As human pluripotent stem cells (hPSCs) exit pluripotency, they reportedly switch from glycolytic energy production to primarily mitochondrial metabolism. Here, we show that upon ectoderm differentiation to neural precursor cells (NPCs), hPSCs increase glycolytic rate, ultimately producing more carbon as lactate than is consumed as glucose. However, glucose, lactate and pyruvate utilization decrease to half their PSC levels by the NPC stage, establishing a more quiescent metabolic state. Furthermore, we characterize a metabolic exit event within the first 24 h of differentiation, plausibly necessary to transition hPSCs out of the pluripotent state. Contrary to current thinking, mitochondrial mass does not increase during NPC induction. Instead, mitochondrial DNA copies and mitochondrial activity decrease, suggesting that mitochondrial metabolism either requires suppression, or is not required, for nascent ectoderm differentiation. Our work, therefore, contrasts with the dogma that the hPSC state is primarily glycolytic, transitioning to an oxidative metabolism upon the loss of the pluripotent state. Instead, we show that heightened glycolytic metabolism is acquired, indicating that metabolic modulation of both glycolysis and mitochondrial metabolism occurs during exit from pluripotency in hPSCs.

Human pluripotent stem cells (hPSCs) and human induced pluripotent stem cells (hiPSCs) exhibit a heavy dependency on glycolysis, with 70-90% of the consumed glucose accounted for as lactate (Zhang et al., 2011; Zhou et al., 2012; Lees et al., 2015; Harvey et al., 2016). In contrast, mitochondrial oxidative phosphorylation (OxPhos) occurs at relatively low levels in hPSCs compared with their differentiated counterparts (Cho et al., 2006; Varum et al., 2011). It is generally accepted that as PSCs exit pluripotency, they undergo a metabolic switch from a primarily glycolytic pluripotent state, to an OxPhos-driven differentiated state (St John et al., 2005; Chen et al., 2010; Chung et al., 2010; Hattori et al., 2010; Prigione and Adjaye, 2010; Varum et al., 2011; Zhang et al., 2011; Mathieu and Ruohola-Baker, 2017).

However, this metabolic switch is primarily based on studies comparing hPSCs with terminally differentiated somatic cells, and is dominated by mesoderm and endoderm protocols. Interestingly, in vitro mesoderm and endoderm development is unaffected by inhibition of glycolysis, whereas, in contrast, hPSCs differentiated with glycolytic inhibitors fail to make neural tissue (Cliff et al., 2017). Notably, of the few studies to examine metabolic events following the exit from pluripotency, none has examined both the glycolytic and mitochondrial transition with the necessary sampling resolution to determine when the changes in metabolic state occur. Collectively, these studies suggest that glycolytic metabolism plays an important role in neural differentiation, consistent with the metabolic profile of ectoderm development in vivo.

The hPSC-derived NPC state corresponds to embryonic day (E) 8.5 in the developing mouse egg cylinder when neural crest development takes place (Chambers et al., 2009; Chen et al., 2017). From implantation at E5, through neural tube, neural crest, and brain vesicle development at E7-E9.5, mouse embryo metabolism is almost exclusively glycolytic (Clough and Whittingham, 1983). Notably, the concentration of oxygen, a key nutrient in mitochondrial OxPhos, declines throughout embryonic development from ∼5% in the uterus to ∼2.5% in the uterine wall at implantation (Maas et al., 1976; Genbacev et al., 1997). Significantly, these data cast doubt on the adoption of an oxidative metabolism during hPSC ectoderm differentiation, suggesting instead that glycolytic metabolism increases by necessity.

To determine the metabolic requirements of ectoderm differentiation, we differentiated hPSCs to neural precursor cells (NPCs) in a defined medium, sampling mitochondrial and glycolytic metabolism every 24 h. Glucose, lactate and pyruvate utilization declined to half their PSC levels by the NPC stage, and mitochondrial metabolism, assessed by mitochondrial membrane potential, mitochondrial DNA (mtDNA), and reactive oxygen species (ROS) production, decreased with nascent ectoderm differentiation. Overall, our findings show that modulation of both glycolysis and mitochondrial metabolism occurs during exit from pluripotency in hPSCs, and that glycolysis is the preferred metabolic pathway during nascent ectoderm differentiation.

