Next Article in Journal
Inflammation, Fibrosis and Cancer: Mechanisms, Therapeutic Options and Challenges
Next Article in Special Issue
Expression of 3-Methylcrotonyl-CoA Carboxylase in Brain Tumors and Capability to Catabolize Leucine by Human Neural Cancer Cells
Previous Article in Journal
Kras Gene Analysis Using Liquid-Based Cytology Specimens Predicts Therapeutic Responses and Prognosis in Patients with Pancreatic Cancer
Previous Article in Special Issue
Resveratrol’s Anti-Cancer Effects through the Modulation of Tumor Glucose Metabolism
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Cancers
Volume 14
Issue 3
10.3390/cancers14030553
cancers-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Review

Advancing Cancer Treatment by Targeting Glutamine Metabolism—A Roadmap

by
Anna Halama
* and
Karsten Suhre
*
Department of Physiology and Biophysics, Weill Cornell Medicine-Qatar, Doha 24144, Qatar
*
Authors to whom correspondence should be addressed.
Cancers 2022, 14(3), 553; https://doi.org/10.3390/cancers14030553
Submission received: 1 January 2022 / Revised: 19 January 2022 / Accepted: 19 January 2022 / Published: 22 January 2022
(This article belongs to the Special Issue Significance of Altered (Glucose) Metabolism in Cancers)
Review Reports Versions Notes

Abstract

:

Simple Summary

Dysregulated glutamine metabolism is one of the metabolic features evident in cancer cells when compared to normal cells. Cancer cells utilize glutamine for energy generation as well as the synthesis of other molecules that are critical for cancer growth and progression. Therefore, drugs targeting glutamine metabolism have been extensively investigated. However, inhibition of glutamine metabolism in cancer cells results in the activation of other metabolic pathways enabling cancer cells to survive. In this review, we summarize and discuss the targets in glutamine metabolism, which has been probed in the development of anticancer drugs in preclinical and clinical studies. We further discuss pathways activated in response to glutamine metabolism inhibition, enabling cancer cells to survive the challenge. Finally, we put into perspective combined treatment strategies targeting glutamine metabolism along with other pathways as potential treatment options.

Abstract

Tumor growth and metastasis strongly depend on adapted cell metabolism. Cancer cells adjust their metabolic program to their specific energy needs and in response to an often challenging tumor microenvironment. Glutamine metabolism is one of the metabolic pathways that can be successfully targeted in cancer treatment. The dependence of many hematological and solid tumors on glutamine is associated with mitochondrial glutaminase (GLS) activity that enables channeling of glutamine into the tricarboxylic acid (TCA) cycle, generation of ATP and NADPH, and regulation of glutathione homeostasis and reactive oxygen species (ROS). Small molecules that target glutamine metabolism through inhibition of GLS therefore simultaneously limit energy availability and increase oxidative stress. However, some cancers can reprogram their metabolism to evade this metabolic trap. Therefore, the effectiveness of treatment strategies that rely solely on glutamine inhibition is limited. In this review, we discuss the metabolic and molecular pathways that are linked to dysregulated glutamine metabolism in multiple cancer types. We further summarize and review current clinical trials of glutaminolysis inhibition in cancer patients. Finally, we put into perspective strategies that deploy a combined treatment targeting glutamine metabolism along with other molecular or metabolic pathways and discuss their potential for clinical applications.

