Highly efficient RNA-guided genome editing in human cells via delivery of purified Cas9 ribonucleoproteins

Genome editing in maize and sorghum: A comprehensive review of CRISPR/Cas9 and emerging technologies

The increasing changes in the climate patterns across the globe have deeply affected food systems where unparalleled and unmatched challenges are created. This jeopardizes food security due to an ever-increasing population. The extreme efficiency of C4 crops as compared to C3 crops makes them incredibly significant in securing food safety. C4 crops, maize (Zea mays L.) and sorghum (Sorghum bicolor L. Moench) in particular, have the ability to withstand osmotic stress induced by oxidative stress. Osmotic stress causes a series of physical changes in a plant thus facilitating reduced water uptake and photosynthesis inhibition, such as membrane tension, cell wall stiffness, and turgor changes. There has been a great advancement in plant breeding brought by introduction of clustered regularly interspaced short palindromic repeats (CRISPR) gene editing technology. This technology offers precise alterations to an organism's DNA through targeting specific genes for desired traits in a wide number of crop species. Despite its immense opportunities in plant breeding, it faces limitations such as effective delivery systems, editing efficiency, regulatory concerns, and off-target effects. Future prospects lie in optimizing next-generation techniques, such as prime editing, and developing novel genotype-independent delivery methods. Overall, the transformative role of CRISPR/Cas9 in sorghum and maize breeding underscores the need for responsible and sustainable utilization to address global food security challenges.

Lipid Nanoparticles for Delivery of CRISPR Gene Editing Components

Gene editing has emerged as a promising therapeutic option for treating genetic diseases. However, a central challenge in the field is the safe and efficient delivery of these large editing tools, especially in vivo. Lipid nanoparticles (LNPs) are attractive nonviral vectors due to their low immunogenicity and high delivery efficiency. To maximize editing efficiency, LNPs should efficiently protect gene editing components against multiple biological barriers and release them into the cytoplasm of target cells. In this review, the widely used CRISPR gene editing systems are first overviewed. Then, each component of LNPs, as well as their effects on delivery, are systematically discussed. Following this, the current LNP engineering strategies to achieve non-liver targeting are summarized. Finally, preclinical and clinical applications of LNPs for in vivo genome editing are highlighted, and perspectives for the future development of LNPs are provided.

Application of CRISPR/Cas gene editing for infectious disease control in poultry

The poultry industry faces multifaceted challenges, including escalating demand for poultry products, climate change impacting feed availability, emergence of novel avian pathogens, and antimicrobial resistance. Traditional disease control measures are costly and not always effective, prompting the need for complementary methods. Gene editing (GE, also called genome editing) technologies, particularly CRISPR/Cas9, offer promising solutions. This article summarizes recent advancements in utilizing CRISPR/Cas GE to enhance infectious disease control in poultry. It begins with an overview of modern GE techniques, highlighting CRISPR/Cas9's advantages over other methods. The potential applications of CRISPR/Cas in poultry infectious disease prevention and control are explored, including the engineering of innovative vaccines, the generation of disease-resilient birds, and in vivo pathogen targeting. Additionally, insights are provided regarding regulatory frameworks and future perspectives in this rapidly evolving field.

Modulating binding affinity of aptamer-based loading constructs enhances extracellular vesicle-mediated CRISPR/Cas9 delivery

The CRISPR/Cas9 toolbox consists of modular nucleases that can be employed to efficiently modify genomic sequences with high specificity. However, delivery of the large Cas9-sgRNA ribonucleoprotein (RNP) complexes remains challenging due to their immunogenicity, size, and overall negative charge. An approach to overcome these limitations is the use of extracellular vesicles (EVs) as intracellular delivery vehicles. EVs exhibit the natural ability to carry and deliver RNA and proteins across biological barriers, and can be engineered to load and deliver a variety of biotherapeutic molecules. Previous studies have shown that efficient EV-mediated cargo delivery does not only require active loading strategies, but also benefits from strategies to release cargo from the EV membrane. Here, we load Cas9 RNP complexes into EVs by expressing sgRNAs containing MS2 aptamers (MS2-sgRNAs), alongside Cas9 and a fusion protein of CD63 and tandem MS2 coat proteins (MCPs). We demonstrate that efficient Cas9 RNP delivery can also be facilitated by modulating the binding affinity between MS2 aptamers and the MCPs. To study the effect of altering the binding affinity between the MS2 hairpin and the MCP on Cas9 RNP delivery, various mutations affecting the binding affinity were made in both the interacting MS2-hairpin and the RNA-binding domain of the MCPs. Comparing Cas9 RNP delivery of the modulated MS2-sgRNAs revealed that adapting binding affinity highly affects functional RNP delivery. Mutations resulting in high affinity did not facilitate efficient RNP delivery unless combined with a photo-inducible release strategy, showing that cargo release was a limiting factor in RNP delivery. Mutations that decreased affinity resolved this issue, resulting in Cas9 RNP delivery without the requirement of additional release strategies. However, further decreasing affinity resulted in decreased Cas9 gene-editing efficiency due to decreased levels of Cas9 RNP loading into EVs. A similar effect on functional delivery was seen after modification of the RNA-binding domain of the MCPs. Our results demonstrate that EVs are capable of functional Cas9-sgRNA complex delivery, and that modulation of binding affinity can be used to increase efficient functional delivery with non-covalent loading constructs, without the need for additional engineering strategies for cargo release.

Lipopeptide-mediated delivery of CRISPR/Cas9 ribonucleoprotein complexes for gene editing and correction

CRISPR/Cas gene editing is a highly promising technology for the treatment and even potential cure of genetic diseases. One of the major challenges for its therapeutic use is finding safe and effective vehicles for intracellular delivery of the CRISPR/Cas9 ribonucleoprotein (RNP) complex. In this study, we tested and characterized a series of novel fatty acid-modified versions of a previously reported Cas9 RNP carrier, consisting of a complex of the cell-penetrating peptide (CPP) LAH5 with Cas9 RNP and homology-directed DNA repair templates. Comparative experiments demonstrated that RNP/peptide nanocomplexes showed various improvements depending on the type of fatty acid modification. These improvements included enhanced stability in serum, improved membrane disruption capability and increased transfection efficacy. Cas9 RNP/oleic acid LAH5 peptide nanocomplexes showed the overall best performance for both gene editing and correction. Moreover, Cas9 RNP/oleic acid LAH5 nanocomplexes significantly protected the Cas9 protein cargo from enzymatic protease digestion. In addition, in vivo testing demonstrated successful gene editing after intramuscular administration. Despite the inherent barriers of the tightly organized muscle tissues, we achieved approximately 10 % gene editing in the skeletal muscle tissues when targeting the CAG-tdTomato locus in the transgenic Ai9 Cre-LoxP reporter mouse strain and 7 % gene editing when targeting the Ccr5 gene, without any observable short-term toxicity. In conclusion, the oleic acid-modified LAH5 peptide is an effective delivery platform for direct Cas9/RNP delivery, and holds great potential for the development of new CRISPR/Cas9-based therapeutic applications for the treatment of genetic diseases.

Therapeutic Application of mRNA for Genetic Diseases

While gene therapy has been at the center of an active research field for decades, messenger RNA (mRNA) has long been considered unsuited for therapeutic application due to challenges with stability, immunogenicity, and delivery. Where gene therapy focuses on providing the desired genetic code, mRNA can directly provide the instructions encoded in the corresponding gene. This review aims to explore recent advances in mRNA therapies, building on the success of mRNA COVID-19 vaccines, and extend these insights to the potential treatment of rare genetic diseases. We follow the "outside-in" trajectory of mRNA therapies from administration to intracellular function, focusing on carrier systems such as lipid nanoparticles and virus-like particles, mRNA modifications, and the potential and challenges for clinical applications. To treat rare diseases, different approaches can be envisioned, including chronic or acute delivery of mRNAs encoding functional enzymes for enzyme deficiencies and delivery of CRISPR/Cas9-based gene-editing tools for gene correction. These different approaches determine safety and immunological considerations. By exploring genetic, technical, and therapeutic aspects, this review highlights the potential and current challenges of mRNA therapies to address the large unmet needs in rare genetic disorders.

