搜索结果：Frontiers in genome editing

The advent of genetics, molecular biology, and genome sequencing has rapidly accelerated the development of elite genetic lines across various species, including poultry. It is now possible to introduce intra- or inter-species single nucleotide polymorphisms into chicken lines to enhance productivity. This advancement may mark the beginning of a new agricultural revolution, dramatically reducing the time required to improve poultry lines for commercial production environments. Transgenic technologies, including lentiviral vectors and piggyBac transposition, have enabled the generation of animals expressing exogenous genes. The emergence of new genome-editing tools is transforming avian biotechnology, allowing the creation of customized organisms for specific traits. Genome editing has become the most efficient method for studying gene function. First and second generation tools, such as zinc finger nucleases and transcription activator-like effector nucleases (TALENs), are limited by complex design and off-target effects. In contrast, the third generation Clustered Regularly Interspaced Short Palindromic Repeats/CRISPR-associated protein 9 (CRISPR/Cas9), represents a significant breakthrough. It encompasses guided RNA (gRNA) and the Cas9 endonuclease which together target specific DNA sequences and induces double-strand breaks that are repaired via error-prone non-homologous end joining, frequently causing insertions or deletions that disrupt gene function. Targeting specificity is achieved through gRNA-DNA base pairing and recognition of a protospacer adjacent motif by Cas9. Beyond gene knockout, CRISPR/Cas9 enables functional analysis of non-coding elements such as enhancers and insulators. Delivered via plasmid systems, Cas9 and gRNA are transiently expressed and degrade within 48-72 h, leaving no permanent genetic footprint. Since no exogenous DNA is integrated, this approach is generally considered less contentious than traditional transgenic methods in the context of genetically modified organism regulation. CRISPR/Cas9 has diverse applications in poultry, including enhancing disease resistance to avian influenza and Marek's disease, improving productivity traits such as growth, feed efficiency, and egg-laying, and enabling early in-ovo sexing to address ethical concerns around male chick culling. It also allows control of reproductive traits for breeding management, supports bio-pharming by producing therapeutic proteins or vaccines in eggs, and facilitates functional genomics through precise gene knockouts to study development, immunity, and metabolism.

Plant genome editing has undergone a transformative shift with the advent of advanced molecular tools, offering unprecedented levels of precision, flexibility and efficiency in modifying genetic material. While classical site-directed nucleases such as ZFNs, TALENs and CRISPR-Cas9 have revolutionized genome engineering by enabling targeted mutagenesis and gene knockouts, the landscape is now rapidly evolving with the emergence of novel systems that go beyond the conventional double strand break (DSB)-mediated approaches. Advanced and recent tools include LEAPER, SATI, RESTORE, RESCUE, ARCUT, SPARDA, helicase-based approaches like HACE and Type IV-A CRISPR system, and transposon-based techniques like TATSI and piggyBac. These tools unlock previously inaccessible avenues of genome and transcriptome modulation. Some of these technologies allow DSB-free editing of DNA, precise base substitutions and RNA editing without altering the genomic DNA, a significant advancement for regulatory approval and for species with complex genomes or limited regeneration capacity. While LEAPER, RESCUE and RESTORE are the new advents in the RNA editing tool, SATI allows DSB-free approach for DNA editing, ARCUT offers less off-target and cleaner DNA repairs and Type IV-A CRISPR system induces gene silencing rather than editing. The transposon-based approaches include TATSI, piggyBac and TnpB, and helicases are used in HACE and Type IV-A CRISPR system. The prokaryotic Argonaute protein is used in SPARDA tool as an endonuclease to edit DNA. The transient and reversible nature of RNA editing tools such as RESTORE and LEAPER introduces a new layer of epigenetics-like control in plant systems, which could be harnessed for tissue-specific and environmentally-responsive trait expression. Simultaneously, innovations like ARCUT and SPARDA utilize chemically-guided editing, minimizing reliance on biological nucleases and reducing off-target risks. Their modularity and programmability are enabling gene function studies, synthetic pathway designs and targeted trait stacking. These advances represent a novel synthesis of genome engineering and systems biology, positioning plant genome editing not just as a tool of modification but as a platform for designing adaptive and intelligent crops, tailored to future environmental and nutritional challenges. Although, many of these recent tools remain to be applied on plant systems, they are proven to be effective elsewhere and hold a great potential to be effective in creating climate-resilient crops.

