Glucose-independent glutamine metabolism via TCA cycling for proliferation and survival in B-cells

Summary

Because MYC plays a causal role in many human cancers, including those with hypoxic and nutrient-poor tumor microenvironments, we have determined the metabolic responses of a MYC-inducible human Burkitt lymphoma model P493 cell line to aerobic and hypoxic conditions, and to glucose deprivation, using Stable Isotope Resolved Metabolomics. Using [U-¹³C]-glucose as the tracer, both glucose consumption and lactate production were increased by MYC expression and hypoxia. Using [U-¹³C,¹⁵N]-glutamine as the tracer, glutamine import and metabolism through the TCA cycle persisted under hypoxia, and glutamine contributed significantly to citrate carbons. Under glucose deprivation, glutamine-derived fumarate, malate, and citrate were significantly increased. Their ¹³C labeling patterns demonstrate an alternative energy-generating glutaminolysis pathway involving a glucose-independent TCA cycle. The essential role of glutamine metabolism in cell survival and proliferation under hypoxia and glucose deficiency, makes them susceptible to the glutaminase inhibitor BPTES, and hence could be targeted for cancer therapy.

Introduction

Mutations of genes involved in the tricarboxylic acid (TCA) cycle such as fumarate hydratase, succinate dehydrogenase or isocitrate dehydrogenase 1 or 2 are causally linked to familial cancer syndromes (Bensaad et al., 2006) or spontaneous low grade gliomas and acute myelogenous leukemia (Dang et al., 2010). Together with the well known Warburg effect (Koppenol et al., 2011; Vander Heiden et al., 2009; Warburg, 1956; Warburg et al., 1924) and numerous other alterations in the central metabolism of cancers (King et al., 2006; Samudio et al., 2009), these all point to the important role of metabolism in the development of many cancers and its therapeutic opportunities (Vander Heiden, 2011). Further, tumor suppressors such as p53 and oncogenes such as MYC and RAS have been directly linked to regulating metabolic pathways (Dang et al., 2009a; Telang, 2007) for initiating tumorigenesis and tumor progression. These genetic changes correlate with an increase in glucose consumption and lactate production when MYC is high. However, solid tumors contain regions that are both hypoxic and glucose depleted (Schroeder et al., 2005), requiring alternative strategies for survival and/or proliferation.

Here, we report stable isotope-resolved metabolomic (SIRM) studies of MYC-induced alterations in glucose and glutamine metabolism, in which we find persistent, MYC-dependent hypoxic metabolism of glutamine, even in the absence of glucose. Using NMR and MS, we have traced the fates of individual atoms from uniformly ¹³C-labeled glucose ([U-¹³C]-Glc) or ¹³C,¹⁵N-labeled glutamine ([U-¹³C,¹⁵N]-Gln) in human B-cell P493 cells carrying an inducible MYC vector. Overexpressed MYC resulted in the concurrent conversion of glucose to lactate and the oxidation of glutamine via the TCA cycle. Under hypoxic conditions with high MYC, a substantial fraction of the glucose consumed was converted to excreted lactate, and glutamine continued to be utilized by the TCA cycle, which was used for cell survival. We further document the finding of a fully ¹³C-labeled citrate isotopologue that contained carbons deriving completely from labeled glutamine, suggesting the existence of a glucose-independent TCA cycle. We also found under glucose-depleted culture conditions that a glutamine-dependent and glucose-independent TCA cycle may operate under both aerobic and hypoxic conditions. Moreover, we observed an enhanced conversion of glutamine to glutathione under hypoxia; glutathione is an important reducing agent for controlling the accumulation of mitochondrial reduced oxygen species (ROS). Under moderate hypoxia, excess ROS is generated at Complex II owing to a mismatch between NADH production and terminal oxidase activity (Wu et al., 2007). We therefore tested whether inhibition of glutamine metabolism could induce oxidative stress under hypoxia. We found that inhibition of glutaminase (GLS) by the glutaminase selective inhibitor BPTES (Robinson et al., 2007) elevated ROS levels and diminished ATP levels in hypoxic cells. In fact, we found that inhibition of glutaminase effectively kills hypoxic cancer cells in vitro and delays tumor xenograft growth in vivo.

Results

Co-existence of oxidative and aerobic glycolysis

Our genomic analysis of MYC target genes indicates that the expression of genes involved in glycolysis and in mitochondrial respiration is co-regulated by MYC (Dang, 2010; Kim et al., 2008; Kim et al., 2007; Li et al., 2005). We thus determined the metabolic consequences of MYC-activation in a model cell line (P493) of human Burkitt lymphoma grown in uniformly labeled [U-¹³C]-Glc under aerobic (21% O₂) or hypoxic (1% O₂) conditions. Although the levels of metabolites at steady state are the result of the balance between production and consumption, the use of SIRM enabled us to determine not only steady levels of metabolites, but also the isotopomer and isotopologue distributions of metabolites derived from ¹³C-labeled glucose for reconstruction of metabolic pathways. Time course isotopomer data on extracellular metabolites further provided flux measurement for substrate import and product release (Figures 1A, S2C, S3B and Table S1). P493 cells contain a tetracycline-repressible MYC construct, such that tetracycline withdrawal results in rapid induction of MYC, and tetracycline treatment results in MYC suppression (Figure S1A). Induction of MYC resulted in an increased of ¹³C-glucose consumption and ¹³C-lactate production, which were further accentuated by hypoxia (Figure 1A). NMR analysis of ¹³C-labeled metabolites derived from [U-¹³C]-Glc in cell extracts also corroborated the finding that overexpressed MYC resulted in lactic fermentation even under aerobic conditions (Figure 1B). The same cell extracts were further analyzed by GC-MS to quantify glucose-derived ¹³C isotopologues of lactate (lactate with different number of ¹³C atoms) (Figure 1D). Levels of the m+3 isotopologue of lactate (triply ¹³C labeled lactate or ¹³C₃-lactate) derived from glucose shown with MYC ON or OFF in aerobic (A) or hypoxic (H) conditions (Figure 1D), also support the ability of MYC to increase aerobic glycolysis (Figure 1C). Under hypoxic conditions, glucose-derived lactate was increased but was less dependent on MYC. The production of lactate (Table S1) accounts for only part of the glucose consumed. Although a complete carbon inventory has not been achieved, we estimate that a significant fraction of the glucose enters new biomass, which with respiration is associated with carbon loss as CO₂.

