Cell Biology

The conventional Ploton silver method employs a high-concentration 50% (w/v; 2.943 mol/L) silver nitrate solution for histological staining and characterization of the osteocyte lacuno-canalicular system (LCS). However, it is limited by prolonged staining times (55 min) and by risks of LCS ultrastructural damage and/or incomplete impregnation. To address these limitations, we developed the Wu–Wang silver nitrate staining method, which uses a 1 mol/L silver nitrate solution under elevated temperature (50–70 °C) to achieve rapid, effective, and high-contrast visualization of the osteocyte LCS within 10 min. We further demonstrate that this novel method enables robust LCS visualization across multiple vertebrate species. Compared with the Ploton method, the Wu–Wang method substantially reduces staining time and overcomes staining limitations inherent to prolonged exposure to concentrated silver nitrate solutions. This rapid and efficient staining method supports more accurate quantitative analysis of LCS morphology and facilitates systematic investigation of osteocyte and LCS morphogenesis, as well as the pathological mechanisms underlying bone and joint disease.

Accurate identification of cell cycle stages is essential for investigating fundamental biological processes such as proliferation, differentiation, and tumorigenesis. While flow cytometry remains a widely used technique for such analyses, it is limited by its lack of single-cell resolution and its requirement for large sample sizes due to its population-based approach. These limitations underscore the need for alternative or complementary methods that offer single-cell precision with compatibility for small-scale applications. We present ImmunoCellCycle-ID, an immunofluorescence-based method that leverages the spatial distribution of endogenous markers, such as DNA, proliferating cell nuclear antigen (PCNA), centromere protein F (CENP-F), and centromere protein C (CENP-C), to reliably distinguish G1, early S, late S, early G2, late G2, and all mitotic sub-stages. This technique does not rely on precise signal quantification and utilizes standard immunofluorescence protocols alongside conventional laboratory microscopes, ensuring broad accessibility. Importantly, ImmunoCellCycle-ID detects endogenous proteins without the need for genetic modification, making it readily applicable to a wide range of human cell lines. Beyond its utility for single-cell resolution, the method can be scaled for population-level analyses, similar to flow cytometry. With its precision, versatility, and ease of implementation, ImmunoCellCycle-ID offers a powerful tool for high-resolution cell cycle profiling across diverse experimental platforms.

Cultured mammalian cells are a common model system for the study of epithelial biology and mechanics. Epithelia are often considered as pseudo–two dimensional and thus imaged and analyzed with respect to the apical tissue surface. We found that the three-dimensional architecture of epithelial monolayers can vary widely even within small culture wells, and that layers that appear organized in the plane of the tissue can show gross disorganization in the apical-basal plane. Epithelial cell shapes should be analyzed in 3D to understand the architecture and maturity of the cultured tissue to accurately compare between experiments. Here, we present a detailed protocol for the use of our image analysis pipeline, Automated Layer Analysis (ALAn), developed to quantitatively characterize the architecture of cultured epithelial layers. ALAn is based on a set of rules that are applied to the spatial distributions of DNA and actin signals in the apical-basal (depth) dimension of cultured layers obtained from imaging cultured cell layers using a confocal microscope. ALAn facilitates reproducibility across experiments, investigations, and labs, providing users with quantitative, unbiased characterization of epithelial architecture and maturity.

Key features

• This protocol was developed to spatially analyze epithelial monolayers in an automated and unbiased fashion.

• ALAn requires two inputs: the spatial distributions of nuclei and actin in cultured cells obtained using confocal fluorescence microscopy.

• ALAn code is written in Python3 using the Jupyter Notebook interactive format.

• Optimized for use in Marbin-Darby Canine Kidney (MDCK) cells and successfully applied to characterize human MCF-7 mammary gland–derived and Caco-2 colon carcinoma cells.

• This protocol utilizes Imaris software to segment nuclei but may be adapted for an alternative method. ALAn requires the centroid coordinates and volume of nuclei.

Graphical overview

Cell signaling is highly integrated for the process of various cell activities. Although previous studies have shown how individual genes contribute to cell migration, it remains unclear how the integration of these signaling pathways is involved in the modulation of cell migration. In our two-hit migration screen, we revealed that serine-threonine kinase 40 (STK40) and mitogen-activated protein kinase (MAPK) worked synergistically, and the suppression of both genes could further lead to suppression in cell migration. Furthermore, based on our analysis of cellular focal adhesion (FA) parameters using MATLAB analysis, we are able to find out the synergistic reduction of STK40 and MAPK that further abolished the increased FA by shSTK40. While FA identification in previous studies includes image analysis using manual selection, our protocol provides a semi-automatic manual selection of FAs using MATLAB. Here, we provide a method that can shorten the amount of time required for manual identification of FAs and increase the precision for discerning individual FAs for various analyses, such as FA numbers, area, and mean signals.

