Genome editing tools have rapidly been adopted by plant scientists for crop improvement. Genome editing using a multiplex sgRNA-CRISPR/Cas9 genome editing system is a useful technique for crop improvement in monocot species. In this study, we utilized precise gene editing techniques to generate wheat 3'(2'), 5'-bisphosphate nucleotidase (TaSal1) mutants using a multiplex sgRNA-CRISPR/Cas9 genome editing system. Five active TaSal1 homologous genes were found in the genome of Giza168 in addition to another apparently inactive gene on chromosome 4A. Three gRNAs were designed and used to target exons 4, 5 and 7 of the five wheat TaSal1 genes. Among the 120 Giza168 transgenic plants, 41 lines exhibited mutations and produced heritable TaSal1 mutations in the M1 progeny and 5 lines were full 5 gene knock-outs. These mutant plants exhibit a rolled-leaf phenotype in young leaves and bended stems, but there were no significant changes in the internode length and width, leaf morphology, and stem shape. Anatomical and scanning electron microscope studies of the young leaves of mutated TaSal1 lines showed closed stomata, increased stomata width and increase in the size of the bulliform cells. Sal1 mutant seedlings germinated and grew better on media containing polyethylene glycol than wildtype seedlings. Our results indicate that the application of the multiplex sgRNA-CRISPR/Cas9 genome editing is efficient tool for mutating more multiple TaSal1 loci in hexaploid wheat.

The recent approval of the first gene editing therapy for sickle cell disease and transfusion-dependent beta-thalassemia by the U.S. Food and Drug Administration (FDA) demonstrates the immense potential of CRISPR (clustered regularly interspaced short palindromic repeats) technologies to treat patients with genetic disorders that were previously considered incurable. While significant advancements have been made with ex vivo gene editing approaches, the development of in vivo CRISPR/Cas gene editing therapies has not progressed as rapidly due to significant challenges in achieving highly efficient and specific in vivo delivery. Adeno-associated viral (AAV) vectors have shown great promise in clinical trials as vehicles for delivering therapeutic transgenes and other cargos but currently face multiple limitations for effective delivery of gene editing machineries. This review elucidates these challenges and highlights the latest engineering strategies aimed at improving the efficiency, specificity, and safety profiles of AAV-packaged CRISPR/Cas systems (AAV-CRISPR) to enhance their clinical utility.

Early diagnosis of hepatocellular carcinoma (HCC) is crucial for improving patient survival and treatment outcomes and the early detection of biomarkers for HCC is key to achieving this goal. However, conventional detection methods often lack sufficient specificity and sensitivity. In recent years, CRISPR/Cas12a-based biosensing has gained significant attention due to its ease of use and high sensitivity, demonstrating its potential to address the limitations of conventional detection methods. This paper primarily reviews the research progress of CRISPR/Cas12a-based biosensors for HCC detection, introducing their fluorescence, electrochemical, colorimetric, and other detection principles, as well as practical applications in detail. Additionally, the differences in sensitivity, specificity, and detection speed among different types of CRISPR/Cas12a biosensors are comparatively analyzed. Finally, the potential future directions for the development and application of CRISPR/Cas12a technology in clinical settings are explored.

Yeast Saccharomyces cerevisiae (S. cerevisiae) is a crucial industrial platform for producing a wide range of chemicals, fuels, pharmaceuticals, and nutraceutical ingredients. It is also commonly used as a model organism for fundamental research. In recent years, the CRISPR-Cas (clustered regularly interspaced short palindromic repeats-CRISPR-associated proteins) system has become the preferred technology for genetic manipulation in S. cerevisiae owing to its high efficiency, precision, and user-friendliness. This system, along with its extensive toolbox, has significantly accelerated the construction of pathways, enzyme optimization, and metabolic engineering in S. cerevisiae. Furthermore, it has allowed researchers to accelerate phenotypic evolution and gain deeper insights into fundamental biological questions, such as genotype-phenotype relationships. In this review, we summarize the latest advancements in the CRISPR-Cas toolbox for S. cerevisiae and highlight its applications in yeast cell factory construction and optimization, enzyme and phenotypic evolution, genome-scale functional interrogation, gene drives, and the advancement of biotechnologies. Finally, we discuss the challenges and potential for further optimization and applications of the CRISPR-Cas system in S. cerevisiae.

The trans-cleavage activity of the CRISPR-Cas system offers tremendous potential for developing highly sensitive and selective molecular diagnostic tools. However, conventional methods often face challenges such as limited catalytic efficiency of single Cas proteins and the necessity of complex multi-enzyme preamplification steps. To address these limitations, we present a novel defective PAM-mediated CRISPR-Cas12a self-catalytic signal amplification strategy, termed DEP-Cas-APE, for the rapid, sensitive, and specific detection of apurinic/apyrimidinic endonuclease 1 (APE1) activity. This approach integrates defective PAM-modified DNA probes to synergize Cas12a trans-cleavage with self-catalytic circuit, achieving efficient signal transformation and amplification under isothermal, one-step conditions. We systematically investigated the influence of defective PAM sequences containing apurinic/apyrimidinic (AP) sites on Cas12a activation and validated the feasibility of the DEP-Cas-APE strategy in detecting APE1. Under optimized conditions, DEP-Cas-APE achieved a detection limit as low as 7.66 × 10-8 U μL-1 within 30 min using a simple isothermal reaction. Additionally, we developed a point-of-care testing (POCT) platform by integrating DEP-Cas-APE with a colorimetric assay based on gold nanoparticles (AuNPs), enabling portable, equipment-free detection. This sensitive and selective strategy successfully detected APE1 in complex biological samples, including serum from lung cancer patients, and demonstrated the ability to distinguish cancerous from normal samples. DEP-Cas-APE represents a robust and versatile platform for advancing CRISPR-Cas12a biosensing technologies, offering new opportunities for molecular diagnostics and clinical research.