NPC induction in a defined medium

hPSC differentiation is reported to coincide with loss of aerobic glycolysis and acquisition of mitochondrial oxidative metabolism (Varum et al., 2011; Zhang et al., 2011; Gu et al., 2016; Mathieu and Ruohola-Baker, 2017). This process, however, has been evaluated primarily in mesoderm inductions and typically assesses only the pluripotent and somatic cell states, raising questions about the coupling of metabolism to pluripotency exit in ectoderm. To address this, we differentiated two hPSC lines that have undergone extensive metabolic characterization, MEL1 and MEL2 (Lees et al., 2015; Harvey et al., 2016), to NPCs (Fig. 1A). To examine metabolism in response to differentiation, rather than in response to changes in nutrient concentrations, an established dual SMAD-inhibition neural induction protocol (Chambers et al., 2009; Drury-Stewart et al., 2011, 2013) was modified replacing KOSR with the hPSC maintenance medium mTeSR1. Induction of NPC identity by day 11 using the modified protocol was confirmed by the presence of the neural stem cell markers PAX6, SOX1 and SOX2, and loss of pluripotency marker OCT4 (POU5F1), at levels equivalent to those obtained by KOSR induction (Fig. 1B). The potency of the derived NPC populations was validated by the formation of neurospheres from day 11 NPCs with high expression of neural development genes (Fig. 1C,D; Denham and Dottori, 2011).

Fig. 1.

Neural progenitor cell (NPC) induction in a defined medium. (A) Neural differentiation scheme. hPSCs were treated with dorsomorphin for 11 days and SB431542 for 5 days. On days 5, 7 and 9, N2 medium was incorporated at 25%, 50% and 75%, respectively. Cells were harvested for analysis on the days marked with an asterisk. Images show representative examples of cell morphology. (B) Immunostaining of PAX6, SOX1, SOX2 and OCT4 in mTeSR-cultured cells, and quantification (mean±s.e.m.) of these markers in mTeSR-cultured cells compared with KOSR-cultured cells. (C) Changes in gene expression in neurospheres from day 11 NPCs presented as minimum, maximum, median, 25% and 75% quartiles. Data are log10 (fold change relative to PSCs) ±s.e.m. n=4. (D) Representative neurospheres after 1 (i), 4 (ii), 8 (iii) and 11 (iv) days. (E,F) Expression of the pluripotency markers TG30 (E) and GCTM2 (F) in hPSCs cultured under physiological (5%) versus atmospheric (20%) levels. Mean±s.e.m. are shown for each time point. Gradients for the line of best fit were tested for a significantly non-zero slope. (G) Mature neurons derived from NPCs on day 21 of differentiation. Red, neuronal marker type III β-tubulin; blue, nuclear marker DAPI. Scale bars: 200 μm (A); 75 μm (B); 250 μm (D); 200 μm (G). ***P<0.001, **P<0.01, *P<0.05.

Fig. 1.

Neural progenitor cell (NPC) induction in a defined medium. (A) Neural differentiation scheme. hPSCs were treated with dorsomorphin for 11 days and SB431542 for 5 days. On days 5, 7 and 9, N2 medium was incorporated at 25%, 50% and 75%, respectively. Cells were harvested for analysis on the days marked with an asterisk. Images show representative examples of cell morphology. (B) Immunostaining of PAX6, SOX1, SOX2 and OCT4 in mTeSR-cultured cells, and quantification (mean±s.e.m.) of these markers in mTeSR-cultured cells compared with KOSR-cultured cells. (C) Changes in gene expression in neurospheres from day 11 NPCs presented as minimum, maximum, median, 25% and 75% quartiles. Data are log10 (fold change relative to PSCs) ±s.e.m. n=4. (D) Representative neurospheres after 1 (i), 4 (ii), 8 (iii) and 11 (iv) days. (E,F) Expression of the pluripotency markers TG30 (E) and GCTM2 (F) in hPSCs cultured under physiological (5%) versus atmospheric (20%) levels. Mean±s.e.m. are shown for each time point. Gradients for the line of best fit were tested for a significantly non-zero slope. (G) Mature neurons derived from NPCs on day 21 of differentiation. Red, neuronal marker type III β-tubulin; blue, nuclear marker DAPI. Scale bars: 200 μm (A); 75 μm (B); 250 μm (D); 200 μm (G). ***P<0.001, **P<0.01, *P<0.05.

