Cell Biology

FlipNanoLuc is a highly sensitive protease biosensor based on the β-strand-flipping principle of NanoLuc luciferase, which is derived from Oplophorus gracilirostris. In the inactive configuration, one β-strand of NanoLuc is repositioned, thereby suppressing luciferase activity. Upon cleavage of the embedded protease recognition sequence by a target protease, the flipped β-strand is released, and luciferase activity is reconstituted. Incorporation of the LgBiT fragment (NanoBiT technology) yields strong luminescent output once the reporter is reconstituted, whereas the CL1-PEST1 degradation tag lowers background luminescence by promoting the degradation of the uncleaved, inactive form; together, these two modifications widen the dynamic range. A firefly luciferase normalization cassette connected via a P2A self-cleaving peptide is encoded in the same reporter plasmid, thereby eliminating the need for separate co-transfection. Because the readout directly reports intracellular protease activity in living cells, the system is suitable for detecting protease activation during apoptosis or viral infection and, in principle, for evaluating protease inhibitors and antiviral compounds. This protocol describes the following: (1) generation of HEK293T cells stably expressing FlipNanoLuc by retroviral transduction; (2) validation of reporter activity by protease overexpression; (3) detection of apoptosis using staurosporine; (4) detection of human coronavirus OC43 infection; and (5) detection of SARS-CoV-2 infection in BHK cells stably expressing hACE2. All luminescence assays employ dual-luciferase normalization and are compatible with standard 384-well plate readers.

The mucus layer lining the human stomach is a critical barrier that protects the underlying epithelium from gastric acid and harmful pathogens such as Helicobacter pylori. The efficacy of this barrier relies on the structural integrity of the mucus, which is determined by various biochemical and biophysical features. Human gastric organoids—3D cellular models that resemble the stomach—contain mucus and have been used to investigate gastric disease. The luminal compartment of three-dimensional epithelial organoids represents a physiologically relevant but experimentally inaccessible microenvironment. In gastric organoids, luminal accumulation of mucus creates a confined viscoelastic hydrogel that mimics native gastric mucus. However, the small volume and topological confinement of organoids preclude conventional bulk rheometry. Here, we describe a particle tracking microrheology (PTM) protocol to measure the viscoelastic properties of the mucus within intact organoid lumina following microinjection of fluorescent microspheres. High-speed fluorescence imaging and particle trajectory analysis enable the quantification of viscous and elastic properties of the mucus through calculation of mean squared displacement (MSD), diffusive scaling exponent (alpha), and frequency-dependent storage (G’) and loss (G’’) moduli. This method enables rheological measurements in nanoliter-scale compartments without disrupting organoid architecture. We further discuss the impact of mucus heterogeneity and microstructure on scale-dependent mechanical behavior. This protocol is broadly applicable to other organoid systems and can be adapted to Transwell or organ-on-chip platforms for in situ luminal measurements.

The aorta–gonad–mesonephros (AGM) region is the site where hematopoietic stem cells (HSCs) first emerge during development, and is therefore widely used to study in vivo hematopoiesis and to discover novel regulatory mechanisms. The endothelial-to-hematopoietic transition (EHT) process can be directly observed via immunofluorescence on frozen sections of the AGM region. However, the mouse AGM region is extremely delicate and lies deep within the embryo, between the notochord and the somatic mesoderm. Here, we present a step-by-step protocol covering embryo collection, fixation, dehydration, and embedding with a defined orientation, followed by frozen sectioning, immunofluorescence staining, and confocal imaging. The protocol is highly reproducible and easy to follow and provides clear instructions on orienting the embryo and anatomically locating AGM. By filling a technical gap, the protocol can enable researchers to reliably study HSC emergence and EHT in the mouse embryonic AGM.

