细胞生物学

Quantitative analysis of biological membrane morphology is essential for understanding fundamental cellular processes such as organelle biogenesis and remodeling. While manual annotation has been the standard for complex structures, it is laborious and subjective, and conventional automated methods often fail to accurately delineate overlapping objects in 2D projected microscopy images. This protocol provides a complete, step-by-step workflow for the quantitative analysis of overlapping prospore membranes (PSMs) in sporulating yeast. The procedure details the synchronous induction of sporulation, acquisition of 3D fluorescence images and their conversion to 2D maximum intensity projections (MIPs), and the generation of a custom-annotated dataset using a semi-automated pipeline. Finally, it outlines the training and application of our mask R-CNN-based model, DeMemSeg, for high-fidelity instance segmentation and the subsequent extraction of morphological parameters. The primary advantage of this protocol is its ability to enable accurate and reproducible segmentation of individual, overlapping membrane structures from widely used 2D MIP images. This framework offers an objective, efficient, and scalable solution for the detailed quantitative analysis of complex membrane morphologies.

Musculoskeletal pathologies present challenges in athletic horses, often leading to functional impairment. The slow or limited regenerative capacity of bone, joint, and tendon/ligament injuries, coupled with the limitations of conventional treatments, highlights the need for innovative therapies such as ortho-biologics and mesenchymal stem/stroma cells. Traditional 2D cell culture systems with fetal bovine serum (FBS) fail to replicate the complexity of the in vivo environment, whereas 3D cultures more accurately mimic native tissue architecture and cell–cell interactions. This study describes a novel method for isolating muscle-derived progenitor cells in a 3D environment using an autologous plasma-based gel and an innovative cell retrieval solution. The cultured cells exhibit immunomodulatory effects on T lymphocytes, trilineage differentiation potential, and immunophenotypic characteristics consistent with conventional mesenchymal stem/stromal cells. This streamlined 3D culture technique offers a promising platform for generating minimally manipulated autologous cell products tailored for equine regenerative medicine.

Detecting the proliferation of cells with copper(I)-catalyzed azide-alkyne cycloaddition (click chemistry) and the thymidine analogue, 5-ethynyl-2’-deoxyuridine (EdU), is a simpler and more versatile method than traditional antibody-based approaches. Instead of the harsh series of steps typically used for 5-bromo-2’-deoxyuridine (BrdU) detection, detecting EdU does not require DNA denaturation and is suitable for use with other applications. This approach was implemented in an animal model of ischemic stroke. The following protocol details how to use EdU to label, track, and visualize leukocyte recruitment for flow cytometry and fluorescence microscopy, including the processes for EdU injection and blood and tissue sample preparation. Considerations for timing, dosing, and cell viability are also outlined to tailor the protocol to experimental needs. This method could be applied to various models that require extended tracking periods, as the signal from EdU can last several cell divisions, depending on cell type and condition.

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.

Microglia, the resident immune cells of the central nervous system, play a crucial role in maintaining neural homeostasis and in regulating neurodevelopment, neuroinflammation, tissue repair, and neurotoxicity. They are also key contributors to the pathogenesis of various neurodegenerative disorders, underscoring the need for in vitro models that accurately recapitulate disease-relevant conditions. Among the available isolation methods, the classical mixed glial culture shaking technique remains the most commonly employed, while alternatives such as magnetic bead separation and fluorescence-activated cell sorting (FACS) offer higher purity but are often constrained by technical complexity and cost. In this study, we refined the traditional shaking method by supplementing specific cytokines during culture to enhance microglial viability and proliferation. Our optimized protocol produced primary microglia with higher purity, greater yield, and improved viability compared with the conventional approach, thereby increasing experimental efficiency while substantially reducing time, animal usage, and overall cost.

Lipid droplets have emerged as dynamic organelles involved in diverse cellular processes beyond simple lipid storage. In plants and cyanobacteria, growing evidence highlights their importance in stress adaptation and signaling, yet methods to study their structure and purity remain limited. Traditionally, in situ transmission electron microscopy (TEM) has been used to visualize lipid droplets within intact cells. While powerful, this approach cannot easily evaluate isolated lipid droplets or confirm their purity. In this protocol, we describe a rapid method for preparing and visualizing cyanoglobule lipid droplets isolated from cyanobacteria. The isolated droplets are directly processed for TEM using negative staining with uranyl acetate, providing a straightforward and efficient workflow. The procedure can be applied broadly to lipid droplets from diverse organisms, independent of species or cellular origin. This protocol offers a simple, fast, and widely applicable approach to assessing lipid droplets, expanding the toolkit for researchers studying their structure and function.

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.

Zebrafish are a powerful model for investigating vascular and lymphatic biology due to their genetic tractability and optical transparency. While translating ribosome affinity purification (TRAP) has been widely applied in other systems, its application in zebrafish has remained limited. Here, we present an optimized TRAP protocol for isolating ribosome-associated mRNAs from endothelial cells in vivo, without the need for cell dissociation or sorting. Using a novel transgenic zebrafish line, which expresses HA-tagged Rpl10a under the mrc1a promoter, we enriched actively translating endothelial transcripts. Differential expression analysis revealed robust upregulation of vascular and lymphatic genes including flt4, kdrl, and lyve1b. This approach captures the endothelial cell translatome with high specificity and offers a robust platform for investigating the molecular mechanisms of endothelial biology under genetic, environmental, or toxicological perturbations.

Cerebrospinal fluid-contacting neurons (CSF-cNs) are a specialized group of multifunctional neurons located around the central canal of the spinal cord. They play critical roles in motor regulation, postural maintenance, and spinal cord injury repair. However, the molecular mechanisms underlying the multifunctionality of CSF-cNs remain poorly understood, partly due to the lack of established in vitro methods for their efficient selection and purification, which significantly hinders mechanistic investigations. In this study, we describe a standardized method using a PKD2L1 promoter-driven lentiviral system, which enables effective enrichment and identification of CSF-cNs in vitro through GFP labeling and puromycin selection. This protocol includes key steps such as construction of the PKD2L1 promoter-driven lentiviral vector, spinal cord tissue collection and digestion from neonatal mice, lentiviral infection, antibiotic selection, and immunofluorescence-based identification of CSF-cNs. Our method provides a reliable platform for obtaining high-purity CSF-cNs (>99%), which facilitates their functional and mechanistic studies for regenerative approaches in vitro.

Following myocardial infarction (MI), myocardial cells undergo cell death, and the necrotic region is replaced by extracellular matrix (ECM) proteins such as collagens. Myofibroblasts are responsible for producing these ECM proteins. Cardiac myofibroblasts are differentiated from resident fibroblasts in response to inflammation. To date, genetically modified mice driven by the Periostin promoter and adeno-associated virus 9 (AAV9) carrying the Periostin promoter have been used for gene transfer into cardiac myofibroblasts. However, these methods require multiple steps and are time-consuming and expensive. Therefore, we developed a method for delivering genes into cardiac myofibroblasts using retroviruses. Specifically, the DNA of the target gene was transfected into Plat-E cells, which are packaging cells, to generate retroviruses. The virus-containing supernatant was then harvested, and the viruses were pelleted by centrifugation and suspended in PBS-containing polybrene. Subsequently, permanent occlusion of the left coronary artery was performed, and 20 μL of viral solution was immediately administered using a 29G needle at a position 1–2 mm below the ligation site in the heart of mice maintained in an open chest state. Using this method, we were able to introduce genes into the myofibroblasts of interest surrounding the MI site.