DNA epigenetic modifications play crucial roles in regulating gene expression and cellular function across diverse organisms. Among them, 5-glyceryl-methylcytosine (5gmC), a unique DNA modification first discovered in Chlamydomonas reinhardtii, represents a novel link between redox metabolism and epigenetic regulation. Accurate genome-wide detection of 5gmC is essential for investigating its biological functions, yet no streamlined method has been available. Here, we present deaminase-assisted sequencing (DEA-seq), a simple and robust approach for base-resolution mapping of 5gmC. DEA-seq employs a single DNA deaminase that efficiently converts unmodified cytosines (C) and 5-methylcytosine (5mC) into uracils or thymines, while leaving 5gmC intact. This selective resistance generates a clear sequence signature that enables precise identification of 5gmC sites across the genome. The method operates under mild reaction conditions and is compatible with low-input DNA, minimizing sample loss and improving detection sensitivity. Overall, DEA-seq provides an accessible, efficient, and highly accurate protocol for profiling 5gmC, offering clear advantages in workflow simplicity, DNA integrity, and analytical performance.

This article presents an efficient protocol for refolding recombinant proteins that are prone to aggregation and form inclusion bodies during expression in Escherichia coli. As a model system, the homolog of CRISPR-associated effector protein CasV-M was investigated. The key element of the developed approach is refolding directly on a metal-affinity Ni-TED (N,N,N´-tris(carboxymethyl)ethylendiamine) resin using a dual-gradient system: a stepwise reduction in the concentration of the chaotropic agent combined with a simultaneous increase in the concentration of a mild nonionic detergent. This combination ensures spatial separation of protein molecules, minimizes aggregation, and promotes the recovery of the native conformation. The resulting method appears to be an alternative to conventional refolding strategies, with potential improvements in the reproducibility and yield of soluble protein compared to dialysis or dilution. The proposed approach can be extended to a broad range of aggregation-prone proteins and is considered a promising strategy for obtaining otherwise insoluble recombinant proteins.

Lipid peroxidation (LPO) is a major indicator of oxidative stress and cellular damage, frequently associated with environmental and toxicological stressors and mechanistically linked to ferroptotic regulated cell death (RCD). This protocol describes a simple and reproducible method for the qualitative in situ visualization of LPO in mosquito larvae using Schiff’s reagent, which histochemically labels reactive aldehyde groups [such as malondialdehyde (MDA)] generated during lipid degradation. Although Schiff’s reagent detects aldehydes commonly associated with lipid peroxidation, these compounds are not exclusive to LPO and may also arise from other oxidative processes. The method preserves tissue integrity, enabling direct, spatially resolved observation of oxidative damage in whole larvae. Following staining, larvae are rinsed in a stabilizing sulfite solution to maintain the characteristic magenta coloration. Using this assay, Culex quinquefasciatus larvae exposed to ferroptotic cyanobacteria, such as Synechocystis sp., exhibit a marked accumulation of lipid-derived aldehydes consistent with lipid ROS–mediated damage. This oxidative response is specifically suppressed by pre-treatment with the canonical ferroptosis inhibitor Ferrostatin-1 (Fer-1), which inhibits lipid peroxidation and significantly reduces larval mortality. As a complementary approach to traditional spectrophotometric assays such as thiobarbituric acid reactive substances (TBARS), this qualitative method enables in situ visualization of lipid peroxidation without tissue homogenization, providing a rapid and biologically informative screening tool for assessing ferroptosis-associated oxidative damage in Cx. quinquefasciatus and other biological models exposed to multiple stressors.

Protoplast systems are widely used in plant research as versatile platforms for studying cellular processes and validating gene editing tools. In maize, they are particularly valuable because stable transformation in immature embryos is slow and labor-intensive, often requiring months to regenerate plants. However, existing protocols often yield inconsistent results in protoplast recovery, transfection efficiency, and viability. We present an optimized protocol for maize mesophyll protoplast isolation and PEG-mediated transfection. Two-week-old etiolated seedlings are processed using vertical cutting, improving the yield and viability of protoplasts. Protoplasts are then immediately transformed with a CRISPR/Cas9 construct after isolation, via PEG4000 with only 10 μg of plasmid DNA, reducing the resource demands of standard methods. Modified washing and storage conditions extend transformed protoplast viability to seven days, enabling longer-term monitoring and expanded downstream analyses. Editing outcomes are quantified by sequencing target sites and calculating efficiency with Cas-Analyzer. This protocol provides a rapid, efficient, and reproducible method for the rapid evaluation of gene editing in maize. This protocol offers a methodology to accelerate agricultural crop studies and broader plant molecular biology.