Figure 1. MYC induces aerobic glycolysis that is heightened in hypoxia.

A. Time course of glucose consumption and lactate secretion into the medium. P493 cells were treated with 0.1 μg/ml tetracycline (MYC OFF) or without tetracycline (MYC ON) for 48 hours in hypoxic (H) or aerobic (A) conditions and were grown in RPMI containing 10 mM [U-¹³C] glucose (¹³C-Glc₆) for 24 hours. Media metabolites and ¹³C enrichments were measured by 1D ¹H NMR. The metabolite amounts are expressed as μmole. Each time data point is an average of duplicate samples. The rates were calculated from linear regression of the time courses, and normalized to cell mass, as μmole/h/g cells. Open symbols are ¹³C lactate, filled symbols are ¹³C glucose. Filled red circles MYC ON aerobic; blue diamonds MYC OFF aerobic , green squares MYC ON hypoxic, black triangles MYC OFF hypoxic. B. 1-D ¹H{¹³C} HSQC NMR spectra of cell extracts of P493 MYC ON versus MYC OFF. Intracellular ¹³C-lactate synthesized from ¹³C glucose was attenuated when MYC OFF, which was also observed by GC-MS analysis of the same extracts (D). NMR spectra were recorded at 800 MHz and 20 °C. Each ¹H peak arose from protons directly attached to ¹³C and the peak assignment denotes the ¹³C-carbon. Thus, the peak intensity reflects ¹³C abundance of the attached carbon. C. Diagram of ¹³C labeling patterns of glycolytic products with ¹³C₆-Glc as tracer. Glucose with carbons (red circles) labeled at all 6 positions (blue) generates ¹³C₃-lactate via pyruvate (three carbons), which also produces ¹³C₂-acetyl CoA. D. GC-MS analysis of MYC and/or hypoxia effect on ¹³C-Glc₆ metabolism to different ¹³C isotopologues of lactate. Overexpressed MYC enhanced lactate production (most notably ¹³C₃-lactate; m+3) and hypoxia further increased labeled incorporation into lactate. Each value is an average of duplicate samples. The error bars represent standard error of the mean (SEM).

As shown in Figure 2, glucose-derived TCA cycle intermediates under aerobic condition displayed a dependence on MYC, such that the doubly ¹³C-labeled isotopologue of citrate (¹³C₂-citrate), succinate, fumarate and malate (m+2 forms, circled red), increased when MYC was ON. Glucose-derived α-ketoglutarate (m+2), which was at very low cellular concentration, also demonstrated a dependence on MYC; it was only detectable when MYC was ON. Hypoxia decreased the MYC-induced conversion of glucose to citrate (m+2) and to other m+2 isotopologues of TCA cycle intermediates (malate, fumarate, and succinate), but these activities were independent of MYC expression (MYC ON-H versus MYC OFF-H). In addition to the synthesis of ¹³C₂-citrate, there was a significant production of ¹³C₅-citrate (m+5, circled green) with MYC ON under aerobic condition. This citrate isotopologue could be produced from m+2 acetyl-CoA plus m+3 oxaloacetate (product of pyruvate carboxylation, green arrow and circles, Figure 2) (Fan et al., 2010), and its level appeared to be attenuated by hypoxia and the absence of ectopic MYC (Tet treatment). It is notable that a large fraction (up to 70%) of these TCA metabolites (m+0) were not derived from the labeled glucose, suggesting an alternative source and/or prolonged half-lives of these metabolites that could have existed prior to the administration of labeled glucose.

Figure 2. Glucose entry into the TCA cycle is induced by MYC and suppressed by hypoxia.

The cycle reactions are depicted without or with pyruvate carboxylation (green arrow) and the ¹³C isotopomer patterns are the result of one cycle turn. The isotopologue distributions were determined by GC-MS. The incorporation of ¹³C atoms from ¹³C₆-Glc into citrate, α-ketoglutarate (α-KG), succinate, fumarate, and malate are denoted as m+n, where n is the number of ¹³C atoms. The m+5 (¹³C₅)-citrate (green circle) is produced by condensation of m+2 acetyl CoA with m+3 OAA (from pyruvate carboxylation), while m+2 (¹³C₂)-citrate is synthesized without input of pyruvate carboxylation. Red and green circles: ¹³C atoms derived from ¹³C₆-Glc without or with pyruvate carboxylation, respectively. GPT2, glutamate-pyruvate transaminase; PC, pyruvate carboxylase; CO₂, indicates where carbon dioxide is released. A= Aerobic, H=Hypoxic. The error bars represent standard error of the mean (SEM).

