Homology-directed Repair Template Research Articles

Peptide-enabled ribonucleoprotein delivery for CRISPR engineering (PERC) in primary human immune cells and hematopoietic stem cells.

Peptide-enabled ribonucleoprotein delivery for CRISPR engineering (PERC) is a new approach for ex vivo genome editing of primary human cells. PERC uses a single amphiphilic peptide reagent to mediate intracellular delivery of the same pre-formed CRISPR ribonucleoprotein enzymes that are broadly used in research and therapeutics, resulting in high-efficiency editing of stimulated immune cells and cultured hematopoietic stem and progenitor cells (HSPCs). PERC facilitates nuclease-mediated gene knockout, precise transgene knock-in, and base editing. PERC involves mixing the CRISPR ribonucleoprotein enzyme with peptide and then incubating the formulation with cultured cells. For efficient transgene knock-in, adeno-associated virus (AAV) bearing homology-directed repair template DNA may be included. In contrast to electroporation, PERC is appealing as it requires no dedicated hardware and has less impact on cell phenotype and viability. Due to the gentle nature of PERC, delivery can be performed multiple times without substantial impact to cell health or phenotype. Here we report methods for improved PERC-mediated editing of T cells as well as novel methods for PERC-mediated editing of HSPCs, including knockout and precise knock-in. Editing efficiencies can surpass 90% using either Cas9 or Cas12a in primary T cells or HSPCs. Because PERC calls for only three readily available reagents - protein, RNA, and peptide - and does not require dedicated hardware for any step, PERC demands no special expertise and is exceptionally straightforward to adopt. The inherent compatibility of PERC with established cell engineering pipelines makes this approach appealing for rapid deployment in research and clinical settings.

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.

Recombinant Production of Glycoengineered Mucins in HEK293-F Cells.

Recombinant mucins are attractive polymeric building blocks for new biomaterials, biolubricants, and therapeutics. Advances in glycoengineered host cell systems now enable the recombinant production of mucins with tailored O-glycan side chains, offering new opportunities to tune the functionality of mucins and investigate the biology of specific O-glycan structures. Here, we provide a protocol for the scalable production of glycoengineered mucins and mucin-like glycoproteins in suspension-adapted HEK293-F cells. The protocol includes the preparation of engineered cell lines with homozygous knockout (KO) of glycosyltransferases using CRISPR/Cas9 and homology-directed repair (HDR) templates designed for efficient screening of clones. Strategies are provided for the stable introduction of mucin expression cassettes into the HEK293-F genome and the subsequent isolation of high-expressing cell populations. The high-titer production of recombinant mucins in conventional shaker flasks is described as an example production strategy using these cell lines.

CRISPR-Cas9-Mediated Bioluminescent Tagging of Endogenous Proteins by Fluorescent Protein-Assisted Cell Sorting.

Oncogenic fusion genes are attractive therapeutic targets because of their tumor-specific expression and central "driver" roles in various human cancers. However, oncogenic fusions involving transcription factors such as PAX3-FOXO1 in alveolar fusion gene-positive rhabdomyosarcoma (FP-RMS) have been difficult to inhibit due to the apparent lack of tractable drug-like binding sites comparable to that recognized by Gleevec (imatinib mesylate) on the BCR-ABL1 tyrosine kinase fusion protein. Toward the identification of novel small molecules that selectively target PAX3-FOXO1, we used CRISPR-Cas9-mediated knock-in to append the pro-luminescent HiBiT tag onto the carboxy terminus of the endogenous PAX3-FOXO1 fusion protein in two human FP-RMS cell lines (RH4 and SCMC). HiBiT is an 11-amino acid peptide derived from the NanoLuc luciferase that produces a luminescence signal which is ~100-fold brighter than firefly or Renilla luciferases through high-affinity binding to a complementary NanoLuc peptide fragment called LgBiT. To facilitate single-cell clonal isolation of knock-ins, the homology-directed repair template encoding HiBiT was followed by a P2A self-cleaving peptide for coexpression of an mCherry fluorescent protein as a fluorescence-activated cell sorter (FACS)-selectable marker. HiBiT tagging thus allows highly sensitive luminescence detection of endogenous PAX3-FOXO1 levels permitting quantitative high-throughput screening of large compound libraries for the discovery of PAX3-FOXO1 inhibitors and degraders.

Genetic Knockin Approaches to Reconstructing DLBCL-Subtypes from Human Germinal Center B-Cells

