The role of the human microbiota in host health is an important area of research. Conventional characterization of the complex microbial communities colonizing the human body is primarily performed using culture-based or high-throughput sequencing approaches, which have improved general understanding yet lack cellular features relevant for interaction and function at the single-cell level. Flow cytometry is an established tool for single-cell analysis, and recent improvements, such as increased resolution of small cells and particles, offer possibilities for characterizing complex microbial communities. Here, we describe an updated protocol to characterize complex microbial communities derived from human stool samples at the single-cell level by multiparametric microbiota flow cytometry. Our protocol covers the process from the isolation of bacteria from stool, preparation of cryo-stocks, staining procedure for phenotypic features of the bacteria, and flow cytometric analysis. Our phenotypic characterization includes light scattering properties, quantitative DNA staining, isotype-specific staining of host-antibody coating, and profiling of bacterial cell surface sugar moieties. In addition, we highlight the importance of determining the bacterial load for improved biological interpretation. External staining controls ensure reproducibility and quality controls. Additionally, we suggest a downstream analysis pipeline involving segmentation of multivariate data by a self-organized map (SOM) and machine learning to generate specific microbiota fingerprints. Optionally, bacteria with specific phenotypic characteristics can be isolated by fluorescence-activated cell sorting (FACS) for further investigation. Our protocol can be applied to any microbial community or sample source and offers the flexibility to be expanded with additional phenotypic markers. © 2026 The Author(s). Current Protocols published by Wiley Periodicals LLC. Basic Protocol 1: Preparation of cryo-preserved bacterial stocks from a single-cell microbiota suspension derived from native human stool sample Basic Protocol 2: Staining protocol for cryo-preserved microbiota stocks and acquisition using a flow cytometer.
Single-nucleotide polymorphisms (SNPs) may influence gene expression and contribute to interindividual variability in immune responses and disease susceptibility. Interleukin-33 (IL33), a cytokine involved in inflammation, immune regulation, epithelial barrier homeostasis, and tissue remodeling, has been implicated in allergic, autoimmune, inflammatory, and neoplastic disorders. The IL33 rs1929992 (T > C) polymorphism, located in intron 3, has been associated with altered IL33 expression, with the T allele linked to increased cytokine production compared with the C allele. This article describes a standardized sequence-specific primer polymerase chain reaction (PCR-SSP) protocol for genotyping the human IL33 rs1929992 polymorphism, with validation by Sanger sequencing. The protocol includes optimization of annealing temperature, MgCl2 concentration, primer balance, deoxynucleotide triphosphates (dNTPs), and DNA input to ensure amplification specificity and reproducibility. Allele-specific amplification is achieved using forward primers specific for the T and C alleles in independent reactions and analyzed by agarose gel electrophoresis, with the human growth hormone (HGH) gene used as an internal control. This protocol provides a simple, reproducible, and accessible method for rs1929992 genotyping, suitable for population-based and genetic association studies involving immune-mediated and inflammatory diseases. © 2026 The Author(s). Current Protocols published by Wiley Periodicals LLC. Basic Protocol 1: PCR-SSP genotyping of IL33 rs1929992 Support Protocol 1: Extraction of genomic DNA by salting-out.
Phages remain severely under-sampled, largely due to the limitations of current isolation methods, which are labor-intensive, low-throughput, and often restricted to a narrow set of bacterial hosts. To address this gap, we developed a robust and scalable field-to-genome workflow that leverages laboratory automation to screen environmental samples against a comprehensive 188-strain bacterial host library using a 384-well plate format. We demonstrate the workflow using Acinetobacter baumannii as the primary model host. Phage activity is detected through Bacterial Lysis Observation over Time assay (BLOTS) using spectrophotometry. Subsequently, isolation and subculture of the phages are done with a single layer agar spot assay. Phage genomes are then sequenced using long-read technology and annotated. This enables systematic and reproducible phage detection and enrichment from diverse sample types. The use of large numbers of host strains streamlines phage discovery and substantially increases the chances of isolating novel phages. The BLOTS high-throughput approach can also be used for screening phage collections against specified clinical hosts of interest for phage research and candidate selection for phage therapy applications. © 2026 The Author(s). Current Protocols published by Wiley Periodicals LLC. Basic Protocol 1: Preparation of host libraries and multi-host primary enrichment Basic Protocol 2: Preparation of the BLOTS screening plate using an automated liquid handler Alternate Protocol 1: Manual phage screening without laboratory automation Basic Protocol 3: Phage screening using BLOTS Basic Protocol 4: Phage validation using the single-layer agar spot assay Basic Protocol 5: Phage amplification for downstream applications Basic Protocol 6: Phage concentration using PEG-8000 precipitation Alternate Protocol 2: Phage concentration using centrifugal filtration Basic Protocol 7: Automated DNA extraction with magnetic beads Basic Protocol 8: Whole-genome sequencing using long reads Basic Protocol 9: De novo genome assembly using the Phage Galaxy platform Basic Protocol 10: Annotation and characterization of phage genome Basic Protocol 11: Host range testing and characterization of isolated phages using BLOTS.
