Review Article | Published:

Metabolic signatures of cancer cells and stem cells

Abstract

In contrast to terminally differentiated cells, cancer cells and stem cells retain the ability to re-enter the cell cycle and proliferate. To proliferate, cells must increase their uptake and catabolism of nutrients to support anabolic cell growth. Intermediates of central metabolic pathways have emerged as key players that influence cell-differentiation ‘decisions’, processes relevant to both oncogenesis and normal development. Consequently, how cells rewire metabolic pathways to support proliferation can have profound consequences for cellular identity. Here, we discuss the metabolic programs that support proliferation, and we explore how metabolic states are intimately entwined with the cell-fate decisions that characterize stem cells and cancer cells. By comparing the metabolism of pluripotent stem cells and cancer cells, we hope to illuminate common metabolic strategies as well as distinct metabolic features that may represent specialized adaptations to unique cellular demands.

Main

At their core, cell survival and growth are metabolic problems. Cells catabolize nutrients to generate the energy and reducing equivalents required to maintain basic cellular processes. Likewise, anabolic metabolic pathways convert nutrients into the macromolecules necessary for cell growth and proliferation. This intimate relationship between metabolism and cellular fitness is best exemplified by the observation that the growth of most unicellular organisms is directly linked to nutrient availability1. In contrast, the cells of multicellular organisms must cooperate to share relatively constant nutrient supplies; consequently, metazoan cell proliferation is regulated by growth factors that license the acquisition of extracellular nutrients and the activation of anabolic growth programs. During development, growth-factor signalling pathways direct the proliferation, migration and death of selected populations, thus ensuring proper organ size and function. These same pathways frequently become subverted in cancer: oncogenic activation of growth-factor signalling or inhibition of cell death enables the pathological proliferation that drives tumour growth. Therefore, that cancer cells share many metabolic features with normal developmental programs is perhaps unsurprising. For example, just as folate deficiency is a major cause of early-embryonic growth defects2, therapies interfering with folate metabolism are key components of many successful chemotherapeutic regimens3.

Metabolites play many roles beyond serving as substrates for energy generation and anabolic growth. Metabolites contribute to the regulation of intracellular redox balance4, directly alter the activity of intracellular signalling cascades5 and serve as co-substrates for enzymes that modify macromolecules such as DNA and proteins6. As a result, intracellular metabolic pathways may influence many cellular programs beyond proliferation. The dual role of metabolites as substrates for both anabolic and regulatory processes raises the possibility that the utilization of nutrients for cell proliferation might inherently affect the availability of metabolites for other non-anabolic roles. This metabolic convergence between proliferation and cell-fate regulation may be particularly relevant in stem cells, which accomplish the dual feat of retaining the abilities to proliferate rapidly and to differentiate into specialized cell types. As a result, there is great interest in elucidating the metabolic networks that sustain stem-cell self-renewal and identifying metabolic nodes that influence lineage-specific differentiation.

Pluripotent stem cells (PSCs) provide an ideal model system to study the intersection of proliferation, metabolism and differentiation. Although pluripotency—the ability to give rise to cells of all three embryonic germ layers—exists only transiently during early mammalian development, the pluripotent state can be captured indefinitely in vitro, thus providing an invaluable resource to study principles of dynamic cell-fate transitions, elucidate pathways critical for lineage-specific differentiation and explore possibilities for regenerative medicine7. PSCs can be isolated from the inner cell mass (ICM) of preimplantation epiblast (embryonic stem cells, ESCs) or the early post-implantation epiblast (epiblast stem cells); alternatively, somatic cells can be reprogrammed to pluripotency through the expression of key pluripotency-associated transcription factors (induced PSCs). Intriguingly, the functional identity of PSCs is largely determined not by the cell of origin but instead by the culture conditions under which the cells are generated and propagated7. Naïve PSCs share molecular characteristics with cells of the ICM and can readily give rise to all germ layers and contribute to chimaeras8. In contrast, primed PSCs exhibit similarities to post-implantation epiblast cells and, while retaining pluripotency, represent a more committed developmental state8. By and large, PSCs can be coaxed into either the naïve or primed state depending on the exogenous cues present in the culture medium7. Therefore, understanding how specific nutrients and growth factors in PSC culture regulate cellular phenotypes such as growth and gene expression programs is likely to yield critical insights into the regulation of cell identity.

A key question when comparing the metabolism of specialized cell types is how to distinguish the metabolic alterations accompanying differences in growth and proliferation from the metabolic alterations that support cell-type-specific fate decisions. Therefore, comparing the emerging literature describing the metabolism of stem cells with the vast literature on the metabolism of cancer cells is useful. Like stem cells, cancer cells are capable of rapid proliferation; unlike stem cells, cancer cells are relatively locked into a malignant identity. In this Review, we will discuss emerging evidence that cancer cells and stem cells engage similar metabolic pathways to support anabolic proliferation. We will further examine how specific metabolic alterations influence the balance between self-renewal and differentiation in cancer cells and stem cells. Because both stem cells and cancer cells face many similar metabolic challenges, lessons from one system may provide generalizable insight into the potential metabolic avenues that support growth, survival and differentiation across multiple cell types.

Metabolic requirements of proliferation

Glucose and glutamine are fundamental substrates for mammalian cells growing in culture

Regardless of the identity of the proliferating cell, proliferation requires a net increase in biomass. To generate the nucleic acids, lipids and proteins required for proliferation, cells increase the uptake and catabolism of nutrients that provide the raw materials for macromolecular synthesis. Glucose and glutamine, two of the most abundant metabolites in the serum and in common tissue culture media, are the major sources of energy and reducing equivalents required to assemble metabolic building blocks into macromolecules9 (Fig. 1). Beyond generating ATP and NAD(P)H, the catabolism of glucose and glutamine generates precursors for the biogenesis of non-essential amino acids required to sustain the synthesis of proteins and nucleic acids. Additionally, glucose provides the ribose, and glutamine provides the obligate source of reduced nitrogen for nucleic acid biosynthesis. In cultured cells, glucose-derived acetyl-CoA is the primary source of carbon for de novo lipid biosynthesis, whereas protein biosynthesis, which makes up most of the cellular biomass, is sustained by the combination of direct amino acid uptake and de novo synthesis of non-essential amino acids from glucose and glutamine10. Therefore, although the relative dependency on each of these metabolic precursors can vary depending on the cell line, culture conditions and nutrient availability9,10, a fundamental feature of mammalian cells proliferating in culture is the central role of glucose and glutamine in supporting anabolic growth.

Fig. 1: Glucose and glutamine are critical inputs in major anabolic pathways.
figure1

In proliferating cells, glucose and glutamine (blue) are taken up from the extracellular environment and catabolized through major metabolic pathways, including glycolysis, the pentose-phosphate pathway (PPP) and the TCA cycle, and they provide the reducing equivalents (purple) and high-energy carriers (ATP, red) required to synthesize major macromolecules (green). A subset of the non-essential amino acids synthesized from glucose and glutamine are shown. Reducing equivalents (NADH and FADH2) in the mitochondria fuel the electron-transport chain and enable synthesis of ATP through oxidative phosphorylation (oxphos). TCA cycle intermediates such as citrate and oxaloacetate (OAA, converted to aspartate) likewise contribute to lipid and nucleotide biosynthesis, respectively.

The critical role of glucose and glutamine in cancer-cell proliferation is well established and has been extensively reviewed elsewhere11,12,13. Like cancer cells, PSCs can proliferate indefinitely in culture and are also heavily reliant on exogenous glucose and glutamine14,15. Although the inherent flexibility of metabolic networks ensures that cells have multiple mechanisms to cope with diminished abundance of either nutrient16, proliferation is maximized when both glucose and glutamine are abundant. Therefore, a major paradox of rapidly proliferating cells is their tendency to discard most glucose carbons as lactate. The conversion of glucose to lactate despite the presence of sufficient oxygen to sustain complete oxidation of glucose-derived carbons, commonly referred to as the ‘Warburg effect’ or aerobic glycolysis, is a hallmark of all rapidly proliferating mammalian cells. Indeed, compared with their slower-growing differentiated counterparts, PSCs—regardless of their cell of origin or stage of pluripotency—engage in aerobic glycolysis, consuming high levels of glucose and secreting large quantities of lactate17,18,19,20. This glycolytic flux is contingent on cells maintaining their pluripotent identity and is rapidly reversible: glycolytic lactate production is decreased early during PSC differentiation19,20,21,22, and reprograming differentiated cells to the pluripotent state results in the re-acquisition of glycolytic phenotypes23,24,25. Accordingly, interfering with glycolytic metabolism impairs reprogramming and maintenance of the pluripotent state20,21,22,23,24,25. Mechanistically, transcription factors establishing the pluripotent state may directly regulate metabolic pathways: for example, the core pluripotency transcription factor Oct4 binds loci encoding glycolytic genes and consequently promotes glycolysis26. Not all differentiation results in downregulation of aerobic glycolysis: whereas human PSC differentiation to mesoderm and endoderm decreases glycolytic flux, differentiation to early neuroectoderm does not22. Therefore, a key open question is whether the links between aerobic glycolysis and pluripotency are a result of the bioenergetic constraints of particular cell types or of altered accumulation of metabolites with the ability to regulate cell-fate programs.

Despite the ubiquity of the Warburg effect in proliferating cells, the potential benefits of this metabolic hallmark remain controversial27. Warburg’s initial hypothesis that the aerobic conversion of pyruvate to lactate results from defective mitochondrial respiration has since been disproven in both cancer cells and normal cells12. Mitochondria are notably distinct in PSCs compared with their differentiated counterparts: PSCs tend to have fewer mitochondria per cell, and these mitochondria exhibit immature morphology marked by sparse cristae18,19,28,29. Nevertheless, PSCs are not defective in mitochondrial-substrate oxidation. Respiratory complexes are fully functional in PSCs19, and PSCs consequently have the flexibility to engage in oxidative or glycolytic programs depending on culture conditions18,30. Indeed, even under basal culture conditions, PSCs engage in substantial glutamine oxidation, thereby sustaining bioenergetics14,15,30,31, and enhanced respiration may be a common feature of PSCs in the naïve state18,32,33. Therefore, the aerobic glycolysis that characterizes PSCs, like that in other proliferating cell types, does not appear to result from a defect in mitochondrial function.

