Cancer Biology

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.

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 production of reactive oxygen species (ROS) and endoplasmic reticulum (ER) stress are tightly linked. The generation of ROS can be both the cause and a consequence of ER stress pathways, and an increasing number of human diseases are characterized by tissue atrophy in response to ER stress and oxidative injury. For the assessment of modulators of ER luminal ROS generation and for mechanistic studies, methods to monitor changes in ER reduction-oxidation (redox) states in a time-resolved and organelle-specific manner are needed. This has been greatly facilitated by the development of genetically encoded fluorescent probes, which can be targeted to different subcellular locations by specific amino acid extensions. One of these probes is the yellow fluorescent protein-based redox biosensor, HyPer. Here, we provide a protocol for the time-resolved monitoring of the oxidizing milieu in the ER of adherent mammalian cells using the ratiometric sensor, HyPerER, which is specifically targeted to the ER lumen.

Mammalian cells generate ATP by mitochondrial (oxidative phosphorylation) and non-mitochondrial (glycolysis) metabolism. Cancer cells are known to reprogram their metabolism using different strategies to meet energetic and anabolic needs (Koppenol et al., 2011; Zheng, 2012). Additionally, each cancer tissue has its own individual metabolic features. Mitochondria not only play a key role in energy metabolism but also in cell cycle regulation of cells. Therefore, mitochondria have emerged as a potential target for anticancer therapy since they are structurally and functionally different from their non-cancerous counterparts (D'Souza et al., 2011). We detail a protocol for measurement of oxygen consumption rate (OCR) and extracellular acidification rate (ECAR) measurements in living cells, utilizing the Seahorse XF24 Extracellular Flux Analyzer (Figure 1). The Seahorse XF24 Extracellular Flux Analyzer continuously measures oxygen concentration and proton flux in the cell supernatant over time (Wu et al., 2007). These measurements are converted in OCR and ECAR values and enable a direct quantification of mitochondrial respiration and glycolysis. With this protocol, we sought to assess basal mitochondrial function and mitochondrial stress of three different cancer cell lines in response to the cytotoxic test lead compound mensacarcin in order to investigate its mechanism of action. Cells were plated in XF24 cell culture plates and maintained for 24 h. Prior to analysis, the culture media was replaced with unbuffered DMEM pH 7.4 and cells were then allowed to equilibrate in a non-CO₂ incubator immediately before metabolic flux analysis using the Seahorse XF to allow for precise measurements of Milli-pH unit changes. OCR and ECAR were measured under basal conditions and after injection of compounds through drug injection ports. With the described protocol we assess the basic energy metabolism profiles of the three cell lines as well as key parameters of mitochondrial function in response to our test compound and by sequential addition of mitochondria perturbing agents oligomycin, FCCP and rotenone/antimycin A.


Figure 1. Overview of seahorse experiment

This is a flow cytometry-based protocol to measure glucose uptake of mouse embryonic fibroblasts (MEFs) and breast cancer cells in vitro. The method is a slightly modified and updated version as previously described (Dong et al., 2017). Briefly, the target cells are incubated with the fluorescently tagged 2-(N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl)amino)-2-deoxyglucose (2-NBDG) for 2 h or 30 min, and the efficiency of glucose uptake is examined using a flow cytometer. This method can be adapted to measure a variety of adipocytes, immune cells, MEFs and cancer cells.

NADH and NADPH are redox cofactors, primarily involved in catabolic and anabolic metabolic processes respectively. In addition, NADPH plays an important role in cellular antioxidant defence. In live cells and tissues, the intensity of their spectrally-identical autofluorescence, termed NAD(P)H, can be used to probe the mitochondrial redox state, while their distinct enzyme-binding characteristics can be used to separate their relative contributions to the total NAD(P)H intensity using fluorescence lifetime imaging microscopy (FLIM). These protocols allow differences in metabolism to be detected between cell types and altered physiological and pathological states.

Like all animals, Drosophila shows robust fat (triglyceride) turnover, i.e., they synthesize, store and utilize triglyceride for their daily metabolic needs. The protocol describes a simple assay to measure this turnover of triglycerides in Drosophila.