Persistence of glutamine oxidation via the TCA cycle under hypoxia

The attenuation of glucose entry into the TCA cycle under hypoxia (Figure 2) is consistent with the hypoxia inducible factor (HIF)-mediated diversion of pyruvate to lactate (away from acetyl-CoA) through the induction of LDHA (which increases the relative flux from pyruvate to lactate) and PDK1 (which decreases the relative flux from pyruvate to acetyl-CoA) (Kim et al., 2006). Because it was previously documented that MYC induces glutamine metabolism under aerobic conditions (Gao et al., 2009; Wise et al., 2008a), we sought to determine whether glutamine entry into the TCA cycle would also be compromised by hypoxia.

Using [U-¹³C,¹⁵N]-Gln (¹³C₅¹⁵N₂-Gln) as the tracer with SIRM analysis, the fates of glutamine as a function of MYC induction and oxygen availability were determined (Figures 3B-D). Glutamine is transported into cells by transporters, such as the direct targets of MYC SLC1A5 or ASCT2 (Figure S1C), and then converted to glutamate by glutaminase (GLS, kidney isoform, which is also a target of MYC) (Figure S1D). ¹³C₅¹⁵N₂-Gln (m+7) is converted by glutaminase into ¹³C₅¹⁵N-Glu (m+6) plus ¹⁵NH ⁺₄ (Figure 3A). NMR studies of biological replicate experiments revealed a MYC-dependent conversion of labeled glutamine to glutamate, which unexpectedly persisted in hypoxia (Figure 3B). This result was corroborated by the GC-MS analysis of the same set of polar extracts. Intracellular glutamine was converted to glutamate (m+6 isotopologue or ¹³C₅¹⁵N₁-Glu, Figure 3D) in a MYC-dependent fashion that persisted under hypoxia. A large fraction of the m+5 glutamate isotopologue was also present and displayed a similar MYC- and hypoxia-dependence as the m+6 isotopologue. The m+5 isotopologue of glutamate was largely ¹³C₅-Glu as determined by high-resolution FT-ICR-MS (Figure S1E), which resolved the neutron mass from ¹³C and ¹⁵N. ¹³C₅-Glu should be a transamination product of ¹³C-labeled glutamine-derived α-ketoglutarate (α-KG) with unlabeled nitrogen sources (Figure 3A). α-ketoglutarate levels also tracked MYC expression (Figure 4). Furthermore, the dependence of glutaminase activity on MYC expression measured in extracts suggests that the intracellular conversion of labeled glutamine to glutamate is at least partly regulated by GLS1 activity in response to MYC (Figure S2A). This was further supported by the higher level of ammonium ions - the other product of the glutaminase reaction - that were released into the medium under MYC ON conditions (Figure S2B).

Figure 3. MYC induces glutamine to glutamate conversion that persists in hypoxia.

A. Diagram of ¹³C labeling patterns products with ¹³C₅,¹⁵N₂-Gln as tracer. Glutamine is labeled at all five carbon and both nitrogen atoms. Glutaminase (GLS) activity produces glutamate ¹³C¹⁵N₁ with the loss of the amido nitrogen as ammonia. Red and green circles: respective ¹³C and ¹⁵N labeling of glutamate from the glutaminase reaction; light blue circles: ¹³C labeling of glutamate from the transamination of α-ketoglutarate; black: unlabeled N; GS: glutamine synthetase. B. 1-D ¹H{¹³C} HSQC NMR spectra of cell extracts of P493 MYC ON versus MYC OFF. Intracellular ¹³C-4-glutamate production from glutamine was reduced when MYC OFF in aerobic condition. The conversion of glutamine to glutamate in hypoxia was as high as aerobic condition regardless of MYC expression. NMR spectra were recorded at 800 MHz and 20°C. Each ¹H peak arose from protons directly attached to ¹³C and the peak assignment denotes the ¹³C-carbon. The peak intensity reflects ¹³C abundance of the attached carbon. C. Glutamine levels in the media. P493 cells were treated with 0.1 μg/ml tetracycline (MYC OFF) or without tetracycline (MYC ON) for 48 hours in hypoxic (H) or aerobic (A) conditions and were grown in ¹³C₅,¹⁵N₂-Gln (m+7) medium in presence or absence of tetracycline for 24 hours. Media metabolites were analyzed by GC-MS. Under aerobic condition, glutamine consumption was decreased when MYC OFF. Under hypoxia, glutamine consumption was decreased. The metabolite concentration was expressed as μmole/g protein dry weight. D. GC-MS analysis of intracellular conversion of ¹³C₅,¹⁵N₂-glutamine to glutamate. The m+5 glutamate was primarily ¹³C₅-glutamate (cf. Figure S1E) with no ¹⁵N label, which is derived from ¹³C₅-αKG via transamination. The data shown at B, C and D were performed three times. The error bars represent standard error of the mean (SEM).

Figure 4. P493 cells were grown in ¹³C₅,¹⁵N₂-glutamine medium with glucose. MYC induces glutamine entry into the TCA cycle, which persists in hypoxia.

¹³C₅,¹⁵N₂ glutamine enters the Krebs cycle to produce α-ketoglutarate, succinate, fumarate and malate. Citrate can be generated by reductive carboxylation (RedCarb) of glutamine-derived α-ketoglutarate (m+5) (orange circles) or via the forward reactions of the cycle. Malic enzyme (ME) catalyzes the oxidative decarboxylation of malate to pyruvate (light blue). This pyruvate pool (m+3) can be precursor for both m+2 acetyl CoA and m+3 OAA (from pyruvate carboxylation, green circles) to make m+5 citrate. The citrate m+6 isotopologue could only arise from glutamine-derived m+4 oxaloacetate (from forward cycle reactions, red circles) and m+2 acetyl-CoA (mediated by malic enzyme). The isotopologue distributions were determined by GC-MS. Each value is an average of duplicate samples. The data shown were performed three times. GPT2, glutamate-pyruvate transaminase; ME; malic enzyme; PC, pyruvate carboxylase; GLS, glutaminase; CO₂, indicates where carbon dioxide is released. A= Aerobic, H=Hypoxic. The error bars represent standard error of the mean (SEM).

