Manganese (Mn) is an essential trace element whose intracellular homeostasis is tightly controlled by specialized membrane transporters. Dysregulation of Mn transport leads to pathological Mn accumulation and severe human disease; however, efficient and quantitative cell-based methods for assessing Mn²⁺ transporter activity remain limited. Here, we present an optimized cellular Fura-2 manganese extraction assay (CFMEA) that enables robust quantification of cellular Mn content and provides a normalized framework for assessing relative Mn²⁺ transport activity in a high-throughput format. This protocol integrates Fura-2-based fluorescence detection of Mn²⁺ at the Ca²⁺ isosbestic excitation wavelength with dsDNA quantification to normalize dsDNA levels in cell extracts and immunoblotting to account for transporter protein expression levels. Cells expressing Mn²⁺ transporters are exposed to MnCl₂ in 96-well plates, washed to remove extracellular Mn²⁺, and lysed in a Fura-2-containing extraction buffer. Fluorescence quenched by Mn²⁺ is quantified and converted to cellular Mn content using a cell-free Mn-Fura-2 standard curve and then normalized to dsDNA content and protein abundance to determine relative transporter activity. This workflow provides a relatively sensitive, reproducible, and low-cost approach for comparative analysis of Mn²⁺ transporters and their variants across multiple cell types. The protocol is demonstrated using the Mn²⁺ efflux transporter SLC30A10 in HEK293T cells and is readily adaptable for studying other Mn²⁺ transport pathways.

Divalent metal ion transporters are conserved across all domains of life and play essential roles in diverse processes such as manganese acquisition during nutritional immunity in bacteria and iron homeostasis in higher eukaryotes [1–3]. Traditional techniques, such as electrophysiological assays, are often unsuitable due to the slow kinetics of many membrane transporters, electroneutral nature of certain transporter types, and the influence of other proteins with similar activity. To overcome these limitations and to investigate both the activity and ion selectivity of transporters, also including those normally expressed intracellularly, we have developed a fluorescence-based transport assay using purified proteins. This in vitro assay uses encapsulated fluorophores to monitor the movement of divalent metal ions (e.g., Mn²⁺, Ca²⁺, Mg²⁺) or protons across liposomal membranes reconstituted with purified transporter proteins. This approach provides detailed functional insight that complements structural and cellular data.

Stochastic optical reconstruction microscopy (STORM) is a single-molecule localization microscopy technique that enables visualization of cellular structures beyond the diffraction limit. This approach has revealed previously inaccessible ultrastructural details in a wide range of cellular components, including the actin cytoskeleton, clathrin-coated pits, mitochondria, and bacterial nucleoid-associated proteins. STORM relies on the sequential emission of single photons from photosensitive fluorophores, which are precisely localized before entering a dark state or undergoing photobleaching. By activating fluorophores individually and fitting their point spread functions (PSFs), the center of mass can be calculated with a localization precision of up to ~20 nm. The parallel detection of thousands of single-molecule events, each assigned to distinct spatial coordinates, enables the reconstruction of a high-resolution image. Here, we describe a simple and efficient STORM workflow—including sample preparation, image acquisition, and quality control measurements—that we used to visualize various subcellular structures, such as mitochondria, microtubules, and lysosomes labeled with the commonly employed cyanine dye Alexa Fluor 647, as well as the actin cytoskeleton stained with Alexa Fluor 488–conjugated phalloidin. Image acquisition was performed using a conventional epifluorescence/total internal reflection (TIRF) microscope adapted for STORM imaging. Key adaptations included the use of a 160×/1.43 NA oil-immersion objective and a high-power mode, which concentrates the laser beam onto a small region of the sample, ensuring sufficient light intensity to drive fluorophores into the dark state. In addition, implementing a 1.6× magnification lens and a 4×4 binning camera mode allowed us to achieve a 100-nm pixel size optimal for reliable molecule detection. We believe that this protocol will be highly valuable to the microscopy community, as it lowers technical barriers to performing STORM on widely available microscopy platforms, thereby facilitating broader implementation of this powerful super-resolution technique.

