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The bioenergetics of somatic dedifferentiation into induced pluripotent stem cells remains largely unknown. Here, stemness factor mediated nuclear reprogramming reverted mitochondrial networks into cristae-poor structures. Metabolomic footprinting and fingerprinting distinguished derived pluripotent progeny from parental fibroblasts according to elevated glucose utilization and production of glycolytic end products. Temporal sampling demonstrated glycolytic gene potentiation prior to induction of pluripotent markers. Functional metamorphosis of somatic oxidative phosphorylation into acquired pluripotent glycolytic metabolism conformed to an embryonic-like archetype. Stimulation of glycolysis promoted, while blockade of glycolytic enzyme activity blunted, reprogramming efficiency. Metaboproteomics resolved upregulated glycolytic enzymes and downregulated electron transport chain complex I subunits underlying cell fate determination. Thus, the energetic infrastructure of somatic cells transitions into a required glycolytic metabotype to fuel induction of pluripotency.
Stemness transcription factors reprogram somatic cell fate to achieve embryonic-like pluripotency, the hallmark of induced pluripotent stem cells (iPSC) (Takahashi and Yamanaka, 2006; Meissner et al., 2007; Nelson et al., 2010). Developmental metabolic plasticity is associated with pluripotent stem cell fate specification (Dzeja et al., 2011), yet the metabolic dynamics that match the bioenergetic demand of nuclear reprogramming have not been established.
Somatic cells largely depend on mitochondrial oxidative phosphorylation as the primary source of energy production (DeBerardinis et al., 2008). In contrast, embryonic stem cells (ESC) rely on glycolytic ATP generation regardless of oxygen availability (Chung et al., 2007; Kondoh et al., 2007). The reliance on glycolysis reflects a low copy number of mitochondrial DNA (mtDNA) as well as low numbers of nascent mitochondria (St John et al., 2005; Cho et al., 2006). Differentiation increases mtDNA abundance and promotes mitochondrial biogenesis to form networks of elongated and cristae-rich mitochondria in support of competent oxidative metabolism (Facucho-Oliveira et al., 2007; Chung et al., 2008). Divergent energetic requirements of somatic versus pluripotent phenotypes implicate a distinguishing metabolic profile characterizing dedifferentiation.
We here demonstrate by high-resolution metabolomics that in nuclear reprogramming energy metabolism of iPSC progeny is transformed away from the parental source. Upregulated glycolytic enzymes and downregulated electron transport chain subunits enabled a metabolic switch converting somatic oxidative metabolism into a glycolytic flux-dependent, mitochondria-independent state underlying pluripotent induction.
Reprogramming by four stemness transcription factors (4F) restructured mouse embryonic fibroblasts (MEF), characterized by organized mitochondrial networks, to a primordial cytotype featuring an increased nuclear-to-cytosol ratio with few perinuclear mitochondria (Figures 1A-D). Mature tubular and cristae-rich somatic mitochondria transitioned into immature spherical and cristae-poor structures in 4F iPSC, indicative of bioenergetic remodeling (Figures 1B and D). High-resolution 1H NMR quantification of 18 extracellular metabolites (Figures (Figures1E1E and S1) decoded the metabolic consequences of dedifferentiation by separating the 4F iPSC metabolomic footprint from the parental MEF landscape (Figure 1F). Accordingly, glucose and lactate were identified as distinguishing metabolites (Figure 1G). Rates of glucose utilization (2.3 ± 0.1 and 2.2 ± 0.1 nmol/μg protein/h) and lactate production (4.4 ± 0.1 and 4.8 ± 0.1 nmol/μg protein/h) were significantly elevated in two 4F iPSC lines (4F iPS1 and 4F iPS2) compared to MEF (1.8 ± 0.1 and 3.1 ± 0.1 nmol/μg protein/h, respectively, n=6, p<0.05; Figure 2H and 2I). Thus, nuclear reprogramming induces mitochondrial regression and extracellular metabolome resetting.
