Cell Biology

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.

Phosphatase and tensin homolog-induced kinase 1 (PINK1) is a serine/threonine kinase that plays a key role in mitophagy initiation. Loss-of-function autosomal recessive mutations in PINK1 cause early onset Parkinson’s disease (EOPD). Current approaches for studying PINK1 function depend on bulk techniques that can only provide snapshots of activity and could miss the dynamics and cell-to-cell heterogeneity of PINK1 activity or provide an indirect readout of PINK1 activity. Here, we present a protocol using our newly developed phase separation–based PINK1 biosensor (PINK1-SPARK) to observe real-time activity of endogenous PINK1 in single cells. Following transfection of live cells with PINK1-SPARK, cells are treated with mitochondrial depolarizing agents and visualized using widefield or confocal fluorescence microscopy, either following the same cells over time for time-lapse imaging of PINK1 activity or end-point measurements. Thus, PINK1-SPARK is a new tool that enables the measurement of PINK1 activity in single live cells, allowing for further elucidation of the role of PINK1 in mitophagy and cell function.

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.

Thin membrane protrusions in cells help them communicate, create traction forces during their movement, and coordinate complex development in multicellular organisms. These structures include cytonemes, tunneling nanotubes, and microtubule-based nanotubes (MT-nanotubes), each with a different cytoskeletal constitution and function. Actin-based cytonemes help deliver signaling molecules, while microtubule-based nanotubes assist with transporting vesicles and organelles. Despite their physiological role, we still do not fully understand how these thin membrane protrusions form and function. In this study, we introduce an improved live-cell imaging method to observe polar cell protrusions during micropyle morphogenesis in developing Drosophila eggs. This technique combines precise developmental staging, careful dissection, and optimized ex vivo culture conditions to maintain tissue health during extended imaging. We also fine-tuned the imaging settings to reduce phototoxicity and thermal stress. This allows for continuous, high-resolution tracking of protrusion dynamics in real time. Our protocol addresses major drawbacks of fixed-tissue methods by capturing the entire process of protrusion formation, extension, and remodeling in intact living tissue. Additionally, it works well with drugs, making it a useful tool for functional studies. Overall, this approach builds a strong foundation for exploring membrane protrusion biology. It can also be applied to investigate similar developmental processes in other systems, aiding our understanding of normal development and diseases.

In vivo imaging of brown algal cells in 3D is extremely challenging because of the presence of pigments, such as fucoxanthin and chlorophyll, that diffract light. Moreover, brown algae live in seawater, a high ionic environment that can change the fluorochrome behavior or cause aggregates. Despite the importance of in vivo monitoring the developmental process of brown algal tissues, 4D imaging (x, y, z, t) on a conventional fluorescence microscope is limited. Here, we propose a detailed protocol using a new orange-emitting fluorochrome, styryl benzoindoleninium sulfonate (SBIS), suitable for labeling the plasma membrane of brown algal cells and multicolor in vivo imaging in 3D using confocal and light sheet microscopy. Unlike calcofluor white (CFW), SBIS enables the observation of brown algal cells at thicknesses up to 25 μm and over periods up to 7 days on brown algae such as Ectocarpus sp., Sphacelaria rigidula, and Saccharina latissima. This step-by-step protocol includes labeling of brown algal tissues, mounting for 3D confocal time-lapse microscopy, and mounting for 3D time-lapse light sheet microscopy. The imaging setup and parameters have been optimized for minimizing toxicity for brown algal tissues, improving signal-to-noise ratio, and enabling detailed visualization of cell shape. Therefore, this protocol provides robust and multiplexed imaging with 4D visualization of brown algal cell shape throughout the brown algae growth, offering broad applications to brown algae study at the cellular level.

RNA imaging techniques enable researchers to monitor RNA localization, dynamics, and regulation in live or fixed cells. While the MS2-MCP system—comprising the MS2 RNA hairpin and its binding partner, the MS2 coat protein (MCP)—remains the most widely used approach, it relies on a tag containing multiple fluorescent proteins and has several limitations, including the potential to perturb RNA function due to the tag’s large mass. Alternative methods using small-molecule binding aptamers have been developed to address these challenges. This protocol describes the synthesis and characterization of RNA-targeting probes incorporating a peptide nucleic acid (PNA)-based linker within the cobalamin (Cbl)-based probe of the Riboglow platform. Characterization in vitro involves a fluorescence turn-on assay to determine binding affinity (K_D) and selective 2′-hydroxyl acylation analyzed by primer extension (SHAPE) footprinting analysis to assess RNA-probe interactions at a single nucleotide resolution. To show the advancement of PNA probes in live cells, we present a detailed approach to perform both stress granule (SG) and U-body assays. By combining sequence-specific hybridization with structure-based recognition, our approach enhances probe affinity and specificity while minimizing disruption to native RNA behavior, offering a robust alternative to protein-based RNA imaging systems.