In the absence of understanding the origin or consequences of the Warburg effect in PSCs, it is difficult to ascribe any particular benefit or function to aerobic glycolysis in the regulation or maintenance of pluripotent-cell growth. Intriguingly, aerobic glycolysis has recently been suggested to arise in part as a secondary consequence of the metabolic requirements of proliferating cells rather than as a primary driver of proliferation. For example, the conversion of pyruvate to lactate becomes favoured when the cytosolic NADH/NAD+ ratio rises, as may occur in proliferating cells as a result of the rapid consumption of NAD+ used to drive the oxidative reactions required for nucleotide production13. PSCs are notable for their rapid proliferation (doubling time of 12–24 h) and high nuclear-to-cytoplasmic ratio; therefore, they may be disproportionally reliant on nucleotide biosynthesis to sustain proliferation. Alternatively, because high mitochondrial-substrate oxidation produces reactive oxygen species that promote differentiation and exert deleterious effects on cellular fitness, discarding reduced electrons in the form of lactate rather than transferring these reducing equivalents to the mitochondria for oxidation may be particularly beneficial to PSCs that must resist spontaneous differentiation and are primed for apoptosis34,35. In support of this hypothesis, the ESC metabolome becomes more oxidized after differentiation36.

The emerging literature on PSC metabolism has collectively demonstrated that cultured PSCs meet the anabolic demands of proliferation in a manner largely similar to that in cancer cells and other proliferating mammalian cells. This correspondence between the metabolic profiles of PSCs and cancer cells may arise simply as a consequence of the conserved metabolic requirements of proliferation. Nevertheless, emerging evidence suggests subtle differences in how different PSC types use glucose and glutamine. For example, both mouse and human PSCs in the naïve state of pluripotency exhibit higher basal respiration than their primed counterparts18,32,33. Moreover, whereas more glycolysis occurs in human naïve PSCs than primed PSCs, the opposite trend is true in mouse PSCs18,20. The extent to which these differences arise from variations in cell size, proliferation rate and culture conditions (including medium nutrient composition, growth-factor availability and choice of extracellular matrix) rather than functional variation among cell types remains an important open question. Certainly, simply culturing the same cell type in different medium formulations can substantially alter how cells engage metabolic pathways to fulfil anabolic demands30. However, because PSC identity is intimately linked to culture conditions7, these metabolic differences could also result from—or contribute to—the establishment of different functional states. Therefore, a critical avenue of future work will be to unravel metabolic phenotypes that are secondary to altered growth conditions from metabolic phenotypes specifically resulting from variations in PSC identity.

Beyond glucose and glutamine: alternative metabolic strategies

Although aerobic glycolysis and glutamine oxidation are nearly ubiquitous features of cancer cells proliferating in culture, recent studies examining tumour metabolism in vivo have illuminated the diverse metabolic strategies and substrates that support cancer growth12,37 (Fig. 2a). For example, several types of brain and lung tumours tend to increase oxidation of glucose-derived carbons and to rely on glucose for replenishment of tricarboxylic acid (TCA) cycle intermediates38,39,40, whereas clear cell renal carcinomas retain profiles consistent with aerobic glycolysis41. Likewise, several tumour types have been reported to have minimal reliance on glutamine oxidation to sustain TCA cycle metabolite pools and may even synthesize glutamine de novo from glucose-derived carbons38,39,42,43. Increasingly, substrates beyond glucose and glutamine are being recognized to play critical roles as bioenergetic substrates in vivo. Lactate, produced in high quantities in the tumour microenvironment, may function as a major source of carbon for growing tumours40,44,45. Similarly, acetate has been reported to serve as a fuel for brain and liver tumours46,47, and circulating branched-chain amino acids can contribute to biomass accumulation in non-small cell lung carcinoma48. Clearly, the myriad metabolic strategies used by tumours to sustain proliferation are only just beginning to be understood. Diverse factors such as local nutrient availability43, transforming oncogenes49 and tissue of origin48 will probably converge in determining the unique metabolic preferences and liabilities of a given tumour.

Fig. 2: Metabolic strategies used by cancer cells and pluripotent stem cells.
figure2

a, Left, cancer cells in vitro take up high levels of glucose and glutamine. Much of the glucose is converted to lactate via aerobic glycolysis, and glutamine-derived carbons provide TCA cycle anaplerosis, thereby fuelling oxidative phosphorylation. Right, a sampling of the diverse metabolic strategies used by cancer cells in vivo. (i) Many tumours, including lung and brain tumours, exhibit high glucose uptake and catabolism in the TCA cycle, often in addition to aerobic glycolysis. Glucose-derived carbons can fuel the TCA cycle through forward pyruvate dehydrogenase flux or via pyruvate carboxylase–mediated anaplerotic entry as oxaloacetate. In this scenario, glutamine is a relatively minor contributor to TCA cycle metabolites, and often tumours can net produce glutamine de novo from glucose-derived carbons. (ii) Increasingly, substrates beyond glucose and glutamine are recognized as major substrates for anabolic pathways and the TCA cycle; these substrates include acetate, lactate and branched-chain amino acids (BCAAs) in lung, brain and liver tumours. (iii) In human clear cell renal carcinoma, tumours exhibit high glycolysis and lactate production consistent with the canonical Warburg effect. Glutamine metabolism has not been assessed and is therefore shaded in grey. (iv) Studies in mouse pancreatic cancer have revealed that glucose, lactate and glutamine all contribute to TCA cycle intermediates. b, Top, PSCs exhibit a variety of metabolic strategies depending on culture condition or stage of differentiation. Naïve mouse PSCs in the ground state of pluripotency can synthesize glutamine de novo from glucose-derived carbons; naïve mouse PSCs also can catabolize threonine (Thr). Both naïve and primed PSCs in culture exhibit high consumption of glucose and glutamine as well as extensive production of lactate. After differentiation, the proliferative hallmark of aerobic glycolysis decreases along with the proliferation rate. Bottom, although little is known about pluripotent-cell metabolism in vivo, studies of embryos developing ex vivo have identified two major metabolic stages: from zygote to morula, embryos are dependent on the monocarboxylates pyruvate and lactate, and engage in oxidative metabolism; around the morula stage, embryos begin to use glycolysis to fuel metabolic pathways, and the trophectoderm exhibits features consistent with oxidative phosphorylation, whereas the ICM may rely more on aerobic glycolysis. Green, blastomeres; light blue, trophectoderm; purple, ICM; orange, primitive endoderm; dark blue, epiblast.

Early evidence suggests that PSCs may exhibit substantial metabolic diversity, even under routine culture conditions (Fig. 2b). Indeed, many of the unique metabolic phenotypes of cancer cells that become revealed only under tumour growth in vivo are readily apparent in PSCs in vitro. For example, driving the normally metastable and heterogeneous population of naïve mouse ESCs into a homogenous ground state of pluripotency substantially increases the contribution of glucose-derived carbons to TCA cycle metabolites14,50. As a result, naïve, ground-state ESCs can generate glutamine de novo from glucose-derived carbons and therefore have the unique ability to proliferate in the absence of exogenous glutamine14. Analogously to the distinct utilization of branched-chain amino acids by certain tumours in vivo48, ESCs may also be able to catabolize amino acids in unusual ways. Mouse ESCs express an active version of the enzyme threonine dehydrogenase, which enables robust catabolism of threonine to glycine and acetyl-CoA; consequently, mouse ESCs are exquisitely reliant on exogenous threonine to sustain viability and cell identity51,52. Moreover, just as tumour cells may efficiently scavenge macromolecular nutrients from the microenvironment53, PSCs appear to preferentially utilize exogenously supplied lipids over those synthesized de novo30. Furthermore, acetate is one of the most changed metabolites during early differentiation, thus suggesting that alterations in acetate production or consumption may follow changes in pluripotent identity21.

Because the state of pluripotent self-renewal is inherently intertwined with culture conditions, whether these metabolic phenotypes of PSCs translate to the transient pluripotent state in vivo remains an open question. Relative to their ICM counterparts, PSCs have undergone substantial adaptation to cell culture and have acquired the ability to sustain continuous self-renewal; consequently, the extent to which the metabolism of cultured PSCs reflects the growth of the ICM or epiblast cells in vivo remains unclear54,55. Very little is known about metabolism during early mammalian development. As embryos progress from the zygote to the blastocyst stage, there is no substantial increase in cell mass: instead, these early cell divisions are ‘reductive’ and are marked by an increase in nuclei at the expense of cytoplasm56. Therefore, the metabolic demands of proliferation during early development are likely to be quite different from those of most proliferating cells, which must generate net biomass. Although assessing how embryos developing in situ meet their metabolic needs is difficult, studies on embryos developing in culture have revealed that pyruvate and lactate are the primary sources of energy in early-cleavage embryos57 (Fig. 2b). At the morula stage, cells become competent to utilize glucose as an energy source, and studies have suggested that the cells of the ICM may be more glycolytic relative to the oxidative cells of the trophectoderm58. Intriguingly, studies examining amino acid turnover in mouse blastocysts and dissociated ICM cells have not identified glutamine as a highly consumed amino acid58, thus suggesting that early blastocyst development may impose very different requirements for amino acids compared with those in cultured cells. Indeed, perhaps because of a lower requirement for net protein synthesis, mouse blastocysts can develop in vitro with just the presence of certain single amino acids or even in the absence of any free amino acids59,60. Future studies, perhaps using tools such as single-cell metabolite sensors, heterologous enzymes and genetic modification of metabolic pathways combined with lineage tracing, may begin to unravel the metabolic pathways required for early mammalian development.

Regulation of anabolic metabolic pathways

Together, these studies suggest that cultured cancer cells and PSCs engage a core set of common metabolic pathways to support rapid proliferation. In normal cells, growth-factor-mediated activation of receptor tyrosine kinases engage signalling pathways, such as those involving Ras and phosphatidylinositol 3-kinase, that in turn activate downstream effectors including MEK/ERK, AKT and mTOR. Collectively, these pathways enhance nutrient uptake and promote anabolic metabolic pathways that drive lipid, protein and nucleic acid synthesis53. Mutations that drive aberrant activation of these gatekeepers of anabolic metabolism are among the most common alterations in human cancer, thus underscoring the deep mechanistic links between metabolic reprogramming and malignant transformation. Other commonly mutated genes can also activate anabolic pathways. MYC, a transcription factor that positively regulates glucose and glutamine metabolism and coordinates anabolic gene expression programs, undergoes frequent amplification in human cancer61. Similarly, loss of wild-type p53, the most common event in human cancer, results in a constitutive shift from nutrient catabolism to anabolism, increased glycolysis and altered redox regulation62.