CRISPR-Cas9 and CRISPR-Cas12a are endonuclease systems widely used for genome editing, but their mechanisms of DNA cleavage, particularly in the presence of nucleotide mismatches, remain incompletely understood. This study deals with thermodynamic parameters governing the cleavage of DNA substrates-containing a mismatch in the region complementary to RNA-by Cas9 and Cas12a. Using a series of 55 bp DNA substrates with various mismatches, we investigated the cleavage efficiency, reaction kinetics, and thermodynamic stability of the Cas12a-crRNA complex and compared it with Cas9-sgRNA on the same substrates. Cas12a manifested strict specificity, with a mismatch leading to a significant reduction in cleavage efficiency or to nonspecific trans-cleavage, whereas Cas9 showed higher tolerance to each mismatch, especially in internal and distal regions. Thermodynamic calculations indicated that Cas12a-crRNA complexes are generally stabler with fully complementary DNA but are more destabilized by a mismatch than Cas9-sgRNA complexes are. Molecular dynamics simulations revealed that a mismatch causes greater structural destabilization in Cas12a than in Cas9, correlating with reduced cleavage efficiency. These findings highlight distinct mechanisms of mismatch recognition by Cas9 and Cas12a, provide insights into their enzymatic behavior, and inform the design of more precise genome-editing tools.

Filamentous fungi are essential industrial microorganisms that can serve as sources of enzymes, organic acids, terpenoids, and other bioactive compounds with significant applications in food, medicine, and agriculture. However, the underdevelopment of gene editing tools limits the full exploitation of filamentous fungi, which still present numerous untapped potential applications. In recent years, the CRISPR/Cas (clustered regularly interspaced short palindromic repeats) system, a versatile genome-editing tool, has advanced significantly and been widely applied in filamentous fungi, showcasing considerable research potential. This review examines the development and mechanisms of genome-editing tools in filamentous fungi, and contrasts the CRISPR/Cas9 and CRISPR/Cpf1 systems. The transformation and delivery strategies of the CRISPR/Cas system in filamentous fungi are also examined. Additionally, recent applications of CRISPR/Cas systems in filamentous fungi are summarized, such as gene disruption, base editing, and gene regulation. Strategies to enhance editing efficiency and reduce off-target effects are also highlighted, with the aim of providing insights for the future construction and optimization of CRISPR/Cas systems in filamentous fungi.

The harmful algal species Karlodinium veneficum (K. veneficum) poses a significant threat to aquatic ecosystems, economic stability, and human health due to its toxin production and widespread occurrence. Rapid climatic changes and eutrophication have intensified harmful algal blooms (HABs), making the timely detection of K. veneficum critical. To address this need, we developed a rapid and accurate detection method of K. veneficum by combining Recombinase Aided Amplification (RAA) with CRISPR/LbCas12a. This method targets the internal transcribed spacer (ITS) sequence of K. veneficum and utilizes the "collateral activity" of CRISPR/LbCas12a for visualization. Our method can detect plasmid DNA as low as 5.9 × 102 copies/µL and genomic DNA as low as 3.6 × 10-2 ng/µL, achieving a detection limit of 10 cells of K. veneficum through a simplified DNA extraction process. The entire detection process, from DNA crude extract to result visualization, can be completed in as fast as 90 min, making it suitable for field applications requiring a rapid response. In addition, our method was validated against a wide range of non-target microalgae species, confirming its specificity to K. veneficum and eliminating the risk of cross-reactivity. Overall, the RAA-CRISPR/LbCas12a system is simple, accurate, and sensitive, showing great potential for field applications in monitoring K. veneficum.

Aujeszky's disease (AD) is an acute infectious disease that infects pigs and other animals, resulting in significant economic losses and posing a threat to human health. Reliable and rapid detection methods are essential for the prevention of AD. In this study, a RAA-Cas12b assay based on recombinase-aided amplification (RAA) and CRISPR-Cas12b system was established, optimized and evaluated for the rapid detection of wild-type Pseudorabies Virus (PRV). The results can not only be detected by real-time fluorescence readout, but also can be visualized by a portable blue light instrument. There was no cross-reaction with PRV Bartha-K61 strain or other swine infectious viruses. The analytical sensitivities of the real-time PRV RAA-Cas12b assay and visual PRV RAA-Cas12b assay were determined to be 15 copies/μL with 95 % confidence interval and 140 copies/μL with 95 % confidence interval, respectively. A total of 31 clinical samples were detected and compared with PRV qPCR assay to evaluate the diagnostic performance of the PRV RAA-Cas12b assay. The diagnostic coincidence rate of the two assays was 100 %. In summary, this convenient and reliable assay has great potential for rapid detection of wild type PRV in point-of-care testing (POCT).

CRISPR-based diagnostics (CRISPR/Dx) have revolutionized the field of molecular diagnostics. It enables home self-test, field-deployable, and point-of-care testing (POCT). Despite the great potential of CRISPR/Dx in diagnoses of biologically complex diseases, preamplification of the template often is required for the sensitive detection of low-abundance nucleic acids. Various amplification-free CRISPR/Dx systems were recently developed to enhance signal detection at sufficient sensitivity. Broadly, these amplification-free CRISPR/Dx systems are classified into five groups depending on the signal enhancement strategies employed: CRISPR/Cas12a and/or CRISPR/Cas13a are integrated with: (1) other catalytic enzymes (Cas14a, Csm6, Argonaute, duplex-specific nuclease, nanozyme, or T7 exonuclease), (2) rational-designed oligonucleotides (multivalent aptamer, tetrahedral DNA framework, RNA G-quadruplexes, DNA roller machine, switchable-caged guide RNA, hybrid locked RNA/DNA probe, hybridized cascade probe, or "U" rich stem-loop RNA), (3) nanomaterials (nanophotonic structure, gold nanoparticle, micromotor, or microbeads), (4) electrochemical and piezoelectric plate biosensors (SERS nanoprobes, graphene field-effect transistor, redox probe, or primer exchange reaction), or (5) cutting-edge detection technology platforms (digital bioanalysis, droplet microfluidic, smartphone camera, or single nanoparticle counting). Herein, we critically discuss the advances, pitfalls and future perspectives for these amplification-free CRISPR/Dx systems in nucleic acids detection. The continued refinement of these CRISPR/Dx systems will pave the road for rapid, cost-effective, ultrasensitive, and ultraspecific on-site detection without resorting to target amplification, with the ultimate goal of establishing CRISPR/Dx as the paragon of diagnostics.

The Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR) sensing technology proved to be valuable during the COVID-19 pandemic through its sensitivity, specificity, robustness, and versatility. However, issues such as overreliance on amplification, susceptibility to false positives, lack of quantification strategies, and complex operation procedures have hindered its broader application in bioanalysis and clinical diagnostics. The collision between micronano-collaborative chips and CRISPR technology has effectively addressed these bottlenecks, offering innovative solutions for diagnosis and treatment. Unlike conventional micronano chips, micronano digital chips enhance CRISPR's response to trace amounts of target molecules by leveraging highly controllable local environments and compartmentalized microreactors. This advancement improves detection efficiency and revolutionizes traditional in vitro bioanalytical processes. First, the working principles, fabrication techniques, and performance metrics of CRISPR-based digital droplet microfluidics and microarray chips are examined. Then, the applications of CRISPR digital sensing chips in bioassays are reviewed, emphasizing their importance in advancing in vitro detection systems for gene editing. Finally, the prospects of CRISPR digital sensing technology are explored, particularly its potential for body surface biomonitoring and its broader development opportunities in the biomedical field.

The commencement of Highly Active Antiretroviral Therapy almost completely stopped viral replication, enabling the immune system to restore its full functionality. The rise in life expectancy has resulted in a decrease in the incidence of classical infections and HIV-associated cancers. HAART has raised concerns, including its exorbitant cost (which hinders its implementation in developing nations), the need for strict adherence, and the potential for both immediate and prolonged ill effects. Lipodystrophy is a significant long-term consequence of HIV that may result in central fat accumulation and severe peripheral fat depletion. Current initiatives to tackle these difficulties include the global expansion of access to HAART, the development of novel drugs that mitigate early side effects, and the introduction of once-daily drug combinations that enhance adherence. The CRISPR-Cas9 system has facilitated the creation of a powerful instrument for precise gene editing. This method has lately established itself as the gold standard for efficient HIV-1 genome editing in HIV therapy, owing to progress in related disciplines. CRISPR may be customized to cleave specific sequences by altering Cas9. This article offers a concise overview of promising CRISPR-Cas9 technology. This technique has the potential to halt the transmission of HIV-1 and alleviate its symptoms. CRISPR-Cas9 technology will be significant in the fight against HIV-1 in the future.

Precise identification and detection of single nucleotide variation (SNV) concomitant with excess wild-type DNA is greatly needed for invasive disease diagnosis, pathogens detection and early prediction of drug responsiveness. Many variants of the Severe Acute Respiratory Syndrome Coronavirus 2 (SARS-CoV-2), notably the D614G and N501Y mutations, have been shown to significantly increase the infectivity of pandemics. We herein investigated CRISPR/Cas12a integrated three types fluorescent reporters and two crRNAs for SNV detection by taking D614G and N501Y variants of SARS-CoV-2 as model examples. We systematically screened all possible base substitutions from positions 0 to 19 and identified the middle position of crRNA could efficiently increase the specificity from both theoretical and experimental standpoints. With selected mutation location of crRNA, we then investigated the specificity of ssDNA, dsDNA and molecular beacon (MB) fluorescent reporters and proved the MB reporters can efficiently increase the discriminatory factors. Furthermore, we designed an additional mutation site on crRNA to increase the specificity. For user convenience, we engineered the lateral flow strips to present the results visualized with the naked eyes. Results of specific variants from Omicron proved the feasibility of clinical applications. These findings indicated that the proposed method is a powerful tool for monitoring the key mutations in pathogens and allows for modifications to incorporate newer upcoming variants.

The advent of clustered regularly interspaced short palindromic repeats (CRISPR)-based technologies has revolutionized genome editing, with continued interest in expanding the CRISPR-associated proteins (Cas) toolbox with diverse, efficient, and specific effectors. CRISPR-Cas12a is a potent, programmable RNA-guided dual nickase, broadly used for genome editing. Here, we mined dairy cow microbial metagenomes for CRISPR-Cas systems, unraveling novel Cas12a enzymes. Using in silico pipelines, we characterized and predicted key drivers of CRISPR-Cas12a activity, encompassing guides and protospacer adjacent motifs for five systems. We next assessed their functional potential in cell-free transcription-translation assays with GFP-based fluorescence readouts. Lastly, we determined their genome editing potential in vivo in Escherichia coli by generating 1 kb knockouts. Unexpectedly, we observed natural sequence variation in the bridge-helix domain of the best-performing candidate and used mutagenesis to alter the activity of Cas12a orthologs, resulting in increased gene editing capabilities of a relatively inefficient candidate. This study illustrates the potential of underexplored metagenomic sequence diversity for the development and refinement of genome editing effectors.

CRISPR/Cas12a systems have emerged as versatile tools for molecular diagnostics, but directly detecting RNA and identifying specific DNA point mutations remain challenging. Herein, we report a simple engineering approach with a split site in the spacer sequence, enabling activation of CRISPR/Cas12a (LbCas12a) for trans-cleavage with similar efficiency to wild-type crRNA. The engineered crRNA facilitated RNA target recognition by replacing the 3'-end with RNA fragments, enhancing point mutation specificity for ssDNA targets. Based on this, we achieved amplification-free detection of microRNAs and DNA point mutations with high sensitivity and specificity. For clinical sample validation, we constructed reverse fluorescence-enhanced lateral flow test strips (rLFTS), which achieved femtomole-level detection. Moreover, the engineered crRNA-based CRISPR/Cas12a system also effectively recognized tumor cells via intracellular and in vivo imaging of miRNA-21. In conclusion, this engineered crRNA platform enhances CRISPR/Cas12a-based nucleic acid detection, promoting its wide application in molecular diagnostics and bioimaging.

The ability to robustly predict guide RNA (gRNA) activity is a long-standing goal for CRISPR applications, as it would reduce the need to pre-screen gRNAs. Quantification of formation of short insertions and deletions (indels) after DNA cleavage by transcribed gRNAs has been typically used to measure and predict gRNA activity. We evaluate the effect of chemically synthesized Cas9 gRNAs on different cellular DNA cleavage outcomes and find that the activity of different gRNAs is largely similar and often underestimated when only indels are scored. We provide a simple linear model that reliably predicts synthetic gRNA activity across cell lines, robustly identifies inefficient gRNAs across different published datasets, and is easily accessible via online genome browser tracks. In addition, we develop a homology-directed repair efficiency prediction tool and show that unintended large-scale repair events are common for Cas9 but not for Cas12a, which may be relevant for safety in gene therapy applications.

