🧬 PubMed RNA Editing Daily Digest

RNA-binding proteins (RBPs) are essential regulators of gene expression at the post-transcriptional level, yet obtaining quantitative insights into RBP-RNA interactions in vivo remains challenging. Here, we developed RBP specificity and contextual analysis via nucleotide editing (RBPscan), which integrates RNA editing with massively parallel reporter assays to profile RBP binding in vivo. In RBPscan, fusion of an RBP to the adenosine deaminase acting on RNA (ADAR) catalytic domain induces RNA editing of a recorder mRNA carrying the tested RBP-binding site, serving as a readout of the RBP-RNA interaction. We demonstrate the utility of RBPscan in zebrafish embryos, human cells, and yeast, showing that it quantifies binding strength, resolves dissociation constants, identifies binding motifs for various RBPs, and links binding affinities to their impact on mRNA stability. RBPscan also provides positional mapping of Pumilio-binding sites in the long non-coding RNA NORAD. With its simplicity, scalability, and cross-system compatibility, RBPscan is a versatile tool for investigating protein-RNA interactions and complements established methods for studying post-transcriptional regulatory networks.

Unlike genome editing, RNA editing offers the ability to transiently alter cells with minimal risk from off-target effects. While exon-skipping technologies can influence splice site selection, many desired perturbations to the transcriptome require replacement or addition of exogenous exons to target mRNAs, such as replacing disease-causing exons, repairing truncated proteins, or engineering protein fusions. Here, we report the development of RNA-guided trans-splicing with Cas editor (RESPLICE). RESPLICE uses two orthogonal RNA-targeting CRISPR effectors to co-localize a trans-splicing pre-mRNA and to inhibit the cis-splicing reaction, respectively. We demonstrate efficient, specific, and programmable trans-splicing of RNA cargo (up to 2.1 kb) into 11 endogenous transcripts across 3 cell types, achieving up to 45% trans-splicing efficiency in bulk or 90% when sorting for high effector expression. Our results present RESPLICE as a mode of RNA editing that could provide fine-tuned and transient control of cellular programs.

Achieving stable and efficient transgene expression is a key challenge in advancing avian genome engineering. Although viral vector-based and piggyBac-mediated transgenesis have been widely used in chickens, both approaches are prone to epigenetic silencing, leading to inconsistent, tissue-specific, and often diminished expression over time. This variability limits used of transgenes requiring robust and long-term expression across multiple tissues. In mammals, site-specific integration into genomic safe harbor loci, such as Rosa26, has enabled stable and predictable transgene expression without disrupting endogenous gene function; however, such strategy has not been established in birds. In this research, we hypothesized that integrating Cas9 into endogenous housekeeping genes (the ACTB and GAPDH) could achieve efficient gene editing in chickens through stable and ubiquitous transgene expression. Using two different approaches, 3'-targeted gene insertion and gene tagging, we inserted Cas9 and GFP cassettes into defined genomic loci in chicken DF-1 cells. Both approaches exhibited stable expression of transgenes in the cells, and functional assays confirmed that Cas9 showed highly efficient nuclease activity following guide RNA delivery. Additionally, we derived single-cell clones stably expressing Cas9, enabling uniform and reproducible genome editing in downstream applications. Targeted insertion of transgenes into active housekeeping genes as candidate safe harbor loci mitigates the limitations of random integration and promoter silencing, offering a robust platform for consistent transgene expression in poultry biotechnology and genome engineering.

Babesia vogeli is a protozoan parasite causing canine babesiosis, a tick-borne disease prevalent in tropical and subtropical regions. Its microscopic identification is challenging due to morphological similarity with other Babesia spp., and serological assays often yield inaccurate results. To address this issue, we developed a rapid, equipment-minimal diagnostic method combining recombinase polymerase amplification (RPA) with CRISPR/Cas12a (RPA/CRISPR-cas12a) for B. vogeli-specific detection. The RPA assay enables DNA amplification for both B. vogeli and Hepatozoon canis, while CRISPR/Cas12a using gRNA_Bab ensures specificity for B. vogeli, even in co-infections and other pathogens. This approach detects as few as 10⁵ copies within two hours for both readout platforms such as fluorescence and lateral flow dipstick (LFD). Forty canine blood samples were detected by RPA/CRISPR-cas12a to examine its performance. Results showed high concordance with qPCR-high resolution melting (HRM) (Cohen's kappa: 0.93 for fluorescence, 0.81 for LFD), outperforming conventional PCR. The clinical sensitivity and specificity of RPA/CRISPR-cas12a were 100 % and 96.8 %, respectively and the concordance with qPCR-HRM was 97.5 %. RPA/CRISPR-cas12a for Babesia spp. detection provided a simple, rapid, and accurate method, demonstrating promise for point-of-care diagnosis of canine babesiosis in resource-limited settings. This method showed high potential as a practical diagnostic tool in veterinary clinics, with accelerated surveillance to control outbreaks of Babesia-associated canine babesiosis.

Cotton leafroll dwarf virus (CLRDV) poses an increasing threat to global cotton production. Transmitted by the cotton aphid (Aphis gossypii) in a persistent, circulative manner, CLRDV exhibits a wide geographical distribution, with documented presence in South America, Africa, Asia, and the USA. Infection can result in either cotton blue disease (CBD) in South America or cotton leafroll dwarf disease (CLRDD) in the USA, both of which are associated with CLRDV. The considerable genetic diversity and frequent recombination events within CLRDV populations contribute to this symptom variability and complicate both diagnosis and management. While resistant cultivars have reduced disease impact in South America, these lines remain susceptible to emerging US strains, underscoring the urgent need for region-specific resistance breeding. Current molecular diagnostics rely on RT-PCR, but there is a need for rapid, field-deployable detection tools. Recent advances, such as CRISPR-Cas13a based SHERLOCK assays, offer sensitive and specific detection of CLRDV, with potential for on-site applications. Efficient screening techniques, supported by next-generation sequencing and transcriptomics, are essential for identifying novel resistance sources and elucidating virus-host interactions. CRISPR-based genome editing holds significant promise, as demonstrated in other crops. Targeted disruption of host susceptibility genes using CRISPR-Cas9, or direct degradation of viral genomes with RNA-targeting systems such as Cas12/Cas13, could offer durable, broad-spectrum resistance. By integrating molecular virology, high-throughput genomics, and precision gene editing, this review outlines a roadmap for translating these advances into sustainable, field-level solutions for CLRDV management and long-term cotton productivity.