Intestinal glucose absorption has been studied for several decades. However, the different methods available for investigating absorption are often the reason for variability in the results, and it is difficult to measure the relative contribution of paracellular absorption using existing methods. Thus, we have established a new model for measuring glucose absorption. In the isolated in situ vascularly perfused small intestine, the intestinal epithelium is completely preserved, and the entire transport pathway is intact. In the present model, we use radioactive labeled ¹⁴C-d-glucose, which allows for sensitive quantification of glucose absorption even with low luminal concentrations. The described method is optimized for intestinal glucose absorption but can be applied to other macro/micronutrients that can be radioactively labeled. The described procedure is a novel approach for measurements of intestinal nutrient absorption and gut permeability in which luminal nutrient concentrations resemble physiological concentrations.

Cerebrospinal fluid-contacting neurons (CSF-cNs) are a specialized group of multifunctional neurons located around the central canal of the spinal cord. They play critical roles in motor regulation, postural maintenance, and spinal cord injury repair. However, the molecular mechanisms underlying the multifunctionality of CSF-cNs remain poorly understood, partly due to the lack of established in vitro methods for their efficient selection and purification, which significantly hinders mechanistic investigations. In this study, we describe a standardized method using a PKD2L1 promoter-driven lentiviral system, which enables effective enrichment and identification of CSF-cNs in vitro through GFP labeling and puromycin selection. This protocol includes key steps such as construction of the PKD2L1 promoter-driven lentiviral vector, spinal cord tissue collection and digestion from neonatal mice, lentiviral infection, antibiotic selection, and immunofluorescence-based identification of CSF-cNs. Our method provides a reliable platform for obtaining high-purity CSF-cNs (>99%), which facilitates their functional and mechanistic studies for regenerative approaches in vitro.

Preserving biological samples in the field is essential for ensuring high-quality nucleic acid extraction and reliable downstream molecular analyses. Broadly, two main preservation strategies are available: physical preservation, such as flash freezing in liquid nitrogen, which halts enzymatic activity by rapid cooling, and chemical preservation, using stabilizing reagents that inactivate nucleases and protect nucleic acids even at ambient temperatures. This protocol presents a comparative approach using liquid nitrogen and a commercial stabilizing reagent (DNA/RNA Shield, Zymo Research) to preserve tissue from five marine invertebrate species: two cold-water corals, two sponges, and one bivalve. Samples preserved by each method were processed with the AllPrep DNA/RNA Mini kit (Qiagen) to extract both RNA and DNA. RNA quality was assessed using RNA Integrity Number (RIN) scores. The stabilizing reagent preserved high-quality RNA in sponge and bivalve samples but did not prevent RNA degradation in coral tissues, which showed lower RIN scores compared to those preserved in liquid nitrogen. DNA yields were also consistently lower in tissues preserved with DNA/RNA Shield across all species. These findings suggest that DNA/RNA Shield can be a viable alternative to liquid nitrogen for some marine invertebrates, particularly in field conditions where cryopreservation is impractical. However, for cold-water corals, liquid nitrogen remains essential to ensure RNA integrity for transcriptomic analyses and other sensitive molecular applications (e.g., RT-qPCR).

Protein phosphorylation is a dynamic post-translational modification that regulates fundamental processes, including signal transduction, cell proliferation, differentiation, and effector function of immune cells. The Janus Kinase/Signal Transducer and Activator of Transcription (JAK/STAT) pathway is a key mediator of cytokine responses, essential for maintaining immune cell homeostasis and determining cell fate across diverse immune subsets. Dysregulation of JAK/STAT signaling has been linked to a broad spectrum of pathologies, including monogenic immune disorders, autoimmunity, and cancer. Platforms facilitating single-cell analysis of protein phosphorylation offer the ability to reveal subtle signaling defects and dissect the pleiotropy in cellular composition and phosphorylation status, providing insights into immune phenotype and function, while identifying potential therapeutic targets. While an application of cytometry-by-time-of-flight, termed phospho-CyTOF, has proven invaluable for studying protein phosphorylation in cryopreserved peripheral blood mononuclear cells (cPBMCs), its application is limited by cell loss and signaling artifacts stemming from isolation and cryopreservation. Conversely, whole blood (WB) approaches, preserving the native immune cell composition and signaling context, offer a more physiological representation but necessitate robust and consistent protocols for broad application. Herein, we present optimized dual phospho-CyTOF workflows tailored for both cPBMCs and whole blood, building upon established protocols for cytokine stimulation of both samples. These workflows facilitate comprehensive, high-dimensional profiling of JAK/STAT signaling in response to pleiotropic cytokines such as Type I interferons (IFN-α), Type II interferons (IFN-γ), and Interleukin-21 (IL-21). By leveraging CyTOF's capacity for high-dimensional profiling using pure heavy metal–labeled antibodies, these protocols aim to identify pathway-specific alterations in STAT phosphorylation across major immune subsets that may be overlooked by traditional flow cytometry. Together, these optimized dual workflows provide scalable, translationally relevant tools for dissecting the subtle and differential JAK/STAT-driven immune responses in both clinical and research settings, while also being compatible with the simultaneous assessment of crosstalk with alternative immune cell signaling pathways.