CRISPR/Cas12a, a promising gene editing technology, faces limitations due to its requirement for a thymine (T)-rich protospacer adjacent motif (PAM). Despite the development of Cas12a variants with expanded PAM profiles, many genomic loci, especially those with guanine-cytosine (GC)-rich PAMs, have remained inaccessible. This study develops a small RNA toxin-aided strategy to evolve ErCas12a for targeting GC-rich PAMs, resulting in the creation of enhanced ErCas12a (enErCas12a). EnErCas12a demonstrates the ability to recognize GC-rich PAMs and target five times more PAM sequences than the wild-type ErCas12a. Furthermore, enErCas12a achieves efficient gene editing in both bacterial and mammalian cells at various sites with non-canonical PAMs, including GC-rich PAMs such as GCCC, CGCC, and GGCC, which are inaccessible to previous Cas12a variants. Moreover, enErCas12a effectively targets PAM sequences with a GC content exceeding 75% in mammalian cells, providing a valuable alternative to the existing Cas12a toolkit. Importantly, enErCas12a maintains high specificity at targets with canonical PAMs, while also demonstrating enhanced specificity at targets with non-canonical PAMs. Collectively, this work establishes enErCas12a as a promising tool for gene editing in both eukaryotes and prokaryotes.

Eukaryotic transposon-encoded Fanzor proteins hold great promise for genome-engineering applications as a result of their compact size and mechanistic resemblance to TnpB. However, the unmodified Fanzor systems show extremely low activity in mammalian cells. Guided by the predicted structure of a Fanzor2 complex using AlphaFold3, we engineered the NlovFz2 nuclease and its cognate ωRNA to create an evolved enNlovFz2 system, with an expanded target-adjacent motif (TAM) recognition scope (5'-NMYG) and a substantially improved genome-editing efficiency, achieving an 11.1-fold increase over the wild-type NlovFz2, comparable to two previously reported IS200 or IS605 transposon-encoded TnpBs and two CRISPR-Cas12f1 nucleases. Notably, enNlovFz2 efficiently mediated gene disruption in mouse embryos and restored dystrophin expression in a humanized Duchenne muscular dystrophy mouse model with single adeno-associated virus delivery. Our findings underscore the potential of eukaryotic RNA-guided Fanzor2 nucleases as a versatile toolbox for both biological research and therapeutic applications.

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.

The development of highly efficient genome editing tools has revolutionized developmental biology and genetic studies in silkworm. Although methods based on CRISPR/Cas9 are currently popular, the Cas12a system has emerged as a promising option. However, it has not yet been applied to target the silkworm genome in vivo, and its activity in silkworm has not yet been characterized. In this study, we established a ribonucleoprotein-based CRISPR/Cas12a system, and compared it to the CRISPR/Cas9 system using 19 crRNA and 17 sgRNAs to target three different genes in vivo. Although Cas12a generates mutants less efficiently than Cas9, we used it successfully to generate transmissible indels, and demonstrated its application by targeting the FibH and mp genes to produce mutants with the expected phenotypes. We also assessed the influence of temperature (37 °C vs. 25 °C) on Cas12a activity, and demonstrated that the effects are target dependent. In summary, we have established a ribonucleoprotein-based CRISPR/Cas12a system in silkworm that offers a practical alternative to CRISPR/Cas9 and extends the genome editing tool box available for silkworm research.

Gene regulatory networks, which control gene expression patterns in development and in response to stimuli, use regulatory logic modules to coordinate inputs and outputs. One example of a regulatory logic module is the gene regulatory cascade (GRC), where a series of transcription factor genes turn on in order. Synthetic biologists have derived artificial systems that encode regulatory rules, including GRCs. Furthermore, the development of single-cell approaches has enabled the discovery of gene regulatory modules in a variety of experimental settings. However, the tools available for validating these observations remain limited. Based on a synthetic GRC using DNA cutting-defective Cas9 (dCas9), we designed and implemented an alternative synthetic GRC utilizing DNA cutting-defective Cas12a (dCas12a). Comparing the ability of these two systems to express a fluorescent reporter, the dCas9 system was initially more active, while the dCas12a system was more streamlined. Investigating the influence of individual components of the systems identified nuclear localization as a major driver of differences in activity. Improving nuclear localization for the dCas12a system resulted in 1.5-fold more reporter-positive cells and a 15-fold increase in reporter intensity relative to the dCas9 system. We call this optimized system the "Synthetic Gene Regulatory Network" (SGRN, pronounced "sojourn").

Clustered regularly interspaced short palindromic repeats/CRISPR-associated protein 12a (CRISPR/Cas12a) detection system is now widely used for nucleic acid detection and disease diagnosis. However, there are still fewer detections for single nucleotide polymorphisms (SNPs) and limited diversified detection systems for pathogen and SNP sites detection, which greatly limits their applications. Obviously, the development of a more diversified and convenient suite of detection tools is essential to unlock the full potential of CRISPR/Cas12a technology and to expand its applications across a wider range of scenarios. We have successfully developed an integrated CRISPR/Cas12a assay system. This system introduces crRNA during protein expression, reducing the number of steps and reaction time by adding only a fluorescent reporter gene and target DNA during subsequent detection. It enables on-site visualization of the assay in combination with a Recombinase polymerase amplification (RPA) reaction. Combined with the RPA reaction, we are able to rapidly detect African swine fever virus (ASFV) pathogens with high specificity. The system also enables genotyping of the SNP site of the porcine prolificacy-associated estrogen receptor (ESR) gene and the sheep prolificacy-associated Fecundity booroola (FecB) gene. Visualization is possible up to a final concentration of 3 nM, and effective differentiation of low concentrations within the concentration range of the assay. The integrated CRISPR/Cas12a assay system we developed has a robust design that ensures high-fidelity genotyping and pathogen detection are no longer restricted to the lab, allowing for rapid field analysis, which is crucial for timely interventions in agricultural and clinical settings. In addition, it has the advantages of low cost, easy operation and visualization of results.

Characterization of clustered regularly interspaced short palindromic repeat (CRISPR)-associated (Cas) trans-cleavage activities has initiated the era of next-generation CRISPR diagnostics. By using the trans-cleavage reaction for signal output, CRISPR systems have been engineered to detect non-nucleic acids (NNAs), including ions, inorganic small molecules, organic compounds, proteins, and bacteria. Diverse strategies are being used to specifically recognize NNAs and regulate Cas trans-cleavage activities, via generation or depletion of output signals. In this review, we introduce the principles and advantages of CRISPR-based NNA detection. We then classify CRISPR-based NNA detection strategies into three classes: the generation or depletion of free activators, synthesis of crRNAs, and reconstruction of active Cas effectors. Finally, we discuss the challenges and potential strategies to advance both clinical and nonclinical applications of CRISPR-based NNA detection.

