Optimized design parameters for CRISPR Cas9 and Cas12a homology-directed repair

Oncogenic KRAS Mutations Confer a Unique Mechanotransduction Response to Peristalsis in Colorectal Cancer Cells

Colorectal cancer tumors start as polyps on the inner lining of the colorectum, in which they are exposed to the mechanics of peristalsis. Our previous work leveraged a custom-built peristalsis bioreactor to demonstrate that colonic peristalsis led to cancer stem cell enrichment in colorectal cancer cells. However, this malignant mechanotransductive response was confined to select colorectal cancer lines that harbored an oncogenic mutation in the Kirsten rat sarcoma virus (KRAS) gene. In this study, we explored the involvement of activating KRAS mutations on peristalsis-associated mechanotransduction in colorectal cancer. Peristalsis enriched cancer stem cell marker Leucine-rich repeat-containing G protein-coupled receptor 5 (LGR5) in KRAS mutant lines in a Wnt ligand-independent manner. Conversely, LGR5 enrichment in wild-type KRAS lines exposed to peristalsis were minimal. LGR5 enrichment downstream of peristalsis translated to increased tumorigenicity in vivo. Differences in mechanotransduction were apparent via unbiased gene set enrichment analysis, in which many unique pathways were enriched in wild-type versus mutant lines. Peristalsis also triggered β-catenin nuclear localization independent of Wnt ligands, particularly in KRAS mutant lines. The involvement of KRAS was validated via gain and loss of function strategies. Peristalsis-induced β-catenin activation and LGR5 enrichment depended on the activation of the MEK/ERK cascade. Taken together, our results demonstrated that oncogenic KRAS mutations conferred a unique peristalsis-associated mechanotransduction response to colorectal cancer cells, resulting in cancer stem cell enrichment and increased tumorigenicity. These mechanosensory connections can be leveraged in improving the sensitivity of emerging therapies that target oncogenic KRAS. Implications: Oncogenic KRAS empowers colorectal cancer cells to harness the mechanics of colonic peristalsis for malignant gain independent of other cooperating signals.

CRISPR/Cas9-editing of PRNP in Alpine goats

Misfolding of the cellular PrP (PrPc) protein causes prion disease, leading to neurodegenerative disorders in numerous mammalian species, including goats. A lack of PrPc induces complete resistance to prion disease. The aim of this work was to engineer Alpine goats carrying knockout (KO) alleles of PRNP, the PrPc-encoding gene, using CRISPR/Cas9-ribonucleoproteins and single-stranded donor oligonucleotides. The targeted region preceded the PRNPTer mutation previously described in Norwegian goats. Genome editors were injected under the zona pellucida prior to the electroporation of 565 Alpine goat embryos/oocytes. A total of 122 two-cell-stage embryos were transferred to 46 hormonally synchronized recipient goats. Six of the goats remained pregnant and naturally gave birth to 10 offspring. Among the 10 newborns, eight founder animals carrying PRNP genome-edited alleles were obtained. Eight different mutated alleles were observed, including five inducing KO mutations. Three founders carried only genome-edited alleles and were phenotypically indistinguishable from their wild-type counterparts. Among them, one male carrying a one base pair insertion leading to a KO allele is currently used to rapidly extend a PRNP-KO line of Alpine goats for future characterization. In addition to KO alleles, a PRNPdel6 genetic variant has been identified in one-third of founder animals. This new variant will be tested for its potential properties with respect to prion disease. Future studies will also evaluate the effects of genetic background on other characters associated with PRNP KO, as previously described in the Norwegian breed or other species.

Comprehensive analysis of off-target and on-target effects resulting from liver-directed CRISPR-Cas9-mediated gene targeting with AAV vectors

Comprehensive genome-wide studies are needed to assess the consequences of adeno-associated virus (AAV) vector-mediated gene editing. We evaluated CRISPR-Cas-mediated on-target and off-target effects and examined the integration of the AAV vectors employed to deliver the CRISPR-Cas components to neonatal mice livers. The guide RNA (gRNA) was specifically designed to target the factor IX gene (F9). On-target and off-target insertions/deletions were examined by whole-genome sequencing (WGS). Efficient F9-targeting (36.45% ± 18.29%) was apparent, whereas off-target events were rare or below the WGS detection limit since only one single putative insertion was detected out of 118 reads, based on >100 computationally predicted off-target sites. AAV integrations were identified by WGS and shearing extension primer tag selection ligation-mediated PCR (S-EPTS/LM-PCR) and occurred preferentially in CRISPR-Cas9-induced double-strand DNA breaks in the F9 locus. In contrast, AAV integrations outside F9 were not in proximity to any of ∼5,000 putative computationally predicted off-target sites (median distance of 70 kb). Moreover, without relying on such off-target prediction algorithms, analysis of DNA sequences close to AAV integrations outside the F9 locus revealed no homology to the F9-specific gRNA. This study supports the use of S-EPTS/LM-PCR for direct in vivo comprehensive, sensitive, and unbiased off-target analysis.

Efficient non-viral immune cell engineering using circular single-stranded DNA-mediated genomic integration

The use of adeno-associated viruses (AAVs) as donors for homology-directed repair (HDR)-mediated genome engineering is limited by safety issues, manufacturing constraints and restricted packaging limits. Non-viral targeted genetic knock-ins rely primarily on double-stranded DNA (dsDNA) and linear single-stranded DNA (lssDNA) donors. dsDNA is known to have low efficiency and high cytotoxicity, while lssDNA is challenging for scaled manufacture. In this study, we developed a non-viral genome writing catalyst (GATALYST) system that allows production of circular single-stranded DNAs (cssDNAs) up to approximately 20 kilobases as donor templates for highly efficient precision transgene integration. cssDNA donors enable knock-in efficiency of up to 70% in induced pluripotent stem cells (iPSCs) and improved efficiency in multiple clinically relevant primary immune cell types and at multiple genomic loci implicated for clinical applications with various nuclease editor systems. The high precision and efficiency in chimeric antigen receptor (CAR)-T and natural killer (NK) cells, improved safety, payload flexibility and scalable manufacturability of cssDNA shows potential for future applications of genome engineering.