Many of these same pathways may regulate pluripotent cell growth. Hypoxia-inducible factor-1 (HIF1), which induces glycolysis and is frequently activated by a combination of genetic and environmental factors in tumours, triggers a glycolytic switch that promotes the metabolic transition from naïve to primed pluripotency and provides a selective advantage during early reprogramming18,25. Likewise, naïve human PSCs exhibit signatures of elevated MYC activity, and accumulation of N-MYC in the nucleus correlates with enhanced glycolytic programs20. Accordingly, sustained N-MYC activity promotes glycolysis during hPSC self-renewal and early ectoderm differentiation, whereas loss of MYC activity coincides with decreased glycolysis during endoderm and mesoderm differentiation22. Deletion of both c- and N-Myc from mouse ESCs decreases anabolic programs without affecting markers of pluripotency and triggers a state of growth arrest reminiscent of embryonic diapause, a reversible state of arrest during preimplantation embryonic development63. Similarly, partial inhibition of mTOR induces a reversible ‘paused’ state of low anabolic activity but sustained pluripotency in cultured ESCs and is sufficient to induce diapause in cultured mouse embryos64. Together, these studies raise the intriguing possibility that cellular ability to engage in anabolic metabolic programs is a prerequisite for exit from pluripotency and successful development of peri-implantation embryos.

Beyond these exciting studies, however, relatively little is known about the regulation of metabolism in PSCs by signal-transduction pathways, although many of the pathways that regulate tumour-cell metabolism have been implicated in aspects of pluripotent-cell biology. For example, loss of wild-type p53 function accelerates reprogramming, and induced PSCs tend to acquire p53 mutations under prolonged culture65,66, although these phenotypes may simply be the result of the proliferative advantage conferred by loss of p53-mediated cell-cycle regulation rather than any specific metabolic effect per se. Similarly, Akt activation promotes pluripotent self-renewal67,68, but whether this effect results from any metabolic consequences of Akt signalling remains unexplored. Nevertheless, growth-factor-mediated metabolic regulation seems likely to contribute to PSC fitness. Leukaemia inhibitory factor (LIF) sustains the undifferentiated state of naïve PSCs, and JAK/STAT3 activation downstream of LIF signalling can induce a subset of STAT3 to translocate to the mitochondria. Mitochondrial STAT3 enhances oxidative phosphorylation that supports cell proliferation but does not appear to influence self-renewal33. Likewise, fibroblast growth factor–mediated activation of MEK/ERK signalling provides a critical trigger for exit from naïve pluripotency, and thus inhibition of MEK signalling is a common component of formulations that capture PSCs in the naïve pluripotent state7. However, MEK inhibition may impose a metabolic limitation on cells that can be rescued by combined inhibition of MEK and GSK3β (ref. 69). Although there are clearly key signalling differences between cancer cells and stem cells—notably, the ability to maintain proliferation despite impaired MEK/ERK signalling appears to be a unique and defining feature of naïve ESCs8—further study into how these networks regulate metabolism in stem cells and cancer cells will continue to reveal the strategies that cells use to wire metabolic pathways to meet cell-type and context-specific demands.

Metabolic regulation of chromatin and differentiation

The influence of metabolites on gene expression programs

In addition to serving as critical substrates that fuel cell growth and proliferation, metabolites can also actively influence differentiation in both stem cells and cancer cells6,11. Several key metabolic intermediates function as obligate substrates or cofactors for enzymes that deposit or remove chemical modifications on DNA and histones. This biochemical link between metabolites and chromatin-modifying enzymes has led to the intriguing hypothesis that metabolites exert effects on cell-fate decisions primarily through altering chromatin modifications, which in turn regulate gene expression programs involved in self-renewal and lineage differentiation70. Acetylation of histones and methylation of both histones and DNA may be particularly responsive to metabolic inputs and will therefore be the focus of this discussion71 (Fig. 3).

Fig. 3: Metabolic regulation of chromatin marks.
figure3

Methyltransferase (MT) enzymes transfer methyl (Me) groups from SAM to histones and DNA. Methylation reactions generate S-adenosylhomocysteine (SAH) as a product; SAH inhibits methylation reactions, thus making MT activity responsive to the relative ratio of SAM/SAH. Methionine is the direct precursor of SAM, and methionine pools are regulated by both cellular uptake (not shown) and the methionine cycle, which regenerates methionine from homocysteine. The methionine cycle is intimately connected to the folate cycles, in which serine, glycine and (in mouse PSCs) threonine provide one-carbon units to tetrahydrofolate (THF) for transfer to homocysteine. Lysine demethylases (KDM) and TET enzymes catalyse demethylation of histones and DNA, respectively, by using αKG, oxygen and ferrous iron (not shown) as substrates. Succinate, fumarate and 2HG are competitive inhibitors of lysine demethylase and TET enzymes, thereby promoting accumulation of methyl marks. Histone acetyltransferases (HAT) transfer acetyl (Ac) groups from acetyl-CoA (Ac-CoA) to histones. Ac-CoA derives from cytosolic and/or nuclear citrate, pyruvate and acetate, which are substrates for the acetyl-CoA-generating enzymes ACL, pyruvate dehydrogenase (PDH) or ACSS2. Moreover, class I, II and IV histone deacetylase (HDAC) reactions generate acetate as a product, which can be captured by nuclear ACSS2 and used to regenerate Ac-CoA.

Acetylation of histones

Histone acetylation generally increases chromatin accessibility and activates gene transcription72. Histone acetyltransferases catalyse the transfer of acetyl groups from acetyl-CoA to histones, whereas histone deacetylases remove acetyl marks (Fig. 3). Despite its abundant production in the mitochondria to fuel the TCA cycle, acetyl-CoA cannot diffuse across the mitochondrial membrane and is thus unavailable for histone acetylation reactions73. Instead, mitochondrial acetyl-CoA (derived from pyruvate, acetate, fatty acids or amino acids) condenses with oxaloacetate, thus producing citrate, which can undergo transport out of the mitochondria. In the cytosol, ATP-citrate lyase (ACL) cleaves citrate into acetyl-CoA and oxaloacetate, thereby providing a major source of acetyl-CoA for acetylation reactions and lipid biosynthesis. In addition to cytosolic ACL, nuclear localization of ACL and the pyruvate dehydrogenase complex provide localized sources of acetyl-CoA production, thereby facilitating histone acetylation74,75,76,77. Activation of oncogenic signalling, most notably the phosphatidylinositol 3-kinase/AKT pathway, increases the generation of glucose-derived citrate and enhances activity of ACL through AKT-mediated phosphorylation74,78. This process results in increased cytosolic acetyl-CoA pools and globally enhanced histone acetylation, thus aiding in the maintenance of oncogenic programs of gene expression74,78,79. Histone acetylation may in turn further influence the metabolic state of the cell by regulating the expression of metabolic enzymes in both cancer cells and stem cells80,81,82.

Under conditions of hypoxic stress, cellular pools of citrate become limiting, and acyl-CoA synthetase short-chain family member 2 (ACSS2) provides an important source of cytosolic acetyl-CoA through the capture of free acetate83. Accordingly, tumours growing in vivo exhibit high consumption of acetate47 and dependence on ACSS2 and acetate for the maintenance of acetyl-CoA production, lipid synthesis and histone acetylation46,84. In particular, nuclear translocation of ACSS2 in response to oxygen and nutrient limitation may play a key role in maintaining histone acetylation46,85. Hypoxia and defective mitochondrial respiration also promote reductive carboxylation of glutamine-derived α-ketoglutarate (αKG) to citrate, which in turn can serve as an additional source of cytosolic acetyl-CoA for lipid production in cancer cells cultured in vitro86,87,88. The potential contribution of reductive carboxylation to histone acetylation remains unknown, but isotopic labelling experiments suggest that utilization of this pathway may be limited in tumours growing in vivo38,47.

Although histone acetylation therefore is likely to respond to defined metabolic inputs that regulate local acetyl-CoA pools, the effects of histone acetylation on stem-cell and cancer-cell differentiation vary depending on the cellular context. During early embryonic development, several TCA cycle enzymes including pyruvate dehydrogenase localize to the nucleus, where they may provide the acetyl-CoA and other metabolites that collectively help to establish the permissive chromatin landscape that facilitates zygotic genome activation77 (Fig. 4). Likewise, metabolic changes may initiate decreases in histone acetylation that allow exit from the pluripotent state. PSCs induced to differentiate experience a rapid decline in histone acetylation as a result of decreased glycolytic flux and diminished generation of glucose-derived acetyl-CoA; forced restoration of histone acetylation through supplementation with exogenous acetate impairs the onset of differentiation21. In contrast, when multipotent progenitor cells from adult tissues differentiate into specialized lineages, increased histone acetylation drives the expression of lineage-specific genetic programs. Consequently, the disruption of acetyl-CoA synthesis through ACL or ACSS2 can impair the differentiation of progenitor cells into adipocytes or neurons, respectively74,89. The degree to which metabolic regulation of histone acetylation influences cancer-cell differentiation remains uncertain. However, inhibitors of histone deacetylases exhibit particular efficacy in hematologic malignancies, at least in part through promoting terminal differentiation of transformed stem or progenitor cells90. Interestingly, histone acetylation may be intimately linked to the regulation of intracellular pH; intracellular acidification decreases global histone acetylation, whereas inhibition of histone deacetylation increases intracellular pH91. Whether coordinate regulation of histone acetylation and pH might contribute to phenotypic heterogeneity within tumours remains to be determined. Likewise, it will be important to determine whether metabolic manipulation of histone acetylation might serve as a more broadly applicable strategy to target cancer-cell differentiation.

