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Blood test uses DNA strands of dying cells

Curators:  Larry H. Bernstein, MD, FCAP and Aviva Lev-Ari, PhD, RN

LPBI

Hadassah-Developed Blood Test Detects Multiple Sclerosis, Cancer & Brain Damage

http://www.hadassah.org/news-stories/blood-test-detects-neurodegenerative-disease.html

Glutamine and cancer: cell biology, physiology, and clinical opportunities

Christopher T. Hensley,¹ Ajla T. Wasti,^1,2 

J Clin Invest 2013   https://www.jci.org/articles/view/69600

Glutamine is an abundant and versatile nutrient that participates in energy formation, redox homeostasis, macromolecular synthesis, and signaling in cancer cells. These characteristics make glutamine metabolism an appealing target for new clinical strategies to detect, monitor, and treat cancer. Here we review the metabolic functions of glutamine as a super nutrient and the surprising roles of glutamine in supporting the biological hallmarks of malignancy. We also review recent efforts in imaging and therapeutics to exploit tumor cell glutamine dependence, discuss some of the challenges in this arena, and suggest a disease-focused paradigm to deploy these emerging approaches.

It has been nearly a century since the discovery that tumors display metabolic activities that distinguish them from differentiated, non-proliferating tissues and presumably contribute to their supraphysiological survival and growth (1). Interest in cancer metabolism was boosted by discoveries that oncogenes and tumor suppressors could regulate nutrient metabolism, and that mutations in some metabolic enzymes participate in the development of malignancy (2, 3). The persistent appeal of cancer metabolism as a line of investigation lies both in its ability to uncover fundamental aspects of malignancy and in the translational potential of exploiting cancer metabolism to improve the way we diagnose, monitor, and treat cancer. Furthermore, an improved understanding of how altered metabolism contributes to cancer has a high potential for synergy with translational efforts. For example, the demonstration that asparagine is a conditionally essential nutrient in rapidly growing cancer cells paved the way for L-asparaginase therapy in leukemia. Additionally, the avidity of some tumors for glucose uptake led to the development of ¹⁸fluoro-2-deoxyglucose imaging by PET; this in turn stimulated hundreds of studies on the biological underpinnings of tumor glucose metabolism.

There continue to be large gaps in understanding which metabolic pathways are altered in cancer, whether these alterations benefit the tumor in a substantive way, and how this information could be used in clinical oncology. In this Review, we consider glutamine, a highly versatile nutrient whose metabolism has implications for tumor cell biology, metabolic imaging, and perhaps novel therapeutics.

Glutamine in intermediary metabolism

Glutamine metabolism has been reviewed extensively and is briefly outlined here (4, 5). The importance of glutamine as a nutrient in cancer derives from its abilities to donate its nitrogen and carbon into an array of growth-promoting pathways (Figure 1). At concentrations of 0.6–0.9 mmol/l, glutamine is the most abundant amino acid in plasma (6). Although most tissues can synthesize glutamine, during periods of rapid growth or other stresses, demand outpaces supply, and glutamine becomes conditionally essential (7). This requirement for glutamine is particularly true in cancer cells, many of which display oncogene-dependent addictions to glutamine in culture (8). Glutamine catabolism begins with its conversion to glutamate in reactions that either donate the amide nitrogen to biosynthetic pathways or release it as ammonia. The latter reactions are catalyzed by the glutaminases (GLSs), of which several isozymes are encoded by human genes GLS and GLS2 (9). Classical studies revealed that GLS isozymes, particularly those encoded by GLS, are expressed in experimental tumors in rats and mice, where their enzyme activity correlates with growth rate and malignancy. Silencing GLS expression or inhibiting GLS activity is sufficient to delay tumor growth in a number of models (10–13). The role of GLS2 in cancer appears to be context specific and regulated by factors that are still incompletely characterized. In some tissues, GLS2 is a p53 target gene and seems to function in tumor suppression (14). On the other hand, GLS2 expression is enhanced in some neuroblastomas, where it contributes to cell survival (15). These observations, coupled with the demonstration that c-Myc stimulates GLS expression (12, 16), position at least some of the GLS isozymes as pro-oncogenic.

Figure 1Glutamine metabolism as a target for diagnostic imaging and therapy in cancer. Glutamine is imported via SLC1A5 and other transporters, then enters a complex metabolic network by which its carbon and nitrogen are supplied to pathways that promote cell survival and growth. Enzymes discussed in the text are shown in green, and inhibitors that target various aspects of glutamine metabolism are shown in red. Green arrows denote reductive carboxylation. ¹⁸F-labeled analogs of glutamine are also under development as PET probes for localization of tumor tissue. AcCoA, acetyl-CoA; DON, 6-diazo-5-oxo-L-norleucine; GSH, glutathione; NEAA, nonessential amino acids; ME, malic enzyme; OAA, oxaloacetate; TA, transaminase; 968, compound 968; α-KG, α-ketoglutarate.