Real-time quantitative PCR (qPCR) is a pivotal technique for analyzing gene expression and DNA copy number variations. However, the limited availability of user-friendly software tools for qPCR data analysis presents a significant challenge for experimental biologists with limited computational skills. To address this issue, we developed Click-qPCR, a user-friendly and web-based Shiny application for qPCR data analysis. Click-qPCR streamlines ΔCq and ΔΔCq calculations using user-uploaded CSV data files. The interactive interface of the application allows users to select genes and experimental groups and perform Welch’s t-tests and one-way analysis of variance with Dunnett’s post-hoc test for pairwise and multi-group comparisons, respectively. Results are visualized via interactive bar plots (mean ± standard deviation with individual data points) and can be downloaded as publication-quality images, along with summary statistics. Click-qPCR empowers researchers to efficiently process, interpret, and visualize qPCR data regardless of their programming experience, thereby facilitating routine analysis tasks. Click-qPCR Shiny application is available at https://kubo-azu.shinyapps.io/Click-qPCR/, while its source code and user guide are available at https://github.com/kubo-azu/Click-qPCR.

In neuropharmacology and drug development, in silico methods have become increasingly vital, particularly for studying receptor–ligand interactions at the molecular level. Membrane proteins such as GABA (A) receptors play a central role in neuronal signaling and are key targets for therapeutic intervention. While experimental techniques like electrophysiology and radioligand binding provide valuable functional data, they often fall short in resolving the structural complexity of membrane proteins and can be time-consuming, costly, and inaccessible in many research settings. This study presents a comprehensive computational workflow for investigating membrane protein–ligand interactions, demonstrated using the GABA (A) receptor α5β2γ2 subtype and mitragynine, an alkaloid from Mitragyna speciosa (Kratom), as a case study. The protocol includes homology modeling of the receptor based on a high-resolution template, followed by structure optimization and validation. Ligand docking is then used to predict binding sites and affinities at known modulatory interfaces. Finally, molecular dynamics (MD) simulations assess the stability and conformational dynamics of receptor–ligand complexes over time. Overall, this workflow offers a robust, reproducible approach for structural analysis of membrane protein–ligand interactions, supporting early-stage drug discovery and mechanistic studies across diverse membrane protein targets.

Vestibulo-ocular reflexes (VORs) are compensatory ocular reflexes that maintain stable vision during head movements. In research, VORs encompass angular VOR (aVOR) and off-vertical axis rotation (OVAR) tests, which various groups have employed to assess vestibular function in mice. This protocol outlines the process for measuring VORs in mice, including eye rotation calibration, immobilizing the mouse with a noninvasive setup, configuring the aVOR and OVAR stimulus modes, and interpreting the obtained waveforms to derive VOR values. As technology advances, VORs are expected to yield more qualitative and quantitative insights into the function of the horizontal semicircular canal cristae (HSCC) and the otolith organs. This methodology can serve as a standard for evaluating common vestibular deficits in mice.