Weaponizing CRISPR/Cas9 for selective elimination of cells with an aberrant genome

The CRISPR/Cas9 technology is a powerful and versatile tool to disrupt genes' functions by introducing sequence-specific DNA double-strand breaks (DSBs). Here, we repurpose this technology to eradicate aberrant cells by specifically targeting silent and non-functional genomic sequences present only in target cells to be eliminated. Indeed, an intrinsic challenge of most current therapies against cancer and viral infections is the non-specific toxicity that they can induce in normal tissues because of their impact on important cellular mechanisms shared, to different extents, between unhealthy and healthy cells. The CRISPR/Cas9 technology has potential to overcome this limitation; however, so far effectiveness of these approaches was made dependent on the targeting and inactivation of a functional gene product. Here, we generate proof-of-principle evidence by engineering HeLa and RKO cells with a promoterless Green Fluorescent Protein (GFP) construct. The integration of this construct simulates either a genomic alteration, as in cancer cells, or a silent proviral genome. Cas9-mediated DSBs in the GFP sequence activate the DNA damage response (DDR), reduce cell viability and increase mortality. This is associated with increased cell size, multinucleation, cGAS-positive micronuclei accumulation and the activation of an inflammatory response. Pharmacological inhibition of the DNA repair factor DNA-PK enhances cell death. These results demonstrate the therapeutic potential of the CRISPR/Cas9 system in eliminating cells with an aberrant genome, regardless of the expression or the function of the target DNA sequence.

Efficient in vivo labeling of endogenous proteins with SMART delineates retina cellular and synaptic organization

A key application of CRISPR/Cas9-based genomic editing is modification of genes to introduce engineered sequences. However, the editing flexibility is severely constrained by the requirement for targeting sites in proximity to the desired modification site, which makes many modifications intractable. Here, we develop a strategy that overcomes this key limitation to allow CRISPR-based editing at any position with high efficiency. It relies on reconstructing the targeted gene using Silently Mutate And Repair Template (SMART) where we mutate the gap sequence in the repair template to prevent its base pairing with the target DNA while maintaining the same amino acid coding. Using vertebrate retina as a neuronal model system we document the application of SMART editing for labeling endogenous proteins in vivo with high efficiency. We show that SMART editing allows us to access numerous cell types in the retina and address fundamental cell biological questions pertaining to its organization. We propose that this approach will facilitate functional genomic studies in a wide range of systems and increase the precision of corrective gene therapies.

Gene Editing in Ganoderma lucidum: Development, Challenges, and Future Prospects

As an emerging and innovative technology, gene-editing technology has been widely applied in crop breeding, human disease treatment, animal model research, drug and vaccine development, and microbial engineering. We mainly introduce the development of gene-editing technology, the application of clustered regularly interspaced short palindromic repeat/Cas9 (CRISPR/Cas9) in Ganoderma lucidum breeding, the current challenges and optimization strategies in the use of gene-editing technology in Ganoderma breeding, as well as the current status of gene-editing technology in Ganoderma breeding. Finally, the future research directions and innovative strategies that gene editing may explore in Ganoderma breeding are prospects given the existing background, future research directions, and innovative strategies that gene editing may explore in Ganoderma breeding prospects.

Fibromodulin selectively accelerates myofibroblast apoptosis in cutaneous wounds by enhancing interleukin 1β signaling

Activated myofibroblasts deposit extracellular matrix material to facilitate rapid wound closure that can heal scarlessly during fetal development. However, adult myofibroblasts exhibit a relatively long life and persistent function, resulting in scarring. Thus, understanding how fetal and adult tissue regeneration differs may serve to identify factors that promote more optimal wound healing in adults with little or less scarring. We previously found that matricellular proteoglycan fibromodulin is one such factor promoting more optimal repair, but the underlying molecular and cellular mechanisms for these effects have not been fully elucidated. Here, we find that fibromodulin induces myofibroblast apoptosis after wound closure to reduce scarring in small and large animal models. Mechanistically, fibromodulin accelerates and prolongs the formation of the interleukin 1β-interleukin 1 receptor type 1-interleukin 1 receptor accessory protein ternary complex to increase the apoptosis of myofibroblasts and keloid- and hypertrophic scar-derived cells. As the persistence of myofibroblasts during tissue regeneration is a key cause of fibrosis in most organs, fibromodulin represents a promising, broad-spectrum anti-fibrotic therapeutic.

Therapeutic promise of CRISPR-Cas9 gene editing in sickle cell disease and β-thalassemia: A current review

Sickle cell disease (SCD and β-thalassemia (BT) affects millions of people worldwide. In addition, around 500,000 infants are born with SCD and 60,000 people are diagnosed with BT every year. Mutations in the hemoglobin subunit beta (HBB) gene are responsible for causing both BT and SCD. Indeed, the diversity of potential mutations in the HBB gene elucidates the diversity in clinical severity observed in individuals with BT and related morbidities. On the other hand, SCD takes place because of the alteration in a single amino acid at position 6 in the beta-globin chain, where a base substitution occurs from glutamic acid to valine, which eventually results in abnormal sickle hemoglobin. Conventional therapies for BT and SCD including pharmaceutical drugs and blood transfusion might temporarily improve the clinical severity of these diseases, however these therapies cannot cure the diseases. CRISPR-Cas9 (CC9) is revolutionizing genome engineering, offering promising therapeutic avenues for genetic diseases. Therefore, CC9-mediated gene therapy provides great hope in the treatment of both BT and SCD. CC9-mediated gene therapy has already demonstrated its effectiveness in correcting both SCD and BT-causing mutations. Moreover, CC9-mediated gene editing was found to be effective in reactivating the expression of hemoglobin F (HbF) and regulating LRF and BCL11A. A number of clinical trials with CC9 gene-edited therapies are being carried out to elucidate their potential in treating BT and SCD. Genetics and pathophysiological mechanisms of SCD and BT, the mechanism of CC9-mediated gene editing, and common delivery methods of the CC9 system have been discussed in this review. Moreover, an in-depth discussion on applications and the current status of CC9-mediated gene editing in SCD and BT along with current challenges and future perspectives have been provided.

An innovative approach using CRISPR-ribonucleoprotein packaged in virus-like particles to generate genetically engineered mouse models

Genetically engineered mouse models (GEMMs) are crucial for investigating disease mechanisms, developing therapeutic strategies, and advancing fundamental biological research. While CRISPR gene editing has greatly facilitated the creation of these models, existing techniques still present technical challenges and efficiency limitations. Here, we establish a CRISPR-VLP-induced targeted mutagenesis (CRISPR-VIM) strategy, enabling precise genome editing by co-culturing zygotes with virus-like particle (VLP)-delivered gene editing ribonucleoproteins (RNPs) without requiring physical manipulation or causing cellular damage. We generate Plin1- and Tyr-knockout mice through VLP-based SpCas9 or adenine base editor (ABE)/sgRNA RNPs and characterize their phenotype and germline transmission. Additionally, we demonstrate cytosine base editor (CBE)/sgRNA-based C-to-T substitution or SpCas9/sgRNA-based knock-in using VLPs. This method further simplifies and accelerates GEMM generation without specialized techniques or equipment. Consequently, the CRISPR-VIM method can facilitate mouse modeling and be applied in various research fields.

Studying the cellular efficacy and tolerability of using CRISPR-gRNA ribonucleoprotein (RNP) complex for in-vitro knockdown of TRIB2 in acute myeloid leukaemia (AML) cells as preliminary clinical evaluations

BACKGROUND

Acute Myeloid Leukemia (AML) is a complex, genetically driven cancer disease with several mutations that complicate therapy regimes. The Tribbles gene family, specifically the TRIB2 gene, has garnered substantial interest as a crucial oncogenic factor of AML progression. Studying cancer through gene expression studies and focusing on the primary oncogenes provide accurate information for future therapies. Here, CRISPR stands as the most exceptional tool used to explore both oncogene functionalities and therapeutic utilities. The ribonucleoprotein (RNP) mode of CRISPR formation and usage are preferred compared to plasmid-mediated CRISPR systems, however, it can lead to complications post-transfection to sensitive cellular entities such as human cancer cells in-vitro, transcribing to similar outcomes in-vivo as well. Therefore, this study describes the use of in-house designed CRISPR-RNP systems targeting the TRIB2 oncogene and evaluates their post-transfection cellular safety and efficacy aspects for future clinical applications.