Glutamate, the product of the GLS reaction, is a precursor of glutathione, the major cellular antioxidant. It is also the source of amino groups for nonessential amino acids like alanine, aspartate, serine, and glycine, all of which are required for macromolecular synthesis. In glutamine-consuming cells, glutamate is also the major source of α-ketoglutarate, a TCA cycle intermediate and substrate for dioxygenases that modify proteins and DNA. These dioxygenases include prolyl hydroxylases, histone demethylases, and 5-methylcytosine hydroxylases. Their requirement for α-ketoglutarate, although likely accounting for only a small fraction of total α-ketoglutarate utilization, makes this metabolite an essential component of cell signaling and epigenetic networks.

Conversion of glutamate to α-ketoglutarate occurs either through oxidative deamination by glutamate dehydrogenase (GDH) in the mitochondrion or by transamination to produce nonessential amino acids in either the cytosol or the mitochondrion. During avid glucose metabolism, the transamination pathway predominates (17). When glucose is scarce, GDH becomes a major pathway to supply glutamine carbon to the TCA cycle, and is required for cell survival (17, 18). Metabolism of glutamine-derived α-ketoglutarate in the TCA cycle serves several purposes: it generates reducing equivalents for the electron transport chain (ETC) and oxidative phosphorylation, becoming a major source of energy (19); and it is an important anaplerotic nutrient, feeding net production of oxaloacetate to offset export of intermediates from the cycle to supply anabolism (20). Glutamine oxidation also supports redox homeostasis by supplying carbon to malic enzyme, some isoforms of which produce NADPH (Figure 1). In KRAS-driven pancreatic adenocarcinoma cells, a pathway involving glutamine-dependent NADPH production is essential for redox balance and growth (21). In these cells, glutamine is used to produce aspartate in the mitochondria. This aspartate is then trafficked to the cytosol, where it is deaminated to produce oxaloacetate and then malate, the substrate for malic enzyme.

Recent work has uncovered an unexpected role for glutamine in cells with reduced mitochondrial function. Despite glutamine’s conventional role as a respiratory substrate, several studies demonstrated a persistence of glutamine dependence in cells with permanent mitochondrial dysfunction from mutations in the ETC or TCA cycle, or transient impairment secondary to hypoxia (22–25). Under these conditions, glutamine-derived α-ketoglutarate is reductively carboxylated by NADPH-dependent isoforms of isocitrate dehydrogenase to produce isocitrate, citrate, and other TCA cycle intermediates (Figure 1). These conditions broaden glutamine’s utility as a carbon source because it becomes not only a major source of oxaloacetate, but also generates acetyl-CoA in what amounts to a striking rewiring of TCA cycle metabolism.

Glutamine promotes hallmarks of malignancy

Deregulated energetics. One hallmark of cancer cells is aberrant bioenergetics (26). Glutamine’s involvement in the pathways outlined above contributes to a phenotype conducive to energy formation, survival, and growth. In addition to its role in mitochondrial metabolism, glutamine also suppresses expression of thioredoxin-interacting protein, a negative regulator of glucose uptake (27). Thus, glutamine contributes to both of the energy-forming pathways in cancer cells: oxidative phosphorylation and glycolysis. Glutamine also modulates hallmarks not traditionally thought to be metabolic, as outlined below. These interactions highlight the complex interplay between glutamine metabolism and many aspects of cell biology.

Sustaining proliferative signaling. Pathological cancer cell growth relies on maintenance of proliferative signaling pathways with increased autonomy relative to non-malignant cells. Several lines of evidence argue that glutamine reinforces activity of these pathways. In some cancer cells, excess glutamine is exported in exchange for leucine and other essential amino acids. This exchange facilitates activation of the serine/threonine kinase mTOR, a major positive regulator of cell growth (28). In addition, glutamine-derived nitrogen is a component of amino sugars, known as hexosamines, that are used to glycosylate growth factor receptors and promote their localization to the cell surface. Disruption of hexosamine synthesis reduces the ability to initiate signaling pathways downstream of growth factors (29).

Enabling replicative immortality. Some aspects of glutamine metabolism oppose senescence and promote replicative immortality in cultured cells. In IMR90 lung fibroblasts, silencing either of two NADPH-generating isoforms of malic enzyme (ME1, ME2) rapidly induced senescence, while malic enzyme overexpression suppressed senescence (30). Both malic enzyme isoforms are repressed at the transcriptional level by p53 and contribute to enhanced levels of glutamine consumption and NADPH production in p53-deficient cells. The ability of p53-replete cells to resist senescence required the expression of ME1 and ME2, and silencing either enzyme reduced the growth of TP53^+/+ and, to a lesser degree, TP53^–/– tumors (30). These observations position malic enzymes as potential therapeutic targets.

