Cas9 Ribonucleoprotein Complexes Research Articles

Bone morphogenetic protein 15 gene disruption affects the in vitro maturation of porcine oocytes by impairing spindle assembly and organelle function

Bone morphogenetic protein 15 (BMP15) plays a crucial role in the porcine follicular development. However, its exact functions in the in vitro maturation (IVM) of porcine oocytes remain largely unknown. Here, through cytoplasmic injection of a preassembled crRNA–tracrRNA–Cas9 ribonucleoprotein complex, we achieved BMP15 disruption in approximately 54 % of the cultured porcine oocytes. Editing BMP15 impaired the IVM of porcine oocytes, as indicated by the significantly increased abnormal spindle assembly and reduced first polar body (PB1) extrusion. The editing also impaired cytoplasmic maturation of porcine oocytes, as reflected by reduced abundant of Golgi apparatus and impaired functions of mitochondria. The impaired IVM of porcine oocytes by editing BMP15 possibly was associated with the attenuated SMAD1/5 and EGFR-ERK1/2 signaling in the cumulus granulosa cells (CGCs) and the inhibited MOS/ERK1/2 signaling in oocytes. The attenuated MOS/ERK1/2 signaling may contribute to the inactivation of maturation promoting factor (MPF) and the increased abnormal spindle assembly, leading to reduced PB1 extrusion. It also may contribute to reduced Golgi apparatus formation, and impaired functions of mitochondria. These findings expand our understanding of the regulatory role of BMP15 in the IVM of porcine oocytes and provide a basis for manipulation of porcine reproductive performance.

Near-Infrared Light Activated Formulation for the Spatially Controlled Release of CRISPR-Cas9 Ribonucleoprotein for Brain Gene Editing.

The CRISPR/Cas9 system has emerged as a promising platform for gene editing; however, the lack of an efficient and safe delivery system to introduce it into cells continues to hinder clinical translation. Here, we report a rationally designed gene-editing nanoparticle (NP) formulation for brain applications: an sgRNA:Cas9 ribonucleoprotein complex is immobilized on the NP surface by oligonucleotides that are complementary to the sgRNA. Irradiation of the formulation with a near-infrared (NIR) laser generates heat in the NP, leading to the release of the ribonucleoprotein complex. The gene-editing potential of the formulation was demonstrated in vitro at the single-cell level. The safety and gene editing of the formulation were also demonstrated in the brains of reporter mice, specifically in the subventricular zone after intracerebral administration and in the olfactory bulb after intranasal administration. The formulation presented here offers a new strategy for the spatially controlled delivery of the CRISPR system to the brain.

Near‐Infrared Light Activated Formulation for the Spatially Controlled Release of CRISPR‐Cas9 Ribonucleoprotein for Brain Gene Editing

AbstractThe CRISPR/Cas9 system has emerged as a promising platform for gene editing; however, the lack of an efficient and safe delivery system to introduce it into cells continues to hinder clinical translation. Here, we report a rationally designed gene‐editing nanoparticle (NP) formulation for brain applications: an sgRNA:Cas9 ribonucleoprotein complex is immobilized on the NP surface by oligonucleotides that are complementary to the sgRNA. Irradiation of the formulation with a near‐infrared (NIR) laser generates heat in the NP, leading to the release of the ribonucleoprotein complex. The gene‐editing potential of the formulation was demonstrated in vitro at the single‐cell level. The safety and gene editing of the formulation were also demonstrated in the brains of reporter mice, specifically in the subventricular zone after intracerebral administration and in the olfactory bulb after intranasal administration. The formulation presented here offers a new strategy for the spatially controlled delivery of the CRISPR system to the brain.

Comparative analysis of lipid Nanoparticle-Mediated delivery of CRISPR-Cas9 RNP versus mRNA/sgRNA for gene editing in vitro and in vivo