The placenta is a metabolically active organ whose mitochondrial activity is tightly linked to fetal growth, oxygenation, and nutrient transport, mediating fetal susceptibility to environmental exposures. Accordingly, aberrant mitochondrial function has been implicated in the progression of placental dysfunction. However, existing respirometry platforms require primarily fresh or cryopreserved placental tissue and offer limited throughput, rendering these platforms impractical in the context of large-scale placental dissections. Here, we describe and validate a Seahorse XF approach for measuring mitochondrial respiration in previously frozen placentae, enabling the functional interrogation of placental mitochondria in prenatal studies. Our protocol fundamentally relies on the restoration of matrix substrates that are depleted due to increased mitochondrial membrane permeability following freeze-thaw cycles. We provide a strategy to assess complex I and II-associated respiration adapted for the Seahorse XFe24 Analyzer and further demonstrate comparable oxygen consumption readouts between fresh and frozen placentae. We further demonstrate distinct differences in the magnitude of oxygen consumption between fresh and frozen placentae in the absence of exogenous NADH. Taken together, we present a simplified and convenient protocol for the assessment of respiratory enzyme complex-associated respiration from archived placental tissue.

3-nitro-tyrosine (nitroTyr) is one of numerous oxidative protein modifications implicated in diseases such as cardiovascular disease, cancer, and amyotrophic lateral sclerosis (ALS). Because of this, the ability to site-specifically encode nitroTyr into recombinant proteins is a powerful approach for studying these disease pathways. However, producing proteins with defined nitration sites is technically challenging due to the limitations of traditional chemical nitration via peroxynitrite, which lacks residue and site-specificity. Genetic code expansion (GCE) offers a solution by enabling precise incorporation of nitroTyr at designated TAG codons using engineered aminoacyl-tRNA synthetase/tRNA pairs from Methanocaldococcus jannaschii and Methanomethylophilus alvus. This protocol provides a reliable, optimized workflow for incorporating nitroTyr into proteins in E. coli using GCE. It guides users through key considerations in selecting cell lines, media conditions, and GCE systems to minimize off-target effects such as release factor 1 competition, near-cognate suppression, and chemical reduction of nitroTyr. The method is demonstrated using wild-type and TAG-containing superfolder GFP but is broadly applicable to other proteins of interest.

DNA language models, such as DNABERT-2, have recently enabled the accurate prediction of functional sequence elements across species. However, the practical, protocol-style steps needed to transform these resources into training datasets, fine-tune the official DNABERT-2 model, and evaluate classifier performance have not been explicitly described. Herein, we present a step-by-step computational protocol for preparing training data, fine-tuning DNABERT-2, and evaluating sequence-level binary classifiers using readily available command-line tools. The protocol has been demonstrated using RNA off-target sites induced by cytosine base editors, detected by our PiCTURE pipeline from RNA sequencing (RNA-seq) data, and extended to core promoter prediction using the EPDnew database. We describe how to derive positive and negative sequence sets into DNABERT-2 compatible datasets, and fine-tune the official pretrained model of DNABERT-2 using the datasets. We also demonstrate how to compute the standard performance metrics and compare the model outputs with the baselines. This protocol will help researchers adapt DNA foundation models to new genomic tasks, including the safety assessment of genome editing tools and the functional annotation of regulatory sequences.