Resisting cell death. Although many cancer cells require glutamine for survival, cells with enhanced expression of Myc oncoproteins are particularly sensitive to glutamine deprivation (8, 12, 16). In these cells, glutamine deprivation induces depletion of TCA cycle intermediates, depression of ATP levels, delayed growth, diminished glutathione pools, and apoptosis. Myc drives glutamine uptake and catabolism by activating the expression of genes involved in glutamine metabolism, including GLSand SLC1A5, which encodes the Na⁺-dependent amino acid transporter ASCT2 (12, 16). SilencingGLS mimicked some of the effects of glutamine deprivation, including growth suppression in Myc-expressing cells and tumors (10, 12). MYCN amplification occurs in 20%–25% of neuroblastomas and is correlated with poor outcome (31). In cells with high N-Myc levels, glutamine deprivation triggered an ATF4-dependent induction of apoptosis that could be prevented by restoring downstream metabolites oxaloacetate and α-ketoglutarate (15). In this model, pharmacological activation of ATF4, inhibition of glutamine metabolic enzymes, or combinations of these treatments mimicked the effects of glutamine deprivation in cells and suppressed growth of MYCN-amplified subcutaneous and transgenic tumors in mice.

The PKC isoform PKC-ζ also regulates glutamine metabolism. Loss of PKC-ζ enhances glutamine utilization and enables cells to survive glucose deprivation (32). This effect requires flux of carbon and nitrogen from glutamine into serine. PKC-ζ reduces the expression of phosphoglycerate dehydrogenase, an enzyme required for glutamine-dependent serine biosynthesis, and also phosphorylates and inactivates this enzyme. Thus, PKC-ζ loss, which promotes intestinal tumorigenesis in mice, enables cells to alter glutamine metabolism in response to nutrient stress.

Invasion and metastasis. Loss of the epithelial cell-cell adhesion molecule E-cadherin is a component of the epithelial-mesenchymal transition, and is sufficient to induce migration, invasion, and tumor progression (33, 34). Addiction to glutamine may oppose this process because glutamine favors stabilization of tight junctions in some cells (35). Furthermore, the selection of breast cancer cells with the ability to grow without glutamine yielded highly adaptable subpopulations with enhanced mesenchymal marker expression and improved capacity for anchorage-independent growth, therapeutic resistance, and metastasis in vivo (36). It is unknown whether this result reflects a primary role for glutamine in suppressing these markers of aggressiveness in breast cancer, or whether prolonged glutamine deprivation selects for cells with enhanced fitness across a number of phenotypes.

Organ-specific glutamine metabolism in health and disease

As a major player in carbon and nitrogen transport, glutamine metabolism displays complex inter-organ dynamics, with some organs functioning as net producers and others as consumers (Figure 2). Organ-specific glutamine metabolism has frequently been studied in humans and animal models by measuring the arteriovenous difference in plasma glutamine abundance. In healthy subjects, the plasma glutamine pool is largely the result of release from skeletal muscle (37–39). In rats, the lungs are comparable to muscle in terms of glutamine production (40, 41), and human lungs also have the capacity for marked glutamine release, although such release is most prominent in times of stress (42, 43). Stress-induced release from the lung is regulated by an induction of glutamine synthase expression as a consequence of glucocorticoid signaling and other mechanisms (44, 45). Although this results in a small arteriovenous difference, the overall release of glutamine is significant because of the large pulmonary perfusion. In rats and humans, adipose tissue is a minor but potentially important source of glutamine (46, 47). The liver has the capacity to synthesize or catabolize glutamine, with these activities subject both to regional heterogeneity among hepatocytes and regulatory effects of systemic acidosis and hyperammonemia. However, the liver does not appear to be a major contributor to the plasma glutamine pool in healthy rats and humans (39, 48, 49).

Figure 2Model for inter-organ glutamine metabolism in health and cancer. Organs that release glutamine into the bloodstream are shown in green, and those that consume glutamine are in red; the shade denotes magnitude of consumption/release. For some organs (liver, kidneys), evidence from model systems and/or human studies suggests that there is a change in net glutamine flux during tumorigenesis.

Glutamine consumption occurs largely in the gut and kidney. The organs of the gastrointestinal tract drained by the portal vein, particularly the small intestine, are major consumers of plasma glutamine in both rats and humans (37, 38, 49, 50). Enterocytes oxidize more than half of glutamine carbon to CO₂, accounting for a third of the respiration of these cells in fasting animals (51). The kidney consumes net quantities of glutamine to maintain acid-base balance (37, 38, 52, 53). During acidosis, the kidneys substantially increase their uptake of glutamine, cleaving it by GLS to produce ammonia, which is excreted along with organic acids to maintain physiologic pH (52, 54). Glutamine is also a major metabolic substrate in lymphocytes and macrophages, at least during mitogenic stimulation of primary cells in culture (55–57).

Importantly, cancer seems to cause major changes in inter-organ glutamine trafficking (Figure 2). Currently, much work in this area is derived from studies in methylcholanthrene-induced fibrosarcoma in the rat, a model of an aggressively growing, glutamine-consuming tumor. In this model, fibrosarcoma induces skeletal muscle expression of glutamine synthetase and greatly increases the release of glutamine into the circulation. As the tumor increases in size, intramuscular glutamine pools are depleted in association with loss of lean muscle mass, mimicking the cachectic phenotype of humans in advanced stages of cancer (52). Simultaneously, both the liver and the kidneys become net glutamine exporters, although the hepatic effect may be diminished as the tumor size becomes very large (48, 49, 52). Glutamine utilization by organs supplied by the portal vein is diminished in cancer (48). In addition to its function as a nutrient for the tumor itself, and possibly for cancer-associated immune cells, glutamine provides additional, indirect metabolic benefits to both the tumor and the host. For example, glutamine was used as a gluconeogenic substrate in cachectic mice with large orthotopic gliomas, providing a significant source of carbon in the plasma glucose pool (58). This glucose was taken up and metabolized by the tumor to produce lactate and to supply the TCA cycle.