The discovery that the bacterial defense mechanism, CRISPR-Cas9, can be reprogrammed as a gene editing tool has revolutionized the field of gene editing. CRISPR-Cas9 can introduce a double-strand break at a specific targeted site within the genome. Subsequent intracellular repair mechanisms repair the double strand break that can either lead to gene knock-out (via the non-homologous end-joining pathway) or specific gene correction in the presence of a DNA template via homology-directed repair. With the latter, pathological mutations can be cut out and repaired. Advances are being made to utilize CRISPR-Cas9 in patients by incorporating its components into non-viral delivery vehicles that will protect them from premature degradation and deliver them to the targeted tissues. Herein, CRISPR-Cas9 can be delivered in the form of three different cargos: plasmid DNA, RNA or a ribonucleoprotein complex (RNP). We and others have recently shown that Cas9 RNP can be efficiently formulated in lipid-nanoparticles (LNP) leading to functional delivery in vitro. In this study, we compared LNP encapsulating the mRNA Cas9, sgRNA and HDR template against LNP containing Cas9-RNP and HDR template. Former showed smaller particle sizes, better protection against degrading enzymes and higher gene editing efficiencies on both reporter HEK293T cells and HEPA 1–6 cells in in vitro assays. Both formulations were additionally tested in female Ai9 mice on biodistribution and gene editing efficiency after systemic administration. LNP delivering mRNA Cas9 were retained mainly in the liver, with LNP delivering Cas9-RNPs additionally found in the spleen and lungs. Finally, gene editing in mice could only be concluded for LNP delivering mRNA Cas9 and sgRNA. These LNPs resulted in 60 % gene knock-out in hepatocytes. Delivery of mRNA Cas9 as cargo format was thereby concluded to surpass Cas9-RNP for application of CRISPR-Cas9 for gene editing in vitro and in vivo.

Efficient Editing of the CXCR4 Locus Using Cas9 Ribonucleoprotein Complexes Stabilized with Polyglutamic Acid.

Gene editing using the CRISPR/Cas9 system provides new opportunities to treat human diseases. Approaches aimed at increasing the efficiency of genome editing are therefore important to develop. To increase the level of editing of the CXCR4 locus, which is a target for gene therapy of HIV infection, the Cas9 protein was modified by introducing additional NLS signals and ribonucleoprotein complexes of Cas9 and guide RNA were stabilized with poly-L-glutamic acid. The approach allowed a 1.8-fold increase in the level of CXCR4 knockout in the CEM/R5 T cell line and a 2-fold increase in the level of knock-in of the HIV-1 fusion peptide inhibitor MT-C34 in primary CD4+ T lymphocytes.

The strategy of knock-in with homology-directed genome editing in the model ornamental plant Petunia using CRISPR/Cas9 ribonucleoprotein complex

The clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated system 9 (Cas9) ribonucleoprotein (RNP) complex offers numerous advantages, including increased genome editing efficiency, reduced off-target effects, minimized risk of genomic integration, expanded applicability, and the ability to address specific ethical concerns. These benefits establish the CRISPR/Cas9 RNP complex as a valuable technique for precise genome editing in plant research and applications. Nevertheless, achieving precise knock-in using CRISPR/Cas9-mediated precision knock-in of genes by homology-directed repair (HDR) remains challenging due to the infrequent occurrence of HDR and the limited availability of DNA repair templates near double-strand breaks (DSBs). In this study, we presented the efficient utilization strategy of HDR-mediated gene knock-in through co-infection of the CRISPR/Cas9 RNP complex and donor plasmid in Petunia protoplast. We accomplish this by designing a donor plasmid targeting the chalcone synthase (CHS) in Petunia, consisting of a CRISPR RNA (crRNA)-PAM sequences flanked by a donor DNA cassette with homology arms. These findings offer insights into the application of these techniques in ornamental crops to enhance CRISPR/Cas9 RNA complex-mediated gene knock-in strategies.

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].

Gene knock-outs in human CD34+ hematopoietic stem and progenitor cells and in the human immune system of mice.

Human CD34+ hematopoietic stem and progenitor cells (HSPCs) are a standard source of cells for clinical HSC transplantations as well as experimental xenotransplantation to generate "humanized mice". To further extend the range of applications of these humanized mice, we developed a protocol to efficiently edit the genomes of human CD34+ HSPCs before transplantation. In the past, manipulating HSPCs has been complicated by the fact that they are inherently difficult to transduce with lentivectors, and rapidly lose their stemness and engraftment potential during in vitro culture. However, with optimized nucleofection of sgRNA:Cas9 ribonucleoprotein complexes, we are now able to edit a candidate gene in CD34+ HSPCs with almost 100% efficiency, and transplant these modified cells in immunodeficient mice with high engraftment levels and multilineage hematopoietic differentiation. The result is a humanized mouse from which we knocked out a gene of interest from their human immune system.