Membrane transporters mediate the selective movement of ions and molecules across biological membranes and are essential for cellular homeostasis. However, their functional characterization in living cells is often complicated by the complexity of the native membrane environment. Reconstitution into model membrane systems provides a powerful alternative by enabling precise control over lipid composition and experimental conditions. Giant unilamellar vesicles (GUVs) are particularly well suited for transporter studies, as their cell-sized dimensions allow direct microscopic observation and fluorescence-based measurements of protein activity. Here, we describe a two-step reconstitution protocol in which transport proteins are first incorporated into large unilamellar vesicles and then used to generate protein-containing giant unilamellar vesicles (proteo-GUVs) via the poly(vinyl alcohol) swelling method. This two-step approach enhances protein incorporation efficiency and preserves transporter functionality. The method is exemplified using the P3-type ATPase Arabidopsis thaliana plasma membrane H⁺-ATPase isoform 2 (AHA2). We further describe a fluorescence-based assay to assess proton transport activity in proteo-GUVs. Our approach provides a versatile and controlled platform for biochemical, biophysical, and single-molecule analysis of membrane transporters.

Engineering of microbial cells, including E. coli, is essential in prototyping genetic designs used in numerous applications throughout synthetic biology. While many advanced genome editing tools, such as CRISPR-based tools, offer new capabilities with genetically recalcitrant organisms, these tools often do not offer an immediate advantage in readily manipulated microbes, such as E. coli, especially when scarless modifications are not critical. We describe a comprehensive recombineering tutorial that we commonly use for multiplex engineering of E. coli using antibiotic markers. We leverage a group of 15 antibiotic resistance cassettes, most of which can be readily included when designing double-stranded DNA donors intended for recombineering and purchased from several vendors. Using these methods, 10–15 defined modifications to a single host strain can be achieved in less than three weeks, using two-day editing cycles. We discuss sequences and protocols as well as the optimal design of genetic modifications and the associated DNA.

The integrity of epithelial barriers is essential for maintaining tissue homeostasis, particularly in the intestinal tract, where it separates the host from the complex luminal environment. Two complementary, standard methods for assessing this barrier are transepithelial electrical resistance (TEER), which provides a rapid, non-destructive measure of ionic conductance across tight junctions, and the fluorescein isothiocyanate (FITC)-dextran assay, which directly quantifies paracellular macromolecule flux. This protocol details a robust and reproducible method for performing both assays using intestinal epithelial cell monolayers (e.g., Caco-2, T84) cultured on permeable Transwell supports. We outline the procedure from cell culture and monolayer differentiation to TEER measurement with an Epithelial Volt/Ohm Meter 3 (EVOM3) and the subsequent FITC-dextran permeability assay. By combining these techniques, this protocol provides a comprehensive assessment of barrier function, making it ideal for studying tight junction dynamics and regulation under various experimental conditions, such as cytokine stimulation, drug screening, or microbial challenges.

Epigenetic modifications play essential roles in regulating gene expression and maintaining cellular identity. Accumulating evidence suggests that chemical agents can contribute to carcinogenesis through epigenetic alterations, such as changes in DNA methylation and histone modifications, even in the absence of direct DNA damage. Here, we have developed a simple, cost-effective, and quantitative reporter assay, termed the epi-TK assay, to evaluate chemically induced epigenetic alterations. The assay is built upon the thymidine kinase (TK) gene mutation assay, a standardized and widely used in vitro genotoxicity assay for chemical safety evaluation. This system is based on an engineered human lymphoblastoid cell line (mTK6), in which the promoter region of the endogenous housekeeping TK gene is site-specifically methylated using epigenome-editing technology, resulting in stable transcriptional repression. Following chemical exposure, epigenetic perturbations at the TK locus are detected by culturing cells under hypoxanthine–aminopterin–thymidine selection and quantifying the frequency of TK revertant colonies, which reflects restoration of TK gene expression. Using the DNA methyltransferase 1 inhibitor GSK3484862 as a model compound, this protocol demonstrates that the epi-TK assay enables sensitive and quantitative detection of epigenetic state transitions. Importantly, this assay allows bi-directional detection of epigenetic changes, including DNA demethylation events and broader alterations in histone modification landscapes. Together, the epi-TK assay provides a practical and quantitative platform for evaluating epigenetic toxicity, with potential applications in chemical safety assessment frameworks.

