UCP2 regulates energy metabolism and differentiation potential of human pluripotent stem cells

Article15 November 2011free access UCP2 regulates energy metabolism and differentiation potential of human pluripotent stem cells Jin Zhang Jin Zhang Department of Pathology and Laboratory Medicine, University of California, Los Angeles, CA, USA Search for more papers by this author Ivan Khvorostov Ivan Khvorostov Department of Pathology and Laboratory Medicine, University of California, Los Angeles, CA, USA Search for more papers by this author Jason S Hong Jason S Hong Department of Pathology and Laboratory Medicine, University of California, Los Angeles, CA, USA Search for more papers by this author Yavuz Oktay Yavuz Oktay Department of Chemistry and Biochemistry, University of California, Los Angeles, CA, USA Search for more papers by this author Laurent Vergnes Laurent Vergnes Department of Human Genetics, University of California, Los Angeles, CA, USA Search for more papers by this author Esther Nuebel Esther Nuebel Department of Chemistry and Biochemistry, University of California, Los Angeles, CA, USA Search for more papers by this author Paulin N Wahjudi Paulin N Wahjudi Department of Pediatrics, LA Biomedical Research Institute, Torrance, CA, USA Search for more papers by this author Kiyoko Setoguchi Kiyoko Setoguchi Department of Pathology and Laboratory Medicine, University of California, Los Angeles, CA, USA Search for more papers by this author Geng Wang Geng Wang Department of Chemistry and Biochemistry, University of California, Los Angeles, CA, USA Search for more papers by this author Anna Do Anna Do Department of Pathology and Laboratory Medicine, University of California, Los Angeles, CA, USA Search for more papers by this author Hea-Jin Jung Hea-Jin Jung Department of Pathology and Laboratory Medicine, University of California, Los Angeles, CA, USA Search for more papers by this author J Michael McCaffery J Michael McCaffery Department of Biology, Integrated Imaging Center, Johns Hopkins University, Baltimore, MD, USA Search for more papers by this author Irwin J Kurland Irwin J Kurland Department of Medicine, Stable Isotopes and Metabolomics Core Facility, Albert Einstein College of Medicine Diabetes Center, Bronx, NY, USA Search for more papers by this author Karen Reue Karen Reue Department of Human Genetics, University of California, Los Angeles, CA, USA Molecular Biology Institute, University of California, Los Angeles, CA, USA Search for more papers by this author Wai-Nang P Lee Wai-Nang P Lee Department of Pediatrics, LA Biomedical Research Institute, Torrance, CA, USA Search for more papers by this author Carla M Koehler Corresponding Author Carla M Koehler Department of Chemistry and Biochemistry, University of California, Los Angeles, CA, USA Molecular Biology Institute, University of California, Los Angeles, CA, USA Search for more papers by this author Michael A Teitell Corresponding Author Michael A Teitell Department of Pathology and Laboratory Medicine, University of California, Los Angeles, CA, USA Molecular Biology Institute, University of California, Los Angeles, CA, USA Broad Stem Cell Research Center, Jonsson Comprehensive Cancer Center, Center for Cell Control, and California NanoSystems Institute, University of California, Los Angeles, CA, USA Search for more papers by this author Jin Zhang Jin Zhang Department of Pathology and Laboratory Medicine, University of California, Los Angeles, CA, USA Search for more papers by this author Ivan Khvorostov Ivan Khvorostov Department of Pathology and Laboratory Medicine, University of California, Los Angeles, CA, USA Search for more papers by this author Jason S Hong Jason S Hong Department of Pathology and Laboratory Medicine, University of California, Los Angeles, CA, USA Search for more papers by this author Yavuz Oktay Yavuz Oktay Department of Chemistry and Biochemistry, University of California, Los Angeles, CA, USA Search for more papers by this author Laurent Vergnes Laurent Vergnes Department of Human Genetics, University of California, Los Angeles, CA, USA Search for more papers by this author Esther Nuebel Esther Nuebel Department of Chemistry and Biochemistry, University