Single-cell RNA sequencing (scRNA-seq) reveals the transcriptional heterogeneity of cells, revolutionizing our understanding of cellular processes. However, the static snapshots obtained from scRNA-seq fail to reveal the time-resolved dynamics of transcription, which impedes critical insights into various biological processes, such as cellular differentiation, embryonic development, disease progression, and responses to external stimuli. Here, we describe Well-TEMP-seq, a protocol for massively parallel profiling of the temporal dynamics of single-cell gene expression. Well-TEMP-seq combines metabolic RNA labeling with a microwell-based scRNA-seq method, Well-paired-seq, to distinguish newly transcribed RNAs marked by T-to-C substitutions from pre-existing RNAs in each of thousands of single cells. Well-TEMP-seq is high-throughput, cost-effective, accurate, and provides a low cell loss rate and high single cell/bead pairing efficiency. More importantly, Well-TEMP-seq can be easily set up in other labs, and the loading of cells and beads can be easily accomplished by an optical microscope and a pipette. We believe that Well-TEMP-seq will be widely adopted and help researchers perform transformative research to unveil the dynamics of single-cell gene expression in diverse biological processes. © 2026 Wiley Periodicals LLC. Basic Protocol 1: Well-paired-seq chip fabrication Basic Protocol 2: Well-TEMP-seq sample processing Basic Protocol 3: Bioinformatics analysis.
Mitochondria are essential for maintaining the high energetic demands of the heart and brain, generating ATP required for contractile function, neuronal signaling, and ionic homeostasis. In both tissues, metabolic flexibility is critical for maintaining bioenergetic efficiency, redox balance, and cellular viability. Despite their importance, existing experimental approaches to assess mitochondrial bioenergetic function present notable limitations. Isolated mitochondria and permeabilized cell assays provide precise control over substrates and respiratory states but disrupt organelle integrity and remove native cellular and extracellular context. Conversely, measurements in isolated or cultured cells preserve intact mitochondria but introduce phenotypic and metabolic artifacts. These constraints highlight the need for an intermediate platform that preserves native tissue architecture and cellular diversity while enabling quantitative assessment of mitochondrial bioenergetics. Building on prior demonstrations of respiration measurements in intact cardiac and neural tissue, we describe a tissue punch-based approach that enables region-specific analysis of mitochondrial function in small ex vivo tissue slices of intact heart and brain. This method preserves cytoarchitecture, and intercellular interactions while remaining compatible with high-resolution Seahorse respirometry analysis. Tissue punches allow multiple technical replicates from individual organs, reduce variability associated with isolation procedures, and enable assessment of regional metabolic heterogeneity, such as atrial versus ventricular myocardium or discrete brain regions. Here we present detailed and reproducible workflow protocols for brain and cardiac tissue punch preparation and extracellular flux analysis, including guidance on sample acquisition, punch sizing, normalization strategies, and data interpretation, with considerations for adapting the protocols across multiple pre-clinical models. © 2026 Wiley Periodicals LLC. Support Protocol: Preparation, Reagent Setup, and Instrumentation Basic Protocol 1: Cardiac tissue preparation Basic Protocol 2: Brain tissue preparation: rodents Basic Protocol 3: Brain tissue preparation: nonhuman primates Basic Protocol 4: Placement of tissue punches on Seahorse organoid plate Basic Protocol 5: Mitochondrial respiration measurements Basic Protocol 6: Tissue disruption for total protein quantification.