Levels of fully labeled glutamine in the media were measured to determine the rate of consumption of glutamine. Glutamine consumption rates were in the order: MYC ON aerobic ≈ MYC ON hypoxic > MYC OFF hypoxic > MYC OFF aerobic. (Figure 3C, Figure S2C). The m+5 and m+6 isotopologues of glutamate were also present in the medium (Figure 3C), which reflect exchange of intracellular glutamine-derived glutamate for other amino acids such as cystine (see below).

As depicted in Figure 3A, labeled glutamine catabolism by glutaminase led to the production of ¹³C₅-αKG, which can enter the TCA cycle for further oxidation. As shown in Figure 4, the synthesis of ¹³C₄-succinate, -fumarate, and -malate (m+4) is consistent with the oxidation of ¹³C₅-αKG via the forward reactions of the TCA cycle (red circles, Figure S2D). These labeled TCA intermediates all responded to MYC status by increasing 60 to >100% when MYC was ON, regardless of O₂ availability (Figure 4). The levels of ¹³C₄-citrate, which is synthesized in the 2^nd turn from ¹³C₄-oxaloacetate (OAA, derived from labeled glutamine in the 1^st turn) and acetyl-CoA (from unlabeled glucose or other unlabeled sources) by citrate synthase (CS), also responded to MYC expression under both aerobic and hypoxic conditions (Figure 4). These results show that MYC can drive glutamine metabolism around the TCA cycle even under hypoxia.

Glucose-independent oxidation of glutamine for survival and proliferation

In addition to the production of the ¹³C₄-citrate isotopologue, there was a significant formation of other citrate isotopologues, e.g. ¹³C₃- (m+3), ¹³C₅- (m+5), and ¹³C₆-citrate (m+6) (Figure 4). These labeled species indicate that the production of labeled acetyl-CoA and OAA isotopologues from the glutamine tracer are using pathways external to the TCA cycle. Figure S2D depicts the pathways that can lead to the synthesis of ¹³C₃-, ¹³C₅-, and ¹³C₆-citrate. These include the cytoplasmic ATP-citrate lyase (ACL) plus malic enzyme (ME) reactions, which produce respectively ¹³C₄/¹³C₂-OAA and ¹³C₃/¹³C₂-pyruvate; the latter yields ¹³C₂/¹³C₁-acetyl CoA via pyruvate dehydrogenase (PDH). Condensation of the labeled OAA and acetyl CoA species derived from the ACL-ME pathway, glycolysis and the TCA cycle produces ¹³C₃ to ¹³C₆-citrate (light blue circles, Figure S2D). In particular, the presence of the ¹³C₆-citrate isotopologue unambiguously confirmed the production of fully labeled acetyl CoA from the glutamine tracer via the ACL-ME-PDH pathway. Labeled acetyl CoA production from the ACL reaction is further supported by ¹³C label incorporation into lipids such as triacylglycerides (TAG) and phosphatidylcholines (PC) (Figure S2E).

The ¹³C₃- and ¹³C₅-citrate isotopologues can also be formed from the ACL-ME plus pyruvate carboxylase (PC) reactions (green circles, Figure S2D). Pyruvate carboxylase is active in P493 cells, as evidenced from the labeled glucose tracer experiment described above (Figure 2). The operation of the ACL-ME1-PDH and ACL-ME1-PC pathways is further corroborated by the production of ¹³C₃-succinate, - fumarate, and -malate, which cannot be formed from labeled glutamine via the TCA cycle activity alone. Finally, the production of ¹³C₅-citrate from the glutamine tracer can also be explained by reductive carboxylation of αKG to form citrate (orange circles, Figure S2D), a reversal of the citrate to αKG reaction catalyzed by aconitase and IDH as recently reported in other cells (Metallo et al., 2011; Mullen et al., 2011; Wise et al., 2011; Yoo et al., 2008) and driven by the hydrolysis of ATP via citrate lyase and ACC. There is abundant CO₂/HCO₃⁻ in cell culture for reductive carboxylation and presumably in tissue from a number of decarboxylation reactions. However, the relative proportion of the m+5 versus m+3, m+4 and m+6 species which are characteristic of the forward reactions in the Krebs cycle plus pyruvate carboxylase activity, indicates that reductive carboxylation is not the major pathway in P493 cells, especially under aerobic conditions where the ratio of αKG to citrate concentration was very low (Figure 4); a high ratio is important for driving this thermodynamically uphill reaction.

We were also intrigued by the apparent up-regulation of such TCA cycle-mediated glutamine metabolism by MYC and its persistence under hypoxia (Figure 4). Such a glucose-independent TCA cycle activity would be advantageous for cancer cells subjected to glucose deficiency and/or hypoxia in the tumor microenvironment. Thus, we next determined whether the glutamine-mediated TCA cycle can operate in the absence of glucose and whether glutamine metabolism alone can sustain cell growth and survival.