of California, Los Angeles, CA, USA Search for more papers by this author Paulin N Wahjudi Paulin N Wahjudi Department of Pediatrics, LA Biomedical Research Institute, Torrance, CA, USA Search for more papers by this author Kiyoko Setoguchi Kiyoko Setoguchi Department of Pathology and Laboratory Medicine, University of California, Los Angeles, CA, USA Search for more papers by this author Geng Wang Geng Wang Department of Chemistry and Biochemistry, University of California, Los Angeles, CA, USA Search for more papers by this author Anna Do Anna Do Department of Pathology and Laboratory Medicine, University of California, Los Angeles, CA, USA Search for more papers by this author Hea-Jin Jung Hea-Jin Jung Department of Pathology and Laboratory Medicine, University of California, Los Angeles, CA, USA Search for more papers by this author J Michael McCaffery J Michael McCaffery Department of Biology, Integrated Imaging Center, Johns Hopkins University, Baltimore, MD, USA Search for more papers by this author Irwin J Kurland Irwin J Kurland Department of Medicine, Stable Isotopes and Metabolomics Core Facility, Albert Einstein College of Medicine Diabetes Center, Bronx, NY, USA Search for more papers by this author Karen Reue Karen Reue Department of Human Genetics, University of California, Los Angeles, CA, USA Molecular Biology Institute, University of California, Los Angeles, CA, USA Search for more papers by this author Wai-Nang P Lee Wai-Nang P Lee Department of Pediatrics, LA Biomedical Research Institute, Torrance, CA, USA Search for more papers by this author Carla M Koehler Corresponding Author Carla M Koehler Department of Chemistry and Biochemistry, University of California, Los Angeles, CA, USA Molecular Biology Institute, University of California, Los Angeles, CA, USA Search for more papers by this author Michael A Teitell Corresponding Author Michael A Teitell Department of Pathology and Laboratory Medicine, University of California, Los Angeles, CA, USA Molecular Biology Institute, University of California, Los Angeles, CA, USA Broad Stem Cell Research Center, Jonsson Comprehensive Cancer Center, Center for Cell Control, and California NanoSystems Institute, University of California, Los Angeles, CA, USA Search for more papers by this author Author Information Jin Zhang1, Ivan Khvorostov1, Jason S Hong1, Yavuz Oktay2, Laurent Vergnes3, Esther Nuebel2, Paulin N Wahjudi4, Kiyoko Setoguchi1, Geng Wang2, Anna Do1, Hea-Jin Jung1, J Michael McCaffery5, Irwin J Kurland6, Karen Reue3,7, Wai-Nang P Lee4, Carla M Koehler 2,7 and Michael A Teitell 1,7,8 1Department of Pathology and Laboratory Medicine, University of California, Los Angeles, CA, USA 2Department of Chemistry and Biochemistry, University of California, Los Angeles, CA, USA 3Department of Human Genetics, University of California, Los Angeles, CA, USA 4Department of Pediatrics, LA Biomedical Research Institute, Torrance, CA, USA 5Department of Biology, Integrated Imaging Center, Johns Hopkins University, Baltimore, MD, USA 6Department of Medicine, Stable Isotopes and Metabolomics Core Facility, Albert Einstein College of Medicine Diabetes Center, Bronx, NY, USA 7Molecular Biology Institute, University of California, Los Angeles, CA, USA 8Broad Stem Cell Research Center, Jonsson Comprehensive Cancer Center, Center for Cell Control, and California NanoSystems Institute, University of California, Los Angeles, CA, USA *Corresponding authors: Department of Chemistry and Biochemistry, University of California, Los Angeles, 607 Charles E. Young Drive East, Los Angeles, CA 90095, USA. Tel.: +1 310 794 4834; Fax: +1 310 206 4038; E-mail: [email protected] of Pathology and Laboratory Medicine, University of California, Los Angeles, 675 Charles E. Young Drive South, Los Angeles, CA 90095, USA. Tel.: +1 310 206 6754; Fax: +1 310 267 0382; E-mail: [email protected] The EMBO Journal (2011)30:4860-4873https://doi.org/10.1038/emboj.2011.401 Correction(s) for this article UCP2 regulates energy metabolism and differentiation potential of human pluripotent stem cells18 April 2016 There is a Have you seen? (December 2011) associated with this Article. PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info