Cell Biology

Calcium signaling is a universal, versatile process in which ionized or free calcium (Ca²⁺) acts as a second messenger to regulate various cellular activities, including hormone secretion, contraction, proliferation, gene expression, and apoptosis. Changes in the cytoplasmic free Ca²⁺ concentration ([Ca²⁺]_cyt) in hepatocytes play a central role in mediating the actions of insulin, glucagon, catecholamines, and other hormones on carbohydrate, lipid, and protein metabolism in the liver. Ratiometric chemical Ca²⁺ indicators are fluorescent dyes that change their emission or excitation spectrum upon binding to calcium, allowing for precise, quantitative measurements of changes in the intracellular Ca²⁺ concentration. They enable calibration by calculating the ratio of two fluorescence intensities, correcting for artifacts such as uneven dye loading, photobleaching, and cell volume variations. Fura-2 acetoxymethyl ester (AM) (hereinafter referred to as Fura-2), a ratiometric and sensitive indicator dye, is a popular fluorescent Ca²⁺ reporter for measuring intracellular calcium. Here, we describe a comprehensive and detailed protocol for Ca²⁺ imaging of the H4IIE cell line and primary rodent hepatocytes in vitro via the chemical reporter Fura-2, which can also be employed on a wide variety of cell types. Unlike previously published protocols, this protocol addresses the challenge of facilitating the attachment of liver cell lines and primary hepatocytes to glass coverslips for imaging using an inverted fluorescence microscope. Our protocol describes two different loading/labeling strategies for Fura-2 dye: one is cost-effective but requires skillful pipettor handling, and the second one is easy but expensive as it needs a large volume of Krebs-Ringer HEPES (KRH)-Fura-2 solution. If the coverslips are handled properly, the cost-effective coverslip-only loading approach produces similar quality results as the large volume method. Finally, we describe a simple and user-friendly procedure to analyze Ca²⁺ signals over time using Microsoft Excel’s functional equations.

Ribonucleoprotein (RNP) condensates are membraneless organelles that exist alongside many RNA-driven processes, such as transcription and splicing. Despite their ubiquity, the biological necessity of forming a condensed phase remains unclear, particularly because the same RNP components exist both within these organelles and in the surrounding dilute phase. Most current methods for studying biochemical interaction dynamics within condensates rely on in vitro reconstitution of minimal factors or low-throughput single-molecule studies. However, RNP condensates are complex organelles containing tens to hundreds of proteins and hundreds to thousands of different RNAs. Here, we describe a scalable, high-throughput fluorescence microscopy–based approach to analyze protein–protein interaction networks, allowing for the rigorous assessment of dynamic, process-critical interactions within RNP condensates from live cells. This method takes advantage of fluorescence lifetime imaging (FLIM) and phasor plot analysis to automate segmentation of condensate-localized fluorescence signals. Using suitable FLIM–Förster resonant energy transfer (FLIM-FRET) fluorescent pairs fused to proteins of interest, protein–protein interactions can be actively monitored throughout various conditions via changes in fluorescence lifetime. Results from this assay yield valuable insight into the organization and assembly of essential factors for different condensate-associated processes to infer the functional consequences of RNP granule partitioning. Although this protocol is tailored for studying protein interactions within condensates, the design and execution framework can be adapted to investigate protein–protein interactions across a wide variety of compartments within different biological systems.

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.

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.

The study of whole organs or tissues and their cellular components and structures has been historically limited by their natural opacity, which is caused by the optical heterogeneity of the tissue components that scatter light as it traverses through the tissue, making 3D tissue imaging highly challenging. In recent years, tissue clearing techniques have received widespread attention and undergone rapid development. We recently demonstrated the synthesis of a 2-hydroxyethyl methacrylate (HEMA)-acrylamide (AAm) copolymer. This was achieved using antipyrine (ATP) and 2,2′-thiodiethanol (TDE) as solvents. The resulting solution rapidly embedded tissue samples with a high degree of transparency and is compatible with multiple fluorescence labeling techniques. The method exhibits significant transparency effects across a range of organs, comprising the heart, liver, spleen, lung, kidney, brain (whole and sectioned), esophagus, and small intestine. It can enable volumetric imaging of tissue up to the scale of mouse organs, decrease the duration of the clearing, and preserve emission from fluorescent proteins and dyes. To facilitate the use of this powerful tool, we have provided here a detailed step-by-step protocol that should allow any laboratory to use tissue transparency technology to achieve transparency of tissues and organs.

