparing gene-specific gRNA Support Protocol 2: Preparation of conditioned medium and culture vessels for single-cell FACS Alternate Protocol 1: Transient CRISPR Cas9-gRNA delivery for gene knockout by Nepagene electroporator Basic Protocol 2: FACS and single-cell clone generation Alternate Protocol 2: Manual cell dilution to obtain single-cell clones Basic Protocol 3: Confirming indels status in single-cell clones by PCR on genomic DNA and Sanger sequencing.

Duchenne muscular dystrophy cell culture models created by CRISPR/Cas9 gene editing and their application in drug screening\xa0

Gene editing methods are an attractive therapeutic option for Duchenne muscular dystrophy, and they have an immediate application in the generation of research models. To generate myoblast cultures that could be useful in in vitro drug screening, we have optimised a CRISPR/Cas9 gene edition protocol. We have successfully used it in wild type immortalised myoblasts to delete exon 52 of the dystrophin gene, modelling a common Duchenne muscular dystrophy mutation; and in patient’s immortalised cultures we have deleted an inhibitory microRNA target region of the utrophin UTR, leading to utrophin upregulation. We have characterised these cultures by demonstrating, respectively, inhibition of dystrophin expression and overexpression of utrophin, and evaluating the expression of myogenic factors (Myf5 and MyH3) and components of the dystrophin associated glycoprotein complex (α-sarcoglycan and β-dystroglycan). To demonstrate their use in the assessment of DMD treatments, we have performed exon skipping on the DMDΔ52-Model and have used the unedited DMD cultures/ DMD-UTRN-Model combo to assess utrophin overexpression after drug treatment. While the practical use of DMDΔ52-Model is limited to the validation to our gene editing protocol, DMD-UTRN-Model presents a possible therapeutic gene edition target as well as a useful positive control in the screening of utrophin overexpression drugs.