细胞生物学

Membrane transporters mediate the selective movement of ions and molecules across biological membranes and are essential for cellular homeostasis. However, their functional characterization in living cells is often complicated by the complexity of the native membrane environment. Reconstitution into model membrane systems provides a powerful alternative by enabling precise control over lipid composition and experimental conditions. Giant unilamellar vesicles (GUVs) are particularly well suited for transporter studies, as their cell-sized dimensions allow direct microscopic observation and fluorescence-based measurements of protein activity. Here, we describe a two-step reconstitution protocol in which transport proteins are first incorporated into large unilamellar vesicles and then used to generate protein-containing giant unilamellar vesicles (proteo-GUVs) via the poly(vinyl alcohol) swelling method. This two-step approach enhances protein incorporation efficiency and preserves transporter functionality. The method is exemplified using the P3-type ATPase Arabidopsis thaliana plasma membrane H⁺-ATPase isoform 2 (AHA2). We further describe a fluorescence-based assay to assess proton transport activity in proteo-GUVs. Our approach provides a versatile and controlled platform for biochemical, biophysical, and single-molecule analysis of membrane transporters.

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.

Stochastic optical reconstruction microscopy (STORM) is a single-molecule localization microscopy technique that enables visualization of cellular structures beyond the diffraction limit. This approach has revealed previously inaccessible ultrastructural details in a wide range of cellular components, including the actin cytoskeleton, clathrin-coated pits, mitochondria, and bacterial nucleoid-associated proteins. STORM relies on the sequential emission of single photons from photosensitive fluorophores, which are precisely localized before entering a dark state or undergoing photobleaching. By activating fluorophores individually and fitting their point spread functions (PSFs), the center of mass can be calculated with a localization precision of up to ~20 nm. The parallel detection of thousands of single-molecule events, each assigned to distinct spatial coordinates, enables the reconstruction of a high-resolution image. Here, we describe a simple and efficient STORM workflow—including sample preparation, image acquisition, and quality control measurements—that we used to visualize various subcellular structures, such as mitochondria, microtubules, and lysosomes labeled with the commonly employed cyanine dye Alexa Fluor 647, as well as the actin cytoskeleton stained with Alexa Fluor 488–conjugated phalloidin. Image acquisition was performed using a conventional epifluorescence/total internal reflection (TIRF) microscope adapted for STORM imaging. Key adaptations included the use of a 160×/1.43 NA oil-immersion objective and a high-power mode, which concentrates the laser beam onto a small region of the sample, ensuring sufficient light intensity to drive fluorophores into the dark state. In addition, implementing a 1.6× magnification lens and a 4×4 binning camera mode allowed us to achieve a 100-nm pixel size optimal for reliable molecule detection. We believe that this protocol will be highly valuable to the microscopy community, as it lowers technical barriers to performing STORM on widely available microscopy platforms, thereby facilitating broader implementation of this powerful super-resolution technique.

Super-resolution imaging of synapses in intact brain tissue remains challenging because light scattering, photobleaching, and limited probe penetration, along with antigen accessibility within the densely packed postsynaptic densities (PSDs), constrain resolution and labeling efficiency. Here, we present a protocol utilizing thin brain cryosections and tau-stimulated emission depletion (STED) nanoscopy to visualize the intricate nano-architecture of excitatory synapses in situ. Slicing the brain into 6 μm sections allows for highly efficient and even penetration of probes throughout sections while ensuring that the resolution is not significantly impacted by the imaging depth of the tissue. We outline step-by-step instructions for labeling pre- and postsynaptic nano-architecture using antibodies and nanobodies, highlighting how fixative choice influences the labeling efficiency of synaptic proteins. While this protocol is compatible with both confocal and super-resolution imaging, when combined with rapid image acquisition times of tau-STED, it enables clear separation of key synaptic features in three dimensions with minimal photobleaching. Thus, this approach enables robust multiplex imaging of fluorescently labeled synaptic proteins in the brain, providing exceptional spatial resolution for visualization and quantification of synaptic nanoarchitecture in its native environment.

Cellulose synthase complexes (CSCs) play a central role in plant cell wall formation. Their dynamic behavior at the plasma membrane leads to the deposition of cellulose microfibrils into the apoplastic space, thereby shaping the architecture and mechanical properties of the cell wall. Although previous imaging studies have provided important insights into CSC dynamics and localization, standardized and reproducible workflows for quantitative measurements of CSC speed and density remain limited. Here, we present a reproducible live-cell imaging and analysis workflow for quantifying the speed and density of fluorescently labeled CSCs at the plasma membrane in Arabidopsis thaliana. The protocol integrates optimized spinning-disk confocal imaging, surface-based projection of z-stack recordings, automated detection of diffraction-limited CSCs foci, and kymograph-based speed measurements using freely available tools in Fiji. While selected steps, such as region of interest definition and parameter selection for spot detection or trajectory analysis, remain user-guided, these decisions are constrained to well-defined stages within an otherwise standardized pipeline, thereby reducing variability and improving reproducibility across experiments. The workflow has been validated across multiple tissues, reporter lines, genetic backgrounds, and perturbation conditions in Arabidopsis and enables robust comparative analysis of CSC dynamics. Beyond CSCs, this workflow is expected to be adaptable to other fluorescently labeled proteins that appear as diffraction-limited foci at or near the plasma membrane.

