DNA-editing 'pencils' probe early human development.
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CRISPR-Cas9 genome editing↗
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Inheritance of the epigenetic signature and reduced intermuscular bone phenotype acquired via DNA methylation editing of the runx2 b promoter in zebrafish.
The presence of intermuscular bones (IBs) can directly affect the economic value of aquaculture fish. Although genome editing can create IB-free fish by knocking out key IB-related genes, such as runx2b, the associated DNA sequence alterations raise food safety and health concerns, limiting its breeding applications. In this study, we used CRISPR/dCas9-mediated epigenome-editing technology targeting the runx2 b promoter in zebrafish to alter DNA methylation patterns without changing the DNA sequence. Our results showed that higher runx2 b promoter methylation patterns significantly inhibited eGFP mRNA expression levels in the recombinant plasmid. Using the CRISPR/dCas9-Dnmt7 system to enhance methylation of the zebrafish runx2b promoter, we observed a significant decrease in runx2 b mRNA expression levels in the F0 generation. The IBs in the 11 th-16 th muscle segments of the adult F0 fish were significantly shorter compared with the controls. Inbreeding of fish was used to produce F1 and F2 offspring that retained these high promoter methylation levels, along with persistent runx2b expression suppression and IB development inhibition. Transcriptome sequencing analysis suggested that increasing runx2 b promoter methylation levels may synergistically induce additional epigenetic modifications, potentially affecting the PPAR signaling pathway and FoxO transcription factor regulation, which appears to inhibit osteoblast proliferation and differentiation. Overall, this study demonstrates an innovative application of epigenetic editing technology for aquaculture breeding. By precisely regulating the expression patterns of key genes for economically important traits while preserving genomic DNA integrity, this approach provides a theoretical foundation and technical support for improving fish economic traits.
Animals↗
DNA-guided CRISPR-Cas12 for cellular RNA targeting.
Here, we present ΨDNA, a DNA-based guide that enables RNA targeting by Cas12 nucleases, overcoming the traditional reliance on RNA-guided systems. We engineer ΨDNA to mimic a CRISPR RNA (crRNA) scaffold in reverse orientation, allowing AsCas12a and Cas12i1 to recognize RNA and trigger strong single-stranded DNA trans-cleavage for sensitive detection of diverse RNA species, including 100% accurate hepatitis C virus RNA detection in clinical samples. ΨDNA also achieves 70-95% multiplex knockdown of endogenous intracellular RNA transcripts through ribosome stalling across multiple human cell lines. Mechanistic studies reveal that activity depends on a stem loop that stabilizes a catalytically competent Cas12-ΨDNA-RNA complex. Lastly, codelivery of crRNA and ΨDNA enables simultaneous DNA editing and RNA knockdown with a single effector and modular fusions of different enzymes to AsCas12a extend ΨDNA to RNase H-mediated RNA degradation and METTL3-based epitranscriptomic editing. Together, ΨDNA guides constitute an adaptable toolkit that extends Cas12 systems beyond genome editing and diagnostics to enable precise, programmable control of cellular transcriptomes and their epitranscriptomic marks.
Journal Article↗
Emerging and potential use of CRISPR in human liver disease.
CRISPR is a gene editing tool adapted from naturally occurring defense systems from bacteria. It is a technology that is revolutionizing the interrogation of gene functions in driving liver disease, especially through genetic screens and by facilitating animal knockout and knockin models. It is being used in models of liver disease to identify which genes are critical for liver pathology, especially in genetic liver disease, hepatitis, and in cancer initiation and progression. It holds tremendous promise in treating human diseases directly by editing DNA. It could disable gene function in the case of expression of a maladaptive protein, such as blocking transthyretin as a therapy for amyloidosis, or to correct gene defects, such as restoring the normal functions of liver enzymes fumarylacetoacetate hydrolase or alpha-1 antitrypsin. It is also being studied for treatment of hepatitis B infection. CRISPR is an exciting, evolving technology that is facilitating gene characterization and discovery in liver disease and holds the potential to treat liver diseases safely and permanently.
Humans↗
Ultra-high field strength electroporation enables efficient DNA transformation and genome editing in nontuberculous mycobacteria.