P493 cells were grown in the absence of glucose using the tracer [U-¹³C,¹⁵N]-Gln to determine whether TCA cycle intermediates could be derived solely from glutamine (Figure 5). In the absence of glucose, the P493 cells completed one doubling in three days under aerobic condition with MYC ON compared with doubling every 34±2 h in the presence of glucose (Figure S3A). The cells consumed glutamine (Figure S3B) to produce ¹³C₅-αKG (m+5) in a MYC-dependent fashion (circled red, Figure 5). The continued functioning of the TCA cycle under glucose-deprived conditions was identified by the production of various isotopologues of fumarate, malate, and aspartate, particularly the ¹³C₄-isotopologues (m+4, Figure 5). However, these labeled isotopologues accumulated to much higher levels (> 100 fold for ¹³C₄-Asp) than under glucose-replete conditions (compare Figures 4, 5 or Figures 1B, S3C). This could result from a lower supply of acetyl CoA under glucose-deprived conditions such that excess glutamine-derived OAA was transaminated to form aspartate. This is also consistent with the lower levels of labeled citrate and αKG isotopologues in glucose-deprived cells than in glucose-replete cells (Figures 4 and 5). In the absence of glucose, it is also notable that ¹³C incorporation from the glutamine tracer into all TCA cycle intermediates decreased under hypoxia but increased with MYC OFF (Figure 5). Furthermore, the ¹³C alanine isotopologues (e.g. ¹³C₃-Ala or m+3 in Figure 5) showed the opposite behavior in response to MYC expression, regardless of the O₂ conditions. The significant buildup of labeled alanine was in contrast to a small production of lactate from glutamine (Figure S3C), which again argues against the operation of the canonical glutamine to lactate pathway in P493 cells.

Figure 5. Glutamine-driven glucose-independent TCA cycle. P493 cells were grown in ¹³C₅,¹⁵N₂-glutamine medium without glucose.

The isotopologue distributions were determined by GC-MS. Under glucose-deprived conditions, the m+5 α-ketoglutarate from labeled glutamine was produced in a MYC-dependent fashion. Labeled glutamine was also incorporated into m+4 isotopologues of fumarate and malate downstream of α-ketoglutarate. These isotopologues decreased under hypoxia but not with decreased MYC. The m+3 alanine isotopologue, which is derived from the transamination of glutamine-derived m+3 pyruvate (via malic enzyme), was highly elevated under high MYC and aerobic conditions. Each value is an average of duplicate samples. Colored circles denote the same sets of reactions as in Figure 4. GPT2, glutamate-pyruvate transaminase; ME, malic enzyme; PC, pyruvate carboxylase; GLS, glutaminase; CO₂, indicates where carbon dioxide is released. A= Aerobic, H=Hypoxic. The error bars represent standard error of the mean (SEM).

The relatively low accumulation of labeled TCA metabolites with MYC ON under glucose deprivation could be caused by a combination of limited TCA cycling and the demands for cell proliferation and cell maintenance. The same argument could also apply to aerobic versus hypoxic conditions. In Figure S3A, the highest proliferation rate was observed under aerobic conditions with MYC ON, followed by MYC OFF under aerobic conditions, whereas under hypoxia there was little proliferation regardless of the MYC status. With a slowdown of TCA cycling due to glucose deficiency, a higher consumption rate of labeled TCA intermediates for cell proliferation could deplete these metabolites for both MYC ON and MYC OFF, but more so for MYC ON than MYC OFF and with more depletion under aerobic than hypoxic condition (Figure 5). Diversion of glutamine for maintenance purposes, e.g. glutathione synthesis for ROS detoxification (inferred by increased ROS production with BPTES inhibition of glutaminolysis, Figure S4A) could also lead to less flux through the TCA cycle. This is consistent with a higher buildup of glutamine-derived glutathione with aerobic and MYC ON conditions (Figure S3C).

When TCA cycling is faster, as in glucose-replete cells, the production rate for the labeled TCA intermediates may be higher than their consumption rate, leading to a higher buildup of these labeled metabolites when MYC is over-expressed (Figure 4). The bottleneck in TCA cycling could also contribute to the significant buildup of ¹³C₃-Ala (m+3) alanine under MYC ON and glucose deprivation (Figure 5) via excess production of glutamine-derived pyruvate (by way of the ACL-ME1 pathway, Figure S4), which is transaminated to form alanine via glutamic pyruvic transaminase 1 or 2. Again, this did not occur under glucose-replete conditions, where glucose, not glutamine, was the main source of alanine production (Figure 2). In the absence of glucose and under hypoxia, P493 cells did not proliferate, but continued to consume glutamine, and remained viable (Figure S3A).

Under aerobic conditions with MYC ON, the percentage of viable (78% versus 76%) and proliferating (R2, 30.6% versus 30.3%) cell populations were similar between glucose-replete (Figure S6) and deplete conditions (Figure S7). This can be mediated by the ability of glutamine metabolism to alleviate oxidative stress (Figure S4A) and by support cell bioenergetics (Figure S4B). These data show that when driven by MYC, glutamine plays a crucial role for both cell survival and proliferation under glucose-deprivation.