Structural proteomics methods allow for the proteome-wide interrogation of protein structural differences between two different conditions. Limited proteolysis mass spectrometry (LiP-MS), as originally implemented by the Picotti lab, utilizes a promiscuous protease to cleave at solvent-exposed regions of a protein to encode structural information, which is then read out with mass spectrometry proteomics. Here, we present a protocol that details experimental steps and data analysis for a LiP-MS workflow. First, tissue is homogenized under native conditions and then subjected to limited proteolysis using proteinase K (PK). The samples are prepared for mass spectrometry, and data are acquired using either data-dependent acquisition (DDA) or data-independent acquisition (DIA). Raw data is processed using FragPipe, and raw ion abundances are processed in FragPipe Limited-Proteolysis Processor (FLiPPR). Proteins with structural changes between the two conditions are identified in a proteome-wide manner.

Anti-CD19 chimeric antigen receptor (CAR)-natural killer (NK) cells are expected to demonstrate anti-CD19 CAR-T-cell-like efficacy against relapsed and refractory B-cell malignancies and autoimmune diseases, with fewer adverse events and the added advantage of permitting the use of allogeneic cells. However, the methodology for generating CAR-NK cells remains under development. Although various cell sources and expansion methods are available, feeder cells derived from cancerous tissue have been most commonly employed to promote ex vivo expansion of NK cells. In the protocol described herein, NK cells are expanded from adult peripheral blood mononuclear cells using CD2- and NKp46-specific stimulating antibodies in combination with multiple cytokines. The activated NK cells can be genetically modified using a retroviral vector. Subsequent culture of these cells yields large numbers of anti-CD19 CAR-NK cells. The current method, which enables feeder-free, large-scale generation of anti-CD19 CAR-NK cells, eliminates the risk of tumor cell contamination and may facilitate safer clinical application.

Natural killer (NK) cells are crucial innate immune effectors, mediating cytotoxicity against cancer and infected cells through receptors such as NKG2D. Reliable quantification of NK cell subsets is essential for evaluating NK cell-based immune responses in cancer research. Unlike other assays, including traditional flow cytometry used in assessing NK cells, imaging flow cytometry (IFC) is a simple and direct method for quantitative analysis of NK cells. This protocol describes the necessary procedures, including harvesting splenocytes, acquiring these cells labeled with NKG2D antibodies, and analyzing IFC data with IDEAS^® software. We applied this protocol to quantitatively assess the number of splenic NKG2D⁺ NK cells in mice injected with SVTneg2 cancer cells (which carry the p53 G242A missense mutation) and compared them to mice injected with EMT6 cancer cells (which have wild-type p53) or normal fibroblasts. We found that the SVTneg2 cancer cells significantly decreased the number of NKG2D⁺ NK cells in mice by approximately 2-fold (933 cells vs. 2360 cells, p < 0.001) compared with mice injected with EMT6 cancer cells. This IFC protocol can be applied to directly quantify NK cells in vivo. This quantitative protocol allows novices to quickly handle the analysis of cytotoxic NK cells with a single NKG2D marker. Further multicolor flow cytometry and cytokine assay may be required to precisely define the subtypes and effects of NK cells in anticancer immunity.

Plant genome editing is a powerful approach for modifying plant DNA to investigate gene function and to engineer desirable traits. Several genome-editing technologies have been developed, among which CRISPR/Cas systems and transcription activator-like effector nucleases (TALENs) are widely used to introduce targeted double-stranded DNA breaks. While CRISPR/Cas systems are highly efficient for nuclear genome editing, their application to plant organellar genomes remains limited, largely due to difficulties in guide RNA delivery into mitochondria and chloroplasts. Here, we present a detailed and reproducible protocol for constructing TALEN-based binary vectors for targeted genome editing in Arabidopsis thaliana. This protocol describes the assembly of TALE repeat arrays, the generation of nuclear-, mitochondrial-, and plastid-targeted TALEN expression vectors using MultiSite Gateway cloning, and subsequent Agrobacterium-mediated plant transformation and genotyping. The workflow enables the production of nTALENs, mitoTALENs, and ptpTALENs using a unified vector design strategy. In addition, the protocol briefly outlines the construction principles of TALE-based cytidine deaminases (TALECDs) for targeted C-to-T base editing in plant organellar genomes. The protocol provides a flexible and robust framework for plant nuclear and organellar genome editing and can be readily adapted to different target genes and experimental purposes. Its modular design and compatibility with standard molecular cloning techniques make it accessible to laboratories aiming to perform precise genome manipulation in plants.