1. Introduction

The molecular landscape of cancer cells strongly differs from that of normal cells and contributes to features that are known as cancer hallmarks [1]. The dysregulated metabolism, adjusted to meet the increased energy and biomass production demands, enabling sustained cell division, was described as one of the vital cancer hallmarks [1]. There is a wide spectrum of metabolic processes activated by cancer cells [2]. These extend beyond the initial observation of increased glucose consumption with simultaneous lactate production, independent of the oxygen availability, known as the Warburg effect [3]. For instance, accelerated purine and pyrimidine metabolism and enhanced lipid synthesis are features significantly contributing to the proliferative capacity of cancer cells [4,5]. To cope with this demand, cancer cells modulate their nutrient acquisition mode and redirect their metabolic pathways [2]. The enhanced fluxes of glucose, various amino acids (e.g., glutamine, arginine, serine, and branched-chain amino acids (BCAAs)), folate, and unsaturated fatty acids were previously described along with altered tricarboxylic acid (TCA) cycle metabolism [2,5,6,7,8,9]. Yet, dysregulated glutamine (Gln) metabolism was recognized as central to cancer cell fitness due to its contribution to various metabolic pathways including synthesis of nucleotides, lipids, nonessential amino acids (NEAAs), energy generation, and redox homeostasis.
The metabolic alterations attributed to cancer cells have become an attractive treatment target. One of the first treatment strategies utilizing dysregulated cancer metabolism was targeting elevated nucleotide synthesis [10], proposed by Sidney Farber, the father of modern chemotherapy [11].
Identification of Gln metabolism as central to cancer cells resulted in multiple attempts probing it as a treatment target. Molecules predominantly hampering glutaminolysis by inhibiting glutaminase (GLS), the enzyme catalyzing metabolism of Gln to glutamate, were shown to attenuate cancer growth and proliferation. One of the GLS inhibitors CB-839 (telaglenastat) is currently being investigated in multiple different clinical trials, frequently in combination with chemotherapeutics. Although inhibition of GLS reduces cancer proliferation, we and others have shown that such interventions result in activation of alternative metabolic compensations enabling cancer cell survival under the treatment [12,13]. Therefore, to design rational treatment strategies efficiently targeting cancer, it is critical to carefully analyze those compensatory pathways.
In this review, we focus on various aspects of dysregulated glutamine metabolism observed in cancer cells and its interaction with molecular signaling. We further summarize the treatment strategies proposed to target dysregulated Gln metabolism and will comment on pathways activated in response to such a treatment as potential cancer survival options. Finally, we put in perspective combined treatment options targeting glutamine metabolism along with other molecular or metabolic pathways and discuss their potential for clinical applications. The interplay between Gln metabolism and the immune system, along with Gln metabolism targets contributing to enhanced cancer immunotherapy, was recently reviewed [14] and will not be covered in this review.

2. Overview of Glutamine Metabolism

The amino acid glutamine (Gln) can be synthesized by the human body and is thus considered as a nonessential amino acid (NEAA) [15]. Nevertheless, due to the involvement of Gln in multiple processes, under specific conditions, this amino acid was recognized as conditionally essential [16]. Among all the amino acids, Gln is the most abundant in the body [17] and its concentration depends on the overall body metabolism; the balance between exogenous supply, endogenous synthesis, and release; and its utilization by various tissues. The Gln metabolism across different organs was captured for the first time by Adolf Krebs in 1935 [15]. The Gln synthesis preliminary occurs in the skeletal muscle, lungs, and adipose tissue and is orchestrated by glutamate ammonia ligase (GLUL), also known as glutamine synthase (GS), an enzyme that resides in the cytosol and that catalyzes Gln synthesis from glutamate and ammonium ions (NH4+). The catabolism of Gln is preliminary conducted by the intestinal mucosa, immune cells, renal tubule cells, and liver, where glutaminase (GLS), located in mitochondria, catalyzes the conversion of Gln into glutamate [18].

2.1. Organ-Dependent Glutamine Fate

The role of Gln depends on the tissue of origin. For instance, Gln synthesized in the muscle is released into the bloodstream to support visceral organs with amino nitrogen. Corticosteroids stimulate Gln synthesis and its release by the skeletal muscle, which can be seen during stress conditions such as fasting, injury, and illness when increased Gln amount is released [19]. The primary site for the utilization of released Gln is the gastrointestinal tract and the liver. The cells of the gastrointestinal tract, e.g., colonocytes and enterocytes, utilize Gln as the main respiratory fuel, rather than glucose [20]. The nitrogen generated from Gln during this process is used to synthesize ammonia, alanine, proline, and citrulline, which are released into the bloodstream and then utilized by the liver and kidney [21]. In the liver, the absorbed Gln supports the proliferation of hepatocytes and generation of energy, as well as contributing to gluconeogenesis (de novo glucose synthesis) [15,22]. Additionally, the liver utilizes nitrogen from Gln to fuel the urea cycle and urea formation, which serves as detoxification and regulation of the blood pH [23]. In the kidney, Gln-dependent ammoniagenesis enables the generation of bicarbonate and thus maintains the acid–base balance in the body [24]. The Gln absorbed by the kidney is also utilized for gluconeogenesis and contributes to blood glucose homeostasis [25]. Importantly, the cells of the immune system, including lymphocytes, macrophages, and neutrophils, depend more on Gln than on glucose [26,27,28]. Immunological challenges increase the demand for Gln in those cells [29]. Immune cells utilize Gln both for energy generation and synthesis of macromolecules [30]. In summary, Gln metabolism is regulated in an organ-dependent manner and contributes to a broad spectrum of processes in the body.

