细胞生物学

Gap junctions are transmembrane protein channels that enable the exchange of small molecules such as ions, second messengers, and metabolites between adjacent cells. Gap junctions are found in various mammalian organs, including skin, endothelium, liver, pancreas, muscle, and central nervous system (CNS). In the CNS, they mediate coupling between neural cells including glial cells, and the resulting panglial networks are vital for brain homeostasis. Tracers of sufficiently small molecular mass can diffuse across gap junctions and are used to visualize the extent of cell-to-cell coupling in situ by delivering them to a single cell through sharp electrodes or patch-clamp micropipettes. Here, we describe a protocol for pre-labeling and identification of astrocytes in acute mouse forebrain slices using Sulforhodamine 101 (SR101). Fluorescent cells can then be targeted for whole-cell patch-clamp, which allows for further confirmation of astroglial identity by assessing their electrophysiological properties, as well as for passive dialysis with a tracer such as biocytin. Slices can then be subjected to chemical fixation and immunostaining to detect dye-coupled networks. This protocol provides a method for the identification of astrocytes in live tissue through SR101 labeling. Alternatively, transgenic reporter mice can also be used to identify astrocytes. While we illustrate the use of this protocol for the study of glial networks in the mouse brain, the general principles are applicable to other species, tissues, and cell types.

Mitochondrial cristae, formed by folding the mitochondrial inner membrane (IM), are essential for cellular energy supply. However, the observation of the IM is challenging due to the limitations in spatiotemporal resolution offered by conventional microscopy and the absence of suitable in vitro probes specifically targeting the IM. Here, we describe a detailed imaging protocol for the mitochondrial inner membrane using the Si-rhodamine dye HBmito Crimson, which has excellent photophysical properties, to label live cells for imaging via stimulated emission depletion (STED) microscopy. This allows for STED imaging over more than 500 frames (approximately one hour), with a spatial resolution of 40 nm, enabling the observation of cristae dynamics during various mitochondrial processes. The protocol includes detailed steps for cell staining, image acquisition, image processing, and resolution analysis. Utilizing the superior resolution of STED microscopy, the structure and complex dynamic changes of cristae can be visualized.

Plants use CO₂, water, and light energy to generate carbohydrates through photosynthesis. During daytime, these carbohydrates are polymerized, leading to the accumulation of starch granules in chloroplasts. The catabolites produced by the degradation of these chloroplast starch granules are used for physiological responses and plant growth. Various staining methods, such as iodine staining, have previously been used to visualize the accumulation of chloroplast starch granules; however, these staining methods cannot be used to image live cells and/or provide confocal images with non-specific signals. In this study, we developed a new imaging method for the fluorescent observation of chloroplast starch granules in living plant cells by staining with fluorescein, a widely available fluorescent dye. This simple staining method, which involves soaking a leaf disk in staining solution, shows high specificity in confocal images. Fluorescent images of the stained tissue allow the cellular starch content of living cells to be quantified with the same level of accuracy as a conventional biochemical method (amyloglucosidase/α-amylase method). Fluorescein staining thus not only enables the easy and clear observation of chloroplast starch granules but also allows for precise quantification in living cells.

Here, we describe immunofluorescent (IF) staining assay of 3D cell culture colonoids isolated from mice colon as described previously. Primary cultures developed from isolated colonic stem cells are called colonoids. Immunofluorescence can be used to analyze the distribution of proteins, glycans, and small molecules—both biological and non-biological ones. Four-day-old colonoid cell cultures grown on Lab-Tek 8-well plate are fixed by paraformaldehyde. Fixed colonoids are then subjected to antigen retrieval and blocking followed by incubation with primary antibody. A corresponding secondary antibody tagged with desired fluorescence is used to visualize primary antibody–marked protein. Counter staining to stain actin filaments and nucleus to assess cell structure and DNA in nucleus is performed by choosing the other two contrasting fluorescences. IF staining of colonoids can be utilized to visualize molecular markers of cell behavior. This technique can be used for translation research by isolating colonoids from colitis patients’ colons, monitoring the biomarkers, and customizing their treatments.

Key features

• Analysis of molecular markers of cell behavior.

Protocol to visualize proteins in 3D cell culture.

• This protocol requires colonoids isolated from mice colon grown on matrigel support.

• Protocol requires at least eight days to complete.

Graphical overview

B cells play a critical role in host defense, producing antibodies in response to microbial infection. An inability to produce an effective antibody response leaves affected individuals prone to serious infection; therefore, proper B-cell development is essential to human health. B-cell development begins in the bone marrow and progresses through various stages until maturation occurs in the spleen. This process involves several sequential, complex events, starting with pre- and pro-B cells, which rearrange the heavy and light chain genes responsible for producing clonally diverse immunoglobulin (Ig) molecules. These cells then differentiate into immature B cells, followed by mature B cells. The bone marrow is a complex ecological niche of supporting stromal cells, extracellular matrix components, macrophages, and hematopoietic precursor cells influencing B-cell development, maturation, and differentiation. Once fully mature, B cells circulate in peripheral lymphoid organs and can respond to antigenic stimuli. As specific cell surface markers are expressed during each stage of B-cell development, researchers use flow cytometry as a powerful tool to evaluate developmental progression. In this protocol, we provide a step-by-step method for bone marrow isolation, cell staining, and data analysis. This tool will help researchers gain a deeper understanding of the progression of B-cell development and provide a pertinent flow gating strategy.