Intracellular metabolite fingerprinting validated the glycolytic capacity of 4F iPSC, segregating the acquired metabolomic pattern away from parental MEF and towards the pluripotent ESC standard (Figures (Figures2A2A and S1B). The resolved 18 intracellular metabolite panel distinguished iPSC based upon acetate, lactate, fumarate and taurine concentrations (Figure (Figure2B2B and S1C). 4F iPSC accumulation of acetate was similar to ESC (31.6 ± 0.9, 32.5 ± 1.0 and 29.2 ± 0.7 pmol/μg protein, n=3), and distinct from MEF (21.3 ± 0.5 pmol/μg protein, n=3 populations, p<0.05, Figure 2C). Comparably, lactate was equivalent in 4F iPSC and ESC (127 ± 3, 152 ± 5 and 140 ± 10 pmol/μg protein, n=3), yet significantly different from MEF (95 ± 5 pmol/μg protein, n=3, p<0.05; Figure 2D). Lactate efflux (5.3 ± 0.3 and 5.8 ± 0.1 nmol/μg protein/h) at a rate double that of MEF (2.8 ± 0.2 nmol/μg protein/h, n=6, p<0.05; Figure 2E) indicated functional glycolysis in 4F iPSC. Consistent with the lower efficiency of glycolytic ATP production compared to oxidative phosphorylation, the ADP/ATP ratio, an index of cellular energy turnover, was reduced in 4F iPSC and ESC compared to MEF (0.046 ± 0.002, 0.052 ± 0.001 and 0.053 ± 0.001 versus 0.070 ± 0.001, respectively, n=3, p<0.05, Figure 2F). Overall, oxygen consumption was low in 4F iPSC and ESC, compared to MEF both at baseline (0.51 ± 0.04, 0.42 ± 0.07 and 0.36 ± 0.02 versus 1.8 ± 0.13 nmol/106 cells/min, n=3, p<0.05, Figure 2G) and under electron transport chain uncoupling (0.98 ± 0.10, 1.22 ± 0.09 and 0.95 ± 0.13 versus 6.51 ± 1.05 nmol/106 cells/min, n=3, p<0.05, Figure 2H). 4F iPSC preserved the ability to generate mitochondrial membrane potential and demonstrated mitochondrial hyperpolarization (Figure S3A-C), indicative of reduced ATP utilization.
Treatment of MEF undergoing reprogramming with 2-deoxyglucose (2DG), a general inhibitor of glycolysis, blunted induction of the pluripotent marker alkaline phosphatase (Figure 2I), implicating a glycolytic requirement for iPSC generation. Beyond 2DG, the hexokinase 2 inhibitor 3-bromopyruvic acid (BrPA) (Ko et al., 2001) and pyruvate dehydrogenase kinase inhibitor dichloroacetate (DCA) (Stacpoole, 1989) reduced reprogramming efficiency (0.9 ± 0.3 in 2DG, 19.8 ± 1.0 in BrPA, 17.3 ± 1.5 in DCA versus 27.8 ± 3.1% SSEA1 positive cells in vehicle, n=3, p<0.05, Figure 2J) without impairing cell growth (Figure 2K). Extracellular metabolite profiles of MEF treated with 2DG or BrPA were consistent with inhibition of glycolysis and stimulation of oxidative metabolism, while DCA treatment stimulated mitochondrial function and accelerated consumption of oxidative substrates (Figure S2A-C). Glucose supplementation stimulated glycolytic flux, reduced mitochondrial function (Figure S2 D-F) and increased SSEA1 positive cell density (0.5 ± 0.3, 16.6 ± 5.3, 27.8 ± 1.1 and 17.1 ± 4.7% SSEA1 positive cells in 0, 5, 12.5 and 25 mM glucose, n=3, p<0.05). Conversely, few MEF underwent nuclear reprogramming in the absence of glucose (Figure 2L-M). Transition from somatic oxidative metabolism to pluripotent glycolysis thus contributes to nuclear reprogramming efficiency.