Targeting structured RNA elements with small molecules has emerged as a promising yet technically challenging strategy for antiviral drug discovery. Here, we present a comprehensive and experimentally validated workflow for the integrative in silico and in vitro screening of RNA-binding small molecules. The approach is exemplified using conserved RNA elements from the SARS-CoV-2 genome, including the 5'-terminal stem-loop 1 and the programmed -1 ribosomal frameshift pseudoknot, but is broadly applicable to other structured RNAs. The workflow integrates high-resolution RNA structural ensemble generation with virtual screening (VS) and nuclear magnetic resonance (NMR)-based experimental validation. Conformational ensembles generated by fragment-assembly approaches serve as targets for docking chemically diverse fragments and lead-like libraries. Top-ranked compounds are prioritized through consensus scoring and evaluated using ligand- and RNA-observed NMR experiments to confirm binding, characterize interaction modes, and assess specificity. Such ranking allows for NMR-guided fragment optimization, which enables systematic improvement of solubility, affinity, and selectivity through iterative medicinal chemistry in the pharmaceutical pipeline. Detailed procedures are provided for library preparation, ensemble-based VS, hit validation, data interpretation, and progression toward functional assays, together with practical considerations, quality-control parameters, and troubleshooting guidance to ensure reproducibility. By combining computational and experimental strategies that select for high-specificity ligands within a unified framework, this set of protocols accelerates the discovery and optimization of RNA-targeting small molecules and provides a scalable platform for RNA-focused drug discovery. © 2026 The Author(s). Current Protocols published by Wiley Periodicals LLC. Basic Protocol 1: Target and library preparation for virtual screening Basic Protocol 2: Ensemble-based virtual screening of low-molecular-weight compounds against RNA targets Basic Protocol 3: Comparative hit prioritization and selectivity filtering for RNA-binding small molecules Basic Protocol 4: Preparation of RNA samples for in vitro validation of small-molecule binding Basic Protocol 5: NMR-based in vitro screening of RNA-small molecule interactions Support Protocol: Preparation of ligand stocks and NMR-based quality control Basic Protocol 6: Characterization and prioritization of validated RNA-binding hits.
Although a drying technology majorly aims at producing dried pineapple slices for enhancing their shelf-life, it also simultaneously affects other characteristics of pineapple slices, e.g., nutrients, bioactive compounds, quality attributes, microbiological and enzymatic activities, physical properties, and sensory attributes, such as taste, texture, appearance, and flavor. The optimization of drying technologies requires consideration of overall attributes of the slices. Thus, it is important for pineapple drying industries to have knowledge of the techniques that can be used for characterization of dried pineapple slices. In this overview article, detailed procedures have been compiled for characterization of dried pineapple slices. Articles were extracted from Scopus (search phrase: "pineapple" AND "slices" AND "drying" OR "dehydration") and techniques were compiled for raw material characteristics, sample preparation, and the characteristics of dried pineapple slices namely as nutritional (moisture, proximate, vitamins C and B-complex, pectin, minerals), physicochemical (water activity, total soluble solids, pH, and water activity), functional (phenolic compounds, flavonoids, carotenoids, antioxidant and enzymatic activity), physical (mechanical properties, microstructure, shrinkage and porosity, shape), quality (color, texture, volatiles, rehydration, sensory), and shelf-life (microbiological, sorption, packaging, and storage) characteristics. © 2026 Wiley Periodicals LLC.
Tissues are highly ordered structures, with cellular localization essential to the optimal function of the tissue. In lymph nodes the spatial arrangement of immune cells and their associated with stromal cell architecture is essential for the generation of optimal immune responses. Circumstances such as aging and chronic disease can alter lymph node architecture in a manner that impacts immune function. Immunologists have developed many tools for analyzing the cellular content of lymphoid tissue, with flow cytometry a central workhorse in the field. While flow cytometry is a powerful high-throughput tool for analyzing multiple cell types in a tissue, it cannot assess where those cells are located. Imaging, on the other hand, is relatively low throughput, with objective image analysis often presenting a significant hurdle for many immunologists. Here we describe two analysis tools built in Fiji (ImageJ) that semi-automate the analysis of lymph node spatial architecture, can be adapted to different antibody staining panels, and are straightforward to implement with little computational experience. © 2026 The Author(s). Current Protocols published by Wiley Periodicals LLC. Basic Protocol 1: Quantifying changes in lymph node regional architecture Alternate Protocol: Adapting KVA1 for use on spleen tissue sections Basic Protocol 2: Measuring follicular dendritic cell network area within individual B cell follicles.