Two major processes are important for genome editing in plants: transformation by stable transfection, in which nucleic acids encoding genome-editing enzymes are introduced into plant cells and the regeneration of plant individuals from cells harboring mutations by genome-editing enzymes. The efficiency of transformation and regeneration by tissue culture varies across plant species, and is low in some practical crop species. In planta methods have been developed to exclude the need for tissue culture. However, few reports are available on methods that do not require stable transfection. Therefore, this study aimed to develop a new protocol for delivery genome editing tools that does not require transformation or tissue culture, by combining the in planta method with transient genome editing tools instead of stable transfection. Cas9, guide RNAs, and developmental regulators, which are factors involved in mitotic tissue induction, were transiently expressed by agroinfiltration of the stem tissue cut surfaces of tomatoes. New chimeric mutants, containing a mixture of cells with mutations introduced at or near the target sequence, were obtained. After examining conditions such as the concentration of Agrobacterium used for infection and post-infection treatment, we succeeded in obtaining chimeric mutants with an efficiency of 11.7%. In addition, most of the observed mutations were single base substitutions. These results indicate that the in planta method with transient expression of genome editing tools and induction of meristematic tissue can be used to introduce genome-edited mutations in tomatoes.

Legumes are among the most important protein-rich crops in global agri-food systems. To meet the rising protein demand of a growing population, significant efforts are underway to enhance legume yield, nutritional quality, and resilience to environmental stresses through the manipulation of key genetic traits. Advanced technologies-including genetic engineering, gene editing, genomic selection, next-generation sequencing, single-cell genomics, and multi-omics-are accelerating legume improvement due to their high precision and efficiency. This review focuses on major gene-editing technologies, namely, CRISPR/Cas9 (Clustered Regularly Interspaced Short Palindromic Repeats/CRISPR-associated protein 9), TALENs (Transcription Activator-Like Effector Nucleases), ZFNs (Zinc Finger Nucleases), base editing (BE), and prime editing (PE), and their applications in key legume crops such as soybean (Glycine max), cowpea (Vigna unguiculata), chickpea (Cicer arietinum), groundnut (Arachis hypogaea), pea (Pisum sativum), barrel clover (Medicago truncatula), alfalfa (Medicago sativa), and Lotus japonicus. Among these platforms, CRISPR/Cas9 is the most widely adopted in legumes, largely due to its simplicity, versatility, and dependence on accurate genome sequence information and guide RNA (gRNA) design. Advances in next-generation sequencing and the growing availability of intuitive online gRNA design tools have streamlined CRISPR workflows, improving accessibility and precision. The present review indicates that CRISPR-P is the most used gRNA design tool in legume research, likely due to its early development for plant systems and integrated off-target prediction features. Therefore, alongside reviewing gene-editing applications, we emphasized the critical role of robust gRNA design tools as a foundation for successful genome editing. Future integration of artificial intelligence and large language models is expected to further enhance target prediction accuracy, minimize off-target effects, and enable more precise genome-editing strategies in legumes.

