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CRISPR-mediated cancer therapies: Approaches to direct tumor targeting.
CRISPR-Cas9 technologies have opened new possibilities for precision cancer treatment, addressing limitations inherent in conventional therapies such as chemotherapy and radiation. This review examines CRISPR-based strategies for direct tumor targeting, including oncogene inactivation, tumor suppressor gene reactivation, and tumor microenvironment (TME) modification. Key advances include KRASG12D inactivation via base editing, in which engineered deaminases introduce precise single-nucleotide changes without generating double-strand breaks; TP53 correction through homologous recombination, which uses a donor DNA template to repair mutant sequences at the targeted locus; and CDKN2A epigenetic remodeling using CRISPR-dCas9-TET1 demethylation, where catalytically inactive Cas9 guides the TET1 demethylase to hypermethylated promoters to restore gene expression. CRISPR screening has identified synthetic lethal interactions, such as PARP1 dependency in BRCA1-/- tumors. TME editing strategies, including modification of cancer-associated fibroblasts, demonstrate enhanced antitumor responses. Delivery challenges are being addressed through viral vectors, including adenovirus, AAV, and lentivirus. Non-viral approaches include lipid nanoparticles, gold nanoparticles, exosomes, and stimuli-responsive systems such as MMP-cleavable and hypoxia-responsive nanoparticles. Clinical trials with CRISPR-engineered T-cells (e.g., CTX130) have demonstrated remission rates in hematologic malignancies. However, significant challenges remain, including cytokine release syndrome, immunotoxicity, tumor heterogeneity, and limited delivery efficiency in solid tumors. Overcoming these barriers requires interdisciplinary innovation, ethical oversight, and technological refinement to support the safe and effective integration of CRISPR-based strategies into precision oncology.
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TIGER: A tdTomato in vivo genome-editing reporter mouse for investigating precision-editor delivery approaches.
In vivo genome editing has the potential to address many inherited and environmental disorders. However, a major hurdle for the clinical translation of genome editing is safe, efficient delivery to disease-relevant tissues. A modality-agnostic reporter animal model that facilitates rapid, precise, and quantifiable assessment of functional delivery and editing could greatly enhance the evaluation and translation of delivery technologies. Here, we present the development of the tdTomato in vivo genome-editing reporter (TIGER) mouse, a reporter strain that harbors an integrated and constitutively expressed mutated tdTomato gene in the Polr2a locus. The mutations (Q115X, Q357X) abolish fluorescence, but successful adenine base editing (ABE) or prime editing (PE) restores tdTomato fluorescence. This mouse model facilitates the tissue- and cell type-specific assessment of genome editing agent delivery. We describe several editing strategies validated in vitro and demonstrate efficient ABE and PE in vivo using viral and nonviral delivery vectors targeting four cell types within the mouse eye: the retinal pigment epithelium, photoreceptors, Müller glia, and the trabecular meshwork. We show direct editing characterization in the ocular tissues via in vivo and ex vivo two-photon confocal microscopy and verify the spectral and fluorescence lifetime properties of tdTomato reporter in other mouse tissues. Additionally, we demonstrate successful adeno-associated virus (AAV)-mediated PE of extraocular tissues, including hepatocytes, skeletal muscle, and brain neurons by intravenous injection. Thus, the TIGER mouse facilitates the direct development, comparison, and optimization of delivery platforms for efficient and productive ABE or PE broadly applicable in vivo across multiple tissues tested in this study.
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Precision genome engineering through adenine base editing in plants.
The recent development of adenine base editors (ABEs) has enabled efficient and precise A-to-G base conversions in higher eukaryotic cells. Here, we show that plant-compatible ABE systems can be successfully applied to protoplasts of Arabidopsis thaliana and Brassica napus through transient transfection, and to individual plants through Agrobacterium-mediated transformation to obtain organisms with desired phenotypes. Targeted, precise A-to-G substitutions generated a single amino acid change in the FT protein or mis-splicing of the PDS3 RNA transcript, and we could thereby obtain transgenic plants with late-flowering and albino phenotypes, respectively. Our results provide 'proof of concept' for in planta ABE applications that can lead to induced neo-functionalization or altered mRNA splicing, opening up new avenues for plant genome engineering and biotechnology.
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Molecular correction of Duchenne muscular dystrophy by splice modulation and gene editing.