Breast cancer remains one of the most prevalent and deadly malignancies affecting women worldwide. Its progression and metastatic behavior are driven by complex mechanisms. To develop more effective therapeutic strategies, it is crucial to understand tumor growth, angiogenesis, and microenvironmental interactions. Although traditional in vivo models such as murine xenografts have long been used to study tumor biology, these approaches are often time-consuming, costly, and ethically constrained. In contrast, the chick embryo chorioallantoic membrane (CAM) assay offers a rapid, cost-effective, and ethically flexible alternative for evaluating tumor development and angiogenesis. This protocol describes an in ovo CAM-based xenograft model in which human breast cancer cells are implanted onto the vascularized CAM of chick embryos. This method enables real-time evaluation of tumor growth. Furthermore, the model allows for manipulation of experimental conditions, including pharmacological treatments or genetic modifications, to study specific molecular mechanisms involved in breast cancer progression. The major advantages of this protocol lie in its simplicity, reduced cost, and capacity for high-throughput screening, making it a valuable tool for translational cancer research.

A prompt and accurate diagnosis of respiratory viral diseases in intensive poultry production is essential to safeguard animal health and ensure the economic sustainability of farms. Currently, much effort is being devoted to preventing the spread of the avian influenza virus in farms. However, the diagnosis of other relevant respiratory viruses, as infectious laryngotracheitis virus (ILTV), is also crucial. Indeed, infection by ILTV does lead to substantial economic losses due to high morbidity, reduced growth, and decreased productivity, making rapid detection a critical aspect of disease control. Conventional diagnostics, including PCR and qPCR, while sensitive and specific, require expensive laboratory infrastructure and well-trained personnel, limiting their deployment in field settings where immediate intervention is most valuable. To address these limitations, this protocol describes a portable molecular diagnostic workflow based on loop-mediated isothermal amplification (LAMP) combined with gold nanoparticle–DNA nanoprobes for specific and visual detection of ILTV directly at the point of need. Gold nanoparticles synthesized via the Turkevich method are functionalized with thiolated DNA probes, which undergo full-length, sequence-specific hybridization to LAMP amplicons, enabling a naked-eye colorimetric readout. The procedure integrates streamlined steps for DNA probe preparation, nanoparticle synthesis and assembly, and minimal sample processing, compatible with on-farm deployment. Results obtained with this workflow on field samples demonstrated 100% sensitivity and specificity, matching the performance of gold-standard assays. This approach offers a rapid, cost-effective, and equipment-free detection system of viral pathogens, enabling timely decision-making for disease containment and biosecurity. By overcoming the barriers of conventional diagnostics, this protocol enables producers with powerful tools for efficient monitoring and response to respiratory outbreaks in poultry farms.

The CRISPR/Cas9 system is a cornerstone technology in genome editing. Delivery of pre-assembled Cas9 ribonucleoprotein (RNP) complexes exhibits distinct advantages, including reduced off-target effects and lower immunogenicity. Conventional methods for purifying Cas9 protein typically involve multi-step chromatography and the cleavage of fusion tag, which are time-consuming and result in diminished yields. In this study, we present a simplified, one-step purification strategy for functional Streptococcus pyogenes Cas9 (SpCas9) using the ubiquitin (Ub) fusion system in Escherichia coli. The N-terminal Ub fusion not only improves protein solubility but also facilitates high-yield production of the His-Ub-Cas9 fusion protein. Importantly, the Ub tag does not require proteolytic removal during purification, allowing direct one-step purification of the fusion protein via nickel-affinity chromatography. The purified His-Ub-Cas9 retains robust DNA cleavage activity in vivo, as validated in zebrafish embryos. This protocol greatly simplifies the production of functional Cas9 protein, facilitating its broad application in genome editing.