It will be valuable to extend work in human inter-organ glutamine trafficking, both in healthy subjects and in cancer patients. Such studies will likely produce a better understanding of the pathophysiology of cancer cachexia, a major source of morbidity and mortality. Research in this area should also aid in the anticipation of organ-specific toxicities of drugs designed to interfere with glutamine metabolism. Alterations of glutamine handling in cancer may induce a different spectrum of toxicities compared with healthy subjects.

Tumors differ according to their need for glutamine

One important consideration is that not all cancer cells need an exogenous supply of glutamine. A panel of lung cancer cell lines displayed significant variability in their response to glutamine deprivation, with some cells possessing almost complete independence (59). Breast cancer cells also demonstrate systematic differences in glutamine dependence, with basal-type cells tending to be glutamine dependent and luminal-type cells tending to be glutamine independent (60). Resistance to glutamine deprivation is associated with the ability to synthesize glutamine de novo and/or to engage alternative pathways of anaplerosis (10, 60).

Tumors also display variable levels of glutamine metabolism in vivo. A study of orthotopic gliomas revealed that genetically diverse, human-derived tumors took up glutamine in the mouse brain but did not catabolize it (58). Rather, the tumors synthesized glutamine de novo and used pyruvate carboxylation for anaplerosis. Cells derived from these tumors did not require glutamine to survive or proliferate when cultured ex vivo. Glutamine synthesis from glucose was also a prominent feature of primary gliomas in human subjects infused with ¹³C-glucose at the time of surgical resection (61). Furthermore, an analysis of glutamine metabolism in lung and liver tumors revealed that both the tissue of origin and the oncogene influence whether the tumor produces or consumes glutamine (62). MET-induced hepatic tumors produced glutamine, whereas Myc-induced liver tumors catabolized it. In the lung, however, Myc expression was associated with glutamine accumulation.

This variability makes it imperative to develop ways to predict which tumors have the highest likelihood of responding to inhibitors of glutamine metabolism. Methods to image or otherwise quantify glutamine metabolism in vivo would be useful in this regard (63). Infusions of pre-surgical subjects with isotopically labeled glutamine, followed by extraction of metabolites from the tumor and analysis of ¹³C enrichment, can be used to detect both glutamine uptake and catabolism (58, 62). However, this approach requires a specimen of the tumor to be obtained. Approaches for glutamine-based imaging, which avoid this problem, include a number of glutamine analogs compatible with PET. Although glutamine could in principle be imaged using the radioisotopes ¹¹C, ¹³N, or ¹⁸F, the relatively long half-life of the latter increases its appeal. In mice, ¹⁸F-(2S, 4R)4-fluoroglutamine is avidly taken up by tumors derived from highly glutaminolytic cells, and by glutamine-consuming organs including the intestine, kidney, liver, and pancreas (64). Labeled analogs of glutamate are also taken up by some tumors (65, 66). One of these, (4S)-4-(3-[¹⁸F] fluoropropyl)-L-glutamate (¹⁸F-FSPG, also called BAY 94-9392), was evaluated in small clinical trials involving patients with several types of cancer (65, 67). This analog enters the cell through the cystine/glutamate exchange transporter (xC^–transport system), which is linked to glutathione biosynthesis (68). The analog was well tolerated, with high tumor detection rates and good tumor-to-background ratios in hepatocellular carcinoma and lung cancer.

PET approaches detect analog uptake and retention but cannot provide information about downstream metabolism. Analysis of hyperpolarized nuclei can provide a real-time view of enzyme-catalyzed reactions. This technique involves redistribution of the populations of energy levels of a nucleus (e.g., ¹³C, ¹⁵N), resulting in a gain in magnetic resonance signal that can temporarily exceed 10,000-fold (69). This gain in signal enables rapid detection of both the labeled molecule and its downstream metabolites. Glutamine has been hyperpolarized on ¹⁵N and ¹³C (70, 71). In the latter case, the conversion of hyperpolarized glutamine to glutamate could be detected in intact hepatoma cells (70). If these analogs are translated to clinical studies, they might provide a dynamic view of the proximal reactions of glutaminolysis in vivo.