Fig. 4: Metabolic control of differentiation in stem cells and cancer cells.
figure4

Naïve PSC self-renewal and differentiation potential depend on a characteristic open (‘poised’) chromatin structure. Metabolic inputs from αKG and acetyl-CoA (Ac-CoA) drive active demethylation and histone acetylation, thereby maintaining naïve PSC identity. In contrast, cancer cells generally exhibit a hypermethylated chromatin landscape that represses expression of both tumour suppressors and gene expression programs required for normal lineage differentiation. The hypermethylated state in cancer cells can be driven at least in part by metabolic rewiring that results in accumulation of serine, methionine, 2HG, succinate and/or fumarate.

Methylation of histones and DNA

Methylation of specific histone residues can either activate or repress transcription, whereas DNA methylation results in transcriptional repression92. The net balance of histone and DNA methylation depends on the relative rates of deposition and removal of methyl marks, a metabolically responsive process discussed in further detail below. Hypermethylation of DNA and repressive histone marks is a well-described hallmark of cancer that contributes to oncogenesis by silencing tumour suppressors and repressing gene expression programs required for normal differentiation93. Methylation changes in tumours can be heterogeneous; for example, whereas cancer-associated hypermethylation generally localizes to promoter elements, cancer cells often exhibit hypomethylation of repetitive DNA elements and gene bodies94,95,96,97,98,99. In contrast to the often-hypermethylated phenotype of cancer, early mammalian development is marked by extremely dynamic chromatin methylation100. Naïve PSCs exhibit a globally hypomethylated landscape similar to that of preimplantation blastocysts, whereas primed PSCs exhibit levels of DNA methylation consistent with postimplantation development7.

S-adenosylmethionine

Methyltransferase enzymes catalyse the transfer of methyl groups from S-adenosylmethionine (SAM) to lysine and arginine residues on histones and (primarily) cytosine nucleotides in DNA. Analogously to histone acetylation, the methylation of cancer cells and stem cells may depend on the metabolic inputs that produce and recycle the methyl donor SAM71 (Fig. 3). Intracellular SAM pools are regulated by both cellular import of methionine through amino acid transporters and the methionine cycle, which salvages the homocysteine produced by methyltransferase reactions to regenerate methionine. Moreover, remethylation of homocysteine occurs via one-carbon donation from the folate cycle, which in turn is intimately related to serine and glycine metabolism71. Physiological or pathological alterations to any of these metabolic inputs may influence histone and DNA methylation and have consequent effects on the differentiation of cancer cells and stem cells.

Many oncogenic lesions lead to enhanced uptake and/or synthesis of amino acids involved in SAM metabolism101,102,103,104,105, which in turn can influence methylation patterns and dependencies in cancer cells and in some cases affect differentiation status106,107. In cultured cancer cells, methionine restriction causes global decreases in histone methylation with decreased expression of genes implicated in cancer-cell proliferation and lineage identity108,109. In lung cancer models, increased expression of the methionine transporter LAT1 enhances intracellular SAM pools, thereby maintaining the histone methyltransferase activity of EZH2 (ref. 110). Restriction of methionine or targeting of the SAM-generating enzyme MAT2A de-represses expression of the differentiation-promoting transcription factor RXRα (ref. 110). Oncogenic cooperativity between loss of liver kinase B 1 (LKB1) and mutant KRAS results in an enhancement of serine biosynthesis to maintain DNA methylation, thus conferring a therapeutic vulnerability to inhibitors of the methionine-salvage pathway106. Acute myeloid leukaemias (AMLs) overexpress the one-carbon folate-pathway enzyme methylenetetrahydrofolate dehydrogenase-cyclohydrolase 2 (MTHFD2), and inhibiting MTHFD2 promotes myeloid differentiation of AML blasts111. These examples suggest that perturbations in SAM homeostasis may contribute to cancer progression, and it will be important for future work to elucidate the combination of factors that enable specific phenotypes to emerge from general perturbations in the availability of the required methyl donor.

Mouse PSCs derive much of their methyl-donating SAM pools from the catabolism of threonine, a metabolic feature absent in humans51,52. Threonine deprivation or inhibition of threonine dehydrogenase in mouse ESCs impairs self-renewal and potentiates differentiation. These effects appear to result from depletion of SAM pools and loss of histone H3 Lys 4 methylation, a modification important for the maintenance of the transcriptionally ‘poised’ state of pluripotency112. Analogously, human PSCs critically depend on methionine uptake and recycling for the maintenance of intracellular SAM levels. Interference with methionine metabolism causes a loss of methylation on histones (primarily H3 Lys 4) and DNA and potentiates ESC differentiation113. To date, these studies have relied on the removal of essential amino acids from PSC culture. Future work will be important to determine whether dynamic regulation of SAM metabolic pathways and/or physiological changes in nutrient availability in developing embryos are sufficient to alter gene expression programs by modulating SAM availability.

α-ketoglutarate

Although passive loss of methylation on histones and DNA can occur during cell division or as a result of nucleosome turnover114, the enzymatic removal of methyl marks plays a critical role in regulating the methylation status of cancer cells and stem cells93. The lysine-specific demethylase enzymes catalyse the removal of some histone marks in a flavin-dependent oxidative reaction. However, most demethylation reactions are catalysed by the large family of αKG-dependent dioxygenases, including the Jumonji-domain-containing histone demethylases (JHDM) and the ten-eleven-translocation (TET) enzymes, which mediate the iterative oxidation and demethylation of methylated cytosine in DNA. The activity of these enzymes depends on the availability of the co-substrates—αKG, ferrous iron and oxygen—as well as the presence of competitive inhibitors with structural similarity to αKG, such as 2-hydroxyglutarate, succinate and fumarate115. Thus, the dynamic interplay among the metabolic pathways that regulate αKG, succinate, fumarate and 2-hydroxyglutarate production and consumption collectively influences the activity of αKG-dependent dioxygenases and has potential implications for cancer cells and stem cells (Fig. 3).

In myeloid leukaemia stem cells, overexpression of the enzyme branched-chain amino acid transaminase 1 (BCAT1) contributes to oncogenesis through direct effects on αKG levels. Specifically, BCAT1 depletes intracellular αKG, thus resulting in impaired TET enzyme activity, DNA hypermethylation and blockade of normal myeloid differentiation116. Ascorbate (vitamin C) enhances the activity of αKG-dependent dioxygenases by facilitating the regeneration of ferrous iron117. Several recent studies have demonstrated that ascorbate supplementation enhances TET-mediated DNA demethylation, which in turn promotes differentiation of leukaemia cells118,119. Enhanced ascorbate uptake has further been implicated in the self-renewal of normal haematopoietic stem cells via the regulation of TET-mediated demethylation118. In human melanoma tumours, hypoxic central regions exhibit substantial depletion of extracellular glutamine and a concomitant decrease in intracellular αKG levels and increase in repressive histone methylation120. Importantly, the hypermethylated phenotype promotes cancer-cell dedifferentiation and therapeutic resistance, effects that can be reversed by inhibition of the histone methyltransferase EZH2.

In contrast to the pro-differentiation effects observed in cancer cells, metabolic activation of αKG-dependent dioxygenases has notably context-specific roles in PSCs, probably as a consequence of the vastly different chromatin states in naïve and primed PSCs. Metabolic pathways in naïve mouse PSCs are wired to promote accumulation of intracellular αKG, perhaps in part to promote activity of the chromatin-modifying enzymes that sustain the open-chromatin landscape characteristic of naïve pluripotency14,121. Interventions that increase αKG drive the loss of repressive chromatin modifications and favour self-renewal over differentiation14,121. In another recent example, the protein prohibitin was identified as a key regulator of human ESC self-renewal. Mechanistic studies have demonstrated that prohibitin interacts with histone H3.3 and consequently regulates the transcription of the isocitrate dehydrogenase enzymes, which produce the αKG required to maintain demethylation and pluripotency122. In contrast to the preceding examples, αKG accelerates differentiation of the more heavily methylated primed human PSCs, in agreement with the requirement to demethylate and de-repress lineage-specific regions as part of the differentiation process31. These data suggest that metabolic pathways may facilitate major transitions in the global chromatin landscape. In support of this hypothesis, ascorbic acid–mediated activation of αKG-dependent dioxygenases, including both JHDM and TET enzymes, has been well established to enhance reprogramming to pluripotency, a process requiring erasure of somatic methylation patterns123.

Antagonists of α-ketoglutarate

Perhaps the best examples of metabolic regulation of chromatin come from the seminal discovery of cancer-associated mutations in the isocitrate dehydrogenase (IDH) enzymes. IDH1/2 enzymes normally catalyse the interconversion of isocitrate and αKG in an NADP(H)-dependent manner. Oncogenic mutations in IDH1/2 occur at specific arginine residues in the active site of the enzyme and confer a neomorphic enzymatic activity that efficiently reduces αKG to the structurally similar metabolite d-2-hydroxyglutarate (d-2HG)124,125. d-2HG is a competitive inhibitor of αKG-dependent dioxygenases126,127,128, and exogenous d-2HG is sufficient to induce many of the oncogenic effects of IDH mutations129. Although the relative importance of each enzyme inhibited by d-2HG is still being elucidated, inhibition of the TET and JHDM enzymes appears to play a key role in oncogenesis mediated by mutant IDH and d-2HG130,131. By poisoning the function of the TET and JHDM enzymes, d-2HG promotes hypermethylation of DNA and repressive histone marks, thus locking IDH-mutant cancer cells in an undifferentiated stem-cell-like state and conferring a differentiation ‘hit’ that contributes to oncogenesis130,131,132,133. In both preclinical models and patients with AML, targeted inhibition of the function of the mutant IDH enzyme suppresses d-2HG production, thereby reversing repressive methylation marks and inducing the differentiation of IDH-mutant stem or progenitor cells134,135. Ongoing efforts seek to define physiological sources and the biological effects of d-2HG and its l-2HG enantiomer; these studies should determine whether targeted inhibition of 2HG metabolism might be a more broadly applicable strategy to modulate chromatin structure, cell differentiation and oncogenesis136,137,138.