NPC induction under physiological and atmospheric oxygen conditions

The earliest neural developments of the implanted mouse embryo, starting at E7 (Chen et al., 2017), take place at physiological (<5%) oxygen concentrations (Genbacev et al., 1997; Caniggia et al., 2000). To determine whether physiological oxygen would alter hPSC metabolism or differentiation kinetics throughout NPC induction, we induced hPSCs adapted for over ten passages at 5% and 20% oxygen to differentiate to NPCs at the same oxygen level. hPSCs displayed a pronounced reduction in expression of the pluripotency markers TG30 and GCTM2, which detect CD9 and PODXL, respectively (Laslett et al., 2007) within the first 24 h, followed by an almost commensurate increase by 48 h regardless of oxygen levels (Fig. 1E,F). Following this, pluripotency markers were progressively lost at 5% and 20% oxygen reaching near-zero levels by day 7, consistent with an exit from pluripotency and acquisition of a differentiated NPC state. No effect of oxygen on exit from pluripotency was observable in pluripotency cell surface marker expression. To confirm the in vitro maturation potential of the derived NPCs, day 11 NPCs were cultured for a further 10 days in monolayer culture conditions in neural maturation medium. By day 21, class III β-tubulin-positive neural processes and complex network formation were observable in both cell lines at 5% and 20% oxygen (Fig. 1G). These data confirm that induction of NPCs from hPSCs using a dual small molecule inhibitor approach in a defined mTeSR1-base medium is equivalent to induction protocols using KOSR and noggin and/or dorsomorphin and SB431542, and is capable of generating mature populations of neurons.

NPC induction reduces mitochondrial activity and investment

During PSC differentiation, mitochondria reportedly develop into a reticulated, mature network (St John et al., 2005; Chung et al., 2007), similar to that observed in fibroblast or liver cells (Varum et al., 2011). Notably, there are very few studies examining the role of the mitochondria in ectoderm differentiation from PSCs (Birket et al., 2011), and none has been conducted at physiological oxygen levels. To examine mitochondrial metabolism as hPSC exit pluripotency to ectoderm, we assessed mitochondrial investment and activity every 24 h during NPC differentiation. mtDNA copy number peaked 24 h after the initiation of neural induction in all treatments, corresponding to an average increase of 20% relative to the pluripotent hPSC stage on day 0, followed by a significant linear decline to the NPC stage (Fig. 2A, Table S1). Total mtDNA copy number over the differentiation period, corresponding to the investment in mitochondrial metabolism, was significantly higher under 5% oxygen conditions compared with 20% oxygen (Fig. 2B). Consistent with observations of mtDNA, peak mitochondrial mass occurred 24 h into the neural induction, corresponding to an average increase of 25% relative to the pluripotent hPSC stage (Fig. 2C). Following this, mitochondrial mass underwent a significant decrease of ∼50% by day 5; however, levels plateaued from day 5 onward. Total mitochondrial mass throughout NPC induction, calculated as area under the curve (AUC), was 30-50% higher at 5% oxygen (Fig. 2D). These observations confirm that mitochondrial investment declines during the very early stages of hPSC differentiation, and that the requirement for mitochondrial machinery remains low during the formation of nascent ectoderm lineages.

Fig. 2.

NPC induction reduces mitochondrial activity and investment. (A,C,E,G) Changes in mtDNA copy number (A), mitochondrial mass (C), MMP (E) and mitochondrial superoxide levels (G) during differentiation of hPSCs to NPCs under physiological or atmospheric oxygen levels. Gradients for the line of best fit were tested for a significantly non-zero slope. (B,D) Total mtDNA copy number (B) and total mitochondrial mass (D) over the time course of differentiation (calculated as AUC). (F,H) Average MMP (F) and average mitochondrial superoxide levels (H) throughout differentiation. Data are mean±s.e.m. n=3. ****P<0.0001, **P<0.01, *P<0.05.

Fig. 2.