It has been assumed, based largely on morphologic evidence, that human pluripotent stem cells (hPSCs) contain underdeveloped, bioenergetically inactive mitochondria. In contrast, differentiated cells harbour a branched mitochondrial network with oxidative phosphorylation as the main energy source. A role for mitochondria in hPSC bioenergetics and in cell differentiation therefore remains uncertain. Here, we show that hPSCs have functional respiratory complexes that are able to consume O2 at maximal capacity. Despite this, ATP generation in hPSCs is mainly by glycolysis and ATP is consumed by the F1F0 ATP synthase to partially maintain hPSC mitochondrial membrane potential and cell viability. Uncoupling protein 2 (UCP2) plays a regulating role in hPSC energy metabolism by preventing mitochondrial glucose oxidation and facilitating glycolysis via a substrate shunting mechanism. With early differentiation, hPSC proliferation slows, energy metabolism decreases, and UCP2 is repressed, resulting in decreased glycolysis and maintained or increased mitochondrial glucose oxidation. Ectopic UCP2 expression perturbs this metabolic transition and impairs hPSC differentiation. Overall, hPSCs contain active mitochondria and require UCP2 repression for full differentiation potential. Introduction A distinguishing feature of human pluripotent stem cells (hPSCs) compared with differentiated cells is the capacity for self-renewal to maintain pluripotency. Self-renewal is supported by unique chromatin modifications (Meshorer and Misteli, 2006) and a regulatory circuit comprised of OCT4, NANOG, and SOX2 transcription factors (Boyer et al, 2005). hPSCs proliferate relatively fast, with shortened cell cycle times and a higher proportion of cells in S phase compared with most differentiated cell types (Becker et al, 2006). Differences in energy status and biosynthesis from distinct physiologies also distinguish hPSCs from differentiated cells, although the extent and regulation of these differences is unknown. Thus, metabolism could provide a relatively unexplored distinguishing feature between hPSCs and differentiated cells. Mitochondria are central organelles in carbohydrate, lipid, and amino-acid metabolism. Studies suggest that there are few, rounded, and non-fused mitochondria with underdeveloped cristae in human or mouse embryonic stem cells (ESCs) (St John et al, 2005; Cho et al, 2006; Chung et al, 2010; Prigione et al, 2010) Also, hPSCs produce more lactate than differentiated cells (Chung et al, 2010; Prigione et al, 2010), suggesting that glycolysis supercedes oxidative phosphorylation (OXPHOS) and that hPSC mitochondria may be metabolically inactive. However, this important assumption has not been thoroughly tested and it remains unclear how glycolysis and respiration contribute to the hPSC metabolic profile. Uncoupling proteins (UCPs) are mitochondrial inner membrane proteins that regulate cell metabolism (Nicholls and Rial, 1999; Klingenberg and Echtay, 2001; Brand and Esteves, 2005). UCP1 mediates proton movement from the mitochondrial intermembrane space to the matrix, which uncouples electron transport from ATP synthesis in brown fat to generate heat (Klingenberg and Huang, 1999). In contrast, the function(s) of the widely expressed UCPs, including UCP2 and UCP3, are still controversial (Brand and Esteves, 2005; Bouillaud, 2009). UCP2 and UCP3 show uncoupling activity with in vitro proteoliposome assays (Echtay et al, 2001) or when activated by fatty acids and free radical-derived alkenals (Considine et al, 2003; Brand et al, 2004a, 2004b). However, UCP2 and UCP3 have not shown physiological uncoupling activity (Cadenas et al, 2002; Couplan et al, 2002). Instead, studies show that UCP2 augments fatty acid or glutamine oxidation and decreases glucose-derived pyruvate oxidation in mitochondria (Pecqueur et al, 2008, 2009; Bouillaud, 2009; Emre and Nubel, 2010). UCP2 blocking of pyruvate entry into the tricarboxylic acid (TCA) cycle has been postulated but not experimentally validated as a mechanism for enhancing aerobic glycolysis, consistent with UCP2 expression mainly in glycolytic tissues (Pecqueur et al, 2001) and in glycolysis-switched cancer cells (Pecqueur et al, 2001; Samudio et al, 2008, 2009; Ayyasamy et al, 2011). Therefore, UCP2 could be a gatekeeper for the oxidation of carbon substrates, such as glucose. Notably, no role has been described for UCPs in hPSC bioenergetics to date. Here, we evaluated bioenergetics in human ESCs (hESCs), human-induced pluripotent stem cells (hIPSCs), and differentiated cells. We report an important role for UCP2 in regulating hPSC energy metabolism and the fate of early differentiating hPSCs. Results A conserved mitochondrial mass ratio for hPSCs and differentiated cells The distribution, abundance, fusion–fission status, and cristae structure of mitochondria regulates O2 consumption, bioenergetics, apoptosis, and autophagy (Frank et al, 2001; Narendra et al, 2008; Chan et al, 2010; Sauvanet et al, 2010). Therefore, the morphology and abundance of hPSC and fibroblast mitochondria were assessed for changes with differentiation or reprogramming. Confocal microscopy with MitoTracker Red CMXRos showed that the mitochondria in hESCs (HSF1 and H1 lines), and to a slightly lesser extent in hIPSCs (HIPS2 and HIPS18 lines), are punctate, or fragmented, compared with the well-developed filamentous network of normal human dermal fibroblasts (NHDFs, also called fibroblasts) (Figure 1A). hESCs differentiated by bFGF withdrawal also establish a filamentous mitochondrial network within days (Supplementary Figure S1A). Notably, hIPSC lines HIPS2 and HIPS18 were reprogrammed from NHDFs (Lowry et al, 2008), indicating that mitochondrial morphology is reversible with de-differentiation. Combined, the data show that mitochondrial fusion–fission and network morphology reflect the extent of cell differentiation. Figure 1.hPSC mitochondrial morphology and abundance. (A) A fragmented hPSC mitochondrial network is shown by confocal microscopy. (B) Ratio of citrate synthase enzyme activity to total cellular protein is plotted, normalized to 1.0 for HSF1. Data are expressed as mean±s.d. (n=3). (C) Representative western blot with 50 μg protein loaded per lane. Download figure Download PowerPoint The F1F0 ATP synthase appears to play a scaffolding role for the mitochondrial inner membrane cristae structure and impacts OXPHOS activity and cellular ATP levels (Giraud et al, 2002; Paumard et al, 2002; Minauro-Sanmiguel et al, 2005; Buzhynskyy et al, 2007; Campanella et al, 2008; Strauss et al, 2008). Therefore, the cristae structure of hPSCs was examined as a potential indicator of mitochondrial function. Transmission electron microscopy shows that hPSC cristae are swollen, circular, and disorganized compared with the linear, stacked cristae observed in many differentiated cell types, including fibroblasts (Supplementary Figure S1B). These features are similar to those seen in mouse ESCs and for a different set of hPSCs (Baharvand and Matthaei, 2003; Prigione et al, 2010). Less orderly cristae and a fragmented network suggests that PSC mitochondria are less functional than differentiated cell mitochondria. Prior studies reported few mitochondria in hESCs (St John et al, 2005; Facucho-Oliveira et al, 2007). However, the ratio of mitochondrial to total cell protein mass is a more informative comparison between cell types and stages. Citrate synthase marks the mitochondrial matrix (Morgunov and Srere, 1998) and is commonly used as an indicator of mitochondrial function and mass. Citrate synthase enzymatic activity normalized to total protein is similar between hPSCs, hPSCs induced to differentiate by retinoic acid (RA) (Pan et al, 2007), and fibroblasts (Figure 1B; Supplementary Figure S1C). Also, the translocons of the inner (TIM23) and outer (TOM40) mitochondrial membranes accurately reflect mitochondrial mass and both are expressed at similar levels relative to the total protein content in hPSCs, RA-differentiated hPSCs, and fibroblasts (Figure 1C; Supplementary Figure S1D). Fibroblast cell diameters are larger (∼25–35 μm) and cell volumes (assuming a spherical shape) are much larger (∼16 000 μm3) than hPSCs (∼15–20 μm and ∼3000 μm3, respectively) (Supplementary Figure S1E). Fibroblasts also have 1.5- to 2-fold more protein than hPSCs, suggesting on a per cell basis there will be more