Development of an optimized protocol for generating knockout cancer cell lines using the CRISPR/Cas9 system, with emphasis on transient transfection

The clustered regularly interspaced short palindromic repeats (CRISPR) system offers cost-effectiveness, high efficiency, precision, and ease of use compared to traditional gene editing techniques. In this study, we employed findings from prestigious investigations to develop an optimized approach for generating knockout cancer cell lines using a transient transfection method. This protocol introduces a distinctive approach that follows rigorous guidelines for designing gRNA to reduce off-target effects, a major challenge in CRISPR applications. Our step-by-step instructions allow researchers, particularly those with limited laboratory equipment and funding, as well as those undertaking CRISPR projects for the first time, to generate knockout cell lines using CRISPR technology in just ten weeks. This protocol covers all needs for enhancing various yields, such as transfection efficiency, and includes leveraging robust bioinformatics tools, conducting essential assays, isolating monoclonal cells via limiting dilution, validating knockout cells, and providing comprehensive troubleshooting recommendations. Using this method, we successfully created several new generations of colorectal cancer cell lines with monoallelic and biallelic knockouts of the epithelial cell adhesion molecule (EpCAM) gene. Our method, optimized for a wide spectrum of cancer cell lines, makes CRISPR more accessible for applications in personalized and precision medicine. It expands opportunities for novel investigations into cancer mechanisms and paves the way for potential therapeutic interventions.

Current Advances and Material Innovations in the Search for Novel Treatments of Phenylketonuria

Phenylketonuria (PKU) is a genetically inherited disease caused by a mutation of the gene encoding phenylalanine hydroxylase (PAH) and is the most common inborn error of amino acid metabolism. A deficiency of PAH leads to increased blood and brain levels of phenylalanine (Phe), which may cause permanent neurocognitive symptoms and developmental delays if untreated. Current management strategies for PKU consist of early detection through neonatal screening and implementation of a restrictive diet with minimal amounts of natural protein in combination with Phe-free supplements and low-protein foods to meet nutritional requirements. For milder forms of PKU, oral treatment with synthetic sapropterin (BH4), the cofactor of PAH, may improve metabolic control of Phe and allow for more natural protein to be included in the patient's diet. For more severe forms, daily injections of pegvaliase, a PEGylated variant of phenylalanine ammonia-lyase (PAL), may allow for normalization of blood Phe levels. However, the latter treatment has considerable drawbacks, notably a strong immunogenicity of the exogenous enzyme and the attached polymeric chains. Research for novel therapies of PKU makes use of innovative materials for drug delivery and state-of-the-art protein engineering techniques to develop treatments which are safer, more effective, and potentially permanent.

Use of paired Cas9-NG nickase and truncated sgRNAs for single-nucleotide microbial genome editing

The paired nickases approach, which utilizes clustered regularly interspaced short palindromic repeats (CRISPR)-CRISPR-associated proteins (Cas) nickase and dual guide RNA, has the advantage of reducing off-target effects by being able to double the target sequence. In this study, our research utilized the Cas9-NG nickase variant to minimize PAM sequence constraints, enabling the generation of paired nicks at desired genomic loci. We performed a systematic investigation into the formation sites for double nicks and the design of donor DNA within a bacterial model system. Although we successfully identified the conditions necessary for the effective formation of double nicks in vivo, achieving single-nucleotide level editing directly at the target sites in the genome proved challenging. Nonetheless, our experiments revealed that efficient editing at the single-nucleotide level was achievable on target DNA sequences that are hybridized with 5'-end-truncated dual single-guide RNAs (sgRNAs). Our findings contribute to a deeper understanding of the paired nickases approach, offering a single-mismatch intolerance design strategy for accurate nucleotide editing. This strategy not only enhances the precision of genome editing but also marks a significant step forward in the development of nickase-derived genome editing technologies.

CRISPR-Cas assisted diagnostics of plant viruses and challenges

Plant viruses threaten global food security by infecting commercial crops, highlighting the critical need for efficient virus detection to enable timely preventive measures. Current techniques rely on polymerase chain reaction (PCR) for viral genome amplification and require laboratory conditions. This review explores the applications of CRISPR-Cas assisted diagnostic tools, specifically CRISPR-Cas12a and CRISPR-Cas13a/d systems for plant virus detection and analysis. The CRISPR-Cas12a system can detect viral DNA/RNA amplicons and can be coupled with PCR or isothermal amplification, allowing multiplexed detection in plants with mixed infections. Recent studies have eliminated the need for expensive RNA purification, streamlining the process by providing a visible readout through lateral flow strips. The CRISPR-Cas13a/d system can directly detect viral RNA with minimal preamplification, offering a proportional readout to the viral load. These approaches enable rapid viral diagnostics within 30 min of leaf harvest, making them valuable for onsite field applications. Timely identification of diseases associated with pathogens is crucial for effective treatment; yet developing rapid, specific, sensitive, and cost-effective diagnostic technologies remains challenging. The current gold standard, PCR technology, has drawbacks such as lengthy operational cycles, high costs, and demanding requirements. Here we update the technical advancements of CRISPR-Cas in viral detection, providing insights into future developments, versatile applications, and potential clinical translation. There is a need for approaches enabling field plant viral nucleic acid detection with high sensitivity, specificity, affordability, and portability. Despite challenges, CRISPR-Cas-mediated pathogen diagnostic solutions hold robust capabilities, paving the way for ideal diagnostic tools. Alternative applications in virus research are also explored, acknowledging the technology's limitations and challenges.

protoSpaceJAM: an open-source, customizable and web-accessible design platform for CRISPR/Cas insertional knock-in

CRISPR/Cas-mediated knock-in of DNA sequences enables precise genome engineering for research and therapeutic applications. However, designing effective guide RNAs (gRNAs) and homology-directed repair (HDR) donors remains a bottleneck. Here, we present protoSpaceJAM, an open-source algorithm to automate and optimize gRNA and HDR donor design for CRISPR/Cas insertional knock-in experiments, currently supporting SpCas9, SpCas9-VQR and enAsCas12a Cas enzymes. protoSpaceJAM utilizes biological rules to rank gRNAs based on specificity, distance to insertion site, and position relative to regulatory regions. protoSpaceJAM can introduce 'recoding' mutations (silent mutations and mutations in non-coding sequences) in HDR donors to prevent re-cutting and increase knock-in efficiency. Users can customize parameters and design double-stranded or single-stranded donors. We validated protoSpaceJAM's design rules by demonstrating increased knock-in efficiency with recoding mutations and optimal strand selection for single-stranded donors. An additional module enables the design of genotyping primers for deep sequencing of edited alleles. Overall, protoSpaceJAM streamlines and optimizes CRISPR knock-in experimental design in a flexible and modular manner to benefit diverse research and therapeutic applications. protoSpaceJAM is available open-source as an interactive web tool at protospacejam.czbiohub.org or as a standalone Python package at github.com/czbiohub-sf/protoSpaceJAM.