Mitochondrial membrane potential served as a surrogate of metabolic transition during nuclear reprogramming. Prospective live cell imaging distinguished ESC-like compact clusters with high tetramethylrhodamine methyl ester (TMRM) fluorescence from transfected non-reprogrammed MEF with low mitochondrial membrane fluorescence (Figure 3A). FACS sampling of high TMRM cells at day 7 and 14 selected an emergent subpopulation within small cell clusters, consistent with mitochondrial membrane potential polarization during dedifferentiation (Figure S3A-D). At day 7, high TMRM cells had significantly elevated glycolytic gene expression (Glut1, Hxk2, Pfkm and Ldha), which met that of ESC by 2-weeks of reprogramming (Figure 3B). In contrast, at 1-week of reprogramming pluripotent gene expression (Fgf4, Nanog, Oct4 and Sox2) remained low in high TMRM cells, similar to starting MEF, with pluripotent gene induction apparent only after 2-weeks (Figure 3C). Metabolic reprogramming thus precedes pluripotent gene expression during somatic cell dedifferentiation from an oxidative to a glycolytic iPSC phenotype (Figure 3D).
As c-Myc gene targets control rates of glycolysis and mitochondrial biogenesis (Dang, 2007), a cell line was derived without c-Myc (3F iPSC). Similar to 4F iPSC, NMR metabolomic footprinting and fingerprinting segregated 3F iPSC away from parental MEF (Figures 4A-B), based upon greater intracellular and extracellular accumulation of glycolytic end products, acetate (intracellular: 30.4 ± 0.8 versus 21.3 ± 0.5 pmol/μg protein; extracellular: 0.41 ± 0.03 versus 0.035 ± 0.001 nmol/μg protein/h, n=3, p<0.05, Figure 4C) and lactate (intracellular: 188 ± 11 versus 95 ± 5 pmol/μg protein; extracellular: 4.1 ± 0.1 versus 3.0 ± 0.1 nmol/μg protein/h n=3, p<0.05, Figure 4D). Consistent with accelerated glycolysis, 3F iPSC displayed reduced energy turnover compared to MEF (0.039 ± 0.005 versus 0.068 ± 0.006, n=3, p<0.05, Figure 4E). Both 3F and 4F iPSC demonstrated limited oxidative capacity and elevated lactate production (Figure 4F). Thus, nuclear reprogramming-induced metabolic remodeling is characteristic of the iPS phenotype, independent of c-Myc transduction.
Metaboproteome dissection delineated a transformed molecular signature in iPSC, distinct from parental MEF, yet synonymous to the metabolic protein profile of ESC (Figures 4G-J and S4, Table S1 and S2). A label-free, quantitative proteomics approach revealed the identities of glycolytic enzymes consistently upregulated in pluripotent cytotypes relative to MEF (Figures (Figures4G4G and S4A), confirmed independently by 2-D gel based analysis (Figure 4I). Concomitantly, 65% of detected complex 1 subunits were downregulated in iPSC extracts (Figures (Figures4H4H and S4B, Table S1) as validated by 2-D gel based analysis (Figure 4J), underscoring departure from MEF patterns and acquisition of a glycolytic-dependent profile (Figure 4K). In addition, reprogramming was associated with selective downregulation of the reducing equivalent entry points (complex I and II) (Figure S4C, Table S1). Thus, the resolved iPSC metaboproteome unmasks a bioenergetic switch in energy utilization infrastructure underlying metabolic remodeling in nuclear reprogramming.
Nuclear reprogramming sets in motion dedifferentiation processes leading to acquisition of pluripotency. This study resolved the metabolic attributes underpinning iPSC generation. Metaboproteome restructuring was demonstrated to underlie conversion from somatic oxidative bioenergetics to glycolysis of pluripotent progeny. Promotion of a glycolytic flux-dependent, mitochondria-independent metabotype fostered nuclear reprogramming.
Regression of the parental somatic network of abundant tubular and cristae-rich mitochondria into sparse spherical and cristae-poor structures characterized iPSC progeny. A mitochondria-poor capacity would enable iPSC to revert reactive oxygen species-dependent telomere shortening associated with cellular aging (Passos et al., 2007; Marion et al., 2009). In fact, mitochondrial regression and reduction of mtDNA as a result of nuclear reprogramming is consistent with the undeveloped mitochondrial morphology of ESC (Kondoh et al., 2007; Armstrong et al., 2010; Prigione et al., 2010; Zeuschner et al., 2010). Induced reduction in mitochondria number and maturity suggests metamorphosis of the bioenergetic infrastructure underlying dedifferentiation.