mitochondrial mass in fibroblasts than in hPSCs, as reported (St John et al, 2005; Facucho-Oliveira et al, 2007). However, a similar mitochondrial mass per unit protein for hESCs, hIPSCs, and fibroblasts indicates that hPSCs and differentiated cells are equally committed to generating or maintaining mitochondria. hPSC and differentiated cell mitochondria show similar O2 consumption rates hPSCs were examined to determine how much O2 they consume. O2 consumption rate (OCR) studies showed that intact hPSCs consume O2 in room air at ∼5–8 nmol O2/min/5 × 106 cells (or ∼1.0–1.6 fmol/min/cell) (Figure 2A). By contrast, fibroblasts consume O2 at ∼13–15 nmol O2/min/5 × 106 cells (or ∼2.6–3.0 fmol/min/cell) in room air, an ∼2-fold higher rate than hPSCs. However, because there is ∼1.5- to 2-fold more mitochondrial mass per fibroblast than hPSC, the OCR normalized to mitochondrial mass is ∼equivalent between fibroblasts and hPSCs. Figure 2.hPSCs consume O2 and are programmed for glycolysis. (A) OCR in room air measured with a Clark-type O2 electrode in intact cells. Data are expressed as mean±s.d. (n=3), *P<0.05. (B) OCR to ECAR ratios measured by the XF24 extracellular flux analyser is shown. Data are expressed as mean±s.d. (n=3), *P<0.005. (C) ECAR normalized to protein concentration is shown. Data are expressed as mean±s.d. (n=3), *P<0.05. (D) Cells were incubated with 150 mM 2-DG for 45 min, followed by flow cytometry and cell-cycle profile analyses. (E) HSF1 cells were induced to differentiate by 10 μM RA for 4 days and ECAR determined. Data are expressed as mean±s.d. (n=3), *P<0.05. Download figure Download PowerPoint Electron transport chain (ETC) complexes I–IV transfer electrons from substrates to O2 and the activity of each complex can vary between cell types. Substrate feeding into each ETC complex was performed with digitonin-treated HSF1 and HEK293 cells to determine relative complex activities. HEK293 cells were chosen because of a similar mitochondrial content (Figure 1B) and size (Supplementary Figure S1E) relative to hPSCs and for ease of substrate feeding compared with fibroblasts. Results show that ETC complex I–IV activities are similar between HSF1 and HEK293 cells (Supplementary Figure S2A) and, overall, the data indicate that hPSCs have a functional respiratory chain and OCR similar to fibroblasts when normalized to mitochondrial mass. hPSCs are programmed for glycolysis Different cell types in distinct microenvironments rely upon different bioenergetic processes for ATP production (Jezek et al, 2010). hPSCs and fibroblasts were therefore analysed for respiration and glycolysis in the same environment. To do this, OCR and the extracellular acidification rate (ECAR), which can approximate glycolysis from lactate production (Xie et al, 2009), were determined using a XF24 Extracellular Flux Analyser (Wu et al, 2007). The XF24 measures the rates of change in pmol O2 and mpH simultaneously for cultured cells, providing a comparison for a defined cell population. For accurate analyses, hPSCs were plated with ROCK inhibitor (Watanabe et al, 2007) to form monolayers. Control or ROCK inhibitor-treated fibroblasts showed no statistical difference in the OCR/ECAR ratio (Supplementary Figure S2B). The number of cells seeded per well was adjusted for the linear range of measurements (Supplementary Figure S2C and D). The OCR/ECAR ratios at two cell densities for hESCs and hIPSCs are ∼50–65% lower than for fetal-derived NHDFs (Figure 2B) and ECAR values for hPSCs are ∼1.5- to 2-fold higher than for fibroblasts (Figure 2C). Analysis of adult-origin fibroblasts (NHDFs) showed results that were statistically similar to those obtained with fetal-derived fibroblasts (Supplementary Figure S2E). These data suggest that hPSC bioenergetics favours glycolysis over OXPHOS, in contrast to fibroblasts. Addition of the competitive glycolysis inhibitor, 2-deoxyglucose (2-DG), causes hPSCs to G2 arrest (Figure 2D) and to minimally increase apoptosis (Supplementary Figure S2F) rather than shifting to respiration in room air, suggesting an immutable energetic preference. To determine a glycolytic transition with differentiation, HSF1 cells were exposed to RA and ECAR measured