The antibody-uptake assay is a commonly used technique to monitor endocytosis of integral membrane proteins including transmembrane and glycosylphosphatidylinositol-anchored proteins (GPI-APs). The antibody-uptake assay typically involves incubating live cells with fluorophore-conjugated antibodies directed against the extracellular domain of the integral membrane protein of interest. Antibody uptake is then detected by flow cytometry or confocal microscopy. However, these detection modalities may be inaccessible to some labs or require extensive training to operate. Thus, we developed an easy and novel sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and western blot-based approach to the antibody-uptake assay that exploits the strong affinity between biotin and streptavidin. Instead of incubating cells with fluorophore-conjugated antibodies to monitor antibody uptake, our assay involves incubating cells with biotinylated antibodies, processing the cell lysates for western blot, and probing the membrane with detectably conjugated streptavidin. From preparation to quantification, this protocol requires less hands-on time than other approaches and is amenable to small-scale drug or siRNA screens. Here, we demonstrate the utility of our approach using the well-characterized misfolded GPI-AP, YFP-tagged C179A mutant of prion protein (YFP-PrP*), as our model substrate. YFP-PrP* constitutively traffics to the plasma membrane (PM), where it binds to anti-GFP antibody, and immediately undergoes endocytosis to lysosomes. To validate our protocol, we present measurements of antibody uptake under conditions known to enhance or inhibit YFP-PrP*’s traffic to the PM. Using this assay, we present new evidence that, under certain conditions, YFP-PrP* is able to undergo degradation via a pathway that does not involve exposure on the cell surface.

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.

Immunohistochemistry (IHC) and immunofluorescence (IF) are fundamental molecular biology techniques to assess protein expression. However, the melanin present normally in the eye in the uveal tract (choroid, iris, and ciliary body) and the retinal pigment epithelium (RPE) poses a significant challenge for IHC and IF. This is because melanin interferes with both chromogenic and fluorescent detection methods. Additionally, formalin fixation, which is commonly used for IHC, can result in shrinkage and loss of cellular detail in the eye. This protocol provides an optimized approach using Davidson’s fixative with a hydrogen peroxide bleaching step to eliminate melanin interference in the mouse eye, improving the quality and interpretability of IHC analyses of the uveal tract and RPE. It is particularly useful for the analysis of uveal melanoma.

Oxygen tension is a key regulator of early human neurogenesis; however, quantifying intra-tissue O₂ in 3D models for an extended period remains difficult. Existing approaches, such as needle-type fiber microsensors and intensity-based oxygen probes or time-domain lifetime imaging, either perturb the organoids or require high excitation doses that limit the measurement period. Here, we present a step-by-step protocol to measure intra-organoid oxygen in human cerebral organoids (hCOs) using embedded ruthenium-based CPOx microbeads and widefield frequency-domain fluorescence lifetime imaging microscopy (FD-FLIM). The workflow covers dorsal/ventral cerebral organoid patterning, organoid fusion at day 12 with co-embedded CPOx beads, standardized FD-FLIM acquisition (470-nm external modulation, 16 phases at 50 kHz, dual-tap camera), automated bead detection and lifetime extraction in MATLAB, and session-matched Stern–Volmer calibration with Ru(dpp)₃(ClO₄)₂ to convert lifetimes to oxygen concentration. The protocol outputs per-bead oxygen maps and longitudinal patterns stratified by bead location (intra-organoid vs. gel) and sample state (healthy vs. abnormal), enabling direct linkage between developmental growth and oxygen dynamics.

Immunopeptidomics enables the identification of peptides presented by major histocompatibility complex (MHC) molecules, offering insights into antigen presentation and immune recognition. Understanding these mechanisms in hypoxic conditions is crucial for deciphering immune responses within the tumor microenvironment. Current immunopeptidomics approaches do not capture hypoxia-induced changes in the repertoire of MHC-presented peptides. This protocol describes the isolation of MHC class I-bound peptides from in vitro hypoxia-treated cells, followed by liquid chromatography-tandem mass spectrometry (LC-MS/MS) analysis. It describes optimized steps for cell lysis, immunoaffinity purification, peptide elution, and MS-compatible preparation under controlled low-oxygen conditions. The method is compatible with various quantitative mass spectrometry approaches and can be adapted to different cell types. This workflow provides a reliable and reproducible approach to studying antigen presentation under hypoxic conditions, thereby enhancing physiological relevance and facilitating deeper immunological insights.

DNA methylation is a crucial epigenetic modification that influences gene expression and plays a role in various biological processes. High-throughput sequencing techniques, such as bisulfite sequencing (BS-seq) and enzymatic methyl sequencing (EM-seq), enable genome-wide profiling of DNA methylation patterns with single-base resolution. In this protocol, we present a bioinformatics pipeline for analyzing genome-wide DNA methylation. We outline the step-by-step process of the essential analyses, including quality control using FASTQ for BS- and EM-seqs raw reads, read alignment with commonly used aligners such as Bowtie2 and BS-Seeker2, DNA methylation calling to generate CGmap files, identification of differentially methylated regions (DMRs) using tools including MethylC-analyzer and HOME, data visualization, and post-alignment analyses. Compared to existing workflows, this pipeline integrates multiple steps into a single protocol, lowering the technical barrier, improving reproducibility, and offering flexibility for both plant and animal methylome studies. To illustrate the application of BS-seq and EM-seq, we demonstrate a case study on analyzing a mutant in Arabidopsis thaliana with a mutation in the met1 gene, which encodes a DNA methyltransferase, and results in global CG hypomethylation and altered gene regulation. This example highlights the biological insights that can be gained through systematic methylome analysis. Our workflow is adaptable to any organism with a reference genome and provides a robust framework for uncovering methylation-associated regulatory mechanisms. All scripts and detailed instructions are provided in GitHub repository: https://github.com/PaoyangLab/Methylation_Analysis.