RNA is now recognized as a highly diverse and dynamic class of molecules whose localization, processing, and turnover are central to cell function and disease. Live-cell RNA imaging is therefore essential for linking RNA behavior to mechanism. Existing approaches include quenched hybridization probes that directly target endogenous transcripts but face delivery and sequestration issues, protein-recruitment tags such as MS2/PP7 that add large payloads and can perturb localization or decay, and CRISPR–dCas13 imaging that requires substantial protein cargo and careful control of background and off-target effects. Here, we present a protocol for live-cell RNA imaging using the Spinach^TM family of fluorogenic RNA aptamers. The method details the design and cloning of Spinach^TM-tagged RNA constructs, selection and handling of cognate small-molecule fluorophores, expression in mammalian cell lines, dye loading, and image acquisition on standard fluorescence microscopes, followed by quantitative analysis of localization and dynamics. We include controls to verify aptamer expression and signal specificity, guidance for multiplexing with related variants (e.g., Broccoli, Corn, Squash, Beetroot), and troubleshooting for dye permeability and signal optimization. Application examples illustrate use in tracking cellular delivery of mRNA therapeutics, monitoring transcription and decay in response to perturbations, and the forming of toxic RNA aggregates. Compared with prior methods, Spinach^TM tags are compact, genetically encodable, and fluorogenic, providing high-contrast imaging in both the nucleus and cytoplasm with single-vector simplicity and multiplexing capability. The protocol standardizes key steps to improve robustness and reproducibility across cell types and laboratories.

Larval zebrafish are often mounted laterally to ensure consistent anatomical positioning and to standardize imaging of body axes across early development. However, this conventional approach often tethers sample orientation to a single microscope configuration and limits optical accessibility. We present a mounting protocol for larval zebrafish that enables optical access from both dorsal and ventral orientations while preserving lateral sample position. This approach uses common laboratory consumables to establish a mounting platform that eliminates any need to remount samples between the use of upright and inverted microscopes. By establishing a hydrophobic seal, mounted embryos can be inverted with ease to access the sample from either orientation. A seamless transition here facilitates reliable identification and longitudinal tracking of the same biological region of interest across microscope configurations. This protocol is broadly applicable to live imaging experiments requiring flexibility in imaging geometry, minimal sample handling, and high reproducibility.

Calcium ions serve as a universal secondary messenger, integrating diverse external signals, such as light, herbivory, and mechanical stimuli, within plant cells. However, the visualization and mechanistic dissection of calcium signaling specifically in response to mechanical stimulation remain technically challenging and underexplored in most plants. Previous studies have been largely confined to a few model systems, including Arabidopsis; here, we introduce a live-cell imaging approach using the stigmas of Torenia fournieri. This in vitro system enables multiscale observation of calcium signal patterns following controlled mechanical stimulation. This versatile platform not only simplifies the design of calcium imaging assays but also provides a tractable system for functionally validating other key molecular components in this signaling pathway.

Metastasis is initiated by cell invasion of the basement membrane, facilitating cell migration and colonization at a secondary tumor site. Cancer cells remodel the cytoskeleton to form ventral protrusions, termed invadopodia, that traffic and deliver matrix metalloproteases to degrade the extracellular matrix. Traditional efforts have utilized immunolabeling to measure protein localization within invadopodia, an approach limited by reduced temporal resolution, logistical challenges in orienting invadopodia within the focal plane of the objective lens, and impaired ability to reconstitute physiological conditions. Here, we describe a protocol for constructing and utilizing the axial invasion chamber (AIC) to perform live-cell 3D visualization of mature elongating invadopodia under physiological conditions. The AIC is simple to build, using standard 35 mm glass-bottom dishes that suit most microscope stage holders. A polyester membrane is used to uniformly orient and promote invadopodia formation and restrict cell migration. The AIC extracellular matrix is composed of readily available reagents that have been optimized to facilitate cell adhesion and invadopodia maturation. Critical advances of the AIC include imaging and measurements of protein localization without immunolabeling, imaging of live cell invadopodia using conventional inverted microscopes, and production of a fully operational apparatus within 28 h from initial assembly. While the protocol has been used for live-cell invadopodia protein localization and structure, it provides an opportunity to interchange components of the polyester membrane and/or the extracellular matrix to optimize the device for a variety of different cell types and cell invasion studies.