细胞生物学

Current ischemic models strive to replicate ischemia-mediated injury. However, they face challenges such as inadequate reproducibility, difficulties in translating rodent findings to humans, and ethical, financial, and practical constraints that limit the accuracy of extensive research. This study introduces a novel approach to inducing persistent ischemia in 3-day-old chicken embryos using endothelin-1. The protocol targets the right vitelline arteries, validated with Doppler blood flow imaging and molecular biology experiments. This innovative approach facilitates the exploration of oxidative stress, inflammatory responses, cellular death, and potential drug screening suitability utilizing a 3-day-old chicken embryo.

Cultured mammalian cells are a common model system for the study of epithelial biology and mechanics. Epithelia are often considered as pseudo–two dimensional and thus imaged and analyzed with respect to the apical tissue surface. We found that the three-dimensional architecture of epithelial monolayers can vary widely even within small culture wells, and that layers that appear organized in the plane of the tissue can show gross disorganization in the apical-basal plane. Epithelial cell shapes should be analyzed in 3D to understand the architecture and maturity of the cultured tissue to accurately compare between experiments. Here, we present a detailed protocol for the use of our image analysis pipeline, Automated Layer Analysis (ALAn), developed to quantitatively characterize the architecture of cultured epithelial layers. ALAn is based on a set of rules that are applied to the spatial distributions of DNA and actin signals in the apical-basal (depth) dimension of cultured layers obtained from imaging cultured cell layers using a confocal microscope. ALAn facilitates reproducibility across experiments, investigations, and labs, providing users with quantitative, unbiased characterization of epithelial architecture and maturity.

Key features

• This protocol was developed to spatially analyze epithelial monolayers in an automated and unbiased fashion.

• ALAn requires two inputs: the spatial distributions of nuclei and actin in cultured cells obtained using confocal fluorescence microscopy.

• ALAn code is written in Python3 using the Jupyter Notebook interactive format.

• Optimized for use in Marbin-Darby Canine Kidney (MDCK) cells and successfully applied to characterize human MCF-7 mammary gland–derived and Caco-2 colon carcinoma cells.

• This protocol utilizes Imaris software to segment nuclei but may be adapted for an alternative method. ALAn requires the centroid coordinates and volume of nuclei.

Graphical overview

Structural and functional changes in vascular networks play a vital role during development, causing or contributing to the pathophysiology of injury and disease. Current methods to trace and image the vasculature in laboratory settings have proven inconsistent, inaccurate, and labor intensive, lacking the inherent three-dimensional structure of vasculature. Here, we provide a robust and highly reproducible method to image and quantify changes in vascular networks down to the capillary level. The method combines vasculature tracing, tissue clearing, and three-dimensional imaging techniques with vessel segmentation using AI-based convolutional reconstruction to rapidly process large, unsectioned tissue specimens throughout the body with high fidelity. The practicality and scalability of our protocol offer application across various fields of biomedical sciences. Obviating the need for sectioning of samples, this method will expedite qualitative and quantitative analyses of vascular networks. Preparation of the fluorescent gel perfusate takes < 30 min per study. Transcardiac perfusion and vasculature tracing takes approximately 20 min, while dissection of tissue samples ranges from 5 to 15 min depending on the tissue of interest. The tissue clearing protocol takes approximately 24–48 h per whole-tissue sample. Lastly, three-dimensional imaging and analysis can be completed in one day. The entire procedure can be carried out by a competent graduate student or experienced technician.

Key features

• This robust and highly reproducible method allows users to image and quantify changes in vascular networks down to the capillary level.

• Three-dimensional imaging techniques with vessel segmentation enable rapid processing of large, unsectioned tissue specimens throughout the body.

• It takes approximately 2–3 days for sample preparation, three-dimensional imaging, and analysis.

• The user-friendly pipeline can be completed by experienced and non-experienced users.

Graphical overview

The retina is a thin neuronal multilayer responsible for the detection of visual information. The first step in visual transduction occurs in the photoreceptor outer segment. The studies on photoreception and visual biochemistry have often utilized rod outer segments (OS) or OS disks purified from mammalian eyes. Literature reports several OS and disk purification procedures that rarely specify the procedure utilized to collect the retina from the eye. Some reports suggest the use of scissors, while others do not mention the issue as they declare to utilize frozen retinas. Because the OS are deeply embedded in the retinal pigmented epithelium (RPE), the detachment of the retina by a harsh pull-out can cause the fracture of the photoreceptor cilium. Here, we present a protocol maximizing OS yield. Eye semi-cups, obtained by hemisecting the eyeball and discarding the anterior chamber structures and the vitreous, are filled with Mammalian Ringer. After 10–15 min of incubation, the retinas spontaneously detach with their wealth of OS almost intact. The impressive ability of the present protocol to minimize the number of OS stuck inside the RPE, and therefore lost, compared with the classic procedure, is shown by confocal laser scanning microscopy analysis of samples stained ex vivo with a dye (MitoTracker deep red) that stains both retinal mitochondria and OS. Total protein assay of OS disks purified by either procedure also shows a 300% total protein yield improvement. The advantage of the protocol presented is its higher yield of photoreceptor OS for subsequent purification procedures, while maintaining the physiological features of the retina.