over time. Results showed a gradual and significant decrease in glycolysis with differentiation, with decreased ECAR (Figure 2E) and lactate in the culture media (Supplementary Figure S2G), indicating a progressive metabolic reprogramming during differentiation. Notably, hPSC OCRs and ECARs are mainly unaffected by 5–25 mM glucose (Supplementary Figure S2H) or by 0–17.5 mM insulin (Supplementary Figure S2I) incubations, indicating that the glucose supplied in these studies is not limiting. Furthermore, glucose transporters GLUT1 (widely expressed) and GLUT4 (insulin-regulated) are expressed at ∼equivalent high or barely detectable levels in hPSCs and fibroblasts, respectively (Supplementary Figure S2J and K), indicating an equivalent ability to bind and uptake glucose. Combined, the data strongly suggest that differences in OCRs and ECARs between hPSCs and differentiated cells are not from differences in glucose uptake but rather from unique intracellular metabolic programmes. hPSCs are more sensitive to glycolysis inhibitors than are differentiated cells OXPHOS was inhibited with oligomycin, a F1F0 ATP synthase inhibitor. Fibroblasts showed an ∼80% reduction in O2 consumption within 10 min (Figure 3A). By contrast, hPSCs showed an ∼50% decrease in OCR, suggesting that hPSC are less dependent on OXPHOS than fibroblasts. Notably, there is a greater compensatory increase in ECAR for differentiated cells than for hPSCs with oligomycin exposure (Supplementary Figure S3A). Supplementary glucose (from 2.5 mM to 25 mM) maximized ECAR, with fibroblasts increasing ∼55% in 20 min, whereas hPSC ECAR increased by a more modest ∼20–30% (Figure 3B), indicating that hPSCs are closer to their maximal glycolytic capacity compared with fibroblasts. Following additions of 2-DG, which competes for uptake with glucose, hPSC glycolysis was repressed ∼65% from the ECAR maximum within 34 min, whereas fibroblast glycolysis did not change for 6 min and then fell by only ∼45% from the ECAR maximum after 34 min (Figure 3B), indicating that the hPSC glycolysis level is greater than in fibroblasts. Combined, the data indicate that hPSCs are less sensitive to OXPHOS inhibition, more sensitive to glycolysis disruption, and function closer to their maximum glycolytic capacity than do fibroblasts. Figure 3.hPSCs require glycolysis for ATP production despite maximal O2 consumption. (A) OCR change with 1 μM oligomycin for 10 min. Data are expressed as mean±s.d. (n=3), *P<0.05. (B) Cells were cultured in assay medium with 2.5 mM glucose. ECAR change with glucose (25 mM final) and 2-DG (150 mM final) additions is shown. Arrows indicate percentage reduction from maximal ECAR (n=3), *P<0.05. (C) Change in cellular ATP with increasing doses of oligomycin at 45 min (n=3), *P<0.05. (D) Change in cellular ATP with increasing doses of sodium oxamate at 45 min (n=3), *P<0.05. (E) OCR changes in response to F1F0 ATP synthase inhibitor (1 μM oligomycin), uncoupler (0.3 μM FCCP), and ETC blockade (1 μM rotenone plus 1 μM antimycin) (n=3), *P<0.05. OCR due to non-mitochondrial sources is the distance between the post-rotenone+antimycin OCR (time points 127–146 min) and the abscissa. (F) OCR change in response to increasing doses of palmitate (n=3), *P<0.05. Download figure Download PowerPoint ATP in hPSCs is mainly produced by glycolysis The steady-state ATP level is ∼2-fold higher per cell in fibroblasts than in hPSCs, reflecting ATP production, consumption, and the size difference between these two cell types (Supplementary Figure S3B). OXPHOS inhibition with 0.1 μM oligomycin caused an ∼30% ATP drop in fibroblasts, whereas the ATP drop in hPSCs was <5% (Figure 3C), suggesting that hPSCs produce minimal ATP by OXPHOS. By contrast, sodium oxamate, a lactate dehydrogenase inhibitor that blocks ATP production by glycolysis, caused the ATP level in hPSCs to drop >60%, whereas fibroblasts showed <40% ATP reduction (Figure 3D), indicating that hPSCs depend more on glycolysis for ATP production than do fibroblasts. ATP reductions with short-term inhibitor exposures are not due to a significant increase in cell death (Supplementary Figure S3C). Combined, the data indicate that the ATP level in hPSCs is sensitive