Efficient DNA delivery is essential for genetic manipulation of mycobacteria and for dissecting their physiology, pathogenesis, and drug resistance. Although electroporation enables transformation efficiencies exceeding 10⁵ CFU per µg DNA in Mycobacterium smegmatis and Mycobacterium tuberculosis, it remains highly inefficient in many nontuberculous mycobacteria (NTM), including Mycobacterium abscessus. Here, we discovered that NTM such as M. abscessus exhibit exceptional tolerance to ultra-high electric field strengths and that hypertonic preconditioning partially protects cells from electroporation-induced damage. Using ultra-high electric field strength (3 kV/mm) electroporation, we achieved dramatic improvements in plasmid transformation efficiency-up to 106-fold in M. abscessus, 83-fold in Mycobacterium marinum, and 37-fold in Mycobacterium kansasii-compared to standard conditions (1.25  kV/mm). Transformation efficiency was further influenced by the choice of selectable marker. Ultra-high field strength electroporation also markedly enhanced allelic exchange in M. abscessus expressing Che9c RecET recombinases, increasing the recovery of gene deletion mutants by over 1,000-fold relative to conventional electroporation. In parallel, oligonucleotide-mediated recombineering for targeted point mutations produced nearly 10,000-fold more mutants under ultra-high field conditions. Together, these findings establish ultra-high field electroporation as a robust, broadly applicable platform for genetic engineering of NTMs. This method substantially enhances transformation efficiency and enables construction of advanced genetic tools-including expression libraries and CRISPRi knockdown libraries-in species that have historically resisted genetic manipulation.IMPORTANCEInfections caused by nontuberculous mycobacteria (NTM), including Mycobacterium abscessus, are increasing globally, yet genetic manipulation of these pathogens remains technically challenging due to inefficient DNA delivery and low gene editing success. The ultra-high electric field strength electroporation strategy described here overcomes these barriers, enabling dramatic improvements in both transformation and genome editing efficiency. This advance paves the way for high-throughput functional genomics in NTMs, including the construction of genome-wide knockout, CRISPRi knockdown, and expression libraries. Broad adoption of this approach will accelerate discovery of genetic determinants of virulence and drug resistance, facilitating the development of antimicrobials and vaccines.
Electroporation↗
Reducing competition between msd and genomic DNA improves retron editing efficiency.
Retrons, found in bacteria and used for defense against phages, generate a unique molecule known as multicopy single-stranded DNA (msDNA). This msDNA mimics Okazaki fragments during DNA replication, making it a promising tool for targeted gene editing in prokaryotes. However, existing retron systems often exhibit suboptimal editing efficiency. Here, we identify the msd gene in Escherichia coli, which encodes the noncoding RNA template for msDNA synthesis and carries the homologous sequence of the target gene to be edited, as a critical bottleneck. Sequence homology causes the msDNA to bind to the msd gene, thereby reducing its efficiency in editing the target gene. To address this issue, we engineer a retron system that tailors msDNA to the leading strand of the plasmid containing the msd gene. This strategy minimizes msd gene editing and reduces competition with target genes, significantly increasing msDNA availability. Our optimized system achieves very high retron editing efficiency, enhancing performance and expanding the potential for in vivo techniques that rely on homologous DNA synthesis.
Gene Editing↗
Editing Approaches to Treat Alpha-1 Antitrypsin Deficiency.
TOPIC IMPORTANCE: Alpha-1 antitrypsin (AAT) deficiency is a genetic disorder most commonly due to a single G to A point mutation (E342K), leading to debilitating lung and/or liver disorders and is associated with increased mortality. The E342K point mutation causes a conformational change of the AAT protein resulting in its retention in liver hepatocytes. This reduces AAT secretion into the serum resulting in higher protease activities due to the lack of inhibition from AAT, causing damage to healthy lung tissue. The current standard of care for lung manifestations involves weekly IV augmentation therapy and is considered suboptimal for these patients. Furthermore, there is currently no approved treatment for liver manifestations. The unmet medical need for patients with AAT deficiency remains high, and new treatment options are needed to treat the underlying disease etiology. REVIEW FINDINGS: Advances in genomic medicines may enable treatment by editing the DNA or RNA sequence to produce wild-type AAT instead of the mutated AAT caused by the E342K mutation. One approach can be achieved by directing endogenous adenosine deaminases that act on RNA to the E342K RNA site, where they catalyze adenosine to inosine conversion through a process known as RNA editing. The A-I RNA change will be read as a G during protein translation, resulting in an altered amino acid and restoration of wild-type AAT secretion and function. SUMMARY: In this review, we will discuss the pathophysiology of AAT deficiency and emerging treatment options with particular focus on RNA editing as a disease-modifying treatment for both liver and lung disease.
alpha 1-Antitrypsin Deficiency↗
Global quantification of off-target activity by base editors.