Neuroinflammation disrupts blood–brain barrier (BBB) integrity, promoting leukocyte recruitment into the central nervous system and contributing to the progression of neurological disorders. This protocol describes a reproducible macrophage adhesion assay to evaluate interactions between immune cells and brain endothelial cells and to screen compounds with potential anti-inflammatory activity. Murine brain endothelial cells (bEnd.3) were cultured to confluency and exposed to inflammatory stimuli, such as lipopolysaccharide (LPS) or pilocarpine, a cholinergic muscarinic receptor agonist reported to induce inflammatory responses through seizure-associated neuroinflammatory mechanisms, in the presence or absence of candidate therapeutic compounds. In these studies, the natural flavonoid quercetin and the synthetic alkyl-lysophospholipid edelfosine were tested for their effects on macrophage adhesion. After 48 h of treatment, fluorescently labeled murine macrophages (RAW 264.7) were added to the endothelial monolayer, and adherent cells were quantified by fluorescence microscopy. The assay was validated using dexamethasone as an anti-inflammatory control and inflammatory stimulation with LPS or pilocarpine. As expected, dexamethasone reduced macrophage adhesion, whereas both LPS and pilocarpine significantly increased adhesion, demonstrating the assay's sensitivity to changes in endothelial inflammatory status. Overall, this protocol provides a reliable and accessible platform for investigating endothelial–immune cell interactions under neuroinflammatory conditions and for evaluating therapeutic compounds that may preserve BBB function and reduce inflammatory cell recruitment in neurological disease models.

Platelets originate from megakaryocytes, whose generation involves a series of biological processes including directed differentiation, proliferation, polyploidization, and maturation of hematopoietic stem cells. Abnormalities in megakaryocyte development and maturation can lead to quantitative and functional defects in platelets, thereby contributing to hemostatic or thrombotic disorders as well as the development of malignancies. Investigating megakaryocyte development and maturation and platelet production can provide important theoretical foundations for the diagnosis and treatment of thrombocytopenia, thrombotic diseases, and myeloproliferative neoplasms. Currently, there are three main clinical sources of hematopoietic stem cells (HSCs): bone marrow (BM), peripheral blood (PBSC), and umbilical cord blood (UCB). Among these, umbilical cord blood (UCB)-derived HSCs, due to their higher differentiation efficiency and stronger proliferative capacity, are the preferred starting cell source for studying megakaryocyte (MK) development and maturation and the mechanisms of platelet production. This article describes a detailed protocol covering all necessary steps for isolating CD34⁺ hematopoietic stem cells from umbilical cord blood, followed by in vitro induction culture with stem cell factor (SCF) and thrombopoietin (TPO) to generate mature megakaryocytes that highly express early megakaryocyte markers (CD41a, CD61) and late maturation markers (CD42a, CD42b). This protocol provides an effective tool for studying megakaryocyte development and platelet production and holds potential value for application in research on megakaryocyte-related diseases.

The transient receptor mucolipin subtype 1 (TRPML1) is a ubiquitously expressed ion channel involved in lysosomal homeostasis. Recent pharmaceutical interest in developing agonist ligands has emerged due to beneficial effects in neurodegenerative diseases. The major high-throughput screening techniques to investigate this ion channel involve fluorescent calcium imaging and electrophysiology. Despite their high capacity for screening compounds, it is well known that both methods face hurdles, such as the need for expensive, specialized equipment. Here, we present a novel technique to screen for ligands of TRPML1 using a bioluminescence resonance energy transfer (BRET) assay. This assay consists of a target engagement assay in live cells, which permits the determination of binding constants between ligands and the target of interest in equilibrium or time-dependently. We employ a full-length TRPML1 C-terminally tagged with the small bioluminescent protein nanoluciferase. This ensures the correct localization of the ion channel in the lysosomal membrane and an optimal placement of the luciferase in the cytoplasm. We also developed a cell- and lysosome-permeable fluorescent BRET tracer that gives a BRET signal only when bound to the ion channel. This new protocol allows researchers worldwide to screen compounds that would interact with TRPML1 by using any plate reader with luminescent and fluorescence filters.

