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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.
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.
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.
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.
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).
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 2nd turn from 13C4-oxaloacetate (OAA, derived from labeled glutamine in the 1st 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.
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, ,55 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 and5).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.
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.
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.
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.
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.
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 and5)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.
Detailed materials and methods are available online in Supplemental Information.
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).
ATP levels were determined by luciferin–luciferase-based assay (Promega).
Cells treated with BPTES and/or tetracycline were then incubated with [3H] glutamine to assess glutaminase activity.
The presence of dead cells was determined by propidium iodide staining at 2μg/ml using flow cytometry.
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.
This work was supported by in part by grant numbers 1R21NS074151-01 (to TT); The Sol Goldman Pancreatic Cancer Research fund (to AL); 5R01CA051497-21, 5R01CA057341-20, Leukemia Lymphoma Society grant LLS-6363-11, Stand-Up-to-Cancer/American Association for Cancer Research translational grant (to CVD); 1R01CA118434-01, 3R01CA118434-02, University of Louisville CTSPGP grant numbers 20044 (to TWMF) and 20061 (to ANL), NIH NCRR Grant 5P20RR018733, Kentucky Challenge for Excellence, the Brown Foundation, and Kentucky Lung Cancer Research Program (postdoctoral fellowship to PL). The FT-ICR-MS instrumentation was supported by NSF/EPSCoR grant number EPS-0447479. We thank Ramani Dinavahi, Julie Tan and Radhika Burra for excellent technical assistance and Stephen Rayport for the heterozygous glutaminase mice.
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SUPPLEMENTAL INFORMATION Supplemental Information includes Additional Experimental Procedures, 7 Supplemental figures and their legends and 1 table.
Author Contributions: AL, ANL, TWMF and CVD: designed research and wrote the manuscript. AL, ANL, MH, SB, JB, CJR, HZ, LJZ, RJCS, PL, RMH, and TWMF: performed research; all authors interpreted the data and commented on the manuscript.
The authors disclose no financial conflict of interest. There is no sponsored research or technology licensing activities involving the company.