NPC induction reduces mitochondrial activity and investment. (A,C,E,G) Changes in mtDNA copy number (A), mitochondrial mass (C), MMP (E) and mitochondrial superoxide levels (G) during differentiation of hPSCs to NPCs under physiological or atmospheric oxygen levels. Gradients for the line of best fit were tested for a significantly non-zero slope. (B,D) Total mtDNA copy number (B) and total mitochondrial mass (D) over the time course of differentiation (calculated as AUC). (F,H) Average MMP (F) and average mitochondrial superoxide levels (H) throughout differentiation. Data are mean±s.e.m. n=3. ****P<0.0001, **P<0.01, *P<0.05.

hPSCs undergo a metabolic exit event following the onset of differentiation

The onset of differentiation and concurrent exit from pluripotency appear to coincide with a peak in mitochondrial activity and a trough in pluripotency markers. Alterations to metabolism and the mitochondria precede lineage marker changes during hPSC differentiation (Mandal et al., 2011; Zhou et al., 2012), suggesting that metabolic state drives differentiation and exit from pluripotency. To assess the potential regulatory role of the mitochondria in hPSC differentiation, we assessed mitochondrial activity by examining mitochondrial membrane potential (MMP) and superoxide production. In support of the decreasing mitochondrial investment, mitochondrial activity declined significantly throughout nascent ectoderm specification at 5% oxygen (Fig. 2E, Table S1), whereas at 20% oxygen, mitochondrial activity increased until day 7 before a sharp decline; average MMP was also higher at 20% oxygen (Fig. 2F). Peak mitochondrial superoxide levels were observed 24 h into differentiation at both 5% and 20% oxygen (Fig. 2G), though 5% cultures had higher average levels of mitochondrial superoxides throughout differentiation (Fig. 2H). Elevated mitochondrial ROS may reflect the disorganized cristae structure of pluripotent mitochondria (Cho et al., 2006; Lees et al., 2017); however, ROS are also well-established signaling molecules, proposed to regulate pluripotency and signaling at low oxygen (Lees et al., 2017). Despite the clear profile of increasing quiescence during hPSC ectoderm differentiation, a sharp peak in metabolic parameters, including ROS, and a trough in pluripotency markers occurred within the first 24 h of differentiation. This metabolic exit event has been missed by other metabolic assessments of differentiation that only assess every 48 h or longer (Cho et al., 2006; Fernandes et al., 2010; Prigione and Adjaye, 2010; Varum et al., 2011; Mondragon-Teran et al., 2013). Similar to the waves of transcriptional and proteomic remodeling that take place during reprogramming to iPSCs (Hansson et al., 2012; Polo et al., 2012), this metabolic exit event may be necessary to shift PSCs out of their stable primed state (Enver et al., 2009), thus marking the onset of differentiation.

Glucose-derived carbon utilization decreases with nascent ectoderm differentiation

The switch from aerobic glycolysis to mitochondrial OxPhos is believed to be an obligatory event during the differentiation of PSCs (Varum et al., 2011; Zhang et al., 2011; Gu et al., 2016; Mathieu and Ruohola-Baker, 2017). However, we have already established during nascent ectoderm specification that mitochondrial metabolism does not increase to support the altered cell state. A recent study has determined that hPSC differentiation to the NPC state requires glycolytic metabolism (Cliff et al., 2017). Thus, we set out to understand the contribution of glycolytic metabolism to hPSC ectoderm differentiation by assessing glucose and lactate utilization.

