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Current Issue in 2026
Volume: 16, Issue: 14
Biochemistry
A Practical Experimental Protocol for Identification and Validation of UFMylation Substrate in Human Cells
UFMylation is an evolutionarily conserved ubiquitin-like modification that covalently conjugates UFM1 to lysine residues of substrates via a sequential E1-E2-E3 enzymatic cascade. UFMylation plays a pivotal role in maintaining cellular homeostasis, and its dysregulation is closely linked to multiple major diseases, including malignant tumors, hematopoietic defects, neurodegenerative disorders, and congenital developmental defects, highlighting its important biological significance. However, few substrates of UFMylation have been reported to date, limiting our deep understanding of the mechanistic functions of this modification. This major bottleneck stems from two major technical limitations: the overwhelming abundance of ribosomal protein L26 (RPL26)-UFM1 conjugates masks signals from low-abundance substrates, and conventional methods rely on cumbersome cotransfection of multiple pathway components with poor efficiency and specificity in UFMylated peptides enrichment. To address these challenges, we have developed an effective and specific experimental protocol for UFMylation detection and large-scale substrate identification. This protocol employs CRISPR-Cas9-mediated gene editing to generate UFSP1/UFSP2 double-knockout (UFSP1KO/UFSP2KO, DKO) HEK293T cells, which completely abrogate de-UFMylation and thus significantly elevate global protein UFMylation levels upon exogenous introduction of mature UFM1-ΔC2. In addition, exogenous co-expression of the E3 ligase core components UFL1 and DDRGK1 can further improve the sensitivity of substrate detection. This protocol enables large-scale identification of UFMylation substrates with modification sites via high-efficiency enrichment with the K-ε-VG antibody and LC-MS/MS analysis.
Ex Vivo Assessment of Extracellular Acidification Rate in Murine Intestinal Tissue
Seahorse metabolic assays are now widely utilized across numerous fields for performing functional assessments of glycolysis and mitochondrial function in adherent or suspension cell culture samples. Seahorse assays measure extracellular acidification rate (ECAR) and oxygen consumption rate (OCR) as a means of assessing glycolysis and mitochondrial function, respectively. Currently, the vast majority of Seahorse metabolic assays are performed using in vitro samples due to the current established standardized method. However, a uniform approach to assess real-time functional measurements of glycolysis and mitochondrial function in ex vivo tissue samples remains elusive. In particular, this protocol was designed to assess glycolysis in ex vivo murine intestinal samples through ECAR measurements using the Agilent Seahorse XFe24 platform with corresponding Islet Capture microplates and screens. This protocol was developed to provide functional measurements of glycolytic metabolism in murine intestinal tissue samples. This protocol details a method to assess glycolysis in tissue samples and represents the next stage of ex vivo metabolic methods to complement existing standardized in vitro approaches. While this protocol was developed to assess ECAR in ex vivo murine intestinal samples, the same approach can be applied to assessing mitochondrial respiration through measurements of OCR in other tissue types. Overall, this protocol expands the purview of Seahorse metabolic assays through the inclusion of tissue samples and provides the framework to interrogate organ-level metabolism in the context of systemic nutrient metabolism and physiology.
Iodine Staining of Glycogen Storage in Caenorhabditis elegans
Glycogen is a highly conserved macromolecule across species, and its visualization provides critical insights into both physiological processes and disease states. Existing approaches for glycogen imaging in Caenorhabditis elegans rely primarily on traditional microscopy slides, which introduce variability in image acquisition and downstream data analysis, limit throughput, and require substantial hands-on time and technical expertise.Here, we present a standardized, cost-effective, and high-throughput imaging method that enables efficient visualization and quantification of glycogen in C. elegans. Our approach utilizes a custom-designed three-dimensional pad containing two to four chambers, allowing control and experimental samples to be processed simultaneously under identical conditions. Worms are exposed to iodine crystals, ensuring uniform staining while minimizing reagent use and handling variability. Imaging is performed using a simple binocular microscope, and analysis is conducted in Fiji, making the workflow accessible to laboratories with minimal specialized equipment or training.This method also reduces technical variability, shortens turnaround time, and requires only basic reagents and expertise, making it well-suited for both research and teaching laboratories. Importantly, the platform is readily adaptable to other nematode species and scalable for large-scale genetic or pharmacological screening applications. Together, this workflow minimizes technical variability and provides a robust platform for comparative glycogen analysis in C. elegans.
