细胞生物学

In nature, filamentous fungi interact with plants. These fungi are characterized by rapid growth in numerous substrates and under minimal nutrient requirements. Investigating the interaction of these fungi with their plant hosts under controlled conditions is of importance for many researchers aiming to proceed with molecular or microscopical investigations of their favorite plant–fungus interaction system. The speed of growth of these fungi complicates transferring plant–fungal interaction systems in laboratory conditions. The issue is more complicated when monoxenic conditions are desired, to ensure that only two members (a fungus and a plant) are present in the system under study. Here, two simple closed systems for investigating plant–filamentous fungi associations under laboratory, monoxenic conditions are described, along with their limitations. The plant and fungal growth conditions, methods for sampling, staining, sectioning, and subsequent microscopical imaging of colonized plant tissues with affordable, common laboratory tools are described.

The organ of Corti, located in the inner ear, is the primary organ responsible for animal hearing. Each hair cell has a V-shaped or U-shaped hair bundle composed of actin-filled stereocilia and a kinocilium supported by true transport microtubules. Damage to these structures due to noise exposure, drug toxicity, aging, or environmental factors can lead to hearing loss and other disorders. The challenge when examining auditory organs is their location within the bony labyrinth and their small and fragile nature. This protocol describes the dissection procedure for the cochlear organ, followed by confocal imaging of immunostained endogenous and fluorescent proteins. This approach can be used to understand hair cell physiology and the molecular mechanisms required for normal hearing.

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.

With the growth of the quantum biology field, the study of magnetic field (MF) effects on biological processes and their potential therapeutic applications has attracted much attention. However, most biologists lack the experience needed to construct an MF exposure apparatus on their own, no consensus standard exists for exposure methods, and protocols for model organisms are sorely lacking. We aim to provide those interested in entering the field with the ability to investigate static MF effects in their own research. This protocol covers how to design, build, calibrate, and operate a static MF exposure chamber (MagShield apparatus), with instructions on how to modify parameters to other specific needs. The MagShield apparatus is constructed of mu-metal (which blocks external MFs), allowing for the generation of experimentally controlled MFs via 3-axial Helmholtz coils. Precise manipulation of static field strengths across a physiologically relevant range is possible: nT hypomagnetic fields, μT to < 1 mT weak MFs, and moderate MFs of several mT. An integrated mu-metal partition enables different control and experimental field strengths to run simultaneously. We demonstrate (with example results) how to use the MagShield apparatus with Xenopus, planarians, and fibroblast/fibrosarcoma cell lines, discussing the modifications needed for cell culture systems; however, the apparatus is easily adaptable to zebrafish, C. elegans, and 3D organoids. The operational methodology provided ensures uniform and reproducible results, affording the means for rigorous examination of static MF effects. Thus, this protocol is a valuable resource for investigators seeking to explore the intricate interplay between MFs and living organisms.

Tissue-engineered constructs combine the mechanical properties of biomaterials with biological agents to serve as scaffolds that direct the wound-healing process and promote tissue regeneration. A limitation to studying wound healing in vivo is that mouse skin contracts to heal rather than exhibiting granulation tissue formation and epithelialization like human skin. Therefore, it became necessary to develop a mouse model to better recapitulate human wound healing. The first splinted excisional wound healing model in mice, described in 2004, utilized silicone splints to prevent skin contracture. This model has been used to test a variety of wound healing strategies; however, to our knowledge, this model has not been adapted to test the effect of implants on wound healing. In our established protocol, circular bilateral excisional wounds are made on the mouse’s dorsum. A circular implant made of porous polyethylene is sutured to the skin within the wound. A thin, donut-shaped silicone splint is secured to the skin surrounding the wound, and a thick, donut-shaped splint is placed on top to tent the wound dressing. Finally, the mouse’s abdomen is wrapped in a bandage and tape to protect the implants. Our protocol offers a significant enhancement to the existing model by enabling the testing of implants for wound healing, as well as using an additional splint that prevents direct contact between the wound dressing and the wound bed. This model can be used to study tissue-engineered implant designs in a relatively low-cost, simple, and high-throughput manner before advancing to larger animal studies.

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

Neovascular diseases of the retina, such as diabetic retinopathy (DR) and age-related macular degeneration (AMD), are proliferative retinopathies involving the growth of new blood vessels on the retina, which in turn causes impairment and potential loss of vision. A drawback of conventional angiogenesis assays is that they are not representative of the angiogenic processes in the retina. In the retina, the new blood vessels grow (from pre-existing blood vessels) and migrate into a non-perfused region of the eye including the inner limiting membrane of the retina and the vitreous, both of which contribute to vision loss. The Matrigel Duplex Assay (MDA) measures the migration of angiogenic capillaries from a primary Matrigel layer to a secondary Matrigel layer, which resembles the pathological angiogenesis in AMD and DR. The methodology of MDA is comprised of two steps. In the first step, the human retinal microvascular endothelial cells (HRMECs) are mixed with phenol red–containing Matrigel (in a 1:1 ratio) and seeded in the center of an 8-well chamber slide. After 24 h, a second layer of phenol red–free Matrigel is overlaid over the first layer. Over the course of the next 24 h, the HRMECs invade from the primary Matrigel layer to the secondary layer. Subsequently, the angiogenic sprouts are visualized by brightfield phase contrast microscopy and quantified by ImageJ software. The present manuscript measures the angiogenesis-inhibitory activity of the Src kinase inhibitor PP2 in primary HRMECs using the MDA. The MDA may be used for multiple applications like screening anti-angiogenic drugs, measuring the pro-angiogenic activity of growth factors, and elucidating signaling pathways underlying retinal angiogenesis in normal and disease states.

Graphical overview