2.2. Glutamine Metabolism on the Cellular Level

Extracellular Gln cannot cross the plasma membrane and requires transporters to enter the cells. There are in total 14 such transporters, classified into four different families: SLC1, SLC6, SLC7, and SLC38, which can all support either Gln influx into the cell or efflux from the cell into the extracellular space [31]. These transporters differ in their substrate specificity (as most of the transporters contribute to the transport of other neutral or cationic amino acids), ion- and pH-dependence, and role under physiological and cancer conditions, as described in great detail by Bhutia et al. [31].
Once in the cell, Gln can be incorporated into a versatile spectrum of metabolic pathways in which Gln-derived nitrogen or carbon are utilized. As a nitrogen source, Gln is particularly used for the synthesis of nucleotides (pyrimidines and purines), NEAAs, and glucosamine, whereas carbon from Gln is used for gluconeogenesis, TCA cycle, and glutathione metabolism.
De novo nucleotide synthesis, critical for DNA and RNA, is supported by Gln, which contributes along with nitrogen to the assembly of purines and pyrimidines [32]. In the purine synthesis pathway, Gln serves as a substrate for three enzymes: (1) phosphoribosyl pyrophosphate amidotransferase (PPAT), involved in the conversion of 5-phosphoribosyl-1-pyrophosphate (PRPP) into 5-phosphoribosyl-1-amine (PRA) by deploying nitrogen from Gln [33]; (2) 5-phosphoribosylformylglycinamidine synthase (FGAM synthase), which catalyzes the formation of formylglycinamidine ribonucleotide in an ATP-dependent reaction in which the amino group of glutamine is transferred into formylglycinamidine ribonucleotide (FGAR) [34]; and (3) guanosine monophosphate (GMP) synthetase, which catalyzes the formation of GMP in an ATP-dependent reaction in which amino group from glutamine is transferred to replace the oxygen in the C-2 position of xanthine monophosphate (XMP) [35]. In the pyrimidine pathway, Gln serves as a substrate in two reactions: (1) ATP-dependent synthesis of carbamoyl phosphate, the first step in pyrimidine synthesis, catalyzed by the enzyme carbamoyl phosphate synthetase II (CPS II) [36], and (2) synthesis of cytidine triphosphate (CTP) by amination of uridine triphosphate (UTP), catalyzed by CTP synthase in an ATP-dependent reaction [37].
Gluconeogenesis is a critical process for maintaining glucose homeostasis in the body under different conditions, including fasting [38]. It was shown that Gln is a major gluconeogenic precursor [39]. The process of glucose formation from Gln requires two deamination steps; the first is catalyzed by GLS and results in the formation of glutamate [15], and the second is catalyzed by glutamate dehydrogenase (GDH) and leads to the synthesis of α-ketoglutarate [40]. There are two GLS isoforms, one characterized as kidney (also known as brain)-type (GLS1) and the other defined as liver-type (GLS2) [15,41]. The acquired α-ketoglutarate is incorporated into the TCA cycle where it is metabolized into malate. The malate can be either further oxidized in the mitochondria by malate dehydrogenase (MDH) into oxaloacetate or transported into the cytoplasm and converted into oxaloacetate by the cytosolic form of MDH [42]. The obtained oxaloacetate is decarboxylated to phosphoenolpyruvate (PEP) by the mitochondrial or cytosolic form of phosphoenolpyruvate carboxykinase (PEPCK), dependent on the reaction site [43]. The PEP, synthesized in mitochondria, is transported into the cytosol, and the cytosolic PEP is used for gluconeogenesis [44].
Alternatively, under conditions where glucose is not required, PEP can be metabolized by pyruvate kinase (PK) into pyruvate [45], which further can be incorporated into the lipogenic pathway.
In fact, Gln can serve as a source of lipid synthesis via two routes: (1) the oxidative route, involving metabolism into pyruvate and further formation of acetyl-CoA, precursor for palmitic acid synthesis, in the reaction catalyzed by pyruvate dehydrogenase complex (PDC), and (2) the reductive route, involving incorporation into TCA cycle in the form of α-ketoglutarate which in the reaction of reductive carboxylation, catalyzed by isocitrate dehydrogenase (IDH), is metabolized into citrate and further utilized for lipogenesis [46].
Redox homeostasis, predominantly regulated by NADPH and reduced glutathione (GSH) molecules that neutralize reactive oxygen species (ROS), is critical for proper cell function [47]. The glutamate metabolized from Gln is a substrate in the first step of GSH synthesis, where γ-glutamylcysteine is obtained from glutamate and cysteine in a reaction catalyzed by cysteine ligase [48].
Additionally, glutamate generated from Gln can be utilized as a substrate for the synthesis of other NEAAs, including alanine, aspartate, proline, and serine, as well as γ-aminobutyric acid (GABA) [49].
The NEAA asparagine is the only one generated from Gln and not glutamate in the reaction catalyzed by the asparagine synthetase (ASNS) [50].