Glucose and lactate utilization decreased throughout NPC induction, generally reaching a plateau after day 5 (Fig. 3A,B). Consistent with the metabolic exit event observed in the mitochondria and pluripotency markers, a peak in glucose, lactate and pyruvate utilization was observed 24 h into NPC differentiation. Initial glucose consumption and lactate production by undifferentiated hPSCs were 2-fold higher at 5% than 20% oxygen, consistent with the accepted regulatory effect of oxygen on glucose transporters and lactate dehydrogenase enzymes (Harvey et al., 2016). Furthermore, the impact of oxygen was maintained at the NPC stage, with glucose and lactate utilization at 5% oxygen ∼2-fold and ∼2.7-fold higher, respectively, than those at 20% oxygen. As a result, total glucose and lactate utilization over the induction period was ∼3-fold higher under physiological oxygen conditions (Fig. 3C,D). Similar to the glucose profile, pyruvate consumption decreased throughout differentiation (Fig. 3E), and approximately 3-fold more pyruvate was consumed under 5% oxygen than under 20% oxygen conditions (Fig. 3F). A decline in glucose consumption has been observed in mouse PSCs and mouse naïve PSC neural differentiation has been charted every 48 h (Fernandes et al., 2010; Mondragon-Teran et al., 2013); however, neither study contrasted mitochondrial metabolism with glycolytic metabolism, and both altered the medium base for neural inductions. Our findings establish the acquisition of a more quiescent metabolism upon PSC exit from pluripotency, in which carbohydrate utilization is halved by the NPC state.

Fig. 3.

Carbohydrate utilization decreases, but the glycolytic contribution to metabolism increases, with nascent ectoderm differentiation. (A,B,E,G) Glucose consumption (A), lactate production (B), pyruvate consumption (E) and glycolytic rate (G) during hPSC to NPC differentiation under physiological or atmospheric oxygen levels. Gradients for the line of best fit were tested for a significantly non-zero slope. (C,D,F,H) Total glucose consumption (C), lactate production (D) and pyruvate consumption (F) (calculated as AUC) and average glycolytic rate (H) throughout NPC differentiation. Data are mean±s.e.m. n=3. ****P<0.0001, ***P<0.001, **P<0.01, *P<0.05.

Fig. 3.

Carbohydrate utilization decreases, but the glycolytic contribution to metabolism increases, with nascent ectoderm differentiation. (A,B,E,G) Glucose consumption (A), lactate production (B), pyruvate consumption (E) and glycolytic rate (G) during hPSC to NPC differentiation under physiological or atmospheric oxygen levels. Gradients for the line of best fit were tested for a significantly non-zero slope. (C,D,F,H) Total glucose consumption (C), lactate production (D) and pyruvate consumption (F) (calculated as AUC) and average glycolytic rate (H) throughout NPC differentiation. Data are mean±s.e.m. n=3. ****P<0.0001, ***P<0.001, **P<0.01, *P<0.05.

The glycolytic contribution to metabolism increases with nascent ectoderm differentiation

The decline in glucose and lactate utilization described herein contrasts with the reported glycolytic requirement for ectoderm specification (Cliff et al., 2017). Specifically, Cliff et al. report that total inhibition of glycolysis with 3-bromo-pyruvate, or rendering glycolysis ineffective with 2-deoxyglucose, inhibits ectoderm formation. Glycolytic rate, determined by the relative concentrations of glucose and lactate (Lane and Gardner, 1996; Harvey et al., 2016; Lees et al., 2015), consistently increased at both 5% and 20% oxygen over the NPC induction period (Fig. 3G), making it the sole metabolic parameter to increase during ectoderm differentiation of hPSCs in this study. At physiological oxygen, both cell lines sharply increased glycolytic rate until day 4, followed by a gradual increase from days 7 to 11, whereas at 20% oxygen there was a distinct peak at 24 h followed by a gradual increase over the remaining differentiation period. The increasing dominance of glycolysis throughout in vitro ectoderm differentiation is consistent with the metabolic development of the post-implantation mammalian embryo in vivo, where oxygen consumption drops following implantation (Houghton et al., 1996) and glycolysis becomes the primary metabolic pathway (Clough and Whittingham, 1983). NPCs proliferate up to twice as fast as their parental hPSCs (Koch et al., 2009; Birket et al., 2011), plausibly generating lactate to regenerate cytosolic NAD+ to support proliferation (Gardner and Harvey, 2015). The percentage of glucose converted to lactate during NPC induction reached ∼100% under atmospheric oxygen and over 125% under physiological oxygen conditions (Fig. 3G,H) indicating contributions from either glutaminolysis or glycogen catabolism, and consistent with rates observed in post-implantation mouse and rat embryos (>90% and 100%, respectively) at the time of neural tube and neural crest development (Gunberg, 1976; Clough and Whittingham, 1983; Ellington, 1987). Overall, these results indicate that nascent ectoderm differentiation increases the glycolytic contribution to metabolism to levels greater than those observed in pluripotent cells.