METHODS

This study uses a designed guide RNA targeting the TRIB2 gene assembled via In-vitro synthesis. The gRNA with Cas9 protein leads to the formation of CRISPR-RNP structures, which target and cleave the TRIB2 gene. The assembled CRISPR-RNP system is transfected into target AML cell and control cell lines (i.e. HEK cells), and the subsequent gene cleavage and resulting changes to the AML cells in terms of cellular safety/tolerability and gene knockdown efficacy were studied via RT-qPCR, flow cytometry, and cell viability analysis.

RESULTS

The outcome demonstrates the well-tolerated transfection of the in-vitro assembled CRISPR RNP system with no signs of cellular toxicity and disruptions towards the AML cell's metabolic activities, promoting the safety aspects of CRISPR RNP post-transfection to human cells. The study further highlights the in-vitro efficacy of the CRISPR RNP in targeting the TRIB2 oncogene, where a statistically significant gene knockdown of more than 80% was detected via qPCR analysis of TRIB2 gene expression with minimal to no background effects from individual RNP components, equating to their targeted gene cleavage effects. In addition, the CRISPR TRIB2 gene knockdown also indicated the possibilities of induced AML cell death measured via flow cytometry markers, translating to favourable outcomes in eliminating cancerous growths.

CONCLUSIONS

This study research contributes to the refinement of the CRISPR-RNP strategy and preliminary evaluation for future clinical uses.

Dasatinib-resistant universal CAR-T cells proliferate in the presence of host immune cells and exhibit antitumor activity

The universal chimeric antigen receptor T cell (UCAR-T) immunotherapy derived from healthy donors holds great promise in pan-cancer treatment. However, UCAR-T cell therapy faces a challenge in the rapid elimination of allogeneic cells by the host immune system. To address this, we introduced a T316I mutation in the leukocyte-specific protein tyrosine kinase (LCK) locus in CAR-T cells using the cytosine base editor (CBE) system. Concurrently, we disrupted endogenous T cell receptor alpha chain (TRAC) and beta-2 microglobulin (B2M) with the CRISPR-Cas9 system, along with dasatinib to overcome host immune rejection, an Src family kinase (SFK) inhibitor. The resulting LCK mutated UCAR-T (KM UCAR-T) cells exhibited normal phenotypes in activation, proliferation, differentiation, and tumor cytotoxicity in vitro. Moreover, KM UCAR-T cells demonstrated sustained expansion in mixed lymphocyte reactions (MLR) when incubated with T cells or peripheral blood mononuclear cells (PBMCs) from HLA-mismatched donors upon dasatinib treatment. Additionally, we illustrated that KM UCAR-T cells displayed antitumor activity in a xenograft murine model and verified the expansion and cytotoxicity of KM UCAR-T over traditional UCAR-T in the presence of allogeneic PBMCs when treated with dasatinib in vivo. These findings offer a novel strategy for UCAR-T cells to resist host immune rejection and achieve sustained expansion.

CRISPR-Cas9-driven antigen conversion of clinically relevant blood group systems

The common practice of blood transfusion entirely relies on blood donations from the population. Ensuring blood group compatibility between a donor and a recipient is paramount to prevent critical adverse reactions. Finding compatible blood can be challenging given the high diversity of blood group antigens, especially for chronically transfused patients at higher risk of alloimmunization owing to repeated exposures to foreign RBCs. In addition, due to the immunogenicity of the ABO blood group and the highly polymorphic nature of the Rhesus (Rh) system, they both remain of prime importance in transfusion medicine. Cultured red blood cells (cRBCs) may eventually provide an alternative for blood donations-at least in some circumstances. Combining cRBCs with blood group gene editing could broaden transfusion accessibility by making antigen expression compatible with rare phenotypes, thus meeting the needs of more patients. Starting from mobilized, erythroid-primed hematopoietic stem and progenitor cells (HSPCs), we used virus- and selection-free, CRISPR-Cas9-mediated knockouts to produce erythroid cells devoid of AB and Rh antigen. The approach yielded almost complete conversion to O- and RhNull phenotypes, as determined by standard hemagglutination and flow cytometry analyses. Combined with robust cRBC protocols, these clinically relevant phenotypic changes could eventually expand the accessibility of blood transfusion for specific and unmet clinical needs.

TOPO-seq reveals DNA topology-induced off-target activity by Cas9 and base editors

With the increasing use of CRISPR-Cas9, detecting off-target events is essential for safety. Current methods primarily focus on guide RNA (gRNA) sequence mismatches, often overlooking the impact of DNA topology in regulating off-target activity. Here we present TOPO-seq, a high-throughput and sensitive method that identifies genome-wide off-target effects of Cas9 and base editors while accounting for DNA topology. TOPO-seq revealed that topology-induced off-target sites frequently harbor higher mismatches than the relaxed DNA sequence, with over 50% of off-target sites containing six mismatches, which are usually overlooked using previous methods. Applying TOPO-seq to three therapeutic gRNAs in hematopoietic stem cells identified 47 bona fide off-target loci, six of which are specifically induced by DNA topology. These findings highlight DNA topology as a regulator of off-target editing rates, establish TOPO-seq as a robust method for capturing DNA topology-induced off-target events and underscore its importance in off-target detection for developing safe genome-editing therapies.

Hairpin Internal Nuclear Localization Signals in CRISPR-Cas9 Enhance Editing in Primary Human Lymphocytes

The incorporation of nuclear localization signal (NLS) sequences at one or both termini of CRISPR enzymes is a widely adopted strategy to facilitate genome editing. Engineered variants of CRISPR enzymes with diverse NLS sequences have demonstrated superior performance, promoting nuclear localization and efficient DNA editing. However, limiting NLS fusion to the CRISPR protein's termini can negatively impact protein yield via recombinant expression. Here we present a distinct strategy involving the installation of hairpin internal NLS sequences (hiNLS) at rationally selected sites within the backbone of CRISPR-Cas9. We evaluated the performance of these hiNLS Cas9 variants by editing genes in human primary T cells following the delivery of ribonucleoprotein enzymes via either electroporation or co-incubation with amphiphilic peptides. We show that hiNLS Cas9 variants can improve editing efficiency in T cells compared with constructs with terminally fused NLS sequences. Furthermore, many hiNLS Cas9 constructs can be produced with high purity and yield, even when these constructs contain as many as nine NLS. These hiNLS Cas9 constructs represent a key advance in optimizing CRISPR effector design and may contribute to improved editing outcomes in research and therapeutic applications.