Homology-directed repair (HDR) holds great promise for plant genetic engineering but remains challenging due to its inherently low efficiency in gene editing applications. While studies in animal systems suggest that the structure of the donor repair template (DRT) influences HDR efficiency, this parameter remains largely unexplored in plants. In this study, we combined protoplast transfection with next-generation sequencing to analyse the impact of DRT structure on HDR efficiency in potato. A highly efficient ribonucleoprotein (RNP) complex targeting the soluble starch synthase 1 (SS1) gene was used in combination with various DRTs, differing in structural factors such as homology arm (HA) length, strandedness (i.e., ssDNA vs. dsDNA), and sequence orientation in ssDNA donors. Our results indicate that a ssDNA donor in the target orientation outperformed other configurations, achieving a HDR efficiency of 1.12% of the sequencing reads in the pool of protoplasts. Interestingly, HDR efficiency appeared independent of HA length. Notably, a ssDNA donor with HAs as short as 30 nucleotides led to targeted insertions in up to 24.89% of reads on average, but predominantly via alternative imprecise repair pathways, such as microhomology-mediated end joining (MMEJ). This donor structure also consistently yielded the highest HDR and targeted insertion rates at two out of three additional loci tested, offering valuable insights for future genome editing strategies in potato. We further assessed strategies to favour HDR over alternative repair outcomes, including the use of small molecules known to inhibit competing pathways in animal systems, and modifications to DRTs to enhance their availability in the vicinity of the target site. However, these approaches did not improve HDR efficiency. Overall, this study presents an effective platform for rapidly assessing gene editing components in potato and provides insights for achieving high-frequency, targeted insertions of short DNA fragments, especially relevant for efficient knock-in integration in non-coding genomic regions.

Protoplast-based systems have been utilised in a wide variety of plant species to enable genome editing without chromosomal introgression of foreign DNA into plant genomes. DNA-free genome editing followed by protoplast regeneration allows elite cultivars to be edited without further genetic segregation, preserving their unique genetic composition and their regulatory status as non-transgenic. However, protoplast isolation presents a barrier to the development of advanced breeding technologies in raspberry and no protocol has been published for DNA-free genome editing in the species. Pre-assembled ribonucleoprotein complexes (RNPs) do not require cellular processing and the commercial availability of Cas9 proteins and synthetic guide RNAs has streamlined genome editing protocols. This study presents a novel high-yielding protoplast isolation protocol from raspberry stem cultures and RNP-mediated transfection of protoplast with CRISPR-Cas9. Targeted mutagenesis of the phytoene desaturase gene at two intragenic loci resulted in an editing efficiency of 19%, though estimated efficiency varied depending on the indel analysis technique. Only amplicon sequencing was sensitive enough to confirm genome editing in a low efficiency sample. To our knowledge, this study constitutes the first use of DNA-free genome editing in raspberry protoplast. This protocol provides a valuable platform for understanding gene function and facilitates the future development of precision breeding in this important soft fruit crop.

CRISPR/Cas9 technology has gained popularity due to its efficient, widely applicable, and relatively easy genome editing. Furthermore, the removal of regulation on site-directed nuclease1- (SDN1) and SDN2-developed products in many countries has made it a more revolutionary technology for adoption in crop improvement. Designing accurate guide RNA (gRNA) is the initial and most crucial step that decides the success of the editing. Although the gene editing technique is widely used in crops, a detailed and comprehensive method for designing efficient gRNA in wheat is still lacking. By virtue of wheat being a hexaploid crop and having a large genome size with repetitive DNA, a tailor-made strategy for designing the gRNA is crucial. The manuscript explains the comprehensive strategies and methods for efficient gRNA designing by considering the physical and structural expression of the target gene in the genome and explains the on-target and off-target effects of gRNA for its precise editing through the CRISPR/Cas9-mediated SDN1 method of genome editing in wheat. The present manuscript is first of its kind to address the holistic approach, starting from efficient gene selection, gRNA designing, and post-gRNA designing issues like gRNA stability, binding efficiency, and functionality for SDN1-CRISPR/Cas9 genome editing in wheat. This manuscript will be a ready reference for wheat researchers designing effective gRNA for wheat improvement to meet future food demand.