Pharmacological strategies to inhibit glutamine metabolism in cancer

Efforts to inhibit glutamine metabolism using amino acid analogs have an extensive history, including evaluation in clinical trials. Acivicin, 6-diazo-5-oxo-L-norleucine, and azaserine, three of the most widely studied analogs (Figure 1), all demonstrated variable degrees of gastrointestinal toxicity, myelosuppression, and neurotoxicity (72). Because these agents non-selectively target glutamine-consuming processes, recent interest has focused on developing methods directed at specific nodes of glutamine metabolism. First, ASCT2, the Na⁺-dependent neutral amino acid transporter encoded by SLC1A5, is broadly expressed in lung cancer cell lines and accounts for a majority of glutamine transport in those cells (Figure 1). It has been shown that γ-L-glutamyl-p-nitroanilide (GPNA) inhibits this transporter and limits lung cancer cell growth (73). Additional interest in GPNA lies in its ability to enhance the uptake of drugs imported via the monocarboxylate transporter MCT1. Suppressing glutamine uptake with GPNA enhances MCT1 stability and stimulates uptake of the glycolytic inhibitor 3-bromopyruvate (3-BrPyr) (74, 75). Because enforced MCT1 overexpression is sufficient to sensitize tumor xenografts to 3-BrPyr (76), GPNA may have a place in 3-BrPyr–based therapeutic regimens.

Two inhibitors of GLS isoforms have been characterized in recent years (Figure 1). Compound 968, an inhibitor of the GLS-encoded splice isoform GAC, inhibits the transformation of fibroblasts by oncogenic RhoGTPases and delays the growth of GLS-expressing lymphoma xenografts (13). Bis-2-(5-phenylacetamido-1,2,4-thiadiazol-2-yl)ethyl sulfide (BPTES) also potently inhibits GLS isoforms encoded by GLS (77). BPTES impairs ATP levels and growth rates of P493 lymphoma cells under both normoxic and hypoxic conditions and suppresses the growth of P493-derived xenografts (78).

Evidence also supports a role for targeting the flux from glutamate to α-ketoglutarate, although no potent, specific inhibitors yet exist to inhibit these enzymes in intact cells. Aminooxyacetate (AOA) inhibits aminotransferases non-specifically, but milliomolar doses are typically used to achieve this effect in cultured cells (Figure 1). Nevertheless, AOA has demonstrated efficacy in both breast adenocarcinoma xenografts and autochthonous neuroblastomas in mice (15, 79). Epigallocatechin gallate (EGCG), a green tea polyphenol, has numerous pharmacological effects, one of which is to inhibit GDH (80). The effects of EGCG on GDH have been used to kill glutamine-addicted cancer cells during glucose deprivation or glycolytic inhibition (17, 18) and to suppress growth of neuroblastoma xenografts (15).

A paradigm to exploit glutamine metabolism in cancer

Recent advances in glutamine-based imaging, coupled with the successful application of glutamine metabolic inhibitors in mouse models of cancer, make it possible to conceive of treatment plans that feature consideration of tumor glutamine utilization. A key challenge will be predicting which tumors are most likely to respond to inhibitors of glutamine metabolism. Neuroblastoma is used here as an example of a tumor in which evidence supports the utility of strategies that would involve both glutamine-based imaging and therapy (Figure 3). Neuroblastoma is the second most common extracranial solid malignancy of childhood. High-risk neuroblastoma is defined by age, stage, and biological features of the tumor, including MYCN amplification, which occurs in some 20%–25% of cases (31). Because MYCN-amplified tumor cells require glutamine catabolism for survival and growth (15), glutamine-based PET at the time of standard diagnostic imaging could help predict which tumors would be likely to respond to inhibitors of glutamine metabolism. Infusion of ¹³C-glutamine coordinated with the diagnostic biopsy could then enable inspection of ¹³C enrichment in glutamine-derived metabolites from the tumor, confirming the activity of glutamine catabolic pathways. Following on evidence from mouse models of neuroblastoma, treatment could then include agents directed against glutamine catabolism (15). Of note, some tumors were sensitive to the ATF4 agonist fenretinide (FRT), alone or in combination with EGCG. Importantly, FRT has already been the focus of a Phase I clinical trial in children with solid tumors, including neuroblastoma, and was fairly well tolerated (81).

Figure 3A strategy to integrate glutamine metabolism into the diagnosis, classification, treatment, and monitoring of neuroblastoma. Neuroblastoma commonly presents in children as an abdominal mass. A standard evaluation of a child with suspected neuroblastoma includes measurement of urine catecholamines, a bone scan, and full-body imaging with meta-iodobenzylguanidine (MIBG), all of which contribute to diagnosis and disease staging. In animal models, a subset of these tumors requires glutamine metabolism. This finding implies that approaches to image, quantify, or block glutamine metabolism (highlighted in red) in human neuroblastoma could be incorporated into the diagnosis and management of this disease. In particular, glutamine metabolic studies may help predict which tumors would respond to therapies targeting glutamine metabolism. HVA, homovanillic acid; VMA, vanillylmandelic acid.

Conclusions

Glutamine is a versatile nutrient required for the survival and growth of a potentially large subset of tumors. Work over the next several years should produce a more accurate picture of the molecular determinants of glutamine addiction and the identification of death pathways that execute cells when glutamine catabolism is impaired. Advancement of glutamine-based imaging into clinical practice should soon make it possible to differentiate tumors that take up glutamine from those that do not. Finally, the development of safe, high-potency inhibitors of key metabolic nodes should facilitate therapeutic regimens featuring inhibition of glutamine metabolism.