Type V-H CRISPR-Cas system, an important subtype of type V CRISPR-Cas systems, has remained enigmatic in terms of its structure and function despite being discovered several years ago. Here, we comprehensively characterize the type V-H CRISPR-Cas system and elucidate its role as a DNA nicking system. The unique CRISPR RNA (crRNA) employed by Cas12h effector protein enables specific targeting of double-stranded DNA (dsDNA), while its RuvC domain is responsible for cleaving the non-target strand (NTS) of dsDNA. We present the structure of Cas12h bound to crRNA and target DNA. Our structural analysis reveals that the RuvC domain possesses a narrow active pocket that facilitates recognition of NTS but potentially hinders access to the target strand. Furthermore, we demonstrate that Cas12h confers adaptive immunity against invading mobile genetic elements through transcriptional gene inhibition. We have engineered an adenine base editor by fusing Cas12h with an adenine deaminase, achieving effective A-to-G substitution.

PfAgo, a thermophilic Argonaute nuclease from Pyrococcus furiosus, is widely used in various fields due to its high DNA-guided DNA cleavage activity. However, its high-temperature-dependent cleavage activity largely restricts its applications in moderate-temperature scenarios. In this study, PfAgo is engineered for cold adaptation based on its ternary complex structure and the attributes of cold-adapted enzymes, yielding a series of variants with better performance at moderate temperatures. Among those, mPfAgo (K617G, L618G) exhibits significantly promoted cleavage activity at 37 °C and a wider catalytic temperature range of 30-95 °C. Its high-temperature cleavage activity is also greatly improved, enabling its application in DNA detection with attomolar sensitivity in the presence of Mg2+. Additionally, mPfAgo shows versatile cleavage activities, including DNA cleavage guided by 5'OH-gDNA, 5'P-gDNA, or 5'COOH-gDNA, as well as RNA cleavage with 5'OH-gDNA, 5'P-gDNA, 5'P-gRNA, or 5'COOH-gDNA as guides. Further analysis through far-UV CD spectra and DSF indicates that mPfAgo has a more flexible structure than wild-type PfAgo. Furthermore, this established strategy is applied to engineer TtdAgo, likewise obtaining its variants with enhanced moderate-temperature activity and expanded substrate spectrum. In summary, this work provides a novel method for the rational design of thermophilic Agos, thereby greatly expanding their application scopes.

Cyanobacteria of the genera Synechocystis and Synechococcus have emerged as promising platforms for metabolic engineering endeavors aimed at converting carbon dioxide into valuable fuels and chemicals, thus addressing the pressing energy demand and mitigating global climate change. Notably, Synechocystis sp. strain PCC 6803 (Synechocystis) has been engineered to produce isobutanol (IB) and 3-methyl-1-butanol (3M1B) via heterologous expression of α-ketoisovalerate decarboxylase (Kivd). Despite these advances, the achieved IB/3M1B titers remain low. CRISPR interference (CRISPRi), an emerging tool for targeted gene repression, has demonstrated success in various cellular systems to enhance biochemical productivity.

In this study, we developed a dCas12a-mediated CRISPRi system (CRISPRi-dCas12a) that effectively blocked the transcriptional initiation/elongation of essential gene(s), resulting in up to 60% gene repression in Synechocystis. Subsequently, the CRISPRi-dCas12a system was successfully integrated into an IB/3M1B producer strain, where it exhibited target gene repression under optimal cultivation conditions. To identify gene targets involved in metabolic pathways potentially limiting IB/3M1B biosynthesis, we initially designed a CRISPR RNA (crRNA) library targeting fifteen individual gene(s), where repression of ten genes significantly increased IB/3M1B production per cell. Moreover, a synergetic effect was observed on IB/3M1B production by designing a single crRNA targeting multiple genes for simultaneous repression. A final strain HX106, featuring dual repression of ppc and gltA, both involved in the TCA cycle, resulted in 2.6-fold and 14.8-fold improvement in IB and 3M1B production per cell, respectively.

Our findings underscore the effectiveness of the CRISPRi-dCas12a system in Synechocystis for identifying competing pathways and redirecting carbon flux to enhance IB/3M1B production. Furthermore, this study established a solid groundwork for utilizing an expanded CRISPRi-crRNA library to undertake genome-wide exploration of potential competing pathways not only for IB/3M1B biosynthesis but also for other diverse biofuels and biochemical production processes.

CRISPR-mediated endogenous tagging is a powerful tool in biological research. Inhibiting the non-homologous end joining (NHEJ) pathway has been shown to improve the low efficiency of accurate knock-in via homology-directed repair (HDR). However, the influence of alternative double-stranded break (DSB) repair pathways on knock-in remains to be fully explored. In this study, our long-read amplicon sequencing analysis reveals various patterns of imprecise repair in CRISPR-mediated knock-in, even with NHEJ inhibition. Further suppressing either microhomology-mediated end joining (MMEJ) or single-strand annealing (SSA) reduces nucleotide deletions around the cut site, thereby elevating knock-in accuracy. Additionally, imprecise donor integration is reduced by inhibiting SSA, but not MMEJ. Particularly, SSA suppression reduced asymmetric HDR, a specific imprecise integration pattern, which we further confirm using a novel reporter system. These findings demonstrate the complex interplay of multiple DSB repair pathways in CRISPR-mediated knock-in and offer novel strategies, including SSA pathway targeting, to improve precise gene editing efficiency.

The advent of genome-editing technologies, particularly the RNA-guided the clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated system (Cas) 9, which originates from prokaryotic CRISPR/Cas adaptive immune mechanisms, has revolutionized molecular biology. Renowned for its simplicity, cost-effectiveness, and capacity for multiplexed gene editing, CRISPR/Cas9 has emerged as the most versatile and widely adopted genome-editing platform. Its applications span fundamental research, biotechnology, medicine, and therapeutics. This review highlights recent advancements in CRISPR-based technologies, focusing on CRISPR/Cas9, CRISPR/Cas12a, and CRISPR/Cas12f. It emphasizes precision editing methods like base editing and prime editing, which enable targeted nucleotide changes without double-strand breaks. The specificity of these tools, including on-target accuracy and off-target risks, is critically evaluated. Additionally, recent preclinical and clinical efforts to treat diseases such as cancer and sickle cell disease using CRISPR are summarized. Finally, the challenges and future directions of CRISPR-mediated gene therapy are discussed, emphasizing its potential to integrate with other molecular approaches to address unmet medical needs.

