Cell Biology

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.

Phosphatidylinositol 4,5-bisphosphate [PI(4,5)P₂] is a phospholipid enriched on the cytoplasmic leaflet of the plasma membrane, where it plays important roles in membrane trafficking and cytoskeletal dynamics through proteins that directly bind to it. PI(4,5)P₂ can be metabolized to other phosphorylated forms of phosphatidylinositol to regulate numerous processes such as cell growth and development. PI(4,5)P₂ can also be hydrolyzed to generate the second messengers diacylglycerol (DAG) and inositol triphosphate (IP₃). Altered metabolism or mislocalization of PI(4,5)P₂ can perturb one or more of its functions and contribute to disease states. Here, we present a protocol to visualize and quantify the localization of PI(4,5)P₂ in live cells. The protocol uses a highly specific PI(4,5)P₂ protein binding domain coupled to enhanced green fluorescence protein (PH-PLCD1-GFP), enabling localization and quantification of cytosol-facing PI(4,5)P₂ to be determined. Localization and quantification of the PH-PLCD1-GFP, PI(4,5)P₂ specific probe, is enabled by fluorescence imaging and confocal microscopy. This approach can be used to study the dynamics of PI(4,5)P₂ localization temporally in live cells under both physiological and pathological conditions.

Neurons and oligodendrocytes are the building blocks of the brain. Neurons form synaptic connections and transmit signals, while oligodendrocytes, including oligodendrocyte precursor cells (OPCs) and their derivatives, are vital for central nervous system maintenance and myelination. The demand for human-specific neuron-oligodendrocyte model systems to study these interactions has grown, yet co-culture protocols remain limited. Recent advancements in the field provide methods for deriving co-cultures of neurons and OPCs from human induced pluripotent stem cells (hiPSC), each with distinct benefits and challenges. This study presents a time-efficient, reproducible method to derive neurons and O4-expressing oligodendrocytes, followed by a straightforward co-culture system that minimizes astrocyte differentiation and ensures robust neuron and oligodendrocyte populations.

Calcium-permeable AMPA receptors (CP-AMPARs) and kainate receptors (CP-KARs) play crucial roles in synaptic plasticity and are implicated in various neurological processes. Current methods for identifying neurons expressing these receptors, such as electrophysiological recordings and immunostaining, have limitations in throughput or inability to distinguish functional receptors. This protocol describes a novel approach for the vital identification of neurons containing CP-AMPARs and CP-KARs using calcium imaging. The method involves loading neurons with Fura-2 AM, a calcium-sensitive fluorescent probe, KCl application to identify all neurons, and further addition of specific AMPAR agonists (e.g., 5-fluorowillardiine) in the presence of voltage-gated calcium channel blockers and NMDAR/KAR antagonists to identify CP-AMPAR-containing neurons. CP-KAR-containing neurons are identified using domoic acid applications in the presence and absence of NASPM (a CP-AMPAR antagonist). This technique offers several advantages over existing methods, including the ability to assess large neuronal populations simultaneously, distinguish between different receptor types, and provide functional information about CP-AMPAR and CP-KAR expression in living neurons, making it a valuable tool for studying synaptic plasticity and neurological disorders.

Protein carbonylation has been known as the major form of irreversible protein modifications and is also widely used as an indicator of oxidative stress in the biological environment. In the presence of oxidative stress, biological systems tend to produce large amounts of carbonyl moieties; these carbonyl groups do not have particular UV-Vis and fluorescence spectroscopic characteristics that we can differentiate, observe, and detect. Thus, their detection and quantification can only be performed using specific chemical probes. Commercially available fluorescent probes to detect specific carbonylation in biological systems have been used, but their chemical portfolio is still very limited. This protocol outlines the methods and procedures employed to synthesize a probe, (E,Z)-2-(2-(2-hydroxybenzylidene)hydrazonyl)-5-nitrophenol (2Hzin5NP), and assess its impact on carbonylation in human cells. The synthesis involves several steps, including the preparation of its hydrazone compounds mimicking cell carbonyls, 2-Hydrazinyl 5-nitrophenol, (E,Z)-2-(2-ethylidenehydrazonyl)-5-nitrophenol, and the final product (E,Z)-2-(2-(2-hydroxybenzylidene)hydrazonyl)-5-nitrophenol. The evaluation of fluorescence quantum yield and subsequent cell culture experiments are detailed for the investigation of 2Hzin5NP effects on cell proliferation and carbonylation.