Immunohistochemistry (IHC) is a highly specific and widely used laboratory technique for assessing protein localization and expression in tissue samples. Interpretation of 3,3’ diamino benzidine (DAB)-based IHC is often based on observer-dependent manual scoring or traditional imaging software, which may show variability in DAB staining quantification. Furthermore, conventional image analysis tools often face limitations in precisely defining cell boundaries and quantifying membrane-specific signals. In this study, we present a standardized image analysis workflow using CellProfiler, an open-source software for image analysis for the quantification of membrane staining intensity in IHC images captured from slides prepared using formalin-fixed paraffin-embedded (FFPE) human cervical cancer tissue sections. The image analysis workflow was demonstrated using ASCT2 (SLC1A5), a membrane-localized amino acid transporter, as a representative biomarker for membrane-associated protein expression. This protocol involves image preprocessing, object identification, segmentation, and intensity measurement modules to distinguish cell membranes from cytoplasmic regions, enabling automated quantification of membrane intensity signals. The CellProfiler pipeline demonstrated improved accuracy in cell boundary identification and quantification of membrane-specific staining intensity. This is a rapid quantification process, since processing of each image only takes a few seconds; therefore, the analysis for 100 images can be performed within 10–15 min. This segmentation and quantification strategy is applicable to other membrane-based biomarkers after appropriate optimization of segmentation parameters. Following further minor modifications to the object identification modules, this pipeline can be used to detect and quantify cytoplasm- or nuclei-localized DAB-IHC markers across different tissue types. Overall, this protocol provides a standardized, user-friendly, and reproducible workflow for quantitative IHC image analysis that can be broadly applied to the study of protein biomarkers of different localizations, such as nuclei, cytoplasm, and cell membranes from different tissue types.

Cytoplasmic protein aggregation is a defining feature of multiple neurodegenerative diseases, including amyotrophic lateral sclerosis, frontotemporal dementia, Huntington’s disease, and certain forms of motor neuron disease. Recent evidence indicates that promyelocytic leukemia protein (PML) and engineered PML-derived variants can act as versatile aggregate-remodeling factors. In particular, cytoplasmically redirected PML variants recognize pathological cytoplasmic inclusions and promote their clearance. Here, we describe a protocol to generate and validate two engineered cytoplasmic PML variants: full-length mPML, which is redirected to the cytoplasm by disruption of its nuclear localization sequence, and the truncated mPML^ΔRBC variant, which lacks the RING, B-box, and coiled-coil domain but retains aggregate-reducing activity. The protocol integrates fluorescence-based imaging, bimolecular fluorescence complementation, detergent-soluble/insoluble fractionation, and validation in primary rat cortical neurons. This workflow provides a practical platform for assessing cytoplasmic aggregate burden and for comparing the aggregate-remodeling activities of PML-derived constructs. It can also be adapted to other disease-associated aggregation-prone proteins, including TDP-43, SOD1, FUS, tau, polyGA, and polyQ-expanded proteins.

Isogenic populations of Saccharomyces cerevisiae exhibit significant proliferative heterogeneity, with individual cells within a clonal culture displaying divergent growth rates and metabolic states. Investigating the origins of this variation requires a method to reconstruct the individual histories of cells within the population. This protocol describes a method for single-cell microencapsulation in alginate microspheres to create a physically stable, traceable, three-dimensional genealogical environment. By utilizing the alginate matrix to prevent daughter cell migration, the replicative history of a founder cell can be mathematically reconstructed. This is achieved by correlating the total cell count (N) within a developed microcolony with the total number of accumulated bud scars (n) visualized via confocal microscopy.

Amyotrophic lateral sclerosis (ALS) is characterized by early and spatially restricted pathology in motor axons, including distal degeneration and accumulation of aggregation-prone proteins such as TDP-43. However, a major limitation in the field has been the lack of approaches that enable robust, quantitative, and compartment-specific analysis of these early axonal events, particularly in human-relevant systems. Here, we describe an integrated experimental and analytical framework that enables quantitative dissection of axonal degeneration and protein aggregation, specifically within distal motor axons. By combining compartmentalized human co-cultures with a dedicated image analysis strategy, this approach enables selective and quantitative analysis of pathological processes specifically within axons, independent of surrounding tissues such as muscle and other cellular compartments. This framework captures both structural degeneration and protein aggregation dynamics at subcellular resolution, enabling spatially resolved quantitative analysis of disease-relevant changes along axons. Importantly, the analytical framework is not limited to TDP-43 but is broadly applicable to diverse aggregation-prone proteins, thereby providing a generalizable platform to study axonal pathology across neurodegenerative diseases. Together, this work provides a scalable approach for investigating axonal pathology as an early and measurable feature of neurodegeneration, with potential applications in mechanistic studies and therapeutic targeting in ALS and related disorders.