Centrosomes are dynamic organelles critical for mitotic spindle assembly and cilia formation. Here, I describe a protocol for quantifying relative centrosomal protein abundance in Drosophila melanogaster embryos using radial profile analysis of fluorescence intensity. The method involves embryo collection, manual dechorionation, mounting for live imaging, confocal microscopy, and subsequent image analysis. Radial profiling allows quantification of relative protein abundance together with its spatial distribution at the centrosome, providing either relative or normalized intensity profiles. I then outline how this approach can be integrated with complementary techniques such as fluorescence recovery after photobleaching (FRAP) and super-resolution imaging, in this case, three-dimensional structured illumination microscopy (3D-SIM). Combining radial fluorescence profiling with these imaging modalities enables high-resolution, quantitative analysis of dynamic centrosome assembly in a genetically tractable system.

Time-lapse into immunofluorescence (TL into IF) imaging combines the wealth of information acquired during live-cell imaging with ease of access for static immunofluorescence markers. In the field of mechanobiology, connecting live and static imaging to visualize cell biology dynamics is often troublesome. For instance, nuclear blebs are deformations of the nucleus that often rupture spontaneously, leading to changes in the molecular composition of the nucleus and the nuclear bleb. Current techniques to connect cellular dynamics and their downstream effects via live-cell imaging, followed by immunofluorescence, often require third-party analysis programs or stage position measurements to accurately track cells. This protocol simplifies the connection between live and static imaging by utilizing a gridded imaging dish. In our protocol, cells are plated on a dish with an engraved coordinate plane. Individual cells are then matched from when the time-lapse ends to the immunofluorescence images simply by their known coordinate location. Overall, TL into IF offers a straightforward method for connecting dynamic live-cell with static immunofluorescence imaging, in an easy and accessible tool for cell biologists.

The deep learning revolution has accelerated discovery in cell biology by allowing researchers to outsource their microscopy analyses to a new class of tools called cell segmentation models. The performance of these models, however, is often constrained by the limited availability of annotated data for them to train on. This limitation is a consequence of the time cost associated with annotating training data by hand. To address this bottleneck, we developed Cell-APP (cellular annotation and perception pipeline), a tool that automates the annotation of high-quality training data for transmitted-light (TL) cell segmentation. Cell-APP uses two inputs—paired TL and fluorescence images—and operates in two main steps. First, it extracts each cell’s location from the fluorescence images. Then, it provides these locations to the promptable deep learning model μSAM, which generates cell masks in the TL images. Users may also employ Cell-APP to classify each annotated cell; in this case, Cell-APP extracts user-specified, single-cell features from the fluorescence images, which can then be used for unsupervised classification. These annotations and optional classifications comprise training data for cell segmentation model development. Here, we provide a step-by-step protocol for using Cell-APP to annotate training data and train custom cell segmentation models. This protocol has been used to train deep learning models that simultaneously segment and assign cell-cycle labels to HeLa, U2OS, HT1080, and RPE-1 cells.

Repetitive increases of intracellular calcium ions (Ca²⁺ oscillations) control cellular functions in various biological events, including meiotic resumption after fertilization. Sperm-derived substances enter the cytoplasm of mature oocytes by sperm fusion, causing Ca²⁺ oscillations. Sperm-independent Ca²⁺ oscillations are also induced in immature oocytes isolated from the ovaries of neonatal to adult mice. The presence of Ca²⁺ oscillations may contribute to subsequent oocyte quality; however, its physiological role and molecular mechanism are unclear. Here, we describe a method of collecting immature oocytes from the ovaries of juvenile (12, 15, and 21 days after birth) and adult mice and monitoring their Ca²⁺ oscillations. Since mouse oocytes are larger than other types of cells, they are a useful model for studying spatiotemporal patterns and the mechanism of Ca²⁺ oscillations in various types of cells. This method can be applied to other rodents due to similarities in oocyte size and developmental processes. Furthermore, the use of various fluorescent probes enables visualization of organelle rearrangement. The mechanism of interaction between oocytes and somatic cells differs between juvenile and adult mice. Therefore, two distinct methods are employed for oocyte collection.