Protocol for genomic recombineering in Yersinia ruckeri using CRISPR Cas12a coupled with the λ Red system

Genomic manipulation of Yersinia ruckeri, a pathogen of salmonid fish species, is essential for understanding bacterial physiology and virulence. Here, we present a protocol for genomic recombineering in Y. ruckeri, a species reluctant to standard genomic engineering, using CRISPR Cas12a coupled with the λ Red system. We describe steps for identifying protospacer guides, preparing repair template plasmids, and electroporating Yersinia cells with Cpf1 and protospacer plasmids with homologous arms. We then detail procedures for genome editing and plasmid curing.

Circumvention of Topoisomerase IIα Intron 19 Intronic Polyadenylation in Acquired Etoposide-Resistant Human Leukemia K562 Cells

DNA topoisomerase IIα (TOP2α; 170 kDa, TOP2α/170) is an essential enzyme for proper chromosome dysjunction by producing transient DNA double-stranded breaks and is an important target for DNA damage-stabilizing anticancer agents, such as etoposide. Therapeutic effects of TOP2α poisons can be limited due to acquired drug resistance. We previously demonstrated decreased TOP2α/170 levels in an etoposide-resistant human leukemia K562 subline, designated K/VP.5, accompanied by increased expression of a C-terminal truncated TOP2α isoform (90 kDa; TOP2α/90), which heterodimerized with TOP2α/170 and was a determinant of resistance by exhibiting dominant-negative effects against etoposide activity. Based on 3'-rapid amplification of cDNA ends, we confirmed TOP2α/90 as the translation product of a TOP2α mRNA in which a cryptic polyadenylation site (PAS) harbored in intron 19 (I19) was used. In this report, we investigated whether the resultant intronic polyadenylation (IPA) would be attenuated by blocking or mutating the I19 PAS, thereby circumventing acquired drug resistance. An antisense morpholino oligonucleotide was used to hybridize/block the PAS in TOP2α pre-mRNA in K/VP.5 cells, resulting in decreased TOP2α/90 mRNA/protein levels in K/VP.5 cells and partially circumventing drug resistance. Subsequently, CRISPR/CRISPR-associated protein 9 with homology-directed repair was used to mutate the cryptic I19 PAS (AATAAA→ACCCAA) to prevent IPA. Gene-edited clones exhibited increased TOP2α/170 and decreased TOP2α/90 mRNA/protein and demonstrated restored sensitivity to etoposide and other TOP2α-targeted drugs. Together, results indicated that blocking/mutating a cryptic I19 PAS in K/VP.5 cells reduced IPA and restored sensitivity to TOP2α-targeting drugs. SIGNIFICANCE STATEMENT: The results presented in this study indicate that CRISPR/CRISPR-associated protein 9 gene editing of a cryptic polyadenylation site (PAS) within I19 of the TOP2α gene results in the reversal of acquired resistance to etoposide and other TOP2-targeted drugs. An antisense morpholino oligonucleotide targeting the PAS also partially circumvented resistance.

Engineering plants using diverse CRISPR-associated proteins and deregulation of genome-edited crops

The CRISPR/Cas system comprises RNA-guided nucleases, the target specificity of which is directed by Watson-Crick base pairing of target loci with single guide (sg)RNA to induce the desired edits. CRISPR-associated proteins and other engineered nucleases are opening new avenues of research in crops to induce heritable mutations. Here, we review the diversity of CRISPR-associated proteins and strategies to deregulate genome-edited (GEd) crops by considering them to be close to natural processes. This technology ensures yield without penalties, advances plant breeding, and guarantees manipulation of the genome for desirable traits. DNA-free and off-target-free GEd crops with defined characteristics can help to achieve sustainable global food security under a changing climate, but need alignment of international regulations to operate in existing supply chains.

Impact of CRISPR/HDR editing versus lentiviral transduction on long-term engraftment and clonal dynamics of HSPCs in rhesus macaques

For precise genome editing via CRISPR/homology-directed repair (HDR), effective and safe editing of long-term engrafting hematopoietic stem cells (LT-HSCs) is required. The impact of HDR on true LT-HSC clonal dynamics in a relevant large animal model has not been studied. To track the output and clonality of HDR-edited cells and to provide a comparison to lentivirally transduced HSCs in vivo, we developed a competitive rhesus macaque (RM) autologous transplantation model, co-infusing HSCs transduced with a barcoded GFP-expressing lentiviral vector (LV) and HDR edited at the CD33 locus. CRISPR/HDR-edited cells showed a two-log decrease by 2 months following transplantation, with little improvement via p53 inhibition, in comparison to minimal loss of LV-transduced cells long term. HDR long-term clonality was oligoclonal in contrast to highly polyclonal LV-transduced HSCs. These results suggest marked clinically relevant differences in the impact of current genetic modification approaches on HSCs.

Protocol for the electroporation of CRISPR-Cas for DNA and RNA targeting in Bos taurus zygotes

The use of CRISPR-Cas9 ribonucleoproteins has revolutionized manipulation of genomes. Here, we present a protocol for the electroporation of CRISPR-Cas for DNA and RNA targeting in Bos taurus zygotes. First, we describe steps for production and preparation of presumptive zygotes for electroporation. The first electroporation introduces ribonucleoproteins formed by Cas9D10A with two guide RNAs to target DNA, and the second introduces the same ribonucleoprotein complex to target DNA plus Cas13a with one guide RNA to target RNAs. For complete details on the use and execution of this protocol, please refer to Nix et al.1.

Improving the Efficiency of CRISPR Ribonucleoprotein-Mediated Precise Gene Editing by Small Molecules in Porcine Fibroblasts

The aim of this study was to verify whether small molecules can improve the efficiency of precision gene editing using clustered regularly interspaced short palindromic repeats (CRISPR) ribonucleoprotein (RNP) in porcine cells. CRISPR associated 9 (Cas9) protein, small guide RNA (sgRNA), phosphorothioate-modified single-stranded oligonucleotides (ssODN), and different small molecules were used to generate precise nucleotide substitutions at the insulin (INS) gene by homology-directed repair (HDR) in porcine fetal fibroblasts (PFFs). These components were introduced into PFFs via electroporation, followed by polymerase chain reaction (PCR) for the target site. All samples were sequenced and analyzed, and the efficiencies of different small molecules at the target site were compared. The results showed that the optimal concentrations of the small molecules, including L-189, NU7441, SCR7, L755507, RS-1, and Brefeldin A, for in vitro-cultured PFFs' viability were determined. Compared with the control group, the single small molecules including L-189, NU7441, SCR7, L755507, RS-1, and Brefeldin A increased the efficiency of HDR-mediated precise gene editing from 1.71-fold to 2.28-fold, respectively. There are no benefits in using the combination of two small molecules, since none of the combinations improved the precise gene editing efficiency compared to single small molecules. In conclusion, these results suggested that a single small molecule can increase the efficiency of CRISPR RNP-mediated precise gene editing in porcine cells.