Bioinformatics and Computational Biology
Actin Quantification Using the Filamentous Actin Segmentation Tool (FAST)
Studying actin-filament assembly into distinct subcellular structures can provide insights into both physiological cellular processes and the mechanisms of disease. However, there are a limited number of tools that can quantify the organization and abundance of different actin structures from confocal microscopy images of cells expressing Lifeact or fixed and stained with phalloidin. Filamentous actin segmentation tool (FAST) is a deep learning model trained with a unique approach of antibody-assisted annotation, resulting in accurate and efficient quantification of distinct classes of actin structures. Here, we detail the protocol for using antibody-assisted annotation to generate datasets that could be applied to train machine learning models. Additionally, we provide step-by-step instructions for applying FAST on phalloidin-stained or live-cell confocal imaging data using our pretrained model. FAST is open source and freely available, with user-friendly notebooks that enable quantification of different classes of actin structure, without the need for structure-specific antibodies. As such, FAST can be a practical tool for researchers investigating the role of cytoskeletal organization in a range of processes.
Biological Engineering
Preparation and Characterization of Neutrophil Membrane-Fused Mitochondria (nMITO)
Mitochondrial transplantation is an emerging strategy for cellular repair, yet its efficiency is often limited by poor targeting and environmental instability. This protocol details the fabrication and comprehensive characterization of neutrophil membrane-fused mitochondria (nMITO), a hybrid organelle platform designed to combine the metabolic vigor of natural mitochondria with the targeting and anti-inflammatory properties of neutrophil membranes. We describe an optimized workflow for mouse heart mitochondrial isolation, lipopolysaccharide (LPS)-activated neutrophil membrane (NEM) extraction, and the subsequent sonication-mediated fusion process. Characterization techniques include dynamic light scattering (DLS) for size and zeta potential, transmission electron microscopy (TEM) for ultrastructural integrity, and bioenergetic assays [ATP synthesis and tetramethylrhodamine methyl ester (TMRM)-based membrane potential] to ensure functional preservation.
Cancer Biology
Light-Regulated Cancer Immunotherapy Using Individually Encapsulated Synthetic Circuit–Engineered Cells
Cell therapy holds great promise for cancer immunotherapy, but its clinical efficacy is severely hindered by poor post-transplant cell survival, low homing efficiency, and host immune clearance. To address these challenges, this study develops a novel light-controlled immunotherapy strategy that integrates a red/far-red light genetic switch with single-cell encapsulation engineering. The red/far-red light (660/730 nm) reversible regulatory system enables precise spatiotemporal control over the expression of therapeutic proteins in engineered cells (e.g., CAR-T or engineered HEK 293T cells), allowing on-demand activation of anti-tumor immune responses. On this basis, a mild enzyme-mediated single-cell encapsulation technique is further employed to rapidly form a protective hydrogel coating in situ on the cell surface, thereby enhancing the survival of transplanted cells under hostile in vivo microenvironments. This strategy combines precise gene expression regulation with physical protection, improving therapeutic outcomes without the need for genomic modification of the cells. It provides a new paradigm for developing safe, controllable, and efficient cancer immunotherapy.
Cell Biology
A Universal Resazurin-Based Viability Assay for Prokaryotic and Eukaryotic Cells in 2D and 3D Cultures
In vitro cytotoxicity assessments frequently rely on staining-based methods that indirectly estimate viable cell numbers. A major limitation of many such techniques is their endpoint nature, requiring cell lysis or irreversible processing that precludes longitudinal monitoring of cellular responses following treatment. An ideal assay for evaluating cell viability and proliferation should be simple, rapid, cost-effective, reproducible, and highly sensitive, while also enabling accurate quantification with minimal interference from test compounds. The resazurin reduction assay satisfies these criteria, offering a sensitive and economical alternative to conventional tetrazolium-based methods. Although both assay types depend on the metabolic reduction of a dye by viable cells, they differ mechanistically. Tetrazolium salts (e.g., MTT) are reduced by cellular dehydrogenases to insoluble formazan crystals that require solubilization before detection. In contrast, resazurin-a cell-permeable, non-fluorescent blue dye-is reduced to resorufin, a highly fluorescent compound detectable without additional processing steps. This property renders the resazurin assay broadly applicable to viability testing in eukaryotic cells cultured in both 2D and 3D formats, as well as in bacterial systems. Here, we present a resazurin-based reduction assay across diverse experimental models, emphasizing its practicality, reproducibility, and adaptability for real-time viability monitoring.
