细胞生物学

Glycogen is a highly conserved macromolecule across species, and its visualization provides critical insights into both physiological processes and disease states. Existing approaches for glycogen imaging in Caenorhabditis elegans rely primarily on traditional microscopy slides, which introduce variability in image acquisition and downstream data analysis, limit throughput, and require substantial hands-on time and technical expertise.

Here, we present a standardized, cost-effective, and high-throughput imaging method that enables efficient visualization and quantification of glycogen in C. elegans. Our approach utilizes a custom-designed three-dimensional pad containing two to four chambers, allowing control and experimental samples to be processed simultaneously under identical conditions. Worms are exposed to iodine crystals, ensuring uniform staining while minimizing reagent use and handling variability. Imaging is performed using a simple binocular microscope, and analysis is conducted in Fiji, making the workflow accessible to laboratories with minimal specialized equipment or training.

This method also reduces technical variability, shortens turnaround time, and requires only basic reagents and expertise, making it well-suited for both research and teaching laboratories. Importantly, the platform is readily adaptable to other nematode species and scalable for large-scale genetic or pharmacological screening applications. Together, this workflow minimizes technical variability and provides a robust platform for comparative glycogen analysis in C. elegans.

Polyploid hepatocytes are one of the unique features of the liver. Some polyploid hepatocytes have chromosomes in a single nucleus (e.g., 1x4n, 1x8n), while others separate their chromosomes into two nuclei (e.g., 2x2n, 2x4n). In ploidy research, hepatocytes are typically sorted according to their cellular ploidy, revealing their contribution to tumorigenesis and cellular senescence. However, the conventional sorting method fails to distinguish 1x4n from 2x2n, or 1x8n from 2x4n cells, leaving it unclear whether hepatocytes with the same cellular ploidy but different nuclear configurations are identical or phenotypically different. Here, we describe a detailed protocol for fractionating mononucleated and binucleated hepatocytes. First, we present the method for isolating primary mouse hepatocytes and staining them with the DNA dye Hoechst 33342. Flow cytometry is then used to detect fluorescence differences between mononucleated and binucleated hepatocytes. This protocol enables the discrimination of hepatocyte subpopulations with identical cellular ploidy, providing a useful tool to investigate the functional heterogeneity of polyploid hepatocytes.

Fluorescence in situ hybridization (FISH) is a cytological method used to visualize specific oligonucleotide sequences within the cell. This method relies on the specific binding of a fluorescence-tagged probe, a short stretch of single-stranded polynucleotide, to its complementary sequence in the DNA or RNA, forming stable double-stranded hybrids. Fluorochromes, such as fluorescein, Alexa Fluor, cyanine dyes, or rhodamine, are attached to these probes to help in detecting their presence within the cell. Based on sequence complementarity, FISH allows for the visualization of the DNA or RNA with which they have hybridized. The distribution of these fluorochrome-tagged probes can be observed under a fluorescence or confocal microscope. The oligo(dT) FISH technique specifically utilizes a fluorochrome-tagged stretch of 40–50 thymidine (T) oligonucleotides that binds to the poly(A) tails of mature mRNAs within the cell. Newly transcribing pre-mRNAs and certain non-coding RNAs may not have poly(A) tails and therefore cannot be detected by this method. This step-by-step protocol outlines the oligo(dT) FISH technique for visualizing the cellular distribution of polyadenylated mRNAs in the tissues of Drosophila and other related model organisms.

