Cell Biology

Acid ceramidase (aCDase) is a lysosomal amidase that catalyzes the hydrolysis of sphingolipids (SphL), including ceramides and glucosylceramides. Altered expressions of aCDase are associated with several pathological conditions, such as cancer, inflammation, pain, and pulmonary disorders. aCDase activity is reduced in Farber disease, spinal muscular atrophy with progressive myoclonic epilepsy, diabetes, and cardiovascular disease. Recent reports suggest that aCDase inhibition may be an emerging strategy for treating several SphL-related neurodegenerative conditions, such as Krabbe, Gaucher, and Parkinson’s disease, due to its role in the accumulation of glycosphingolipids. Therefore, the development of a tissue-based aCDase activity assay has potential applications in clinical diagnostics and drug discovery, enabling the evaluation of the onset and progression of disease from biological samples of patients, drug-target engagement analysis, and identification of biomarkers. Here, we report a detailed protocol for detecting aCDase activity in tissue lysates, using Rbm14-12 as a specific fluorogenic substrate for aCDase. Assay protocol optimization, including a procedure for the preparation and storage of tissue lysates and the identification of optimal protein tissue lysate amounts and substrate concentrations based on kinetic enzymatic parameter analyses, is described.

Calcium signaling is a universal, versatile process in which ionized or free calcium (Ca²⁺) acts as a second messenger to regulate various cellular activities, including hormone secretion, contraction, proliferation, gene expression, and apoptosis. Changes in the cytoplasmic free Ca²⁺ concentration ([Ca²⁺]_cyt) in hepatocytes play a central role in mediating the actions of insulin, glucagon, catecholamines, and other hormones on carbohydrate, lipid, and protein metabolism in the liver. Ratiometric chemical Ca²⁺ indicators are fluorescent dyes that change their emission or excitation spectrum upon binding to calcium, allowing for precise, quantitative measurements of changes in the intracellular Ca²⁺ concentration. They enable calibration by calculating the ratio of two fluorescence intensities, correcting for artifacts such as uneven dye loading, photobleaching, and cell volume variations. Fura-2 acetoxymethyl ester (AM) (hereinafter referred to as Fura-2), a ratiometric and sensitive indicator dye, is a popular fluorescent Ca²⁺ reporter for measuring intracellular calcium. Here, we describe a comprehensive and detailed protocol for Ca²⁺ imaging of the H4IIE cell line and primary rodent hepatocytes in vitro via the chemical reporter Fura-2, which can also be employed on a wide variety of cell types. Unlike previously published protocols, this protocol addresses the challenge of facilitating the attachment of liver cell lines and primary hepatocytes to glass coverslips for imaging using an inverted fluorescence microscope. Our protocol describes two different loading/labeling strategies for Fura-2 dye: one is cost-effective but requires skillful pipettor handling, and the second one is easy but expensive as it needs a large volume of Krebs-Ringer HEPES (KRH)-Fura-2 solution. If the coverslips are handled properly, the cost-effective coverslip-only loading approach produces similar quality results as the large volume method. Finally, we describe a simple and user-friendly procedure to analyze Ca²⁺ signals over time using Microsoft Excel’s functional equations.

Ornithine decarboxylase (ODC) is a rate-limiting enzyme in polyamine biosynthesis that plays a critical role in cell proliferation and tumorigenesis. Reliable quantification of ODC activity is essential for mechanistic and therapeutic studies. Traditional assays often rely on radiolabeled substrates or discontinuous endpoint measurements. Here, we describe a non-radioactive, continuous spectrophotometric assay for measuring ODC activity in cell lysates using a commercially available liquid-stable CO₂ detection reagent. In this assay, CO₂ generated by ODC is captured as bicarbonate and utilized in a coupled enzymatic system containing phosphoenolpyruvate carboxylase (PEPC) and malate dehydrogenase (MDH), leading to oxidation of thio-NADH. The decrease in absorbance at 405 nm due to thio-NADH oxidation is monitored in real time and is proportional to ODC activity. The protocol is performed in a 96-well plate format, requires minimal reagent preparation, and is suitable for medium- to high-throughput applications.

Surface receptor engagement governs T-cell activation. Since these surface receptors undergo extensive glycosylation, lectin-mediated crosslinking of these glycosylated surface receptors has the potential to modulate signaling. Here, we systematically evaluate the abilities of recombinant human galectins in triggering immune responses. We describe how to apply the human galectins to modulate Jurkat E6-1 cell activation by measuring the expression level of cellular surface CD69 and the mRNA of IL-2. To validate the protocol, we confirmed that galectin-3 and galectin-8 variants 1 and 2 reproducibly induce CD69 and IL-2 expression on Jurkat E6-1 cells. Our approach offers a galectin-based toolset to study how glycosylation modulates human adaptive immunity.

Progressive neurodegeneration linked to the accumulation of misfolded proteins is a hallmark of several neurodegenerative disorders, including Parkinson’s disease, Huntington’s disease, and Alzheimer’s disease. Dysfunction in the protein homeostasis machinery correlates with pathology. The chaperone protein DNAJB6 is expressed in neurons and oligodendrocytes and has been shown to play a key role in preventing amyloid aggregation by binding to amyloidogenic proteins and facilitating their refolding or degradation, in cooperation with other chaperones. Here, we describe a simple and feasible assay that enables high-throughput screening for DNAJB6 activity in a plate reader format. We use genetically engineered HEK293 cells that stably express DNAJB6 fused to either CFP or YFP. These cells can be plated into multi-well plates, and the fluorescence resonance energy transfer (FRET) signal can be measured for analysis of DNAJB6 dimerization, which is linked to DNAJB6 activity. The protocol can be used for drug screening and to identify compounds that increase DNAJB6 dimerization, and can serve as a starting point for finding new medicines that act through modulating DNAJB6 activity.