Sphingolipids are important structural components of cellular membranes. They also function as prominent signaling molecules to control a variety of cellular events, such as cell growth, differentiation, and apoptosis. Impaired sphingolipid metabolism, particularly defects in sphingolipid degradation, has been associated with many human diseases. Fluorescence sphingolipid analogs have been widely used as efficient probes to study sphingolipid metabolism and intracellular trafficking in living mammalian cells. Compared with nitrobenzoxadiazole fluorophores (NBD FL), the boron dipyrromethene difluoride fluorophores (BODIPY FL) have much higher absorptivity and fluorescence quantum. These features allow more intensive labeling of cells for fluorescence microscopy imaging and flow cytometry analysis. Here, we describe a protocol employing BODIPY FL-labeled sphingolipid analogs to elucidate sphingolipid internalization, trafficking, and endocytosis in mouse embryonic stem cells.

Graphical abstract:

Employing a novel mouse model of immune related adverse events (irAEs) induced by combination of anti-PD1 and anti-CTLA-4 antibodies, we visualized immune infiltration into the liver, lung, pancreas, and colon. Here, we describe the avidin-biotin conjugate (ABC) method used to stain T cells (CD4 and CD8), B cells (CD19), macrophages (F4/80), and cells bound by the in vivo administered rat anti-mouse antibodies for chromogenic immunohistochemistry (IHC). Using a biotinylated goat anti-rat antibody, we detected the localization of cells bound to the in vivo antibodies for PD-1 and CTLA-4. IHC has advantages over other techniques, namely antibody availability, resistance to photobleaching, and greater sensitivity. Additionally, detection and localization of in vivo antibodies can be used in mice models to infer their therapeutic efficacy, stability, and function.

Graphical abstract:

Macrophages are important for host defense against intracellular pathogens like Salmonella and can be differentiated into two major subtypes. M1 macrophages, which are pro-inflammatory and induce antimicrobial immune effector mechanisms, including the expression of inducible nitric oxide synthase (iNOS), and M2 macrophages, which exert anti-inflammatory functions and express arginase 1 (ARG1). Through the process of phagocytosis, macrophages contain, engulf, and eliminate bacteria. Therefore, they are one of the first lines of defense against Salmonella. Infection with Salmonella leads to gastrointestinal disorders and systemic infection, termed typhoid fever. For further characterization of infection pathways, we established an in vitro model where macrophages are infected with the mouse Salmonella typhi correlate Salmonella enterica serovar Typhimurium (S.tm), which additionally expresses red fluorescent protein (RFP). This allows us to clearly characterize macrophages that phagocytosed the bacteria, using multi-color flow cytometry.

In this protocol, we focus on the in vitro characterization of pro- and anti-inflammatory macrophages displaying red fluorescent protein-expressing Salmonella enterica serovar Typhimurium, by multi-color flow cytometry.

The ability to stain lipid stores in vivo allows for the facile assessment of metabolic status in individuals of a population following genetic and environmental manipulation or pharmacological treatment. In the animal model Caenorhabditis elegans, lipids are stored in and mobilized from intracellular lipid droplets in the intestinal and hypodermal tissues. The abundance, size, and distribution of these lipids can be readily assessed by two staining methods for neutral lipids: Oil Red O (ORO) and Nile Red (NR). ORO and NR can be used to quantitatively measure lipid droplet abundance, while ORO can also define tissue distribution and lipid droplet size. C. elegans are a useful animal model in studying pathways relating to aging, fat storage, and metabolism, as their transparent nature allows for easy microscopic assessment of lipid droplets. This is done by fixation and permeabilization, staining with NR or ORO, image capture on a microscope, and computational identification and quantification of lipid droplets in individuals within a cohort. To ensure reproducibility in lipid measurements, we provide a detailed protocol to measure intracellular lipid dynamics in C. elegans.

Graphic abstract:

Flow chart depicting the preparation of C. elegans for fat staining protocols.

Although the advent of genetically-encoded fluorescent markers, such as the green fluorescent protein (GFP; Chalfie et al., 1994), has enabled convenient visualization of gene expression in vivo, this method is generally not effective for detecting post-translational modifications because they are not translated from DNA sequences. Genetically-encoded, fluorescently-tagged transgene products can also be misleading for observing expression patterns because transgenes may lack endogenous regulatory DNA elements needed for precise regulation of expression that could result in over or under expression. Fluorescently-tagged proteins created by CRISPR genome editing are less prone to defective expression patterns because the loci retain endogenous DNA elements that regulate their transcription (Nance and Frøkjær-Jensen, 2019). However, even CRISPR alleles encoding heritable fluorescently-tagged protein markers can result in defects in function or localization of the gene product if the fluorescent tag obstructs or otherwise interferes with important protein interaction domains or affects the protein structure.

Indirect immunofluorescence is a method for detecting endogenous gene expression or post-translational modifications without the need for transgenesis or genome editing. Here, we present a reliable protocol in which C. elegans nematodes are fixed, preserved, and permeabilized for staining with a primary antibody to bind proteins or post-translational modifications, which are then labeled with a secondary antibody conjugated to a fluorescent dye. Use of this method may be limited by the availability of (or ability to generate) a primary antibody that binds the epitope of interest in fixed animals. Thousands of animals are simultaneously subjected to a series of chemical treatments and washes in a single centrifuge tube, allowing large numbers of identically-treated stained animals to be examined. We have successfully used this protocol (O’Hagan et al., 2011 and 2017; Power et al., 2020) to preserve and detect post-translational modifications of tubulin in C. elegans ciliated sensory neurons and to detect non-modified endogenous protein (Topalidou and Chalfie, 2011).