Novel application of ribonucleoprotein-mediated CRISPR-Cas9 gene editing in plant pathogenic oomycete species

CRISPR-Cas9 gene editing has become an important tool for the study of plant pathogens, allowing researchers to functionally characterize specific genes involved in phytopathogenicity, virulence, and fungicide resistance. Protocols for CRISPR-Cas9 gene editing have already been developed for Phytophthoras, an important group of oomycete plant pathogens; however, these efforts have exclusively focused on agricultural pathosystems, with research lacking for forest pathosystems. We sought to develop CRISPR-Cas9 gene editing in two forest pathogenic Phytophthoras, Phytophthora cactorum and P. ramorum, using a plasmid-ribonucleoprotein (RNP) co-transformation approach. Our gene target in both species was the ortholog of PcORP1, which encodes an oxysterol-binding protein that is the target of the fungicide oxathiapiprolin in the agricultural pathogen P. capsici. We delivered liposome complexes, each containing plasmid DNA and CRISPR-Cas9 RNPs, to Phytophthora protoplasts using a polyethylene glycol-mediated transformation protocol. We obtained two ORP1 mutants in P. cactorum but were unable to obtain any mutants in P. ramorum. The two P. cactorum mutants exhibited decreased resistance to oxathiapiprolin, as measured by their radial growth relative to wild-type cultures on oxathiapiprolin-supplemented medium. Our results demonstrate the potential for RNP-mediated CRISPR-Cas9 gene editing in P. cactorum and provide a foundation for future optimization of our protocol in other forest pathogenic Phytophthora species.IMPORTANCECRISPR-Cas9 gene editing has become a valuable tool for characterizing the genetics driving virulence and pathogenicity in plant pathogens. CRISPR-Cas9 protocols are now well-established in several Phytophthora species, an oomycete genus with significant economic and ecological impact globally. These protocols, however, have been developed for agricultural Phytophthora pathogens only; CRISPR-Cas9 systems have not yet been developed for any forest pathogenic Phytophthoras. In this study, we sought to establish CRISPR-Cas9 gene editing in two forest Phytophthora pathogens that cause widespread tree mortality: P. cactorum and P. ramorum. We successfully obtained gene mutations in P. cactorum and demonstrated a decrease in fungicide resistance, a trait that could impact the pathogen's ability to cause disease. However, the same protocol did not yield any mutants in P. ramorum. The results of our study will serve as a baseline for the development of CRISPR-Cas9 gene editing in forest Phytophthoras and other oomycetes.

The mosaicism of Cas-induced mutations and pleiotropic effects of scarlet gene in an emerging model system

The effective use of CRISPR technologies in emerging model organisms faces significant challenges in efficiently generating heritable mutations and in understanding the genomic consequences of induced DNA damages and the inheritance patterns of induced mutations. This study addresses these issues by 1) developing an efficient microinjection delivery method for gene editing in the microcrustacean Daphnia pulex; 2) assessing the editing dynamics of Cas9 and Cas12a nucleases in the scarlet knock-out mutants; and 3) investigating the transcriptomes of scarlet mutants to understand the pleiotropic effects of scarlet gene. Our reengineered microinjection method results in efficient biallelic editing with both nucleases. Our data suggest site-specific DNA cleavage mostly occurs in a stepwise fashion. Indels dominate the induced mutations. A few, unexpected on-site large deletions (>1 kb) are also observed. Notably, genome-wide analyses reveal no off-target mutations. Knock-in of a stop codon cassette to the scarlet locus was successful, despite complex induced mutations surrounding the target site. Moreover, extensive germline mosaicism exists in some mutants, which unexpectedly produce different phenotypes/genotypes in their asexual progeny. Lastly, our transcriptomic analyses unveil significant gene expression changes associated with scarlet knock-out and altered swimming behavior in mutants, including several genes involved in human neurodegenerative diseases.

Proteogenomic discovery of RB1-defective phenocopy in cancer predicts disease outcome, response to treatment, and therapeutic targets

Genomic defects caused by truncating mutations or deletions in the Retinoblastoma tumor suppressor gene (RB1) are frequently observed in many cancer types leading to dysregulation of the RB pathway. Here, we propose an integrative proteogenomic approach that predicts cancers with dysregulation in the RB pathway. A subset of these cancers, which we term as "RBness," lack RB1 genomic defects and yet phenocopy the transcriptional profile of RB1-defective cancers. We report RBness as a pan-cancer phenomenon, associated with patient outcome and chemotherapy response in multiple cancer types, and predictive of CDK4/6 inhibitor response in estrogen-positive breast cancer. Using RNA interference and a CRISPR-Cas9 screen in isogenic models, we find that RBness cancers also phenocopy synthetic lethal vulnerabilities of cells with RB1 genomic defects. In summary, our findings suggest that dysregulation of the RB pathway in cancers lacking RB1 genomic defects provides a molecular rationale for how these cancers could be treated.

Optimizing in vitro Transcribed CRISPR-Cas9 Single-Guide RNA Libraries for Improved Uniformity and Affordability

We describe a scalable and cost-effective sgRNA synthesis workflow that reduces costs by over 70% through the use of large pools of microarray-derived oligos encoding unique sgRNA spacers. These subpool oligos are assembled into full-length dsDNA templates via Golden Gate Assembly before in vitro transcription with T7 RNA polymerase. RNA-seq analysis reveals severe biases in spacer representation, with some spacers being highly overrepresented while others are completely absent. Consistent with previous studies, we identify guanine-rich sequences within the first four nucleotides of the spacer, immediately downstream of the T7 promoter, as the primary driver of this bias. To address this issue, we introduced a guanine tetramer upstream of all spacers, which reduced bias by an average of 19% in sgRNA libraries containing 389 spacers. However, this modification also increased the presence of high-molecular-weight RNA species after transcription. We also tested two alternative bias-reduction strategies: compartmentalizing spacers within emulsions and optimizing DNA input and reaction volumes. Both methods independently reduced bias in 2,626-plex sgRNA libraries, though to a lesser extent than the guanine tetramer approach. These advancements enhance both the affordability and uniformity of sgRNA libraries, with broad implications for improving CRISPR-Cas9 screens and optimizing guide RNA design for other CRISPR and nuclease systems.

Recent advances in therapeutic gene-editing technologies

The advent of gene-editing technologies, particularly CRISPR-based systems, has revolutionized the landscape of biomedical research and gene therapy. Ongoing research in gene editing has led to the rapid iteration of CRISPR technologies, such as base and prime editors, enabling precise nucleotide changes without the need for generating harmful double-strand breaks (DSBs). Furthermore, innovations such as CRISPR fusion systems with DNA recombinases, DNA polymerases, and DNA ligases have expanded the size limitations for edited sequences, opening new avenues for therapeutic development. Beyond the CRISPR system, mobile genetic elements (MGEs) and epigenetic editors are emerging as efficient alternatives for precise large insertions or stable gene manipulation in mammalian cells. These advances collectively set the stage for next-generation gene therapy development. This review highlights recent developments of genetic and epigenetic editing tools and explores preclinical innovations poised to advance the field.

Efficient DNA- and virus-free engineering of cellular transcriptomic states using dCas9 ribonucleoprotein (dRNP) complexes

For genome editing, the use of CRISPR ribonucleoprotein (RNP) complexes is well established and often the superior choice over plasmid-based or viral strategies. RNPs containing dCas9 fusion proteins, which enable the targeted manipulation of transcriptomes and epigenomes, remain significantly less accessible. Here, we describe the production, delivery, and optimization of second generation CRISPRa RNPs (dRNPs). We characterize the transcriptional and cellular consequences of dRNP treatments in a variety of human target cells and show that the uptake is very efficient. The targeted activation of genes demonstrates remarkable potency, even for genes that are strongly silenced, such as developmental master transcription factors. In contrast to DNA-based CRISPRa strategies, gene activation is immediate and characterized by a sharp temporal precision. We also show that dRNPs allow very high-target multiplexing, enabling undiminished gene activation of multiple genes simultaneously. Applying these insights, we find that intensive target multiplexing at single promoters synergistically elevates gene transcription. Finally, we demonstrate in human stem and differentiated cells that the preferable features of dRNPs allow to instruct and convert cell fates efficiently without the need for DNA delivery or viral vectors.

Modeling and correction of protein conformational disease in iPSC-derived neurons through personalized base editing

Altered protein conformation can cause incurable neurodegenerative disorders. Mutations in SERPINI1, the gene encoding neuroserpin, can alter protein conformation resulting in cytotoxic aggregation leading to neuronal death. Familial encephalopathy with neuroserpin inclusion bodies (FENIB) is a rare autosomal dominant progressive myoclonic epilepsy that progresses to dementia and premature death. We developed HEK293T and induced pluripotent stem cell (iPSC) models of FENIB, harboring a patient-specific pathogenic SERPINI1 variant or stably overexpressing mutant neuroserpin fused to GFP (MUT NS-GFP). Here, we utilized a personalized adenine base editor (ABE)-mediated approach to correct the pathogenic variant efficiently and precisely to restore neuronal dendritic morphology. ABE-treated MUT NS-GFP cells demonstrated reduced inclusion size and number. Using an inducible MUT NS-GFP neuron system, we identified early prevention of toxic protein expression allowed aggregate clearance, while late prevention halted further aggregation. To address several challenges for clinical applications of gene correction, we developed a neuron-specific engineered virus-like particle to optimize neuronal ABE delivery, resulting in higher correction efficiency. Our findings provide a targeted strategy that may treat FENIB and potentially other neurodegenerative diseases due to altered protein conformation such as Alzheimer's and Huntington's diseases.