Current Strategies for Increasing Knock-In Efficiency in CRISPR/Cas9-Based Approaches

Since its discovery in 2012, the clustered regularly interspaced short palindromic repeats (CRISPR) and CRISPR-associated protein 9 (Cas9) system has supposed a promising panorama for developing novel and highly precise genome editing-based gene therapy (GT) alternatives, leading to overcoming the challenges associated with classical GT. Classical GT aims to deliver transgenes to the cells via their random integration in the genome or episomal persistence into the nucleus through lentivirus (LV) or adeno-associated virus (AAV), respectively. Although high transgene expression efficiency is achieved by using either LV or AAV, their nature can result in severe side effects in humans. For instance, an LV (NCT03852498)- and AAV9 (NCT05514249)-based GT clinical trials for treating X-linked adrenoleukodystrophy and Duchenne Muscular Dystrophy showed the development of myelodysplastic syndrome and patient's death, respectively. In contrast with classical GT, the CRISPR/Cas9-based genome editing requires the homologous direct repair (HDR) machinery of the cells for inserting the transgene in specific regions of the genome. This sophisticated and well-regulated process is limited in the cell cycle of mammalian cells, and in turn, the nonhomologous end-joining (NHEJ) predominates. Consequently, seeking approaches to increase HDR efficiency over NHEJ is crucial. This manuscript comprehensively reviews the current alternatives for improving the HDR for CRISPR/Cas9-based GTs.

Functional assay for assessment of pathogenicity of BAP1 variants

BACKGROUND

Pathogenic germline variants in BRCA1-Associated Protein 1 (BAP1) cause BAP1 tumor predisposition syndrome (BAP1-TPDS). Carriers run especially a risk of uveal (UM) and cutaneous melanoma, malignant mesothelioma, and clear cell renal carcinoma. Approximately half of increasingly reported BAP1 variants lack accurate classification. Correct interpretation of pathogenicity can improve prognosis of the patients through tumor screening with better understanding of BAP1-TPDS.

METHODS

We edited five rare BAP1 variants with differing functional characteristics identified from patients with UM in HAP1 cells using CRISPR-Cas9 and assayed their effect on cell adhesion/spreading (at 4 h) and proliferation (at 48 h), measured as cell index (CI), using xCELLigence real-time analysis system.

RESULTS

In BAP1 knockout HAP1 cultures, cell number was half of wild type (WT) cultures at 48 h (p = 0.00021), reaching confluence later, and CI was 78% reduced (p < 0.0001). BAP1-TPDS-associated null variants c.67+1G>T and c.1780_1781insT, and a likely pathogenic missense variant c.281A>G reduced adhesion (all p ≤ 0.015) and proliferation by 74%-83% (all p ≤ 0.032). Another likely pathogenic missense variant c.680G>A reduced both by at least 50% (all p ≤ 0.032), whereas cells edited with likely benign one c.1526C>T grew similarly to WT.

CONCLUSIONS

BAP1 is essential for optimal fitness of HAP1 cells. Pathogenic and likely pathogenic BAP1 variants reduced cell fitness, reflected in adhesion/spreading and proliferation properties. Further, moderate effects were quantifiable. Variant modelling in HAP1 with CRISPR-Cas9 enabled functional analysis of coding and non-coding region variants in an endogenous expression system.

Deep orange gene editing triggers temperature-sensitive lethal phenotypes in Ceratitis capitata

BACKGROUND

The Mediterranean fruit fly, Ceratitis capitata, is a significant agricultural pest managed through area-wide integrated pest management (AW-IPM) including a sterile insect technique (SIT) component. Male-only releases increase the efficiency and cost-effectiveness of SIT programs, which can be achieved through the development of genetic sexing strains (GSS). The most successful GSS developed to date is the C. capitata VIENNA 8 GSS, constructed using classical genetic approaches and an irradiation-induced translocation with two selectable markers: the white pupae (wp) and temperature-sensitive lethal (tsl) genes. However, currently used methods for selecting suitable markers and inducing translocations are stochastic and non-specific, resulting in a laborious and time-consuming process. Recent efforts have focused on identifying the gene(s) and the causal mutation(s) for suitable phenotypes, such as wp and tsl, which could be used as selectable markers for developing a generic approach for constructing GSS. The wp gene was recently identified, and efforts have been initiated to identify the tsl gene. This study investigates Ceratitis capitata deep orange (Ccdor) as a tsl candidate gene and its potential to induce tsl phenotypes.

RESULTS

An integrated approach based on cytogenetics, genomics, bioinformatics, and gene editing was used to characterize the Ccdor. Its location was confirmed on the right arm of chromosome 5 in the putative tsl genomic region. Knock-out of Ccdor using CRISPR/Cas9-NHEJ and targeting the fourth exon resulted in lethality at mid- and late-pupal stage, while the successful application of CRISPR HDR introducing a point mutation on the sixth exon resulted in the establishment of the desired strain and two additional strains (dor 12del and dor 51dup), all of them expressing tsl phenotypes and presenting no (or minimal) fitness cost when reared at 25 °C. One of the strains exhibited complete lethality when embryos were exposed at 36 °C.

CONCLUSIONS

Gene editing of the deep orange gene in Ceratitis capitata resulted in the establishment of temperature-sensitive lethal mutant strains. The induced mutations did not significantly affect the rearing efficiency of the strains. As deep orange is a highly conserved gene, these data suggest that it can be considered a target for the development of tsl mutations which could potentially be used to develop novel genetic sexing strains in insect pests and disease vectors.

