细胞生物学

Macrophages are at the center of innate immunity and iron metabolism. In the case of an infection, macrophages adapt their cellular iron metabolism to deprive iron from invading bacteria to combat intracellular bacterial proliferation. A concise evaluation of the cellular iron content upon an infection with bacterial pathogens and diverse cellular stimuli is necessary to identify underlying mechanisms concerning iron homeostasis in macrophages. For the characterization of cellular iron levels during infection, we established an in vitro infection model where the murine macrophage cell line J774A.1 is infected with Salmonella enterica serovar Typhimurium (S.tm), the mouse counterpart to S. enterica serovar Typhi, under normal and iron-overload conditions using ferric chloride (FeCl₃) treatment. To evaluate the effect of infection and iron stimulation on cellular iron levels, the macrophages are stained with FerroOrange. This fluorescent probe specifically detects Fe²⁺ ions and its fluorescence can be quantified photometrically in a plate reader. Importantly, FerroOrange fluorescence does not increase with chelated iron or other bivalent metal ions. In this protocol, we present a simple and reliable method to quantify cellular Fe²⁺ levels in cultured macrophages by applying a highly specific fluorescence probe (FerroOrange) in a TECAN Spark microplate reader. Compared to already established techniques, our protocol allows assessing cellular iron levels in innate immune cells without the use of radioactive iron isotopes or extensive sample preparation, exposing the cells to stress.

Key features

• Easy quantification of Fe²⁺ in cultured macrophages with a fluorescent probe.

• Analysis of iron in living cells without the need for fixation.

• Performed on a plate reader capable of 540 nm excitation and 585 nm emission by trained employees for handling biosafety level 2 bacteria.

Graphical overview

The mitochondrial electron transport chain (ETC) is a multi-component pathway that mediates the transfer of electrons from metabolic reactions that occur in the mitochondrion to molecular oxygen (O₂). The ETC contributes to numerous cellular processes, including the generation of cellular ATP through oxidative phosphorylation, serving as an electron sink for metabolic pathways such as de novo pyrimidine biosynthesis and for maintaining mitochondrial membrane potential. Proper functioning of the mitochondrial ETC is necessary for the growth and survival of apicomplexan parasites including Plasmodium falciparum, a causative agent of malaria. The mitochondrial ETC of P. falciparum is an attractive target for antimalarial drugs, due to its essentiality and its differences from the mammalian ETC. To identify novel P. falciparum ETC inhibitors, we have established a real-time assay to assess ETC function, which we describe here. This approach measures the O₂ consumption rate (OCR) of permeabilized P. falciparum parasites using a Seahorse XFe96 flux analyzer and can be used to screen compound libraries for the identification of ETC inhibitors and, in part, to determine the targets of those inhibitors.

Key features

• With this protocol, the effects of candidate inhibitors on mitochondrial O₂ consumption in permeabilized asexual P. falciparum parasites can be tested in real time.

• Through the sequential injection of inhibitors and substrates into the assay, the molecular targets of candidate inhibitors in the ETC can, in part, be determined.

• The assay is applicable for both drug discovery approaches and enquiries into a fundamental aspect of parasite mitochondrial biology.

Graphical overview

Seahorse assay experimental workflow. Prior to the assay, coat the cell culture microplate with Cell-Tak to help adhere the parasites to the wells; hydrate the cartridge wells to ensure proper sensor functionality and design the assay template using the Agilent Seahorse Wave Desktop software (Analyze Seahorse data files, Seahorse Wave desktop software|Agilent). On the day of the assay, prepare the inhibitors/substrates that are to be injected into the ports. Then, separate 3 × 10⁸ trophozoite-stage parasites from the uninfected red blood cells (RBCs) and ring-stage parasites using a MACS^® magnetic column. Check the purity of the parasites with Giemsa-stained smears. Determine the concentration of infected RBCs in the sample using a hemocytometer and dilute to approximately 5 × 10⁷ parasites per milliliter. Treat infected RBCs with saponin to permeabilize the host cell membrane and seed approximately 5 × 10⁶ parasites (100 μL) per well in mitochondria assay solution (MAS) buffer. Supplement MAS buffer with digitonin to permeabilize the parasite plasma membrane. Load the ports with the prepared inhibitors/substrates and run the assay using a Seahorse XFe96 analyzer. Once the assay is completed, analyze the data using the Wave desktop software. Further data processing can be done using statistical analysis software.