Research on brain disorders, particularly in the field of oncology, requires in vivo models to evaluate various therapeutic approaches, including intracerebral drug delivery. To meet this requirement, the implantation of intracerebral cannulas offers a reliable method for administering candidate therapeutics directly into the brain. This protocol describes a surgical technique for cannula implantation in mice, enabling repeated administration of therapeutic compounds in the context of glioblastoma treatment. The method was designed with an emphasis on using accessible, easy-to-handle, and sterilized tools to optimize surgical outcomes. Particular attention was also given to animal welfare, notably through refined procedures for asepsis, anesthesia, and postoperative care.

In plants, the apoplast contains a diverse set of proteins that underpin mechanisms for maintaining cell homeostasis, cell wall remodeling, cell signaling, and pathogen defense. Apoplast protein composition is highly regulated, primarily through the control of secretory traffic in response to endogenous and environmental factors. Dynamic changes in apoplast proteome facilitate plant survival in a changing climate. Even so, the apoplast proteome profiles in plants remain poorly characterized due to technological limitations. Recent progress in quantitative proteomics has significantly advanced the resolution of proteomic profiling in mammalian systems and has the potential for application in plant systems. In this protocol, we provide a detailed and efficient protocol for tandem mass tag (TMT)-based quantitative analysis of Arabidopsis thaliana secretory proteome to resolve dynamic changes in leaf apoplast proteome profiles. The protocol employs apoplast flush collection followed by protein cleaning using filter-aided sample preparation (FASP), protein digestion, TMT-labeling of peptides, and mass spectrometry (MS) analysis. Subsequent data analysis for peptide detection and quantification uses Proteome Discoverer software (PD) 3.0. Additionally, we have incorporated in silico–generated spectral libraries using PD 3.0, which enables rapid and efficient analysis of proteomic data. Our optimized protocol offers a robust framework for quantitative secretory proteomic analysis in plants, with potential applications in functional proteomics and the study of trafficking systems that impact plant growth, survival, and health.

Following myocardial infarction (MI), myocardial cells undergo cell death, and the necrotic region is replaced by extracellular matrix (ECM) proteins such as collagens. Myofibroblasts are responsible for producing these ECM proteins. Cardiac myofibroblasts are differentiated from resident fibroblasts in response to inflammation. To date, genetically modified mice driven by the Periostin promoter and adeno-associated virus 9 (AAV9) carrying the Periostin promoter have been used for gene transfer into cardiac myofibroblasts. However, these methods require multiple steps and are time-consuming and expensive. Therefore, we developed a method for delivering genes into cardiac myofibroblasts using retroviruses. Specifically, the DNA of the target gene was transfected into Plat-E cells, which are packaging cells, to generate retroviruses. The virus-containing supernatant was then harvested, and the viruses were pelleted by centrifugation and suspended in PBS-containing polybrene. Subsequently, permanent occlusion of the left coronary artery was performed, and 20 μL of viral solution was immediately administered using a 29G needle at a position 1–2 mm below the ligation site in the heart of mice maintained in an open chest state. Using this method, we were able to introduce genes into the myofibroblasts of interest surrounding the MI site.

