Cell Biology

The placenta is a metabolically active organ whose mitochondrial activity is tightly linked to fetal growth, oxygenation, and nutrient transport, mediating fetal susceptibility to environmental exposures. Accordingly, aberrant mitochondrial function has been implicated in the progression of placental dysfunction. However, existing respirometry platforms require primarily fresh or cryopreserved placental tissue and offer limited throughput, rendering these platforms impractical in the context of large-scale placental dissections. Here, we describe and validate a Seahorse XF approach for measuring mitochondrial respiration in previously frozen placentae, enabling the functional interrogation of placental mitochondria in prenatal studies. Our protocol fundamentally relies on the restoration of matrix substrates that are depleted due to increased mitochondrial membrane permeability following freeze-thaw cycles. We provide a strategy to assess complex I and II-associated respiration adapted for the Seahorse XFe24 Analyzer and further demonstrate comparable oxygen consumption readouts between fresh and frozen placentae. We further demonstrate distinct differences in the magnitude of oxygen consumption between fresh and frozen placentae in the absence of exogenous NADH. Taken together, we present a simplified and convenient protocol for the assessment of respiratory enzyme complex-associated respiration from archived placental tissue.

Manganese (Mn) is an essential trace element whose intracellular homeostasis is tightly controlled by specialized membrane transporters. Dysregulation of Mn transport leads to pathological Mn accumulation and severe human disease; however, efficient and quantitative cell-based methods for assessing Mn²⁺ transporter activity remain limited. Here, we present an optimized cellular Fura-2 manganese extraction assay (CFMEA) that enables robust quantification of cellular Mn content and provides a normalized framework for assessing relative Mn²⁺ transport activity in a high-throughput format. This protocol integrates Fura-2-based fluorescence detection of Mn²⁺ at the Ca²⁺ isosbestic excitation wavelength with dsDNA quantification to normalize dsDNA levels in cell extracts and immunoblotting to account for transporter protein expression levels. Cells expressing Mn²⁺ transporters are exposed to MnCl₂ in 96-well plates, washed to remove extracellular Mn²⁺, and lysed in a Fura-2-containing extraction buffer. Fluorescence quenched by Mn²⁺ is quantified and converted to cellular Mn content using a cell-free Mn-Fura-2 standard curve and then normalized to dsDNA content and protein abundance to determine relative transporter activity. This workflow provides a relatively sensitive, reproducible, and low-cost approach for comparative analysis of Mn²⁺ transporters and their variants across multiple cell types. The protocol is demonstrated using the Mn²⁺ efflux transporter SLC30A10 in HEK293T cells and is readily adaptable for studying other Mn²⁺ transport pathways.

Epigenetic modifications play essential roles in regulating gene expression and maintaining cellular identity. Accumulating evidence suggests that chemical agents can contribute to carcinogenesis through epigenetic alterations, such as changes in DNA methylation and histone modifications, even in the absence of direct DNA damage. Here, we have developed a simple, cost-effective, and quantitative reporter assay, termed the epi-TK assay, to evaluate chemically induced epigenetic alterations. The assay is built upon the thymidine kinase (TK) gene mutation assay, a standardized and widely used in vitro genotoxicity assay for chemical safety evaluation. This system is based on an engineered human lymphoblastoid cell line (mTK6), in which the promoter region of the endogenous housekeeping TK gene is site-specifically methylated using epigenome-editing technology, resulting in stable transcriptional repression. Following chemical exposure, epigenetic perturbations at the TK locus are detected by culturing cells under hypoxanthine–aminopterin–thymidine selection and quantifying the frequency of TK revertant colonies, which reflects restoration of TK gene expression. Using the DNA methyltransferase 1 inhibitor GSK3484862 as a model compound, this protocol demonstrates that the epi-TK assay enables sensitive and quantitative detection of epigenetic state transitions. Importantly, this assay allows bi-directional detection of epigenetic changes, including DNA demethylation events and broader alterations in histone modification landscapes. Together, the epi-TK assay provides a practical and quantitative platform for evaluating epigenetic toxicity, with potential applications in chemical safety assessment frameworks.

The integrity of epithelial barriers is essential for maintaining tissue homeostasis, particularly in the intestinal tract, where it separates the host from the complex luminal environment. Two complementary, standard methods for assessing this barrier are transepithelial electrical resistance (TEER), which provides a rapid, non-destructive measure of ionic conductance across tight junctions, and the fluorescein isothiocyanate (FITC)-dextran assay, which directly quantifies paracellular macromolecule flux. This protocol details a robust and reproducible method for performing both assays using intestinal epithelial cell monolayers (e.g., Caco-2, T84) cultured on permeable Transwell supports. We outline the procedure from cell culture and monolayer differentiation to TEER measurement with an Epithelial Volt/Ohm Meter 3 (EVOM3) and the subsequent FITC-dextran permeability assay. By combining these techniques, this protocol provides a comprehensive assessment of barrier function, making it ideal for studying tight junction dynamics and regulation under various experimental conditions, such as cytokine stimulation, drug screening, or microbial challenges.