The internal mechanics of living tissues, cells, and nuclei exhibit exquisite complexity, depend on dynamic structural changes, and influence biological functions such as tissue remodeling, cell homeostasis and migration, and gene expression. Full-field methods to reveal the internal mechanics of extracellular matrix, cells, and nuclei are emerging and can uncover new discoveries in mechanobiology. Here we detail a noninvasive technology to probe the internal mechanics of extracellular matrix, cells, and nuclei using a combination of imaging and deformation-matching computational approaches. Important to this technology is the acquisition of image data showing sufficient spatial and temporal resolution to capture a motion event, which can be readily evaluated using automated assessment of structural features. With careful consideration of image acquisition and processing parameters, micron- to nanometer-scale intracellular and intranuclear mechanics can be reliably measured with small displacement errors (<0.04 µm) using a conventional confocal or wide-field microscope. Further, the technology can be used to spatially correlate a multitude of biological events in complex biological problems. © 2026 Wiley Periodicals LLC. Basic Protocol: Quantifying cellular and nuclear deformation using deformation microscopy.
BACKGROUND:Non-specific chronic low back pain (NSCLBP) is a common musculoskeletal condition characterized by impaired neuromuscular control, reduced postural stability, and functional deficits. Core stabilization exercises (CSE) are widely used but may have limitations in coordinated, multi-muscle activation. Dynamic neuromuscular stabilization (DNS), based on developmental kinesiology, seeks to restore neuromuscular control by engaging integrated diaphragm and core muscle activation. Despite their clinical use, direct comparative evidence between CSE and DNS in NSCLBP remains scarce. OBJECTIVE:To compare the effects of DNS and CSE on pain, lumbar muscle electromyography (EMG) activity, balance, posture, kinesiophobia, and quality of life in patients with NSCLBP Methods: This study will include two parallel groups. A total of 34 participants aged between 20 and 50 years diagnosed with NSCLBP will be recruited on the basis of inclusion and exclusion criteria. The participants will be divided into two groups: (1) DNS with interferential therapy (IFT) and hot pack (n = 17) and (2) core stabilization exercises with IFT and hot pack (n = 17). The intervention will be given 3 times per week for 6 weeks. Outcome measures will include pain intensity, EMG activity of erector spinae and multifidus muscles, balance, posture analysis, kinesiophobia, and health-related quality of life. Assessments will be conducted at baseline and post-intervention. RESULTS:Normality will be assessed using the Shapiro-Wilk test. Baseline differences will be analyzed by independent samples t-tests, and intervention effects by mixed-design ANOVA. DISCUSSION:Both DNS and CSE are expected to improve pain, neuromuscular activity, balance, posture, kinesiophobia, and quality of life. Comparing objective outcomes across groups will identify which intervention offers greater clinical benefit. CONCLUSION:This trial will provide evidence to guide clinicians in selecting the most effective exercise intervention for NSCLBP rehabilitation. © 2026 Wiley Periodicals LLC.
Hydrogen peroxide (H2O2) plays an important role in liver metabolism as a second messenger under physiological conditions, as well as in the development of oxidative stress and pathology. The major metabolic functions of the liver are carried out by hepatocytes, the parenchymal cells that constitute most of the liver mass. The study of oxidative stress in the liver, in particular the determination of a causal relationship between pathological progression and H2O2 production, requires an optimal model system. The chemogenetic system based on yeast d-amino acid oxidase (DAAO) fused to the C-terminus of the HyPer7 biosensor has been established as a tool for controlled H2O2 production and examination of intracellular H2O2 dynamics in vitro and in vivo. This article details adeno associated virus (AAV)-mediated targeted delivery of the HyPer7-DAAO transgene into mouse liver, followed by isolation of primary hepatocytes for quantitative evaluation of intracellular H2O2 dynamics based on ratiometric HyPer7 fluorescence acquired by wide-field microscopy and subsequent image analysis. © 2026 Wiley Periodicals LLC. Basic Protocol 1: AAVs production and purification Basic Protocol 2: Intraparenchymal injection of AAVs into mouse liver Basic Protocol 3: Primary mouse hepatocyte culture preparation Basic Protocol 4: Wide-field fluorescence microscopy of HyPer7 in primary mouse hepatocyte culture Basic Protocol 5: Time series processing.
