Cell Biology

Primary cilia are evolutionarily conserved organelles that play critical roles in brain development. In the developing cortex, neural progenitors extend their primary cilia into the ventricular surface, where the cilia act as key signaling hubs. However, visualizing these cilia in a systematic and intact manner has been challenging. The commonly used cryostat sectioning only provides a limited snapshot of cilia on individual sections, and this process often disrupts the ciliary morphology. By contrast, the previously established whole-mount technique has been shown to preserve ciliary architecture in the adult mouse cortex. Here, we adapt and optimize the whole-mount approach for embryonic and neonatal brain, allowing robust visualization of ciliary morphology at the ventricular surface during development. This protocol describes step-by-step procedures for whole-mounting and immunostaining delicate embryonic and neonatal mouse cortices, enabling direct visualization of cilia in neural progenitors in the developing brain.

Expansion microscopy (ExM) enables nanoscale imaging of biological structures using standard fluorescence microscopes. Accurate labeling of cytoskeletal filaments, such as microtubules, remains challenging due to structural distortion and labeling inaccuracy during sample preparation. This protocol describes an optimized method combining detergent extraction and NHS-ester labeling for high-precision visualization of microtubules in expanded samples. Cytoplasmic components and membranes are selectively removed, preserving the ultrastructure of the microtubule network. Microtubules are digested into peptides during expansion and subsequently labeled at their N-termini using NHS-ester dyes, eliminating the need for antibodies. Effective fluorophore displacement of ~1 nm or lower is achieved, depending on the applied expansion factor. The protocol is compatible with both in vitro and cellular samples and can be integrated into a wide range of ExM workflows. Labeled microtubules can serve as internal reference standards for correcting expansion factors in ExM datasets.

The pancreatic islet, the only type of tissue that secretes insulin in response to elevated blood glucose, plays a vital role in diabetes development and treatment. While various islet vascularization strategies have been developed, they have been hindered by major limitations such as relying on pre-patterning and the inability to span long distances. Furthermore, few strategies have demonstrated robust enough vascularization in vivo to support therapeutic subcutaneous islet transplantation. Using adaptive endothelial cells (ECs) reprogrammed by transient expression of the ETS Variant Transcription Factor 2 (ETV-2) gene, we have physiologically vascularized human islets within a generic microchamber and have achieved functional engraftment of human islets in the subcutaneous space of mice. Such adaptive ECs, which we term reprogrammed vascular ECs (R-VECs), have been proven to be a suitable tool for both in vitro disease modeling and in vivo transplantation of not only islets but also other organoids.

The tissue explant culture (histoculture) is a method that involves maintaining small pieces taken from an organ ex vivo or post mortem in a controlled laboratory setting. Such a technique has a number of advantages: unlike the 2D, organoid, or on-chip cultures, tissue explants preserve the whole complexity of the original tissue in vivo, its structure, extracellular matrix, and the diverse cell populations, including resident immune cells. The explant culture method can be applied to human tissue specimens obtained from biopsies or autopsies, provided that proper ethical protocols are followed. This avoids the difficulties that may arise in translating results obtained on animal models into biomedical research for humans. This advantage makes histocultures especially desirable for studying human pathogenesis in the course of infectious diseases. The disadvantage of the method is the limited lifespan of the cultured tissues; however, a number of approaches allow extending tissue viability to a period sufficient for observing the infection onset and development. Here, we provide a protocol for lung explant maintenance that allows tracing the local effects of infection with SARS-CoV-2 in humans. Further applications of the lung tissues cultured according to this protocol include, but are not limited to, histochemical and immunohistochemical studies and microscopy, FACS, qPCR, and ELISA-based analysis of the conditioned culture media.

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.

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.