Similarly to 2HG, succinate and fumarate are competitive inhibitors of αKG-dependent dioxygenases139. Catabolism of succinate and fumarate depends on the activity of the TCA cycle enzymes succinate dehydrogenase (SDH) and fumarate hydratase (FH), respectively. Inactivating lesions in components of the SDH complex or FH have been implicated in the pathogenesis of a variety of rare cancers and inherited cancer-predisposition syndromes140. Loss of SDH or FH activity results in cellular accumulation of succinate or fumarate, thus leading to stabilization of HIF-1 via inhibition of αKG-dependent prolyl hydroxylases that normally target HIF-1 for degradation in the presence of oxygen141,142. This ‘pseudohypoxic’ state of SDH- and FH-deficient tumours contributes to oncogenesis by driving glycolytic metabolism and tumour angiogenesis140. More recent evidence indicates that the accumulation of fumarate and succinate also drives widespread hypermethylation of DNA and repressive histone marks143,144, thus altering cancer-cell differentiation, at least in part by inducing genetic programs that promote an invasive mesenchymal phenotype145. Importantly, restoration of the balance of αKG relative to succinate/fumarate reverses the hypermethylated phenotype and promotes HIF-1 degradation143,146.

By coupling chromatin modifications to the metabolic state, cellular differentiation decisions may be influenced by nutrient availability and microenvironmental conditions (for example, oxygen availability and pH). It will be interesting for future work to examine how competition for scarce resources in the tumour microenvironment and the adult stem-cell niche affects chromatin regulation and gene expression programs. The consequences of metabolic regulation of chromatin structure often result in divergent effects on the differentiation state of cancer cells and stem cells (Fig. 4). This context specificity probably arises as a consequence of the many factors that collectively determine gene expression programs, including local metabolite levels, the relative sensitivity of chromatin-modifying enzymes to metabolic fluctuations and the expression and recruitment of relevant transcription factors and co-activators to key genetic loci6. Understanding how global shifts in nutrient availability translate into specific gene expression programs is therefore a major area for future investigation.

Conclusions and future directions

Metabolites that are key intermediates of central metabolic pathways supporting bioenergetics and anabolic growth can also influence the regulation of gene expression programs and cell-fate decisions. Therefore, the particular bioenergetic requirements of a given cell—and the pathways engaged to meet those requirements—may have important consequences for the regulation of cell identity. Indeed, although all cells proliferating in vitro appear to share several canonical hallmarks of proliferative metabolism, accumulating evidence demonstrates that there is no single metabolic profile of a cancer cell, just as there is no single profile of a stem cell. The diversity of metabolic strategies that support cell proliferation and the factors that regulate these myriad metabolic profiles are probably only just beginning to be understood. Clearly, both cell-intrinsic and cell-extrinsic factors have critical roles in determining cellular metabolic phenotypes.

Interestingly, to date, the metabolic plasticity of cancer cells is largely revealed in vivo, whereas PSCs may engage a wider array of metabolic strategies in vitro. Because many oncogenes and tumour suppressors regulate cellular metabolism, the very oncogenic lesions that drive malignant identity may also lock cultured cancer cells into a relatively inflexible metabolic identity. In contrast, the diversity of extracellular stimuli experienced in vivo may enable rewiring of both intracellular signalling and metabolic pathways even in malignant cells. Similarly, although several key signalling pathways sustain the growth of PSCs, these pathways are dynamic and largely tied to the regulation of cell identity. Therefore, the inherent metabolic plasticity of PSCs might possibly arise as a consequence of their functional plasticity.

Several avenues of research may provide valuable insight into the connections among metabolic pathways, gene regulation and cell-fate decisions in cancer and development. First, inborn errors of metabolism are a largely untapped opportunity for elucidating how defined changes in cellular metabolite levels influence cellular phenotypes and disease susceptibility147. Some intriguing examples are beginning to emerge. Heterozygous mutations in genes encoding FH or subunits of SDH are associated with susceptibility to specific cancer syndromes and widespread changes in chromatin modifications, in agreement with inhibition of αKG-dependent dioxygenases mediated by high levels of fumarate or succinate, respectively140,143. Similarly, biallelic mutation of the gene encoding the l-2HG dehydrogenase, which converts l-2HG to αKG, drives high levels of l-2HG and predisposes patients to central-nervous-system malignancies in addition to other adverse metabolic consequences138,148,149,150. Intriguingly, inborn errors of metabolism that result in elevated d-2HG, including mutations in the d-2HG dehydrogenase and gain-of-function mutations in IDH2, induce phenotypes including developmental delays and cardiomyopathy but do not appear to be associated with an increased risk of cancer148,151,152. How specific metabolic alterations contribute to developmental phenotypes, why certain metabolic perturbations predispose individuals to tumorigenesis and why some tissues are more or less sensitive to transformation in the background of these germline mutations remain key open questions that may be illuminated by the continued study of inborn errors of metabolism.

Second, cancer stem cells (or tumour-initiating cells) are an intriguing example of cells that can exhibit features of both normal stem cells and aberrantly proliferating tumour cells153. To date, the metabolism of these tumour-initiating cells remains poorly understood—indeed, the possibility of one single metabolic signature of a cancer stem cell seems unlikely, just as there is no single metabolic phenotype of adult stem cells from which these cancer stem cells may arise153,154. Some studies suggest that cancer stem cells exhibit decreased mitochondrial function and increased dependence on glycolytic flux155,156,157,158,159,160, whereas other studies have reported enhanced dependence on mitochondrial function and oxidative phosphorylation161,162,163,164,165,166. Lipid metabolism also appears to play an important role in cancer stem-cell maintenance by supplying bioenergetic substrates and potentially influencing cell-fate decisions156,167,168,169,170,171. Cell lineage, environmental niche and dynamic cellular phenotypes—including the switch from self-renewal to transient amplification and differentiation—are likely to result in a heterogeneous metabolic profile. Nevertheless, continued study of the metabolic requirements of adult stem cells and their transformed counterparts may yield critical insights into how metabolism supports both normal and aberrant stem-cell phenotypes and may provide opportunities for targeted therapeutic intervention.

The intersection between metabolic activity and the regulation of cell-fate decisions is only beginning to be understood, and several outstanding questions will shape future research in this area. First, the mechanisms linking metabolic profiles to cellular phenotypes remain largely unidentified. Although the most compelling hypothesis is that metabolic regulation of chromatin mechanistically links metabolic profiles to gene expression programs, careful genetic experiments will be required to determine the extent to which individual metabolites influence cell fate through specific chromatin modifications. Second, the aspects of cellular metabolic phenotypes that result from bioenergetic demands as opposed to intrinsically determined metabolic preferences linked to cell identity remain an open question. Third, determining whether physiological changes in nutrient abundance contribute to altered cell-fate decisions during cancer progression or embryonic development will be important. Addressing these questions will require new, sensitive techniques to assess the metabolism of rare cell populations in vivo. Together, these studies should enable a better understanding of whether individual metabolic features are causes or consequences of changing cellular programs, and should reveal opportunities for targeted manipulation of cellular metabolism that hold promise for anticancer therapies and regenerative-medicine approaches.

References

  1. 1.

    Vander Heiden, M. G., Cantley, L. C. & Thompson, C. B. Understanding the Warburg effect: the metabolic requirements of cell proliferation. Science 324, 1029–1033 (2009).

  2. 2.

    Beaudin, A. E. & Stover, P. J. Insights into metabolic mechanisms underlying folate-responsive neural tube defects: a minireview. Birth Defects Res. A. Clin. Mol. Teratol. 85, 274–284 (2009).

  3. 3.

    Luengo, A., Gui, D. Y. & Vander Heiden, M. G. Targeting metabolism for cancer therapy. Cell Chem. Biol. 24, 1161–1180 (2017).

  4. 4.

    Kong, H. & Chandel, N. S. Regulation of redox balance in cancer and T cells. J. Biol. Chem. 293, 7499–7507 (2018).

  5. 5.

    Chantranupong, L., Wolfson, R. L. & Sabatini, D. M. Nutrient-sensing mechanisms across evolution. Cell 161, 67–83 (2015).

  6. 6.

    Schvartzman, J. M., Thompson, C. B. & Finley, L. W. S. Metabolic regulation of chromatin modifications and gene expression. J. Cell Biol. 217, 2247–2259 (2018).

  7. 7.

    Weinberger, L., Ayyash, M., Novershtern, N. & Hanna, J. H. Dynamic stem cell states: naive to primed pluripotency in rodents and humans. Nat. Rev. Mol. Cell Biol. 17, 155–169 (2016).

  8. 8.

    Martello, G. & Smith, A. The nature of embryonic stem cells. Annu. Rev. Cell Dev. Biol. 30, 647–675 (2014).

  9. 9.

    Fan, J. et al. Glutamine-driven oxidative phosphorylation is a major ATP source in transformed mammalian cells in both normoxia and hypoxia. Mol. Syst. Biol. 9, 712 (2013).

  10. 10.

    Hosios, A. M. et al. Amino acids rather than glucose account for the majority of cell mass in proliferating mammalian cells. Dev. Cell 36, 540–549 (2016).

  11. 11.

    Pavlova, N. N. & Thompson, C. B. The emerging hallmarks of cancer metabolism. Cell Metab. 23, 27–47 (2016).

  12. 12.

    DeBerardinis, R. J. & Chandel, N. S. Fundamentals of cancer metabolism. Sci. Adv. 2, e1600200 (2016).

  13. 13.

    Vander Heiden, M. G. & DeBerardinis, R. J. Understanding the intersections between metabolism and cancer biology. Cell 168, 657–669 (2017).

  14. 14.

    Carey, B. W., Finley, L. W., Cross, J. R., Allis, C. D. & Thompson, C. B. Intracellular α-ketoglutarate maintains the pluripotency of embryonic stem cells. Nature 518, 413–416 (2015).

  15. 15.

    Tohyama, S. et al. Glutamine oxidation is indispensable for survival of human pluripotent stem cells. Cell Metab. 23, 663–674 (2016).

  16. 16.

    Boroughs, L. K. & DeBerardinis, R. J. Metabolic pathways promoting cancer cell survival and growth. Nat. Cell Biol. 17, 351–359 (2015).

  17. 17.

    Chung, S. et al. Mitochondrial oxidative metabolism is required for the cardiac differentiation of stem cells. Nat. Clin. Pract. Cardiovasc. Med. 4(Suppl. 1), S60–S67 (2007).