Aloe vera has long been used for its diverse pharmacological properties, motivating continued interest in isolating and preserving the bioactive molecules responsible for its therapeutic potential. More recently, Aloe vera–derived extracellular vesicles (Av-EVs) have emerged as nanoscale, cell-free carriers capable of retaining and delivering these properties, making them attractive for various biomaterials, nanomedicine, and regenerative medicine applications. Multiple techniques are available for extracellular vesicle isolation. These include ultracentrifugation, polymer-based precipitation, size-exclusion chromatography, immunoaffinity capture, ultrafiltration, density gradient separation, and emerging microfluidic platforms. Each method presents distinct trade-offs in purity, yield, scalability, and downstream compatibility. Despite this diversity, standardized workflows tailored to Av-EV isolation remain limited, and the influence of homogenization-induced shear forces and plant maturity on vesicle recovery and characterization has not been systematically addressed. Here, we present a reproducible protocol for isolating Av-EVs from Aloe vera gel employing two distinct homogenization strategies: manual, no-shear force (NB EVs), and blender-based shear-force homogenization (B EVs). The workflow covers gel preparation, serial centrifugation for debris removal, ultracentrifugation as the gold standard for vesicle enrichment, and final sterile filtration. This protocol enables consistent recovery of Av-EVs suitable for physicochemical characterization and functional analyses. It is simple and relies on commonly available laboratory equipment, facilitating broad adoption by ultracentrifugation users and offering adaptability to diverse research projects involving purified Aloe vera gel and Av-EVs, including studies focused on wound healing, fibrotic scarring, and regenerative processes, where coordinated antioxidant, anti-inflammatory, antimicrobial, immunomodulatory, and moisturizing responses are of interest.

Protein–protein interactions (PPIs) govern nearly all aspects of cellular physiology, yet identifying these interactions under native conditions remains challenging. Here, we present TIE-UP-SIN (targeted interactome experiment for unknown proteins by stable isotope normalization), a robust method for in vivo identification and quantification of PPIs in bacterial systems. The protocol combines metabolic labeling with ¹⁵N isotopes, reversible formaldehyde crosslinking, affinity purification, and quantitative mass spectrometry. TIE-UP-SIN preserves transient or weak interactions during purification and quantifies interaction partners using internal light/heavy peptide ratios, reducing experimental variability. The method employs a triple-sample design to distinguish specific from nonspecific interactors and can be adapted to various bacterial species and affinity tags. Data analysis is streamlined through a user-friendly web application (https://shiny-fungene.biologie.uni-greifswald.de/TIE_UP_SIN_app) that automates statistical analysis, normalization, and visualization, requiring no programming expertise. The entire workflow from cell culture to mass spectrometry data acquisition takes approximately 4–5 days, with data analysis completed in 1–2 days using the web application.

We established a step-by-step approach for generating a single-nucleotide mutation in the promoter region of an immune regulatory gene in human monocyte THP-1 cells by employing a plasmid-based CRISPR-Cas9 system delivered via transfection with a homology-directed repair template DNA (HDR). Key steps include designing a single-guide RNA (sgRNA), cloning it into a CRISPR plasmid encoding the Cas9 protein, transfection of the plasmid constructs along with single-stranded oligonucleotide repair template (ssODNs) into THP-1 cells, followed by selection and validation. This approach provides a precise and relevant model to investigate the role of single polymorphisms in the regulation of inflammatory gene expression in human monocytes. In addition to the rs1024611 single-nucleotide polymorphism (SNP), this CRISPR/Cas9-based strategy is broadly applicable to functional studies of noncoding and coding variants in innate immune genes.