A Simplified Langendorff-Based Method for Mouse Cardiac Myocyte Isolation
Isolation of adult mouse ventricular myocytes is essential for studying cardiac physiology and cellular function. Traditional methods commonly rely on Langendorff perfusion systems, which provide continuous retrograde coronary perfusion but require specialized equipment and can be complex to operate. Here, we describe a simplified Langendorff-based protocol that uses a syringe pump–driven system to achieve constant-flow retrograde aortic perfusion during enzymatic digestion. The setup incorporates an inline heater for precise temperature control and uses widely available laboratory components, enabling consistent delivery of digestion enzymes. This approach maintains stable perfusion despite changes in coronary resistance and reduces variability associated with conventional gravity-driven systems. The protocol yields high-quality adult ventricular myocytes suitable for downstream functional analyses, including electrophysiology, contractility, and calcium imaging. Compared with traditional systems, this method is more accessible, reduces technical complexity, and improves reproducibility, facilitating adoption in laboratories without dedicated isolated-heart perfusion infrastructure.
Isolation of Mononucleated and Binucleated Hepatocytes by Flow Cytometry
Polyploid hepatocytes are one of the unique features of the liver. Some polyploid hepatocytes have chromosomes in a single nucleus (e.g., 1x4n, 1x8n), while others separate their chromosomes into two nuclei (e.g., 2x2n, 2x4n). In ploidy research, hepatocytes are typically sorted according to their cellular ploidy, revealing their contribution to tumorigenesis and cellular senescence. However, the conventional sorting method fails to distinguish 1x4n from 2x2n, or 1x8n from 2x4n cells, leaving it unclear whether hepatocytes with the same cellular ploidy but different nuclear configurations are identical or phenotypically different. Here, we describe a detailed protocol for fractionating mononucleated and binucleated hepatocytes. First, we present the method for isolating primary mouse hepatocytes and staining them with the DNA dye Hoechst 33342. Flow cytometry is then used to detect fluorescence differences between mononucleated and binucleated hepatocytes. This protocol enables the discrimination of hepatocyte subpopulations with identical cellular ploidy, providing a useful tool to investigate the functional heterogeneity of polyploid hepatocytes.
Developmental Biology
In Vivo Light-Sheet Imaging of Senescence Reporter Activity in a Transparent Killifish
Aging is associated with progressive accumulation of senescent cells, which contribute to tissue dysfunction and organismal decline. Conventional approaches for assessing cellular senescence, such as histological or immunofluorescence analyses of fixed tissue sections and flow cytometry, require tissue collection, thereby precluding longitudinal in vivo studies. To enable the analysis of cellular senescence in a living vertebrate model, we have previously generated a cdkn1a (p21)-driven GFP reporter line that was established in the transparent klara background of Nothobranchius furzeri. Here, we describe a protocol for in vivo light-sheet microscopy of the reporter activity as readout for senescence-associated cell cycle arrest with single-cell resolution. The procedure involves anesthesia and mounting of fish for stable positioning within the imaging chamber, with particular attention to animal welfare considerations. It further includes the acquisition of three-dimensional image stacks and subsequent image processing. The workflow allows monitoring of GFP-positive cells in intact living killifish at different developmental stages. Although imaging depth remains limited despite organismal transparency, this method provides high-resolution volumetric imaging with minimal phototoxicity and enables analysis of senescence dynamics in a short-lived vertebrate model. It is currently performed as a terminal procedure under approved ethical regulations, but longitudinal imaging would also be possible with additional ethical authorization.