Focused ultrasound and microbubble-mediated delivery of CRISPR-Cas9 ribonucleoprotein to human induced pluripotent stem cells

CRISPR-Cas9 ribonucleoproteins (RNPs) have been heavily considered for gene therapy due to their high on-target efficiency, rapid activity, and lack of insertional mutagenesis relative to other CRISPR-Cas9 delivery formats. Genetic diseases such as hypertrophic cardiomyopathy currently lack effective treatment strategies and are prime targets for CRISPR-Cas9 gene editing technology. However, current in vivo delivery strategies for Cas9 pose risks of unwanted immunogenic responses. This proof-of-concept study aimed to demonstrate that focused ultrasound (FUS) in combination with microbubbles can be used to deliver Cas9-sgRNA (single-guide RNA) RNPs and functionally edit human induced pluripotent stem cells (hiPSCs) in vitro, a model system that can be expanded to cardiovascular research via hiPSC-derived cardiomyocytes. Here, we first determine acoustic conditions suitable for the viable delivery of large proteins to hiPSCs with clinical Definity microbubble agents using our customized experimental platform. From here, we delivered Cas9-sgRNA RNP complexes targeting the EGFP (enhanced green fluorescent protein) gene to EGFP-expressing hiPSCs for EGFP knockout. Simultaneous acoustic cavitation detection during treatment confirmed a strong correlation between microbubble disruption and viable FUS-mediated protein delivery in hiPSCs. This study shows for the first time the potential for an FUS-mediated technique for targeted and precise CRISPR-Cas9 gene editing in human stem cells.

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

Peptide-enabled ribonucleoprotein delivery for CRISPR engineering (PERC) is a new approach for ex vivo genome editing of primary human cells. PERC uses a single amphiphilic peptide reagent to mediate intracellular delivery of the same pre-formed CRISPR ribonucleoprotein enzymes that are broadly used in research and therapeutics, resulting in high-efficiency editing of stimulated immune cells and cultured hematopoietic stem and progenitor cells (HSPCs). PERC facilitates nuclease-mediated gene knockout, precise transgene knock-in and base editing. The protocol involves mixing the CRISPR ribonucleoprotein enzyme with peptide and then incubating with cultured cells. For efficient transgene knock-in, adeno-associated virus (AAV) homology-directed repair template (HDRT) DNA may be included. In contrast to electroporation, PERC is appealing because it needs no dedicated hardware and has less impact on cell phenotype and viability. Because of the gentle nature of PERC, delivery can be performed multiple times without substantial impact to cell health or phenotype. Editing efficiencies can surpass 90% when using either Cas9 or Cas12a in primary T cells or HSPCs. After 3 h dedicated to reagent preparation, the PERC delivery step can be completed in 1 h, with the associated cell culture steps taking 3-7 d total. Because the protocol calls for only three readily available reagents (protein, RNA and peptide) and does not require dedicated hardware for any step, PERC demands no special expertise and is exceptionally straightforward to adopt. The inherent compatibility of PERC with established cell engineering pipelines makes the protocol appealing for rapid deployment in research and clinical settings.

CRISPR-Cas9 genome editing reveals that the Pgs gene of Fusarium circinatum is involved in pathogenicity, growth and sporulation

Fusarium circinatum, the causal agent of pine pitch canker, is one of the most destructive pathogens of Pinus species worldwide. Infections by this pathogen result in serious mortality of seedlings due to root and root collar disease, and growth reduction in trees due to canker formation and dieback. Although much is known about the population biology, genetics, and genomics of F. circinatum, relatively little is known regarding the molecular basis of pathogenicity in F. circinatum. In this study, a protoplast-based transformation using CRISPR-Cas9-mediated genome editing was utilized to functionally characterize a putative pathogenicity gene in three different strains of the fungus. In silico analyses suggested the gene likely encodes a small secreted protein, and all isolates in which it was deleted displayed significantly reduced vegetative growth and asexual spore production compared to the wild-type isolates. In pathogenicity tests, lesions induced by the deletion mutants on detached Pinus patula branches were significantly shorter than those produced by the wild-types. The putative pathogenicity gene was named Pgs reflecting its role in pathogenicity, growth, and sporulation. Future research will seek to explore the molecular mechanisms underlying the mutant phenotypes observed. Overall, this study represents a significant advance in F. circinatum research as the development and application of a Cas9-mediated gene deletion process opens new avenues for functional gene characterization underlying many of the pathogen's biological traits.

The transcriptomic architecture of common cancers reflects synthetic lethal interactions

To maintain cell fitness, deleterious genetic alterations are buffered by compensatory changes in additional genes. In cancer, buffering processes could be targeted by synthetic lethality. However, despite the large-scale identification of synthetic lethal effects in preclinical models, evidence that these operate clinically is limited. This impedes the application of synthetic lethal approaches. By integrating molecular profiling data from >9,000 cancers with synthetic lethal screens, we show that transcriptomic buffering of tumor suppressor gene (TSG) loss by hyperexpression of synthetic lethal partners is a common phenomenon, extending to multiple TSGs and histotypes. Transcriptomic buffering is also notable in cancers that phenocopy TSG loss, such as BRCAness cancers, where expression of BRCA1/2 synthetic lethal genes correlates with clinical outcome. Synthetic lethal genes that exhibit transcriptomic buffering also represent more robust synthetic lethal effects. These observations have implications for understanding how tumor cells tolerate TSG loss, in part explain transcriptomic architectures in cancer and provide insight into target selection.

Preparation of high-purity RNPs of CRISPR-based DNA base editors

Since their introduction, CRISPR-based DNA base editors (BEs) have become essential in the field of precision genome editing, revolutionizing the correction of pathogenic SNPs for both basic research and therapeutic applications. As this technology advances, more laboratories are implementing these tools into their workflow. The delivery of BEs as BE-guide RNA complexes (RNPs), rather than as mRNA or plasmids, has been shown to exhibit lower off-target effects, establishing it as the preferred method of delivery. However, there are no protocols describing in detail how to obtain high-purity and highly active BE RNPs. Here, we offer a comprehensive guide for the expression, purification, RNP reconstitution, and in vitro activity assessment of TadA-based BEs. The protocol includes guidance on performing activity assays using commercial denaturing gels, which is convenient and uses standard molecular biology equipment. This allows for rapid quality control testing of reconstituted BE RNPs prior to more expensive and time-consuming in vivo genome editing experiments. Overall, this protocol aims to empower more laboratories to generate tailored BE RNPs for diverse in vitro and in vivo applications.

Packaged delivery of CRISPR-Cas9 ribonucleoproteins accelerates genome editing

Effective genome editing requires a sufficient dose of CRISPR-Cas9 ribonucleoproteins (RNPs) to enter the target cell while minimizing immune responses, off-target editing, and cytotoxicity. Clinical use of Cas9 RNPs currently entails electroporation into cells ex vivo, but no systematic comparison of this method to packaged RNP delivery has been made. Here we compared two delivery strategies, electroporation and enveloped delivery vehicles (EDVs), to investigate the Cas9 dosage requirements for genome editing. Using fluorescence correlation spectroscopy, we determined that >1300 Cas9 RNPs per nucleus are typically required for productive genome editing. EDV-mediated editing was >30-fold more efficient than electroporation, and editing occurs at least 2-fold faster for EDV delivery at comparable total Cas9 RNP doses. We hypothesize that differences in efficacy between these methods result in part from the increased duration of RNP nuclear residence resulting from EDV delivery. Our results directly compare RNP delivery strategies, showing that packaged delivery could dramatically reduce the amount of CRISPR-Cas9 RNPs required for experimental or clinical genome editing.