Genome-wide association studies (GWAS) have identified numerous single nucleotide polymorphisms (SNPs) associated with complex traits in poultry. However, most GWAS-identified variants reside in non-coding regions, making their functional relevance to their phenotypes unclear. Emerging evidence suggests that many of these markers overlap cis-regulatory elements, yet experimental validation of their biological function remains limited. Here, we investigated non-coding GWAS variants associated with nucleotide-related compounds in chicken breast muscle by targeting SNP-containing genomic regions using a CRISPR activation (CRISPRa) system in DF-1 cells and profiling transcriptomic responses via bulk RNA sequencing to assess the functional impact of activating these regions. Based on chicken muscle-specific epigenetic profiles and chromatin state annotations, we identified three significant GWAS variants on chromosome five associated with nucleotide metabolism. These variants are situated within cis-regulatory elements, specifically in intron three of DUSP8, intron one of SLC25A22, and upstream of FBXO3. To understand their functional impact, we employed an in vitro CRISPRa system with targeted guide RNAs to activate each non-coding SNP region in DF-1 cells. This activation resulted in significant changes at the transcriptomic level. Subsequent functional enrichment analysis of the differentially expressed genes consistently highlighted muscle-related pathways across all SNPs, including MAPK signaling, cytoskeletal remodeling, and ECM-receptor interactions, which are potentially involved in regulating nucleotide metabolism and deposition in muscle. Furthermore, transcript-level analysis of RNA-seq reads revealed that the non-coding SNP region within the intron three of DUSP8 may function as an alternative promoter, resulting in significantly higher expression of a shorter transcript that could generate a non-canonical protein isoform. Our study demonstrates that activating genomic regions harboring specific non-coding GWAS SNPs can modulate gene expression, suggesting that these SNPs may contribute to gene regulatory functions. Importantly, this work underscores the powerful utility of CRISPRa as a functional genomics tool for linking GWAS signals to their biological roles in chickens by targeting SNP-containing regions and uncovering consequential molecular phenotypes.

Targeted insertion (TIN) of transgenic trait cassettes has the potential to reduce timeline and cost for GM product development and commercialization. Precise genome engineering has made remarkable progress over the last several decades, particularly with the use of site-directed nucleases as core editing machinery. However, there are still many critical factors that can impact TIN efficiency including insertion site selection, nuclease optimization and expression, donor vector design, gene delivery, and stable event regeneration. Here, we develop workflows for target site sequence identification and gRNA screening for CRISPR-Cas12a system and demonstrate its successful application for TIN in maize with donor sequences up to 10 kilobase pairs (kb) in size. We first prioritize genomic regions for inserting transgenes in silico using bioinformatics tools and then test gRNA performance using a leaf protoplast transient assay. Despite its known low efficiency, we identify homology-directed repair (HDR) as the preferential pathway for directing targeted insertions of large sequences in immature embryos and demonstrate double-junction integrations at a rate of up to 4%. We further apply a molecular analysis workflow using large amplicon TaqMan assays and nanopore sequencing for streamlined identification and characterization of high-quality insertion events with intact large inserts. Analysis of TIN events across generations suggests that efficiency bottlenecks are not limited to donor targeted integration; attrition in efficiency also results from partial or additional donor insertion, chimerism, and close linkage with undesired sequence insertions such as those encoding the editing machinery. This work represents a major step forward in realizing the potential of precise genome engineering in maize for basic research and biotech trait development applications.

Advancements in genome editing technologies, notably CRISPR/Cas9, base editing (BE), and prime editing (PE), have revolutionized plant biotechnology, offering unprecedented precision in crop improvement to address the ongoing global warming challenge. This review provides a critical analysis of recent developments in SpCas9-based editing tools, emphasizing enhancements in editing efficiency and specificity and follow the chronological development of editing tools. We explore methodological innovations, including dual pegRNA strategies and site-specific integrases, that have expanded the potential of PE for precise gene insertions. By integrating insights into DNA repair mechanisms and leveraging SpCas9 enhancements, we outline future directions for the application of genome editing in plant breeding.