Base editors are engineered deaminases combined with CRISPR components. These engineered deaminases are designed to target specific sites within DNA or RNA to make a precise change in the molecule. In therapeutics, they hold promise for correcting mutations associated with genetic diseases. However, a key challenge is minimizing unintended edits at off-target sites, which could lead to harmful mutations. Researchers are actively addressing this concern through a variety of optimization efforts that aim to improve the precision of base editors and minimize off-target activity. Here, we examine the various types of off-target activity, and the methods used to evaluate them. Current methods for finding off-target activity focus on identifying similar sequences in the genome or in the transcriptome, assuming the guide RNA misdirects the editor. The main method presented here, that was originally developed to quantify editing levels mediated by the ADAR enzyme, takes a different approach, investigating the inherent activity of base editors themselves, which might lead to off-target edits beyond sequence similarity. The editing index tool quantifies global off-target editing, eliminates the need to detect individual off-target sites, and allows for assessment of the global load of mutations.
Gene Editing↗
RNA-DNA hybrid binding domain broadens the editing window of base editors.
Adenine base editors (ABEs) and cytosine base editors (CBEs) are prominent tools for precise genome editing but are hindered by limited editing activity at positions proximal to the protospacer adjacent motif (PAM). This study investigates the potential of enhancing base editors editing activity by fusing them with RNA-DNA hybrid binding domains (RHBDs). Specifically, fusing ABE8e with the RHBD of Homo sapiens RNaseH1 (RHBD1) significantly increased A-to-G editing efficiency in the PAM-proximal region (A9-A15) by up to 3.5-fold, while reducing off-target cytosine editing. Additionally, RHBD1 is compatible with ABEmax, BE4max, and dual base editor (eA&C-BEmax), enhancing their editing activity at the PAM-proximal bases. Notably, RHBD1-fused BE4max led to a 3.1-fold improvement in C-to-T editing efficiency at PAM-proximal region (C9-C12). Furthermore, we demonstrated that RHBD1-fused ABE8e could effectively edit disease-related single nucleotide variations (SNVs) in human cells and validated its efficacy in adult mouse liver. These findings highlight the significance of the RHBD in expanding editing window and the applicability of base editors for gene therapy and disease modeling.
Gene Editing↗
CRISPR RNP-Mediated Transgene-Free Genome Editing in Plants: Advances, Challenges and Future Directions for Tree Species.
CRISPR ribonucleoprotein (RNP)-mediated genome editing offers a transgene-free platform for precise genetic modification in diverse herbaceous and tree species, including rice, wheat, apple, poplar, oil palm, rubber tree and grapevine. However, its application in woody plants faces distinct challenges, notably inefficient delivery and regeneration difficulties, particularly in species such as bamboo. While some of these issues also occur in herbaceous plants, they are often significantly more complex in woody species due to factors such as intricate cell wall architecture, widespread recalcitrant genotypes and inherent limitations of current delivery platforms. This review presents the first in-depth, critical re-evaluation of recent advancements in RNP-mediated editing in woody plants, highlighting these obstacles that warrant focused attention. Unlike plasmid-based CRISPR systems, RNP editing utilises Cas9/Cas12a protein-guide RNA complexes without integrating foreign DNA. This enables a DNA-free editing strategy that simplifies regulatory approval and minimises off-target effects due to the transient presence and rapid degradation of RNPs within plant cells. While PEG-mediated protoplast transfection and particle bombardment remain the primary reported methods for RNP delivery in trees, we evaluate promising alternative strategies such as lipofection, electroporation, cell-penetrating peptides and nanoparticle-based systems for targeted RNP delivery. Despite their promise, these advanced methods remain largely untested in woody species. Finally, we outline future research directions, including the development of tree-specific RNP delivery systems and regeneration protocols to enhance efficiency and minimise cytotoxicity. These innovations are essential for unlocking the full potential of RNP-mediated genome editing in long-lived tree species. This review provides a focused and timely roadmap for expanding the application of RNP technology across diverse woody plants.
Gene Editing↗
Stepwise DNA-unwinding gates TnpB genome-editing activity.
TnpB is a compact RNA-guided endonuclease and an evolutionary ancestor of CRISPR-Cas12 that offers a promising platform for genome engineering. However, the genome-editing activity of TnpBs remains limited, and its underlying determinants are poorly understood. Here, we used biochemical and single-molecule assays to examine the DNA-unwinding mechanism of Youngiibacter multivorans TnpB (Ymu1 TnpB). DNA unwinding proceeds through a discrete, long-lived, partially unwound intermediate state before reaching a fully unwound open state. The open state forms inefficiently and collapses readily in the absence of negative supercoiling. An optimized variant, H4W-L304F-V305R (Ymu1-WFR), stabilizes formation of both the intermediate and open states, resulting in enhanced DNA cleavage in vitro and increased genome editing in plants. These findings identify the physical basis for the observed minimal activities of natural TnpBs, revealing how stabilizing specific unwinding states enables efficient DNA targeting.