An optimized SpCas9 high-fidelity variant for direct protein delivery

Electroporation of the Cas9 ribonucleoprotein (RNP) complex offers the advantage of preventing off-target cleavages and potential immune responses produced by long-term expression of the nuclease. Nevertheless, the majority of engineered high-fidelity Streptococcus pyogenes Cas9 (SpCas9) variants are less active than the wild-type enzyme and are not compatible with RNP delivery. Building on our previous studies on evoCas9, we developed a high-fidelity SpCas9 variant suitable for RNP delivery. The editing efficacy and precision of the recombinant high-fidelity Cas9 (rCas9HF), characterized by the K526D substitution, was compared with the R691A mutant (HiFi Cas9), which is currently the only available high-fidelity Cas9 that can be used as an RNP. The comparative analysis was extended to gene substitution experiments where the two high fidelities were used in combination with a DNA donor template, generating different ratios of non-homologous end joining (NHEJ) versus homology-directed repair (HDR) for precise editing. The analyses revealed a heterogeneous efficacy and precision indicating different targeting capabilities between the two variants throughout the genome. The development of rCas9HF, characterized by an editing profile diverse from the currently used HiFi Cas9 in RNP electroporation, increases the genome editing solutions for the highest precision and efficient applications.

Cellular Delivery of Large Functional Proteins and Protein-Nucleic Acid Constructs via Localized Electroporation.

Delivery of proteins and protein-nucleic acid constructs into live cells enables a wide range of applications from gene editing to cell-based therapies and intracellular sensing. However, electroporation-based protein delivery remains challenging due to the large sizes of proteins, their low surface charge, and susceptibility to conformational changes that result in loss of function. Here, we use a nanochannel-based localized electroporation platform with multiplexing capabilities to optimize the intracellular delivery of large proteins (β-galactosidase, 472 kDa, 75.38% efficiency), protein-nucleic acid conjugates (protein spherical nucleic acids (ProSNA), 668 kDa, 80.25% efficiency), and Cas9-ribonucleoprotein complex (160 kDa, ∼60% knock-out and ∼24% knock-in) while retaining functionality post-delivery. Importantly, we delivered the largest protein to date using a localized electroporation platform and showed a nearly 2-fold improvement in gene editing efficiencies compared to previous reports. Furthermore, using confocal microscopy, we observed enhanced cytosolic delivery of ProSNAs, which may expand opportunities for detection and therapy.

Double-gene targeting with preassembled Cas9 ribonucleoprotein for safe genome editing in the edible mushroom Pleurotus ostreatus.

CRISPR/Cas9 has potential for efficient molecular breeding. Recently, a foreign-DNA-free gene-targeting technology was established by introducing a preassembled Cas9 ribonucleoprotein (RNP) complex into the oyster mushroom Pleurotus ostreatus. However, the target gene was restricted to such a gene like pyrG, since screening of a genome-edited strain was indispensable and could be performed via examination of 5-fluoroorotic acid (5-FOA) resistance caused by the disruption of the target gene. In this study, we simultaneously introduced the Cas9 RNP complex targeting fcy1, a mutation that conferred P. ostreatus resistance to 5-fluorocytosine (5-FC), together with that targeting pyrG. A total of 76 5-FOA resistant strains were isolated during the first screening. Subsequently, a 5-FC resistance examination was conducted, and three strains exhibited resistance. Genomic PCR experiments followed by DNA sequencing revealed that mutations were successfully introduced into fcy1 and pyrG in the three strains. The results indicated that double gene-edited mutants could be obtained in one experiment employing 5-FOA resistance screening for strains with Cas9 RNP incorporation. This work may pave the way for safe CRISPR/Cas9 technology to isolate mutant strains in any gene of interest without an ectopic marker gene.

CRISPR-Cas9-Based Functional Analysis in Amphibians: Xenopus laevis, Xenopus tropicalis, and Pleurodeles waltl.

Amphibians have made many fundamental contributions to our knowledge, from basic biology to biomedical research on human diseases. Current genome editing tools based on the CRISPR-Cas system enable us to perform gene functional analysis in vivo, even in non-model organisms. We introduce here a highly efficient and easy protocol for gene knockout, which can be used in three different amphibians seamlessly: Xenopus laevis, Xenopus tropicalis, and Pleurodeles waltl. As it utilizes Cas9 ribonucleoprotein complex (RNP) for injection, this cloning-free method enables researchers to obtain founder embryos with a nearly complete knockout phenotype within a week. To evaluate somatic mutation rate and its correlation to the phenotype of a Cas9 RNP-injected embryo (crispant), we also present accurate and cost-effective genotyping methods using pooled amplicon-sequencing and a user-friendly web-based tool.