Our genome is duplicated during every round of cell division through the process of DNA replication, but this fundamental process is subjected to various stresses arising from endogenous or exogenous sources. Thus, studying replication dynamics is crucial for understanding the mechanisms underlying genome duplication in physiological and replication stress conditions. Earlier, radioisotope-based autoradiography and density-labeling methods were used to study replication dynamics, which were limited in spatial resolution, representing only average estimates from many DNA samples. Here, we describe a DNA fiber assay that utilizes different thymidine analog incorporation, like 5-chloro-2’-deoxyuridine (CldU) and 5-iodo-2’-deoxyuridine (IdU), into replicating DNA. Such labeled DNA can be stretched and fixed on silanized glass slides, which are denatured with mild acidic treatment to expose the labeled nascent DNA. This DNA can then be visualized by using primary antibodies against CldU and IdU, followed by fluorophore-conjugated secondary antibodies, and observing them using a fluorescence microscope. The DNA fiber assay allows the visualization of individually replicating DNA at a single-molecular resolution and is highly quantitative, high-throughput, and easily reproducible. This technique offers insights into different replication parameters, like rate of DNA synthesis, extent of reversed fork protection, restart of stalled forks, and fork asymmetry under untreated or replication stress conditions at a single-molecule level.

Tail vein catheterization in mice is a standard technique for precise drug delivery in pharmacological research, offering high accuracy and reproducibility. However, existing techniques face significant limitations in maintaining long-term stable catheter patency in awake, freely moving mice, and there is currently no standardized, detailed protocol for tail vein catheterization. Current methods suffer from high rates of catheter dislodgement, increased animal stress from repeated injections, and movement restrictions, all of which introduce confounding variables in behavioral and pharmacological studies. We have developed a simple and efficient fixation method that maintains stable tail vein catheter patency for more than 60 min while allowing complete freedom of movement. This protocol employs a strain relief loop design and multi-point fixation strategy, effectively preventing catheter dislodgement during extended periods while minimizing animal stress. This protocol has been successfully applied across multiple research areas, including metabolic studies, behavioral assessments, and neuropharmacological research in awake mice, achieving >95% catheter retention with normal animal behavior, providing a reliable technical platform for long-term awake-state research applications.

Plasma membrane–associated condensates driven by liquid–liquid phase separation represent a novel mechanism of receptor-mediated signaling transduction, serving as mesoscale platforms that concentrate signaling molecules and modulate reaction kinetics. Condensate formation is a highly dynamic process that occurs within seconds to minutes following receptor activation. Here, we present methods for de novo reconstituting liquid-like condensates on supported lipid bilayers and assessing the condensate fluidity using fluorescence recovery after photobleaching (FRAP). This protocol encompasses supported lipid bilayer preparation, condensation imaging, and FRAP analysis using total internal reflection fluorescence (TIRF) microscopy. Supported lipid bilayers provide a membrane-mimicking environment for receptor signaling cascades, offering mechanistic insights into protein–protein and lipid–protein interactions amid micron-scale condensates. The protocol can also be adapted to study condensates associated with the internal membranes of the Golgi apparatus, mitochondria, and other organelles.

Repetitive increases of intracellular calcium ions (Ca²⁺ oscillations) control cellular functions in various biological events, including meiotic resumption after fertilization. Sperm-derived substances enter the cytoplasm of mature oocytes by sperm fusion, causing Ca²⁺ oscillations. Sperm-independent Ca²⁺ oscillations are also induced in immature oocytes isolated from the ovaries of neonatal to adult mice. The presence of Ca²⁺ oscillations may contribute to subsequent oocyte quality; however, its physiological role and molecular mechanism are unclear. Here, we describe a method of collecting immature oocytes from the ovaries of juvenile (12, 15, and 21 days after birth) and adult mice and monitoring their Ca²⁺ oscillations. Since mouse oocytes are larger than other types of cells, they are a useful model for studying spatiotemporal patterns and the mechanism of Ca²⁺ oscillations in various types of cells. This method can be applied to other rodents due to similarities in oocyte size and developmental processes. Furthermore, the use of various fluorescent probes enables visualization of organelle rearrangement. The mechanism of interaction between oocytes and somatic cells differs between juvenile and adult mice. Therefore, two distinct methods are employed for oocyte collection.