3. Dysregulated Glutamine Metabolism in Cancer

In cancer cells, same as in normal cells, Gln contributes to multiple different processes. However, to meet the energy and biomass production requirements, Gln metabolism is accelerated in cancer cells in comparison to normal, non-proliferative cells, and thus the demand is higher. One of the first observations showing enhanced glutamine metabolism in cancer was described by Roberts et al., who found identyfied levels of glutamine in tumors in comparison with normal tissue [51]. This observation was followed by multiple other studies in which dysregulated glutamine metabolism, in the context of various cancers, was described [52,53,54,55,56,57,58]. The summary of the Gln-related dysregulations in the context of involved molecules and cancer type is provided in Table 1.
Overall, in cancer cells, there is a significant demand for Gln, which can be either synthesized or obtained from the bloodstream. The enhanced transport of Gln from the system into the cancer cell can be linked with the observed decrease in the plasma level of Gln in cancer patients across various cancer types [54,59,60]. In fact, enhanced Gln uptake by tumors was suggested to be utilized for tumor imaging where glucose-based positron emission tomography (PET) imaging with 18F-fluorodeoxyglucose (18F-FDG) is inaccurate due to the high background, especially in tissue with high glucose demand, e.g., brain [61]. An investigational PET radiotracer, the 18F-(2S,4R)-4-fluoroglutamine (18F-FGln), was proven as a sensitive strategy to monitor Gln transport and metabolism in human malignancies [62].
Table
Table 1. An overview of metabolic fate of glutamine across different cancers.
Table 1. An overview of metabolic fate of glutamine across different cancers.
Pathway Involved Molecules Cancer Type Study Type References
Increased glutamine transport SLC1A5 Lung cancer Clinical and in vitro [63]
Breast cancer
(TNBC)		 In vitro and in vivo [64]
Head and neck cancer In vitro and in vivo [65]
Colorectal cancer In vitro and in vivo [66,67]
SLC6A14 Pancreatic cancer Clinical, in vitro, and in vivo [68]
SLC38A5 Breast cancer
(TNBC)		 Clinical, in vitro, and in vivo [69]
Pancreatic cancer Clinical and in vivo [70]
Increased glutamine/arginine transport SLC6A14 Cervical cancer Clinical [71]
Colorectal cancer Clinical [72]
Breast cancer (ER+) In vitro and in vivo [73]
Increased glutamine efflux SLC7A5 Colorectal cancer (K-Ras mutation) In vivo [74]
Increased glutaminolysis GLS1 Breast cancer Clinical, in vitro, and in vivo [75,76,77]
Prostate cancer Clinical and in vitro [78,79,80]
Colorectal cancer Clinical, in vitro, and in vivo [81]
Lung cancer Clinical, in vitro, and in vivo [82]
Increased glutaminolysis GLS2 Pancreatic cancer In vivo [83]
Controls glutamine metabolism and ROS level GLS2 * Hepatocellular cancer In vitro [84,85]
Glutamine contributes to antioxidative capacity of cancer cell GCL Breast cancer In vitro and in vivo [86]
Lung cancer In vitro and in vivo [87]
Liver cancer In vivo [88]
GDH1 Lung cancer In vitro and in vivo [89]
Breast cancer In vitro and in vivo [90]
GOT1/GOT2 Pancreatic cancer In vitro and in vivo [91]
GOT2 Pancreatic cancer In vitro [92]
Glutamine contributes to citrate and lipid synthesis through reductive carboxylation (RC) of α-ketoglutarate (αKG) as well as contributing to aspartate and pyrimidine synthesis IDH2 Renal cell carcinoma deficient in the von Hippel–Lindau (VHL) tumor suppressor gene In vitro and in vivo [93]
Renal cell carcinoma and glioblastoma In vitro [94]
Glutamine oxidation maintains TCA cycle GDH1 Lung cancer In vitro and in vivo [95]
Glioblastoma In vitro [96]
Glutamine contributes to de novo nucleotide synthesis GMPS Prostate cancer Clinical and in vitro [97]
GLS1, PPAT, and their ratio PPAT/GLS1 Lung cancer/potential role in other cancers In vitro and in vivo [98]
PPAT and PAICS Lung cancer Clinical, in vitro, and in ovo [99]