Expansion microscopy is an innovative method that enables super-resolution imaging of biological materials using a simple confocal microscope. The principle of this method relies on the physical isotropic expansion of a biological specimen cross-linked to a swellable polymer, stained with antibodies, and imaged. Since its first development, several improved versions of expansion microscopy and adaptations for different types of samples have been produced. Here, we show the application of ultrastructure expansion microscopy (U-ExM) to investigate the 3D organization of the green algae Chlamydomonas reinhardtii cellular ultrastructure, with a particular emphasis on the different types of sample fixation that can be used, as well as compatible staining procedures including membranes.

Graphical overview

Highly multiplexed protein measurements from multiple spatial scales using fluorescence microscopy recently emerged as a powerful way to investigate tumor microenvironments in biomedicine and the multivariate nature of complex systems’ interactions. A range of methods for this exist, which either rely on directly labeling the primary antibody with oligonucleotides/rare metals or employing methods to remove fluorescence for cyclic acquisition. Here, we describe a protocol that uses off-the-shelf primary and secondary antibodies without further need for modification and only commonly available chemical reagents. The method harnesses the observation that antibodies can crosslink to bound epitopes during light exposure, thus preventing elution. By utilizing a simple oxygen radical scavenging buffer during imaging and by blocking free sulfhydryl groups before antibody incubation, the presented method can employ comparably mild conditions to remove bound antibodies from epitopes, which preserves sample integrity. Thus, with the stated minor modifications, it allows for a standard immunofluorescence imaging protocol in cyclic fashion, currently permitting staining of up to ~80 unique epitopes.

Graphical overview

In the peripheral nervous system, Schwann cells are the primary type of glia. This protocol describes an in vitro differentiation and dedifferentiation system for rat Schwann cells. These cultures and systems can be used to investigate the morphological and biochemical effects of pharmacological intervention or lentivirus-mediated gene transfer on the process of Schwann cell differentiation or dedifferentiation.

Graphical abstract

Skeletal muscle disorders commonly affect the function and integrity of muscles. Novel interventions bring new potential to rescue or alleviate the symptoms associated with these disorders. In vivo and in vitro testing in mouse models allows quantitative evaluation of the degree of muscle dysfunction, and therefore, the level of potential rescue/restoration by the target intervention. Several resources and methods are available to assess muscle function and lean and muscle mass, as well as myofiber typing as separate concepts; however, a technical resource unifying these methods is missing. Here, we provide detailed procedures for analyzing muscle function, lean and muscle mass, and myofiber typing in a comprehensive technical resource paper.

Graphical abstract

Polarized actin cables in S. cerevisiae are linear bundles of crosslinked actin filaments that are assembled by two formins, Bnr1 (localized to the bud neck), and Bni1 (localized to the bud tip). Actin is polymerized at these two sites, which results in cables extending along the cell cortex toward the back of the mother cell. These cables serve as polarized tracks for myosin-based transport of secretory vesicles and other cargo, from the mother cell to the growing daughter cell. Until recently, descriptions of actin cable morphology and architecture have largely been qualitative or descriptive in nature. Here, we introduce a new quantitative method that enables more precise characterization of actin cable length. This technological advance generates quantitative datasets that can be used to determine the contributions of different actin regulatory proteins to the maintenance of cable architecture, and to assess how different pharmacological agents affect cable arrays. Additionally, these datasets can be used to test theoretical models, and be compared to results from computational simulations of actin assembly.

Graphical abstract:

Illustration of actin cable length and morphology analysis. (A) Representative maximum intensity projection image of S. cerevisiae fixed and stained with fluorescently-conjugated phalloidin to label F-actin (displayed in color), and fluorescently-conjugated Concanavalin A to label the cell wall (displayed in grey scale). Lengths of actin cables traced from the bud neck to their ends are indicated (dashed lines). (B) Inverted grey scale image of F-actin labelled with fluorescently-conjugated phalloidin and the cell wall traced in black. The length (purple) and end-to-end distance (green) of a single actin cable is indicated. Scale bar, 2 µm. (C–E) Actin cable length (C), end-to-end distance (D), and tortuosity (E) from hypothetical datasets, where each data point represents an individual cable and larger symbols represent the mean from each hypothetical experiment. Error bars, 95% confidence intervals.

Targeting receptor-mediated transcytosis (RMT) is a successful strategy for drug delivery of biologic agents across the blood-brain barrier (BBB). The recent development of human BBB organoid models is a major advancement to help characterize the mechanisms of RMT and thus accelerate the design of brain delivery technologies. BBB organoids exhibit self-organization, which resembles the architecture of the neurovascular unit, and low paracellular permeability, due to the formation of tight junctions between endothelial cells. However, current methods of organoid generation have low throughput, exhibit substantial heterogeneity across experiments, and require extensive manual handling. These limitations prevent the use of BBB organoids as a screening tool for discovery and optimization of therapeutic molecules. In this protocol, we use hydrogel-based arrays to generate human BBB organoids, with a 35-fold increase in organoid yield as compared to previous protocols using 96-well plates. We incubate BBB organoid arrays with monoclonal antibody-based constructs and use a custom semi-automated imaging assay to assess RMT within the organoid core. The experimental and analytical tools described in this protocol provide a scalable platform that can be incorporated in the early stages of drug discovery to accelerate the development and optimization of brain delivery technologies to cross the BBB.