Gene Editing↗
Re-arranging the Cis-regulatory Modules of Hox Complex in Drosophila via FLP-FRT and CRISPR/Cas9.
FLP-FRT, a well-established technique for genome manipulation, and the revolutionary CRISPR/Cas9, known for its targeted indels, are combined in a novel approach. This unique method is applied to the Hox genes in the Drosophila melanogaster bithorax complex, which are closely located to the cis-regulatory modules that define their spatial-temporal regulation. The number and position of these genes are directly correlated to their expression pattern. This chapter unveils the exciting potential of this combinatorial use of FLP-FRT and CRISPR-Cas9 to rearrange the cis-regulatory modules of the Hox complex in Drosophila melanogaster.
Animals↗
In vivo genome editing of central nervous system SIV reservoirs in ART-suppressed rhesus macaques.
Latent human immunodeficiency virus type 1 (HIV-1) reservoirs in the central nervous system (CNS) may sustain viral persistence and neuroinflammation contributing to HIV-associated neurocognitive disorders (HAND) despite suppressive ART. AAV9-delivered CRISPR has successfully edited SIV proviral DNA in peripheral tissues with acceptable safety profiles, but the extent of in vivo genome editing in the brain remains unclear. Using SIV-infected rhesus macaques, we mapped intact proviral DNA across CNS regions and tested systemic AAV9-CRISPR-Cas9 targeting conserved sites within Ψ packaging signal and Gag region. Ten adult rhesus macaques were infected with genetically barcoded SIVmac239, suppressed with ART, then randomized to receive intravenous AAV9-SaCas9 with dual gRNAs (Ψ + Gag) or a Cas9-only control. At necropsy after viral rebound, SIV genomes were detected in multiple brain regions as well as lymphoid tissues, confirming the CNS as a persistent reservoir during ART. Barcode analysis revealed region-specific patterns consistent with compartmentalized CNS persistence. In CRISPR-treated animals, proviral editing was measurable across anatomically distinct CNS sites. These findings demonstrate that intact and potentially replication-competent virus persists in the primate brain under ART and that systemic AAV9-CRISPR can reach and edit proviral DNA in this sanctuary, supporting genome editing as a strategy toward durable remission of CNS reservoirs.
ART↗
Discovery and Engineering of a Rat Endogenous Retrovirus Reverse Transcriptase for Efficient Prime Editing.
CRISPR-based prime editors (PEs) install precise edits into genomic DNA without generating double-strand breaks. Their editing efficiency is highly dependent on reverse transcriptases (RTs), but efficient RT candidates remain limited. Here, we identified 19 novel active RTs by screening 558 candidates. Among them, RERV-RT, derived from Rattus norvegicus, exhibited the highest activity. Through structure-guided engineering and deep mutational scanning, we developed an optimized variant, enRERV-RT, which outperforms conventional M-MLV-RT-based PE systems by 1.20-fold in mammalian and plant cells, and by 1.88-fold at hard-to-edit loci, while enabling precise multiplex editing of functionally relevant genes. Additionally, we developed a high-throughput platform, TRAP-seq-PE, to systematically evaluate prime editor performance. Across diverse mutation types, we found that PE systems based on enRERV-RT exhibited higher editing efficiencies than those based on M-MLV-RT. Collectively, our work establishes a versatile, high-efficiency PE system, thereby facilitating advances in clinical gene therapy and precise crop breeding.
Animals↗
Cellular Inflammation-Induced Cleavage of Phosphorothioate DNA Locker Activates CRISPR/Cas9 Regulator for Gene Editing.
The CRISPR technology is a highly promising strategy for developing a versatile toolbox to engineer genetic circuits. However, achieving precise and specific control over the activity of the CRISPR/Cas9 system in response to intracellular processes remains a challenging endeavor. In this study, we present a cellular inflammation-induced activation of an engineered CRISPR/Cas9 regulator for gene regulation. A phosphorothioate (PS)-modified DNA sequence, referred as the "locker," is employed to deactivate single guide RNA (sgRNA), whose locker sequence complements the spacer region of sgRNA. In the presence of myeloperoxidase during cellular inflammation, a halogenation process is triggered, leading to the generation of HClO, specifically cleaving the PS site of locker and activating CRISPR/Cas9 for gene editing. The target GFP gene has been successfully edited, downregulating the GFP protein expression in HeLa cells. This study provides valuable insights into the CRISPR-based gene regulation through specific endogenous processes.