The compound eyes of Drosophila are widely used to gain valuable insights into genetics, developmental biology, cell biology, disease biology, and gene regulation. Various parameters, such as eye size, pigmentation loss, formation of necrotic patches, and disorientation, fusion, or disruption of ommatidial arrays, are commonly assessed to evaluate eye development and degeneration. We developed an improved imaging method named low-angle ring illumination stereomicroscopy (LARIS) to capture high-contrast images of the Drosophila compound eye. Different optical alignments were tested to capture the fly compound eye image under the stereomicroscope; the highest contrast with minimal reflection was achieved through the LARIS method. The images captured using LARIS clearly showed ommatidial fusion, disorientation, and pigmentation loss, which were hardly visible with a conventional imaging method in the degenerating compound eyes of Drosophila. In addition to its research applications, this protocol is cost-effective due to the low expenses associated with supplies and equipment. We anticipate that LARIS will facilitate high-contrast imaging of the compound eyes in Drosophila and other insects.

Biomolecular condensates organize cellular processes through liquid–liquid phase separation, creating membrane-less compartments enriched in specific proteins and RNAs. Understanding their RNA composition is essential for elucidating plant stress responses, yet capturing these transiently associated RNAs remains technically challenging. We present Turbo-RIP (TurboID-based proximity labeling with RNA immunopurification), a comprehensive protocol for identifying condensate-associated RNAs in plants. Turbo-RIP employs the biotin ligase TurboID to label proximal proteins at 22 °C, followed by formaldehyde crosslinking and streptavidin-based capture of protein–RNA complexes. We provide detailed procedures for three cloning strategies, transformation of Nicotiana benthamiana and Arabidopsis thaliana, validation of TurboID activity, and RNA recovery. The protocol successfully captured processing body–associated RNAs with minimal background. Turbo-RIP enables systematic mapping of RNA populations within plant condensates under diverse conditions. The protocol requires 3–5 days from sample preparation to RNA isolation, with construct validation taking 2–4 weeks. All procedures use standard laboratory equipment, making Turbo-RIP accessible for plant molecular biology laboratories.

The extracellular matrix (ECM) critically shapes melanoma progression and therapeutic response, yet commonly used matrices such as Matrigel fail to capture tissue- and disease-specific ECM properties. This protocol provides a streamlined and scalable method for generating murine, tissue-specific ECM hydrogels from skin, lung, and melanoma tumors, therefore overcoming the restricted materials of mouse-derived ECM. The workflow integrates tissue-tailored decellularization, lyophilization, mechanical fragmentation, pepsin digestion, and physiological polymerization to produce hydrogels that reliably preserve fibrillar collagen architecture and organ-specific ECM cues. Decellularization efficiency and ECM integrity are validated by DNA quantification, H&E staining, and Picrosirius Red staining analysis. These hydrogels provide a species- and tissue-matched platform for studying melanoma–ECM–immune interactions, pre-metastatic niche features, and therapy-induced ECM remodeling. Overall, this protocol offers a reproducible and physiologically relevant ECM model that expands experimental capabilities for melanoma biology and treatment-resistance research and that can be easily extended to other tumors and tissues.

Neuronal survival in vitro is usually used as a parameter to assess the effect of drug treatments or genetic manipulation in a disease condition. Easy and inexpensive protocols based on neuronal metabolism, such as 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT), provide a global view of protective or toxic effects but do not allow for the monitoring of cell survival at the single neuronal level over time. By utilizing live imaging microscopy with a high-throughput microscope, we monitored transduced primary cortical neurons from 7–21 days in vitro (DIV) at the single neuronal level. We established a semi-automated analysis pipeline that incorporates data stratification to minimize the misleading impact of neuronal trophic effects due to plating variability; here, we provide all the necessary commands to reproduce it.