CRISPR-Cas9-generated PTCHD1 2489T>G stem cells recapitulate patient phenotype when undergoing neural induction

An estimated 3.5%-5.9% of the global population live with rare diseases, and approximately 80% of these diseases have a genetic cause. Rare genetic diseases are difficult to diagnose, with some affected individuals experiencing diagnostic delays of 5-30 years. Next-generation sequencing has improved clinical diagnostic rates to 33%-48%. In a majority of cases, novel variants potentially causing the disease are discovered. These variants require functional validation in specialist laboratories, resulting in a diagnostic delay. In the interim, the finding is classified as a genetic variant of uncertain significance (VUS) and the affected individual remains undiagnosed. A VUS (PTCHD1 c. 2489T>G) was identified in a child with autistic behavior, global developmental delay, and hypotonia. Loss of function mutations in PTCHD1 are associated with autism spectrum disorder and intellectual disability; however, the molecular function of PTCHD1 and its role in neurodevelopmental disease is unknown. Here, we apply CRISPR gene editing and induced pluripotent stem cell (iPSC) neural disease modeling to assess the variant. During differentiation from iPSCs to neural progenitors, we detect subtle but significant gene signatures in synaptic transmission and muscle contraction pathways. Our work supports the causal link between the genetic variant and the child's phenotype, providing evidence for the variant to be considered a pathogenic variant according to the American College of Medical Genetics and Genomics guidelines. In addition, our study provides molecular data on the role of PTCHD1 in the context of other neurodevelopmental disorders.

Increasing Gene Editing Efficiency via CRISPR/Cas9- or Cas12a-Mediated Knock-In in Primary Human T Cells

T lymphocytes represent a promising target for genome editing. They are primarily modified to recognize and kill tumor cells or to withstand HIV infection. In most studies, T cell genome editing is performed using the CRISPR/Cas technology. Although this technology is easily programmable and widely accessible, its efficiency of T cell genome editing was initially low. Several crucial improvements were made in the components of the CRISPR/Cas technology and their delivery methods, as well as in the culturing conditions of T cells, before a reasonable editing level suitable for clinical applications was achieved. In this review, we summarize and describe the aforementioned parameters that affect human T cell editing efficiency using the CRISPR/Cas technology, with a special focus on gene knock-in.

Understanding genetic heterogeneity in gene-edited hematopoietic stem cell products

CRISPR/Cas gene editing has transformed genetic research and is poised to drive the next generation of gene therapies targeting hematopoietic stem cells (HSCs). However, the installation of the "desired" edit is most often only achieved in a minor subset of alleles. The array of cellular pathways triggered by gene editing tools produces a broad spectrum of "undesired" editing outcomes, including short insertions and deletions (indels) and chromosome rearrangements, leading to considerable genetic heterogeneity in gene-edited HSC populations. This heterogeneity may undermine the effect of the genetic intervention since only a subset of cells will carry the intended modification. Also, undesired mutations represent a potential safety concern as gene editing advances toward broader clinical use. Here, we will review the different sources of "undesired" edits and will discuss strategies for their mitigation and control.

Dissecting the Role of the Ccr4-Not Deadenylase Complex in Pluripotency and Differentiation

In this chapter, we describe methods and assays to examine the role of the Ccr4-Not mRNA deadenylase complex in regulating mouse embryonic stem cell (ESC) pluripotency and differentiation. We present the following procedures: sgRNA design and cloning, the culture of mouse ESCs, transfection and colony picking, genotyping, generation of Cnot3 conditional deletion ESCs, assessment of Cnot3 deletion efficiency, and examination of the impact on ESC maintenance and differentiation. The above approach and protocols can also be readily applied to study other Ccr4-Not subunits in ESC fate regulation, thereby facilitating the systematic dissection of Ccr4-Not function in stem cells and development.

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.

Impact of CRISPR/HDR-editing versus lentiviral transduction on long-term engraftment and clonal dynamics of HSPCs in rhesus macaques

For precise genome editing via CRISPR/homology-directed repair (HDR), effective and safe editing of long-term engrafting hematopoietic stem cells (LT-HSCs) requires both sufficient HDR efficiency and protection of LT-HSC function and number. The impact of HDR on true LT-HSCs clonal dynamics in a relevant large animal model has not previously been studied. To track the HDR-edited cells, autologous rhesus macaque (RM) CD34 + cells were electroporated with the gRNA/Cas9 ribonucleoprotein (RNP) and HDR cassette barcode library structure and reinfused into RMs following myeloablation. For competitive model animals, fractionated CD34 + cells were transduced with a barcoded GFP-expressing lentiviral vector (LV) and electroporated via HDR machinery, respectively. CD33 knockout (KO) neutrophils were prevalent early following engraftment and then rapidly decreased, resulting in less than 1% total editing efficiency. Interestingly, in competitive animals, a higher concentration of i53 mRNA result in a less steep reduction in CD33 KO cells, presented a modest decrease in HDR rate (0.1-0.2%) and total indels (1.5-6.5%). In contrast, the drop off of LV-transduced GFP + cells stabilized at 20% after 2 months. We next retrieved embedded barcodes and revealed that various clones contributed to early hematopoietic reconstitution, then after dominant clones appeared at steady state throughout the animals. In conclusion, CRISPR/HDR edited cells disappeared rapidly after the autologous transplantation in RM despite substantial gene editing outcome, whereas LV-transduced cells were relatively well maintained. Clonality of HDR-edited cells drastically shrank at early stage and then relied on several dominant clones, which can be mildly mitigated by the introduction of i53 mRNA.

Unleashing the potential of CRISPR multiplexing: Harnessing Cas12 and Cas13 for precise gene modulation in eye diseases

Gene therapy is a flourishing field with the potential to revolutionize the treatment of genetic diseases. The emergence of CRISPR-Cas9 has significantly advanced targeted and efficient genome editing. Although CRISPR-Cas9 has demonstrated promising potential applications in various genetic disorders, it faces limitations in simultaneously targeting multiple genes. Novel CRISPR systems, such as Cas12 and Cas13, have been developed to overcome these challenges, enabling multiplexing and providing unique advantages. Cas13, in particular, targets mRNA instead of genomic DNA, permitting precise gene expression control and mitigating off-target effects. This review investigates the potential of Cas12 and Cas13 in ocular gene therapy applications, such as suppression of inflammation and cell death. In addition, the capabilities of Cas12 and Cas13 are explored in addressing potential targets related with disease mechanisms such as aberrant isoforms, mitochondrial genes, cis-regulatory sequences, modifier genes, and long non-coding RNAs. Anatomical accessibility and relative immune privilege of the eye provide an ideal organ system for evaluating these novel techniques' efficacy and safety. By targeting multiple genes concurrently, CRISPR-Cas12 and Cas13 systems hold promise for treating a range of ocular disorders, including glaucoma, retinal dystrophies, and age-related macular degeneration. Nonetheless, additional refinement is required to ascertain the safety and efficacy of these approaches in ocular disease treatments. Thus, the development of Cas12 and Cas13 systems marks a significant advancement in gene therapy, offering the potential to devise effective treatments for ocular disorders.

