Noninvasive quantitation of metabolic flux in cancer cells

Metabolic reprogramming facilitates cancer cell growth, so quantitative metabolic flux measurements could produce useful biomarkers. However, current methods to analyze flux in vivo provide either a steady-state overview of relative activities (infusion of C and analysis of extracted metabolites) or a dynamic view of a few reactions (hyperpolarized C spectroscopy). Moreover, while hyperpolarization has successfully quantified pyruvate-lactate exchanges, its ability to assess mitochondrial pyruvate metabolism is unproven in cancer. Here, we combined C hyperpolarization and isotopomer analysis to quantify multiple fates of pyruvate simultaneously. Two cancer cell lines with divergent pyruvate metabolism were incubated with thermallypolarized [3-C]pyruvate for several hours, then briefly exposed to hyperpolarized [1-C]pyruvate during acquisition of NMR spectra, using selective excitation to maximize detection of H[C]O3 and [1-C]lactate. Metabolites were then extracted and subjected to isotopomer analysis to determine relative rates of pathways involving [3C]pyruvate. Quantitation of hyperpolarized H[C]O3 provided a single definitive metabolic rate, which was then used to convert relative rates derived from isotopomer analysis into quantitative fluxes. This revealed that H[C]O3 appearance reflects activity of pyruvate dehydrogenase (PDH) rather than pyruvate carboxylation followed by subsequent decarboxylation reactions. Glucose substantially altered [1-C]pyruvate metabolism, enhancing exchanges with [1-C]lactate and suppressing H[C]O3 formation. Furthermore, inhibiting Akt, an oncogenic kinase that stimulates glycolysis, reversed these effects, indicating that metabolism of pyruvate by both LDH and PDH is subject to the acute effects of oncogenic signaling on glycolysis. The data suggest that combining C isotopomer analyses and dynamic hyperpolarized C spectroscopy may enable quantitative flux measurements in living tumors. http://www.jbc.org/cgi/doi/10.1074/jbc.M113.543637 The latest version is at JBC Papers in Press. Published on January 10, 2014 as Manuscript M113.543637 Copyright 2014 by The American Society for Biochemistry and Molecular Biology, Inc. by gest on Jne 9, 2017 hp://w w w .jb.org/ D ow nladed from Noninvasive quantitation of metabolic flux in cancer cells 2 Cancer cells display metabolic properties that distinguish them from surrounding, normal tissue. Tumor metabolism is regulated by oncogenes and tumor suppressors (1-4), suggesting that mutations in these genes orchestrate metabolic activities to support an environment conducive to malignancy (5,6). In accordance with this view, mutations in a subset of metabolic enzymes, including fumarate hydratase, the succinate dehydrogenase complex, and isocitrate dehydrogenases-1 and -2 cause significant perturbations in intermediary metabolism and contribute directly to cancer (711). Metabolism is also highly sensitive to effects of the tumor microenvironment, particularly hypoxia, which correlates with chemotherapeutic resistance and poor patient survival (12). Thus, the ability to measure metabolic flux in live tumors would potentially report clinically and prognostically valuable information. Imaging of hyperpolarized C-enriched substrates is emerging as a promising technology for cancer diagnosis and monitoring because it enables the real-time detection of enzyme-catalyzed metabolic activities in tissues (13-18). Recently, it has also been shown to be capable of detecting malignant lesions in the human prostate, even diagnosing masses that were not positively identified by other methods (19). These studies revealed that exchanges between hyperpolarized pyruvate and lactate are readily detected in malignant cells and tend to be enhanced in tumors relative to nonmalignant tissue. Hyperpolarization involves the temporary redistribution of the populations of the available energy levels into a non-equilibrium state, enabling a massive gain in magnetic resonance signal. For C, this gain can exceed 10,000-fold (20), enabling detection of both a Cenriched substrate and some downstream metabolites with a temporal resolution of seconds. Hyperpolarized [1-C]pyruvate is widely used to interrogate cancer metabolism, in part because the long T1 of this carboxyl carbon allows the hyperpolarized state to persist for multiple metabolic steps, and also because pyruvate is located in a pivotal intersection of intermediary metabolism involving lactate, alanine, and oxidation in the tricarboxylic acid (TCA) cycle. Cancer cells often display high rates of glycolysis and lactate secretion even when oxygen is plentiful (the Warburg effect). This oncogenedriven phenomenon contributes to the utility of tumor detection by uptake of the glucose analog [F]-fluoro-2-deoxyglucose (FDG) (3). Abundant expression of lactate dehydrogenase (LDH) and the presence of a large lactate pool in tumor cells allows for rapid exchange between hyperpolarized [1-C]pyruvate and [1-C]lactate, making detection of [1-C]lactate appealing for in vivo detection of cancer and for monitoring response to therapy (21,22). However, cancer cells also oxidize pyruvate in the mitochondria, producing both energy and macromolecular precursors for