to glycolysis inhibition. hPSCs consume O2 at maximal capacity Although hPSC and fibroblast mitochondria consume O2 at an ∼equivalent rate, it is unclear what proportion of their maximal electron transport capacity is being utilized. FCCP is a mitochondrial uncoupler (protonophore) that dissipates the mitochondrial membrane potential (Δψ) to stimulate maximal electron transport and O2 consumption (Heytler, 1979). hPSCs failed to increase OCR, whereas fibroblasts showed an ∼90–100% increase in OCR at 0.1–0.4 μM FCCP exposures (Supplementary Figure S3D). To establish a baseline at which coupled respiration is inhibited, HSF1 cells and fibroblasts were exposed to 1 μM oligomycin to prevent proton movement through the F1F0 ATP synthase (Figure 3E). No significant changes in cell viability were detected during the course of oligomycin exposure (Supplementary Figure S3C). Addition of 0.3 μM FCCP re-establishes proton movement and results in maximal O2 consumption. Then, incubation with rotenone and antimycin (complex I and III inhibitors, respectively) completely blocked mitochondrial respiration. The maximal mitochondrial respiration capacity is the difference between blocked and FCCP-induced OCR. Maximal hPSC respiration is equivalent to its pre-oligomycin respiration, which contrasts with the up to ∼100% increase in OCR measured for fibroblasts (Figure 3E). OCR does not increase further for hPSCs with increased sodium pyruvate supplemented media and FCCP exposure, excluding limited substrate as a cause for an hPSC ceiling (Supplementary Figure S3E). Because oligomycin inhibits and FCCP stimulates OCR in coupled mitochondria, the smaller difference between FCCP-stimulated and oligomycin-inhibited OCR also suggests that hPSC mitochondria could be less coupled for ATP synthesis than fibroblast mitochondria (Supplementary Figure S3E). Finally, free fatty acid palmitate, an OXPHOS substrate, was added to cells incubated with L-carnitine, which transports fatty acids from the cytosol into the mitochondria. In contrast to fibroblasts that showed a dose-dependent OCR increase of ∼35%, HSF1 cells showed only a dose-independent ∼5% OCR increase (Figure 3F). The lack of a robust free fatty acid response in HSF1 cells could be from ETC saturation with glucose or internally produced fatty acids. Notably, the essential enzymes for fatty acid oxidation are equally expressed in HSF1 cells and fibroblasts by QPCR analysis. Combined, these data show that hPSCs consume O2 at maximal capacity. Glycolytic ATP hydrolysis helps to maintain Δψ in hPSCs A reduction in Δψ has been linked to mitochondrial dysfunction and possible mitophagy (Narendra et al, 2008). The presence of PARKIN protein expression in HSF1 cells that increases with RA-induced differentiation is consistent with the need to maintain Δψ in hPSCs (Supplementary Figure S4A), although this suggestion requires further investigation (Narendra et al, 2008, 2010). In differentiated cells, Δψ is maintained by a proton gradient formed during the passage of electrons through the ETC. Alternatively, Δψ is maintained by hydrolysis of glycolytic ATP in the F1F0 ATP synthase when ETC function is reduced (Nicholls and Lindberg, 1972; Hatefi, 1985). For mouse ESCs, the relative level of Δψ has a role in regulating differentiation (Schieke et al, 2008). Therefore, the mechanism(s) for hPSC Δψ maintenance was examined. Immunofluorescence microscopy showed an intact Δψ for all HSF1 mitochondria imaged that was dissipated with 100 μM FCCP (Supplementary Figure S4B). To determine what portion of Δψ is provided by electron transport versus hydrolysis of glycolytic ATP, antimycin was used to dissipate the ETC-generated component of Δψ and sodium oxamate was used to dissipate the glycolysis-generated component of Δψ, followed by quantification using Δψ-sensitive TMRM staining and flow cytometry. Antimycin and FCCP equivalently and robustly dissipate Δψ in fibroblasts, whereas antimycin only partially dissipates Δψ in hPSCs (Figure 4A; Supplementary Figure S4C, with JC-1 staining). Sodium oxamate partially dissipates Δψ in hPSCs but had no effect on Δψ in fibroblasts (Figure 4B). Combined, the data indicate that Δψ in fibroblasts is