Cell Biology

This protocol describes a reproducible workflow for modeling in vitro impact-induced traumatic brain injury (TBI) using a mechanical stretch system applied to differentiated SH-SY5Y human neuroblastoma cells cultured on polydimethylsiloxane (PDMS) substrates. The protocol integrates three primary components: (1) fabrication and surface modification of deformable PDMS chambers to support cellular adhesion, (2) partial differentiation of SH-SY5Y cells using retinoic acid, and (3) induction of controlled mechanical strain to simulate mild to moderate TBI. The stretch-induced injury model enables quantitative assessment of cellular viability and recovery following mechanical insult. This approach provides a versatile platform for studying cellular and molecular mechanisms of TBI, screening neuroprotective compounds, and exploring mechanobiological responses in neural cells under controlled strain magnitudes and rates.

Lipid peroxidation (LPO) is a major indicator of oxidative stress and cellular damage, frequently associated with environmental and toxicological stressors and mechanistically linked to ferroptotic regulated cell death (RCD). This protocol describes a simple and reproducible method for the qualitative in situ visualization of LPO in mosquito larvae using Schiff’s reagent, which histochemically labels reactive aldehyde groups [such as malondialdehyde (MDA)] generated during lipid degradation. Although Schiff’s reagent detects aldehydes commonly associated with lipid peroxidation, these compounds are not exclusive to LPO and may also arise from other oxidative processes. The method preserves tissue integrity, enabling direct, spatially resolved observation of oxidative damage in whole larvae. Following staining, larvae are rinsed in a stabilizing sulfite solution to maintain the characteristic magenta coloration. Using this assay, Culex quinquefasciatus larvae exposed to ferroptotic cyanobacteria, such as Synechocystis sp., exhibit a marked accumulation of lipid-derived aldehydes consistent with lipid ROS–mediated damage. This oxidative response is specifically suppressed by pre-treatment with the canonical ferroptosis inhibitor Ferrostatin-1 (Fer-1), which inhibits lipid peroxidation and significantly reduces larval mortality. As a complementary approach to traditional spectrophotometric assays such as thiobarbituric acid reactive substances (TBARS), this qualitative method enables in situ visualization of lipid peroxidation without tissue homogenization, providing a rapid and biologically informative screening tool for assessing ferroptosis-associated oxidative damage in Cx. quinquefasciatus and other biological models exposed to multiple stressors.

To study gene function in regulating rodent retinal waves during development, an efficient method for gene delivery into whole-mount retinas is required while preserving circuit functionality for physiological studies. We present an optimized electroporation protocol for developing rodent retinal explants. The procedure includes the fabrication of horizontally aligned platinum electrodes and the placement of retinal explants between them to generate a uniform electric field for high transfection efficiency. The entire process—dissection and electroporation—can be completed within 1–2 h. Successful transfection is verified by fluorescence microscopy, and physiological assays such as patch-clamp recordings and live imaging can be performed within 1–4 days following electroporation. This rapid and reliable protocol enables functional analysis for a specific gene in regulating retinal waves and can be adapted to other organotypic slice cultures.

The study of whole organs or tissues and their cellular components and structures has been historically limited by their natural opacity, which is caused by the optical heterogeneity of the tissue components that scatter light as it traverses through the tissue, making 3D tissue imaging highly challenging. In recent years, tissue clearing techniques have received widespread attention and undergone rapid development. We recently demonstrated the synthesis of a 2-hydroxyethyl methacrylate (HEMA)-acrylamide (AAm) copolymer. This was achieved using antipyrine (ATP) and 2,2′-thiodiethanol (TDE) as solvents. The resulting solution rapidly embedded tissue samples with a high degree of transparency and is compatible with multiple fluorescence labeling techniques. The method exhibits significant transparency effects across a range of organs, comprising the heart, liver, spleen, lung, kidney, brain (whole and sectioned), esophagus, and small intestine. It can enable volumetric imaging of tissue up to the scale of mouse organs, decrease the duration of the clearing, and preserve emission from fluorescent proteins and dyes. To facilitate the use of this powerful tool, we have provided here a detailed step-by-step protocol that should allow any laboratory to use tissue transparency technology to achieve transparency of tissues and organs.

Intestinal glucose absorption has been studied for several decades. However, the different methods available for investigating absorption are often the reason for variability in the results, and it is difficult to measure the relative contribution of paracellular absorption using existing methods. Thus, we have established a new model for measuring glucose absorption. In the isolated in situ vascularly perfused small intestine, the intestinal epithelium is completely preserved, and the entire transport pathway is intact. In the present model, we use radioactive labeled ¹⁴C-d-glucose, which allows for sensitive quantification of glucose absorption even with low luminal concentrations. The described method is optimized for intestinal glucose absorption but can be applied to other macro/micronutrients that can be radioactively labeled. The described procedure is a novel approach for measurements of intestinal nutrient absorption and gut permeability in which luminal nutrient concentrations resemble physiological concentrations.