Background and Significance: The clinical and genetic heterogeneity of DLBCL has led to the recent classification of the disease into five genetically distinct subtypes. The MCD/C5 subtype of DLBCL is defined by genetic alterations of CD79B at the first tyrosine residue of its ITAM motif (CD79B Y196H), crucial for signaling downstream of the BCR, and MYD88 (MYD88 L265P), a signaling adaptor protein essential for Toll Like receptor signaling. CD79B Y196H and MYD88 L265P mutations together define the MCD/C5 subtype and were found to be co-dependent for proliferation and oncogenic BCR-signaling in vitro. However, the mechanism of epistatic interaction between CD79B Y196H and MYD88 L265P is unknown. The MCD/C5 DLBCL subtype is associated with particularly poor clinical outcomes, however only a few human cell lines and a genetic mouse model are available for preclinical testing. Here, we developed a novel genetic approach to engineer the MCD/C5 subtype of DLBCL in primary human tonsillar germinal center B cells. To this end, we introduced the CD79B Y196H and MYD88 L265P mutations as genetic knockin alleles utilizing CRISPR-Cas9 homology directed repair templates (HDRT). In addition, we developed humanized NBSGW mice carrying a mouse Tslp-transgene to enable development of lymph nodes and secondary lymphoid organs to propagate genetically engineered GC B-cells carrying CD79B Y196H and MYD88 L265P knockin mutations for lymphoma development ( Figure 1). Results: Primary human germinal center B-cells were isolated from tonsils and grown on follicular dendritic feeder cells expressing CD40L and IL21. Single strand DNA templates were designed for CD79B, with and without Y196H mutation. Constructs were linked to fluorescent reporters by T2A sites and flanked by homology arms of ~280 bp targeting the guide RNA cut site. The target site was selected to disrupt the endogenous gene, while placing the construct under transcriptional control of endogenous promoter/enhancer elements. We achieved a knock-in efficiency of 5-8%, prior to single-cell cloning and genotyping to confirm the presence of heterozygous CD79B Y196H mutant CD79B Y196 wild-type mutations ( Figure 1). Preliminary biochemical studies show a delayed response and reduced intensity of signaling downstream of the BCR upon stimulation in CD79B Y196H mutant, compared to CD79B Y196 wild-type knockin germinal center B-cells. A similar approach was taken to design temples encoding Exon 5 of MYD88, with and without the L265P mutation, utilizing two guide RNAs to excise and replace Exon 5, linked to a mCherry reporter by a T2A site. Upon successfully introducing the MYD88 HDR templates into our CD79B wild-type and mutant clones to generate four distinct genotypes based on wildtype and CD79B Y196H and MYD88 L265P mutations alone or in combination. In addition, traditional NSG mice lack lymph nodes and secondary lymphoid organs. Lack of lymphoid follicles in NSG mice, owing to defective IL7Rα-signaling, contributed to lack of lymphoma formation observed in the previous model. In our model, we substituted the missing IL7Rα-signal with transgenic expression of murine Tslp, which enables development of embryonic lymphoid tissue inducer (LTi) cells that are required for the development of secondary lymphoid organs and germinal center follicles supporting initiation of human DLBCL development. Conclusion: Previous approaches to model DLBCL starting from germinal center B-cells were based on lentiviral overexpression of mutant oncogenes. Thereby, tonsillar germinal center B-cells with lentiviral BCL2 and BCL6 overexpression did not form lymphomas in NSG mice, however, were permissive to malignant transformation by MYC. Building on this previous work, we propose an improved model for preclinical testing for genetically defined DLBCL subtypes. Our two main improvements involve (i) genetic modeling based on knockin-strategies targeting endogenous loci and transcriptional control elements to phenocopy physiological gene expression levels and transcriptional regulation, and (ii) substitution of defective IL7Rα-signaling by transgenic expression of murine Tslp to rescue development of LTi cells. Transgenic expression of mTslp enables LTi-dependent formation of lymph nodes and secondary lymphoid organs supporting lymphoma development.

Platinum Talen-Mediated Knock-in of Single-Stranded DNA Templates Facilitates Manufacturing of Clinical-Scale Non-Virally Gene-Edited T Cells from Peripheral Blood