Xenobiotics, including environmental pollutants such as bisphenols, phthalates, and parabens, are widely present in food, cosmetics, packaging, and water. These compounds can reach the gastrointestinal tract and interact with the gut microbiota (GM), a complex microbial community that plays a key role in host immunity, metabolism, and barrier function. The GM engages in bidirectional communication with the host via the production of bioactive metabolites, including short-chain fatty acids, neurotransmitter precursors, and bile acid derivatives. Dysbiosis induced by xenobiotics can disrupt microbial metabolite production, impair gut barrier integrity, and contribute to the development of systemic disorders affecting distant organs such as the liver or brain. On the other hand, the GM can biotransform xenobiotics into metabolites with altered bioactivity or toxicity. In vitro models of the human GM offer a valuable tool to complement population-based and in vivo studies, enabling controlled investigation of causative effects and underlying mechanisms. Here, we present an optimized protocol for the collection, cryopreservation, and cultivation of human GM under strictly anaerobic conditions for toxicomicrobiomics applications. The method allows the assessment of xenobiotic–GM interactions in a cost-effective and ethically sustainable way. It is compatible with a wide range of downstream applications, including 16S rDNA sequencing, metabolomics, and endocrine activity assays. The protocol has been optimized to minimize oxygen exposure to less than 2 min, ensuring the viability of obligate anaerobes that dominate the gut ecosystem. This approach facilitates reproducible, mechanistic studies on the impact of environmental xenobiotics on human GM.

In the field of osteoarthritis (OA), the identification of reliable diagnostic and prognostic biomarkers in patients with hip lesions such as femoroacetabular impingement (FAI) could have an immeasurable value. Calcium crystal detection in synovial fluids (SFs) is one tool currently available to diagnose patients with rheumatologic disorders. Crystals, such as monosodium urate (MSU) and calcium pyrophosphate (CPP), are identified qualitatively by compensated polarized light, whereas basic calcium phosphate (BCP) crystals are visualized under conventional light microscopy by Alizarin red S (ARS) staining. Here, we present an efficient and straightforward protocol to quantify calcium crystals by spectrophotometric analysis in human osteoarthritic SFs after staining with ARS. The type and size of the different crystal species are confirmed by environmental scanning electron microscopy (ESEM).

Cellular phenomena such as signal integration and transmission are based on the correct spatiotemporal organization of biomolecules within the cell. Therefore, the targeted manipulation of such processes requires tools that can precisely induce the localizations and interactions of the key players relevant to these processes with high temporal resolution. Chemically induced dimerization (CID) techniques offer a powerful means to manipulate protein function with high temporal resolution and subcellular specificity, enabling direct control over cellular behavior. Here, we present the detailed synthesis and application of dual SLIPT and dual SLIPT^NVOC, which expand the SLIPT (self-localizing ligand-induced protein translocation) platform by incorporating a dual-ligand CID system. Dual SLIPT and dual SLIPT^NVOC independently sort into the inner leaflet of the plasma membrane via a lipid-like anchoring motif, where they present the two headgroup moieties trimethoprim (TMP) and HaloTag ligand (HTL), which recruit and dimerize any two ^iK6eDHFR- and HOB-tagged proteins of interest (POIs). Dual-SLIPT^NVOC furthermore enables this protein dimerization of POIs at the inner leaflet of the plasma membrane in a pre-determined order and light-controlled manner. In this protocol, we detail the synthetic strategy to access dual SLIPT and dual SLIPT^NVOC, while also providing the underlying rationale for key design and synthetic decisions, with the aim of offering a streamlined, accessible, and broadly implementable methodology. In addition to the detailed synthesis, we present representative applications and typical experimental outcomes and recommend strategies for data analysis to support effective use of the system. Notably, dual SLIPT and dual SLIPT^NVOC represent the first CID systems to emulate endogenous lipidation-driven membrane targeting, while retaining hallmark advantages of CID technology—the precision over POI identity, recruitment sequence, high spatiotemporal control, and “plug-and-play” flexibility.

Bottom-up tissue engineering using cell spheroids offers many advantages in recapitulating native cell–cell and cell–matrix interactions. Many tissues, such as cartilage, bone, cardiac muscle, intestine, and neural tissues, have been tissue-engineered using cell spheroids. However, previous methods for spheroid assembling, such as mold casting, hydrogel-based bioprinting, or needle array, either lack control over final tissue geometry or face challenges in scalability and throughput. In this protocol, we describe a robust and scalable tissue engineering method for assembling cell spheroids into a thin, planar spheroid sheet. The spheroids are sandwiched between two flexible meshes held by a frame, facilitating uniform spheroid fusion while ensuring nutrient exchange and ease of handling. We demonstrate this method by producing thin cartilage tissue from human mesenchymal stem cells undergoing chondrogenic differentiation. This approach offers a practical platform for producing thin membrane-like tissue constructs for many research and therapeutic applications.