Physiological oxygen exerts a persistent effect on metabolism through differentiation

We have shown that as hPSCs exit the pluripotent state to nascent ectoderm, they actively increase the glycolytic contribution to metabolism while reducing mitochondrial investment. At the post-implantation stage of embryonic development, a reliance on anaerobic metabolism and minimal oxygen consumption by the embryo, suggest that in vivo neural development takes place at low oxygen before adequate vascularization has been established. We did not find that physiological oxygen altered the kinetics of neural differentiation in our system; however, oxygen-induced differences in metabolism persisted through to the NPC stage. Principal component analysis (PCA) using all measured parameters indicates that despite the adoption of the NPC state and a general decline in metabolic activity, 5% and 20% oxygen trajectories remained separate (Fig. 4A). When exiting pluripotency to ectoderm, both glycolytic and mitochondrial metabolism are remodeled in an oxygen-specific manner. Our findings indicate that the nutrient environment in which differentiation occurs has a persistent effect that impacts both the pluripotent cell and its differentiated derivatives (Mills et al., 2017). The metabolic exit event observed in the mitochondria, carbohydrate utilization, and pluripotency markers, was clear at both 5% and 20% oxygen (Fig. 4A) and was evident in both hPSC lines (Fig. 4B). Cell line-specific metabolism converged from day 5 onwards (Fig. 4B), highlighting the robustness of the induction protocol in generating homogeneous NPC populations from discrete hPSCs.

Fig. 4.

A new metabolic model for nascent ectoderm differentiation. (A,B) Principal component analyses (PCA) incorporating pluripotency markers GCTM2 and TG30 (CD9), mitochondrial mass, mtROS, mtDNA, glucose, lactate and pyruvate utilization, and glycolytic rate parameters throughout hPSC to NPC differentiation isolating 5% and 20% oxygen treatments (A), and isolating cell lines (B). (C) Conventional model of metabolism throughout differentiation based primarily on mesoderm/endoderm inductions describing increasing mitochondrial metabolism and declining glycolytic metabolism. (D) Revised nascent ectoderm metabolic model of differentiation. The hPSC state comprises dual glycolytic and mitochondrial metabolism. Upon ectoderm induction, there is a transient peak in glycolytic metabolism and mitochondrial mass, mtDNA and superoxide production, before overall metabolism declines, although the relative proportion of glycolytic metabolism increases. Question marks indicate that it is currently unknown how precursor cell metabolism transitions to somatic cell metabolism.

Fig. 4.

A new metabolic model for nascent ectoderm differentiation. (A,B) Principal component analyses (PCA) incorporating pluripotency markers GCTM2 and TG30 (CD9), mitochondrial mass, mtROS, mtDNA, glucose, lactate and pyruvate utilization, and glycolytic rate parameters throughout hPSC to NPC differentiation isolating 5% and 20% oxygen treatments (A), and isolating cell lines (B). (C) Conventional model of metabolism throughout differentiation based primarily on mesoderm/endoderm inductions describing increasing mitochondrial metabolism and declining glycolytic metabolism. (D) Revised nascent ectoderm metabolic model of differentiation. The hPSC state comprises dual glycolytic and mitochondrial metabolism. Upon ectoderm induction, there is a transient peak in glycolytic metabolism and mitochondrial mass, mtDNA and superoxide production, before overall metabolism declines, although the relative proportion of glycolytic metabolism increases. Question marks indicate that it is currently unknown how precursor cell metabolism transitions to somatic cell metabolism.