Gene therapy development, re-engineering, and application to patients hold promise to revolutionize medicine, including therapies for disorders of the brain. Advances in delivery modalities, expression regulation, and improving safety profiles are of critical importance. Additionally, each inherited disorder has its own unique characteristics as to regions and cell types impacted and the temporal dynamics of that impact that are essential for the design of therapeutic design strategies. Here, we review the current state of the art in gene therapies for inherited brain disorders, summarizing key considerations for vector delivery, gene addition, gene silencing, gene editing, and epigenetic editing. We provide examples from animal models, human cell lines, and, where possible, clinical trials. This review also highlights the various tools available to researchers for basic research questions and discusses our views on the current limitations in the field.

A photo-triggered CRISPR/Cas12a machinery for in vitro and in vivo gene editing is introduced. The system consists of a caged, inactive ortho-nitrobenzyl phosphate ester photo-responsive crRNA, which, upon light-induced deprotection, yields the active CRISPR/Cas12a gene editing machinery (LAC12aGE). The LAC12aGE system induces specific thymidine-rich (TTTN) protospacer-adjacent motif (PAM)-guided double-stranded breaks in genomic DNA, which upon non-homologous end-joining lead to gene repair. The LAC12aGE machinery is applied for gene editing of an exogenous dual fluorescent reporter gene in living cells, as well as the endogenous gene encoding DNA methyltransferase 1. In addition, the LAC12aGE is applied for in vitro gene editing and disruption of the hepatocyte growth factor (HGF) gene in HepG2 cells, where knockout of the HGF gene results in inhibited cell proliferation and migration, as well as enhanced apoptosis. Moreover, the in vivo knockout and disruption of the HGF gene in HepG2 tumors by the LAC12aGE machinery is demonstrated. The cyclic temporal development of the LAC12aGE system in tumors shows effective inhibition of tumor growth and enhanced apoptosis/necrosis of tumor tissues compared to control systems.

For centuries, a competitive evolutionary race between prokaryotes and related phages or other mobile genetic elements has led to the diversification of Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR) and CRISPR‑associated sequence (Cas) genome‑editing systems. Among the different CRISPR/Cas systems, the CRISPR/Cas9 system has been widely studied for its precise DNA manipulation; however, due to certain limitations of direct DNA targeting, off‑target effects and delivery challenges, researchers are looking to perform transient knockdown of gene expression by targeting RNA. In this context, the more recently discovered type VI CRISPR/Cas13 system, a programmable single‑subunit RNA‑guided endonuclease system that has the capacity to target and edit any RNA sequence of interest, has emerged as a powerful platform to modulate gene expression outcomes. All the Cas13 effectors known so far possess two distinct ribonuclease activities. Pre‑CRISPR RNA processing is performed by one RNase activity, whereas the two higher eukaryotes and prokaryotes nucleotide‑binding domains provide the other RNase activity required for target RNA degradation. Recent innovative applications of the type VI CRISPR/Cas13 system in nucleic acid detection, viral interference, transcriptome engineering and RNA imaging hold great promise for disease management. This genome editing system can also be employed by the Specific High Sensitivity Enzymatic Reporter Unlocking platform to identify any tumor DNA. The discovery of this system has added a new dimension to targeting, tracking and editing circulating microRNA/RNA/DNA/cancer proteins for the management of cancer. However, there is still a lack of thorough understanding of the mechanisms underlying some of their functions. The present review summarizes the recent updates on the type VI CRISPR/Cas system in terms of its structural and mechanistic properties and some novel applications of this genome‑editing tool in cancer management. However, some issues, such as collateral degradation of bystander RNA, impose major limitations on its in vivo application. Furthermore, additional challenges and future prospects for this genome editing system are described in the present review.

CRISPR-Cas genome editing technology has transformed genetic research, by enabling unprecedented precision in modifying DNA sequences across various organisms, including fish. This review explores the significant advancements and potential uses of CRISPR-Cas technology in the study and management of fish diseases, which pose serious challenges to aquaculture and wild fish populations. Fish diseases cause significant economic losses and environmental impacts, therefore effective disease control a top priority. The review highlights the pivotal role of CRISPR-Cas in identifying disease-associated genes, which is critical to comprehending the genetic causes of disease susceptibility and resistance. Some studies have reported key genetic factors that influence disease outcomes, using targeted gene knockouts and modifications to pave the way for the development of disease-resistant fish strains. The creation of such genetically engineered fish holds great promise for enhancing aquaculture sustainability by reducing the reliance on antibiotics and other conventional disease control measures. In addition, CRISPR-Cas has facilitated in-depth studies of pathogen-host interactions, offering new insights into the mechanisms by which pathogens infect and proliferate within their hosts. By manipulating both host and pathogen genes, this technology provides a powerful tool for uncovering the molecular underpinnings of these interactions, leading to the development of more effective treatment strategies. While CRISPR-Cas has shown great promise in fish research, its application remains limited to a few species, primarily model organisms and some freshwater fish. In addition, challenges such as off-target effects, ecological risks, and ethical concerns regarding the release of genetically modified organisms into the environment must be carefully addressed. This review also discusses these challenges and emphasizes the need for robust regulatory frameworks and ongoing research to mitigate risks. Looking forward, the integration of CRISPR-Cas with other emerging technologies, such as multi-omics approaches, promises to further advance our understanding and management of fish diseases. This review concludes by envisioning the future directions of CRISPR-Cas applications in fish health, underscoring its potential to its growing in the field.

The CRISPR/Cas technology has demonstrated revolutionary potential across various fields, including agriculture, medicine, and food safety detection. However, the utility of CRISPR/Cas12a, a particularly promising gene-editing tool, is constrained by its temperature sensitivity, limiting its application in low-temperature environments. In this study, we developed a gene-editing technique based on the CRISPR/Cas12a system in the poikilothermic species goldfish Carassius auratus. We systematically evaluated the editing efficiencies of LbCas12a and AsCas12a on the tyrosinase (tyr) gene under varying temperature conditions. Our results revealed a pronounced temperature dependence of Cas12a, with elevated temperatures markedly enhancing its editing activity, particularly for AsCas12a. A brief one-hour high-temperature treatment was sufficient to achieve effective gene disruption, underscoring CRISPR/Cas12a as a rapid and efficient gene-editing tool. Temperature was utilized as a conditional trigger for Cas12a-mediated gene knockout, enabling precise modulation of gene disruption at specific embryonic developmental stages. Whole-genome resequencing of the mutants confirmed the absence of off-target effects, further emphasizing the precision of this editing process. These findings indicated that CRISPR/Cas12a represented a viable alternative to the widely utilized CRISPR/Cas9 system and could be applied in conjunction, thereby expanding the potential applications of gene-editing technologies.