Efficient editing BMP15 in porcine oocytes through microinjection of CRISPR ctRNP

Bone morphogenetic protein 15 (BMP15) is an X-linked gene encoding an oocyte secreted factor, which plays varied functions in the female fertility between mono-ovulatory and poly-ovulatory mammalian species. We previously found that knockout of BMP15 completely blocked porcine follicular development at preantral stages. However, the specific function of BMP15 on porcine oocytes in vitro maturation remains largely unknown. Here, we injected the pre-assembled crRNA + tracrRNA + Cas9 ribonucleoprotein (ctRNP) complex into the cytoplasm of germinal vesicle stage porcine oocytes to disrupt BMP15. The ctRNP composed of Cas9 nuclease and crRNA-tracrRNA complex at 1.2/1 content ratio. The tested crRNA-tracrRNA complex concentration ranging from 50 to 200 ng/μL, all presented effective editing of BMP15 in porcine oocytes, and the 125 ng/μL crRNA-tracrRNA complex presented the highest editing efficiency (39.23 ± 3.33%). Surprisingly, we found approximately 95% edited oocytes presented monoallelic mutations, and only 5% edited oocytes harbored biallelic mutations. Interestingly, the coinjected two crRNAs guided the ctRNP complex to concurrently cut within a 10 bp window of the PAM (protospacer adjacent motif), resulting in a precise deletion within BMP15 in 85.9% edited oocytes, and additional deletion happened in 14.1% edited oocytes, which resulted in large fragment deletions in BMP15. Most deletions caused frameshift and introduced premature stop codon in BMP15, resulting in the disruption of BMP15 protein expression, which was confirmed by the Western blot analysis showing the reduced BMP15 protein expression in ctRNP injected oocytes. The disruption of BMP15 attenuated the activation of SMAD1/5/8 signaling, and impaired cumulus expansion of porcine cumulus cell-oocyte complexes (COCs). Our study proved that delivering CRISPR ctRNP into porcine oocytes by microinjection was able to edit BMP15 efficiently, providing a new strategy to investigate the functions of oocyte-specific secreted factors in oocyte in vitro maturation.

High-Throughput and Efficient Intracellular Delivery Method via a Vibration-Assisted Nanoneedle/Microfluidic Composite System.

Intracellular delivery and genetic modification have brought a significant revolutionary to tumor immunotherapy, yet existing methods are still limited by low delivery efficiency, poor throughput, excessive cell damage, or unsuitability for suspension immune cells, specifically the natural killer cell, which is highly resistant to transfection. Here, we proposed a vibration-assisted nanoneedle/microfluidic composite system that uses large-area nanoneedles to rapidly puncture and detach the fast-moving suspension cells in the microchannel under vibration to achieve continuous high-throughput intracellular delivery. The nanoneedle arrays fabricated based on the large-area self-assembly technique and microchannels can maximize the delivery efficiency. Cas9 ribonucleoprotein complexes (Cas9/RNPs) can be delivered directly into cells due to the sufficient cellular membrane nanoperforation size; for difficult-to-transfect immune cells, the delivery efficiency can be up to 98%, while the cell viability remains at about 80%. Moreover, the throughput is demonstrated to maintain a mL/min level, which is significantly higher than that of conventional delivery techniques. Further, we prepared CD96 knockout NK-92 cells via this platform, and the gene-edited NK-92 cells possessed higher immunity by reversing exhaustion. The high-throughput, high-efficiency, and low-damage performance of our intracellular delivery strategy has great potential for cellular immunotherapy in clinical applications.

Highly Efficient One-Step Tagging of Endogenous Genes in Primary Cells Using CRISPR-Cas Ribonucleoproteins.

Genome editing tools have simplified the generation of knock-in gene fusions, which are widely used to study proteins in their natural context. However, strategies for tagging endogenous genes in primary cells are few and inefficient. In this study, we developed a one-step endogenous gene-tagging strategy by co-delivery of clustered regularly interspaced short palindromic repeats (CRISPR)-Cas9 ribonucleoprotein complexes and chemically modified donor DNA into cells. Upon CRISPR-Cas9 blunt-end double-strand breaks, highly efficient site-specific insertion of genetic materials (3 × FLAG or eGFP) was achieved in both cell lines and primary cells. We further optimized the gene-tagging efficiency and precision by using CRISPR-Cas12a, which produces a staggered cut with a 5' overhang and thus enables precise ligation of DNA donors with a complementary 3' overhang. With high efficiency and flexibility, this platform would be extremely useful for multiplex endogenous genes tagging and further exploration of protein functions in various cell types.

