细胞生物学

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.

Whole-mount techniques are widely used in medical and biological research to analyze protein expression and tissue organization in intact specimens. Traditional approaches for protein localization include section-based immunohistochemistry and in situ hybridization; however, these methods can be limited by tissue disruption and loss of spatial context. Whole-mount protocols generally involve tissue fixation, permeabilization, and staining with specific probes, but their effectiveness varies depending on the antigen–antibody combination and the specimen type. Consequently, no universal protocol is suitable for all experimental conditions. This protocol presents a detailed whole-mount immunostaining protocol for evaluating tyrosine hydroxylase (TH) expression, a key marker of dopaminergic neurons, in zebrafish (Danio rerio) larvae. The procedure outlines critical steps from sample preparation to staining optimization to ensure reproducible and specific signal detection. This approach enables accurate visualization and analysis of dopaminergic neuron distribution in intact larvae. The protocol offers a reliable and adaptable approach that preserves tissue integrity, enables three-dimensional visualization, and is particularly well-suited for developmental and neurobiological studies using zebrafish larvae.

Postnatal mouse retinal vascular development is a widely used model for studying retinal vascular diseases and evaluating candidate therapies. This is particularly relevant for inherited disorders such as familial exudative vitreoretinopathy (FEVR), in which impaired vascular growth and organization are central to disease pathogenesis. Numerous approaches have been used to assess retinal vasculature in mouse flat mounts, ranging from qualitative descriptions to limited quantitative measurements of vascular growth. However, phenotypic variability across genetic models, including different models of FEVR, complicates comparisons and underscores the need for standardized, comprehensive multi-parameter analyses that are suitable for rapid and cost-effective screening studies. We describe a standardized morphometric protocol using ImageJ software to quantitatively analyze mouse retinal vasculature in a reproducible manner. The protocol begins with measurement of areas of vascular disorganization (meshes) as well as total vascular and retinal area. Two defined regions in the peripheral and midperipheral retina are then selected to quantify cell clusters, followed by image processing, binarization, and skeletonization. From these processed images, vascular density, branch number, branch length and thickness, junction number, triple points, and box-counting fractal dimension and lacunarity are quantified. Overall, this protocol provides a rapid, cost-effective, and standardized framework for quantifying retinal vascular phenotypes across diverse mouse models. By capturing multiple structural features and accommodating phenotypic variability, it is well-suited for comparative studies and therapeutic screening in retinal vascular disease.

The mammalian central circadian clock resides in the suprachiasmatic nucleus (SCN) of the hypothalamus in the brain and is responsible for coordinating daily rhythms of biological processes spanning from gene expression to behavior. Light, the primary environmental zeitgeber, entrains the SCN via melanopsin-expressing intrinsically photosensitive retinal ganglion cells that project through the retino-hypothalamic tract. Altered circadian rhythms are common in individuals diagnosed with neurodevelopmental and neurodegenerative disorders, and often, associated with structural alterations of the SCN and impaired retinal input; importantly, these anomalies can be recapitulated in animal models. Here, we describe step-by-step protocols for quantitative histomorphometrical analysis of the SCN and the assessment of retinal–SCN connectivity, previously used in mouse models of neurodevelopmental and neurodegenerative disorders. These include measurement of the SCN area, perimeter, height and width using Nissl- or DAPI-stained coronal sections, as well as densitometric and plot profile analyses of cholera toxin β-subunit–labeled retinal projections using Axiovision or Fiji/ImageJ. The protocols incorporate standardized region-of-interest, measurements by masked observers, and consistent scaling procedures to enhance reproducibility. These methods provide a rigorous framework for detecting structural anomalies and connectivity defects in the circadian system and can be broadly applied to other experimental models of circadian dysfunction.

Formalin-fixed paraffin-embedded (FFPE) slides are essential for histological and immunohistochemical analyses of organoids. Conventional preparation of FFPE slides from organoids embedded in basement membrane extract (BME) presents several challenges. During the fixation step, dehydration often causes collapse of the BME, which normally supports the three-dimensional architecture of organoids. As a result, organoids may lose their original morphology, particularly in the case of cystic or structurally delicate types, leading to distortion and reduced reliability in downstream histological evaluation. Here, we introduce a straightforward protocol that improves the reliability of FFPE slide preparation for BME-based organoids by enhancing sample integrity and sectioning quality. By using 2% agarose as a mold during the embedding process, organoids grown in BME were effectively stabilized, enabling reliable preservation of their morphology throughout FFPE slide preparation. This method effectively addresses the difficulties in processing structurally delicate organoids and allows robust preparation of diverse cancer organoid morphologies—such as cystic, dense, and grape-like structures—while maintaining their native three-dimensional architecture. Our approach simplified the technical process while ensuring reliable histopathological analysis, making it a valuable tool for cancer research and personalized medicine.

Skeletal muscle–specific stem cells are responsible for regenerating damaged muscle tissue following strenuous physical activity. These muscle stem cells, also known as satellite cells (SCs), can activate, proliferate, and differentiate to form new skeletal muscle cells. SCs can be identified and visualized utilizing optical and electron microscopy techniques. However, studies identifying SCs using fluorescent imaging techniques vary significantly within their methodology and lack fundamental aspects of the guidelines for rigor and reproducibility that must be included within immunohistochemical studies. Therefore, a standardized method for identifying human skeletal muscle stem cells is warranted, which will improve the reproducibility of future studies investigating satellite activity. Additionally, although it has been suggested that SC shape can change after exercise, there are currently no methods for examining SC morphology. Thus, we present an integrated workflow for three-dimensional visualization of satellite cell nuclei, validated by the spatial context of the fluorescent labeling and multichannel signal overlap. Our protocol includes, from start to finish, post-biopsy extraction and embedding, tissue sectioning, immunofluorescence, imaging steps and acquisition, and three-dimensional data post-processing. Because of the depth volume generated from the confocal microscope z-stacks, this will allow future studies to investigate the morphology of SC nuclei and their activity, instead of traditionally observing them in two-dimensional space (x, y).