Reactive oxygen species (ROS) are central regulators of plant development and stress responses, with hydrogen peroxide (H₂O₂) acting as a key signaling molecule whose spatial distribution determines adaptive versus damaging outcomes. Accurate detection of H₂O₂ at tissue and cellular resolution is therefore essential for understanding redox-dependent regulation of plant growth. A variety of techniques have been used to monitor H₂O₂, including bulk spectrophotometric and fluorometric assays, genetically encoded sensors for real-time measurements, and chemical probes for in situ detection. While these approaches differ in sensitivity, specificity, and temporal resolution, many are limited by a lack of spatial information, technical complexity, or dependence on transgenic material. Here, we present a detailed protocol for 3,3′-diaminobenzidine (DAB)-based histochemical detection of H₂O₂ in seedling roots, covering staining, imaging, and semi-quantitative image analysis using open-source software (FIJI/ImageJ). The method relies on peroxidase-mediated oxidation of DAB, resulting in a stable, light-resistant, and insoluble precipitate that enables visualization of H₂O₂ accumulation with high spatial resolution. This protocol provides a robust, accessible, and genetically independent approach for spatial analysis of H₂O₂ in plant tissues. Its simplicity, compatibility with diverse genotypes and treatments, and suitability for semi-quantitative analysis make it a valuable tool for examining the spatial distribution of H₂O₂, thereby providing spatial insight into redox-related regulatory processes during plant development and stress responses.

The rising global incidence of pancreatitis, pancreatic cancer, and diabetes has increased the need for efficient in vivo gene manipulation approaches to study the pancreas and develop new therapies. Although transgenic mouse models are widely used, they are time-consuming and costly to generate and maintain. Systemic viral delivery methods offer greater flexibility but often lack pancreatic specificity and require high viral doses. Here, we describe a streamlined protocol for intrapancreatic ductal delivery of adeno-associated viruses (AAVs) for targeted gene delivery. Our protocol requires standard surgical equipment and can be implemented in most laboratories. Specifically, we adopted a clamping strategy at the hepatopancreatic duct near the liver, as well as beneath the major duodenal papilla at the duodenum. This strategy exposes the duodenal papilla, facilitating viral delivery, preventing backflow, and enabling efficient pancreatic transduction at lower viral doses. Overall, this method provides a fast, simple, and effective approach for pancreas-targeted gene manipulation, facilitating preclinical studies of pancreatic biology and disease.

Metastasis is initiated by cell invasion of the basement membrane, facilitating cell migration and colonization at a secondary tumor site. Cancer cells remodel the cytoskeleton to form ventral protrusions, termed invadopodia, that traffic and deliver matrix metalloproteases to degrade the extracellular matrix. Traditional efforts have utilized immunolabeling to measure protein localization within invadopodia, an approach limited by reduced temporal resolution, logistical challenges in orienting invadopodia within the focal plane of the objective lens, and impaired ability to reconstitute physiological conditions. Here, we describe a protocol for constructing and utilizing the axial invasion chamber (AIC) to perform live-cell 3D visualization of mature elongating invadopodia under physiological conditions. The AIC is simple to build, using standard 35 mm glass-bottom dishes that suit most microscope stage holders. A polyester membrane is used to uniformly orient and promote invadopodia formation and restrict cell migration. The AIC extracellular matrix is composed of readily available reagents that have been optimized to facilitate cell adhesion and invadopodia maturation. Critical advances of the AIC include imaging and measurements of protein localization without immunolabeling, imaging of live cell invadopodia using conventional inverted microscopes, and production of a fully operational apparatus within 28 h from initial assembly. While the protocol has been used for live-cell invadopodia protein localization and structure, it provides an opportunity to interchange components of the polyester membrane and/or the extracellular matrix to optimize the device for a variety of different cell types and cell invasion studies.