Metabolomic footprinting and fingerprinting indeed demonstrated distinct features of iPSC metabolism. The key metabolic rates contributing to the iPSC phenotype were consistent among multiple clones, and included elevated utilization of glucose and accumulation of intracellular and extracellular lactate (via lactate dehydrogenase) and acetate (via acetyl-CoA synthase or ATP-citrate lyase), suggesting that accelerated glycolysis results in accumulation of metabolic byproducts within the cell, followed by metabolite exportation. iPSC demonstrated diminished basal oxygen consumption and uncoupled oxidative capacity, indicating departure from oxidative metabolism. ESC also rely on glycolytic ATP generation (Cho et al., 2006; Facucho-Oliveira et al., 2007), with pluripotency maintenance sustained under hypoxic conditions (Ezashi et al., 2005). Acquisition of an embryonic-like metabolic state is concomitant with global transcriptional and epigenetic reorganization that includes demethylation and activation of the promoter region of pluripotency inductors (Bock et al., 2011; Plath and Lowry, 2011). A glycolytic burst could promote linkage of energy metabolism with nuclear reprogramming-induced epigenetic resetting (Huangfu et al., 2008; Friis et al., 2009; Wellen et al., 2009; Teperino et al., 2010), priming the cell for pluripotent induction. Indeed, expression of glycolytic genes preceded pluripotent gene expression, implicating metabolic remodeling as a facilitator of nuclear reprogramming.
Complete oxidation of glucose limits generation of substrates for biosynthetic pathways. Transition to glycolytic metabolism would fuel specific cellular processes, such as ATP consumption in the cell nucleus and provide the necessary balance between ATP generation and production of metabolic precursors to meet anabolic requirements of dedifferentiation and associated proliferation (DeBerardinis et al., 2008; Chung et al., 2010). Strategies that in part stimulate glycolysis, including induction of hypoxia or inhibition of the p53 pathway, increase nuclear reprogramming efficiency (Kruse and Gu, 2006; Krizhanovsky and Lowe, 2009; Yoshida et al., 2009). Consistent with the ability of small molecules with known metabolic targets to modulate the appearance of pluripotent colonies (Zhu et al., 2010), here stimulation of glycolytic flux potentiated reprogramming whereas oxidative metabolism promotion preserved the somatic cytotype.
c-Myc, through its regulation of both glycolysis and mitochondrial metabolism, is a key contributor to the metabolic Warburg effect, reflecting the tendency of highly proliferative cell types to convert glucose to lactate despite sufficient oxygen for mitochondrial oxidative metabolism (Dang, 2007). Here 3F iPSC displayed equivalent metabolic profiles to 4F iPSC counterparts, verifying that nuclear reprogramming-induced metabolic remodeling did not require c-Myc transduction. Proteomic dissection resolved a metabolic infrastructure that distinguished 3F and 4F iPSC from fibroblasts. Specifically, reprogramming induced reciprocal changes in glycolytic enzymes and electron transport chain components, indicating a molecular substrate contributing to the switch from oxidative to glycolytic metabolism. The observed decreased expression of complex I and II subunits would uncouple oxidative metabolism and ATP generation, favoring glycolytic dependent ATP production.
In summary, dedifferentiation of parental somatic cells induced mitochondrial regression, electron transport chain downregulation, and glycolysis enzyme upregulation, revealing a pluripotent glycolytic signature with limited dependence on mitochondrial metabolism. Induction of the glycolytic metabotype support anabolic and catabolic requirements for bona fide pluripotency to regulate cell fate.
Mouse embryonic fibroblasts were transduced with in house HIV-based viral vectors encoding OCT3/4, SOX2, and KLF4 in the presence or absence of c-MYC to produce iPSC clones that met pluripotent criteria including expression of stem cell markers, embryoid body differentiation, teratoma formation, diploid aggregation and contribution to organogenesis (Martinez-Fernandez et al., 2009; Nelson et al., 2009a; Nelson et al., 2009b). Alternatively iPSC were derived with a viPS (Open Biosystems) kit. iPSC and mouse ESC (R1) were induced and maintained in DMEM supplemented with 15% FBS, 25 mM glucose, 2 mM Glutamax (Invitrogen) and 1 mM sodium pyruvate. Reprogramming efficiency was quantified using an alkaline phosphatase staining kit (Stemgent) or FACS analysis of SSEA1 expression (Millipore) on a LSR II flow cytometer.