Lysine acetylation is a conserved post-translational modification and a key regulatory mechanism for various cellular processes, including metabolic control, epigenetic regulation, and cellular signaling transduction. Recent advances in mass spectrometry (MS) enable the extensive identification of acetylated lysine residues of histone and non-histone proteins. However, protein enrichment before MS analysis may be necessary to improve the detection of low-abundant proteins or proteins that exhibit low acetylation levels. Fatty acid synthase (FASN), an essential enzyme catalyzing the de novo synthesis of fatty acids, has been found to be acetylated in various species, from fruit flies to humans. Here, we describe a step-by-step process of antibody-based protein enrichment and sample preparation for acetylation identification of endogenous FASN protein by MS-based proteomics analysis. Meanwhile, we provide a protocol for nicotinamide adenine dinucleotide phosphate (NADPH) absorbance assay for FASN activity measurement, which is one of the primary functional readouts of de novo lipogenesis.

Key features

• A comprehensive protocol for protein immunoprecipitation and sample preparation for acetylation site identification by mass spectrometry.

• Step-by-step procedures for measurement of FASN activity of fruit fly larvae using an absorbance assay.

Graphical overview

Macropinocytosis is an evolutionarily conserved process, which is characterized by the formation of membrane ruffles and the uptake of extracellular fluid. We recently demonstrated a role for CYFIP-related Rac1 Interactor (CYRI) proteins in macropinocytosis. High-molecular weight dextran (70kDa or higher) has generally been used as a marker for macropinocytosis because it is too large to fit in smaller endocytic vesicles, such as those of clathrin or caveolin-mediated endocytosis. Through the use of an image-based dextran uptake assay, we showed that cells lacking CYRI proteins internalise less dextran compared to their wild-type counterparts. Here, we will describe a step-by-step experimentation procedure to detect internalised dextran in cultured cells, and an image pipeline to analyse the acquired images, using the open-access software ImageJ/Fiji. This protocol is detailed yet simple and easily adaptable to different treatment conditions, and the analysis can also be automated for improved processing speed.

The ability to stain lipid stores in vivo allows for the facile assessment of metabolic status in individuals of a population following genetic and environmental manipulation or pharmacological treatment. In the animal model Caenorhabditis elegans, lipids are stored in and mobilized from intracellular lipid droplets in the intestinal and hypodermal tissues. The abundance, size, and distribution of these lipids can be readily assessed by two staining methods for neutral lipids: Oil Red O (ORO) and Nile Red (NR). ORO and NR can be used to quantitatively measure lipid droplet abundance, while ORO can also define tissue distribution and lipid droplet size. C. elegans are a useful animal model in studying pathways relating to aging, fat storage, and metabolism, as their transparent nature allows for easy microscopic assessment of lipid droplets. This is done by fixation and permeabilization, staining with NR or ORO, image capture on a microscope, and computational identification and quantification of lipid droplets in individuals within a cohort. To ensure reproducibility in lipid measurements, we provide a detailed protocol to measure intracellular lipid dynamics in C. elegans.

Graphic abstract:

Flow chart depicting the preparation of C. elegans for fat staining protocols.

Three-dimensional (3D) cell culture models are widely used in tumor studies to more accurately reflect cell-cell interactions and tumor growth conditions in vivo. 3D anchorage-independent spheroids derived by culturing cells in ultra-low attachment (ULA) conditions is particularly relevant to ovarian cancer, as such cell clusters are often observed in malignant ascites of late-stage ovarian cancer patients. We and others have found that cells derived from anchorage-independent spheroids vary widely in gene expression profiles, proliferative state, and metabolism compared to cells maintained under attached culture conditions. This includes changes in mitochondrial function, which is most commonly assessed in cultured live cells by measuring oxygen consumption in extracellular flux assays. To measure mitochondrial function in anchorage-independent multicellular aggregates, we have adapted the Agilent Seahorse extracellular flux assay to optimize measurements of oxygen consumption and extracellular acidification of ovarian cancer cell spheroids generated by culture in ULA plates. This protocol includes: (i) Methods for culturing tumor cells as uniform anchorage-independent spheroids; (ii) Optimization for the transfer of spheroids to the Agilent Seahorse cell culture plates; (iii) Adaptations of the mitochondrial and glycolysis stress tests for spheroid extracellular flux analysis; and (iv) Suggestions for optimization of cell numbers, spheroid size, and normalization of oxygen consumption rate (OCR) and extracellular acidification rate (ECAR) values. Using this method, we have found that ovarian cancer cells cultured as anchorage-independent spheroids display altered mitochondrial function compared to monolayer cultures attached to plastic dishes. This method allows for the assessment of mitochondrial function in a more relevant patho/physiological culture condition and can be adapted to evaluate mitochondrial function of various cell types that are able to aggregate into multicellular clusters in anchorage-independence.