Duchenne muscular dystrophy (DMD) is a currently incurable X-linked neuromuscular disorder, characterized by progressive muscle wasting and premature death, typically as a consequence of cardiac failure. DMD-causing mutations in the dystrophin gene are highly diverse, meaning that the development of a universally-applicable therapy to treat all patients is very challenging. The leading therapeutic strategy for DMD is antisense oligonucleotide-mediated splice modulation, whereby one or more specific exons are excluded from the mature dystrophin mRNA in order to correct the translation reading frame. Indeed, three exon skipping oligonucleotides have received FDA approval for use in DMD patients. Second-generation exon skipping drugs (i.e. peptide-antisense oligonucleotide conjugates) exhibit enhanced potency, and also induce dystrophin restoration in the heart. Similarly, multiple additional antisense oligonucleotide drugs targeting various exons are in clinical development in order to treat a greater proportion of DMD patient mutations. Relatively recent advances in the field of genome engineering (specifically, the development of the CRISPR/Cas system) have provided multiple promising therapeutic approaches for the RNA-directed genetic correction of DMD, including exon excision, exon reframing via the introduction of insertion/deletion mutations, disruption of splice signals to promote exon skipping, and the templated correction of point mutations by seamless homology directed repair or base editing technology. Potential limitations to the clinical translation of the splice modulation and gene editing approaches are discussed, including drug delivery, the importance of uniform dystrophin expression in corrected myofibres, safety issues (e.g. renal toxicity, viral vector immunogenicity, and off-target gene editing), and the high cost of therapy.
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Application and future perspective of CRISPR/Cas9 genome editing in fruit crops.
Fruit crops, including apple, orange, grape, banana, strawberry, watermelon, kiwifruit and tomato, not only provide essential nutrients for human life but also contribute to the major agricultural output and economic growth of many countries and regions in the world. Recent advancements in genome editing provides an unprecedented opportunity for the genetic improvement of these agronomically important fruit crops. Here, we summarize recent reports of applying CRISPR/Cas9 to fruit crops, including efforts to reduce disease susceptibility, change plant architecture or flower morphology, improve fruit quality traits, and increase fruit yield. We discuss challenges facing fruit crops as well as new improvements and platforms that could be used to facilitate genome editing in fruit crops, including dCas9-base-editing to introduce desirable alleles and heat treatment to increase editing efficiency. In addition, we highlight what we see as potentially revolutionary development ranging from transgene-free genome editing to de novo domestication of wild relatives. Without doubt, we now see only the beginning of what will eventually be possible with the use of the CRISPR/Cas9 toolkit. Efforts to communicate with the public and an emphasis on the manipulation of consumer-friendly traits will be critical to facilitate public acceptance of genetically engineered fruits with this new technology.
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Small fish, big prospects: using zebrafish to unravel the mechanisms of hereditary hearing loss.
Over the past decade, advancements in high-throughput sequencing have greatly enhanced our knowledge of the mutational signatures responsible for hereditary hearing loss. In its present state, the field has a largely uncensored view of protein coding changes in a growing number of genes that have been associated with hereditary hearing loss, and many more that have been proposed as candidate genes. Sequencing data can now be generated using methods that have become widespread and affordable. The greatest hurdles facing the field concern functional validation of uncharacterized genes and rapid application to human diseases, including hearing and balance disorders. To date, over 30 hearing-related disease models exist in zebrafish. New genome editing technologies, including CRISPR/Cas9 will accelerate the functional validation of hearing loss genes and variants in zebrafish. Here, we discuss current progress in the field and recent advances in genome editing approaches.
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Synergistic optimization enhancing the precision and efficiency of cytosine base editors in poplar.
CRISPR/Cas9 genome editing technology, particularly cytosine base editing (CBE) systems, emerges as a powerful tool for precise genomic modification in plants, offering transformative applications across agricultural and forestry research and breeding programs. However, current CBE systems in poplar exhibit low efficiency and imprecise base substitutions, and optimization of base editing systems specifically for poplar remains a significant challenge. To address these limitations, we engineer a high-efficiency poplar CBE system (hyPopCBE) by integrating the MS2-UGI system, fusing Rad51 DNA-binding domain, and modifying the nuclear localization signal. Through stepwise optimization, we develop hyPopCBE-V4, which exhibits a synergistic effect in woody plants. Compared to the original hyPopCBE-V1, hyPopCBE-V4 improves C to T editing efficiency while reducing byproducts and exhibiting a narrower editing window. The proportion of plants with clean C to T edits (without byproducts) increases from 20.93% to 40.48%, and the efficiency of clean homozygous C to T editing rises from 4.65% to 21.43%. Using hyPopCBE-V1 and its variants, we induce Pro197Leu mutation in the herbicide target gene PagALS. Poplar lines with edits in all four PagALS homologues exhibit high resistance to tribenuron and nicosulfuron. This study employs a multi-component synergistic optimization strategy that specifically enhances the efficiency and precision of CBE editing in poplar while improving synchronous editing of alleles. Through editing the herbicide resistance gene PagALS, we obtain the herbicide-resistant poplar germplasm. Our research provides a more precise and efficient CBE tool for genetic modification in poplar that can also be applied to other forestry species, demonstrating its potential for advancing forestry research and breeding programs.