Accurate labeling of excitatory postsynaptic sites remains a major challenge for high-resolution imaging due to the dense and sterically restricted environment of the postsynaptic density (PSD). Here, we present a protocol utilizing Sylites, 3 kDa synthetic peptide probes that bind with nanomolar affinity to key postsynaptic markers, PSD-95 and Gephyrin. eSylites (excitatory Sylites) specifically target the PDZ1 and PDZ2 domains of PSD-95, enabling precise and efficient labeling of excitatory postsynaptic density (ePSD). In contrast, iSylites (inhibitory Sylites) bind to the dimerizing E-domain of the Gephyrin C-terminus, allowing selective visualization of inhibitory postsynaptic density (iPSD). Their small size reduces linkage error and enhances accessibility compared to conventional antibodies, enabling clear separation of PSD-95 nanodomains in super-resolution microscopy. The protocol is compatible with co-labeling using standard antibodies and integrates seamlessly into multichannel immunocytochemistry workflows for primary neurons and brain tissue. This method enables robust, reproducible labeling of excitatory synapses with enhanced spatial resolution and can be readily adapted for expansion microscopy or live-cell applications.

DNA methylation is a fundamental epigenetic mark with critical roles in epigenetic regulation, development, and genome stability across diverse organisms. Whole genome bisulfite sequencing (WGBS) enables single-base resolution mapping of cytosine methylation patterns and has become a standard method in epigenomics. This protocol provides a detailed, step-by-step workflow for WGBS library construction starting from genomic DNA. It includes steps of RNaseA treatment, DNA shearing, end-repair and A-tailing, adapter ligation, bisulfite conversion, library amplification, and quantification. Notably, the method uses self-prepared reagents and customizable index systems, avoiding the constraints of commercial library preparation kits. This flexibility supports cost-effective, scalable methylome profiling, suitable for diverse experimental designs, including high-throughput multiplexed sequencing.

The rhizosphere, a 2–10 mm region surrounding the root surface, is colonized by numerous microorganisms, known as the rhizosphere microbiome. These microorganisms interact with each other, leading to emergent properties that affect plant fitness. Mapping these interactions is crucial to understanding microbial ecology in the rhizosphere and predicting and manipulating plant health. However, current methods do not capture the chemistry of the rhizosphere environment, and common plant–microbe interaction study setups do not map bacterial interactions in this niche. Additionally, studying bacterial interactions may require the creation of transgenic bacterial lines with markers for antibiotic resistance/fluorescent probes and even isotope labeling. Here, we describe a protocol for both in silico prediction and in vitro validation of bacterial interactions that closely recapitulate the major chemical constituents of the rhizosphere environment using a widely used Murashige & Skoog (MS)-based gnotobiotic plant growth system. We use the auto-fluorescent Pseudomonas, abundantly found in the rhizosphere, to estimate their interactions with other strains, thereby avoiding the need for the creation of transgenic bacterial strains. By combining artificial root exudate medium, plant cultivation medium, and a synthetic bacterial community (SynCom), we first simulate their interactions using genome-scale metabolic models (GSMMs) and then validate these interactions in vitro, using growth assays. We show that the GSMM-predicted interaction scores correlate moderately, yet significantly, with their in vitro validation. Given the complexity of interactions among rhizosphere microbiome members, this reproducible and efficient protocol will allow confident mapping of interactions of fluorescent Pseudomonas with other bacterial strains within the rhizosphere microbiome.

Three-dimensional (3D) human brain tissue models derived from induced pluripotent stem cells (iPSCs) have transformed the study of neural development and disease in vitro. While cerebral organoids offer high structural complexity, their large size often leads to necrotic core formation, limiting reproducibility and challenging the integration of microglia. Here, we present a detailed, reproducible protocol for generating multi-cell type 3D neurospheres that incorporate neurons, astrocytes, and optionally microglia, all derived from the same iPSCs. While neurons and astrocytes differentiate spontaneously from neural precursor cells, generated by dual SMAD-inhibition (blocking BMP and TGF-b signaling), microglia are generated in parallel and can infiltrate the mature neurosphere tissue after plating neurospheres into 48-well plates. The system supports a range of downstream applications, including functional confocal live imaging of GCaMP6f after adeno-associated virus (AAV) transduction of neurospheres or immunofluorescence staining after fixation. Our approach has been successfully implemented across multiple laboratories, demonstrating its robustness and translational potential for studying neuron–glia interactions and modeling neurodegenerative processes.