Therapeutic strategies impacting cancer cell glutamine metabolism

Michael J Lukey, Kristin F Wilson, and Richard A Cerione^*

Future Med Chem. 2013 Sep; 5(14): 1685–1700.   doi:  10.4155/fmc.13.130

A second major change in the metabolic program of many cancer cells, and the primary focus of this review, is the alteration of glutamine metabolism. Glutamine is the major carrier of nitrogen between organs, and the most abundant amino acid in plasma [7]. It is also a key nutrient for numerous intracellular processes including oxidative metabolism and ATP generation, biosynthesis of proteins, lipids and nucleic acids, and also redox homeostasis and the regulation of signal transduction pathways [8–10]. Although most mammalian cells are capable of synthesizing glutamine, the demand for this amino acid can become so great during rapid proliferation that an additional extracellular supply is required; hence glutamine is considered conditionally essential [11]. Indeed, many cancer cells are ‘glutamine addicted’, and cannot survive in the absence of an exogenous glutamine supply [12,13].

An important step in the elevation of glutamine catabolism is the activation of the mitochondrial enzyme glutaminase, which catalyzes the hydrolysis of glutamine to generate glutamate and ammonium. The subsequent deamination of glutamate releases a second ammonium to yield the TCA cycle intermediate α-ketoglutarate (α-KG), a reaction catalyzed by glutamate dehydrogenase (GLUD1). This series of reactions is particularly important in rapidly proliferating cells, in which a considerable proportion of the TCA cycle metabolite citrate is exported from mitochondria in order to generate cytosolic acetyl-CoA for lipid biosynthesis [14]. Replenishment of TCA cycle intermediates (anaplerosis) is therefore required, and glutamine often serves as the key anaplerotic substrate through its conversion via glutamate to α-KG (Figure 1).

Mammals express two genes for glutaminase enzymes [15–17]. The GLS gene encodes a protein initially characterized in kidney and thus called kidney-type glutaminase (KGA), although this enzyme and its shorter splice variant glutaminase C (GAC), collectively referred to as GLS, are now known to be widely distributed [18–20]. The KGA and GAC isoforms share identical N-terminal and catalytic domains, encoded by exons 1–14 of the GLS gene, but have distinct C-termini derived from exon 15 in the case of GAC and exons 16–19 in the case of KGA [21]. Upregulation of GLS, in particular the GAC iso-form, is common in cancer cells and the degree of GLS overexpression correlates with both the degree of malignancy and the tumor grade in human breast cancer samples [22,23]. The GLS2 gene encodes a protein originally discovered and characterized in liver, which has thus been referred to as liver-type glutaminase and, more recently, as glutaminase 2 (GLS2) [15].

Both KGA and GAC can be activated by inorganic phosphate (Pi), and this activation correlates closely with a dimer-to-tetramer transition for each enzyme [7, 22]. As the concentration of Pi is raised the apparent catalytic constant, k_cat^app, increases and simultaneously the apparent Michaelis constant, K_m^app, decreases; consequently the catalytic efficiency rises dramatically, especially in the case of GAC [22]. x-ray crystal structures of GAC and KGA in different states indicate that the positioning of a key loop within each monomer (Glu312 to Pro329), located between the active site and the dimer–dimer interface, is critical for mediating tetramerization-induced activation [22,24]. Given the ability of Pi to promote tetramerization and activation of GAC and KGA, it has been proposed that the elevated mitochondrial Pi levels found under hypoxic conditions, which are commonly encountered in the tumor microenvironment, could be one trigger for GLS activation [22].

Oncogenic alterations affecting glutamine metabolism

At least two classes of cellular signals regulate glutamine metabolism, influencing both the expression level and the enzymatic activity of GLS. The transcription factor c-Myc can suppress the expression of microRNAs miR-23a and miR-23b and, in doing so, upregulates GLS (specifically GAC) expression [13,25]. Independent of changes in GAC expression, oncogenic diffuse B-cell lymphoma protein (Dbl), a GEF for Rho GTPases and oncogenic variants of downstream Rho GTPases are able to signal to activate GAC in a manner that is dependent on NF-κB [23]. Mitochondria isolated from Dbl- or Rho GTPase-transformed NIH-3T3 fibroblasts demonstrate significantly higher basal glutaminase activity than mitochondria isolated from non-transformed cells [23]. Furthermore, the enzymatic activity of GAC immunoprecipitated from Dbl-transformed cells is elevated relative to GAC from non-transformed cells, indicating the presence of activating post-translational modification(s) [23]. Indeed, when GAC isolated from Dbl-transformed cells is treated with alkaline phosphatase, basal enzymatic activity is dramatically reduced [23]. Collectively, these findings point to phosphorylation events underlying the activation of GAC in transformed cells. Similarly, phosphorylation-dependent regulation of KGA activity downstream of the Raf-Mek-Erk signaling axis occurs in response to EGF stimulation [24].