Background: During the past decade, technologies that manufacture gene-modified immune cells, such as chimeric antigen receptor T cells, have been rapidly evolved and introduced in human clinics. T cells reprogrammed to express antigen-specific T-cell receptors (TCR-T) are an up-and-coming tool to treat intractable infection or malignant neoplasm in a personalized manner. However, several caveats still exist in the production of TCR-T for clinical use, because competitive surface expression of endogenous and transduced TCRs might hamper desired expression of transduced TCR. Moreover, mispairing of TCR subunits could mediate unexpected harmful immune complications. Therefore, in this study, we aimed to develop a method to introduce any desired TCRs in human primary T cells independent of interference with endogenous TCR by way of targeted knockout of TRA and TRB gene loci using highly-efficient transcription activator-like effector nuclease (TALEN), named Platinum TALEN (Sakuma T, Sci Rep 2013). Methods: We designed and synthesized two pairs of 5'-Cap-1 Platinum TALEN mRNA targeting the gene sequences that encode constant regions of human TRA ( TRAC) and TRB ( TRBC). TCR knockout efficiencies by these TALEN pairs were assessed by flow cytometric analysis using anti-CD3/TCR antibodies. Next, we constructed single-stranded DNA (ssDNA) homology-directed repair templates for the knock-in of desired TCR transgene into the TALEN target sequence of the TRAC locus. As a model TCR, we selected a 1G4 clone that reacts with NY-ESO-1-derived peptide in an HLA-A*02-restricted manner. We delivered these TALEN mRNA pairs and 1G4-encoding ssDNA by electroporation into primary human T cells collected from healthy volunteer adult donors after 72 hours of anti-CD3/CD28 beads activation. Genome-edited T cells were subsequently cultured in Therapeak TM X-VIVO15 (Lonza) supplemented with 10% human AB serum in the presence of 2-mercaptoethanol and interleukin-2 followed by second anti-CD3/CD28 beads activation. Density of T cells were maintained approximately at 1.0-3.0 x10 6/mL and transferred to larger flasks with additional fresh media as appropriate according to cell counts throughout the culture.1G4-transduced T cells were detected and counted by HLA-A*02:01 NY-ESO-1 tetramers at 17-22 days after electroporation. Additionally, we performed optical genome mapping analysis (Bionano Genomics) of Platinum TALEN-treated cells to screen genome-editing-associated large chromosomal structural variations (SVs) including unintended translocation. Results: Platinum TALEN mRNA pairs targeting TRAC and TRBC yielded a high proportion (>70%) of CD3/TCR negative T cells without the cost of compromised cell viability. By co-electroporation of 1G4 ssDNA and these Platinum TALEN mRNA pairs, we successfully obtained an average of 10.5 (range 2.2-19.0) million (n=5)NY-ESO-1-tetramer-positive cells with high viability from 3 millionprimary T cells until 3 weeks after electroporation. By flow cytometric analysis, a proportion of 1G4-tetramer positive cells that express Vβ chains other than Vβ13 (Vβ of 1G4) was approximately 10%. The PD-1-positive fraction in the expanded 1G4-expressing T cells was less than 10%, while levels of TIM-3 expression were 40-60%. These tetramer-positive cells showed specific cytotoxicities against HLA-A*02-positive cells pulsed with cognate NY-ESO-1 peptides. In an optical genome mapping analysis of genomic DNA obtained from TALEN-treated T cells, we found unique SVs with relatively small sizes and low coverage but no large unique SVs with high coverage when compared with control genomic DNA. Additionally, no interchromosomal translocation was observed. Conclusions: We have successfully designed Platinum TALEN mRNA pairs targeting TRAC and TRBC which facilitate the production of genome-edited human primary T cells. We also developed an electroporation and expansion culture protocol that enables us to produce viable genome-edited 1G4-transduced T cells at a large cell dose equivalent to a clinical scale. Collectively, these results suggest that targeted orthotopic knock-in of antigen-specific TCR-encoding ssDNA into the TCR locus by the use of Platinum TALEN is a promising strategy that can be applied for the clinical manufacturing of therapeutic TCR-T cells.

Generation of iPSC lines with SLC16A2:G401R or SLC16A2 knock out

The X-linked Allan-Herndon-Dudley syndrome (AHDS) is characterized by severely impaired psychomotor development and is caused by mutations in the SLC16A2 gene encoding the thyroid hormone transporter MCT8 (monocarboxylate transporter 8). By targeting exon 3 of SLC16A2 using CRISPR/Cas9 with single-stranded oligodeoxynucleotides as homology-directed repair templates, we introduced the AHDS patient missense variant G401R and a novel knock-out deletion variant (F400Sfs*17) into the male healthy donor hiPSC line BIHi001-B. We successfully generated cerebral organoids from these genome-edited lines, demonstrating the utility of the novel lines for modelling the effects of MCT8-deficency on human neurodevelopment.

445 Awardee Talk: Generation of a BMPR2 Heterozygous Ovine Genetic Model for Heritable Pulmonary Arterial Hypertension

Abstract Pulmonary Arterial Hypertension (PAH) is a rare vascular disorder affecting 1 to 2 cases per million individuals. Monoallelic mutations in the BMPR2 gene are the most common genetic risk factor, with about 20% of carriers affected. BMPR2 is a serine/threonine kinase receptor in the transforming growth factor beta (TGF-β) superfamily. Despite well-known associations with PAH, the role of BMPR2 in the progression of the disorder remains poorly understood due in part to the lack of appropriate preclinical genetic models. The homozygous BMPR2 (-/-) knockout genotype is embryonic lethal, which is an obstacle to creating a suitable genetic biomedical model for PAH. To overcome this barrier, we used a single guide RNA (sgRNA) CRISPR/Cas9 approach to target exon 3 of the ovine BMPR2 gene, in combination with a single-stranded oligo-deoxynucleotide (ssODN) synonymous homology-directed repair (HDR) template to create heterozygous BMPR2 (+/-) sheep. Heterozygote lambs could be generated in two ways with this approach – high editing efficiency coupled with ssODN homology-directed repair of one of the edited alleles, OR intermediate editing efficiency with low to no presence of HDR in the genetic sequence. Our main objectives were to:1.) Determine optimal concentrations of ssODN (100, 300, and 600 ng/ul) to promote HDR and 2.) Identify the sgRNA which maximized the probability of obtaining heterozygous lambs. A ssODN was designed to serve as a synonymous repair template that disrupted the PAM site for a pair of sgRNAs targeting BMPR2 exon 3 on opposite strands. We electroporated ovine zygotes six hours post-fertilization using sgRNA (100 ng/µL)/Cas9 (200 ng/µL) ribonucleoprotein (RNP) complexes alongside three ODN concentrations (100, 300, and 600 ng/µL). Zygotes were cultured for seven days, and blastocysts were lysed, PCR-amplified, and Sanger sequenced. Sequences were analyzed using TIDER (Brinkman et al., Nucl. Acids Res., 2018). TIDER is a computer model which quantifies the frequency of targeted small nucleotide changes introduced by CRISPR in combination with HDR using a donor template. A Kruskal-Wallis test compared the efficacy of HDR at 100, 300, and 600 ng/µL of ODN template. A significant difference was found between groups with 600 ng/µL giving the highest rate of HDR (P < 0.0001). Post-hoc pairwise comparisons using a Dunn's test found that sgRNA1 at 600 ng/ul ODN was significantly more efficient for HDR repair as compared with sgRNA2 (P < 0.05). The editing efficiency of the blastocysts collected using the 600 ng/µLODN and sgRNA1 treatment combination was 44.7% high (>66%), 42.6% intermediate (66-33%), and 8.5% low (< 33%) knockout rate, and 4.2% wild-type. The knockin rate was 27.7% high (>20%), 21.3% intermediate (20-5%), and 51% low to no (< 5%) HDR. These data predict that about one-half the transferred embryos subject to these conditions will have the desired heterozygous genotype [BMPR2 (+/-) sheep].