cell growth (23). This is of particular interest because lung tumors, gliomas, and metastatic brain tumors have all been demonstrated to oxidize pyruvate in vivo in humans and mice (24-28). Therefore, assessment of both pyruvate/lactate exchanges and pyruvate oxidation in the mitochondria would provide a much more comprehensive view of cancer cell metabolism than lactate formation alone. We previously used conventional C NMR spectroscopy to evaluate fluxes through competing metabolic pathways supplied by pyruvate, including LDH and the TCA cycle, in cultured cancer cells (29,30). These same activities were detected in mouse and human tumors by infusing C-enriched glucose prior to surgery, extracting metabolites from surgically-resected tumor tissue, and analyzing C enrichment patterns by NMR (26,28). We also used hyperpolarized [1C]pyruvate to quantify flux into lactate (31). Here, we combined these methods to study two metabolically distinct cancer cell lines. First, we incubated cancer cells with thermally polarized [3C]pyruvate for several hours to produce steadystate labeling of metabolic intermediates. Next, using a selective excitation pulse to maximize detection of H[C]O3 and [1-C]lactate, we subjected cells to hyperpolarized [1-C]pyruvate to measure flux into lactate and the TCA cycle. Combining the rate of pyruvate decarboxylation with steady-state isotopomer data provided a method to evaluate absolute flux rates through a variety of reactions associated with the TCA cycle. EXPERIMENTAL PROCEDURES by gest on Jne 9, 2017 hp://w w w .jb.org/ D ow nladed from Noninvasive quantitation of metabolic flux in cancer cells 3 Cell culture reagents and basic metabolism experiments. Two cell lines, SF188-derived glioblastoma cells overexpressing human Bcl-xL (SFxL) and Huh-7 hepatocellular carcinoma cells were maintained in culture as described previously (30,32,33). Metabolic experiments were performed in Dulbecco’s Modified Eagle Medium (DMEM) prepared from powder lacking glucose, glutamine, phenol red, sodium pyruvate and sodium bicarbonate. This basal medium was supplemented with 4 mmol/L L-glutamine, 10% dialyzed fetal calf serum, 42.5 mmol/L sodium bicarbonate, 25 mmol/L HEPES, 10 units/mL penicillin and 10 μg/mL streptomycin. Glucose and pyruvate were added as indicated for each experiment. To measure rates of metabolite consumption/excretion in the medium, glucose, lactate, glutamine and glutamate were measured using a BioProfile Basic 4 analyzer (NOVA Biomedical) and ammonia was measured using a spectrophotometric assay (Megazyme). For oxygen consumption assays, cells were harvested by trypsinization, suspended in fresh medium at a concentration of 10 cells/ml, and transferred to an Oxygraph water-jacketed oxygen electrode (Hansatech). The Akt inhibitor was Akt Inhibitor VIII (Calbiochem). Pyruvate decarboxylation assay. Decarboxylation of [1-C]pyruvate was measured essentially as described (34). Micro-bridges (Hampton Research) were placed into wells of a 24-well plate with one piece of 0.6 X 1 cm chromatography paper in each. Assay medium was prepared by supplementing DMEM (containing 10% fetal calf serum, 4 mM glutamine and 6 mM sodium pyruvate) with 2.2 μCi of [1-C]pyruvate. This medium was warmed to 37C and incubated for 2 hours to remove any CO2 produced from spontaneous decarboxylation, then an aliquot was used to quantify radioactivity on a scintillation counter. This value was used to determine the specific activity of pyruvate, assuming a total pyruvate concentration of 6 mM. The specific activity ranged from 50-120 CPM/nmole pyruvate. One million cells per well were then suspended in 370 μl assay medium on ice. Each micro-bridge was moistened with 30 μl of 2N NaOH, and the plate was sealed with adhesive film. Pyruvate metabolism was initiated by transferring the plate to a 37C water bath. After 15 minutes, metabolism was terminated by adding 50 μl of 20% trichloroacetic acid. The plate was re-sealed with adhesive film and incubated at 37C for another 60 minutes to release CO2 completely. Then the CO2–containing chromatography papers were collected for scintillation counts. Pyruvate oxidation rates were determined from the total specific activity in the reaction, and were reported as nmol/10 cells/hr. Wells containing culture medium but no cells were used to establish background levels of CO2. This level was subtracted from cell-containing wells. Gas chromatography/mass spectrometry. Mass spectrometry experiments were performed essentially as described previously (30). Cultures of 80-90% confluent cells grown in 60-mm dishes were rinsed twice in phosphate buffered saline, then overlaid with DMEM-based medium containing 10% dialyzed fetal calf serum, 4 mM glutamine, and 6 mM pyruvate (either [1C]pyruvate or [3-C]pyruvate, Cambridge Isotope Laboratories). At the end of the culture period, the medium was removed and the cells were briefly rinsed in ice-cold normal saline. A cold solution of 50% methanol/50% water was added, and the cells were lysed using repeated freeze-thaw cycles. After centrifugation to remove debris, the samples were evaporated and derivatized by trimethylsilylation (Tri-Sil HTP reagent, Thermo). An aliquot of 1-3 μL was injected onto an Agilent 6970 gas chromatograph