Humans↗
Diversity at the HYP1 locus in potato cyst nematodes does not result from developmentally-programmed somatic mutations.
Most genetic diversity stems from spontaneous mutations, that is, errors in DNA repair or replication. But for dozens of organisms across the tree of life, mutations at specific loci are not spontaneous but developmentally programmed: effectively, some organisms edit their own DNA sequences. This is perhaps most common among pathogens and parasites, many of which use editing to diversify genes that produce important antigens. Plant-parasitic potato cyst nematodes are damaging agricultural pests that establish a lifelong feeding site inside the root of their host plant. We previously observed extensive diversity of rare alleles at HYP1, the most highly expressed gene that encodes a protein secreted by potato cyst nematodes during parasitism. Importantly, HYP1 alleles differ from each other by complex, in-frame rearrangements of short repeated sequence motifs within a single exon. Combining several lines of evidence, we previously hypothesized that potato cyst nematodes use developmentally-programmed mutations, or editing, to diversify HYP1 alleles in the soma. In the current work, we now test this hypothesis. We employ highly accurate long-read DNA sequencing of a simplified genetic system to identify potential rare edited alleles, we use a transgenic yeast system to describe large de novo mutations at HYP1, and we interpret our findings in light of key population genetic parameters as well as the genetic diversity surrounding HYP1 and across the genome.
Animals↗
Development of a highly efficient protoplast regeneration and transfection protocol for enhancing CRISPR genome editing of Brassica carinata.
Brassica carinata is an important oil crop with significant potential for food and industrial production. The application of the CRISPR/Cas9 genome editing tool in B. carinata could accelerate its breeding cycle. However, no efficient DNA-free gene editing method currently exists for this species. Protoplast-based CRISPR editing presents a promising solution, though it is often challenging for many crop species. In this study, we investigated several critical factors influencing in vitro shoot regeneration, including genotype, sugar type, selection and combination of plant growth regulators (PGRs), and culture duration on different media throughout various stages of protoplast development. As a result, we developed a highly efficient, five-stage protoplast regeneration protocol for B. carinata based on specific stages of protoplast development. Key findings of this study include the requirement for high concentrations of NAA and 2,4-D in the initial medium (MI) for cell wall formation, while a lower auxin concentration relative to cytokinin was necessary for active cell division (MII). For callus growth and shoot induction, a high cytokinin-to-auxin ratio was essential (MIII), and an even higher cytokinin-to-auxin ratio was optimal for shoot regeneration (MIV). For shoot elongation, low levels of BAP and GA3 were sufficient (MV). Our results also demonstrated that the duration of culture on different media and maintaining appropriate osmotic pressure at the early stages were crucial for successful protoplast regeneration. With this optimized protocol, we achieved an average regeneration frequency of up to 64% and a transfection efficiency of 40% using the GFP marker gene. This efficient protoplast regeneration protocol is now being employed for genome editing in our lab and is expected to significantly enhance the application of the CRISPR system in both basic research and the genetic improvement of B. carinata over the long term.
Brassica carinata↗
Activation of the imprinted Prader-Willi syndrome locus by CRISPR-based epigenome editing.
Epigenome editing with DNA-targeting technologies such as CRISPR-dCas9 can be used to dissect gene regulatory mechanisms and potentially treat associated disorders. For example, Prader-Willi syndrome (PWS) results from loss of paternally expressed imprinted genes on chromosome 15q11.2-q13.3, although the maternal allele is intact but epigenetically silenced. Using CRISPR repression and activation screens in human induced pluripotent stem cells (iPSCs), we identified genomic elements that control the expression of the PWS gene SNRPN from the paternal and maternal chromosomes. We showed that either targeted transcriptional activation or DNA demethylation can activate the silenced maternal SNRPN and downstream PWS transcripts. However, these two approaches function at unique regions, preferentially activating different transcript variants and involving distinct epigenetic reprogramming mechanisms. Remarkably, transient expression of the targeted demethylase leads to stable, long-term maternal SNRPN expression in PWS iPSCs. This work uncovers targeted epigenetic manipulations to reprogram a disease-associated imprinted locus and suggests possible therapeutic interventions.
Prader-Willi Syndrome↗