A new metabolic model for nascent ectoderm differentiation

The overall findings of this study show that the conventional metabolic model of hPSC differentiation is not applicable to nascent ectoderm development (Fig. 4C). Most analyses of PSC metabolic differentiation have been performed in mesoderm or endoderm, describing a switch from an exclusively glycolytic PSC state to a more active and OxPhos-dominated differentiated state (Mathieu and Ruohola-Baker, 2017). In this report, we have established a new metabolic model for nascent ectoderm differentiation (Fig. 4D), contrasting glycolytic metabolism with mitochondrial mass and activity with 24 h resolution. Our findings show that hPSCs exiting pluripotency to nascent ectoderm adopt a more quiescent glycolytic and oxidative metabolism, halving their nutrient intake by the NPC stage. Mitochondrial mass, activity and genome replication do not increase during differentiation; instead, mtDNA copies and mitochondrial activity decline. Despite a continuous decrease in glycolytic carbon utilization, the glucose-to-lactate conversion rate increases, indicating a greater reliance by NPCs on glycolytic metabolism. Finally, we describe a metabolic exit event within the first 24 h of differentiation, characterized by a peak in mitochondrial and glycolytic activity, suggesting that modulation of both glycolytic and mitochondrial metabolism occurs during exit from pluripotency.

Cell culture and neural precursor cell differentiation

hPSCs, MEL1 and MEL2 (Australian Stem Cell Centre), were cultured in mTeSR1 medium (Stem Cell Technologies) on PSC-qualified Matrigel (BD Biosciences)-coated tissue culture plates (BD Biosciences). Cells were cultured in humidified CB 150 incubators (Binder) at 37°C with 5% CO2 in air (20% oxygen), or 5% CO2, 5% O2, and balanced with N2. To minimize exposure to 20% oxygen conditions during handling, medium was pre-equilibrated under each respective oxygen condition. All cell cultures were acclimated to 5% or 20% oxygen conditions for a minimum of two passages before use. hPSC lines are routinely tested for mycoplasma. To induce NPCs, hPSCs were treated with Y-27632 (ROCK inhibitor; 10 μM; AdipoGen) for 1 h, dissociated with TrypLE Select, and seeded as single cells at a density of 40,000 cells/cm2 in 12-well plates in mTeSR+Y-27632 (10 μM). Cells were cultured until >80% confluency was achieved; this was designated as ‘day 0’. From day 0 to day 4 medium was replaced daily with mTeSR+SB431542 (a TGFβ signaling inhibitor; 10 μM; Miltenyi Biotech) +dorsomorphin (a BMP signaling inhibitor; 4 μM; Sigma). From days 5 to 11, dorsomorphin treatment continued but SB431542 was withdrawn. From days 5 to 11, N2 medium (DMEM/F12 base with N2 supplement; Thermo Fisher Scientific) was added to mTeSR every 48 h in 25% increments (Fig. 1). Pluripotent hPSCs were harvested on day 0 and differentiating cells were harvested on days 1, 2, 3, 4, 5, 7, 9 and 11 for RNA, DNA, flow cytometry, cell counts and spent medium. On day 11, NPCs were passaged as single cells for expansion in N2/B27 medium (DMEM/F12 base with 1× N2 and 1× B27 supplement; Thermo Fisher Scientific), fixed for immunofluorescence or passaged for terminal differentiation. To terminally differentiate NPCs into neurons, NPCs were passaged as single cells in N2/B27 medium at a low density and cultured for 2-4 weeks. Neurospheres were formed from day 11 NPCs according to a published protocol (Denham and Dottori, 2011). Neurosphere medium consisted of 2% B27 supplement (Invitrogen), 1% ITS-A (Invitrogen), 1% N2 supplement (Invitrogen), L-glutamine (2 mM) and Penstrep (0.5%; Invitrogen) in Neurobasal medium (Invitrogen) with 20 ng/ml bFGF and EGF (Invitrogen). Small, individual pieces of NPC colonies were dissected and transferred to individual wells of an ultra-low attachment 96-well plate and cultured for 11 days, at which point neurospheres were pooled and harvested for RNA.

mtDNA measurement

mtDNA copy numbers were calculated by triplicate reactions using qPCR on 1 ng of total DNA. MEL1 and MEL2 cells were harvested using TrypLE Select (Invitrogen) for DNA isolation using the QIAamp DNA Mini Prep Kit (Qiagen) according to the manufacturer's instructions. mtDNA copy number was determined using the relative copy number method, whereby the abundance of transcripts for the mitochondrial gene ND1 was given relative to the nuclear gene GAPDH.