Callose is a β-1,3-glucan polysaccharide deposited at the plant cell wall interface. It is involved in numerous plant physiological processes and responses to both biotic and abiotic stresses. Callose deposition under biotic stress conditions is typically associated with pattern-triggered immunity, which is activated upon recognition of pathogen-associated molecular patterns or damage-associated molecular patterns at the cell wall interface. These depositions reinforce compromised and damaged cell walls caused by pathogen invasion. The standard method for visualizing callose deposition in various plant tissues involves aniline blue staining. Aniline blue fluorochrome preferentially binds to β-1,3-glucans, which enables this staining technique to specifically locate callose deposition. Although multiple protocols for callose detection using aniline blue are available in various model plants, such as Arabidopsis, there is no optimized method for lignified lateral root tissues after fungal infection. Lignification of roots can hinder the clear visualization of callose depositions; therefore, it is essential to remove them for improved callose detection. Here, we have optimized a robust and reliable method for detecting callose deposition in soybean lateral roots during Macrophomina phaseolina infections. M. phaseolina is a filamentous, soil-borne, necrotrophic fungus that causes charcoal rot disease in soybean and other crop plants. Here, we also provide a detailed methodology for soybean root infection with M. phaseolina using the root-dip method of inoculation, followed by aniline blue staining. Furthermore, we provide a detailed workflow for employing open-source Fiji software together with the Trainable Weka Segmentation (TWS) plugin to detect and count callose structures in fungal-infected root tissues. This protocol may be applicable for detecting callose deposition in other crop plants during fungal infections. © 2026 Wiley Periodicals LLC. Basic Protocol 1: Infection assay with M. phaseolina using root dip method of inoculation Basic Protocol 2: Staining of M. phaseolina-infected soybean roots with aniline blue and imaging of stained soybean roots using fluorescence microscopy Support Protocol: Callose quantification using ImageJ software combined with the TWS plugin.
Cryogenic electron microscopy (cryo-EM) has become a key method in structural biology for determining macromolecular structures. Numerous computational tools have been developed to build atomic models from cryo-EM density maps. However, relatively few tools are available for modeling protein-nucleic acid complexes. Here, we describe how to use two such methods developed by our group, ComplexModeler and CryoZeta, with a focus on modeling protein-nucleic acid complexes. Both tools are available through the EMSuite web server, a freely accessible platform that hosts multiple methods for cryo-EM structure modeling and validation. ComplexModeler integrates DiffModeler and CryoREAD to construct protein-DNA/RNA complex structures at resolutions of up to 5 Å. DiffModeler employs a diffusion model for backbone tracing, followed by fitting AlphaFold2-predicted protein structures into the traced backbone. CryoREAD identifies nucleotide components (phosphate, sugar, and base), constructs the backbone, assigns sequences, and builds full atomic models of DNA/RNA chains. CryoZeta uses a diffusion-based generative model that integrates sequence-based structure prediction with cryo-EM density features to generate accurate models of proteins, nucleic acids, and their complexes. This article describes how to use these two tools on the EMSuite web server through two modeling examples. © 2026 The Author(s). Current Protocols published by Wiley Periodicals LLC. Basic Protocol 1: Protein-nucleic acid structure modeling using ComplexModeler on the EMSuite server Basic Protocol 2: Protein-nucleic acid structure modeling using CryoZeta on the EMSuite server.