Somatostatin receptor-targeted polymeric nanoplatform for efficient CRISPR/Cas9 gene editing to enhance synergistic hepatocellular carcinoma therapy

INTRODUCTION

The CRISPR/Cas9 system-based gene therapy can fundamentally address the issues of cancer occurrence, development, progression, and metastasis. However, the lack of targeting and effectiveness hinders gene therapy from entering clinical application. Herein, a somatostatin receptor-targeted polymeric nanoplatform is developed for the delivery of a PD-L1-targeted CRISPR/Cas9 system and synergistic treatment of hepatocellular carcinoma. This nanoplatform can effectively incorporate the CRISPR/Cas9 system and the chemotherapeutic drug paclitaxel to simultaneously address the biological safety and packaging capacity issues of viral vectors. After the octreotide-modified polymer (LNA-PEG-OCT) guided the nanoparticle into hepatoma carcinoma cells, the nanoparticle protected the CRISPR/Cas9 ribonucleoprotein complex (RNP) and achieved lysosomal escape. Then, the RNP reached the target gene (PD-L1) under the guidance of the single guide RNA (sgRNA) in the RNP. The PD-L1 gene editing efficiency reached up to 55.8% for HepG2 cells in vitro and 46.0% for tumor tissues in vivo, leading to effective suppression of PD-L1 protein expression. Substantial inhibition of hepatocellular carcinoma cell proliferation and further 79.45% growth repression against subcutaneous xenograft tumors were achieved. Overall, this somatostatin receptor-targeted polymeric nanoplatform system not only provides a promising nanocarrier for CRISPR/Cas9 system delivery, but also expands the potential of combining gene editing and chemotherapy.

Modulation of CRISPR-Cas9 Cleavage with an Oligo-Ribonucleoprotein Design

Clustered regularly interspaced short palindromic repeats (CRISPR) associated protein Cas9 system has been widely used for genome editing. However, the editing or cleavage specificity of CRISPR Cas9 remains a major concern due to the off-target effects. The existing approaches to control or modulate CRISPR Cas9 cleavage include engineering Cas9 protein and development of anti-CRISPR proteins. There are also attempts on direct modification of sgRNA, for example, structural modification via truncation or hairpin design, or chemical modification on sgRNA such as partially replacing RNA with DNA. The above-mentioned strategies rely on extensive protein engineering and direct chemical or structural modification of sgRNA. In this study, we proposed an indirect method to modulate CRISPR Cas9 cleavage without modification on Cas9 protein or sgRNA. An oligonucleotide was used to form an RNA-DNA hybrid structure with the sgRNA spacer, creating steric hindrance during the Cas9 mediated DNA cleavage process. We first introduced a simple and robust method to assemble the oligo-ribonucleoprotein (oligo-RNP). Next, the cleavage efficiency of the assembled oligo-RNP was examined using different oligo lengths in vitro. Lastly, we showed that the oligo-RNP directly delivered into cells could also modulate Cas9 activity inside cells using three model gene targets with reduced off-target effects.

Strategic base modifications refine RNA function and reduce CRISPR-Cas9 off-targets

In contrast to traditional RNA regulatory approaches that modify the 2'-OH group, this study explores strategic base modifications using 5-carboxylcytosine (ca5C). We developed a technique where ca5C is transformed into dihydrouracil via treatment with borane-pyridine complex or 2-picoline borane complex, leading to base mutations that directly impact RNA functionality. This innovative strategy effectively manages CRISPR-Cas9 system activities, significantly minimizing off-target effects. Our approach not only demonstrates a significant advancement in RNA manipulation but also offers a new method for the precise control of gene editing technologies, showcasing its potential for broad application in chemical biology.

Gene editing without ex vivo culture evades genotoxicity in human hematopoietic stem cells

Gene editing the BCL11A erythroid enhancer is a validated approach to fetal hemoglobin (HbF) induction for β-hemoglobinopathy therapy, though heterogeneity in edit allele distribution and HbF response may impact its safety and efficacy. Here, we compare combined CRISPR-Cas9 editing of the BCL11A +58 and +55 enhancers with leading gene modification approaches under clinical investigation. Dual targeting of the BCL11A +58 and +55 enhancers with 3xNLS-SpCas9 and two single guide RNAs (sgRNAs) resulted in superior HbF induction, including in sickle cell disease (SCD) patient xenografts, attributable to simultaneous disruption of core half E-box/GATA motifs at both enhancers. Unintended on-target outcomes of double-strand break (DSB) repair in hematopoietic stem and progenitor cells (HSPCs), such as long deletions and centromere-distal chromosome fragment loss, are a byproduct of cellular proliferation stimulated by ex vivo culture. Editing quiescent HSPCs bypasses long deletion and micronuclei formation and preserves efficient on-target editing and engraftment function.

Nature Inspired Delivery Vehicles for CRISPR-Based Genome Editing

The advent of Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)-based genome editing technologies has opened up groundbreaking possibilities for treating a wide spectrum of genetic disorders and diseases. However, the success of these technologies relies heavily on the development of efficient and safe delivery systems. Among the most promising approaches are natural and synthetic nanocarrier-mediated delivery systems, including viral vectors, extracellular vesicles (EVs), engineered cellular membrane particles, liposomes, and various nanoparticles. These carriers enhance the efficacy of the CRISPR system by providing a unique combination of efficiency, specificity, and reduced immunogenicity. Synthetic carriers such as liposomes and nanoparticles facilitate CRISPR delivery with high reproducibility and customizable functions. Viral vectors, renowned for their high transduction efficiency and broad tropism, serve as powerful vehicles for delivering CRISPR components to various cell types. EVs, as natural carriers of RNA and proteins, offer a stealth mechanism to evade immune detection, allowing for the targeted delivery of genome editors with minimal off-target effects. Engineered cellular membrane particles further improve delivery by simulating the cellular environment, enhancing uptake, and minimizing immune response. This review explores the innovative integration of CRISPR genome editors with various nanocarrier systems, focusing on recent advancements, applications, and future directions in therapeutic genome editing.

β2-tubulin and its promoter in the brown planthopper: A versatile tool for genetic control strategies

At present, the application of CRISPR/Cas9 technology for genetic manipulation in insects is predominantly concentrated on Diptera model species, including Drosophila and mosquitoes. In contrast, non-model insects such as the brown planthoppers (BPH, Nilaparvata lugens), a major insect pest of rice, have received less attention in genetic manipulation due to insufficient tools. Here, the analysis of spatiotemporal expression patterns revealed that β2-tubulin in BPH (NlB2t) was predominantly concentrated in male adults and male testis, exhibiting high expression levels. Knockdown of NlB2t expression by using RNAi resulted in the obstruction of male testis development. Mating between the RNAi-treated males and wild-type females led to a notable reduction in the number of eggs laid and the hatching rate of those eggs by 58.2% and 50.6%, respectively. The longevity of RNAi males significantly increased, and females that had previously mated with RNAi males exhibited a diminished inclination for re-mating with wild-type males. The dual-luciferase reporter assay demonstrated robust promoter activity in the upstream 943 bp of NlB2t, capable of driving Cas9 protein expression in vivo and effectively inducing target gene knockout. These findings elucidated that NlB2t may be a key gene in BPH male testis development and reproduction, as a promising target for sterilization. Its upstream promoter serves as a germline promoter, significantly facilitating the development of genetic control tools based on CRISPR/Cas9 technology in BPH.

Gene therapy for genetic diseases: challenges and future directions

Genetic diseases constitute the majority of rare human diseases, resulting from abnormalities in an individual's genetic composition. Traditional treatments offer limited relief for these challenging conditions. In contrast, the rapid advancement of gene therapy presents significant advantages by directly addressing the underlying causes of genetic diseases, thereby providing the potential for precision treatment and the possibility of curing these disorders. This review aims to delineate the mechanisms and outcomes of current gene therapy approaches in clinical applications across various genetic diseases affecting different body systems. Additionally, genetic muscular disorders will be examined as a case study to investigate innovative strategies of novel therapeutic approaches, including gene replacement, gene suppression, gene supplementation, and gene editing, along with their associated advantages and limitations at both clinical and preclinical levels. Finally, this review emphasizes the existing challenges of gene therapy, such as vector packaging limitations, immunotoxicity, therapy specificity, and the subcellular localization and immunogenicity of therapeutic cargos, while discussing potential optimization directions for future research. Achieving delivery specificity, as well as long-term effectiveness and safety, will be crucial for the future development of gene therapies targeting genetic diseases.