Sheath blight, caused by Rhizoctonia solani, is a major fungal disease of rice that leads to significant yield losses globally. Conventional inoculation methods often fail to achieve consistent and uniform infection, limiting their applicability in antifungal screening studies. This protocol describes a reliable in planta inoculation method for R. solani using mature sclerotia placed at the internodal region of tillering-stage rice seedlings. The procedure includes step-by-step instructions for seed germination, seedling preparation, pathogen culture, artificial inoculation, and post-infection application of antifungal treatments, including botanical compounds such as Ocimum gratissimum essential oil and thymol. Lesion development is monitored and quantified over time, and data are analyzed statistically to evaluate treatment efficacy. The protocol is optimized for reproducibility, scalability, and compatibility with sustainable disease management approaches. It provides a robust platform for evaluating antifungal agents in a biologically relevant and controlled environment.

The RNA-guided Cas enzyme specifically cuts chromosomes and introduces a targeted double-strand break, facilitating multiple kinds of genome editing, including gene deletion, insertion, and replacement. Caulobacter crescentus and its relatives, such as Agrobacterium fabrum and Sinorhizobium meliloti, have been widely studied for industrial, agricultural, and biomedical applications; however, their genetic manipulations are usually characterized as time-consuming and labor-intensive. C. crescentus and its relatives are known to be CRISPR/Cas-recalcitrant organisms due to intrinsic limitations of SpCas9 expression and possible CRISPR escapes. By fusing a reporting gene to the C terminus of SpCas9M and precisely manipulating the expression of SpCas9M, we developed a CRISPR/SpCas9M-reporting system and achieved efficient genome editing in C. crescentus and relatives. Here, we describe a protocol for rapid, marker-less, and convenient gene deletion by using the CRISPR/SpCas9M-reporting system in C. crescentus, as an example.

N⁶-methyladenosine (m⁶A) is the most abundant internal modification in mRNA and is regulated primarily by the balance between the METTL3 methylase complex and two demethylases, FTO (fat mass and obesity-associated protein) and ALKBH5 (α-ketoglutarate-dependent dioxygenase alkB homolog). Reflecting this prevalence, m⁶A participates in virtually every step of RNA metabolism, influencing a wide range of physiological and pathological processes. The first step in studying m⁶A is genome-wide mapping, typically performed by m⁶A-seq, which sequences RNA fragments immunoprecipitated with an m⁶A-specific antibody. This is followed by identification of RRACH motifs (R = A or G; H = A, C, or U) within these sequences, with m⁶A being located at the third nucleotide. The second step involves mutating the putative m⁶A sites to establish a causal link between the modification and downstream biological effects. Since the mapping step has been covered in several detailed protocols, this article focuses on the second step—mutagenesis of RRACH motifs and subsequent functional analysis of the mutations by ectopic expression. The 3′ untranslated region (UTR) of the mouse Runx2 gene is used as an example. The mutant and wild-type sequences are inserted into a luciferase reporter vector and transfected into 293FT cells to evaluate how loss of m⁶A affects luciferase protein levels. The same reporter plasmids are also used in an RNA stability assay with a transcription inhibitor. Although site-specific demethylation of endogenous mRNA would be preferable, it remains technically challenging despite many attempts. Thus, ectopic expression of the mutated target gene remains a widely used and practical alternative.

Nowadays, recombinant proteins are the focus of various research fields, and their use ranges from therapeutic investigations to cellular model systems for the development of therapeutic approaches. Cell systems used for the expression of recombinant proteins should be comparable in terms of yield and expression efficiency. In many research fields, it is desirable to obtain high protein concentrations. A method that combines an easy workflow with rapid results and affordable costs remains missing, and a standardized approach to determining protein concentration in transgenic cell lines is essential for more reliable data analysis. Our protocol demonstrates the cluster fluorescence-linked immunosorbent assay (FLISA), a technique that allows the exact quantification of comparable protein expression amounts. Moreover, it enables the detection of clustered or bound subunits of a protein without necessitating ultracentrifugation. In the present protocol, we demonstrate the utilization of two transgene cell lines, each expressing distinct recombinant proteins, to provide comparability of protein yields and detectable subunit clustering.