In less than a year, SARS-CoV-2 managed to displace Mycobacterium tuberculosis (Mtb) as the leading cause of death worldwide due to a single infectious agent. Both pathogens affect the respiratory tract, mainly the lungs. However, the impact that a possible Mtb + SARS-CoV-2 co-infection can have on the host response is still unknown. Herein we depict a rigorous system to evaluate the complex interaction between two simultaneous infections in vitro in a lung epithelial cell line (A549). Overall, the process includes eight steps: (1) Mtb culture, (2) cell maintenance, (3) preparation of viral stocks, (4) determination of infectious titers, (5) Mtb and SARS-CoV-2 co-infection, (6) determination of intracellular bacterial load, (7) SARS-CoV-2 viability test, and (8) optional UV-C validation assay for infectivity reduction. This comprehensive protocol will allow experimentalists to study the pathogenesis of co-infection in vitro and facilitate collaborative work in the literature. © 2026 The Author(s). Current Protocols published by Wiley Periodicals LLC. Basic Protocol: In vitro coinfection model using Mtb and SARS-CoV-2 in human lung epithelial cells.
Protein-peptide interactions are central to many biological processes and form the basis for many techniques in basic research and therapeutic discovery. A common research challenge is to identify specific peptide ligands or binding motifs where no initial hit is known. Here we describe a protocol for discovering protein-binding peptides using a very large, target-agnostic yeast surface display library containing approximately 6.1 × 109 unique clones and providing broad coverage of short peptide sequence space. To facilitate its use, this library has been made commercially available at low cost. The described workflow entails thawing and expanding the library, validating target protein reagents, enriching target-binding cells through iterative rounds of cell sorting, monitoring enrichment by analytical flow cytometry, and analyzing selected pools by next-generation sequencing. We emphasize practical considerations that affect selection outcomes, including target labeling strategy, appropriate counterselections, and cell sorting parameters. A selection campaign using these protocols can be completed in a matter of weeks, and will typically result in the identification of thousands of candidate target-binding peptide sequences. These protocols are broadly applicable to many diverse protein targets, provided that the target can be purified and labeled in a suitable format. © 2026 The Author(s). Current Protocols published by Wiley Periodicals LLC. Basic Protocol: Selecting protein-binding peptides from the naïve peptide library using MACS and FACS sorting Support Protocol 1: Thawing, expanding, and cryopreserving the peptide library Support Protocol 2: Preparing and validating target protein reagents Support Protocol 3: Performing analytical flow cytometry to assess bulk library binding Support Protocol 4: Extracting plasmid DNA and preparing NGS libraries from post-selection libraries.
Solid-phase RNA synthesis is challenging because the 2'-hydroxyl requires a protecting group that is stable during chain assembly yet easily removable without degrading the product. Driven by the demand for long RNA therapeutics, we report a synthesis method using 2'-O-acetal levulinic ester (ALE) ribonucleoside 3'-O-phosphoramidites. This approach utilizes a rapid, base-labile on-column deprotection strategy that preserves the 5'-O-dimethoxytrityl (DMTr) group, facilitating DMTr-ON reverse-phase (RP) purification. The resulting protocols enable the efficient production of long, functional RNAs, such as single-guide RNAs with superior yield and purity compared to conventional silyl-based methods, while remaining compatible with diverse chemical modifications. © 2026 The Author(s). Current Protocols published by Wiley Periodicals LLC. Basic Protocol 1: Automated solid-phase synthesis of 2'-ALE RNA Basic Protocol 2: Fast on-column deprotection of 2'-ALE RNA Basic Protocol 3: DMTr-ON reverse-phase high-performance liquid chromatography (RP-HPLC) purification of long RNA Support Protocol 1: Automated deprotection setup of 2'-ALE RNA Support Protocol 2: Denaturing polyacrylamide gel electrophoresis (PAGE) purification of RNA.