Small fiber neuropathy (SFN) is an underdiagnosed condition characterized by sensory and autonomic dysfunction due to impairment of small nerve fibers in skin, blood vessels, and internal organs. Various underlying disorders are associated with SFN, and the pathophysiology of nerve fiber damage and functional impairment is the subject of extensive research. Diagnosis of SFN is challenging as standard electrodiagnostic techniques assess large fiber function and therefore are normal in SFN patients. The current gold standard for SFN diagnosis in humans is a skin biopsy, commonly obtained from the distal leg, hairy skin region, with evaluation of intraepidermal nerve fiber density (IENFD) using protein gene product 9.5 (PGP9.5) immunolabeling. While well-established in clinical practice, equivalent standardized, reproducible methods for assessing IENFD in experimental mouse models are lacking, which limits translational research in this field. Previous work in mice has relied on diverse antibodies, variable tissue sampling, and the use of confocal microscopy to trace nerve fibers. Other approaches have used chromogenic precipitate-based staining, which limits the ability to co-label multiple proteins. Here, we present a detailed, simple, and reproducible protocol for IENFD quantification of small nerves in the distal glabrous skin of the mouse hind paw. This protocol uses the two distal footpads, ensuring consistent sampling across animals. Prior to sectioning, the tissue is fixed and cryoprotected. Serial 20-μm sections are mounted on glass slides, dried, permeabilized, blocked, and immunostained with an anti-PGP9.5 monoclonal antibody, and then detected by binding secondary fluorescent-labeled antibodies. Although murine hairy skin analysis may apparently show a higher translational value, as it better reflects human biopsy sites, it is compromised by dense hair shafts and follicles, which interrupt epidermis continuity and thus interfere with sampling consistency. Polyneuropathy sensory symptoms, in fact, begin at the most distal sensory site, which is the glabrous skin of the toes. Thus, evaluation of this anatomical location best represents the clinical realm and may have the best sensitivity for identifying early axonal changes. In this protocol, we focused on IENFD quantification as done in human samples. Mechanoreceptors such as Meissner corpuscles are detectable and quantifiable by this method, and represent additional value since pressure-evoked pain, transmitted by these, is often reported by affected individuals. This immunolabeling protocol can be completed within one day [involving a small number of animals, where all three stages can be performed during a long working day (approximately 12 h)], while the entire workflow, including fixation and cryoprotection, is completed in up to 72 h. Importantly, the dermal and epidermal small fibers can be visualized using a standard fluorescence microscope, thereby avoiding the need for confocal imaging while maintaining high reproducibility. Preliminary validation in several animal models of inflammatory neuropathy and pain demonstrated a reproducible approximately 50% reduction in IENFD compared to controls, reaching statistical significance with n = 4 per group. This method supports SFN research and preclinical evaluation of novel therapeutics.

Immunohistochemistry (IHC) and immunofluorescence (IF) are fundamental molecular biology techniques to assess protein expression. However, the melanin present normally in the eye in the uveal tract (choroid, iris, and ciliary body) and the retinal pigment epithelium (RPE) poses a significant challenge for IHC and IF. This is because melanin interferes with both chromogenic and fluorescent detection methods. Additionally, formalin fixation, which is commonly used for IHC, can result in shrinkage and loss of cellular detail in the eye. This protocol provides an optimized approach using Davidson’s fixative with a hydrogen peroxide bleaching step to eliminate melanin interference in the mouse eye, improving the quality and interpretability of IHC analyses of the uveal tract and RPE. It is particularly useful for the analysis of uveal melanoma.

Accurate labeling of excitatory postsynaptic sites remains a major challenge for high-resolution imaging due to the dense and sterically restricted environment of the postsynaptic density (PSD). Here, we present a protocol utilizing Sylites, 3 kDa synthetic peptide probes that bind with nanomolar affinity to key postsynaptic markers, PSD-95 and Gephyrin. eSylites (excitatory Sylites) specifically target the PDZ1 and PDZ2 domains of PSD-95, enabling precise and efficient labeling of excitatory postsynaptic density (ePSD). In contrast, iSylites (inhibitory Sylites) bind to the dimerizing E-domain of the Gephyrin C-terminus, allowing selective visualization of inhibitory postsynaptic density (iPSD). Their small size reduces linkage error and enhances accessibility compared to conventional antibodies, enabling clear separation of PSD-95 nanodomains in super-resolution microscopy. The protocol is compatible with co-labeling using standard antibodies and integrates seamlessly into multichannel immunocytochemistry workflows for primary neurons and brain tissue. This method enables robust, reproducible labeling of excitatory synapses with enhanced spatial resolution and can be readily adapted for expansion microscopy or live-cell applications.