Understanding gene expression within defined neuronal populations is essential for dissecting the cellular and molecular diversity of the brain. mRNA assays provide a direct readout of gene expression, capturing transcriptional changes that may precede or occur independently of protein abundance, whereas protein assays reflect the cumulative effects of translation, modification, and degradation. Moreover, in histological analysis, immunohistochemical protein detection results in visually diffuse labeling, which makes it difficult to quantitatively assess levels and locations of expression at high resolution. Here, we present a protocol that allows for mRNA detection in single neuronal cell types with a high degree of sensitivity and anatomical resolution. This protocol combines fluorescent in situ hybridization (FISH) with immunohistochemistry (IHC) on the same tissue section. Briefly, FISH is carried out by ACDBio RNAscope^® fluorescent in situ hybridization technology, which involves processing the tissue sections, followed by signal amplification. This involves target retrieval, probe hybridization, and signal enhancement. Then, the tissue section is processed for IHC, which involves blocking nonspecific sites and incubation with primary antibodies, followed by development of a fluorescent signal with secondary antibodies. Typically, visual mRNA detection with FISH can be seen as individual puncta, whereas targeting the protein with an antibody results in filled cells or processes. The variation in staining pattern allows for the quantification of distinct mRNA transcripts within different neuronal populations, which renders co-localization analyses easy and efficient.

Evaluating RNA–protein interactions is key to understanding post-transcriptional gene regulation. Electrophoretic mobility shift assays (EMSAs) remain a widely used technique to study these interactions, revealing information about binding affinities and binding modalities, including cooperativity and complex formation. Here, we detail, in a step-by-step protocol, how to perform EMSAs. We describe how to generate, purify, and quantitate ³²P-radiolabeled RNA by in vitro transcription, as well as the expression and purification of recombinant RNA-binding proteins in E. coli using ELAV as an example. We then describe how to set up binding reactions using serial dilutions in a microtiter plate format of recombinant ELAV and in vitro–transcribed RNA and how to perform EMSAs using native low-crosslinked acrylamide gels, with detailed graphically supported instructions and troubleshooting guides.

This protocol describes a reproducible workflow for modeling in vitro impact-induced traumatic brain injury (TBI) using a mechanical stretch system applied to differentiated SH-SY5Y human neuroblastoma cells cultured on polydimethylsiloxane (PDMS) substrates. The protocol integrates three primary components: (1) fabrication and surface modification of deformable PDMS chambers to support cellular adhesion, (2) partial differentiation of SH-SY5Y cells using retinoic acid, and (3) induction of controlled mechanical strain to simulate mild to moderate TBI. The stretch-induced injury model enables quantitative assessment of cellular viability and recovery following mechanical insult. This approach provides a versatile platform for studying cellular and molecular mechanisms of TBI, screening neuroprotective compounds, and exploring mechanobiological responses in neural cells under controlled strain magnitudes and rates.

Nowadays, the use of 3D cultures (organoids) is considered a valuable experimental tool to model physiological and pathological conditions of organs and tissues. Organoids, retaining cellular heterogeneity with the presence of stem, progenitor, and differentiated cells, allow the faithful in vitro reproduction of structures resembling the original tissue. In this context, the growth of endometrial organoids allows the generation of 3D cultures characterized by a hollow lumen, secretory activity, and apicobasal polarity and displaying phenotypical modification in response to hormone stimulation. However, a limitation in currently used models is the absence of stromal cells in their structure; as a result, they miss epithelial–stromal interactions, which are crucial in endometrial physiology. We developed a novel 3D model to generate endometrial organoids grown in floating Matrigel^TM droplets in the presence of standard culture medium. From a structural point of view, these novel floating 3D cultures develop as gland-like structures constituted by epithelial cells organized around a central lumen and retain the expression of endometrial and decidual genes, like previously published organoids, although with a phenotype resembling hormonally differentiated structures. Importantly, floating organoids retain stromal cells which grow in close contact with the epithelial cells, localized within the internal or external portion of the organoid structure. In summary, we present a simple and rapid model for generating 3D endometrial organoids that preserve epithelial–stromal cell interactions, promoting the formation of differentiated organoids and enabling the study of reciprocal modulation between epithelium and stroma.