Abstract This article describes an efficient acid‐catalyzed cyclization strategy for constructing the tricyclic pyrimido[1,2‐a]purin‐10(3H)‐one scaffold via the reaction of guanine derivatives with 1,1,3,3‐tetramethoxypropane (TMOP) catalyzed by 2,4,5‐trifluorobenzoic acid. The method features mild conditions, eliminates the need for hydroxyl protection, and exhibits excellent functional‐group tolerance, making it applicable to the late‐stage structural diversification of various nucleosides, nucleotides, and oligonucleotides. Using the antiviral drug acyclovir and an azide‐containing guanosine derivative as model substrates, we successfully synthesized a tricyclic acyclovir analogue in 81% yield and an M 1 G‐containing oligonucleotide in 40% yield. This protocol provides a general, convenient, and efficient experimental approach for synthesizing this class of potentially bioactive tricyclic nucleoside analogues. © 2026 Wiley Periodicals LLC. Basic Protocol 1 : Synthesis of tricyclic acyclovir analogue Basic Protocol 2 : Synthesis of M 1 G‐containing oligonucleotide
Abstract This unit describes a method for the synthesis of 2′,4′‐ and 3′,4′‐bridged nucleosides from nucleosides bearing oxime imidate moieties at the 5′ position. Hydrogen abstraction at the 4′‐position of nucleosides by iminyl radicals derived from oxime imidates facilitates the facile generation of 4′‐carbon radicals, which undergo cyclization and subsequent hydrolysis of the imidate residues to afford bridged nucleosides. Using this method, the eight‐step syntheses of S ‐constrained ethyl‐bridged 5‐methyluridine ( S ‐cEt‐T) and 6′ S ‐methyl‐2′‐ O ,4′‐ C ‐ethylene‐bridged 5‐methyluridine (6′ S ‐Me‐ENA‐T), which are 2′,4′‐bridged nucleosides with five‐ and six‐membered bridges, respectively, were achieved. Moreover, 3′‐ O ,4′‐ C ‐bridged 2′‐deoxynucleosides bearing various bases were prepared in five steps. Overall, this H‐abstraction/cyclization strategy provides an efficient route for constructing 2′,4′‐ and 3′,4′‐bridges in fewer steps than with existing methods. © 2026 Wiley Periodicals LLC. Basic Protocol 1 : Synthesis of S ‐constrained ethyl‐bridged 5‐methyluridine ( S ‐cEt‐T) Basic Protocol 2 : Synthesis of 6′ S ‐methyl‐2′‐ O ,4′‐ C ‐ethylene‐bridged 5‐methyluridine (6′ S ‐Me‐ENA‐T) Basic Protocol 3 : Synthesis of 3′‐ O ,4′‐ C ‐bridged nucleosides Support Protocol : Preparation of ( Z )‐ N ‐phenoxy‐4‐(trifluoromethyl)benzimidoyl chloride
This article presents a rigorous and reproducible protocol for the enrichment of plant-derived nanovesicles (PDNVs) from the aquatic plant Wolffia globosa using differential ultracentrifugation combined with sucrose density gradient enrichment. The protocol is specifically optimized to address the technical challenges associated with PDNV enrichment from extremely small aquatic plants that require whole-plant homogenization, a process that inherently increases contamination from intracellular organelles, soluble proteins, and chlorophyll. The workflow integrates gentle homogenization, sequential low- and medium-speed centrifugation, sterile filtration, and an initial ultracentrifugation step to obtain crude PDNVs, followed by density-based separation using a discontinuous sucrose gradient (8%-30% w/v). This combined size- and density-based strategy enables effective enrichment of PDNVs while substantially reducing non-vesicular contaminants. Quality control and characterization are performed in accordance with MISEV2023 recommendations using nanoparticle tracking analysis (NTA) and transmission electron microscopy (TEM). Gradient-enriched PDNVs exhibit a predominantly nanoscale, homogeneous particle population with a mean diameter of approximately 105.3 nm and a particle concentration on the order of 1010 particles/ml. TEM analysis confirms well-preserved membrane-bound vesicles with characteristic round-to-cup-shaped morphology and minimal background contamination, demonstrating a clear improvement over crude PDNV preparations. Quantitative normalization of PDNV yield to plant biomass and assessment of particle-to-protein ratios provide standardized metrics for evaluating preparation quality and reproducibility across independent preparations. Overall, this protocol provides a robust, scalable approach for obtaining enriched PDNVs prepared from W.globosa using commonly available laboratory equipment. The resulting PDNV preparations are suitable for downstream physicochemical characterization, molecular profiling, and functional studies, supporting their application in plant PDNV biology and emerging biomedical research. © 2026 Wiley Periodicals LLC. Basic Protocol 1: Enrichment of plant-derived nanovesicles (PDNVs) from the aquatic plant W. globosa by differential ultracentrifugation and sucrose density gradient.