The persistence of the HIV-1 reservoir remains the ultimate obstacle in achieving a cure. Cure strategies targeting the HIV-1 reservoir are under development, and therefore, finding ways to improve the detection of the reservoir is crucial. Several reservoir detection techniques exist to assess different markers of the HIV-1 reservoir, such as PCR-based assays and protein-based flow cytometric methods. We developed a flow cytometry-fluorescent in situ hybridization (flow-FISH) approach that assesses HIV-1 at the transcriptional level. Using a combination of probes that target either the HIV-1 trans-activation response (TAR) region and 5′ long terminal repeat (LTR) or the Gag sequence, our assay distinguishes between infected cells expressing abortive or elongated HIV-1 RNAs. This assay utilizes the branched-DNA method to amplify the fluorescent signal of the hybridized RNA probes and can be used directly for thawed or cultured cells, with the option to include surface antibody staining. Cellular expression of abortive and/or Gag HIV-1 RNAs is measured by flow cytometry. Our flow-FISH approach gives insight into the transcriptional dynamics of the HIV-1 reservoir and allows for the characterization of latently infected cells.

One of the major factors contributing to aging and age-related diseases is the well-understood decline in the function of adult stem cells. Quantifying the degree of aging in adult stem cells is essential for advancing anti-aging mechanisms and developing anti-aging agents. However, no systematic approach to this exists. In this study, we developed a method to quantitatively assess the degree of aging in adult intestinal stem cells using a Drosophila midgut model and two aging markers. First, aging was induced in Drosophila with the desired genotype, and the anti-aging agent was administered 7 days before dissection. Then, the levels of two intestinal stem cell aging markers found in Drosophila (PH3 and γ-tubulin) were measured using immunohistochemistry. Finally, fluorescence microscopy was employed to count the number of aging markers and take images, which were analyzed using image analysis software. Using this approach, we quantitatively analyzed the effects of anti-aging agents on the aging of adult intestinal stem cells. This methodology is expected to significantly expedite the development of anti-aging agents and substantially reduce the research costs associated with aging-related studies.

In the intestinal epithelium, intraepithelial lymphocytes (IELs) coexist with intestinal epithelial cells (IECs). The IELs have an important role in defending the intestinal tract against pathogens and eliminating tumor cells. Anomalies in the absolute IEL count have been reported in various digestive diseases. IELs are typically counted using histologic techniques or under light microscopy after isolation of the epithelium. However, these techniques can introduce bias, which might account for the discrepancies in counts from one study to another. Here, we describe a flow cytometry assay for determining the absolute IEL count and the IEL/IEC ratio. We combined a conventional epithelial isolation method with a BD TruCount^TM bead-based absolute counting technique to quantify IELs (CD45⁺ CD326/EpCAM- CD103⁺CD3⁺) and IECs (CD45- CD326/EpCAM⁺) in a C57BL/6 mouse model.

The osteocyte lacuno-canalicular system (LCS) plays a crucial role in maintaining bone homeostasis and mediating cellular mechanotransduction. Current histological techniques, particularly the Ploton silver nitrate staining method, face challenges such as variations in solution concentrations and types as well as a lack of standardization, which limits their broader application in osteocyte research. In this study, we present a simplified and more effective silver nitrate staining protocol designed to address these issues. Our method utilizes a 1 mol/L silver nitrate solution combined with optimized gelatin-formic acid solutions at varying concentrations (0.05%–0.5% type-B gelatin and 0.05%–5% formic acid, or 1%–2% type-B gelatin and 0.1%–2% formic acid). Staining is performed for 1 h under 254 nm ultraviolet light or 90 min under room light, followed by washing with Milli-Q water to terminate staining. This novel optimized method yields consistent and distinct staining of the osteocyte LCS across multiple species, demonstrating superior efficiency and reliability compared to the Ploton method. It will significantly advance research in osteocyte biology and provide a valuable tool for exploring the adaptive evolution of osteocyte LCS morphology and function across various taxa.