细胞生物学

Cellular stress induces profound changes in cytoskeletal organization and biomolecular condensate formation. Traditional approaches for monitoring cellular stress often require multi-component biosensors, endpoint staining procedures, or indirect biochemical measurements. Here, we describe a protocol for the use of OptoProfilin, a genetically encoded single-component optogenetic biosensor derived from Profilin-1 fused to Cryptochrome 2 (Cry2) and mCherry. Following transient expression in mammalian cells, OptoProfilin exhibits light-dependent localization to focal adhesions under non-stressed conditions and transitions to punctate condensates under energetic, oxidative, osmotic, and senescence-associated stress conditions. The protocol includes transient transfection, induction of cellular stress, live-cell imaging, immunofluorescence validation, and quantitative image analysis. While this protocol describes imaging on a Leica widefield fluorescence microscope, it can readily be extended to other microscopy platforms. As a stand-alone biosensor that produces visually distinct responses in stressed versus non-stressed cells, OptoProfilin provides a convenient platform for investigating stress-associated cytoskeletal remodeling and biomolecular condensate formation.

Extracellular vesicle (EV) proteomics can be used to study intercellular communication and find biomarkers of non-invasive diseases. Traditional separation methods (ultracentrifugation, size exclusion chromatography) and data-dependent acquisition (DDA) mass spectrometry usually have the drawbacks of copurification of pollutants, poor reproducibility, and insufficient sampling of low-abundance peptides. This protocol describes a workflow for label-free extracellular vesicle proteomics, which combines size exclusion chromatography for vesicle separation, data-independent acquisition (DIA) for deep discovery, and parallel reaction monitoring (PRM) for targeted verification. Plasma-derived extracellular vesicles are treated by standardized lysis, digestion, and LC-MS/MS procedures, so that the protein group of extracellular vesicles can be identified and quantified robustly. The main advantages of this scheme are that it can achieve high quantitative reproducibility, unbiased detection of low-intensity peptides, and seamless transition from discovery to targeted verification, while also being suitable for liquid biopsy samples and various cardiovascular diseases.

Multicellular organization relies on reciprocal interactions between molecular events, such as gene expression and protein state, and higher-scale properties, such as spatial patterning and tissue architecture. Understanding these processes requires methods that enable quantitative measurements at subcellular resolution, while maintaining the three-dimensional tissue organization. Conventional immunofluorescence imaging captures spatial information but is limited to the number of fluorescence markers that can be imaged simultaneously, whereas dissociation-based single-cell approaches can profile multimodal cellular states but lack positional information. Here, we describe 3D in toto iterative immunofluorescence imaging, termed 3D-4i, which enables up to ten-plex protein and protein state measurements in early zebrafish embryos. Leveraging sample immobilization on 96-well plates together with a gentle liquid handling system and high-content spinning disc confocal microscopy, this method comprises repeated rounds of antibody staining, optical clearing, confocal imaging, and antibody elution. Subsequent image analysis allows segmentation of nuclei and cells, extraction of quantitative single-cell features, and integration of molecular measurements with spatial context. Altogether, 3D-4i provides a scalable platform for investigating diverse biological processes in intact embryos, while maintaining both subcellular resolution and three-dimensional context.

Xenopus laevis oocytes are widely used as a heterologous expression system for investigating the function of membrane proteins due to robust expression of heterologous protein and a low endogenous transport background. Traditionally, transporter activity in oocytes has been assessed using electrophysiology or radiolabeled uptake assays, approaches that are constrained by the requirement for electrogenicity of the transport process, availability of radiolabeled compounds, and instrumentation. Here, we describe a fluorescence-based uptake assay that enables direct and rapid quantification of transporter activity using a fluorescence plate reader. The protocol uses the Arabidopsis thaliana sucrose transporter 1 (SUC1) and its fluorescent substrate esculin as a case for how to set up the assay. The workflow includes optimizing assay conditions, sample preparation, fluorescent measurements, and downstream data analysis using R. This method can readily be adapted to other transporter-substrate pairs, and it supports applications such as transporter inhibitor screening, mutational analysis, characterization of kinetic properties, or indirect substrate specificity testing through competition assays. Overall, this protocol provides a simple and scalable alternative to traditional techniques, eliminating the need for radiolabeled compounds or electrophysiology while enabling easy quantitative assessment of transporter activity.