Engineering Base Changes and Epitope-Tagged Alleles in Mice Using Cas9 RNA-Guided Nuclease

Mice carrying patient-associated base changes are powerful tools to define the causality of single-nucleotide variants to disease states. Epitope tags enable immuno-based studies of genes for which no antibodies are available. These alleles enable detailed and precise developmental, mechanistic, and translational research. The first step in generating these alleles is to identify within the target sequence-the orthologous sequence for base changes or the N or C terminus for epitope tags-appropriate Cas9 protospacer sequences. Subsequent steps include design and acquisition of a single-stranded oligonucleotide repair template, synthesis of a single guide RNA (sgRNA), collection of zygotes, and microinjection or electroporation of zygotes with Cas9 mRNA or protein, sgRNA, and repair template followed by screening born mice for the presence of the desired sequence change. Quality control of mouse lines includes screening for random or multicopy insertions of the repair template and, depending on sgRNA sequence, off-target sequence variation introduced by Cas9. © 2025 The Author(s). Current Protocols published by Wiley Periodicals LLC. Basic Protocol 1: Single guide RNA design and synthesis Alternate Protocol 1: Single guide RNA synthesis by primer extension and in vitro transcription Basic Protocol 2: Design of oligonucleotide repair template Basic Protocol 3: Preparation of RNA mixture for microinjection Support Protocol 1: Preparation of microinjection buffer Alternate Protocol 2: Preparation of RNP complexes for electroporation Basic Protocol 4: Collection and preparation of mouse zygotes for microinjection or electroporation Basic Protocol 5: Electroporation of Cas9 RNP into zygotes using cuvettes Alternate Protocol 3: Electroporation of Cas9 RNP into zygotes using electrode slides Basic Protocol 6: Screening and quality control of derived mice Support Protocol 2: Deconvoluting multiple sequence chromatograms with DECODR.

Synthetic Vesicle-Based Drug Delivery Systems for Oral Disease Therapy: Current Applications and Future Directions

Oral diseases such as dental caries, periodontitis, and oral cancer are prevalent and present significant challenges to global public health. Although these diseases are typically treated through procedures like dental preparation and resin filling, scaling and root planning, or surgical excision, these interventions are often not entirely effective, and postoperative drug therapy is usually required. Traditional drug treatments, however, are limited by factors such as poor drug penetration, significant side effects, and the development of drug resistance. As a result, there is a growing need for novel drug delivery systems that can enhance therapeutic efficacy, reduce side effects, and improve treatment outcomes. In recent years, drug-loaded vesicles, such as liposomes, polymersomes, and extracellular vesicles (EVs), have emerged as promising drug delivery platforms due to their high drug encapsulation efficiency, controlled release properties, and excellent biocompatibility. This review provides an in-depth examination of the characteristics, advantages, and limitations of liposomes, polymersomes, and extracellular vesicles in the context of oral disease treatment. It further explores the reasons for their advantages and limitations and discusses the specific applications, development prospects, and strategies for optimizing these vesicle-based systems for improved clinical outcomes.

Direct delivery of Cas-embedded cytosine base editors as ribonucleoprotein complexes for efficient and accurate editing of clinically relevant targets

Recently, cytosine base editors (CBEs) have emerged as a promising therapeutic tool for specific editing of single nucleotide variants and disrupting specific genes associated with disease. Despite this promise, the currently available CBEs have the significant liabilities of off-target and bystander editing activities, partly due to the mechanism by which they are delivered, causing limitations in their potential applications. In this study, we engineered optimized, soluble and stable Cas-embedded CBEs (CE_CBEs) that integrate several recent advances, which were efficiently formulated for direct delivery into cells as ribonucleoprotein (RNP) complexes. Our resulting CE_CBE RNP complexes efficiently target cytosines in TC dinucleotides with minimal off-target or bystander mutations. Delivery of additional uracil glycosylase inhibitor protein in trans further increased C-to-T editing efficiency and target purity in a dose-dependent manner, minimizing indel formation. A single electroporation was sufficient to effectively edit the therapeutically relevant locus BCL11A for sickle cell disease in hematopoietic stem and progenitor cells in a dose-dependent manner without cellular toxicity. Significantly, these CE_CBE RNPs permitted highly efficient editing and engraftment of transplanted cells in mice. Thus, our designed CBE proteins provide promising reagents for RNP-based editing at disease-related sites.

Improving the use of CRISPR/Cas9 gene editing machinery as a cancer therapeutic tool with the help of nanomedicine

CRISPR-Cas9 (clustered regularly interspaced short palindromic repeats-associated protein 9) has revolutionized gene editing tools and paved the way for innovations in medical research for disease diagnosis and treatment. However, better specificity and efficient delivery of this gene machinery make it challenging to successfully edit genes for treating various diseases. This is mainly due to cellular barriers, instability in biological environments, and various off-target effects that prohibit safe and efficient delivery under in vivo conditions. This review examines several delivery modes [plasmid, mRNA, RNP (ribonucleoprotein)] and methods for the CRISPR-Cas9 system delivery, focusing on its potential applications in cancer therapy. Biocompatibility and cytotoxicity are crucial factors determining their safe and effective use. Various nanomaterials have been reviewed for their biocompatibility, limitations, and challenges in treating cancer. Among the reviewed nanoparticles, lipid nanoparticles (LNPs) stand out for their biocompatibility due to their biomimetic lipid bilayer that effectively delivers CRISPR/Cas9 cargoes while reducing toxicity. We discuss challenges in in vivo delivery and associated findings such as encapsulation, target delivery, controlled release, and endosomal escape. Future directions involve addressing limitations and adapting CRISPR-Cas9 for clinical trials, ensuring its safe and effective use.

Rapid generation of homozygous fluorescent knock-in human cells using CRISPR-Cas9 genome editing and validation by automated imaging and digital PCR screening

We previously described a protocol for genome engineering of mammalian cultured cells with clustered regularly interspaced short palindromic repeats and associated protein 9 (CRISPR-Cas9) to generate homozygous knock-ins of fluorescent tags into endogenous genes. Here we are updating this former protocol to reflect major improvements in the workflow regarding efficiency and throughput. In brief, we have improved our method by combining high-efficiency electroporation of optimized CRISPR-Cas9 reagents, screening of single cell-derived clones by automated bright-field and fluorescence imaging, rapidly assessing the number of tagged alleles and potential off-targets using digital polymerase chain reaction (PCR) and automated data analysis. Compared with the original protocol, our current procedure (1) substantially increases the efficiency of tag integration, (2) automates the identification of clones derived from single cells with correct subcellular localization of the tagged protein and (3) provides a quantitative and high throughput assay to measure the number of on- and off-target integrations with digital PCR. The increased efficiency of the new procedure reduces the number of clones that need to be analyzed in-depth by more than tenfold and yields to more than 26% of homozygous clones in polyploid cancer cell lines in a single genome engineering round. Overall, we were able to dramatically reduce the hands-on time from 30 d to 10 d during the overall ~10 week procedure, allowing a single person to process up to five genes in parallel, assuming that validated reagents-for example, PCR primers, digital PCR assays and western blot antibodies-are available.