Multiplex Prime Editing and PASSIGE TM for Non-Viral Generation of an Allogeneic CAR-T Cell Product

While CAR-T cell therapy represents a major advance in personalized immunotherapy, the autologous nature of commercially available CAR-T products have delayed its broad application beyond a subset of hematological malignancies. In particular, manufacturing autologous CAR-T therapies is costly and time-consuming, and success relies on the fitness of a patient's cells. An allogeneic off-the-shelf CAR-T product could overcome these cell quality and quantity issues and could be accessed on-demand. However, a successful allogeneic CAR-T product will require multiplex gene knockout in addition to stable CAR integration to prevent graft-versus-host disease (GvHD) and graft rejection by the patient's immune system. Strategies for the delivery and expression of CAR transgenes include semi-random integration using lentivirus or transposons, which risk unintended gene disruption or activation. Targeted integration can be achieved using nucleases in combination with a template for homology-directed repair (HDR), but efficiency of this approach is generally low and there are risks associated with the induction of double-strand breaks (DSBs), including p53 activation and chromothripsis. This is further complicated in a situation where multiplex editing is required, as multiple DSBs can lead to gross chromosomal rearrangements. Alternatively, base editors have been used for targeted gene disruption without the induction of DSBs via single nucleotide conversion to disrupt a splice site or introduce a single premature stop (pmSTOP) codon. However, this approach is limited to transition mutations and cannot be used to integrate large genetic cargo, which is required to generate CAR-T cells. Prime Editing (PE) can be used for precise and programmable gene disruption via multiple strategies, including introduction of multiple pmSTOP codons, splice site disruption, frameshift mutations, or deletion of large segments of regulatory or coding sequence. Further, Prime-Assisted Site-Specific Integrase Gene Editing (PASSIGE) can be used to precisely and flexibly integrate large genetic cargo at a specific locus. Together, PE-mediated gene knock out and PASSIGE can be tailored to generate a more broadly applicable, potentially safer, and more effective CAR-T cell product. To evaluate the efficacy of an all-PE non-viral approach to allogeneic CAR-T cell generation, PASSIGE and PE mediated gene knockout were used to generate CD19-CAR-T cells. Multiplex PE precisely disrupted expression of the endogenous T cell receptor alpha constant ( TRAC) and beta-2-microglobulin ( B2M) loci in over 90% of human T cells. Co-delivery of a non-viral DNA donor template with PASSIGE editing components resulted in targeted integration of a 3.5 kb CD19-CAR transgene expression cassette at the TRAC locus in over 60% of the T cells, with no observed impact on T cell viability, phenotype, or functionality. CD19 CAR-T cells generated using PASSIGE show potent anti-tumor activity and cytokine production in response to CD19 + tumor cell lines in vitro and in vivo. The PASSIGE-generated CD19 CAR-T cells reduce tumor burden and prolong survival of mice bearing CD19 + tumors. These results show that a PE platform can be used to generate multiplex edited CAR-T cells without the need for viral vectors and without causing DSBs. This modular, one-step, non-viral delivery Prime Editing platform expands the applicability of T cell therapies for the treatment of tumors and immune diseases.