Mitochondrial density and morphology was evaluated in 1% glutaraldehyde and 4% formaldehyde fixed cells, and examined as ultramicrotome sections on a JEOL 1200 EXII electron microscope (Perez-Terzic et al., 2007).
For footprinting of extracellular metabolites (see Supplemental Experimental Procedures), 540 μL of media collected following 24 h of culture was added to 60 μL of D2O (Sigma) containing 5 mM sodium 3-(trimethylsilyl)propionate-2,2,3,3-d4 (TSP) (Sigma) for chemical shift reference and 81.84 mM formate (Sigma) for peak quantification reference (Turner et al., 2008). For intracellular metabolite fingerprinting, neutralized perchloric acid extracts were concentrated with a SpeedVac and suspended in 600 μl of 100 mM phosphate buffer (pH 7.0) in D2O (Sigma) containing 0.5 mM TSP (Beckonert et al., 2007). Identities of 1H NMR spectra peaks obtained on a Bruker Ultrashield 700 MHz spectrometer with a zgpr water suppression pre-saturation pulse were assigned by comparison to reference values for chemical shift and multiplicity, and confirmed by comparison to spectra of pure compounds in the Human Metabolome database (Govindaraju et al., 2000; Wishart et al., 2009).
Lactate efflux rate was assessed in extracellular media using a lactate assay (SUNY). Nucleotide concentrations were determined in neutralized perchloric acid extracts by high performance liquid chromatography, using a 0.1 M phosphate (pH 6.5), 0.01 M tetrabutylammoniumhydrogensulfate, and 40% methanol elution buffer (Chung et al., 2007). Oxygen consumption was assessed using an electrode system (Hansatech) on 5 million trypsinized cells suspended in DMEM. Maximal rate of uncoupled oxygen consumption was assessed by serial additions of 2,4-dinitrophenol (Sigma).
Mitochondrial membrane potential was assessed at day 4-14 of reprogramming by incubating with 20 nM TMRM (Anaspec) for 30 min at 37°C, and imaged with a LSM 510 Axiovert laser confocal microscope. By 1 and 2-weeks of reprogramming, single cell suspensions were incubated in TMRM and separated by a FACS Aria Cell Sorter. Gene expression was examined on an Eco RT-PCR system (Illumina).
Protein extracts were resolved by 2-D gel electrophoresis and 4-15% SDS-PAGE (100 and 30 μg, respectively) and silver stained (Zlatkovic-Lindor et al., 2010). For comparative analysis, SDS-PAGE lanes were excised, destained, and prepared for LC-MS/MS, as were protein species from 2-D gels identified as significantly altered by PDQuest analysis (Zlatkovic-Lindor et al., 2010). Isolated tryptic peptides were analyzed and identified by LTQ-Orbitrap mass spectrometry. Label-free quantitative comparison of SDS-PAGE protein and peptide abundance was carried out on acquired MS spectra using Rosetta Elucidator’s differential workflow, with annotation performed using PeptideTeller and ProteinTeller (Neubert et al., 2008; Lomenick et al., 2009). See Supplemental Experimental Procedures.
Data are presented as mean ± SEM. Metabolic footprinting and fingerprinting were analyzed using principle component analysis and JMP. Student t-test was used to evaluate two group comparisons, and ANOVA with a Bonferroni post-hoc correction for three group comparisons. A value of P<0.05 was considered significant.
Authors thank Mayo Clinic Nuclear Magnetic Resonance Core and Flow Cytometry/Optical Morphology Shared Resource for assistance. This work was supported by National Institutes of Health (R01HL083439, T32HL007111, R01HL085208, R56AI074363), Canadian Institutes of Health Research, Marriott Program, and Mayo Clinic.
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