Isolation of adult mouse ventricular myocytes is essential for studying cardiac physiology and cellular function. Traditional methods commonly rely on Langendorff perfusion systems, which provide continuous retrograde coronary perfusion but require specialized equipment and can be complex to operate. Here, we describe a simplified Langendorff-based protocol that uses a syringe pump–driven system to achieve constant-flow retrograde aortic perfusion during enzymatic digestion. The setup incorporates an inline heater for precise temperature control and uses widely available laboratory components, enabling consistent delivery of digestion enzymes. This approach maintains stable perfusion despite changes in coronary resistance and reduces variability associated with conventional gravity-driven systems. The protocol yields high-quality adult ventricular myocytes suitable for downstream functional analyses, including electrophysiology, contractility, and calcium imaging. Compared with traditional systems, this method is more accessible, reduces technical complexity, and improves reproducibility, facilitating adoption in laboratories without dedicated isolated-heart perfusion infrastructure.

In vitro cytotoxicity assessments frequently rely on staining-based methods that indirectly estimate viable cell numbers. A major limitation of many such techniques is their endpoint nature, requiring cell lysis or irreversible processing that precludes longitudinal monitoring of cellular responses following treatment. An ideal assay for evaluating cell viability and proliferation should be simple, rapid, cost-effective, reproducible, and highly sensitive, while also enabling accurate quantification with minimal interference from test compounds. The resazurin reduction assay satisfies these criteria, offering a sensitive and economical alternative to conventional tetrazolium-based methods. Although both assay types depend on the metabolic reduction of a dye by viable cells, they differ mechanistically. Tetrazolium salts (e.g., MTT) are reduced by cellular dehydrogenases to insoluble formazan crystals that require solubilization before detection. In contrast, resazurin-a cell-permeable, non-fluorescent blue dye-is reduced to resorufin, a highly fluorescent compound detectable without additional processing steps. This property renders the resazurin assay broadly applicable to viability testing in eukaryotic cells cultured in both 2D and 3D formats, as well as in bacterial systems. Here, we present a resazurin-based reduction assay across diverse experimental models, emphasizing its practicality, reproducibility, and adaptability for real-time viability monitoring.

The binding of transmembrane (TM) ligands to their cognate TM receptors on neighboring cells governs intercellular adhesion and direct cell–cell communication. However, these interactions are difficult to study in vitro because they depend on membrane presentation, ligand orientation, receptor clustering, and avidity, features often not captured by soluble recombinant ligands or cell-free assays. Here, we describe a flow cytometry–based assay using fluorescent, lentiviral virus-like particles (VLPs) displaying TM ligands to quantify binding to their receptors on target cells. Fluorescent VLPs are generated in-house by plasmid transfection in HEK293T cells and enable direct fluorescent detection without fluorochrome-conjugated secondary antibodies. The system is modular and readily accommodates engineered ligand constructs, including patient-derived variants. We applied this platform to generate ICAM-1-displaying fluorescent VLPs and to study human LFA-1 function in patient-derived leukocytes. This protocol provides a detailed workflow for VLP production and in vitro binding assays, offering a simple, quantitative, and cost-effective approach for studying TM ligand–receptor interactions in a membrane context. The system is well-suited for mechanistic studies, functional assessment of patient-derived variants, and direct binding assays using patient-derived cells. Integrating the assay into multicolor flow cytometry panels enables simultaneous immunophenotyping and quantification of up to four ligand–receptor interactions at single-cell resolution.

Anoikis resistance, or the ability of cancer cells to evade cell death triggered by immediate detachment from the extracellular matrix, is a critical established hallmark of metastatic cancer. While suspension culture models have been used to study anoikis, most focus on defined single time points or prolonged suspension that may not recapitulate the effects of repeated stress that tumor cells experience during metastatic dissemination. Here, we describe a detailed protocol for generating anoikis-resistant (AnR) cancer cells that have adapted to such stress through exposure to repeated cycles of suspension stress on poly-HEMA-coated plates, followed by recovery under standard attached conditions. The protocol includes methods for determining baseline anoikis sensitivity, generating AnR cells over 7–9 attachment-detachment cycles, assessing the stability and reversion of the anoikis-resistant phenotype, and characterizing AnR cells using Live/Dead staining of spheroids, flow cytometry–based apoptosis assays, and immunofluorescence for proliferation markers. This approach produces a non-genetic, reversible anoikis-resistant state that models the adaptive transcriptional reprogramming underlying metastatic progression, providing a reproducible and physiologically relevant in vitro system for studying anoikis resistance mechanisms and evaluating therapeutic strategies for prevention and reversal of such adaptations.

Antibody therapeutics have demonstrated transformative impacts on improving the quality of life of millions of patients, whereas advances in antibody discovery technologies have imposed a significant production challenge for the generation of a large diversity of therapeutic antibody candidates. A demand for the rapid production of dozens of purified antibodies in 10-mg quantities is entailed for functional screening and molecular assessment studies. Here, we present a robust semi-automated production protocol that bridges the gap between miniaturized high-throughput screenings and conventional custom-scale workflows. This methodology and workflow utilize a simple high-titer transient Chinese hamster ovary (CHO) cell host–CHO4Tx^® expression system, a procedure of magnetic protein-A bead in-culture antibody capturing, and a semi-automated purification process with the GenScript AmMag^TM SA Plus system. This production protocol has been proven to be robust and valuable for the routine production of dozens of antibody constructs per week in sufficient quality and quantity for cell-based and biophysical studies.