Targeted Integration of a Chimeric Antigen Receptor in Key Selected Loci in Primary Natural Killer Cells

Despite promising preclinical studies and early clinical data (Liu et al., NEJM, 2020), chimeric antigen receptor natural killer cells (CAR NK) are not yet on par with CAR T cells for the treatment of malignant diseases. Much effort has gone into developing NK-optimized CAR architectures (Li et al., Cell Stem Cell, 2018) and to increasing the persistence of CAR NK cells. We hypothesized that CAR NK cells could also benefit from the targeted integration of a CAR transgene, as compared with semi-random integration obtained using gammaretroviral vectors. Precise targeting has proven highly advantageous for TRAC CAR T cells owing to the optimal regulation and homogeneous expression conferred by the endogenous TCR alpha promoter (Eyquem et al., Nature, 2017). We first optimized an editing protocol for primary NK cells that uses electroporation of a CRISPR/Cas9 ribonucleoprotein followed by transduction with AAV6 bearing a homology-directed repair template. With this protocol we were able to knock out (KO) genes (including CD38, B2M, and different NK receptors), knock in transgenes (GFP-fusion protein, truncated EGFR, CAR), or both, at high efficiency. There was only moderate cell toxicity, and proliferation and cytotoxicity were conserved. Because of the high constitutive expression of B2M, we initially chose to focus on this locus and targeted a 1928z CAR transgene for regulation under the endogenous B2M promoter; our template design also rescues the B2M gene upon knock-in, enabling intermediate B2M expression (Figure 1). This strategy conferred high expression of the CAR and significantly more homogeneity as compared with gammaretroviral delivery (Figure 1). Furthermore, the rescued B2M expression protected B2M CAR NK cells from fratricide killing, whereas the B2M KO NK cells and the CAR NK cells with disrupted B2M expression were susceptible. B2M CAR NK cells also displayed higher in vitro cytotoxicity as compared with WT NK cells against two different CD19-expressing tumor cell lines (NALM6, SupB15). We tested B2M CAR NK cells in an NSG mouse xenograft model with an intraperitoneally injected human ovarian cancer cell line engineered to express CD19 (CD19 + SKOV3). Unexpectedly, compared to retroviral CAR-NK cells, B2M CAR NK cells conferred less tumor control (assessed by bioluminescence imaging) and less of a survival benefit. Therefore, we screened a panel of other loci for CAR targeting: five for NK activating receptors ( CD16A, CD56, DNAM1, NKp30, NKp46) and four for NK inhibitory receptors ( CD161, LILRB1, NKG2A, TIGIT). For activating receptors, the transgene was designed to use the endogenous promoter and to maintain expression of the targeted locus. For inhibitory receptors, the transgene was placed under the control of an EF1a exogenous promoter and designed to disrupt expression of the targeted locus. We compared these constructs and the initial B2M CAR NK design for CAR expression (by flow cytometry) and in vitro functionality (cytotoxicity assays) and observed wide variation in knock-in efficiency and CAR expression (Figure 2). For the top constructs based on knock-in efficiency, we conducted in vitro cytotoxicity assays against NALM6 and SupB15 cell lines. We found that when normalized for CAR percentage using the respective KO conditions, all tested knock-in loci performed better than B2M CAR NK cells, despite the lower CAR expression. Our current focus is on validating these top loci for the targeted integration of a 1928z CAR using in vivo models for comparison with B2M CAR NK and retroviral CAR NK. In summary, we have identified advantageous in vitro characteristics for the targeted integration of a CAR transgene in primary NK cells and screened 10 loci for CAR expression and in vitro cytotoxicity against tumor target cells. Confirmatory in vivo studies are ongoing and will be presented.

SsDNA is not superior to dsDNA as long HDR donors for CRISPR-mediated endogenous gene tagging in human diploid RPE1 and HCT116 cells

BackgroundRecent advances in CRISPR technology have enabled us to perform gene knock-in in various species and cell lines. CRISPR-mediated knock-in requires donor DNA which serves as a template for homology-directed repair (HDR). For knock-in of short sequences or base substitutions, ssDNA donors are frequently used among various other forms of HDR donors, such as linear dsDNA. However, partly due to the complexity of long ssDNA preparation, it remains unclear whether ssDNA is the optimal type of HDR donors for insertion of long transgenes such as fluorescent reporters in human cells.ResultsIn this study, we established a nuclease-based simple method for the preparation of long ssDNA with high yield and purity, and comprehensively compared the performance of ssDNA and dsDNA donors with 90 bases of homology arms for endogenous gene tagging with long transgenes in human diploid RPE1 and HCT116 cells. Quantification using flow cytometry revealed lower efficiency of endogenous fluorescent tagging with ssDNA donors than with dsDNA. By analyzing knock-in outcomes using long-read amplicon sequencing and a classification framework, a variety of mis-integration events were detected regardless of the donor type. Importantly, the ratio of precise insertion was lower with ssDNA donors than with dsDNA. Moreover, in off-target integration analyses using donors without homology arms, ssDNA and dsDNA were comparably prone to non-homologous integration.ConclusionsThese results indicate that ssDNA is not superior to dsDNA as long HDR donors with relatively short homology arms for gene knock-in in human RPE1 and HCT116 cells.

A Multifunctional and Highly Adaptable Reporter System for CRISPR/Cas Editing.