The plant cell wall is a dynamic and complex extracellular matrix that not only provides structural integrity and determines cell shape but also mediates intercellular communication. Among its major components, pectins play essential roles in cell adhesion, wall porosity, hydration, and flexibility. Rhamnogalacturonan-I (RG-I), a structurally diverse pectic polysaccharide, remains one of the least understood components of the plant cell wall. Its backbone is substituted with arabinan, galactan, and arabinogalactan side chains that vary in length, branching, and composition across tissues, species, and developmental stages. In addition, RG-I can undergo modifications such as backbone acetylation, further contributing to its structural complexity and functional diversity. To advance understanding of RG-I, we present a detailed method for isolating RG-I from the model plant Arabidopsis thaliana. Leveraging Arabidopsis as a model system provides major advantages owing to its well-characterized genome and powerful molecular toolkit, enabling deeper investigation into the roles of RG-I in plant development and responses to environmental stress. Our method consists of two major steps: an initial chemical extraction using oxalate, followed by endo-polygalacturonase (EPG) digestion to fragment the pectic domains. An advantage of this approach is that it produces a dry material that can be stored at room temperature without special handling and does not introduce chemicals that may interfere with downstream analyses. The purified RG-I can be used for detailed compositional and structural analyses, as well as for functional studies of enzymes involved in pectin biosynthesis, modification, and degradation. Although this protocol was developed for isolating RG-I from Arabidopsis rosette leaves, it is also applicable to other Arabidopsis organs and other plant species.

Pinpointing causal genes for complex traits from genome-wide association studies (GWAS) remains a central challenge in crop genetics, particularly in species with extensive linkage disequilibrium (LD) such as rice. Here, we present CisTrans-ECAS, a computational protocol that overcomes this limitation by integrating population genomics and transcriptomics. The method’s core principle is the decomposition of gene expression into two distinct components: a cis-expression component (cis-EC), regulated by local genetic variants, and a trans-expression component (trans-EC), influenced by distal genetic factors. By testing the association of both components with a phenotype, CisTrans-ECAS establishes a dual-evidence framework that substantially improves the reliability of causal inference. This protocol details the complete workflow, demonstrating its power not only to identify causal genes at loci with weak GWAS signals but also to systematically reconstruct gene regulatory networks. It provides a robust and powerful tool for advancing crop functional genomics and molecular breeding.

The genomes of RNA viruses can fold into dynamic structures that regulate their own infection and immune evasion processes. Proximity ligation methods (e.g., SPLASH) enable genome-wide interaction mapping but lack specificity when dealing with low-abundance targets in complex samples. Here, we describe HiCapR, a protocol integrating in vivo psoralen crosslinking, RNA fragmentation, proximity ligation, and hybridization capture to specifically enrich viral RNA–RNA interactions. Captured libraries are sequenced, and chimeric reads are analyzed via a customized computational pipeline to generate constrained secondary structures. HiCapR generates high-resolution RNA interaction maps for viral genomes. We applied it to resolve the in vivo structure of the complete HIV-1 RNA genome, identifying functional domains, homodimers, and long-range interactions. The protocol's robustness has been previously validated on the SARS-CoV-2 genome. HiCapR combines proximity ligation with targeted enrichment, providing an efficient and specific tool for studying RNA architecture in viruses, with broad applications in virology and antiviral development.

Functional enrichment analysis is essential for understanding the biological significance of differentially expressed genes. Commonly used tools such as g:Profiler, DAVID, and GOrilla are effective when applied to well-annotated model organisms. However, for non-model organisms, particularly for bacteria and other microorganisms, curated functional annotations are often scarce. In such cases, researchers often rely on homology-based approaches, using tools like BLAST to transfer annotations from closely related species. Although this strategy can yield some insights, it often introduces annotation errors and overlooks unique species-specific functions. To address this limitation, we present a user-friendly and adaptable method for creating custom annotation R packages using genomic data retrieved from NCBI. These packages can be directly imported as libraries into the R environment and are compatible with the clusterProfiler package, enabling effective gene ontology and pathway enrichment analysis. We demonstrate this approach by constructing an R annotation package for Mycobacterium tuberculosis H37Rv, as an example. The annotation package is then utilized to analyze differentially expressed genes from a subset of RNA-seq dataset (GSE292409), which investigates the transcriptional response of M. tuberculosis H37Rv to rifampicin treatment. The chosen dataset includes six samples, with three serving as untreated controls and three exposed to rifampicin for 1 h. Further, enrichment analysis was performed on genes to demonstrate changes in response to the treatment. This workflow provides a reliable and scalable solution for functional enrichment analysis in organisms with limited annotation resources. It also enhances the accuracy and biological relevance of gene expression interpretation in microbial genomics research.