Genome-edited plants, endowed with climate-smart traits, have been promoted as tools for strengthening resilience against climate change. Successful plant gene editing (GE) requires precise regulation of the GE machinery, a process controlled by the promoters, which drives its transcription through interactions with transcription factors (TFs) and RNA polymerase. While constitutive promoters are extensively used in GE constructs, their limitations highlight the need for alternative approaches. This review emphasizes the promise of tissue/organ specific as well as inducible promoters, which enable targeted GE in a spatiotemporal manner with no effects on other tissues. Advances in synthetic biology have paved the way for the creation of synthetic promoters, offering refined control over gene expression and augmenting the potential of plant GE. The integration of these novel promoters with synthetic systems presents significant opportunities for precise and conditional genome editing. Moreover, the advent of bioinformatic tools and artificial intelligence is revolutionizing the characterization of regulatory elements, enhancing our understanding of their roles in plants. Thus, this review provides novel insights into the strategic use of promoters and promoter editing to enhance the precision, efficiency and specificity of plant GE, setting the stage for innovative crop improvement strategies.

CRISPR-Cas9 has been widely used in research and medical investigations as a pioneering technology. However, challenges such as the large size of the Cas9 sequence and the need for precise control over its activity in specific cell types have impeded its widespread adoption. Various alternatives, such as split-Cas9 technology, have emerged. Split-Cas9 systems allow the large Cas9 sequence to be divided into two segments to aid in the delivery of the enzyme. Nevertheless, challenges persist in achieving precise control over the timing and location of Cas9 reassembly and activity to ensure targeted action. This study presents an enzymatic-based split-Cas9 system, introducing a new approach utilizing the Sortase enzyme for the reconstitution of the full Cas9 protein. The developed method eliminates the need for chemical or physical induction and allows for precise genome editing in specific cells through the utilization of various specific promoters or targeted drug delivery. Experimental validation of the enzymatic system was conducted in E. coli, HEK cells, and Jurkat cells, demonstrating successful assembly and activity of the assembled Cas9 enzyme. In addition, this study explored the incorporation of nuclear localization signals, the evaluation of inducible promoters, and the delivery of the system's components in mRNA or protein form. Furthermore, we investigated the potential of S/MAR minicircle technology instead of viral vectors within the system. Overall, we highlighted the feasibility and utility of the Sortase-based split-Cas9 system to enhance control and efficiency compared to traditional CRISPR-Cas9 approaches. Additionally, this study revealed the potential of using the Sortase enzyme for posttranslational modifications and protein assembly in human cells.

Clustered regularly interspaced short palindromic repeats (CRISPR) tools are revolutionizing the establishment of genotype-phenotype relationships and are transforming cell- and gene-based therapies. In the field of oncology, CRISPR/CRISPR-associated protein 9 (Cas9), Cas12, and Cas13 have advanced the generation of cancer models, the study of tumor evolution, the identification of target genes involved in cancer growth, and the discovery of genes involved in chemosensitivity and resistance. Moreover, preclinical therapeutic strategies employing CRISPR/Cas have emerged. These include the generation of chimeric antigen receptor T (CAR-T) cells and engineered immune cells, and the use of precision anticancer gene-editing agents to inactivate driver oncogenes, suppress tumor support genes, and cull cancer cells in response to genetic circuit output. This review summarizes the collective impact that CRISPR technology has had on basic and applied cancer research, and highlights the promises and challenges facing its clinical translation.

CRISPR-Cas12a is a powerful tool in nucleic acid detection, but the relationship between its trans-cleavage activity and protospacer adjacent motif (PAM) sequences remains incompletely understood. In this study, we synthesized diverse PAM-sequence substrates and conducted systematic cis-cleavage and trans-cleavage experiments with three Cas12a orthologs. We found that double-stranded DNA (dsDNA) can activate Cas12a's trans-cleavage activity even without PAM and this activation occurring independently of cis-cleavage. Notably, our results also revealed that single-stranded RNA (ssRNA) can directly initiate the trans-cleavage activity of Cas12a.We also experimentally validated the feasibility of CRISPR-Cas12a in detecting target dsDNA lacking PAM sequences, including identifying mutated sites in clinical samples. Structural prediction using AlphaFold 3 revealed the potential mechanism of Cas12a's PAM-independent trans-cleavage. Our research expands the understanding of Cas12a's trans-cleavage mechanism and demonstrates its potential for nucleic acid detection beyond PAM-dependent targets. This discovery broadens the application scope of Cas12a, providing new opportunities for developing highly sensitive and versatile diagnostic platforms.

RNA-guided systems provide remarkable versatility, enabling diverse biological functions. Through iterative structural and sequence homology-based mining starting with a guide RNA-interaction domain of Cas9, we identified a family of RNA-guided DNA-targeting proteins in phage and parasitic bacteria. Each system consists of a tandem interspaced guide RNA (TIGR) array and a TIGR-associated (Tas) protein containing a nucleolar protein (Nop) domain, sometimes fused to HNH (TasH)- or RuvC (TasR)-nuclease domains. We show that TIGR arrays are processed into 36-nucleotide RNAs (tigRNAs) that direct sequence-specific DNA binding through a tandem-spacer targeting mechanism. TasR can be reprogrammed for precise DNA cleavage, including in human cells. The structure of TasR reveals striking similarities to box C/D small nucleolar ribonucleoproteins and IS110 RNA-guided transposases, providing insights into the evolution of diverse RNA-guided systems.

This work comprehensively explores the effects of various positions of Cas12a split activators. Based on this, a multi-functional synergistic platform was constructed. The construction of a structural dynamic network, the sensitive detection of APE1, and a time-controlled photo-activation have been achieved, demonstrating the potential for expanding Cas12a applications.