State of the art CRISPR-based strategies for cancer diagnostics and treatment

Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR) technology is a groundbreaking and dynamic molecular tool for DNA and RNA "surgery". CRISPR/Cas9 is the most widely applied system in oncology research. It is a major advancement in genome manipulation due to its precision, efficiency, scalability and versatility compared to previous gene editing methods. It has shown great potential not only in the targeting of oncogenes or genes coding for immune checkpoint molecules, and in engineering T cells, but also in targeting epigenomic disturbances, which contribute to cancer development and progression. It has proven useful for detecting genetic mutations, enabling the large-scale screening of genes involved in tumor onset, progression and drug resistance, and in speeding up the development of highly targeted therapies tailored to the genetic and immunological profiles of the patient's tumor. Furthermore, the recently discovered Cas12 and Cas13 systems have expanded Cas9-based editing applications, providing new opportunities in the diagnosis and treatment of cancer. In addition to traditional cis-cleavage, they exhibit trans-cleavage activity, which enables their use as sensitive and specific diagnostic tools. Diagnostic platforms like DETECTR, which employs the Cas12 enzyme, that cuts single-stranded DNA reporters, and SHERLOCK, which uses Cas12, or Cas13, that specifically target and cleave single-stranded RNA, can be exploited to speed up and advance oncological diagnostics. Overall, CRISPR platform has the great potential to improve molecular diagnostics and the functionality and safety of engineered cellular medicines. Here, we will emphasize the potentially transformative impact of CRISPR technology in the field of oncology compared to traditional treatments, diagnostic and prognostic approaches, and highlight the opportunities and challenges raised by using the newly introduced CRISPR-based systems for cancer diagnosis and therapy.

Lipid Nanoparticles Enable Efficient In Vivo DNA Knock-In via HITI-Mediated Genome Editing

In vivo genome editing holds great therapeutic potential for treating monogenic diseases by enabling precise gene correction or addition. However, improving the efficiency of delivery systems remains a key challenge. In this study, we investigated the use of lipid nanoparticles (LNPs) for in vivo knock-in of ectopic DNA. Our in vitro experiments demonstrated that the homology-independent targeted integration (HITI)-mediated genome-editing method achieved significantly higher knock-in efficiency at the Alb locus in hepatic cells compared to the traditional homology-directed repair (HDR)-mediated approach. By optimizing LNP composition and administration routes, we successfully achieved HITI-mediated GFP knock-in (2.1-2.7%) in the livers of mice through intravenous delivery of LNP-loaded genome editing components. Notably, repeated intravenous dosing led to a twofold increase in liver GFP knock-in efficiency (4.3-7.0%) compared to a single dose, highlighting the potential for cumulative genome editing effects. These findings provide a solid foundation for the use of LNPs in in vivo knock-in strategies, paving the way for future genome-editing therapies.

Combining Off-flow, a Nextflow-coded program, and whole genome sequencing reveals unintended genetic variation in CRISPR/Cas-edited iPSCs

Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)-Cas nucleases and human induced pluripotent stem cell (iPSC) technology can reveal deep insight into the genetic and molecular bases of human biology and disease. Undesired editing outcomes, both on-target (at the edited locus) and off-target (at other genomic loci) hinder the application of CRISPR-Cas nucleases. We developed Off-flow, a Nextflow-coded bioinformatic workflow that takes a specific guide sequence and Cas protein input to call four separate off-target prediction programs (CHOPCHOP, Cas-Offinder, CRISPRitz, CRISPR-Offinder) to output a comprehensive list of predicted off-target sites. We applied it to whole genome sequencing (WGS) data to investigate the occurrence of unintended effects in human iPSCs that underwent repair or insertion of disease-related variants by homology-directed repair. Off-flow identified a 3-base-pair-substitution and a mono-allelic genomic deletion at the target loci, KCNQ2, in 2 clones. Unbiased WGS analysis further identified off-target missense variants and a mono-allelic genomic deletion at the targeted locus, GNAQ, in 10 clones. On-target substitution and deletions had escaped standard PCR and Sanger sequencing analysis, while missense variants at other genomic loci were not detected by Off-flow. We used these results to filter out iPSC clones for subsequent functional experiments. Off-flow, which we make publicly available, works for human and mouse genomes currently and can be adapted for other genomes. Off-flow and WGS analysis can improve the integrity of studies using CRISPR/Cas-edited cells and animal models.

Mechanism of Genome Editing Tools and Their Application on Genetic Inheritance Disorders

In the fields of medicine and bioscience, gene editing is increasingly recognized as a promising therapeutic approach for treating pathogenic variants in humans and other living organisms. With advancements in technology and knowledge, it is now understood that most genetic defects are caused by single-base pair variants. The ability to substitute genes using genome editing tools enables scientists and doctors to cure genetic diseases and disorders. Starting with CRISPR (clustered regularly interspaced short palindromic repeats)/Cas, the technology has evolved to become more efficient and safer, leading to the development of base and prime editors. Furthermore, various approaches are used to treat genetic disorders such as hemophilia, cystic fibrosis, and Duchenne muscular dystrophy. As previously mentioned, most genetic defects leading to specific diseases are caused by single-base pair variants, which can occur at many locations in corresponding gene, potentially causing the same disease. This means that, even when using the same genome editing tool, results in terms of editing efficiency or treatment effectiveness may differ. Therefore, different approaches may need to be applied to different types of diseases. Prevalently, due to the safety of adeno-associated virus (AAV) vectors in gene therapy, most clinical trials of gene therapy are based on AAV delivery methods. However, despite their safety and nonintegration into the host genome, their limitations, such as confined capacity, dosage-dependent viral toxicity, and immunogenicity, necessitate the development of new approaches to enhance treatment effects. This review provides the structure and function of each CRISPR-based gene editing tool and focuses on introducing new approaches in gene therapy associated with improving treatment efficiency.

Efficient Genome Editing Using 'NanoMEDIC' AsCas12a-VLPs Produced with Pol II-Transcribed crRNA

Virus-like particles (VLPs) are an attractive vehicle for the delivery of Cas nuclease and guide RNA ribonucleoprotein complexes (RNPs). Most VLPs are produced by packaging SpCas9 and its sgRNA, which is expressed from the RNA polymerase III (Pol III)-transcribed U6 promoter. VLPs assemble in the cytoplasm, but U6-driven sgRNA is localized in the nucleus, which hinders the efficient formation and packaging of RNPs into VLPs. In this study, using the nuclease packaging mechanism of 'NanoMEDIC' VLPs, we produced VLPs with AsCas12a and exploited its ability to process pre-crRNA. This allowed us to direct crRNA in the cytoplasm as part of a Pol II-driven transcript where AsCas12a excised mature crRNA, thus boosting RNP incorporation into VLPs. CMV-driven crRNA increased Venus and CCR5 transgene knockout levels in 293 cells from 30% to 50-90% and raised the level of endogenous CXCR4 knockout in Jurkat T cells from 1% to 20%. Changing a single crRNA to an array of three or six identical crRNAs improved CXCR4 knockout rates by up to 60-70%. Compared to SpCas9-VLPs, the editing efficiencies of AsCas12a-VLPs were higher, regardless of promoter usage. Thus, we showed that AsCas12a and CMV-driven crRNA could be efficiently packaged into VLPs and mediate high levels of gene editing. AsCas12a-VLPs are a new and promising tool for the delivery of RNPs into mammalian cells that will allow efficient target genome editing and may be useful for gene therapy applications.

In vivo gene editing of T-cells in lymph nodes for enhanced cancer immunotherapy

Immune checkpoint blockade (ICB) therapy, while promising for cancer treatment, faces challenges like unexpected side effects and limited objective responses. Here, we develop an in vivo gene-editing strategy for improving ICB cancer therapy in a lastingly effective manner. The approach uses a conductive hydrogel-based electroporation system to enable nucleofection of programmed cell death protein 1 (PD1) targeted CRISPR-Cas9 DNAs into T-cells directly within the lymph nodes, and subsequently produces PD1-deficient T-cells to combat tumor growth, metastasis and recurrence in different melanoma models in mice. Following in vivo gene editing, animals show enhanced cellular and humoral immune responses along with multi-fold increases of effector T-cells infiltration to the solid tumors, preventing tumor recurrence and prolonging their survival. These findings provide a proof-of-concept for direct in vivo T-cell engineering via localized gene-editing for enhanced cancer immunotherapy, and also unlock the possibilities of using this method to treat more complex human diseases.

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.