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Development of a DNA double-strand break-free base editing tool in Corynebacterium glutamicum for genome editing and metabolic engineering.
As a traditional amino acid producing bacterium, Corynebacterium glutamicum is a platform strain for production of various fine chemicals. Based on the CRISPR (Clustered regularly interspaced short palindromic repeats)-Cas9 system, gene editing tools that enable base conversion in the genome of C. glutamicum have been developed. However, some problems such as genomic instability caused by DNA double-strand break (DSB) and off-target effects need to be solved. In this study, a DSB-free single nucleotide genome editing system was developed by construction of a bi-directional base conversion tool TadA-dCas9-AID. This system includes cytosine base editors (CBEs): activation-induced cytidine deaminase (AID) and adenine deaminase (ABEs): tRNA adenosine deaminase (TadA), which can specifically target the gene through a 20-nt single guide RNA (sgRNA) and achieve the base conversion of C-T, C-G and A-G in the 28-bp editing window upstream of protospacer adjacent motif. Finally, as a proof-of-concept demonstration, the system was used to construct a mutant library of zwf gene in C. glutamicum S9114 genome to improve the production of a typical nutraceutical N-acetylglucosamine (GlcNAc). The GlcNAc titer of the mutant strain K293R was increased by 31.9% to 9.1 g/L in shake flask. Here, the developed bases conversion tool TadA-dCas9-AID does not need DNA double-strand break and homologous template, and is effective for genome editing and metabolic engineering in C. glutamicum.
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One-step base editing in multiple genes by direct embryo injection for pig trait improvement.
The precise and simultaneous acquisition of multiple beneficial alleles in the genome is in great demand for the development of elite pig breeders. Cytidine base editors (CBEs) that convert C:G to T:A have emerged as powerful tools for single-nucleotide replacement. Whether CBEs can effectively mediate C-to-T substitution at multiple sites/loci for trait improvement by direct zygote injection has not been verified in large animals. Here, we determined the editing efficiency of four CBE variants in porcine embryonic fibroblast cells and embryos. The findings showed that hA3A-BE3-Y130F and hA3A-eBE-Y130F consistently resulted in increased base-editing efficiency and low toxic effects in embryonic development. Further, we verified that using a one-step approach, direct zygote microinjection of the CBE system can generate pigs harboring multiple point mutations. Our process resulted in a stop codon in CD163 and myostatin (MSTN) and introduced a beneficial allele in insulin-like growth factor-2 (IGF2). The pigs showed disrupted expression of CD163 and MSTN and increased expression of IGF2, which significantly improved growth performance and infectious disease resistance. Our approach allows immediate introduction of multiple mutations in transgene-free animals to comprehensively improve economic traits through direct embryo microinjection, providing a potential new route to produce elite pig breeders.
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Mutation Knock-in Methods Using Single-Stranded DNA and Gene Editing Tools in Zebrafish.
Introduction or knock-in of precise genomic modifications remains one of the most important applications of CRISPR/Cas9 in all model systems including zebrafish. The most widely used type of donor template containing the desired modification is single-stranded DNA (ssDNA), either in the form of single-stranded oligodeoxynucleotides (ssODN) (<150 nucleotides (nt)) or as long ssDNA (lssDNA) molecules (up to about 2000 nt). Despite the challenges posed by DNA repair after DNA double-strand breaks, knock-in of precise mutations is relatively straightforward in zebrafish. Knock-in efficiency can be enhanced by careful donor template design, using lssDNA as template or tethering the donor template DNA to the Cas9-guide RNA complex. Other point mutation methods such as base editing and prime editing are starting to be applied in zebrafish and many other model systems. However, these methods may not always be sufficiently accessible or may have limited capacity to perform all desired mutation knock-ins which are possible with ssDNA-based knock-in methods. Thus, it is likely that there will be complementarity in the technologies used for generating precise mutants. Here, we review and describe a suite of CRISPR/Cas9 knock-in procedures utilizing ssDNA as the donor template in zebrafish, point out the potential challenges and suggest possible approaches for their solution ultimately leading to successful generation of precise mutant lines.
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