RNA is now recognized as a highly diverse and dynamic class of molecules whose localization, processing, and turnover are central to cell function and disease. Live-cell RNA imaging is therefore essential for linking RNA behavior to mechanism. Existing approaches include quenched hybridization probes that directly target endogenous transcripts but face delivery and sequestration issues, protein-recruitment tags such as MS2/PP7 that add large payloads and can perturb localization or decay, and CRISPR–dCas13 imaging that requires substantial protein cargo and careful control of background and off-target effects. Here, we present a protocol for live-cell RNA imaging using the Spinach^TM family of fluorogenic RNA aptamers. The method details the design and cloning of Spinach^TM-tagged RNA constructs, selection and handling of cognate small-molecule fluorophores, expression in mammalian cell lines, dye loading, and image acquisition on standard fluorescence microscopes, followed by quantitative analysis of localization and dynamics. We include controls to verify aptamer expression and signal specificity, guidance for multiplexing with related variants (e.g., Broccoli, Corn, Squash, Beetroot), and troubleshooting for dye permeability and signal optimization. Application examples illustrate use in tracking cellular delivery of mRNA therapeutics, monitoring transcription and decay in response to perturbations, and the forming of toxic RNA aggregates. Compared with prior methods, Spinach^TM tags are compact, genetically encodable, and fluorogenic, providing high-contrast imaging in both the nucleus and cytoplasm with single-vector simplicity and multiplexing capability. The protocol standardizes key steps to improve robustness and reproducibility across cell types and laboratories.

RNA imaging techniques enable researchers to monitor RNA localization, dynamics, and regulation in live or fixed cells. While the MS2-MCP system—comprising the MS2 RNA hairpin and its binding partner, the MS2 coat protein (MCP)—remains the most widely used approach, it relies on a tag containing multiple fluorescent proteins and has several limitations, including the potential to perturb RNA function due to the tag’s large mass. Alternative methods using small-molecule binding aptamers have been developed to address these challenges. This protocol describes the synthesis and characterization of RNA-targeting probes incorporating a peptide nucleic acid (PNA)-based linker within the cobalamin (Cbl)-based probe of the Riboglow platform. Characterization in vitro involves a fluorescence turn-on assay to determine binding affinity (K_D) and selective 2′-hydroxyl acylation analyzed by primer extension (SHAPE) footprinting analysis to assess RNA-probe interactions at a single nucleotide resolution. To show the advancement of PNA probes in live cells, we present a detailed approach to perform both stress granule (SG) and U-body assays. By combining sequence-specific hybridization with structure-based recognition, our approach enhances probe affinity and specificity while minimizing disruption to native RNA behavior, offering a robust alternative to protein-based RNA imaging systems.

The CRISPR/Cas12a system has revolutionized molecular diagnostics; however, conventional Cas12a-based methods for RNA detection typically require transcription and pre-amplification steps. Our group has recently developed a diagnostic technique known as the SCas12a assay, which combines Cas12a with a split crRNA, achieving amplification-free detection of miRNA. However, this method still encounters challenges in accurately quantifying long RNA molecules with complex secondary structures. Here, we report an enhanced version termed SCas12aV2 (split-crRNA Cas12a version 2 system), which enables direct detection of RNA molecules without sequence limitation while demonstrating high specificity in single-nucleotide polymorphism (SNP) applications. We describe the general procedure for preparing the SCas12a system and its application in detecting RNA targets from clinical samples.