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.

When plants undergo senescence or experience carbon starvation, leaf cells degrade proteins in the chloroplasts on a massive scale via autophagy, an evolutionarily conserved process in which intracellular components are transported to the vacuole for degradation to facilitate nutrient recycling. Nonetheless, how portions of chloroplasts are released from the main chloroplast body and mobilized to the vacuole remains unclear. Here, we developed a method to observe the autophagic transport of chloroplast proteins in real time using confocal laser-scanning microscopy on transgenic plants expressing fluorescently labeled chloroplast components and autophagy-associated membranes. This protocol enabled us to track changes in chloroplast morphology during chloroplast-targeted autophagy on a timescale of seconds, and it could be adapted to monitor the dynamics of other intracellular processes in plant leaves.

Cellular phenomena such as signal integration and transmission are based on the correct spatiotemporal organization of biomolecules within the cell. Therefore, the targeted manipulation of such processes requires tools that can precisely induce the localizations and interactions of the key players relevant to these processes with high temporal resolution. Chemically induced dimerization (CID) techniques offer a powerful means to manipulate protein function with high temporal resolution and subcellular specificity, enabling direct control over cellular behavior. Here, we present the detailed synthesis and application of dual SLIPT and dual SLIPT^NVOC, which expand the SLIPT (self-localizing ligand-induced protein translocation) platform by incorporating a dual-ligand CID system. Dual SLIPT and dual SLIPT^NVOC independently sort into the inner leaflet of the plasma membrane via a lipid-like anchoring motif, where they present the two headgroup moieties trimethoprim (TMP) and HaloTag ligand (HTL), which recruit and dimerize any two ^iK6eDHFR- and HOB-tagged proteins of interest (POIs). Dual-SLIPT^NVOC furthermore enables this protein dimerization of POIs at the inner leaflet of the plasma membrane in a pre-determined order and light-controlled manner. In this protocol, we detail the synthetic strategy to access dual SLIPT and dual SLIPT^NVOC, while also providing the underlying rationale for key design and synthetic decisions, with the aim of offering a streamlined, accessible, and broadly implementable methodology. In addition to the detailed synthesis, we present representative applications and typical experimental outcomes and recommend strategies for data analysis to support effective use of the system. Notably, dual SLIPT and dual SLIPT^NVOC represent the first CID systems to emulate endogenous lipidation-driven membrane targeting, while retaining hallmark advantages of CID technology—the precision over POI identity, recruitment sequence, high spatiotemporal control, and “plug-and-play” flexibility.

Most membrane and secreted proteins are transported from the endoplasmic reticulum (ER) to the Golgi apparatus and subsequently directed to their final destinations in the cell. However, the mechanisms underlying transport and cargo sorting remain unclear. Recent advancements in optical microscopy, combined with synchronized cargo protein release methods, have enabled the direct observation of cargo protein transport. We developed a new optically synchronized cargo release method called retention using the dark state of LOV2 (RudLOV). This innovative technique offers three exceptional control capabilities: spatial, temporal, and quantitative control of cargo release. RudLOV uses illumination to trigger transport and detect cargo. Consequently, the selection of an appropriate laser and filter set for controlling the illumination and/or detection is crucial. The protocol presented here provides step-by-step guidelines for obtaining high-resolution live imaging data using RudLOV, thereby enabling researchers to investigate intracellular cargo transport with unprecedented precision and control.

Translation is a key step in decoding the genetic information stored in DNA. Regulation of translation is an important step in gene expression control and is essential for healthy organismal development and behavior. Despite the importance of translation regulation, its impact and dynamics remain only partially understood. One reason is the lack of methods that enable the real-time visualization of translation in the context of multicellular organisms. To overcome this critical gap, microscopy-based methods that allow visualization of translation on single mRNAs in living cells and animals have been developed. A powerful approach is the SunTag system, which enables real-time imaging of nascent peptide synthesis with high spatial and temporal resolution. This protocol describes the implementation and use of the SunTag translation imaging system in the small round worm Caenorhabditis elegans. The protocol provides details on how to design, carry out, and interpret experiments to image translation dynamics of an mRNA of interest in a cell type of choice of living C. elegans. The ability to image translation live enables better understanding of translation and reveals the mechanisms underlying the dynamics of cell type–specific and subcellular localization of translation in development.

Rapid and uniform labeling of plasma membrane proteins is essential for high-resolution imaging of dynamic membrane topologies and intercellular communication in live mammalian cells. Existing strategies for labeling live cell membranes, such as fluorescent fusion proteins, enzyme-mediated tags, metabolic bioorthogonal labeling, and lipophilic dyes, face trade-offs in the requirement of genetic manipulation, the presence of non-uniform labeling, the need for extensive preparation times, and limited choices of fluorophores. Here, we present a streamlined protocol that leverages N-hydroxysuccinimide (NHS)-ester chemistry to achieve rapid (≤5 min), covalent conjugation of synthetic small-molecule dyes to surface-exposed primary amines, enabling pan-membrane-protein labeling. This workflow covers dye stock preparation, labeling for suspension and adherent cells, multiplex live-cell imaging, fusion protein co-staining (including insulin-triggered receptor endocytosis), 3D membrane visualization, and in vivo assays for visualizing membrane-derived material transfers between donor and recipient cells using a lymphoma T-cell mouse model. This high-density labeling approach is compatible with various cell types across diverse imaging platforms. Its speed, versatility, and stability make it a broadly applicable tool for studying plasma membrane dynamics and intercellular membrane trafficking.

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.

Cell–surface and cell–cell interaction assays are fundamental for studying receptor–ligand interactions and characterizing cellular responses and functions. They play a critical role in diagnostics and in modulating immune system activity for therapeutic applications, notably in cancer immunotherapy. By providing time-lapsed and cell-level direct observation of the sample, optical microscopy offers strong advantages compared to current go-to techniques, which are typically either ensemble methods (e.g., measuring cell populations) or indirect readouts (e.g., impedance for adherent cells). This protocol describes two complementary microscopy-based assays: (1) a cell–surface ligand binding assay to quantify dynamic interactions between human primary Natural Killer (NK) cells and a cancer-mimicking surface, and (2) a cell–cell interaction assay to evaluate antibody-dependent cell cytotoxicity (ADCC) mediated by NK cells targeting tumor cells. Additionally, the protocol uses Celldetective, a new open graphical user interface for quantitative analysis of cell interaction dynamics from 2D time-lapse microscopy datasets. Although applied here to primary immune cells, these methods are adaptable to various cell types, including other immune cells, fibroblasts, and cancer cells. This approach enables direct observation and quantification of cellular morphology, motility, cell–cell interactions, and dynamic behaviors at single-cell resolution over time, facilitating detailed analysis of mechanisms such as cell death, migration, and immune synapse formation.