Potato is an important vegetatively propagated, starch-rich tuber crop. High amylose potatoes containing more resistant starch offer healthier food alternatives. However, the resistant starch content is low in most cultivated potato varieties. In this study, targeted mutation of the starch branching enzyme2 (SBE2.1 & SBE2.2 isoforms) had been done in the commercially significant potato cultivar, Kufri Chipsona-I using Clustered regularly interspaced short palindromic repeats-CRISPR-associated protein 9 (CRISPR-Cas9 system) to develop high-amylose potato lines. SBE2 is one of the key enzymes involved in amylopectin biosynthesis, a starch component. Two isoforms, SBE2.1 & SBE2.2, were mutated using CRISPR-Cas9-mediated genome editing. After Agrobacterium-mediated genetic transformation, fifty transformed lines were generated on herbicide Basta selection medium, out of which 70% were found positive for bar and Cas9 genes. Overall, six mutant lines, viz. K301, K302, K303, K304, K305, K306, derived from distinct events, exhibited deletions and substitutions in the target exons. The CRISPR-Cas9 edited K304 potato line exhibited both insertion-deletion (indel) and substitution mutations in three out of the four selected targets across both genes, and was therefore identified as the most efficiently edited line. The harvested tubers from SBE2.1 & SBE2.2 mutant K304 line showed the highest amylose (95.91%) and resistant starch content (8.69 g/100 g). Evaluation of starch using X-ray crystallography (XRD) illustrated an altered crystallinity index (CI%) in all six mutant events in comparison to the wild study. Furthermore, 1H-NMR study demonstrated a substantial decline in branch chain elongation in amylopectin, and thus a low degree of branching in a range of 1.15%-3.66% was reported in mutant lines, relative to the wild type (5.46%). The present study demonstrated the efficacy of CRISPR-Cas9-mediated mutagenesis of starch biosynthetic genes to develop high-amylose potato lines with elevated resistant starch content for improved health benefits.

Base editing has revolutionized genome engineering by enabling precise single-nucleotide modifications without inducing double-strand breaks. As a powerful and efficient gene-editing tool, base editors (BEs) have been widely applied in various model organisms, including zebrafish (Danio rerio), to facilitate functional genomic studies and disease modeling. Zebrafish, with its genetic similarity to humans and rapid development, provides an excellent platform for testing and optimizing emerging base editing technologies. This review comprehensively explores the advancements of cytosine and adenine base editors in zebrafish, highlighting recent developments that enhance efficiency, specificity, and editing scope. We discuss novel base editor variants tailored for zebrafish applications, improvements in delivery strategies, and methodologies to minimize off-target effects. Furthermore, we compare base editing with other precision genome-editing technologies, such as prime editing and homology-directed repair, to underscore its advantages in achieving targeted mutations with high fidelity. By evaluating the expanding role of base editing in zebrafish, this review provides valuable insights into its potential for translational research, genetic disease modeling, and future therapeutic applications.

Peruvian agriculture is characterize by crops such as potato, maize, rice, asparagus, mango, banana, avocado, cassava, onion, oil palm, chili, papikra, blueberry, coffee, cacao, grapes, quinoa, olive, citrus and others. All of them have challenges in production in their specific agroecosystems under stress due to pests, diseases, salinity, drought, cold among others. Gene editing through CRISPR/Cas is a key tool for addressing critical challenges in agriculture by improving resilience to biotic and abiotic stress, increasing yield and enhancing the nutritional value of the crops. This approach allows precise mutation on site-specific gene at the DNA level, obtaining desirable traits when its function is altered. The CRISPR/Cas system could be used as a transgene-free genome editing tool when the ribonucleoprotein (RNP) acts as a carrier to delivered the CRISPR/Cas components into the plant cell protoplasts, or when the tRNA-like sequence (TLS) motifs are fused to single-guide RNA (sgRNA) and Cas mRNA sequence and expressed in transgenic plants rootstock to produce "mobile" CRISPR/Cas components to upper tissue (scion). Those innovations could be a potential approach to strengthen the Peruvian agriculture, food security and gricultural economy, especially in the tropical, Andean and coastal regions. This review article examines the advances and strategies of gene editing, focusing on transgene-free methodologies that could be adopted for research, development and use, and also identifies potential applications in key crops for Peru and analyzes their impact in the productivity and reduction of agrochemicals dependence. Finally, this review highlights the need to establish regulatory policies that strengthen the use of biotechnological precise innovations, ensuring the conservation and valorization of agrobiodiversity for the benefit of Peruvian farmers.