Nanomaterials-integrated CRISPR/Cas systems have rapidly emerged as powerful next-generation platforms for optical biosensing. These integrated platforms harness the precision of CRISPR/Cas-mediated nucleic acid detection while leveraging the unique properties of nanomaterials to achieve enhanced sensitivity and expanded analytical capabilities, thereby broadening their diagnostic potential. By incorporating a diverse range of nanomaterials, these systems effectively expand the analytical toolbox for optical detection, offering adaptable solutions tailored to various diagnostic challenges. This review provides a comprehensive overview of the nanomaterials successfully integrated into CRISPR/Cas-based optical sensing systems. It examines multiple optical detection modalities, including fluorescence, electrochemiluminescence, colorimetry, and surface-enhanced Raman spectroscopy, highlighting how nanomaterials facilitate signal amplification, enable multiplexing, and support the development of point-of-care applications. Additionally, practical applications of these integrated systems in critical fields such as healthcare diagnostics and environmental monitoring are showcased. While these platforms offer considerable advantages, several real-world challenges such as the complexity of assay workflows, environmental impact of nanomaterials, cost, and regulatory hurdles must be addressed before widespread implementation can be achieved. By identifying these critical obstacles and proposing strategic solutions, we aim to pave the way for the continued advancement and adoption of nanomaterial-integrated CRISPR/Cas optical biosensing technologies.

Amyotrophic lateral sclerosis (ALS) is a neurodegenerative disease characterized by the selective death of motor neurons, which causes muscle atrophy. Genetic forms of ALS are recorded only in 10% of cases. However, over the past decade, studies in genetics have substantially contributed to our understanding of the molecular mechanisms underlying ALS. The identification of key mutations such as SOD1, C9orf72, FUS, and TARDBP has led to the development of targeted therapy that is gradually being introduced into clinical trials, opening up a broad range of opportunities for correcting these mutations. In this review, we aimed to present an extensive overview of the currently known mechanisms of motor neuron degeneration associated with mutations in these genes and also the gene therapy methods for inhibiting the expression of their mutant proteins. Among these, antisense oligonucleotides, RNA interference (siRNA and miRNA), and gene-editing (CRISPR/Cas9) methods are of particular interest. Each has shown its efficacy in animal models when targeting mutant genes, whereas some of them have proven to be efficient in human clinical trials.

The CRISPR-Cas12a system is a groundbreaking tool widely used for genome editing and diagnostics in biotechnology and biomedicine research laboratories. Despite its growing application, studies optimizing Cas12a protein production at the laboratory scale using straightforward protocols remains scarce. This study aimed to enhance the lab-scale recombinant production of Acidaminococcus sp Cas12a protein (AsCas12a) in E. coli. Through targeted adjustments of simple parameters, AsCas12a production was significantly increased. The optimized conditions included the use of E. coli BL21(DE3), TB medium supplemented with 1 % glucose, induction with 0.3 mM IPTG for at least 6-9 h, and incubation at 30 °C. Notably, these conditions differ from conventional protocols typically used for Cas12a and related proteins, such as Streptococcus pyogenes Cas9. Upon combining all optimized parameters, AsCas12a production increased approximately 3-fold, from 0.95 mg/mL of bacterial lysate under non-optimized conditions to 3.73 mg/mL under optimized ones. After chromatographic purification, the final protein yield rose approximately 4.5-fold, from 5.2 to 23.4 mg/L of culture volume, without compromising functional activity. The purified AsCas12a retained full activity for programmable in vitro DNA cis-cleavage and collateral trans-cleavage, which was successfully applied to detect the N gene of SARS-CoV-2. This optimized method provide an efficient and high-yield approach for producing functional AsCas12a protein using accessible materials and conditions available to many research laboratories worldwide.

CRISPR technologies have revolutionized strain engineering of Aspergillus species, and drastically increased the ease and speed at which genomic modifications can be performed. One of the advantages of CRISPR technologies is the possibility of rapid strain engineering using multiplex experiments. This can be achieved by using a set of different guiding RNA molecules (gRNA) to target multiple loci in the same experiment. Two major challenges in such experiments are firstly, the delivery of multiple guides simultaneously, and secondly, ensuring that each target locus is cut efficiently by the CRISPR nuclease. The CRISPR nuclease Cas12a, also known as Cpf1, presents a unique advantage to bypass this challenge. Specifically, and unlike Cas9, Cpf1 is able to release several gRNAs from a common precursor RNA molecule through its own RNase activity, eliminating the need for elements such as ribozymes or tRNA machinery for gRNA maturation. This feature sets the stage for much more straightforward construction of vectors for the delivery of many gRNAs, which in turn allows each locus to be targeted by multiple gRNAs to increase the odds of successfully inducing a break in the DNA.

Here we present a toolbox that can be used to assemble plasmids containing a gRNA multiplex expression cassette, which is able to express a multi gRNA precursor. The precursor can be processed via Cpf1 RNase activity to produce multiple functional gRNAs in vivo. Using our setup, we have constructed plasmids that are able to deliver up to ten gRNAs. In addition, we show that three simultaneous deletions can be introduced robustly in Aspergillus niger by targeting each gene with several gRNAs, without prior gRNA validation or the use of genomically integrated selection markers.

In this study we have established an efficient system for the construction of CRISPR-Cpf1 vectors that are able to deliver a large number of gRNAs for multiplex genome editing in Aspergillus species. Our strategy allows multiple specific genomic modifications to be performed in a time frame of less than two weeks, and we envision this will be able to speed up cell factory construction efforts significantly.

Prompt and accurate diagnosis of infectious pathogens of livestock origin is of utmost importance for epidemiological surveillance and effective therapeutic strategy formulation. Among various methods, nucleic acid-based detection of pathogens is the most sensitive and specific; but the majority of these assays need expensive equipment and skilled workers. Due to the rapid advancement of clustered regularly interspaced short palindromic repeats-CRISPR-associated protein (CRISPR-Cas)-based nucleic acid detection methods, these are now being widely used for pathogen detection. CRISPR-Cas is a bacterial counterpart of "adaptive immunity", generally used for editing genome. Many CRISPR systems have been modified for nucleic acid detection due to their excellent selectivity in detecting DNA and RNA sequences. The combination of CRISPR with suitable isothermal amplification technologies has made it more sensitive, specific, versatile, and reproducible for the detection of pathogen nucleic acids at the point of care. Amplification of pathogen nucleic acid by isothermal amplification followed by CRISPR-Cas-based detection has several advantages, including short sample-to-answer times and no requirement for laboratory set-up. They are also significantly less expensive than the existing nucleic acid detection methods. This review focuses on the recent trends in the use of this precision diagnostic method for diagnosis of a wide range of animal pathogens with or without zoonotic potential, particularly various isothermal amplification strategies, and visualization methods for sensing bacteria, viruses, and parasites of veterinary and public health importance.