Immunology
A Streamlined and Time-Saving Approach to Generate HLA-DR15 MHC Class II Tetramers via In Vivo Biotinylation
This protocol describes an optimized strategy for the efficient generation of peptide-loaded major histocompatibility complex (MHC) class II (pMHC) tetramers, which are essential tools for detecting and characterizing antigen-specific T cells in immunological research. Traditional methods require separate expression of MHC proteins followed by in vitro biotinylation—a multi-step process that is time-consuming and prone to protein loss. Here, we present an integrated approach based on co-expression of MHC monomers and BirA biotin ligase in Expi293F T cells, enabling site-specific biotinylation in vivo during protein synthesis. At the same time, the incorporation of a thrombin-cleavable class II–associated invariant chain peptide (CLIP) peptide into the MHC construct allows flexible loading of any antigenic peptide of interest without the need for re-cloning or re-expression of the MHC molecule. Pre-biotinylated MHC molecules are subsequently purified, loaded with antigenic peptides, and assembled into fluorescent tetramers via streptavidin conjugation. This streamlined workflow significantly reduces handling steps, improves protein yield, and enhances reproducibility. The resulting tetramers are suitable for sensitive detection and isolation of antigen-specific T cells by flow cytometry, supporting applications in T-cell immunogenicity studies, vaccine development, and autoimmune disease research.
Microbiology
Separating Chromera velia Zoospores From Culture and Estimating Their Average Motility Speed and Lifespan
Chromera velia is an apicomplexan alga uniquely positioned as the closest photosynthetic relative to apicomplexan parasites (Sporozoa), which include the human pathogens that cause malaria (Plasmodium) and toxoplasmosis (Toxoplasma). Under favorable conditions, C. velia forms motile zoospores that contribute to dispersal and possibly host interaction. However, zoospores coexist with other developmental stages in culture, making their isolation technically challenging. Previous studies characterized the phototactic behavior of zoospores in several taxa, yet this response has not been used to separate motile zoospores from mixed cultures. Other reported methods for zoospore recovery relied instead on physical or chemical principles such as passive filtration, differential centrifugation, or column-based purification, all of which can compromise zoospore motility and viability through mechanical shear or osmotic changes. To address this limitation, we developed a non-invasive, simple, and effective method for rapid zoospore isolation depending entirely on their negative phototaxis response. Using a directional light gradient, the method enables reliable collection of active, motile zoospores without specialized equipment or chemical treatments. Our protocol is straightforward to reproduce, relies on standard laboratory equipment, can be completed in under two hours, and yields a zoospore fraction of sufficient quality for live-imaging, motility assays, and downstream molecular and -omics applications. It may also be adapted to other flagellated protists with light-responsive motile stages.
Optimized Field Collection and Gut Dissection Workflows for Microbiome Studies of the Citrus Root Weevil, Diaprepes abbreviatus
Careful dissection of insect gut tissues is essential for microbiome studies to ensure accurate characterization of internal microbial communities and preservation of DNA integrity. Because insect-associated microbiomes are highly sensitive to contamination, effective removal of external microbes prior to dissection is critical to minimize bias in downstream analyses. While ethanol- and bleach-based surface sterilization methods are commonly used, standardized workflows integrating field collection, sterilization, and dissection remain limited. Here, we present a step-by-step protocol for the field collection, surface sterilization, and dissection of gut tissues from the agricultural pest Diaprepes abbreviatus (Coleoptera: Curculionidae), optimized for genomic DNA extraction and microbiome analyses. Using wild-caught specimens, this workflow incorporates a rigorous surface sterilization and dissection strategy that minimizes external contamination while preserving biologically relevant microbial signatures and DNA integrity for downstream microbiome analyses. The protocol provides a standardized framework for insect gut microbiome studies and can be broadly adapted to other wild-caught insect species requiring careful collection, disinfection, and sterile dissection prior to molecular analysis. The protocol integrates field collection and laboratory processing steps into a streamlined workflow that minimizes contamination while preserving tissue integrity for downstream applications.