Innovative Strategies of Reprogramming Immune System Cells by Targeting CRISPR/Cas9-Based Genome-Editing Tools: A New Era of Cancer Management

The recent developments in the study of clustered regularly interspaced short palindromic repeats/associated protein 9 (CRISPR/Cas9) system have revolutionized the art of genome-editing and its applications for cellular differentiation and immune response behavior. This technology has further helped in understanding the mysteries of cancer progression and possible designing of novel antitumor immunotherapies. CRISPR/Cas9-based genome-editing is now often used to engineer universal T-cells, equipped with recombinant T-cell receptor (TCR) or chimeric antigen receptor (CAR). In addition, this technology is used in cytokine stimulation, antibody designing, natural killer (NK) cell transfer, and to overcome immune checkpoints. The innovative potential of CRISPR/Cas9 in preparing the building blocks of adoptive cell transfer (ACT) immunotherapy has opened a new window of antitumor immunotherapy and some of them have gained FDA approval. The manipulation of immunogenetic regulators has opened a new interface for designing, implementation and interpretation of CRISPR/Cas9-based screening in immuno-oncology. Several cancers like lymphoma, melanoma, lung, and liver malignancies have been treated with this strategy, once thought to be impossible. The safe and efficient delivery of CRISPR/Cas9 system within the immune cells for the genome-editing strategy is a challenging task which needs to be sorted out for efficient immunotherapy. Several targeting approaches like virus-mediated, electroporation, microinjection and nanoformulation-based methods have been used, but each procedure offers some limitations. Here, we elaborate the recent updates of cancer management through immunotherapy in partnership with CRISPR/Cas9 technology. Further, some innovative methods of targeting this genome-editing system within the immune system cells for reprogramming them, as a novel strategy of anticancer immunotherapy is elaborated. In addition, future prospects and clinical trials are also discussed.

Generation of gene-of-interest double allele knockout clones in primary human T cells by CRISPR

Gene-of-interest (GOI) knockout is an important technique to study the genetic mechanisms of T cells. Here, we present a protocol to generate GOI double allele gene knockouts in primary human T cells by CRISPR, thus depleting proteins of interest expressed intracellularly or extracellularly in primary T cells. We describe steps for gRNA selection and efficiency validation, homology-directed repair (HDR) DNA template design and cloning, and genome editing and HDR gene insertion. We then detail clone isolation and GOI knockout validation. For complete details on the use and execution of this protocol, please refer to Wu et al.1.

Manipulating and studying gene function in human pluripotent stem cell models

Human pluripotent stem cells (hPSCs) are uniquely suited to study human development and disease and promise to revolutionize regenerative medicine. These applications rely on robust methods to manipulate gene function in hPSC models. This comprehensive review aims to both empower scientists approaching the field and update experienced stem cell biologists. We begin by highlighting challenges with manipulating gene expression in hPSCs and their differentiated derivatives, and relevant solutions (transfection, transduction, transposition, and genomic safe harbor editing). We then outline how to perform robust constitutive or inducible loss-, gain-, and change-of-function experiments in hPSCs models, both using historical methods (RNA interference, transgenesis, and homologous recombination) and modern programmable nucleases (particularly CRISPR/Cas9 and its derivatives, i.e., CRISPR interference, activation, base editing, and prime editing). We further describe extension of these approaches for arrayed or pooled functional studies, including emerging single-cell genomic methods, and the related design and analytical bioinformatic tools. Finally, we suggest some directions for future advancements in all of these areas. Mastering the combination of these transformative technologies will empower unprecedented advances in human biology and medicine.

Generation of CRISPR-Cas9-mediated knockin mutant models in mice and MEFs for studies of polymorphism in clock genes

The creation of mutant mice has been invaluable for advancing biomedical science, but is too time- and resource-intensive for investigating the full range of mutations and polymorphisms. Cell culture models are therefore an invaluable complement to mouse models, especially for cell-autonomous pathways like the circadian clock. In this study, we quantitatively assessed the use of CRISPR to create cell models in mouse embryonic fibroblasts (MEFs) as compared to mouse models. We generated two point mutations in the clock genes Per1 and Per2 in mice and in MEFs using the same sgRNAs and repair templates for HDR and quantified the frequency of the mutations by digital PCR. The frequency was about an order of magnitude higher in mouse zygotes compared to that in MEFs. However, the mutation frequency in MEFs was still high enough for clonal isolation by simple screening of a few dozen individual cells. The Per mutant cells that we generated provide important new insights into the role of the PAS domain in regulating PER phosphorylation, a key aspect of the circadian clock mechanism. Quantification of the mutation frequency in bulk MEF populations provides a valuable basis for optimizing CRISPR protocols and time/resource planning for generating cell models for further studies.

Homology-Directed-Repair-Based Genome Editing in HSPCs for the Treatment of Inborn Errors of Immunity and Blood Disorders

Genome engineering via targeted nucleases, specifically CRISPR-Cas9, has revolutionized the field of gene therapy research, providing a potential treatment for diseases of the blood and immune system. While numerous genome editing techniques have been used, CRISPR-Cas9 homology-directed repair (HDR)-mediated editing represents a promising method for the site-specific insertion of large transgenes for gene knock-in or gene correction. Alternative methods, such as lentiviral/gammaretroviral gene addition, gene knock-out via non-homologous end joining (NHEJ)-mediated editing, and base or prime editing, have shown great promise for clinical applications, yet all possess significant drawbacks when applied in the treatment of patients suffering from inborn errors of immunity or blood system disorders. This review aims to highlight the transformational benefits of HDR-mediated gene therapy and possible solutions for the existing problems holding the methodology back. Together, we aim to help bring HDR-based gene therapy in CD34+ hematopoietic stem progenitor cells (HSPCs) from the lab bench to the bedside.

Efficient bi-allelic tagging in human induced pluripotent stem cells using CRISPR

Allelic tagging of endogenous genes enables studying gene function and transcriptional control in the native genomic context. Here, we present an efficient protocol for bi-allelic tagging of protein-coding genes with fluorescent reporters in human iPSCs using the CRISPR-Cas9-mediated homology-directed repair. We detail steps for design, cloning, electroporation, and single-cell clone isolation and validation. The tagging strategy described in this protocol is readily applicable for knockin of other reporters in diverse cell types for biomedical research.