Rare monogenic disorders are caused by mutations in single genes and have an incidence rate of less than 0.5%. Due to their low prevalence, these diseases often attract limited research and commercial interest, leading to significant unmet medical needs. In a therapeutic landscape where treatments are targeted to manage symptoms, gene editing therapy emerges as a promising approach to craft curative and lasting treatments for these patients, often referred to as "one-and-done" therapeutics. CRISPR-dependent base editing enables the precise correction of genetic mutations by direct modification of DNA bases without creating potentially deleterious DNA double-strand breaks. Base editors combine a nickase version of Cas9 with cytosine or adenine deaminases to convert C·G to T·A and A·T to G·C, respectively. Together, cytosine (CBE) and adenine (ABE) base editors can theoretically correct ∼95% of pathogenic transition mutations cataloged in ClinVar. This mini-review explores the application of base editing as a therapeutic approach for rare monogenic disorders. It provides an overview of the state of gene therapies and a comprehensive compilation of preclinical studies using base editing to treat rare monogenic disorders. Key considerations for designing base editing-driven therapeutics are summarized in a user-friendly guide for researchers interested in applying this technology to a specific rare monogenic disorder. Finally, we discuss the prospects and challenges for bench-to-bedside translation of base editing therapies for rare monogenic disorders.

Polyploidy, or whole-genome duplication (WGD), is a significant evolutionary force. Following allopolyploidy, duplicate gene copies (homeologs) have divergent evolutionary trajectories: some genes are preferentially retained in duplicate, while others tend to revert to single-copy status. Examining the effect of homeolog loss (i.e., changes in gene dosage) on associated phenotypes is essential for unraveling the genetic mechanisms underlying polyploid genome evolution. However, homeolog-specific editing has been demonstrated in only a few crop species and remains unexplored beyond agricultural applications. Tragopogon (Asteraceae) includes an evolutionary model system for studying the immediate consequences of polyploidy in nature. In this study, we developed a CRISPR-mediated homeolog-specific editing platform in allotetraploid T. mirus. Using the MYB10 and DFR genes as examples, we successfully knocked out the targeted homeolog in T. mirus (4x) without editing the other homeolog (i.e., no off-target events). The editing efficiencies, defined as the percentage of plants with at least one allele of the targeted homeolog modified, were 35.7% and 45.5% for MYB10 and DFR, respectively. Biallelic modification of the targeted homeolog occurred in the T0 generation. These results demonstrate the robustness of homeolog-specific editing in polyploid Tragopogon, laying the foundation for future studies of genome evolution following WGD in nature.

Tumor-associated antigen (TAA) loss remains a significant mechanism of resistance to chimeric antigen receptor (CAR) T cell therapy, leading to relapse in patients with B-cell malignancies and representing a major clinical challenge. Recent clinical data suggest that CD19 antigen loss triggers relapse in more than 40% of patients undergoing CD19 CAR-T cell therapy. To rigorously validate antigen loss, robust in vitro models that mimic the dynamic process of antigen escape are essential. However, the current absence of these models hampers our ability to fully evaluate and optimize treatment strategies. To model this clinically relevant phenomenon, we generated single (sKO), double (dKO), and triple (tKO) knockout Raji lymphoma cell lines targeting CD19, CD20, and CD22 using CRISPR/Cas9 genome editing. Initially, we established a dual-reporter cell line expressing the fluorescent marker mCherry and the bioluminescent marker Luciferase, enabling a uniform luminescence background across all the knockout cell lines before performing the CRISPR/Cas9 editing. The loss of individual or combinatorial TAAs was validated at the genomic, transcript, and protein levels. Functional co-culture assays with antigen-specific CAR-T cells showed that antigen-deficient Raji cells resisted CAR-T cell-mediated killing, closely mimicking clinical relapse. The triple knockout (tKO) model, in particular, provided a superior system compared to commonly used K562 models, as it retains the same lymphoma background while eliminating the crucial antigenic targets, thus better simulating resistance to CAR-T cell therapy. These antigen-loss models serve as valuable tools for studying mechanisms of CAR-T cell resistance and evaluating next-generation, multi-targeting CAR-T cell therapies.