CRISPR/Cas systems are some of the most promising tools for therapeutic genome editing. The use of these systems is contingent on the optimal designs of guides and homology-directed repair (HDR) templates. While this design can be achieved in silico, validation and further optimization are usually performed with the help of reporter systems. Here, we describe a novel reporter system, termed BETLE, that allows for the fast, sensitive, and cell-specific detection of genome editing and template-specific HDR by encoding multiple reporter proteins in different open-reading frames. Out-of-frame non-homologous end joining (NHEJ) leads to the expression of either secretable NanoLuc luciferase, enabling a highly sensitive and low-cost analysis of editing, or fluorescent mTagBFP2, allowing for the enumeration and tissue-specific localization of genome-edited cells. BETLE includes a site to validate CRISPR/Cas systems for a sequence-of-interest, making it broadly adaptable. We evaluated BETLE using a defective moxGFP with a 39-base-pair deletion and showed spCas9, saCas9, and asCas12a editing as well as sequence-specific HDR and the repair of moxGFP in cell lines with single and multiple reporter integrants. Taken together, these data show that BETLE allows for the rapid detection and optimization of CRISPR/Cas genome editing and HDR in vitro and represents a state-of the art tool for future applications in vivo.

Peptide-mediated delivery of CRISPR enzymes for the efficient editing of primary human lymphocytes.

CRISPR-mediated genome editing of primary human lymphocytes is typically carried out via electroporation, which can be cytotoxic, cumbersome and costly. Here we show that the yields of edited primary human lymphocytes can be increased substantially by delivering a CRISPR ribonucleoprotein mixed with an amphiphilic peptide identified through screening. We evaluated the performance of this simple delivery method by knocking out genes in T cells, B cells and natural killer cells via the delivery of Cas9 or Cas12a ribonucleoproteins or an adenine base editor. We also show that peptide-mediated ribonucleoprotein delivery paired with an adeno-associated-virus-mediated homology-directed repair template can introduce a chimaeric antigen receptor gene at the T-cell receptor α constant locus, and that the engineered cells display antitumour potency in mice. The method is minimally perturbative, does not require dedicated hardware, and is compatible with multiplexed editing via sequential delivery, which minimizes the risk of genotoxicity. The peptide-mediated intracellular delivery of ribonucleoproteins may facilitate the manufacturing of engineered T cells.

Optimization of Electroporation and Adeno-Associated Virus-Mediated Generation of 2.7 kb Knock-In Livestock Blastocysts

The production of genome-edited livestock has traditionally relied on either somatic cell nuclear transfer cloning of an edited cell or microinjection of gene-editing reagents into developing embryos. Electroporation of gene-editing reagents into zygotes offers an alternative that is both high throughput and easy to use. Transfer of large (>1 kb) homology-directed repair (HDR) donor nucleic acid templates into mammalian zygotes is hindered by the thick extracellular zona pellucida (ZP) glycoprotein layer. A potentially scalable approach to deliver HDR templates of up to 4.9 kb into zygotes is to use recombinant adeno-associated virus (rAAV) vectors. In this study, electroporation conditions were optimized to efficiently deliver CRISPR-associated protein 9 (Cas9):single guide RNA (sgRNA) ribonucleoprotein (RNP) complexes to bovine and ovine zygotes, while retaining embryo viability. rAAV vectors were then used to transduce a 3.9-kb HDR template in combination with electroporation of Cas9:sgRNA RNP editing reagents 6 h postinsemination to generate bovine blastocysts with a targeted 2.7 kb knock-in (KI) at the H11 safe harbor locus. With this approach, there was no need to remove or weaken the ZP of the blastocysts that developed, and a KI rate of up to ∼38% was observed. However, to circumvent transduction of the cumulus cells, the oocytes were denuded before transduction that decreased the blastocyst development rate. In addition, genetic mosaicism was observed in the blastocysts. These results highlight that further optimization of editing approaches will be required to achieve nonmosaic embryos with a targeted KI at scale for livestock applications.

CRISPR‐Cas9 Genome Editing in the Moss Physcomitrium (Formerly Physcomitrella) patens

Until recently, precise genome editing has been limited to a few organisms. The ability of Cas9 to generate double stranded DNA breaks at specific genomic sites has greatly expanded molecular toolkits in many organisms and cell types. Before CRISPR-Cas9 mediated genome editing, P. patens was unique among plants in its ability to integrate DNA via homologous recombination. However, selection for homologous recombination events was required to obtain edited plants, limiting the types of editing that were possible. Now with CRISPR-Cas9, molecular manipulations in P. patens have greatly expanded. This protocol describes a method to generate a variety of different genome edits. The protocol describes a streamlined method to generate the Cas9/sgRNA expression constructs, design homology templates, transform, and quickly genotype plants. © 2023 Wiley Periodicals LLC. Basic Protocol 1: Constructing the Cas9/sgRNA transient expression vector Alternate Protocol 1: Shortcut to generating single and pooled Cas9/sgRNA expression vectors Basic Protocol 2: Designing the oligonucleotide-based homology-directed repair (HDR) template Alternate Protocol 2: Designing the plasmid-based HDR template Basic Protocol 3: Inducing genome editing by transforming CRISPR vector into P. patens protoplasts Basic Protocol 4: Identifying edited plants.