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.

Inherited germline variants are now recognized as important contributors to hematologic myeloid malignancies, but their reliable detection depends on obtaining uncontaminated germline DNA. In solid tumors, peripheral blood remains free of tumor cells and therefore serves as a standard source for germline testing. In contrast, peripheral blood often contains neoplastic or clonally mutated cells in hematologic malignancies, making it impossible to distinguish somatic from germline variants. This unique challenge necessitates using an alternative, non-hematopoietic tissue source for accurate germline assessment in patients with hematologic myeloid malignancies. Cultured skin fibroblasts derived from punch biopsies have long been considered the gold standard for this purpose. Nevertheless, most existing protocols are optimized for research settings and lack detailed, patient-centric workflows for routine clinical use. Addressing this translational gap, we present a robust, enzyme-free protocol for culturing dermal fibroblasts from skin punch biopsies collected at the bedside during routine bone marrow procedures. The method details practical bedside collection, sterile transport, mechanical dissection without enzymatic digestion, plating strategy, culture expansion, and high-yield DNA isolation with validated purity. By integrating this standardized approach into routine hematopathology workflows, the protocol ensures reliable germline material with minimal patient discomfort and a turnaround time suitable for clinical diagnostics.

Cell transplantation is a promising strategy for treating age-related muscle atrophy, but its critical application remains limited. Cultured myoblasts, unlike freshly isolated muscle stem cells, show poor engraftment efficiency and fail to contribute effectively to muscle regeneration. Moreover, successful engraftment generally requires prior muscle injury, as skeletal muscle regeneration is typically triggered by a damaged microenvironment. These limitations present major obstacles for applying cell therapy to sarcopenia, where muscle degeneration occurs without injury. In this protocol, we describe a novel approach that enables the transplantation of cultured myoblasts into intact skeletal muscle without the need for preexisting injuries or genetic modification. By combining myoblasts with extracellular matrices (ECM), such as Matrigel, which mimic the native muscle niche and support cell survival, adhesion, proliferation, and differentiation, we achieve efficient engraftment and increased muscle mass without the need for preexisting injury. The ECM also provides a scaffold and retains bioactive factors that enhance the regenerative capacity of transplanted cells. This is the first protocol that enables robust myoblast engraftment in non-injury muscle conditions, offering a practical tool for studying and potentially treating sarcopenia.

Proper brain function depends on the integrity of the blood–brain barrier (BBB), which is formed by a specialized network of microvessels in the brain. Reliable isolation of these microvessels is crucial for studying BBB composition and function in both health and disease. Here, we describe a protocol for the mechanical dissociation and density-based separation of microvessels from fresh or frozen human and murine brain tissue. The isolated microvessels retain their molecular integrity and are compatible with downstream applications, including fluorescence imaging and biochemical analyses. This method enables direct comparisons across species and disease states using the same workflow, facilitating translational research on BBB biology.

Osteoarthritis (OA) is the primary cause of joint impairment, particularly in the knee. The prevalence of OA has significantly increased, with knee OA being a major contributor whose pathogenesis remains unknown. Articular cartilage and the synovium play critical roles in OA, but extracting high-quality RNA from these tissues is challenging because of the high extracellular matrix content and low cellularity. This study aimed to identify the most suitable RNA isolation method for obtaining high-quality RNA from microquantities of guinea pig cartilage and synovial tissues, a relevant model for idiopathic OA. We compared the traditional TRIzol^® method with modifications to spin column–based methods (TRIspin-TRIzol^®/RNeasy^TM, RNeasy^TM kit, RNAqueous^TM kit, and Quick-RNA^TM Miniprep Plus kit), and an optimized RNA isolation protocol was developed to increase RNA yield and purity. The procedure involved meticulous sample collection, specialized tissue processing, and measures to minimize RNA degradation. RNA quality was assessed via spectrophotometry and RT–qPCR. The results demonstrated that among the tested methods, the Quick-RNA^TM Miniprep Plus kit with proteinase K treatment yielded the highest RNA purity, with A_260:280 ratios ranging from 1.9 to 2.0 and A_260:230 ratios between 1.6 and 2.0, indicating minimal to no salt contamination and RNA concentrations up to 240 ng/μL from ⁓20 mg of tissue. The preparation, storage, homogenization process, and choice of RNA isolation method are all critical factors in obtaining high-purity RNA from guinea pig cartilage and synovial tissues. Our developed protocol significantly enhances RNA quality and purity from micro-quantities of tissue, making it particularly effective for RTqPCR in resource-limited settings. Further refinements can potentially increase RNA yield and purity, but this protocol facilitates accurate gene expression analyses, contributing to a better understanding of OA pathogenesis and the development of therapeutic strategies.