Extracellular vesicles (EVs) have emerged as promising carriers for the targeted delivery of therapeutic proteins to specific cells. Previously, we demonstrated that genetically engineered EVs can be used for targeted protein delivery. This protocol details the generation of mannose receptor (CD206)-targeted EVs using a modular plasmid system optimized for production in HEK293T cells. Three plasmids enable customizable EV budding, cargo loading, and surface modification for targeting to antigen-presenting cells (APCs). EVs are isolated via differential centrifugation and chromatography, characterized using transmission electron microscopy (TEM) and nanoparticle tracking analysis (NTA), and validated through functional uptake assays in primary human activated dendritic cells. Our approach combines flexibility in engineering required EVs with robust, reproducible isolation and characterization workflows. Its modularity allows easy adaptation to alternative targets or cargoes, which can be validated immediately through in vitro testing.

This protocol describes the isolation and flow cytometric analysis of extracellular vesicles (EVs) derived from red blood cells, endothelial cells, and platelets in human peripheral blood. The protocol includes steps for preparing platelet-free plasma, fluorescent antibody staining, gating strategies, and technical controls. This protocol was developed within a study on EV release in snakebite-associated thrombotic microangiopathy; the protocol addresses challenges such as variable autofluorescence and heterogeneity in EV origin. It is flexible and can be adapted for alternative antibody panels targeting different cell populations or EV subtypes, including leukocyte-derived EVs.

Nociception is critically shaped by descending modulation of spinal circuits, yet its cellular and synaptic mechanisms remain poorly defined. Elucidating these mechanisms is technically challenging, as it requires simultaneous activation of primary afferents and descending fibers while monitoring the functioning of individual spinal neurons. Here, we present a method to investigate the influence of the rostral ventromedial medulla (RVM), a principal supraspinal structure mediating descending modulation, on the activity of spinal lamina I neurons. Our approach combines electrophysiological recordings in ex vivo intact spinal cord preparation with optogenetics, granting several advantages. First, ex vivo preparation spares rostrocaudal and mediolateral spinal architecture, preserving lamina I as well as primary afferent and descending inputs. Second, virally mediated channelrhodopsin-2 (ChR2) expression enables selective photostimulation of RVM-originating fibers. When coupled with patch-clamp recordings, this photostimulation allows identifying postsynaptic inputs from RVM to spinal neurons and revealing RVM-dependent presynaptic inhibition of primary afferent inputs. Overall, our approach is well-suited for investigating both pre- and postsynaptic mechanisms of descending modulation in physiological and pathological pain conditions.

When plants undergo senescence or experience carbon starvation, leaf cells degrade proteins in the chloroplasts on a massive scale via autophagy, an evolutionarily conserved process in which intracellular components are transported to the vacuole for degradation to facilitate nutrient recycling. Nonetheless, how portions of chloroplasts are released from the main chloroplast body and mobilized to the vacuole remains unclear. Here, we developed a method to observe the autophagic transport of chloroplast proteins in real time using confocal laser-scanning microscopy on transgenic plants expressing fluorescently labeled chloroplast components and autophagy-associated membranes. This protocol enabled us to track changes in chloroplast morphology during chloroplast-targeted autophagy on a timescale of seconds, and it could be adapted to monitor the dynamics of other intracellular processes in plant leaves.

Telomere length maintenance is strongly linked to cellular aging, as telomeres progressively shorten with each cell division. This phenomenon is well-documented in mitotic, or dividing, cells. However, neurons are post-mitotic and do not undergo mitosis, meaning they lack the classical mechanisms through which telomere shortening occurs. Despite this, neurons retain telomeres that protect chromosomal ends. The role of telomeres in neurons has gained interest, particularly in the context of neurodegenerative diseases such as amyotrophic lateral sclerosis (ALS), where aging is a major risk factor. This has sparked interest in investigating telomere maintenance mechanisms in post-mitotic neurons. Nevertheless, most existing telomere analysis techniques were developed for and optimized using mitotic cells, posing challenges for studying telomeres in non-dividing neuronal cells. Thus, this protocol adapts an already established technique, the combined immunofluorescence and telomere fluorescent in situ hybridization (IF-FISH) on mitotic cells to study the processes occurring at telomeres in cortical neurons of the mouse ALS transgenic model, TDP-43 rNLS. Specifically, it determines the occurrence of DNA damage and the alternative lengthening of telomeres (ALT) mechanism through simultaneous labeling of the DNA damage marker, γH2AX, or the ALT marker, promyelocytic leukemia (PML) protein, together with telomeres. Therefore, the protocol enables the visualization of DNA damage (γH2AX) or the ALT marker (PML) concurrently with telomeres. This technique can be successfully applied to brain tissue and enables the investigation of telomeres specifically in cortical neurons, rather than in bulk tissue, offering a significant advantage over Southern blot or qPCR-based techniques.

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.