Thus, the above observations confirm a re-programmed glutamine-dependent TCA cycle that functions in the absence of glucose. Figure S4C outlines the pathways via which ¹³C carbons of glutamine are converted to labeled acetyl CoA and re-enters the TCA cycle. The significant presence of ¹³C₅- (m+5) and ¹³C₆-citrate (m+6, Figure 5) is consistent with an ACL-ME-mediated production of acetyl-CoA and citrate solely from glutamine. The ¹³C labeling of lipid acyl chains, although at relatively low levels, (Figure S4D) and the increased labeling under aerobic with MYC ON confirm the activity of ACL, and is consistent with the higher rate of proliferation under these conditions (Figure S3A). However, since there was a significant level of unlabeled and ¹³C₄-citrate (m+4, Figure 5) under glucose deprivation, there must also be source(s) of unlabeled acetyl CoA that contribute to the continued operation of the TCA cycle. We have observed that fatty acid oxidation, a possible source for unlabeled acetyl-CoA, increased in the absence of ectopic MYC in P493 cells (Figure S5B). This could account for the higher levels of ¹³C₄-citrate when MYC was off (Figure 5). However, it cannot explain the comparable levels of unlabeled citrate (m+0 Figure 5) between aerobic MYC ON and MYC OFF. Therefore other sources of unlabeled acetyl CoA, such as oxidation of amino acids may also exist.

Glutamine-dependent bioenergetics and redox homeostasis, a cell survival pathway in hypoxia

The dependence of P493 cells on glutamine for proliferation and maintenance under aerobic and hypoxic conditions suggests a key role for glutamine in driving anaplerotic and bioenergetic needs of both dividing and resting cells. We found that the interruption of glutamine metabolism with the glutaminase inhibitor BPTES decreased ATP levels under aerobic conditions. In hypoxia, cells maintained a lower ATP level that was further diminished by BPTES treatment (Figure 7B). These results suggest that glutamine metabolism via the glucose-independent TCA cycle supports cellular bioenergetics for cell survival (and proliferation) under both aerobic and hypoxic conditions.

Figure 7.

A. Effect of glutaminase inhibitor, BPTES, on aerobic and hypoxic P493 cell growth. Growth of control cells compared with cells treated with BPTES under both aerobic and hypoxic conditions. All cells were grown at 1 × 10⁵ cells/mL. Cell counts were performed in triplicate and shown as mean ± SD. Live cells were counted daily in a hemocytometer using trypan blue dye exclusion. B. Effect of BPTES on steady state ATP levels. P493 cells were treated with 2 μM BPTES for 20 h and counted. ATP levels (mean ± SD, n = 3 experiments) were determined by luciferin– luciferase-based assay on aliquots containing an equal number of live cells. *p = 0.0006; **p = 0.01 (t-test).C. Effects of hypoxia and glucose deprivation on oxygen consumption rates. Oxygen consumption rates of aerobic (A) or hypoxic (H) P493 cells were determined by a Clark-type oxygen electrode after culture in the presence or absence of glucose for 24 hours. Data are from duplicate experiments with standard deviation and p-values (T-test) shown. D. In vivo efficacy of BPTES. 2.0 × 10⁷ P493 human lymphoma B cells were injected subcutaneously into tumor-bearing SCID mice. When the tumor volume reached 100 mm³, 200 μg BPTES was injected every other day by i.p. administration (12.5 mg/kg bodyweight) for 20 days. Control animals were treated with daily i.p. injection of vehicle (2% (vol/vol) DMSO). BPTES inhibited lymphoma xenograft growth as compared to control. The tumor volumes were measured using digital calipers every 4 days and calculated using the following formula: [length (mm) × width (mm) × width (mm) × 0.52]. The results represent the average ± SEM (n=7 each).

In addition, both dividing and resting cells require redox homeostasis, not only for continuing glycolysis but also for detoxifying ROS. Glutamine, when converted to glutamate, could also be involved in the import of cystine through the anti-porter SLC7A11 (Figure 6A). This is consistent with the significant excretion of labeled glutamate into the culture medium (Figure 3C). Cystine, when converted to cysteine, contributes to the synthesis of glutathione together with glycine and glutamate derived from glutamine. We thus determined the contributions of labeled glutamine and glucose to total de novo glutathione synthesis by quantifying ¹³C labeled reduced plus oxidized glutathione levels derived from respective glucose or glutamine tracers (Figure 6B; *GSH-*GSSG). Relative to glucose, glutamine carbons were more readily incorporated into the glutamate moiety of the glutathione, indicating that glutamine, not glucose, is the main precursor for glutathione synthesis. While the glucose carbon contribution to glutathione diminished in hypoxia, the contribution from glutamine carbons persisted. Furthermore, the ¹³C fractional distribution in oxidized glutathione (GSSG) isotopologues (e.g. m+5 and m+10 in Figure 6C) was in fact higher under hypoxia with MYC ON than under aerobic with MYC OFF, which could reflect a higher ROS production or oxidation of de novo synthesized glutathione in response to MYC expression and hypoxia. This observation is supported by a lower level of reduced glutathione (GSH) (Figure S5D) and higher ROS production (Figure 6D) in P493 cells under hypoxia and inhibition of glutaminase by BPTES. The order of GSH levels were aerobic control > hypoxic control > aerobic BPTES > > hypoxic BPTES while that of ROS production (measured by DCFDA fluorescence) was reversed, which is expected for removal of ROS by oxidation of GSH to GSSG.

Figure 6. Glutamine contributes to glutathione synthesis and redox homeostasis.