  18. 18.

    Zhou, W. et al. HIF1α induced switch from bivalent to exclusively glycolytic metabolism during ESC-to-EpiSC/hESC transition. EMBO J. 31, 2103–2116 (2012).

  19. 19.

    Zhang, J. et al. UCP2 regulates energy metabolism and differentiation potential of human pluripotent stem cells. EMBO J. 30, 4860–4873 (2011).

  20. 20.

    Gu, W. et al. Glycolytic metabolism plays a functional role in regulating human pluripotent stem cell state. Cell Stem Cell 19, 476–490 (2016).

  21. 21.

    Moussaieff, A. et al. Glycolysis-mediated changes in acetyl-CoA and histone acetylation control the early differentiation of embryonic stem cells. Cell Metab. 21, 392–402 (2015).

  22. 22.

    Cliff, T. S. et al. MYC controls human pluripotent stem cell fate decisions through regulation of metabolic flux. Cell Stem Cell 21, 502–516 (2017).

  23. 23.

    Folmes, C. D. et al. Somatic oxidative bioenergetics transitions into pluripotency-dependent glycolysis to facilitate nuclear reprogramming. Cell Metab. 14, 264–271 (2011).

  24. 24.

    Panopoulos, A. D. et al. The metabolome of induced pluripotent stem cells reveals metabolic changes occurring in somatic cell reprogramming. Cell Res. 22, 168–177 (2012).

  25. 25.

    Mathieu, J. et al. Hypoxia-inducible factors have distinct and stage-specific roles during reprogramming of human cells to pluripotency. Cell Stem Cell 14, 592–605 (2014).

  26. 26.

    Kim, H. et al. Core pluripotency factors directly regulate metabolism in embryonic stem cell to maintain pluripotency. Stem Cells 33, 2699–2711 (2015).

  27. 27.

    Liberti, M. V. & Locasale, J. W. The Warburg effect: how does it benefit cancer cells? Trends Biochem. Sci. 41, 211–218 (2016).

  28. 28.

    Prigione, A., Fauler, B., Lurz, R., Lehrach, H. & Adjaye, J. The senescence-related mitochondrial/oxidative stress pathway is repressed in human induced pluripotent stem cells. Stem Cells 28, 721–733 (2010).

  29. 29.

    St John, J. C. et al. The expression of mitochondrial DNA transcription factors during early cardiomyocyte in vitro differentiation from human embryonic stem cells. Cloning Stem Cells 7, 141–153 (2005).

  30. 30.

    Zhang, H. et al. Distinct metabolic states can support self-renewal and lipogenesis in human pluripotent stem cells under different culture conditions. Cell Rep. 16, 1536–1547 (2016).

  31. 31.

    TeSlaa, T. et al. α-Ketoglutarate accelerates the initial differentiation of primed human pluripotent stem cells. Cell Metab. 24, 485–493 (2016).

  32. 32.

    Takashima, Y. et al. Resetting transcription factor control circuitry toward ground-state pluripotency in human. Cell 162, 452–453 (2015).

  33. 33.

    Carbognin, E., Betto, R. M., Soriano, M. E., Smith, A. G. & Martello, G. Stat3 promotes mitochondrial transcription and oxidative respiration during maintenance and induction of naive pluripotency. EMBO J. 35, 618–634 (2016).

  34. 34.

    Setoguchi, K., TeSlaa, T., Koehler, C. M. & Teitell, M. A. P53 regulates rapid apoptosis in human pluripotent stem cells. J. Mol. Biol. 428, 1465–1475 (2016).

  35. 35.

    Madden, D. T., Davila-Kruger, D., Melov, S. & Bredesen, D. E. Human embryonic stem cells express elevated levels of multiple pro-apoptotic BCL-2 family members. PLoS One 6, e28530 (2011).

  36. 36.

    Yanes, O. et al. Metabolic oxidation regulates embryonic stem cell differentiation. Nat. Chem. Biol. 6, 411–417 (2010).

  37. 37.

    Muir, A., Danai, L. V. & Vander Heiden, M. G. Microenvironmental regulation of cancer cell metabolism: implications for experimental design and translational studies. Dis. Model Mech. 11, dmm035758 (2018).

  38. 38.

    Marin-Valencia, I. et al. Analysis of tumor metabolism reveals mitochondrial glucose oxidation in genetically diverse human glioblastomas in the mouse brain in vivo. Cell Metab. 15, 827–837 (2012).

  39. 39.

    Davidson, S. M. et al. Environment impacts the metabolic dependencies of ras-driven non-small cell lung cancer. Cell Metab. 23, 517–528 (2016).

  40. 40.

    Hensley, C. T. et al. Metabolic heterogeneity in human lung tumors. Cell 164, 681–694 (2016).

  41. 41.

    Courtney, K. D. et al. Isotope tracing of human clear cell renal cell carcinomas demonstrates suppressed glucose oxidation In Vivo. Cell Metab. 28, 793–800.e2 (2018).

  42. 42.

    Tardito, S. et al. Glutamine synthetase activity fuels nucleotide biosynthesis and supports growth of glutamine-restricted glioblastoma. Nat. Cell Biol. 17, 1556–1568 (2015).

  43. 43.

    Muir, A. et al. Environmental cystine drives glutamine anaplerosis and sensitizes cancer cells to glutaminase inhibition. eLife 6, e27713 (2017).

  44. 44.

    Hui, S. et al. Glucose feeds the TCA cycle via circulating lactate. Nature 551, 115–118 (2017).

  45. 45.

    Faubert, B. et al. Lactate metabolism in human lung tumors. Cell 171, 358–371.e359 (2017).

  46. 46.

    Comerford, S. A. et al. Acetate dependence of tumors. Cell 159, 1591–1602 (2014).

  47. 47.

    Mashimo, T. et al. Acetate is a bioenergetic substrate for human glioblastoma and brain metastases. Cell 159, 1603–1614 (2014).

  48. 48.

    Mayers, J. R. et al. Tissue of origin dictates branched-chain amino acid metabolism in mutant Kras-driven cancers. Science 353, 1161–1165 (2016).

  49. 49.

    Yuneva, M. O. et al. The metabolic profile of tumors depends on both the responsible genetic lesion and tissue type. Cell Metab. 15, 157–170 (2012).

  50. 50.

    Zhang, J. et al. LIN28 regulates stem cell metabolism and conversion to primed pluripotency. Cell Stem Cell 19, 66–80 (2016).

  51. 51.

    Shyh-Chang, N. et al. Influence of threonine metabolism on S-adenosylmethionine and histone methylation. Science 339, 222–226 (2013).

  52. 52.

    Wang, J. et al. Dependence of mouse embryonic stem cells on threonine catabolism. Science 325, 435–439 (2009).

  53. 53.

    Palm, W. & Thompson, C. B. Nutrient acquisition strategies of mammalian cells. Nature 546, 234–242 (2017).

  54. 54.

    Zhang, J. et al. Metabolism in pluripotent stem cells and early mammalian development. Cell Metab. 27, 332–338 (2018).

  55. 55.

    Tang, F. et al. Tracing the derivation of embryonic stem cells from the inner cell mass by single-cell RNA-Seq analysis. Cell Stem Cell 6, 468–478 (2010).

  56. 56.

    Kaneko, K. J. Metabolism of preimplantation embryo development: a bystander or an active participant? Curr. Top. Dev. Biol. 120, 259–310 (2016).

  57. 57.

    Houghton, F. D., Thompson, J. G., Kennedy, C. J. & Leese, H. J. Oxygen consumption and energy metabolism of the early mouse embryo. Mol. Reprod. Dev. 44, 476–485 (1996).

  58. 58.

    Houghton, F. D. Energy metabolism of the inner cell mass and trophectoderm of the mouse blastocyst. Differentiation 74, 11–18 (2006).

  59. 59.

    Brinster, R. L. Effect of glutathione on the development of two-cell mouse embryos in vitro. J. Reprod. Fertil. 17, 521–525 (1968).

  60. 60.

    Cholewa, J. A. & Whitten, W. K. Development of two-cell mouse embryos in the absence of a fixed-nitrogen source. J. Reprod. Fertil. 22, 553–555 (1970).

  61. 61.

    Stine, Z. E., Walton, Z. E., Altman, B. J., Hsieh, A. L. & Dang, C. V. MYC, metabolism, and cancer. Cancer Discov. 5, 1024–1039 (2015).

  62. 62.

    Kruiswijk, F., Labuschagne, C. F. & Vousden, K. H. p53 in survival, death and metabolic health: a lifeguard with a licence to kill. Nat. Rev. Mol. Cell Biol. 16, 393–405 (2015).

  63. 63.

    Scognamiglio, R. et al. Myc depletion induces a pluripotent dormant state mimicking diapause. Cell 164, 668–680 (2016).

  64. 64.

    Bulut-Karslioglu, A. et al. Inhibition of mTOR induces a paused pluripotent state. Nature 540, 119–123 (2016).

  65. 65.

    Merkle, F. T. et al. Human pluripotent stem cells recurrently acquire and expand dominant negative P53 mutations. Nature 545, 229–233 (2017).

  66. 66.

    Hanna, J. et al. Direct cell reprogramming is a stochastic process amenable to acceleration. Nature 462, 595–601 (2009).

  67. 67.

    Paling, N. R., Wheadon, H., Bone, H. K. & Welham, M. J. Regulation of embryonic stem cell self-renewal by phosphoinositide 3-kinase-dependent signaling. J. Biol. Chem. 279, 48063–48070 (2004).

  68. 68.

    Watanabe, S. et al. Activation of Akt signaling is sufficient to maintain pluripotency in mouse and primate embryonic stem cells. Oncogene 25, 2697–2707 (2006).

  69. 69.

    Ying, Q. L. et al. The ground state of embryonic stem cell self-renewal. Nature 453, 519–523 (2008).

  70. 70.

    Reid, M. A., Dai, Z. & Locasale, J. W. The impact of cellular metabolism on chromatin dynamics and epigenetics. Nat. Cell Biol. 19, 1298–1306 (2017).

  71. 71.

    Su, X., Wellen, K. E. & Rabinowitz, J. D. Metabolic control of methylation and acetylation. Curr. Opin. Chem. Biol. 30, 52–60 (2016).