It is becoming clear that, in addition to c-Myc and Dbl, many other oncogenic signals and environmental conditions can impact cellular glutamine metabolism. Loss of the retinoblastoma tumor suppressor, for example, leads to a marked increase in glutamine uptake and catabolism, and renders mouse embryonic fibroblasts dependent on exogenous glutamine [26]. Cells transformed by KRAS also illustrate increased expression of genes associated with glutamine metabolism and a corresponding increased utilization of glutamine for anabolic synthesis [27]. In fact, KRAS signaling appears to induce glutamine dependence, since the deleterious effects of glutamine withdrawal in KRAS-driven cells can be rescued by expression of a dominant-negative GEF for Ras [28]. Downstream of Ras, the Raf-MEK-ERK signaling pathway has been implicated in the upregulation of glutamine uptake and metabolism [24,29]. A recent study using human pancreatic ductal adenocarcinoma cells identified a novel KRAS-regulated metabolic pathway, through which glutamine supports cell growth [30]. Proliferation of KRAS-mutant pancreatic ductal adenocarcinoma cells depends on GLS-catalyzed production of glutamate, but not on downstream deamination of glutamate to α-KG; instead, transaminase-mediated glutamate metabolism is essential for growth. Glutamine-derived aspartate is subsequently transported into the cytoplasm where it is converted by aspartate transaminase into oxaloacetate, which can be used to generate malate and pyruvate. The series of reactions maintains NADPH levels and thus the cellular redox state [30].

Other recent studies have revealed that another pathway for glutamine metabolism can be essential under hypoxic conditions, and also in cancer cells with mitochondrial defects or loss of the VHL tumor suppressor [31–35]. In these situations, glutamine-derived α-KG undergoes reductive carboxylation by IDH1 or IDH2 to generate citrate, which can be exported from mitochondria to support lipogenesis (Figure 1). Activation of HIF is both necessary and sufficient for driving the reductive carboxylation phenotype in renal cell carcinoma, and suppression of HIF activity can induce a switch from glutamine-mediated lipogenesis back to glucose-mediated lipogenesis [32,35]. Furthermore, loss of VHL and consequent downstream activation of HIF renders renal cell carcinoma cells sensitive to inhibitors of GLS [35]. Evidently, the metabolic routes through which glutamine supports cancer cell proliferation vary with genetic background and with microenvironmental conditions. Nevertheless, it is increasingly clear that diverse oncogenic signals promote glutamine utilization and furthermore that hypoxia, a common condition within poorly vascularized tumors, increases glutamine dependence.

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Mitochondria as biosynthetic factories for cancer proliferation

Christopher S Ahn and Christian M Metallo

Cancer & Metabolism (2015) 3:1      http://dx.doi.org:/10.1186/s40170-015-0128-2

Unchecked growth and proliferation is a hallmark of cancer, and numerous oncogenic mutations reprogram cellular metabolism to fuel these processes. As a central metabolic organelle, mitochondria execute critical biochemical functions for the synthesis of fundamental cellular components, including fatty acids, amino acids, and nucleotides. Despite the extensive interest in the glycolytic phenotype of many cancer cells, tumors contain fully functional mitochondria that support proliferation and survival. Furthermore, tumor cells commonly increase flux through one or more mitochondrial pathways, and pharmacological inhibition of mitochondrial metabolism is emerging as a potential therapeutic strategy in some cancers. Here, we review the biosynthetic roles of mitochondrial metabolism in tumors and highlight specific cancers where these processes are activated.

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Recent characterizations of metabolic enzymes as tumor suppressors and oncogene-driven metabolic reprogramming have reinvigorated interest in cancer metabolism. Although therapies targeting metabolic processes have long been a staple in cancer treatment (e.g. inhibition of folate metabolism via methotrexate), the focused therapeutic potential surrounding these findings have generated a renewed appreciation for Otto Warburg’s work almost a century ago. Warburg observed that tumor cells ferment much of the glucose taken up during growth to lactate, thus using glycolysis as a major means of adenosine triphosphate (ATP) regeneration [1]. However, the observation of decreased respiration in cancer cells and idea that “the respiration of all cancer cells is damaged” belies the critical role of mitochondria in biosynthesis and cell survival [1]. On the contrary, functional mitochondria are present in all proliferative cells within our body (including all tumors), as they are responsible for converting the diverse nutrients available to cells into the fundamental building blocks required for cell growth. These organelles execute numerous functions in cancer cells to promote tumor growth and survival in response to stress. Here, we outline the critical biosynthetic functions served by mitochondria within tumors (Figure 1). Although many of these functions are similarly important in normal, proliferating cells, we have attempted to highlight potential points where mitochondrial metabolism may be therapeutically targeted to slow cancer growth. This review is organized by specific metabolic pathways or processes (i.e., glucose metabolism and lipogenesis, amino acid metabolism, and nucleotide biosynthesis). Tumors or cancer cell types where enzymes in each pathway have been specifically observed to by dysregulated are described within the text and summarized in Table 1.