Genome editing in melon (Cucumis melo L.) remains a significant challenge due to the inefficiencies associated with conventional cell culture-based transformation methods. In the present study, a novel in planta Particle Bombardment (iPB) approach was developed to enable DNA-free genome editing in melon without the need for cell culture. CRISPR/Cas9 ribonucleoproteins (RNPs) were coated onto gold particles and delivered directly into shoot apical meristem tissue, which harbors potential germline cells, via particle bombardment. This method was applied to enhance fruit shelf-life by targeting an ethylene biosynthesis gene (CmACO1). The resulting cmaco1 mutant demonstrated a significantly extended shelf-life, attributable to reduced ethylene production during fruit ripening. This delayed ripening phenotype was reversed upon treatment with exogenous ethylene, confirming the functional impact of CmACO1 disruption. Because this strategy bypasses cell culture, the iPB-RNP method offers a solution to common limitations in genome editing, such as genotype dependence and somaclonal variation. Consequently, this technique holds substantial promise for advancing commercial melon breeding efforts and may be broadly applicable to other species within the Cucurbitaceae family.

The pursuit of sustainable livestock farming to meet the rising global protein demand has positioned myostatin (MSTN) gene editing as a key technology. However, the field's focus on the remarkable double-muscle phenotype has often overshadowed a systematic examination of its concomitant effects. The present review aims to bridge this gap by moving beyond a singular focus on productivity. First, the pleiotropic effects of MSTN gene editing on growth performance, carcass quality, and meat quality in cattle, swine, sheep, poultry, and aquatic species were comprehensively evaluated. Next, the cascading biological effects of MSTN editing on metabolic homeostasis, reproductive performance, and animal health and welfare werAAe analyzed in depth. Finally, the inherent limitations and ethical issues of current editing techniques were critically discussed, and future sustainable breeding programs aimed at balanced multitrait regulation were prospectively proposed. Ultimately, this review affirms that MSTN editing has a multiplicative effect on trait alterations; however, there is also a series of associated health challenges, which demonstrates that the technology's impact is systemic, generating a spectrum of trade-offs that are often species specific. Its responsible application therefore hinges on multitrait balancing strategies to simultaneously secure productivity and sustainability in animal agriculture.

Red leaf lettuce (Lactuca sativa L. cv. 'Red Fire') is a preferred crop in plant factories with artificial light (PFALs) due to its short cultivation cycle and high anthocyanin content, which increases both its nutritional value and visual appeal. However, anthocyanins strongly influence leaf coloration and antioxidant profiles, and their levels are highly responsive to the light environment. Therefore, targeted editing of flavonoid biosynthesis may provide a breeding strategy to diversify pigment composition and associated functional traits under PFAL conditions. In this study, we used CRISPR/Cas9 to knock out DFR (dihydroflavonol 4-reductase), a key enzyme in the anthocyanin pathway. Genome-edited lines were generated via a dual-guide RNA system, resulting in a successfully edited red leaf genotype. The DFR-knockout lines displayed a complete loss of red pigmentation and a visibly distinct green phenotype. Metabolite profiling revealed a significant decrease in anthocyanin levels, accompanied by an increase in total flavonoid levels in some lines. Growth traits, including shoot dry weight and leaf number, were not significantly affected, suggesting that DFR knockout does not compromise growth under PFAL conditions. These findings highlight DFR as a promising target for creating pigment-altered lettuce lines for controlled-environment cultivation, including PFAL systems.