  72. 72.

    Tessarz, P. & Kouzarides, T. Histone core modifications regulating nucleosome structure and dynamics. Nat. Rev. Mol. Cell Biol. 15, 703–708 (2014).

  73. 73.

    Sivanand, S., Viney, I. & Wellen, K. E. Spatiotemporal control of acetyl-CoA metabolism in chromatin regulation. Trends Biochem. Sci. 43, 61–74 (2018).

  74. 74.

    Wellen, K. E. et al. ATP-citrate lyase links cellular metabolism to histone acetylation. Science 324, 1076–1080 (2009).

  75. 75.

    Sutendra, G. et al. A nuclear pyruvate dehydrogenase complex is important for the generation of acetyl-CoA and histone acetylation. Cell 158, 84–97 (2014).

  76. 76.

    Sivanand, S. et al. Nuclear acetyl-CoA production by ACLY promotes homologous recombination. Mol. Cell 67, 252–265.e256 (2017).

  77. 77.

    Nagaraj, R. et al. Nuclear localization of mitochondrial TCA cycle enzymes as a critical step in mammalian zygotic genome activation. Cell 168, 210–223.e211 (2017).

  78. 78.

    Lee, J. V. et al. Akt-dependent metabolic reprogramming regulates tumor cell histone acetylation. Cell Metab. 20, 306–319 (2014).

  79. 79.

    Cai, L., Sutter, B. M., Li, B. & Tu, B. P. Acetyl-CoA induces cell growth and proliferation by promoting the acetylation of histones at growth genes. Mol. Cell 42, 426–437 (2011).

  80. 80.

    Sebastián, C. et al. The histone deacetylase SIRT6 is a tumor suppressor that controls cancer metabolism. Cell 151, 1185–1199 (2012).

  81. 81.

    Zhong, L. et al. The histone deacetylase Sirt6 regulates glucose homeostasis via Hif1α. Cell 140, 280–293 (2010).

  82. 82.

    Yang, J. et al. Inhibiting histone deacetylases suppresses glucose metabolism and hepatocellular carcinoma growth by restoring FBP1 expression. Sci. Rep. 7, 43864 (2017).

  83. 83.

    Kamphorst, J. J., Chung, M. K., Fan, J. & Rabinowitz, J. D. Quantitative analysis of acetyl-CoA production in hypoxic cancer cells reveals substantial contribution from acetate. Cancer Metab. 2, 23 (2014).

  84. 84.

    Schug, Z. T. et al. Acetyl-CoA synthetase 2 promotes acetate utilization and maintains cancer cell growth under metabolic stress. Cancer Cell 27, 57–71 (2015).

  85. 85.

    Bulusu, V. et al. Acetate recapturing by nuclear acetyl-CoA synthetase 2 prevents loss of histone acetylation during oxygen and serum limitation. Cell Rep. 18, 647–658 (2017).

  86. 86.

    Metallo, C. M. et al. Reductive glutamine metabolism by IDH1 mediates lipogenesis under hypoxia. Nature 481, 380–384 (2011).

  87. 87.

    Mullen, A. R. et al. Reductive carboxylation supports growth in tumour cells with defective mitochondria. Nature 481, 385–388 (2011).

  88. 88.

    Wise, D. R. et al. Hypoxia promotes isocitrate dehydrogenase-dependent carboxylation of α-ketoglutarate to citrate to support cell growth and viability. Proc. Natl Acad. Sci. USA 108, 19611–19616 (2011).

  89. 89.

    Mews, P. et al. Acetyl-CoA synthetase regulates histone acetylation and hippocampal memory. Nature 546, 381–386 (2017).

  90. 90.

    de Thé, H. Differentiation therapy revisited. Nat. Rev. Cancer 18, 117–127 (2018).

  91. 91.

    McBrian, M. A. et al. Histone acetylation regulates intracellular pH. Mol. Cell 49, 310–321 (2013).

  92. 92.

    Berger, S. L. The complex language of chromatin regulation during transcription. Nature 447, 407–412 (2007).

  93. 93.

    Plass, C. et al. Mutations in regulators of the epigenome and their connections to global chromatin patterns in cancer. Nat. Rev. Genet. 14, 765–780 (2013).

  94. 94.

    Feinberg, A. P. & Vogelstein, B. Hypomethylation distinguishes genes of some human cancers from their normal counterparts. Nature 301, 89–92 (1983).

  95. 95.

    Kulis, M. et al. Epigenomic analysis detects widespread gene-body DNA hypomethylation in chronic lymphocytic leukemia. Nat. Genet. 44, 1236–1242 (2012).

  96. 96.

    Ehrlich, M. & Lacey, M. DNA hypomethylation and hemimethylation in cancer. Adv. Exp. Med. Biol. 754, 31–56 (2013).

  97. 97.

    Jäkel, C. et al. Genome-wide genetic and epigenetic analyses of pancreatic acinar cell carcinomas reveal aberrations in genome stability. Nat. Commun. 8, 1323 (2017).

  98. 98.

    Ehrlich, M. DNA methylation in cancer: too much, but also too little. Oncogene 21, 5400–5413 (2002).

  99. 99.

    Kulis, M. & Esteller, M. DNA methylation and cancer. Adv. Genet. 70, 27–56 (2010).

  100. 100.

    Smith, Z. D. et al. DNA methylation dynamics of the human preimplantation embryo. Nature 511, 611–615 (2014).

  101. 101.

    DeNicola, G. M. et al. NRF2 regulates serine biosynthesis in non-small cell lung cancer. Nat. Genet. 47, 1475–1481 (2015).

  102. 102.

    Locasale, J. W. et al. Phosphoglycerate dehydrogenase diverts glycolytic flux and contributes to oncogenesis. Nat. Genet. 43, 869–874 (2011).

  103. 103.

    Possemato, R. et al. Functional genomics reveal that the serine synthesis pathway is essential in breast cancer. Nature 476, 346–350 (2011).

  104. 104.

    Maddocks, O. D. et al. Serine starvation induces stress and p53-dependent metabolic remodelling in cancer cells. Nature 493, 542–546 (2013).

  105. 105.

    Edinger, A. L. & Thompson, C. B. Akt maintains cell size and survival by increasing mTOR-dependent nutrient uptake. Mol. Biol. Cell. 13, 2276–2288 (2002).

  106. 106.

    Kottakis, F. et al. LKB1 loss links serine metabolism to DNA methylation and tumorigenesis. Nature 539, 390–395 (2016).

  107. 107.

    Maddocks, O. D., Labuschagne, C. F., Adams, P. D. & Vousden, K. H. Serine metabolism supports the methionine cycle and DNA/RNA methylation through de novo ATP synthesis in cancer cells. Mol. Cell 61, 210–221 (2016).

  108. 108.

    Mentch, S. J. et al. Histone methylation dynamics and gene regulation occur through the sensing of one-carbon metabolism. Cell Metab. 22, 861–873 (2015).

  109. 109.

    Dai, Z., Mentch, S. J., Gao, X., Nichenametla, S. N. & Locasale, J. W. Methionine metabolism influences genomic architecture and gene expression through H3K4me3 peak width. Nat. Commun. 9, 1955 (2018).

  110. 110.

    Dann, S. G. et al. Reciprocal regulation of amino acid import and epigenetic state through Lat1 and EZH2. EMBO J. 34, 1773–1785 (2015).

  111. 111.

    Pikman, Y. et al. Targeting MTHFD2 in acute myeloid leukemia. J. Exp. Med. 213, 1285–1306 (2016).

  112. 112.

    Bernstein, B. E. et al. A bivalent chromatin structure marks key developmental genes in embryonic stem cells. Cell 125, 315–326 (2006).

  113. 113.

    Shiraki, N. et al. Methionine metabolism regulates maintenance and differentiation of human pluripotent stem cells. Cell Metab. 19, 780–794 (2014).

  114. 114.

    Chory, E. J. et al. Nucleosome turnover regulates histone methylation patterns over the genome. Mol. Cell 73, 61–72.e3 (2018).

  115. 115.

    Losman, J. A. & Kaelin, W. G. Jr. What a difference a hydroxyl makes: mutant IDH, (R)-2-hydroxyglutarate, and cancer. Genes Dev. 27, 836–852 (2013).

  116. 116.

    Raffel, S. et al. BCAT1 restricts αKG levels in AML stem cells leading to IDHmut-like DNA hypermethylation. Nature 551, 384–388 (2017).

  117. 117.

    Knowles, H. J., Raval, R. R., Harris, A. L. & Ratcliffe, P. J. Effect of ascorbate on the activity of hypoxia-inducible factor in cancer cells. Cancer Res. 63, 1764–1768 (2003).

  118. 118.

    Agathocleous, M. et al. Ascorbate regulates haematopoietic stem cell function and leukaemogenesis. Nature 549, 476–481 (2017).

  119. 119.

    Cimmino, L. et al. Restoration of TET2 function blocks aberrant self-renewal and leukemia progression. Cell 170, 1079–1095.e1020 (2017).

  120. 120.

    Pan, M. et al. Regional glutamine deficiency in tumours promotes dedifferentiation through inhibition of histone demethylation. Nat. Cell Biol. 18, 1090–1101 (2016).

  121. 121.

    Hwang, I. Y. et al. Psat1-dependent fluctuations in α-ketoglutarate affect the timing of ESC differentiation. Cell Metab. 24, 494–501 (2016).

  122. 122.

    Zhu, Z. et al. PHB associates with the HIRA complex to control an epigenetic-metabolic circuit in human ESCs. Cell Stem Cell 20, 274–289.e277 (2017).

  123. 123.

    Cimmino, L., Neel, B. G. & Aifantis, I. Vitamin C in stem cell reprogramming and cancer. Trends Cell Biol. 28, 698–708 (2018).

  124. 124.

    Dang, L. et al. Cancer-associated IDH1 mutations produce 2-hydroxyglutarate. Nature 462, 739–744 (2009).

  125. 125.

    Ward, P. S. et al. The common feature of leukemia-associated IDH1 and IDH2 mutations is a neomorphic enzyme activity converting alpha-ketoglutarate to 2-hydroxyglutarate. Cancer Cell 17, 225–234 (2010).

  126. 126.