NA Breast cancer with SIRT3 loss In vitro and in vivo [100]
Glutamine contributes to de novo asparagine synthesis ASNS Different cancer cell lines In vitro [101]
Lung cancer Clinical and in vitro [102]
Glutamine synthesis GLUL Pancreatic cancer Clinical, in vitro, and in vivo [103,104]
Glioblastoma Clinical, in vitro, and in vivo [105]
The key metabolic enzymes contributing to Gln metabolism: SLC1A5, neutral amino acid transporter belonging to the solute carrier (SLC) family 1 member 5; SLC6A14, neutral and basic amino acid transporter belonging to SLC family 6 member 14; SLC38A5, neutral amino acid transporter belonging to SLC family 38 member 5; SLC7A5, essential amino acid transporter, neutral amino acid antiporter belonging to SLC family 7 member 5; GLS1, glutaminase (characterized as kidney (also known as brain)-type); GLS2, glutaminase (characterized as liver-type); GCL, glutamate cysteine ligase; GDH1, glutamate dehydrogenase 1; GOT1, glutamate oxaloacetate transaminase 1 (cytosolic); GOT2, glutamate oxaloacetate transaminase 2 (mitochondrial); IDH2, isocitrate dehydrogenase 2 (mitochondrial); GMPS, guanosine monophosphate synthetase; PPAT, phosphoribosyl pyrophosphate amidotransferase; PAICS, phosphoribosylaminoimidazole carboxylase and phosphoribosylaminoimidazole succinocarboxamide synthetase; ASNS, asparagine synthetase; GLUL, glutamate ammonia ligase (also known as glutamine synthase). Other abbreviations: ROS, reactive oxygen species; TCA, tricarboxylic acid; TNBC, triple-negative breast cancer; ER+, estrogen-receptor-positive; K-Ras, Kirsten rat sarcoma virus. SIRT3, sirtuin 3 (mitochondrial). “*” reflects decreased expression of GLS2 supporting growth of hepatocellular cancer.
Many transporters enabling Gln transport but also antiport were found to be altered across different cancers [31]. The role of SLC1A5 in Gln transport mediation was shown in various lung cancers [63]; breast cancers, including triple-negative breast cancer [64]; head and neck cancer [65]; and colorectal cancer [66,67]. A novel variant of SLC1A5 residing in the inner mitochondrial membrane was recently reported as a key contributor to mitochondrial Gln metabolism and metabolic reprogramming in pancreatic cancer, further pinpointing the importance of Gln in mitochondrial metabolism [106]. The interplay between SLC1A5 and SLC7A5, belonging to the SLC7 family of glutamine antiporters, was previously suggested, pointing towards the enhanced entry of essential amino acids via SLC7A5, activated by Gln flux through SLC1A5, as a strategy contributing to cancer progression [66]. In agreement, the critical role of SLC7A5, exporting glutamine in exchange for essential amino acids to meet cellular demands, in colorectal cancer progression and metastasis was recently reported [74]. However, a study by Cormerais et al. showed the capacity of SLC1A5 as an independent transporter promoting lung tumor growth [66], further suggesting that the tumor-specific landscape of Gln transporters adjusts to meet the metabolic demand.
There are also other Gln transporters, including SLC6A14 and SLC38A5, reported to be upregulated in various tumor types (for SLC6A14: cervical cancer [71], colorectal cancer [72], estrogen-receptor-positive breast cancer [73], and pancreatic cancer [68]; for SLC38A5: TNBC [69] and pancreatic cancer [70]) and contributing to the overall Gln pool, which further highlights the importance of this nutrient in cancer cell metabolism. Furthermore, enhanced expression of SLC38A5 was shown to promote glutamine dependence and oxidative stress resistance in breast cancer [107]. The enhanced expression of different Gln transporters could be multifactorial and might depend on the cancer type, stage, and site, as well as the overall metabolic balance and interplay between cancer and its microenvironment.