Assessment of Saccharomyces cerevisiae Survival Upon Exposure to Transient High Pressure and Temperature in a High-Intensity Shock Tube for Astrobiology (HISTA)
Understanding microbial survival under extreme planetary conditions is critical for astrobiology and stress biology. Several experimental platforms, including radiation, desiccation, and microgravity, have been used to mimic extraterrestrial environments; however, controlled simulation of high-intensity shock waves has not been used to assess microbial survival. Here, we describe a detailed protocol for shock processing of Saccharomyces cerevisiae using the high-intensity shock tube for astrochemistry (HISTA), which generates high-Mach-number shock waves under inert gas conditions. Yeast cells are drop-casted onto a metal flange, exposed to transient high-pressure shock waves, and recovered for downstream survival and cellular analyses. Shock intensity can be precisely tuned by adjusting driver pressure, diaphragm thickness, and driven gas pressure. This protocol provides a platform to investigate microbial adaptation to shock waves.
Molecular Biology
An Accurate and Precise ddPCR-Based Method for Determining the Concentration of Plasmid DNA
Transient transfection is commonly used for the commercial production of adeno-associated viral particles for gene therapy. In this process, packaging cells such as HEK293 cells are transfected with three plasmids, including the Rep/Cap plasmid, the Helper plasmid, and the gene-of-interest plasmid containing the transgene/gene therapy product. The combination of these plasmids allows for the robust production of recombinant adeno-associated viral particles. As a result, the concentration of these plasmids plays a critical role in viral production and must be accurately assessed. Typically, A260/A280 readings are utilized to measure plasmid titer; however, this approach lacks accuracy and specificity and is susceptible to matrix interference. To address these shortcomings, a digital droplet PCR method was developed to titer plasmids. This method uses a combined restriction digest/PCR protocol to linearize the plasmid template and evaluate copy numbers of a plasmid-specific gene. Qualification demonstrated that the method is highly accurate, specific to plasmid DNA, and impervious to matrix interference.
Neuroscience
Measuring PINK1 Activity in Single Cells Using a PINK1 Kinase Activity Reporter
Phosphatase and tensin homolog-induced kinase 1 (PINK1) is a serine/threonine kinase that plays a key role in mitophagy initiation. Loss-of-function autosomal recessive mutations in PINK1 cause early onset Parkinson’s disease (EOPD). Current approaches for studying PINK1 function depend on bulk techniques that can only provide snapshots of activity and could miss the dynamics and cell-to-cell heterogeneity of PINK1 activity or provide an indirect readout of PINK1 activity. Here, we present a protocol using our newly developed phase separation–based PINK1 biosensor (PINK1-SPARK) to observe real-time activity of endogenous PINK1 in single cells. Following transfection of live cells with PINK1-SPARK, cells are treated with mitochondrial depolarizing agents and visualized using widefield or confocal fluorescence microscopy, either following the same cells over time for time-lapse imaging of PINK1 activity or end-point measurements. Thus, PINK1-SPARK is a new tool that enables the measurement of PINK1 activity in single live cells, allowing for further elucidation of the role of PINK1 in mitophagy and cell function.
Plant Science
Simple Electroporation of Chlamydomonas reinhardtii Strains With an Intact Cell Wall
Chlamydomonas reinhardtii is a model green alga extensively used to study photosynthesis and cilia using molecular biology and genetics. Electroporation is a very common technique to integrate DNA into the nuclear genome, which is essential to generate mutant collections and express transgenes. Here, we describe a simple, fast, and efficient protocol to transform strains with an intact cell wall. The technique achieves good transformation efficiency without cell wall digestion or the use of commercial kits and is compatible with the widely available Gene Pulser electroporation system.