Figure 1

Biosynthetic nodes within mitochondria. Metabolic pathways within mitochondria that contribute to biosynthesis in cancer and other proliferating cells. TCA metabolism and FOCM enable cells to convert carbohydrates and amino acids to lipids, non-essential amino acids, nucleotides (including purines used for cofactor synthesis), glutathione, heme, and other cellular components. Critical biosynthetic routes are indicated by yellow arrows. Enzymatic reactions that are dependent on redox-sensitive cofactors are depicted in red.  https://static-content.springer.com/image/art%3A10.1186%2Fs40170-015-0128-2/MediaObjects/40170_2015_128_Fig1_HTML.gif

Table 1

Overview of mitochondrial biosynthetic enzymes important in cancer

TCA cycle, anaplerosis, and AcCoA metabolism

Cancers in which three or more mitochondrial enzymes have been studied and found to be differentially regulated (or mutated, as indicated) in cancers vs. control groups are included. Dysregulation of each enzyme was demonstrated in clinical tumors samples, animal models, or cell lines at the levels of genes, mRNA, protein, metabolites, and/or flux.

Figure 2

Coordination of carbon and nitrogen metabolism across amino acids. Glutamate and aKG are key substrates in numerous transamination reactions and can also serve as precursors for glutamine, proline, and the TCA cycle. Mitochondrial enzymes catalyzing these reactions are highlighted in blue, and TCA cycle intermediates are highlighted in orange (pyruvate enters the TCA cycle as acetyl-CoA or oxaloacetate).
https://static-content.springer.com/image/art%3A10.1186%2Fs40170-015-0128-2/MediaObjects/40170_2015_128_Fig2_HTML.gif

Figure 3

Biosynthetic sources for purine and pyrimidine synthesis. Sources and fates of nitrogen, carbon, and oxygen atoms are colored as indicated. Italicized metabolites can be sourced from the mitochondria or cytosol. The double bond formed by the action of DHODH/ubiquinone is also indicated.      https://static-content.springer.com/image/art%3A10.1186%2Fs40170-015-0128-2/MediaObjects/40170_2015_128_Fig3_HTML.gif

Mitochondria operate as both engine and factory in eukaryotes, coordinating cellular energy production and the availability of fundamental building blocks that are required for cell proliferation. Cancer cells must therefore balance their relative bioenergetic and biosynthetic needs to grow, proliferate, and survive within the physical constraints of energy and mass conservation. In contrast to quiescent cells, which predominantly use oxidative mitochondrial metabolism to produce ATP and uptake glucose at much lower rates than proliferating cells, tumor cells exhibit increased glycolytic rates to provide an elevated flux of substrate for biosynthetic pathways, including those executed within mitochondria. Given these higher rates of nutrient utilization, metabolic flux through mitochondrial pathways and the associated ROS production can often be higher in cancer cells. Not surprisingly, activation of cellular antioxidant response pathways is commonly observed in cancer or subpopulations of cells within tumors [46,78]. Cellular compartmentalization affords a degree of protection from such damaging side products of metabolism, and methods which are able to deconvolute the relative contributions of each cellular compartment (e.g. mitochondria, cytosol, peroxisome, etc.) to cancer metabolism will be crucial to more completely understand the metabolism of cancer cells in the future [74,79]. Ultimately, while mitochondrial dysregulation is widely considered to be a hallmark of cancer, numerous mitochondrial functions remain critical for tumor growth and are emerging as clinical targets.

Following this point, it comes as no surprise that mitochondrial metabolism is highly active in virtually all tumors (i.e., cancer cells, stroma, or both), and investigators have begun targeting these pathways to explore potential efficacy. Indeed, some evidence suggests that biguanides such as metformin or phenformin may limit tumor incidence and burden in humans and animals [80,81]. These effects are presumably due, at least in part, to complex I inhibition of the ETC, which significantly perturbs mitochondrial function [82,83]. However, more insights are needed into the mechanisms of these compounds in patients to determine the therapeutic potential of targeting this and other components of mitochondria. In developing new therapies that target cancer metabolism, researchers will face challenges similar to those that are relevant for many established chemotherapies since deleterious effects on normal proliferating cells that also depend on mitochondrial metabolism (and aerobic glycolysis) are likely to arise.

As we acquire a more detailed picture of how specific genetic modifications in a patient’s tumor correlate with its metabolic profile, opportunities for designing targeted or combinatorial therapies will become increasingly apparent. Cancer therapies that address tumor-specific mitochondrial dysregulation and dysfunction may be particularly effective. For example, some cancer cells harbor mutations in TCA enzymes (e.g., FH, SDH, IDH2) or regulatory proteins that control mitophagy (i.e., LKB1) [84]. Such tumors may be compromised with respect to some aspects of mitochondrial biosynthesis and dependent on alternate pathways for growth and/or survival such that synthetically lethal targets emerge. Ultimately, such strategies will require clinicians and researchers to coordinate metabolic, biochemical, and genetic information in the design of therapeutic strategies.

 

David Terrano, M.D., Ph.D. commented on your update “Not well versed in Nat peptides so I could not say. I also hesitate with any PNAS paper because those in their academy tend to have a fast track to publication. It has been that way since at least early 2000’s wh n I began research. I don’t doubt their goal and approach (this same group leads the way in methylation-based diagnosis of CNS neoplasms, which is apparently highly accurate). But when I see “dying cells” I know what that means biochemically and look for those hallmarks. Organ specific oligonucleosomes would be a nice cell death surrogate. “

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