Blockade of Ceramide-Dependent Extracellular Vesicle Secretion Improves Hematopoietic Stem Cell Transplantation

Hematopoietic stem cell (HSC) transplantation serves as a curative therapy for numerous benign and malignant hematopoietic diseases, and as a platform for HSC gene therapy. A main constraint is the limited quantity of HSCs collected from patients or donors and the inherent risk of graft failure, or oligocolonal reconstitution, especially after ex vivo manipulation. Current efforts to mitigate these limitations have predominantly focused on the discovery of extrinsic factors to maintain or expand hematopoietic stem and progenitor cells (HSPCs), whereas the role of the HSPC secretome in modulating niche function and improving occupancy have not been extensively studied. Recently, classes of nanometer-sized extracellular vesicles (EVs) have been shown to regulate tissue function by providing potent proangiogenic signals and remodeling cellular microenvironments. EV secretion additionally regulates homeostasis and stemness properties of HSPCs; however, in-depth studies into HSPC EV biogenesis and the impact on homeostatic and regenerative potency have been hindered by numerous technical challenges. Here, we utilize an optimized ex vivo expansion protocol for human and murine HSPCs to examine the impact of different EV biogenesis pathways on vesicle secretion. Our studies rapidly focused on ceramide-dependent vesicle secretion; blockade of ceramide biosynthesis with a widely used neutral sphingomyelinase inhibitor, GW4869 significantly reduced HSPC EV secretion. Treatment of HSPCs with GW4869 further inhibited mTOR signaling, altered autophagic flux, and decreased cell cycle entry. Increased ratios of lymphoid (MPP4):myeloid (MPP2/3) primed progenitors accumulated in culture. Upon competitive transplantation into myeloablated recipients, GW4869-exposed murine HSPCs demonstrated superior long-term engraftment potential (>8 steady-state fold change, p < 0.001), independent of bone marrow or splenic homing. Ceramide inhibition did not significantly alter long-term lineage output from transplanted HSPCs. Confirmatory genetic knockout (KO) of neutral sphingomyelinase (nSMase2) in murine and human HSPCs using Cas9 ribonucleoprotein complex delivery similarly demonstrated enhanced colony forming potential. To understand the impact of ceramide inhibition on vesicle cargo, comparative mass spectrometry was performed on HSPC nSMase2 KO EVs (EVnSmase2-KO). Nearly 900 proteins were depleted (fold change <2) in EVnSmase2-KO; these underrepresented proteins were noted to be involved in myeloid differentiation and activation, immune response, and exocytosis, suggesting vesicle modulation of these processes. These results suggest that ceramide inhibition not only leads to a reduction in EV release, but also impacts selective trafficking of EV cargo, ultimately altering HSPC function. While other EV biogenesis pathways have been demonstrated to promote HSC-self renewal, these findings implicate ceramide-dependent EV secretion as a novel negative regulator of HSPC repopulation. Transient disruption of ceramide-dependent EV secretion results in a more quiescent and less metabolically active HSPC pool, and additionally prevents bias toward a myeloid-primed progenitor pool seen with ex vivo expansion. These effects culminate in dramatically improved repopulation potential of HSPCs. Furthermore, unlike other small molecular inhibitors proposed to enhance engraftment of HSPCs, including rapamycin, no significant hinderance of ex vivo expansion was seen with GW4869 treatment. We therefore propose broadened exploration of ceramide inhibition for ex vivo maintenance of HSPCs prior to transplantation.

AML-144 Uroporphyrinogen Decarboxylase (UROD) Is a Therapeutically Targetable Dependency in Acute Erythroid Leukemia