Perspectives of gene editing for cattle farming in tropical and subtropical regions

Cattle productivity in tropical and subtropical regions can be severely affected by the environment. Reproductive performance, milk and meat production are compromised by the heat stress imposed by the elevated temperature and humidity. The resulting low productivity contributes to reduce the farmer's income and to increase the methane emissions per unit of animal protein produced and the pressure on land usage. The introduction of highly productive European cattle breeds as well as crossbreeding with local breeds have been adopted as strategies to increase productivity but the positive effects have been limited by the low adaptation of European animals to hot climates and by the reduction of the heterosis effect in the following generations. Gene editing tools allow precise modifications in the animal genome and can be an ally to the cattle industry in tropical and subtropical regions. Alleles associated with production or heat tolerance can be shifted between breeds without the need of crossbreeding. Alongside assisted reproductive biotechnologies and genome selection, gene editing can accelerate the genetic gain of indigenous breeds such as zebu cattle. This review focuses on some of the potential applications of gene editing for cattle farming in tropical and subtropical regions, bringing aspects related to heat stress, milk yield, bull reproduction and methane emissions.

CRISPR/Cas9 mediated gene editing in non-model nematode Panagrolaimus sp. PS1159

The phylum Nematoda harbors a huge diversity of species in a broad range of ecosystems and habitats. Nematodes share a largely conserved Bauplan but major differences have been found in early developmental processes. The development of the nematode model organism Caenorhabditis elegans has been studied in great detail for decades. These efforts have provided the community with a large number of protocols and methods. Unfortunately, many of these tools are not easily applicable in non-Caenorhabditis nematodes. In recent years it has become clear that many crucial genes in the C. elegans developmental toolkit are absent in other nematode species. It is thus necessary to study the developmental program of other nematode species in detail to understand evolutionary conservation and novelty in the phylum. Panagrolaimus sp. PS1159 is a non-parasitic nematode exhibiting parthenogenetic reproduction and we are establishing the species to comparatively study evolution, biodiversity, and alternative reproduction and survival strategies. Here, we demonstrate the first successful application of the CRISPR/Cas9 system for genome editing in Panagrolaimus sp. PS1159 and the closely related hermaphroditic species Propanagrolaimus sp. JU765 applying the non-homologous end joining and the homology-directed repair (HDR) mechanisms. Using microinjections and modifying published protocols from C. elegans and P. pacificus we induced mutations in the orthologue of unc-22. This resulted in a visible uncoordinated twitching phenotype. We also compared the HDR efficiency following the delivery of different single-stranded oligodeoxynucleotides (ssODNs). Our work will expand the applicability for a wide range of non-model nematodes from across the tree and facilitate functional analysis into the evolution of parthenogenesis, changes in the developmental program of Nematoda, and cryptobiosis.

Richardson R. New advances in CRISPR/Cas-mediated precise gene-editing techniques

Over the past decade, CRISPR/Cas-based gene editing has become a powerful tool for generating mutations in a variety of model organisms, from Escherichia coli to zebrafish, rodents and large mammals. CRISPR/Cas-based gene editing effectively generates insertions or deletions (indels), which allow for rapid gene disruption. However, a large proportion of human genetic diseases are caused by single-base-pair substitutions, which result in more subtle alterations to protein function, and which require more complex and precise editing to recreate in model systems. Precise genome editing (PGE) methods, however, typically have efficiencies of less than a tenth of those that generate less-specific indels, and so there has been a great deal of effort to improve PGE efficiency. Such optimisations include optimal guide RNA and mutation-bearing donor DNA template design, modulation of DNA repair pathways that underpin how edits result from Cas-induced cuts, and the development of Cas9 fusion proteins that introduce edits via alternative mechanisms. In this Review, we provide an overview of the recent progress in optimising PGE methods and their potential for generating models of human genetic disease.

In search of an ideal template for therapeutic genome editing: A review of current developments for structure optimization

Gene therapy is a fast developing field of medicine with hundreds of ongoing early-stage clinical trials and numerous preclinical studies. Genome editing (GE) now is an increasingly important technology for achieving stable therapeutic effect in gene correction, with hematopoietic cells representing a key target cell population for developing novel treatments for a number of hereditary diseases, infections and cancer. By introducing a double strand break (DSB) in the defined locus of genomic DNA, GE tools allow to knockout the desired gene or to knock-in the therapeutic gene if provided with an appropriate repair template. Currently, the efficiency of methods for GE-mediated knock-in is limited. Significant efforts were focused on improving the parameters and interaction of GE nuclease proteins. However, emerging data suggests that optimal characteristics of repair templates may play an important role in the knock-in mechanisms. While viral vectors with notable example of AAVs as a donor template carrier remain the mainstay in many preclinical trials, non-viral templates, including plasmid and linear dsDNA, long ssDNA templates, single and double-stranded ODNs, represent a promising alternative. Furthermore, tuning of editing conditions for the chosen template as well as its structure, length, sequence optimization, homology arm (HA) modifications may have paramount importance for achieving highly efficient knock-in with favorable safety profile. This review outlines the current developments in optimization of templates for the GE mediated therapeutic gene correction.

Efficient Homology-Directed Repair with Circular Single-Stranded DNA Donors

While genome editing has been revolutionized by the advent of CRISPR-based nucleases, difficulties in achieving efficient, nuclease-mediated, homology-directed repair (HDR) still limit many applications. Commonly used DNA donors such as plasmids suffer from low HDR efficiencies in many cell types, as well as integration at unintended sites. In contrast, single-stranded DNA (ssDNA) donors can produce efficient HDR with minimal off-target integration. In this study, we describe the use of ssDNA phage to efficiently and inexpensively produce long circular ssDNA (cssDNA) donors. These cssDNA donors serve as efficient HDR templates when used with Cas9 or Cas12a, with integration frequencies superior to linear ssDNA (lssDNA) donors. To evaluate the relative efficiencies of imprecise and precise repair for a suite of different Cas9 or Cas12a nucleases, we have developed a modified traffic light reporter (TLR) system (TLR-multi-Cas variant 1 [MCV1]) that permits side-by-side comparisons of different nuclease systems. We used this system to assess editing and HDR efficiencies of different nuclease platforms with distinct DNA donor types. We then extended the analysis of DNA donor types to evaluate efficiencies of fluorescent tag knockins at endogenous sites in HEK293T and K562 cells. Our results show that cssDNA templates produce efficient and robust insertion of reporter tags. Targeting efficiency is high, allowing production of biallelic integrants using cssDNA donors. cssDNA donors also outcompete lssDNA donors in template-driven repair at the target site. These data demonstrate that circular donors provide an efficient, cost-effective method to achieve knockins in mammalian cell lines.