Glutamine synthetase (GS), which catalyzes the conversion of glutamate and ammonia to glutamine, is widely distributed in animal tissues and cell culture lines. The importance of this enzyme is suggested by the fact that glutamine, the product of GS-catalyzed de novo synthesis reaction, is the most abundant free amino acid in blood (Smith and Wilmore, 1990). Glutamine is involved in many biological processes including serving as the nitrogen donor for biosynthesis, as an exchanger for the import of essential amino acids, as a means to detoxifying intracellular ammonia and glutamate, and as a bioenergetics nutrient to fuel the tricarboxylic acid (TCA) cycle (Bott et al., 2015). The method for the assay of GS enzymatic activity relies on its γ-glutamyl transferase reaction by measuring γ-glutamylhydroxamate synthesized from glutamine and hydroxylamine, and the chromatographic separation of the reaction product from the reactants (Deuel et al., 1978). An overview of the GS glutamyl transferase reaction can be found in Figure 1. GS activity was measured by a spectrophotometric assay at a specific wavelength of 560 nm using a microplate reader. The method is simple, and has a comparable sensitivity with those methods applying radioactively labelled substrates. This modified procedure has been applied to assay/determine GS activity in cultured cell lines including the human mammary epithelial MCF10A cells and the murine pre-B FL5.12 cells, and could be used to measure GS activity in other cell lines.


Figure 1. An overview of the GS glutamyl transferase reaction

Iron in blood plasma is bound to its transport protein transferrin, which delivers iron to most tissues. In iron overload and certain pathological conditions, the carrying capacity of transferrin can become exceeded, giving rise to non-transferrin-bound iron, which is taken up preferentially by the liver, kidney, pancreas, and heart. The measurement of tissue transferrin- and non-transferrin-bound iron (TBI and NTBI, respectively) uptake in vivo can be achieved via intravenous administration of ⁵⁹Fe-labeled TBI or NTBI followed by gamma counting of various organs. Here we describe a detailed protocol for the measurement of TBI and NTBI uptake by mouse tissues.

Two enantiomers of 2-hydroxyglutarate (2HG), L (L2HG) and D (D2HG), are metabolites of unknown function in mammalian cells that were initially associated with separate and rare inborn errors of metabolism resulting in increased urinary excretion of 2HG linked to neurological deficits in children (Chalmers et al., 1980; Duran et al., 1980; Kranendijk et al., 2012). More recently, investigators have shown that D2HG is produced by mutant isocitrate dehydrogenase enzymes associated with a variety of human malignancies, such as acute myeloid leukemia, glioblastoma multiforme, and cholangiocarcinoma (Cairns and Mak, 2013; Dang et al., 2009; Ward et al., 2010). By contrast, we and others have shown that L2HG accumulates in response to cellular reductive stressors like hypoxia, activation of hypoxia inducible factors, and mitochondrial electron transport chain defects (Oldham et al., 2015; Reinecke et al., 2011; Intlekofer et al., 2015; Mullen et al., 2015). Each enantiomer is produced and metabolized in independent biochemical pathways in reactions catalyzed by separate enzymes and utilizing different cofactors with presumably different consequences for cellular metabolism (Kranendijk et al., 2012). Therefore, as research into the roles of D2HG and L2HG in human metabolism continues, it becomes increasingly important for investigators to consider each enantiomer independently (Struys, 2013). Several methods for quantification of biochemically relevant enantiomers in general have been developed and typically include enzymatic assays using enzymes specific for one enantiomeric species or the other, the use of chiral chromatography medium to facilitate chromatographic separation of enantiomers prior to spectroscopy, or the use of chiral derivatization reagents to convert a mixture of enantiomers to diastereomers with differing physical and chemical properties facilitating their chromatographic separation. In this protocol, we report the adaptation of a previously published derivatization method using diacetyl-L-tartaric anhydride (DATAN) for the quantification of 2HG enantiomers (Figure 1) (Oldham et al., 2015; Struys et al., 2004).


Figure 1. Reaction scheme for the derivatization protocol