The spatiotemporal dynamics and density of actin networks are key determinants of actin cytoskeleton–mediated cellular functions. In vitro reconstitution systems have been widely used to study actin cytoskeletal dynamics; however, many existing approaches offer limited flexibility in controlling the geometry, thickness, and density of the assembled actin networks. Here, we present an in vitro optogenetic protocol that enables precise control of actin network assembly on supported lipid bilayers using an improved light-induced dimer (iLID)-SspB-based light-inducible dimerization system. In this system, His-mEGFP-iLID is anchored to a Ni-NTA-containing lipid bilayer, while SspB-mScarlet-I-VCA, a nucleation-promoting factor fused with SspB, together with other actin cytoskeletal proteins, is supplied in bulk solution. Upon blue light illumination, SspB-mScarlet-I-VCA is recruited to the membrane in a spatially and temporally defined manner, inducing localized actin polymerization. By tuning illumination patterns and duration, actin networks with defined density, thickness, and geometry can be generated, and polymerization can be rapidly halted by stopping illumination. This protocol provides a versatile platform for reconstructing actin networks with controlled spatial organization and density, enabling quantitative analysis of density-dependent interactions between actin networks and actin-binding proteins.

Among the biophysical techniques used in fragment-based drug discovery (FBDD) campaigns, crystallography is the most sensitive, allowing for the identification of low-affinity ligands and the characterization of protein–ligand complexes at atomic resolution. Although powerful, the proper application of this technique depends on obtaining crystals capable of diffracting X-rays at high resolution. Additionally, in crystallographic compound screening, the crystals must be resistant to multiple organic solvents used in chemical libraries, such as DMSO. In this protocol, we describe recombinant protein production suitable for crystallization and procedures for X-ray crystallographic screening of a library of 768 fragments. As a case study, we used the Schistosoma mansoni thioredoxin glutathione reductase (SmTGR), a redox enzyme with a key role in controlling oxidative stress in parasites of the Schistosoma genus, which causes schistosomiasis. As a validated pharmacological target, SmTGR is used in the development of new schistosomicidal drugs. The experimental pipeline includes SmTGR expression, purification, and crystallization, crystal soaking, diffraction data collection, and refinement. The 768 fragments from the Diamond-SGC Poised Library (DSPL) were individually soaked onto the crystals, and diffraction data were collected and processed at the I04-1 beamline of the Diamond Light Source synchrotron. Diffraction data were subsequently analyzed using PanDDA to identify fragment-binding events and to enable reliable detection of low-occupancy ligands within the protein crystal structures. In addition to the core experimental steps, this protocol incorporates systematic approaches to overcome limitations frequently encountered in crystallographic screening campaigns, including assessment of crystal solvent tolerance, acceleration of crystal mounting through the use of auxiliary devices, acoustic dispensing–based soaking of hundreds of fragments for low material consumption and high throughput, automated data collection, and efficient data analysis pipeline for the detection of weakly bound ligand. This protocol can be broadly applied to screen diverse compound sets against multiple targets amenable to crystallization.

Fluorescence in situ hybridization (FISH) can be employed to study the expression and subcellular localization of nucleic acids by using labeled antisense strands that hybridize with the target RNA or DNA molecules. Likewise, immunofluorescence antibody staining (IF) takes advantage of the specific interaction between a fluorophore-labeled antibody and its corresponding antigen. This protocol reports the combination of RNA-FISH and IF antibody staining for simultaneous detection of both RNA transcripts and proteins of interest in routine formalin-fixed paraffin-embedded (FFPE) bone marrow biopsy samples. Herein, we provide a detailed description of the methodology that we have developed and optimized to study the spatial expression of two transcripts—TGFB1 and PDGFA1—in human hematopoietic (CD45⁺) and non-hematopoietic (CD271⁺) cells in the bone marrow of patients with acute lymphoblastic leukemia (ALL).

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.

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.