Human induced pluripotent stem cell (hiPSC)-derived liver organoids have emerged as valuable models for studying human liver development. However, existing organoid systems often lack developmentally matched cell populations, particularly fetal hepatic stellate cells (HSCs), limiting their ability to recapitulate key developmental processes. Current approaches for generating HSCs rely on primary cells, immortalized cell lines, or hiPSC differentiation methods that frequently produce activated HSC-like cells and often require cell sorting. Here, we describe an efficient protocol for generating expandable fetal-like HSCs from hiPSCs through a stepwise differentiation strategy that mimics embryonic HSC development. The resulting cells can be robustly expanded while maintaining characteristic molecular and functional features of fetal HSCs. This protocol provides a reproducible and scalable source of fetal-like HSCs without cell sorting and supports the generation of multicellular liver organoids containing developmentally relevant stromal components. Beyond the validation of the protocol in studies of liver maturation and vascularization, it can be applied to investigations of HSC biology and congenital liver diseases.

Molecular tension sensors enable the visualization of forces acting on specific intracellular proteins in living cells. Most established genetically encoded sensors rely on Förster resonance energy transfer (FRET), requiring donor–acceptor imaging and correction for spectral bleed-through, which can complicate their application in heterogeneous tissues. We developed non-FRET α-actinin and α-catenin tension indicators containing a force-responsive tension sensor (TS) module composed of an optimized circularly permuted enhanced green fluorescent protein (cpEGFP) scaffold and an elastic (GPGGA)₈ linker. The TS module is incorporated into α-actinin or α-catenin, while a C-terminal mCherry serves as a force-insensitive reference for indicator abundance. This protocol describes indicator expression in cultured cells, validation using the myosin II inhibitor blebbistatin, two-color time-lapse imaging and live-cell super-resolution imaging using SRRF-Stream reconstruction based on super-resolution radial fluctuations (SRRF), and calculation of the green/red fluorescence ratio and normalized relaxation ratio. The protocol also specifies essential quality-control procedures, including imaging under nonsaturating conditions with fixed channel-specific acquisition settings across comparisons, registration of the green and red channels, and the use of appropriate vehicle and negative controls. For SRRF-Stream analysis, the green and red channels are reconstructed separately using identical reconstruction settings before ratio calculation. Representative reconstructed images should be compared with the corresponding conventional images to check for reconstruction artifacts. Optional procedures describe imaging of isolated cardiomyocytes and freshly isolated organs from tension-indicator mice. Because molecular loading is inferred from changes in TS fluorescence normalized to mCherry rather than from energy transfer between two fluorophores, the method avoids FRET-specific bleed-through correction; nevertheless, controlled acquisition and channel registration remain necessary. The indicators report relative changes in molecular loading and are particularly useful for resolving protein-specific and subcellular heterogeneity in tension dynamics.

Whole-mount 3D imaging of multilayered biological tissues enables quantitative analysis of cell states and organization in their spatial context. However, extracting unbiased and meaningful quantitative information from dense, multilayered samples remains challenging due to imaging artifacts, increased density, and limited signal-to-noise ratio. Open source bioimage analysis workflows tailored to this type of analysis are scarce, and analysis bottlenecks like image curation or cell segmentation are seldom available without coding expertise. Here, we present a step-by-step computational protocol for the analysis of dense 3D organoid datasets using the Tapenade (Thorough Analysis PipEliNe for Advanced DEep imaging) workflow. Starting from multichannel image stacks, the protocol guides users through software installation, registration and fusion of multi-view datasets, preprocessing, and nuclei segmentation. It further details the generation of quantitative outputs, including morphometric measurements, deformation fields, and spatial correlation analyses. The workflow can be executed through open-source Python scripts or user-friendly Napari interfaces, allowing interactive parameter tuning and 3D visualization at each stage. This pipeline provides an accessible and modular framework for nonspecialist users to perform reproducible, multiscale quantitative analysis of 3D organoid images, while retaining flexibility for advanced users to customize individual steps.

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.