Double-strand break toxicity is chromatin context independent

Cells respond to double-strand breaks (DSBs) by activating DNA damage response pathways, including cell cycle arrest. We have previously shown that a single double-strand break generated via CRISPR/Cas9 is sufficient to delay cell cycle progression and compromise cell viability. However, we also found that the cellular response to DSBs can vary, independent of the number of lesions. This implies that not all DSBs are equally toxic, and raises the question if the location of a single double-strand break could influence its toxicity. To systematically investigate if DSB-location is a determinant of toxicity we performed a CRISPR/Cas9 screen targeting 6237 single sites in the human genome. Next, we developed a data-driven framework to design CRISPR/Cas9 sgRNA (crRNA) pools targeting specific chromatin features. The chromatin context was defined using ChromHMM states, Lamin-B1 DAM-iD, DNAseI hypersensitivity, and RNA-sequencing data. We computationally designed 6 distinct crRNA pools, each containing 10 crRNAs targeting the same chromatin state. We show that the toxicity of a DSB is highly similar across the different ChromHMM states. Rather, we find that the major determinants of toxicity of a sgRNA are cutting efficiency and off-target effects. Thus, chromatin features have little to no effect on the toxicity of a single CRISPR/Cas9-induced DSB.

Generation of corrected hiPSC clones from a Cornelia de Lange Syndrome (CdLS) patient through CRISPR-Cas-based technology

Background

Cornelia de Lange syndrome (CdLS) is a rare multisystem genetic disorder which is caused by genetic defects involving the Nipped-B-like protein (NIPBL) gene in the majority of clinical cases (60–70%). Currently, there are no specific cures available for CdLS and clinical management is needed for life. Disease models are highly needed to find a cure. Among therapeutic possibilities are genome editing strategies based on CRISPR-Cas technology.

Methods

A comparative analysis was performed to test the most recent CRISPR-Cas technologies comprising base- and prime-editors which introduce modifications without DNA cleavages and compared with sequence substitution approaches through homology directed repair (HDR) induced by Cas9 nuclease activity. The HDR method that was found more efficient was applied to repair a CdLS-causing mutation in the NIPBL gene. Human-induced pluripotent stem cells (hiPSCs) derived from a CdLS patient carrying the c.5483G > A mutation in the NIPBL were modified through HDR to generate isogenic corrected clones.

Results

This study reports an efficient method to repair the NIPBL gene through HDR mediated by CRISPR-Cas and induced with a compound (NU7441) inhibiting non-homologous end joining (NHEJ) repair. This sequence repair method allowed the generation of isogenic wild-type hiPSCs clones with regular karyotype and preserved pluripotency.

Conclusions

CdLS cellular models were generated which will facilitate the investigation of the disease molecular determinants and the identification of therapeutic targets. In particular, the hiPSC-based cellular models offer the paramount advantage to study the tissue differentiation stages which are altered in the CdLS clinical development. Importantly, the hiPSCs that were generated are isogenic thus providing the most controlled experimental set up between wild-type and mutated conditions.

Supplementary Information

The online version contains supplementary material available at 10.1186/s13287-022-03135-0.

Optimizing CRISPR/Cas9 Editing of Repetitive Single Nucleotide Variants

CRISPR/Cas9, base editors and prime editors comprise the contemporary genome editing toolbox. Many studies have optimized the use of CRISPR/Cas9, as the original CRISPR genome editing system, in substituting single nucleotides by homology directed repair (HDR), although this remains challenging. Studies describing modifications that improve editing efficiency fall short of isolating clonal cell lines or have not been validated for challenging loci or cell models. We present data from 95 transfections using a colony forming and an immortalized cell line comparing the effect on editing efficiency of donor template modifications, concentration of components, HDR enhancing agents and cold shock. We found that in silico predictions of guide RNA efficiency correlated poorly withactivity in cells. Using NGS and ddPCR we detected editing efficiencies of 5–12% in the transfected populations which fell to 1% on clonal cell line isolation. Our data demonstrate the variability of CRISPR efficiency by cell model, target locus and other factors. Successful genome editing requires a comparison of systems and modifications to develop the optimal protocol for the cell model and locus. We describe the steps in this process in a flowchart for those embarking on genome editing using any system and incorporate validated HDR-boosting modifications for those using CRISPR/Cas9.

Molecular and Computational Strategies to Increase the Efficiency of CRISPR-Based Techniques

The prokaryote-derived Clustered Regularly Interspaced Palindromic Repeats (CRISPR)/Cas mediated gene editing tools have revolutionized our ability to precisely manipulate specific genome sequences in plants and animals. The simplicity, precision, affordability, and robustness of this technology have allowed a myriad of genomes from a diverse group of plant species to be successfully edited. Even though CRISPR/Cas, base editing, and prime editing technologies have been rapidly adopted and implemented in plants, their editing efficiency rate and specificity varies greatly. In this review, we provide a critical overview of the recent advances in CRISPR/Cas9-derived technologies and their implications on enhancing editing efficiency. We highlight the major efforts of engineering Cas9, Cas12a, Cas12b, and Cas12f proteins aiming to improve their efficiencies. We also provide a perspective on the global future of agriculturally based products using DNA-free CRISPR/Cas techniques. The improvement of CRISPR-based technologies efficiency will enable the implementation of genome editing tools in a variety of crop plants, as well as accelerate progress in basic research and molecular breeding.

Challenges of CRISPR-Based Gene Editing in Primary T Cells

Adaptive T-cell immunotherapy holds great promise for the successful treatment of leukemia, as well as other types of cancers. More recently, it was also shown to be an effective treatment option for chronic virus infections in immunosuppressed patients. Autologous or allogeneic T cells used for immunotherapy are usually genetically modified to express novel T-cell or chimeric antigen receptors. The production of such cells was significantly simplified with the CRISPR/Cas system, allowing for the deletion or insertion of novel genes at specific locations within the genome. In this review, we describe recent methodological breakthroughs that were important for the conduction of these genetic modifications, summarize crucial points to be considered when conducting such experiments, and highlight the potential pitfalls of these approaches.