    Xu, W. et al. Oncometabolite 2-hydroxyglutarate is a competitive inhibitor of α-ketoglutarate-dependent dioxygenases. Cancer Cell 19, 17–30 (2011).

  127. 127.

    Chowdhury, R. et al. The oncometabolite 2-hydroxyglutarate inhibits histone lysine demethylases. EMBO Rep. 12, 463–469 (2011).

  128. 128.

    Koivunen, P. et al. Transformation by the (R)-enantiomer of 2-hydroxyglutarate linked to EGLN activation. Nature 483, 484–488 (2012).

  129. 129.

    Losman, J. A. et al. 2-hydroxyglutarate is sufficient to promote leukemogenesis and its effects are reversible. Science 339, 1621–1625 (2013).

  130. 130.

    Lu, C. et al. IDH mutation impairs histone demethylation and results in a block to cell differentiation. Nature 483, 474–478 (2012).

  131. 131.

    Figueroa, M. E. et al. Leukemic IDH1 and IDH2 mutations result in a hypermethylation phenotype, disrupt TET2 function, and impair hematopoietic differentiation. Cancer Cell 18, 553–567 (2010).

  132. 132.

    Saha, S. K. et al. Mutant IDH inhibits HNF-4α to block hepatocyte differentiation and promote biliary cancer. Nature 513, 110–114 (2014).

  133. 133.

    Sasaki, M. et al. IDH1(R132H) mutation increases murine haematopoietic progenitors and alters epigenetics. Nature 488, 656–659 (2012).

  134. 134.

    Amatangelo, M. D. et al. Enasidenib induces acute myeloid leukemia cell differentiation to promote clinical response. Blood 130, 732–741 (2017).

  135. 135.

    Chen, C. et al. Cancer-associated IDH2 mutants drive an acute myeloid leukemia that is susceptible to Brd4 inhibition. Genes Dev. 27, 1974–1985 (2013).

  136. 136.

    Fan, J. et al. Human phosphoglycerate dehydrogenase produces the oncometabolite D-2-hydroxyglutarate. ACS Chem. Biol. 10, 510–516 (2015).

  137. 137.

    Intlekofer, A. M. et al. Hypoxia induces production of L-2-hydroxyglutarate. Cell Metab. 22, 304–311 (2015).

  138. 138.

    Ye, D., Guan, K. L. & Xiong, Y. Metabolism, activity, and targeting of D- and L-2-hydroxyglutarates. Trends Cancer 4, 151–165 (2018).

  139. 139.

    Kaelin, W. G. Jr & McKnight, S. L. Influence of metabolism on epigenetics and disease. Cell 153, 56–69 (2013).

  140. 140.

    Gottlieb, E. & Tomlinson, I. P. Mitochondrial tumour suppressors: a genetic and biochemical update. Nat. Rev. Cancer 5, 857–866 (2005).

  141. 141.

    Selak, M. A. et al. Succinate links TCA cycle dysfunction to oncogenesis by inhibiting HIF-alpha prolyl hydroxylase. Cancer Cell. 7, 77–85 (2005).

  142. 142.

    Isaacs, J. S. et al. HIF overexpression correlates with biallelic loss of fumarate hydratase in renal cancer: novel role of fumarate in regulation of HIF stability. Cancer Cell. 8, 143–153 (2005).

  143. 143.

    Xiao, M. et al. Inhibition of α-KG-dependent histone and DNA demethylases by fumarate and succinate that are accumulated in mutations of FH and SDH tumor suppressors. Genes Dev. 26, 1326–1338 (2012).

  144. 144.

    Killian, J. K. et al. Succinate dehydrogenase mutation underlies global epigenomic divergence in gastrointestinal stromal tumor. Cancer Discov. 3, 648–657 (2013).

  145. 145.

    Sciacovelli, M. et al. Fumarate is an epigenetic modifier that elicits epithelial-to-mesenchymal transition. Nature 537, 544–547 (2016).

  146. 146.

    MacKenzie, E. D. et al. Cell-permeating alpha-ketoglutarate derivatives alleviate pseudohypoxia in succinate dehydrogenase-deficient cells. Mol. Cell. Biol. 27, 3282–3289 (2007).

  147. 147.

    Erez, A. & DeBerardinis, R. J. Metabolic dysregulation in monogenic disorders and cancer: finding method in madness. Nat. Rev. Cancer 15, 440–448 (2015).

  148. 148.

    Kranendijk, M., Struys, E. A., Salomons, G. S., Van der Knaap, M. S. & Jakobs, C. Progress in understanding 2-hydroxyglutaric acidurias. J. Inherit. Metab. Dis. 35, 571–587 (2012).

  149. 149.

    Ma, S. et al. L2hgdh Deficiency accumulates l-2-hydroxyglutarate with progressive leukoencephalopathy and neurodegeneration. Mol. Cell Biol. 37, e00492–16 (2017).

  150. 150.

    Rzem, R. et al. A mouse model of L-2-hydroxyglutaric aciduria, a disorder of metabolite repair. PLoS One 10, e0119540 (2015).

  151. 151.

    Kranendijk, M. et al. IDH2 mutations in patients with D-2-hydroxyglutaric aciduria. Science 330, 336 (2010).

  152. 152.

    Akbay, E. A. et al. D-2-hydroxyglutarate produced by mutant IDH2 causes cardiomyopathy and neurodegeneration in mice. Genes Dev. 28, 479–490 (2014).

  153. 153.

    Batlle, E. & Clevers, H. Cancer stem cells revisited. Nat. Med. 23, 1124–1134 (2017).

  154. 154.

    Snyder, V., Reed-Newman, T. C., Arnold, L., Thomas, S. M. & Anant, S. Cancer stem cell metabolism and potential therapeutic targets. Front. Oncol. 8, 203 (2018).

  155. 155.

    Zhou, Y. et al. Metabolic alterations in highly tumorigenic glioblastoma cells: preference for hypoxia and high dependency on glycolysis. J. Biol. Chem. 286, 32843–32853 (2011).

  156. 156.

    Chen, C. L. et al. NANOG metabolically reprograms tumor-initiating stem-like cells through tumorigenic changes in oxidative phosphorylation and fatty acid metabolism. Cell. Metab. 23, 206–219 (2016).

  157. 157.

    Li, Z. et al. Hypoxia-inducible factors regulate tumorigenic capacity of glioma stem cells. Cancer Cell 15, 501–513 (2009).

  158. 158.

    Dong, C. et al. Loss of FBP1 by Snail-mediated repression provides metabolic advantages in basal-like breast cancer. Cancer Cell 23, 316–331 (2013).

  159. 159.

    Wang, Y. H. et al. Cell-state-specific metabolic dependency in hematopoiesis and leukemogenesis. Cell 158, 1309–1323 (2014).

  160. 160.

    Saito, Y., Chapple, R. H., Lin, A., Kitano, A. & Nakada, D. AMPK protects leukemia-initiating cells in myeloid leukemias from metabolic stress in the bone marrow. Cell Stem Cell 17, 585–596 (2015).

  161. 161.

    Janiszewska, M. et al. Imp2 controls oxidative phosphorylation and is crucial for preserving glioblastoma cancer stem cells. Genes Dev. 26, 1926–1944 (2012).

  162. 162.

    Lagadinou, E. D. et al. BCL-2 inhibition targets oxidative phosphorylation and selectively eradicates quiescent human leukemia stem cells. Cell Stem Cell 12, 329–341 (2013).

  163. 163.

    Sancho, P. et al. MYC/PGC-1α balance determines the metabolic phenotype and plasticity of pancreatic cancer stem cells. Cell Metab. 22, 590–605 (2015).

  164. 164.

    Jones, C. L. et al. Inhibition of amino acid metabolism selectively targets human leukemia stem cells. Cancer Cell 34, 724–740.e724 (2018).

  165. 165.

    Vlashi, E. et al. Metabolic state of glioma stem cells and nontumorigenic cells. Proc. Natl Acad. Sci. USA 108, 16062–16067 (2011).

  166. 166.

    Kuntz, E. M. et al. Targeting mitochondrial oxidative phosphorylation eradicates therapy-resistant chronic myeloid leukemia stem cells. Nat. Med. 23, 1234–1240 (2017).

  167. 167.

    Ye, H. et al. Leukemic stem cells evade chemotherapy by metabolic adaptation to an adipose tissue niche. Cell Stem Cell 19, 23–37 (2016).

  168. 168.

    Pascual, G. et al. Targeting metastasis-initiating cells through the fatty acid receptor CD36. Nature 541, 41–45 (2017).

  169. 169.

    Samudio, I. et al. Pharmacologic inhibition of fatty acid oxidation sensitizes human leukemia cells to apoptosis induction. J. Clin. Invest. 120, 142–156 (2010).

  170. 170.

    Li, J. et al. Lipid desaturation is a metabolic marker and therapeutic target of ovarian cancer stem cells. Cell Stem Cell 20, 303–314.e305 (2017).

  171. 171.

    El Helou, R. et al. miR-600 acts as a bimodal switch that regulates breast cancer stem cell fate through WNT signaling. Cell Rep. 18, 2256–2268 (2017).

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Acknowledgements

We thank members of the laboratories of A.M.I. and L.W.S.F. for discussion and J. Schvartzman and S. Vardhana for critical reading of the manuscript. A.M.I. is supported by the NIH/NCI (K08 CA201483), Damon Runyon Cancer Research Foundation (CI 95-18), Burroughs Wellcome Fund (CAMS 1015584), Leukemia & Lymphoma Society (SCOR 7011-16), Susan & Peter Solomon Divisional Genomics Program, Steven A. Greenberg Fund and Cycle for Survival. L.W.S.F. is supported as a Dale F. Frey-William Raveis Charitable Fund Scientist by the Damon Runyon Cancer Research Foundation (DFS-23-17). This work was additionally supported by the Searle Scholars program (to L.W.S.F.), The Starr Foundation (I11-0039 to L.W.S.F.) and Memorial Sloan Kettering Cancer Center Support Grant P30 CA008748.

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A.M.I. and L.W.S.F. conceived the topic and wrote the manuscript.

Correspondence to Andrew M. Intlekofer or Lydia W. S. Finley.

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A.M.I. has previously consulted for Foundation Medicine, Inc.

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