Flow cytometry (JC1, MitoTracker Green, MitoSox Red, TG30, GCTM2)

Cells cultured as described above were trypsinized using TrypLE Select and stained for 30 min at 37°C using the MitoProbe JC-1 Assay Kit for Flow Cytometry (Invitrogen, M34152; 2 μM final) with the addition of a non-stained control for fluorescence baseline identification, MitoSOX Red superoxide indicator (Invitrogen, M36008; 1 μM final) and MitoTracker Green FM (Invitrogen, M7514; 20 nM final). GCTM2 (mouse IgM) and TG30 (anti-CD9, mouse IgG2a) were stained using neat supernatant derived from hybridomas courtesy of Professor Martin Pera (Stem Cells Australia). Primary antibodies were detected using goat anti-mouse IgM (μ chain) AF647 (A21238) and goat anti-mouse IgG2a (γ2a) AF488 (A21131) at 1:1000, respectively (Invitrogen). Samples were split into three to avoid overlapping spectral profiles. JC1 was assayed with DAPI. MitoSOX Red, MitoTracker Green FM and DAPI were assessed simultaneously and pluripotency cell surface antigens GCTM2, TG30 and DAPI were assayed concurrently.

Spent medium collection for metabolite quantification

For the determination of metabolite use, three independent biological replicates of cells (MEL1 and MEL2) were cultured in Matrigel-coated 12-well plates in 5% or 20% oxygen as described in the ‘Cell culture and neural precursor cell differentiation’ section. Precisely 24 h after cell medium was replaced, spent medium was collected, centrifuged (1000 g for 2 min) to remove cellular debris and stored at −80°C until metabolite extraction. To determine cell numbers for each replicate well, cells were trypsinized using TrypLE Select and counted on a hemocytometer (Hausser Scientific). Glucose, lactate and pyruvate concentrations in spent medium were assayed using coupled enzyme reactions based on the formation or consumption of the fluorescent metabolite NADH/NADPH. Concentrations were determined from a standard curve and fluorescence was measured using a FLUOstar Omega plate reader (BMG Labtech). Glycolytic rate is defined here as glucose consumed from the medium relative to lactate produced into the medium assuming that 2 moles of glucose give rise to 1 mole of lactate. Therefore, glycolytic rate=(number of moles of lactate)/(2× number of moles of glucose) (Lane and Gardner, 1996; Lees et al., 2015; Harvey et al., 2016).

Statistical analyses

Data were analyzed using Prism software (GraphPad Prism 5) and SPSS (IBM). Gene expression data were analyzed using an unpaired, two-tailed Student's t-test comparing 5% and 20% oxygen treatments. Variances in carbohydrate utilization, percentage glycolysis and neural timeline data were analyzed using a two-factor ANOVA [factor 1: oxygen (5%; 20%); factor 2: cell line (MEL1; MEL2)]. Linear regressions were performed on timeline data to determine whether the coefficient of change was significantly non-zero. Where biologically applicable, AUC analysis was performed on timeline data and analyzed using a two-factor ANOVA as described. Significant (P<0.05) main effects and interactions were further analyzed using simple main effects calculated using the MS-residual from the initial ANOVA. Alternatively, a planned comparison between 5% and 20% oxygen treatments was performed. Results were considered statistically significant at P<0.05. All data are presented as mean±s.e.m.

Ethics approval

All methods used were approved by the University of Melbourne Human Ethics Committee (approval number 0722502.1).

Author contributions

Conceptualization: J.G.L., A.J.H., D.K.G.; Methodology: J.G.L., A.J.H.; Software: J.G.L.; Validation: J.G.L.; Formal analysis: J.G.L.; Investigation: J.G.L.; Resources: D.K.G.; Data curation: D.K.G.; Writing - original draft: J.G.L.; Writing - review & editing: J.G.L., A.J.H., D.K.G.; Visualization: A.J.H., D.K.G.; Supervision: A.J.H., D.K.G.; Project administration: A.J.H., D.K.G.; Funding acquisition: A.J.H., D.K.G.

Funding

This work was supported by the Australian Research Council Special Research Initiative Stem Cells Australia (SR110001002), and a Jasper Loftus-Hills Award (UTR7.116), an Alfred Nicholas Fellowship Award (UTR6.197), and an F. H. Drummond Travel Award (UTR6.184) from the University of Melbourne.

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Competing interests

The authors declare no competing or financial interests.

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