Graphic abstract:



Workflow of the Extracellular Flux Assay to Measure Respiration of Anchorage-independent Tumor Cell Spheroids.

Reactive oxygen species and reactive nitrogen species (RONS) are involved in programmed cell death in the context of numerous degenerative and chronic diseases. In particular, the ability of cells to maintain redox homeostasis is necessary for an adaptive cellular response to adverse conditions that can cause damage to proteins and DNA, resulting in apoptosis and genetic mutations. Here, we focus on the 2',7'-dichlorodihydrofluorescein diacetate (DCFH₂-DA) assay to detect RONS. Although this fluorescence-based assay is widely utilized due to its high sensitivity to detect changes in cellular redox status that allow measuring alterations in RONS over time, its validity has been a matter of controversy. If correctly carried out, its limitations are understood and results are correctly interpreted, the DCFH₂-DA assay is a valuable tool for cell-based studies.

Once thought to be a mere consequence of the state of a cell, intermediary metabolism is now recognized as a key regulator of mammalian cell fate and function. In addition, cell metabolism is often disturbed in malignancies such as cancer, and targeting metabolic pathways can provide new therapeutic options. Cell metabolism is mostly studied in cell cultures in vitro, using techniques such as metabolomics, stable isotope tracing, and biochemical assays. Increasing evidence however shows that the metabolic profile of cells is highly dependent on the microenvironment, and metabolic vulnerabilities identified in vitro do not always translate to in vivo settings. Here, we provide a detailed protocol on how to perform in vivo stable isotope tracing in leukemia cells in mice, focusing on glutamine metabolism in acute myeloid leukemia (AML) cells. This method allows studying the metabolic profile of leukemia cells in their native bone marrow niche.

The endothelial cells from the microvasculature are key drivers and targets of inflammatory and thrombotic processes in microvascular diseases. The study of bioactive lipids in inflammatory processes has been largely based on two-dimensional endothelial cell cultures. Three-dimensional microvessels-on-a-chip provides an opportunity to monitor the inflammatory phenotype of human microvessels in a more physiological-relevant environment. This protocol describes the culture of endothelial cells as three-dimensional microvessels in the OrganoPlate. The microvessels are treated with tumor necrosis factor alpha to induce inflammation. The collection of samples from the microvessels is optimized for measuring bioactive lipids with liquid chromatography-mass spectrometry, providing a more informative metabolic readout as compared with functional assays.

The activation of the Takeda G-protein receptor 5 (TGR5, also known as the G protein-coupled bile acid receptor 1, GPBAR1) in enteroendocrine L-cells results in secretion of the anti-diabetic hormone Glucagon-Like Peptide 1 (GLP-1) into systemic circulation. Consequently, recent research has focused on identification and development of TGR5 agonists as type 2 diabetes therapeutics. However, the clinical application of TGR5 agonists has been hampered by side effects of these compounds that primarily result from their absorption into circulation. Here we describe an in vitro screening protocol to evaluate the TGR5 agonism, GLP-1 secretion, and gut-restricted properties of small molecules. The protocol involves differentiating gut epithelial and endocrine cells together in transwells to assess both the pharmacodynamics of TGR5 agonists and the toxicity of compounds to the intestinal monolayer. As a proof of concept, we demonstrate the use of the protocol in evaluating properties of naturally occurring bile acid metabolites that are potent TGR5 agonists. This protocol is adapted from Chaudhari et al. (2021).