Assessment of distant-site rescue elements for CRISPR toxin-antidote gene drives.

Gene drive is a genetic engineering technology that can enable super-mendelian inheritance of specific alleles, allowing them to spread through a population. New gene drive types have increased flexibility, offering options for confined modification or suppression of target populations. Among the most promising are CRISPR toxin-antidote gene drives, which disrupt essential wild-type genes by targeting them with Cas9/gRNA. This results in their removal, increasing the frequency of the drive. All these drives rely on having an effective rescue element, which consists of a recoded version of the target gene. This rescue element can be at the same site as the target gene, maximizing the chance of efficient rescue, or at a distant site, which allows useful options such as easily disrupting another essential gene or increasing confinement. Previously, we developed a homing rescue drive targeting a haplolethal gene and a toxin-antidote drive targeting a haplosufficient gene. These successful drives had functional rescue elements but suboptimal drive efficiency. Here, we attempted to construct toxin-antidote drives targeting these genes with a distant-site configuration from three loci in Drosophila melanogaster. We found that additional gRNAs increased cut rates to nearly 100%. However, all distant-site rescue elements failed for both target genes. Furthermore, one rescue element with a minimally recoded sequence was used as a template for homology-directed repair for the target gene on a different chromosomal arm, resulting in the formation of functional resistance alleles. Together, these results can inform the design of future CRISPR-based toxin-antidote gene drives.

Gene Editing of Primary Rhesus Macaque B Cells.

B cells and their progeny are the sources of highly expressed antibodies. Their high protein expression capabilities together with their abundance, easy accessibility via peripheral blood, and amenability to simple adoptive transfers have made them an attractive target for gene editing approaches to express recombinant antibodies or other therapeutic proteins. The gene editing of mouse and human primary B cells is efficient, and mouse models for in vivo studies have shown promise, but feasibility and scalability for larger animal models have so far not been demonstrated. We, therefore, developed a protocol to edit rhesus macaque primary B cells in vitro to enable such studies. We report conditions for in vitro culture and gene-editing of primary rhesus macaque B cells from peripheral blood mononuclear cells or splenocytes using CRISPR/Cas9. To achieve the targeted integration of large (<4.5 kb) cassettes, a fast and efficient protocol was included for preparing recombinant adeno-associated virus serotype 6 as a homology-directed repair template using a tetracycline-enabled self-silencing adenoviral helper vector. These protocols enable the study of prospective B cell therapeutics in rhesus macaques.

Fluorescent PCR-based Screening Methods for Precise Knock-in of Small DNA Fragments and Point Mutations in Zebrafish.

Generation of zebrafish (Danio rerio) models with targeted insertion of epitope tags and point mutations is highly desirable for functional genomics and disease modeling studies. Currently, CRISPR/Cas9-mediated knock-in is the method of choice for insertion of exogeneous sequences by providing a repair template for homology-directed repair (HDR). A major hurdle in generating knock-in models is the labor and cost involved in screening of injected fish to identify the precise knock-in events due to low efficiency of the HDR pathway in zebrafish. Thus, we developed fluorescent PCR-based high-throughput screening methods for precise knock-in of epitope tags and point mutations in zebrafish. Here, we provide a step-by-step guide that describes selection of an active sgRNA near the intended knock-in site, design of single-stranded oligonucleotide (ssODN) templates for HDR, quick validation of somatic knock-in using injected embryos, and screening for germline transmission of precise knock-in events to establish stable lines. Our screening method relies on the size-based separation of all fragments in an amplicon by fluorescent PCR and capillary electrophoresis, thus providing a robust and cost-effective strategy. Although we present the use of this protocol for insertion of epitope tags and point mutations, it can be used for insertion of any small DNA fragments (e.g., LoxP sites, in-frame codons). Furthermore, the screening strategy described here can be used to screen for precise knock-in of small DNA sequences in any model system, as PCR amplification of the target region is its only requirement. Key features This protocol expands the use of fluorescent PCR and CRISPR-STAT for screening of precise knock-in of small insertions and point mutations in zebrafish. Allows validation of selected sgRNA and HDR template within two weeks by somatic knock-in screening. Allows robust screening of point mutations by combining restriction digest with CRISPR-STAT. Graphical overview Overview of the three-phase knock-in pipeline in zebrafish (created with BioRender.com).

CRISPR/Cas9-based Engineering of Immunoglobulin Loci in Hybridoma Cells.