Acute erythroid leukemia (AEL) is a disease of erythroid lineage that commonly exhibits TP53 mutations, a complex karyotype, and poor prognosis irrespective of currently available therapies. To identify AEL dependencies and novel therapeutic targets in AEL using genome-wide CRISPR/Cas9 knockout screens in mouse models of AEL. Analysis of the screens performed using the Brie library identified the Urod gene encoding uroporphyrinogen decarboxylase as a critical dependency in two Trp53/Bcor mutated mouse AEL cell lines. UROD is a cytosolic enzyme in the heme biosynthesis pathway involved in converting cytosolic uroporphyrinogen III into the key heme biosynthetic intermediate, coproporphyrinogen III. In an analysis of RNA-seq data from over 2,000 acute leukemia samples, UROD was significantly overexpressed and active by a data-driven network inference algorithm in AEL compared to other leukemia subtypes. We validated UROD as a dependency by electroporation of Cas9 and synthetic sgRNA ribonucleoprotein complex in the two screened cell lines, additional mouse leukemia cell lines, and two commercially available human AEL cell lines. Targeted knockout of UROD resulted in decreased tumor cell growth and viability compared to the non-targeting guide control cells. Sequencing analysis showed high editing efficiency (>85%) followed by recovery of the wildtype cells, indicating that UROD is a dependency in these cells. These results correlated with a significant increase of necrotic cells preceding loss of editing in the two tested human cell lines. We also observed an increase in ALAS1 expression, which is negatively regulated by cellular heme, indicating that UROD knockout leukemic cells cannot generate heme, leading to necrotic cell death. Moreover, mitochondrial function assays showed significant decreases in basal respiration, ATP production, maximal respiration, and spare capacity in a time-dependent manner, followed by the rescue of mitochondrial function after loss of editing efficiency and re-expression of UROD. We have recently developed homozygous UROD-dTAG engineered human AEL cell lines to perform in-vivo survival studies. Through whole-genome CRISPR knockout screening and functional studies, we show that UROD is a dependency in AEL and provides a basis for exploring its therapeutic inhibition.

A Simple and Efficient CRISPR/Cas9 System Using a Ribonucleoprotein Method for Flammulina filiformis.

CRISPR/Cas9 systems were established in some edible fungi based on in vivo expressed Cas9 and guide RNA. Compared with those systems, the in vitro assembled Cas9 and sgRNA ribonucleoprotein complexes (RNPs) have more advantages, but only a few examples were reported, and the editing efficiency is relatively low. In this study, we developed and optimized a CRISPR/Cas9 genome-editing method based on in vitro assembled ribonucleoprotein complexes in the mushroom Flammulina filiformis. The surfactant Triton X-100 played a critical role in the optimal method, and the targeting efficiency of the genomic editing reached 100% on a selective medium containing 5-FOA. This study is the first to use an RNP complex delivery to establish a CRISPR/Cas9 genome-editing system in F. filiformis. Moreover, compared with other methods, this method avoids the use of any foreign DNA, thus saving time and labor when it comes to plasmid construction.

Exosome-mediated delivery of Cas9 ribonucleoprotein complexes for tissue-specific gene therapy of liver diseases.

CRISPR-Cas9 gene editing has emerged as a powerful therapeutic technology, but the lack of safe and efficient in vivo delivery systems, especially for tissue-specific vectors, limits its broad clinical applications. Delivery of Cas9 ribonucleoprotein (RNP) owns competitive advantages over other options; however, the large size of RNPs exceeds the loading capacity of currently available delivery vectors. Here, we report a previously unidentified genome editing delivery system, named exosomeRNP, in which Cas9 RNPs were loaded into purified exosomes isolated from hepatic stellate cells through electroporation. ExosomeRNP facilitated effective cytosolic delivery of RNP in vitro while specifically accumulated in the liver tissue in vivo. ExosomeRNP showed vigorous therapeutic potential in acute liver injury, chronic liver fibrosis, and hepatocellular carcinoma mouse models via targeting p53 up-regulated modulator of apoptosis (PUMA), cyclin E1 (CcnE1), and K (lysine) acetyltransferase 5 (KAT5), respectively. The developed exosomeRNP provides a feasible platform for precise and tissue-specific gene therapies of liver diseases.

Modulating CRISPR-Cas Genome Editing Using Guide-Complementary DNA Oligonucleotides.

Clustered regularly interspaced short palindromic repeats and CRISPR-associated proteins (CRISPR-Cas) has revolutionized genome editing and has great potential for many applications, such as correcting human genetic disorders. To increase the safety of genome editing applications, CRISPR-Cas may benefit from strict control over Cas enzyme activity. Previously, anti-CRISPR proteins and designed oligonucleotides have been proposed to modulate CRISPR-Cas activity. In this study, we report on the potential of guide-complementary DNA oligonucleotides as controlled inhibitors of Cas9 ribonucleoprotein complexes. First, we show that DNA oligonucleotides inhibit Cas9 activity in human cells, reducing both on- and off-target cleavage. We then used in vitro assays to better understand how inhibition is achieved and under which conditions. Two factors were found to be important for robust inhibition: the length of the complementary region and the presence of a protospacer adjacent motif-loop on the inhibitor. We conclude that DNA oligonucleotides can be used to effectively inhibit Cas9 activity both ex vivo and in vitro.