A. Diagram of glutamine metabolism to glutathione. Glutamine, when converted to glutamate, is involved in the import of cystine through the antiporter SLC7A11. Cystine is converted to cysteine for the synthesis of glutathione, which also requires glycine and glutamine-derived glutamate. B. 1-D ¹H{¹³C} HSQC NMR analysis of ¹³C-reduced (*GSH) plus oxidized (*GSSG) glutathiones indicates that the fraction of glutamine contributing to glutathione production was sustained under hypoxia (H) as compared to aerobic condition (A). ¹³C incorporation was estimated from the ¹³C-4 peak of the glutamate residue in GSH+GSSG by 1D HSQC NMR. C. FT-ICR-MS determination of glutathione oxidation. The ¹³C₅ and ¹³C₁₀ isotopologues of GSSG were most likely derived from the ¹³C₅,¹⁵N₂-Glutamine tracer via respective incorporation of one and two units of ¹³C₅-Glu. D. Glutamine-dependent redox homeostasis. Flow cytometric histograms of ROS levels were determined by DCFDA fluorescence in P493 cells treated with 2, 10, 20 μM BPTES for 24 and 48 hours. E. Mean fluorescence intensity (MFI). MFI for DCFDA fluorescence is shown for the different concentrations of BPTES under aerobic and hypoxic conditions. MFI was determined from triplicate experiments with standard deviation and p-values (T-test) shown.

To determine further the role of glutaminase in redox homeostasis, we isolated T cells from mice that are heterozygous for mitochondrial glutaminase 1 (Gls(+/-)) (Masson et al., 2006), which is the homolog of human kidney type glutaminase rather than liver-specific glutaminase 2. When compared to aerobic and hypoxic wild-type (WT) T cells, the Gls(+/-) T cells had higher basal ROS levels which were further increased under hypoxia (Figure S5E). Altogether, these data suggest that MYC expression can enhance aerobic glutathione biosynthesis from glutamine to maintain redox homeostasis. Glutamine-derived glutathione production was also sustained under hypoxia to cope with heightened ROS production from the perturbed mitochondrial electron transport chain (Chandel et al., 1998; Guzy et al., 2005).

Our findings that MYC-induced glutamine metabolism persisted in hypoxia and in the absence of glucose for cell maintenance led us to test whether targeting glutaminase is feasible for cancer therapy. We determined the consequences of BPTES treatment on aerobic and hypoxic P493 cell proliferation (Figure 7A). While BPTES decreased glutaminase activity (Figure S5C) and the proliferation of aerobic P493 cells, inhibition of glutaminase killed hypoxic P493 cells (Figure 7A). Under hypoxia, the cells proliferated less but continued to import and metabolize glutamine (Figures S2C and 4) as well as survive (Figure S6), even under glucose-deprived conditions (Figure S7). The killing effect of BPTES under hypoxia can be ascribed to the crucial role of glutamine metabolism for survival by supporting cell bioenergetics (Figure 7B) and alleviating oxidative stress (Figure 6). This notion is consistent with the persistent, although diminished, rate of oxygen consumption by hypoxic cells as compared with aerobic cells (Figure 7C). Glucose deprivation increases the rate of oxygen consumption (Figure 7C) as previously reported (Gao et al., 2009). To determine the in vivo significance of our findings, P493 tumor xenograft bearing mice were treated with intraperitoneal injections of BPTES (Figure 7D). As compared with DMSO vehicle treated mice, the BPTES-treated mice demonstrated a significantly diminished tumor progression.

Discussion

Although increased glucose metabolism has been hypothesized to drive the bioenergetic needs of cancer cells, it is clear that highly proliferative cancer cells require additional supplies of biosynthetic precursors not met by glucose metabolism; thus there is the requirement of anaplerosis via glutaminolysis and/or pyruvate carboxylation as well as other sources such as fatty acids (DeBerardinis et al., 2007; Deberardinis et al., 2008; Fan et al., 2009; Gao et al., 2009; Wise et al., 2008b). A significant proportion of the biosynthetic needs under aerobic conditions may be met by glutamine metabolism, which is regulated by the activity of the MYC oncogene (Gao et al., 2009; Wise et al., 2008a). In this regard, our previous studies documented the ability of Myc to induce mitochondrial biogenesis, function, and enhanced oxygen consumption (Li et al., 2005), which persisted under hypoxia (Figure 7C). Our data shows that the import rates of both glucose and glutamine greatly exceed that of other amino acids under these various experimental conditions, suggesting that they are the major sources for energy and anaplerosis. However, it is a long-standing question how cancer cells survive or even proliferate under the hypoxic and nutrient deprived conditions encountered in the tumor microenvironment (Schroeder et al., 2005).

Solid tumors are often poorly vascularized, leading to regions of hypoxia and glucose deficiency (Goel et al., 2011; Schroeder et al., 2005), which must be overcome for cancer cells to continue proliferation and survive. Although Myc-overexpressing cells appear to become addicted to glutamine, the heightened addiction to glutamine under hypoxia had not been previously established (Yuneva et al., 2007). We therefore determined how limited oxygen availability would influence glutamine metabolism in vitro to gain insights into what might happen in vivo. It should be noted, however, that our well-controlled cell line studied in vitro may have adapted epigenetically and metabolically to prolonged cultures in nutrient-rich conditions, and hence metabolic changes under our in vitro experimental conditions may be instructive, but may not fully represent what happens in vivo in the tumor microenvironment. Notwithstanding this caveat, we used uniformly labeled [U-¹³C,¹⁵N]-Gln and traced the fate of glutamine carbon and nitrogen atoms in both aerobic and hypoxic conditions with MYC turned OFF or ON in a human B cell model of Burkitt lymphoma (P493). The P493 system is strictly dependent on Myc function for cell proliferation; as such, this experimental model does not conclusively allow us to distinguish the metabolic consequences of Myc-mediated transcriptional changes versus those that occur due to changes in cell size increase or proliferative status as previously reported (Schuhmacher et al., 1999). We surmise that most of the metabolic changes between MYC ON versus MYC OFF states are consequential to MYC regulation of genes involved in both glucose and glutamine metabolism as previously documented (Dang et al., 2009a). The entry of glutamine, after its conversion to glutamate and then to α-ketoglutarate, into the TCA and oxidation to succinate, fumarate, and malate were highly facilitated by MYC expression. More intriguingly, glutamine metabolism via the TCA cycle persisted under hypoxia and remained responsive to MYC. Based on detailed analyses of ¹³C isotopologues of citrate, succinate, fumarate, and malate (Figure 4), we uncovered a glucose-independent TCA cycle solely supported by glutamine, which involved the supply of acetyl CoA and OAA to the TCA cycle by the combined activity of ATP citrate lyase, malic enzyme, pyruvate dehydrogenase, and/or pyruvate carboxylase (Figure 4). This pathway was operative even when glucose was abundant and remained active under hypoxia. These data collectively define a more efficient pathway of glutaminolysis, in which glutamine can be used either for anabolic purposes for proliferation, or for producing energy. This pathway can produce up to 17.5 mol ATP/mol glutamine oxidized to CO₂. This is in contrast to the canonical glutaminolysis pathway from glutamine to lactate (Helmlinger et al., 2002; Newsholme et al., 1985a, b), where the glutamine carbons are not oxidized via the TCA cycle, and produce only 5 ATP/mol glutamine converted to lactate + CO₂.