Histological techniques to study muscle are crucial for assessing skeletal muscle health. To preserve tissue morphology, samples are usually fixed in formaldehyde or cryopreserved immediately after excision from the body. Freezing samples in liquid nitrogen, using isopentane as a mediator for efficient cooling, preserves the tissue in its natural state. However, this method is highly susceptible to freeze-fracture artifacts, which alter or destroy tissue architecture. Isopentane is most commonly used in a semi-frozen/liquid state that is visually assessed by the experimenter, which can pose a challenge when freezing multiple tissues at a time or maintaining a consistent temperature. Furthermore, tissue size is also a confounding factor; depending on the size, freezing times can vary. In this study, we compare two different options for using isopentane while cryopreserving tissue. We also present an easy and reproducible method of freezing the soleus tissue of mice using frozen isopentane. This method decreased the occurrence of freeze-fractures by an order of magnitude, to ~4%, whereas the traditional method of cryopreservation resulted in ~56% freeze-fracturing.

The mammary gland undergoes functional, developmental, and structural changes that are essential for lactation and reproductive processes. An overview of such unique tissue can offer clearer insights into mammary development and tumorigenesis. Compared to traditional methods, mouse mammary gland whole mount is a pivotal technique that provides three-dimensional structural perspectives on gland morphology and developmental stages, offering an inexpensive and accessible approach. This protocol outlines the tissue isolation of the mouse mammary gland and provides detailed instructions for whole-mount staining and analysis. Mammary gland tissues are carefully dissected from euthanized mice and stained with Carmine Alum to highlight the ductal structures, enabling detailed visualization of the branching patterns and morphological changes. Light microscopy is used to capture a panoramic image of the stained mammary gland, enabling the quantitative analysis of terminal end buds (TEBs) and bifurcated TEBs to further investigate mammary gland remodeling. This method can provide invaluable insights, particularly in the study of mammary gland morphogenesis and tumorigenesis, underscoring its significance in both basic research and clinical applications.

Acute respiratory distress syndrome (ARDS) is a life-threatening, high mortality pulmonary condition characterized by acute lung injury (ALI) resulting in diffuse alveolar damage. Despite progress regarding the understanding of ARDS pathophysiology, there are presently no effective pharmacotherapies. Due to the complexity and multiorgan involvement typically associated with ARDS, animal models remain the most commonly used research tool for investigating potential new therapies. Experimental models of ALI/ARDS use different methods of injury to acutely induce lung damage in both small and large animals. These models have historically played an important role in the development of new clinical interventions, such as fluid therapy and the use of supportive mechanical ventilation (MV). However, failures in recent clinical trials have highlighted the potential inadequacy of small animal models due to major anatomical and physiological differences, as well as technical challenges associated with the use of clinical co-interventions [e.g., MV and extracorporeal membrane oxygenation (ECMO)]. Thus, there is a need for larger animal models of ALI/ARDS, to allow the incorporation of clinically relevant measurements and co-interventions, hopefully leading to improved rates of clinical translation. However, one of the main challenges in using large animal models of preclinical research is that fewer species-specific experimental tools and metrics are available for evaluating the extent of lung injury, as compared to rodent models. One of the most relevant indicators of ALI in all animal models is evidence of histological tissue damage, and while histological scoring systems exist for small animal models, these cannot frequently be readily applied to large animal models. Histological injury in these models differs due to the type and severity of the injury being modeled. Additionally, the incorporation of other clinical support devices such as MV and ECMO in large animal models can lead to further lung damage and appearance of features absent in the small animal models. Therefore, semi-quantitative histological scoring systems designed to evaluate tissue-level injury in large animal models of ALI/ARDS are needed. Here we describe a semi-quantitative scoring system to evaluate histological injury using a previously established porcine model of ALI via intratracheal and intravascular lipopolysaccharide (LPS) administration. Additionally, and owing to the higher number of samples generated from large animal models, we worked to implement a more sustainable and greener histopathological workflow throughout the entire process.

Autoreactive T cells in autoantibody-mediated autoimmune diseases can be divided into two major subsets: (i) follicular T helper cells (Tfh) that provide T cell help in germinal centers (GC) and (ii) effector T (Teff) cells that immigrate into peripheral tissue sites such as the skin and mediate local inflammation. To study the sequence of events leading to the loss of tolerance in autoantibody-mediated autoimmune diseases it is required to investigate both T cell subsets simultaneously. This approach is hampered mainly because the appearance of skin inflammation in mouse models is a random process, which makes it difficult to define the location of inflammation at the right time point. To overcome this problem, we developed a scratching technique for ear skins that leads to the establishment of chronic autoimmune wounds in the mouse model for the pemphigoid-like disease epidermolysis bullosa acquisita. By defining the exact place where the skin wounds should form, this protocol enables a detailed analysis of skin-immigrating Teff cells. Of note, this protocol induces GC in draining lymph nodes in parallel so that Tfh cells in GC can be investigated concurrently. This protocol is not restricted to T cells and can be used for any other skin-immigrating inflammatory cells.