Cell Biology

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.

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.

Neuronal survival in vitro is usually used as a parameter to assess the effect of drug treatments or genetic manipulation in a disease condition. Easy and inexpensive protocols based on neuronal metabolism, such as 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT), provide a global view of protective or toxic effects but do not allow for the monitoring of cell survival at the single neuronal level over time. By utilizing live imaging microscopy with a high-throughput microscope, we monitored transduced primary cortical neurons from 7–21 days in vitro (DIV) at the single neuronal level. We established a semi-automated analysis pipeline that incorporates data stratification to minimize the misleading impact of neuronal trophic effects due to plating variability; here, we provide all the necessary commands to reproduce it.

Plasma membrane–associated condensates driven by liquid–liquid phase separation represent a novel mechanism of receptor-mediated signaling transduction, serving as mesoscale platforms that concentrate signaling molecules and modulate reaction kinetics. Condensate formation is a highly dynamic process that occurs within seconds to minutes following receptor activation. Here, we present methods for de novo reconstituting liquid-like condensates on supported lipid bilayers and assessing the condensate fluidity using fluorescence recovery after photobleaching (FRAP). This protocol encompasses supported lipid bilayer preparation, condensation imaging, and FRAP analysis using total internal reflection fluorescence (TIRF) microscopy. Supported lipid bilayers provide a membrane-mimicking environment for receptor signaling cascades, offering mechanistic insights into protein–protein and lipid–protein interactions amid micron-scale condensates. The protocol can also be adapted to study condensates associated with the internal membranes of the Golgi apparatus, mitochondria, and other organelles.

Flagellate stages of green microalgae such as Trebouxia are only partially characterised, with recent evidence suggesting that they are involved in both sexual and asexual reproduction. Conventional methods based on fixed samples in light, confocal, or electron microscopy provide only static observations and prevent real-time monitoring of living cells. To overcome this limitation, we have developed a simple and cost-effective protocol for observing Trebouxia flagellate cells over several days by coating microscopy slides with Bold’s basal medium. The method preserves cell viability and allows repeated imaging of motile cells in the same areas so that their behaviour and development can be continuously observed. In this way, qualitative observations, such as flagellate cell release, motility, and gamete fusion, can be combined with quantitative analyses of cell morphology. The protocol has proven to be robust and reproducible and was applied to several Trebouxia species. Compared to existing techniques, it allows the monitoring of dynamic processes and provides a powerful tool to study specific life stages not only in Trebouxia but also in other unicellular and colonial green algae.

Microbial life cycles are often reconstructed theoretically from fragmentary pieces of evidence. Protocols for the direct and continuous observation of entire microbial life cycles, including sexual reproduction, are scarce, which limits the study of cellular transitions between different life cycle stages and prevents the visualization of cryptic stages. Although sequence-based techniques, such as -omics approaches, can reconstruct cellular transitions at the genetic and biochemical level, these methods are destructive and do not recover information from the same living cell over time. This protocol provides a solution to directly and continuously observe microbial life cycles, including sexual reproduction, by using microfluidics manipulations that expose single cells to nutritional stimuli and selective pressures. As proof of principle, we triggered a life cycle sequence transition in the model yeast Saccharomyces cerevisiae, starting with an arrest of proliferation in an ancestor cell followed by induction of meiosis through starvation, selection of sexually reproducing cells through exposure to a drug cocktail, germination of haploid spores, and mating of haploid individuals, creating a new descendant generation. This protocol offers the possibility to directly compare molecular and cellular behavior across life cycle stages and across sexually reproducing generations.