Small ubiquitin-related modifiers (SUMOs) are covalently conjugated onto the proteome and serve as signaling molecules in many aspects of eukaryotic cell biology, from S. cerevisiae and C. elegans to H. sapiens. The conjugatable SUMO variants, SUMO1 and the almost identical SUMO2 and SUMO3 (designated SUMO2/3), are processed by an E1(SAE1:SAE2)-E2(UBC9)-E3 enzyme cascade to produce SUMO-modified proteins. The prerogative of the SUMO biology field is to identify and study the specific proteins undergoing SUMOylation, which grants us insights into the biological pathway of interest. This protocol was developed using the human osteosarcoma cell line U2OS to enable the investigation of SUMO conjugates in mitosis, the cell division phase of the cell cycle. We enrich the cell population for mitotic cells, which are isolated and subjected to stringent lysis conditions involving a high concentration of SDS and DTT in RIPA buffer, to promote complete protein denaturation. The lysates in high SDS RIPA buffer are diluted to reduce the overall SDS concentration and undergo conventional immunoprecipitation using SUMO1- or SUMO2/3-specific antibodies bound to protein A/G agarose beads. The samples are then compatible with downstream readouts such as western blots and mass spectrometry. This protocol detects endogenous SUMOylated proteins and avoids exogenous SUMO overexpression, which can alter SUMO conjugate formation. Furthermore, this denaturing protocol ensures only SUMOylated proteins are immunoprecipitated, and not their interactors.

Extracellular vesicles (EVs) are critical mediators of cell–cell communication and play a key role in male reproductive biology by modulating sperm function. This protocol describes a robust and reproducible workflow for isolating EVs from ram seminal plasma using size-exclusion chromatography (SEC) and assessing their uptake by ram spermatozoa. In contrast to ultracentrifugation-based methods, SEC provides a gentle and more efficient isolation approach that preserves EV integrity and functionality. A central innovation of this protocol is the use of carboxyfluorescein succinimidyl ester (CFSE)-labeled seminal plasma EVs (SP-EVs) to evaluate their incorporation into sperm cells through two complementary detection platforms: (i) flow cytometry with standard resolution and (ii) confocal microscopy, for spatial confirmation of EV–sperm interactions. By bridging the gap between EV isolation and functional analysis, this protocol provides a valuable tool for investigating the role of EV–cell interactions. Specifically, it offers potential applications in male fertility preservation, biomarker discovery, and the development of EV-based therapeutic strategies in reproductive medicine.

Endocytosis is an essential membrane transport mechanism that is indispensable for the maintenance of life. It is responsible for the selective internalization and subsequent degradation or recycling of specific extracellular proteins and nutrients, thereby facilitating cellular nutrient supply, modulation of receptor signaling, and clearance of foreign substances. However, methods for the quantitative analysis of lysosomal degradation of extracellular proteins via endocytosis remain limited. This protocol describes a method for purifying the protein-of-interest (POI)–red fluorescent protein (RFP)–green fluorescent protein (GFP) fusion protein, which is modified with specific mammalian cell glycans or other modifications, from the conditioned medium of mammalian cell cultures. Subsequently, the protocol details a quantitative approach for evaluating its internalization and lysosomal degradation within cells using the RFP–GFP tandem fluorescent reporter. Following the addition of POI-RFP-GFP to the medium, cells can be subjected to cell biological assays, such as flow cytometry, as well as biochemical analyses, such as immunoblotting. This protocol is broadly applicable to studies of the internalization of extracellular proteins.

Organelle abundance is a key microscopic readout of organelle formation and, in many cases, function. Quantification of organelle abundance using confocal microscopy requires estimating their area based on the fluorescence intensity of compartment-specific markers. This analysis usually depends on a user-defined intensity threshold to distinguish organelle regions from the surrounding cytoplasm, which introduces potential bias and variability. To address this issue, we present a machine learning–assisted algorithm that allows for the quantification of organelle density using the open-source Fiji platform and WEKA segmentation. Our method enables the automated quantification of organelle number, area, and density by learning from training data. This standardizes threshold selection and minimizes user intervention. We demonstrate the utility of this approach for both membrane and non-membrane organelles, such as peroxisomes, lipid droplets, and stress granules, in human cells and whole fish samples.

Obesity is a risk factor for many diseases. The 3T3-L1 cell line is often used to obtain mature adipocytes, but these lack the inflammatory phenotype observed in obesity. Using a cocktail of cytokines that mimics the secretome of macrophages found in the inflammatory adipose tissue, we developed a protocol for obtaining mature inflammatory adipocytes. This model was validated at gene (RT-qPCR) and protein levels (multiplex adipokine array) as we found a decrease of adipogenic markers (C/EBPα, PPARУ, adiponectin, and CD36) and an increase of pro-inflammatory cytokines (IL-6, IL-1β, CXCL1, CXCL10, TNF-α, ICAM-1, and lipocalin-2). We provide a relevant in vitro model for studying the impact of low-grade chronic inflammation caused by obesity and its downstream effects on metabolic disorders and tumor microenvironments.

Transfecting neurons remains technically challenging due to their sensitivity. Conventional methods, such as Lipofectamine 2000 or Lipofectamine RNAiMAX, often result in significant cytotoxicity, which limits their utility. Although lentiviral transfection offers high efficiency, it is hindered by high costs and complex procedures. This experiment employs a small interfering RNA (siRNA)-specific transfection reagent from the Kermey company. This reagent is a novel nanoparticle-based lipid material designed for the efficient delivery of oligonucleotides, including siRNA, into a wide range of cell types. Its efficacy in achieving high transfection efficiency in neurons, however, has not yet been established. After several days of in vitro neuronal culture, researchers can perform a simple transfection procedure using this reagent to achieve robust transfection efficiency. Notably, the protocol does not require medium replacement 6–8 h post-transfection, streamlining the workflow and minimizing cellular stress.