Development of the hybridoma technology by Köhler and Milstein (1975) has revolutionized the immunological field by enabling routine use of monoclonal antibodies (mAbs) in research and development efforts, resulting in their successful application in the clinic today. While recombinant good manufacturing practices production technologies are required to produce clinical grade mAbs, academic laboratories and biotechnology companies still rely on the original hybridoma lines to stably and effortlessly produce high antibody yields at a modest price. In our own work, we were confronted with a major issue when using hybridoma-derived mAbs: there was no control over the antibody format that was produced, a flexibility that recombinant production does allow. We set out to remove this hurdle by genetically engineering antibodies directly in the immunoglobulin (Ig) locus of hybridoma cells. We used clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9 (Cas9) and homology-directed repair (HDR) to modify antibody's format [mAb or antigen-binding fragment (Fab')] and isotype. This protocol describes a straightforward approach, with little hands-on time, leading to stable cell lines secreting high levels of engineered antibodies. Parental hybridoma cells are maintained in culture, transfected with a guide RNA (gRNA) targeting the site of interest in the Ig locus and an HDR template to knock in the desired insert and an antibiotic resistance gene. By applying antibiotic pressure, resistant clones are expanded and characterized at the genetic and protein level for their ability to produce modified mAbs instead of the parental protein. Finally, the modified antibody is characterized in functional assays. To demonstrate the versatility of our strategy, we illustrate this protocol with examples where we have (i) exchanged the constant heavy region of the antibody, creating chimeric mAb of a novel isotype, (ii) truncated the antibody to create an antigenic peptide-fused Fab' fragment to produce a dendritic cell-targeted vaccine, and (iii) modified both the constant heavy (CH)1 domain of the heavy chain (HC) and the constant kappa (Cκ) light chain (LC) to introduce site-selective modification tags for further derivatization of the purified protein. Only standard laboratory equipment is required, which facilitates its application across various labs. We hope that this protocol will further disseminate our technology and help other researchers. Graphical abstract.

CD25 As Feedback Regulator of Oncogenic TCR Signaling in T-Cell Lymphoma

Introduction: CD25, also known as interleukin-2 receptor α chain (IL-2Rα), forms a heterotrimer with IL-2Rβ and common γ chain, and mediates IL-2 signaling. IL-2 signaling is essential for T- and NK-cell survival and proliferation. Additionally, CD25 is required for development of thymic regulatory T cells (Treg) and their suppressive function. Interestingly, CD25 is upregulated in a variety of lymphoid and myeloid malignancies, which do not depend on IL-2 signaling, suggesting IL-2-independent roles of CD25. Here, we seek to investigate whether CD25 has previously unrecognized functions in T cells and T-cell lymphomas. Results: Our previous study in B cells has discovered the cytoplasmic tail of CD25 as an essential structural element for site-specific recruitment of inhibitory phosphatases. CD25-mediated shuttling of inhibitory phosphatases balanced strength of oncogenic B-cell receptor (BCR)-signaling and was essential for cell-survival in B-cell malignancies. Interestingly, we found that CD25 expressed on B cells was monomeric and showed dynamic recruitment to BCR or its oncogenic mimics within lipid rafts. Studying gene expression data across different hematopoietic lineages, we found CD25 was strongly induced in T cell receptor (TCR)-activated T cells, as well as in T-cell malignancies that harbor oncogenic TCR-mimics, such as PTCL. Our mechanistic studies discovered that the serine residue 268 (S268) in the cytoplasmic tail of CD25 was a principal substrate of protein kinase C (PKC), and that S268 phosphorylation by PKC was critical for cell surface expression of CD25. The proximity-based labeling assays (Bio-ID) revealed that CD25-S268 but not its phospho-dead mutant recruits PKC and its adapter RACK1, which scaffold the inhibitory phosphatases SHIP1 and SHP1 for site-specific activation in lipid rafts. CRISPR-mediated knock-in of CD25-S268A that abrogates the formation of inhibitory phosphatase complexes in primary human T cells enabled us to specifically interrogate the functional relevance of CD25-S268 in the regulation of TCR signaling. By using single stranded homology-directed repair templates (ssHDRT) that contain a Cas9 shuttling sequence, we were able to achieve 70% knock-in efficiency in primary human T cells (Figure 1). Signaling studies with these cells revealed that S268A mutation significantly augmented TCR signaling strength. Additionally, S268A mutation resulted in acute inactivation of inhibitory phosphatases including SHP1. Consistent with these results, CD25 shuttling to TCR complex was promptly induced upon TCR stimulation and was mitigated by S268 mutation. Furthermore, immunophenotyping showed that loss of S268 phosphorylation resulted in increased expression of T-cell activation markers such as CD44 and CD69. Based on the ex vivo results, we generated a transgenic mouse model with germline knock-in of CD25-S268A, which would allow us to study S268-specific roles in CD25-mediated feedback regulation of TCR and oncogenic signaling in vivo. Conclusions: The role of CD25 in mediating IL-2 signaling has been extensively studied. However, increasing evidence suggests that CD25 plays a more sophisticated role in TCR signaling and T-lymphomagenesis. Here we discovered a previously unrecognized function of CD25 as a feedback regulator of TCR signaling. TCR activation or oncogenic TCR-mimics induce CD25-mediated recruitment of inhibitory phosphatases in lipid rafts to mitigate TCR signaling or curb excessive oncogenic signaling. Figure 1. CRISPR-mediated knock-out of CD25 alone, or followed by knock-in of CD25-S268-GFP or CD25-A268-GFP, in primary human T cells. The proportion of CD25-expressing GFP+ cells are analyzed by flow cytometry on day 4 and day 30 post CRISPR. Figure 1View largeDownload PPTFigure 1View largeDownload PPT Close modal