networked to an Agilent 5973 Mass Selective Detector. Fragment ions m/z 334-338 and 465-471 were used to monitor enrichment in aspartate and citrate, respectively (30). The measured distribution of mass isotopomers was corrected for natural abundance of C as described (35). NMR spectroscopy of extracted metabolites. Cells were cultured to 80-90% confluence in eight 150mm dishes. Freshly-prepared DMEM with 10% dialyzed fetal calf serum, 0 mM glucose, 4 mM glutamine, standard concentrations of other amino acids, and 6 mM [3-C]pyruvate was added to each dish, and the cells were cultured for 6 hours, then either harvested immediately for extraction or collected for hyperpolarization experiments. For hyperpolarization, the cells were trypsinized, pelleted by centrifugation, and resuspended at a by gest on Jne 9, 2017 hp://w w w .jb.org/ D ow nladed from Noninvasive quantitation of metabolic flux in cancer cells 4 concentration of 1x10 cells/mL in fresh DMEM lacking glucose and pyruvate, but containing 10% dialyzed fetal calf serum, 4 mM glutamine and standard concentrations of other amino acids. This suspension was mixed thoroughly, then transferred to a 10-mm NMR tube which already contained hyperpolarized [1-C]pyruvate so that the final concentration was 6 mM, precisely mimicking the nutrient availability of the steady-state labeling experiment. Acquisition of spectra was initiated immediately and proceeded for approximately 3.5 minutes (see “Hyperpolarization, shaped pulse and NMR spectroscopy in intact cells” for details on cell transfer and conditions for the hyperpolarization experiment). We previously reported that cells subjected to trypsinization and hyperpolarized [1-C]pyruvate maintain rates of LDH flux similar to rates of adherent cells (31). To protect against depletion of oxygen from the medium during trypsinization and preparation of the cell suspension, the medium was mixed frequently, including immediately before transfer of the cells to the NMR tube, which occurred within a few seconds of introduction of [1C]pyruvate. Immediately after the hyperpolarization experiment, the cells were pelleted by centrifugation and frozen in liquid nitrogen. Frozen cell pellets were homogenized by sonication in 4 mL of 4% ice-cold perchloric acid. After centrifuging to remove debris, the acidsoluble material was neutralized with 8N potassium hydroxide and centrifuged. Supernatants were subjected to lyophilization, reconstituted in deuterium oxide and titrated to pH 7 for NMR analysis in a 3 mm tube. NMR spectroscopy was performed on a Varian INOVA 14.1 T spectrometer (Agilent Instruments, Walnut Creek, CA) equipped with a 3-mm broadband probe with the observe coil tuned to C (150 MHz). Proton decoupling was performed using a standard WALTZ-16 pulse sequence. Carbon spectra were acquired under the following conditions: pulse flip angle 45°, repetition time 1.5 s, spectral width 35 kHz, number of data points 104,986, and number of scans 7,000–30,000, requiring 6–25 h. Free induction decays were zero-filled to 131,072 points and apodized with exponential multiplication. Peak areas were determined using ACD Labs SpecManager (Advanced Chemistry Development, Toronto, Canada). Glutamate isotopomer analysis to calculate FC2 (the fraction of acetyl-CoA labeled in position-2 with C) in Fig. 3 was performed using equations described in (36). Relative flux values for the combined steady-state/hyperpolarization experiments in Table 1, including FC2 and ys (the rate of anaplerosis relative to citrate synthase flux) were calculated by tcaCALC (37). Hyperpolarization, shaped pulses and NMR spectroscopy of intact cells. A solution of [1C]pyruvic acid containing 15 mmol/L OX63 radical was prepared for polarization. 8.6 μL of the solution was polarized for 2 hours in an Oxford HyperSense dynamic nuclear polarization system (Oxford Instruments Molecular Biotools Ltd, Oxfordshire, UK) at 1.4 K with a microwave irradiation frequency of 94.125 GHz at 100 mW. The frozen sample was dissolved with 4 mL of 15.3 mM sodium bicarbonate (heated to 190° C and pressurized to 10 bar) in less than 10 seconds. An aliquot of the resulting solution was added to a 10 mm NMR tube (400 μL for 2 mL total volume or 200 μL for 1 mL total volume). The tube was placed in a Varian 10-mm broadband probe in a 9.4 Tesla magnet equipped with a VNMRS console (Agilent Inc., Walnut Creek, CA). The NMR tube was directly attached by a thin tube to a syringe containing the suspension of cells. At time zero of the acquisition, either 1.6 or 0.8 mL of the cell suspension (for final total volumes of 1 or 2 mL, respectively) was added to the hyperpolarized solution, resulting in a final pyruvate concentration of 6 mM and a cell concentration of 1.0 x10 cells/mL. Frequency selective pulses were utilized to avoid depolarization of the pyruvate C1 resonance. Pulses were created in the PBox software provided with VNMRJ (Agilent). The pulse profile was designed to excite the pyruvate C2 resonance (205.9 ppm), lactate C1 (183.3 ppm), and H[C]O3. (160.1 ppm) with a Gaussian shape of narrow bandwidth (350-500 Hz). Profiles were fit to a triple Gaussian function: by gest on Jne 9, 2017 hp://w w w .jb.org/ D ow nladed from Noninvasive quantitation of metabolic flux in cancer cells