To further explore the role of the glucose-independent TCA cycle, we traced glutamine metabolism under glucose-deprived culture conditions. SIRM analysis of TCA cycle metabolites revealed a relative increase in this glutamine-mediated pathway in glucose-deprived conditions, as evidenced by the increase in the ratios of ¹³C₆-citrate and ¹³C₅-citrate to ¹³C₄-citrate (Figure 5). The metabolites produced through this glutamine pathway provided the anabolic precursors (citrate) for lipid synthesis (Figure S4D) and cell survival precursor (Glu) for glutathione production (Figures 6B, S3C), which was commensurate with the aerobic growth demand (Figure S3A) and the need for ROS quenching under hypoxia (Figure 6). These observations are consistent with our previous findings that knock-down of glutaminase by siRNA resulted in increased ROS and slowed growth that could be partially rescued by N-acetylcysteine (Gao et al., 2009). It is also notable that pyruvate carboxylase plays a significant role in this glutamine pathway (Figures 4 and 5) and that glutamine-independent tumor cells use pyruvate carboxylase to drive a glucose-dependent TCA cycle in the absence of glutamine (Cheng et al., 2011). Thus, the flexibility of the TCA cycle to use either or both anaplerotic pathways may be important for tumor adaptation. More practically, therapeutic approaches targeting metabolism must consider these adaptive strategies.

As a proof-of-concept, we tested the effect of a glutaminase inhibitor (BPTES) on growth and ROS production of P493 cells and showed that blocking glutamine metabolism not only inhibited tumor cell growth under aerobic condition but also led to cell death under hypoxia and reduction of tumor xenograft growth in vivo (Figures 7A and D). The specificity of BPTES was previously documented by our studies in glioma cells, which revealed BPTES-induced metabolic changes, such as diminished glutamate, α-ketoglutarate, succinate, fumarate, and malate levels, consistent with an on-target effect on glutaminase (Seltzer et al., 2010). Further, the uncompetitive nature of BPTES inhibitory activity was recently underscored by the crystal structure of glutaminase (GAC form) co-crystallized with BPTES, which sits at the oligomerization interface of the glutaminase tetramer (Delabarre et al., 2011). Our finding of a glutamine-driven TCA cycle and other recent discoveries of cancer-related metabolic pathways (Dang et al., 2009b; Frezza et al., 2011; Possemato et al., 2011) suggest that metabolic flexibility may be a common feature of tumor cell metabolism. Hence, a broader and deeper understanding of cancer cell metabolism and their ability to reprogram canonical biochemical pathways under metabolic stress can be a rich ground for uncovering strategies for therapeutic targeting of tumors.

Experimental Procedures

Detailed materials and methods are available online in Supplemental Information.

¹³C labeled glutamine or glucose and hypoxic exposure

Briefly, Human lymphoma B P493 cells were grown in [U-¹³C]-labeled glucose or [U-¹³C,¹⁵N]-labeled glutamine for 24 hours. Non hypoxic cells (21% (vol/vol) O₂) were maintained at 37 °C in a 5% (vol/vol) CO₂ and 95% (vol/vol) air incubator. Hypoxic cells (1% O₂) were maintained in a controlled atmosphere chamber (PLAS-LABS) with a gas mixture containing 1% O₂, 5% (vol/vol) CO₂, and 94% (vol/vol) N₂ at 37 °C for 48 hours. Metabolites were extracted from cells and media as previously described (Fan et al., 2011).

Bromobimane and carboxy-H2DCFDA were used to measure reduced glutathione ROS levels, respectively, using flow cytometry

ATP levels were determined by luciferin–luciferase-based assay (Promega).

Glutaminase activity

Cells treated with BPTES and/or tetracycline were then incubated with [³H] glutamine to assess glutaminase activity.

The beta oxidation of ¹⁴C labeled oleic acid was used to assess fatty acid oxidation

Cell viability assay

The presence of dead cells was determined by propidium iodide staining at 2μg/ml using flow cytometry.

Statistical Analysis

Values are shown as mean ± standard deviation (SD) or standard error of the mean (SEM). Data were analyzed using the Student t-test and significance was defined as p < 0.05.

Highlights.

• P493 cells use glutamine for survival and proliferation in hypoxia

• P493 cells display a glucose-independent glutamine-driven TCA cycle

• Glutaminase inhibition diminishes in vitro and in vivo tumor cell growth