Gene Editing in Chlamydomonas Using the SCREAM Technique
In the model alga Chlamydomonas reinhardtii, CRISPR (clustered regularly interspaced short palindromic repeat)-based gene editing using Cas (CRISPR-associated) enzymes enables both (a) insertion of large gene cassettes and (b) the creation of knockouts based on the introduction of indels, and specific mutations via mutation-directing oligonucleotides. Owing to the relatively low efficiency of this process, selection markers are frequently used to enrich the candidate pool prior to screening, which typically employs PCR. Unfortunately, few selection markers are available for Chlamydomonas. Furthermore, each marker requires different selection media, and deletion of the selectable marker can be difficult. When multiple successive gene editing steps are required, the use of these markers becomes onerous. The SCREAM (sequential CRISPR via recycling endogenous auxotrophic markers) technique employs an endogenous gene as a marker, the mutation of which can be selected both in the forward (loss of function) and reverse (gain of function) directions. During the first gene editing step, crRNA and mutation-directing oligonucleotides are provided for both the marker and the first target gene (Target 1). Candidates with edited marker genes are selected by loss of marker function, prior to screening for the desired modification of the first target gene. Using a successful candidate, a subsequent gene editing step directs reversion of the mutant marker gene to wild-type status, with candidates being selected on auxotrophic media to detect the regain of function of the auxotrophic marker to wild type (i.e., reversion). Simultaneously, a second target gene modification is produced using Target 2–specific crRNA and oligonucleotides. Revertants, now with a wild-type auxotrophic marker, are then screened for the specific mutation of Target 2. This reversion strategy enables a single selectable marker to be reused indefinitely, facilitating the creation of many successive mutations in a single cell line. As the marker can be completely reconstituted, strains can be created in which only the target gene is altered. Employment of homology-directed repair, using single-stranded oligonucleotides for mutation creation, enables the creation of site-directed mutants, tag insertion, and gene knockouts or reversion, rather than the insertion of large gene cassettes. In this implementation, nitrate reductase is used as the endogenous auxotrophic marker, and the adenine phosphoribosyltransferase gene is used as an example of a target gene.
Stem Cell
Histological Processing of Organoids for Immunostaining
Organoids are three-dimensional cell structures derived from stem cells that recapitulate the architecture and function of native tissues. Histological analysis of organoids is essential for assessing their structure, cellular composition, and responses to experimental conditions. However, their small size and fragility make standard paraffin embedding workflows difficult. Here, we describe a robust and reproducible protocol for the fixation, paraffin embedding, and sectioning of human organoids, enabling high-quality histological and immunostaining analysis. The method involves direct fixation within the culture matrix and inclusion in HistoGel to prevent organoid loss during processing. The protocol is compatible with hematoxylin–eosin (H&E) staining and multiplex immunofluorescence. Critical steps, troubleshooting, and adaptations for intestinal and cardiac organoids are discussed. This cost-effective and accessible method supports long-term preservation and detailed structural analysis of organoid models.
Protocol for Measuring Drug–Target Engagement in Mouse Colorectal Cancer Organoids Using NanoBRET Assay
Organoids as a drug discovery platform represent an emerging field that continues to refine its tools. NanoBRET (bioluminescence resonance energy transfer) has emerged as a proximity-based and highly sensitive assay to measure protein–protein and protein–ligand interactions. NanoBRET assays were developed and are currently used for 2D cell line experiments. Here, we present the development of the first organoid-compatible Nanoluciferase (Nluc) for 3D model systems. We utilise the Nluc for NanoBRET assays to test drug–target engagement. We describe steps for seeding, transfecting, and replating of mouse colorectal cancer organoids. In addition, we provide detailed procedures for the NanoBRET assay. Various lines of evidence have shown significant difference in drug response between 2D human cell lines and 3D model systems, including patient-derived organoids. Our protocol provides a template for measuring this difference in the context of drug–target engagement.
Satellite Cell Isolation, Culture, and Infection After Retroviral Preparation
Satellite cells are adult skeletal muscle stem cells that play essential roles in muscle regeneration. Understanding their behavior is critical for elucidating the mechanisms of muscle repair and advancing muscle regenerative therapies. This requires efficient methods for genetic manipulation in these cells. Retroviral-mediated gene delivery is commonly used for stable transgene expression in immortalized cell lines. However, existing approaches are not optimized for primary satellite cells, often resulting in variable efficiency and inconsistent outcomes. Here, we describe an optimized protocol for satellite cell isolation and culture, as well as retroviral production and infection of primary satellite cells that achieves high transduction efficiency. The satellite cell isolation procedure enriches for myofiber fragments prior to satellite cell release, thereby reducing contamination by non-myogenic cells and improving cell purity. Another key feature of this protocol is the concentration of retroviral particles and their resuspension in satellite cell growth medium prior to infection, which minimizes satellite cell exposure to packaging cell-conditioned medium. Compared to standard approaches, this protocol improves both infection efficiency and reproducibility. It is readily adaptable to a wide range of downstream applications, including microscopies, biochemical assays, and molecular biology analyses.