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

Here, we present a protocol for implementing the fluorogen-activating protein FAST (fluorescence-activating and absorption-shifting tag) in fluorescence lifetime imaging microscopy (FLIM), which allows separating fluorescent species in the same spectral channel based on fluorescence lifetime properties. Previous studies have demonstrated FLIM multiplexing using various combinations of synthetic probes, fluorescent proteins, or self-labeling tags. In this protocol, we utilize engineered FAST point mutation variants that bind fluorogen HBR-2,5-DM. The designed probes possess nearly identical, compact protein sizes (14 kDa), and the resulting protein–fluorogen complexes demonstrate comparable steady-state optical properties and exhibit distinct fluorescence lifetimes, displaying monoexponential fluorescence decay kinetics. When FAST variants are expressed with localization signals, these properties facilitate robust signal separation in regions with co-localized or spatially overlapping labels (nucleus and cytoskeleton in this protocol) in live mammalian cells. This method can be applied to separate other overlapping cellular compartments, such as the nucleus and Golgi apparatus, or mitochondria and cytoskeleton.

High-content analysis (HCA) is a powerful image-based approach for phenotypic profiling and drug discovery, enabling the extraction of multiparametric data from individual cells. Traditional HCA protocols often rely on fixed-cell imaging, with assays like cell painting widely adopted as standard. While these methods provide rich morphological information, the integration of live-cell imaging expands analytical capabilities by enabling the study of dynamic biological processes and real-time cellular responses. This protocol presents a simple, cost-effective, and scalable method for live-cell HCA using acridine orange (AO), a metachromatic fluorescent dye that highlights cellular organization by staining nucleic acids and acidic compartments. The assay provides visualization of distinct subcellular structures, including nuclei and cytoplasmic organelles, using a two-channel fluorescence readout. Compatible with high-throughput microscopy and computational analysis, the method supports diverse applications such as phenotypic screening, cytotoxicity assessment, and morphological profiling. By preserving cell viability and enabling dynamic, real-time measurements, this live-cell imaging approach complements existing fixed-cell assays and offers a versatile platform for uncovering complex cellular phenotypes.

Counting protein molecules helps reveal the organization of components within cellular structures and the stoichiometries of protein complexes. Existing protein and peptide quantitation methods vary in their complexity. Here, we report a straightforward workflow to measure the absolute number of HaloTag-labeled myosin 10 (Myo10) molecules in U2OS cells. Myo10 is a motor protein that plays a prominent role in cellular protrusion formation. Various biochemical and biological properties of Myo10 are established, but it is not well-defined how many molecules of Myo10 pack into narrow cellular structures called filopodia. We present a workflow for using SDS-PAGE to calibrate Myo10 signal with a reference protein, segmenting epifluorescence microscopy images to map Myo10 intracellular distribution, and interpreting the results to derive biological and functional insights. Our protocol is simple to employ and not only applicable for Myo10 research but also easily adaptable for other biological systems that use HaloTag.

Three-dimensional cell models, such as spheroids, represent a more physiological arrangement in which cells can grow, allowing them to develop cell–cell interactions in all dimensions. The most common methods for growing spheroids are scaffold-based, typically using either extracellular matrix or hydrogels as a physical support for the cellular assembly. One key problem with this approach is that the spheroids that are produced can be highly variable in size and shape. The protocol presented here allows for the systematic production of uniform spheroids in a short time frame by utilising a micropatterned plate. We show that spheroids can be used to investigate fundamental research questions, such as how the endomembrane system is organised in cells. Our protocol can be used in a manual or automated manner, potentially allowing scaling up for screening applications. Furthermore, without the complication of removing the spheroids from the extracellular matrix or hydrogel, as would be required in scaffold-based systems, spheroids can easily be used in other downstream applications.