Senescence-associated beta-galactosidase (SA-β-GAL) is an enzyme that accumulates in the lysosomes of senescent cells, where it hydrolyses β-galactosides. With p16, it represents a well-recognized biomarker used to assess senescence both in vivo and in cell culture. The use of a chromogenic substrate, such as 5-bromo-4-chloro-3-indoyl-β-d-galactopyranoside (X-Gal), allows the detection of SA-β-GAL activity at pH 6.0 by the release of a visible blue product. Senescence occurs during aging and is part of the aging process itself. We have shown that prematurely aged zebrafish accumulate senescent cells detectable by SA-β-GAL staining in different tissues, including testis and gut. Here, we report a detailed protocol to perform an SA-β-GAL assay to detect senescent cell accumulation across the entire adult zebrafish organism (Danio rerio). We also identify previously unreported organs that show increased cell senescence in telomerase mutants, including the liver and the spinal cord.

Single cell RNA sequencing is a powerful tool that can be used to identify distinct cell types and transcriptomic differences within complex tissues. It has proven to be especially useful in tissues of the eye, where investigators have identified novel cell types within the retina, anterior chamber, and iridocorneal angle and explored transcriptomic contribution to disease phenotypes in age-related macular degeneration. However, to obtain high quality results, the technique requires isolation of healthy single cells from the tissue of interest, seeking complete tissue digestion while minimizing stress and transcriptomic changes in the isolated cells prior to library preparation. Here, we present a protocol developed in our laboratory for isolation of live single cells from the murine iridocorneal angle, which includes Schlemm’s canal and the trabecular meshwork, suitable for single cell RNA sequencing, flow cytometry, or other downstream analysis.

Graphical abstract:

Although herpes simplex virus 1 (HSV-1) is a well-studied virus, how the virus invades its human host via skin and mucosa to reach its receptors and initiate infection remains an open question. For studies of HSV-1 infection in skin, mice have been used as animal models. Murine skin infection can be induced after injection or scratching of the skin, which provides insights into disease pathogenesis but is clearly distinct from the natural entry route in human tissue. To explore the invasion route of HSV-1 on the tissue level, we established an ex vivo infection assay using skin explants. Here, we detail a protocol allowing the investigation of how the virus overcomes mechanical barriers in human skin to penetrate in keratinocytes and dermal fibroblasts. The protocol includes the preparation of total skin samples, skin shaves, and of separated epidermis and dermis, which is followed by incubation in virus suspension. The ex vivo infection assay allows the visualization, quantification, and characterization of single infected cells in the epidermis and dermis prior to viral replication and the virus-induced tissue damage. Hence, this experimental approach enables the identification of primary viral entry portals.

Graphical abstract:

Larval zebrafish have been established as an excellent model for examining vertebrate biology, with many researchers using the system for neuroscience. Controlling a fast escape response of the fish, the Mauthner cells and their associated network are an attractive model, given their experimental accessibility and fast development, driving ethologically relevant behavior in the first five days of development. Here, we describe methods for immunostaining electrical and chemical synapse proteins at 3-7 days post fertilization (dpf) in zebrafish using tricholoracetic acid fixation. The methods presented are ideally suited to easily visualize neural circuits and synapses within the fish.

Lymphatic vessels are abundant in the skin where they regulate interstitial fluid uptake and immune surveillance. Defects in dermal lymphatic vessels, such as fewer vessels and abnormal lymphatic vessel coverage with mural cells, are frequently associated with lymphedema and other lymphatic disorders. Whole-mount immunohistochemistry allows the visualization of dermal lymphatic vessels and identifies morphogenetic defects. Most dermal lymphatic vessels start growing during embryogenesis from lymph sacs that are located close to the axilla towards the dorsal and ventral midlines. Here, we present an approach that we have developed to permeabilize, immunolabel, clear, and visualize the lymphatic vessels. These simple and inexpensive techniques reproducibly generate images of dermal lymphatic vessels with great clarity.

Advances in C. elegans research have allowed scientists to recapitulate different human disorders, from neurodegenerative diseases to muscle dysfunction, in these nematodes. Concomitantly, the interest in visualizing organs affected by these conditions has grown, leading to the establishment of different antibody- and dye-based staining protocols to verify tissue morphology. In particular, the quality of muscle tissue has been largely used in nematodes as a readout for fitness and healthspan. Phalloidin derivatives, which are commonly used to stain actin filaments in cells and tissues, have been implemented in the context of C. elegans research for visualization of muscle fibers. However, the majority of the phalloidin-based protocols depend on fixation steps using harmful compounds, preparation of specific buffers, and large amounts of worms. Herein, we implemented a safer and more flexible experimental procedure to stain actin filaments in C. elegans using phalloidin-based dyes. Lyophilization of the worms followed by their acetone permeabilization allows bypassing the fixation process while also providing the opportunity to suspend the experiment at different steps. Moreover, by using conventional buffers throughout our protocol, we avoid the additional preparation of solutions. Finally, our protocol requires a limited number of worms, making it suitable for slow-growing C. elegans strains. Overall, this protocol provides an efficient, fast, and safer method to stain actin filaments and visualize muscle fibers in C. elegans.

Graphic abstract:

Schematic overview of phalloidin staining in C. elegans for assessing muscle fiber morphology.