Cell Biology

The antibody-uptake assay is a commonly used technique to monitor endocytosis of integral membrane proteins including transmembrane and glycosylphosphatidylinositol-anchored proteins (GPI-APs). The antibody-uptake assay typically involves incubating live cells with fluorophore-conjugated antibodies directed against the extracellular domain of the integral membrane protein of interest. Antibody uptake is then detected by flow cytometry or confocal microscopy. However, these detection modalities may be inaccessible to some labs or require extensive training to operate. Thus, we developed an easy and novel sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and western blot-based approach to the antibody-uptake assay that exploits the strong affinity between biotin and streptavidin. Instead of incubating cells with fluorophore-conjugated antibodies to monitor antibody uptake, our assay involves incubating cells with biotinylated antibodies, processing the cell lysates for western blot, and probing the membrane with detectably conjugated streptavidin. From preparation to quantification, this protocol requires less hands-on time than other approaches and is amenable to small-scale drug or siRNA screens. Here, we demonstrate the utility of our approach using the well-characterized misfolded GPI-AP, YFP-tagged C179A mutant of prion protein (YFP-PrP*), as our model substrate. YFP-PrP* constitutively traffics to the plasma membrane (PM), where it binds to anti-GFP antibody, and immediately undergoes endocytosis to lysosomes. To validate our protocol, we present measurements of antibody uptake under conditions known to enhance or inhibit YFP-PrP*’s traffic to the PM. Using this assay, we present new evidence that, under certain conditions, YFP-PrP* is able to undergo degradation via a pathway that does not involve exposure on the cell surface.

Proteomics analysis is crucial for understanding the molecular mechanisms underlying muscle adaptations to different types of exercise, such as concentric and eccentric training. Traditional methods like two-dimensional gel electrophoresis and standard mass spectrometry have been used to analyze muscle protein content and modifications. This protocol details the preparation of muscle samples for proteomics analysis using ultra-high-performance liquid chromatography (UHPLC). It includes steps for muscle biopsy collection, protein extraction, digestion, and UHPLC-based analysis. The UHPLC method offers high-resolution separation of complex protein mixtures, providing more detailed and accurate proteomic profiles compared to conventional techniques. This protocol significantly enhances sensitivity, reproducibility, and efficiency, making it ideal for comprehensive muscle proteomics studies.

Apolipoprotein B (APOB) is the primary structural protein of atherogenic lipoproteins, which drive atherogenesis and thereby lead to deadly cardiovascular diseases (CVDs). Plasma levels of APOB-containing lipoproteins are tightly modulated by LDL receptor–mediated endocytic trafficking and cargo receptor–initiated exocytic route; the latter is much less well understood. This protocol aims to present an uncomplicated yet effective method for detecting APOB/lipoprotein secretion. We perform primary mouse hepatocyte isolation and culture coupled with well-established techniques such as immunoblotting for highly sensitive, specific, and semi-quantitative analysis of the lipoprotein secretion process. Its inherent simplicity facilitates ease of operation, rendering it a valuable tool widely utilized to explore the intricate landscape of cellular lipid metabolism and unravel the mechanistic complexities underlying lipoprotein-related diseases.

Enteroendocrine cells (EECs) are known chemosensors in the gastrointestinal (GI) epithelium. They release a diversity of gut hormones in response to various stimuli. Here, we report an in-vitro assay to measure GLP-1 release from cultured murine EEC’s under fatty acid stimulation.

The mucosal surfaces of the gastrointestinal, respiratory, reproductive, and urinary tracts, and the surface of the eye harbor a resident microflora that lives in symbiosis with their host and forms a complex ecosystem. The protection of the vulnerable epithelium is primarily achieved by mucins that form a gel-like structure adherent to the apical cell surface. This mucus layer constitutes a physical and chemical barrier between the microbial flora and the underlying epithelium. Mucus is critical to the maintenance of a homeostatic relationship between the microbiota and its host. Subtle deviations from this dynamic interaction may result in major implications for health. The protocol in this article describes the procedures to grow low mucus-producing HT29 and high mucus-producing HT29-MTX-E12 cells, maintain cells and use them for mucus quantification by ELISA. Additionally, it is described how to assess the amount of secreted adherent mucus. This system can be used to study the protective effect of mucus, e.g., against bacterial toxins, to test the effect of different culture conditions on mucus production or to analyze diffusion of molecules through the mucus layer. Since the ELISA used in this protocol is available for different species and mucus proteins, also other cell types can be used.

Here we describe two assays to measure dense core vesicle (DCV) exocytosis-mediated cargo secretion in neuroendocrine cells. To conduct siRNA screens for novel genes in regulated DCV exocytosis, we developed a plate reader-based secretion assay using DCV cargo, NPY-Venus, and an orthogonal ³H-serotonin secretion assay. The NPY-Venus secretion assay was successfully used for a high throughput siRNA screen, and the serotonin secretion assay was used to validate hits identified from the screen (Sorensen, 2017; Zhang et al., 2017).

Quantitative analysis of proteins secreted from the cells poses a challenge due to their low abundance and the interfering presence of a large amount of bovine serum albumin (BSA) in the cell culture media. We established assays for detection of mutant huntingtin (mHtt) secreted from Neuro2A cell line stably expressing mHtt and rat primary cortical neurons by Western blotting. Our protocol is based on reducing the amounts of BSA in the media while maintaining cell viability and secretory potential, and concentrating the media prior to analysis by means of ultrafiltration.

Interferon-gamma (IFN-γ) is crucial for immunity against intracellular pathogens and for tumor control. It is produced predominantly by natural killer (NK) and natural killer T cells (NKT) as well as by antigen-specific Th1 CD4⁺ and CD8⁺ effector T cells. When investigating immune responses against pathogens and cancer cells, measuring antigen-specific cytokine-responses by cells of adaptive immunity offers an advantage over total non-specific cytokine responses. Significantly, the measurement of antigen-specific IFN-γ responses against pathogens or cancer cells, when compared to a treatment group, provides a quantitative measure of how well the treatment works. Measuring antigen-specific IFN-γ responses involves culture of the cells being considered (CD4⁺ or CD8⁺ T cells) with antigen presenting cells (APC) and a specific peptide from the target pathogen or cancer cell compared to control cultures without a peptide. After a suitable timeframe, the cytokine released is measured by an ELISPOT assay. The difference in the number of cells secreting IFN-γ, with and without peptide, is a measure of antigen-specific IFN-γ responses. This assay can be applied to other cytokines such as IL-10.