The enhanced Gln catabolism in cancer cells was shown to be streamed predominantly to supply TCA cycle, glutathione metabolism [9], and synthesis of lipids [56] and NEAAs [108]. The enzyme GLS1, enabling further processing of Gln and its incorporation into various pathways, was reported as significantly upregulated in a broad spectrum of cancers, including breast [75,76,77], prostate [78,79,80], colorectal [81], and lung [82] cancers, and was shown to be crucial for cancer cell fitness. Interestingly, the other glutaminase isoform, namely GLS2, was observed to be suppressed during malignant transformation [109]. Moreover, GLS2 overexpression greatly reduced tumor cell colony formation and inhibited cancer cell proliferation, suggesting its antitumoral function [84,85]. However, GLS2 expression was shown to strongly depend on the microenvironment as enhanced GLS2 expression was reported under hypoxia in pancreatic cancer [83]. Those studies further highlight the importance of environmental factors contributing to metabolic alterations and adding to the complexity of the network.
The glutamate generated under enhanced GLS1 was shown to serve as a substrate for glutathione synthesis to protect cancer cells from oxidative stress, a feature important for their acquired drug resistance [86]. The glutamate cysteine ligase, catalyzing the first and rate-limiting reaction in glutathione synthesis, where glutamate serves as a substrate, was reported as upregulated in cancer and associated with drug resistance [87,110]. Interestingly, in liver cancer, depletion of glutamate cysteine ligase was associated with decreased glutathione level and higher sensitivity to oxidative stress [88], further highlighting the importance of redox balance in cancer.
Nevertheless, glutathione metabolism is not the only pathway supported by glutamate, as its enhanced contribution to the TCA cycle in the form of α-ketoglutarate was found in malignant cells [9,95]. The metabolism of glutamate into α-ketoglutarate is catalyzed by glutamate dehydrogenase 1 (GDH1), glutamate pyruvate transaminase 2 (GPT2), or glutamate oxaloacetate transaminase 1 and 2 (GOT1 (cytoplasmic) and GOT2 (mitochondrial)), which were al previously reported to be overexpressed in cancer [89,90,91,92]. The activity of enzymes catalyzing glutamate metabolism and the metabolic route of the produced α-ketoglutarate depends on cancer type and metabolic condition of the cell, e.g., nutritional status, and is optimized towards proliferative advantages to cancer cells. For instance, the overexpression of GDH1, observed in breast and lung cancer, associated with tumor progression [89,111], can be related to redox homeostasis beyond the role of feeding the TCA cycle intermediates. Jin et al. suggested that α-ketoglutarate formation controls the level of fumarate, which binds and activates the ROS-scavenging enzyme GPx1 [89]. However, different metabolic dependencies were found in pancreatic cancer, in which two subsequent reactions occur: (1) noncanonical pathway, metabolizing glutamate to aspartate and α-ketoglutarate through GOT2, followed by (2) metabolism of aspartate to oxaloacetate by GOT1 [91,112]. Interestingly, the activation of these reactions was shown to increase the NADPH/NADP+ ratio, thereby maintaining ROS balance, and was indicated as a key pathway for coping with oxidative stress in pancreatic cancer [91]. In breast cancer, reversible transamination catalyzed by GPT2 was shown to be channeled towards glutamate rather than α-ketoglutarate and was linked with the enhanced need for building block production in those cells [90]. On the other hand, the metabolic rerouting towards glutamate generation can also support the maintenance of the cellular redox balance. However, not all cancers utilize the given pathway in a similar manner. For instance, in colon cancer cells, GPT2 was shown to enhance α-ketoglutarate synthesis and incorporation as a Gln-delivered carbon source for the TCA cycle under the enhanced Warburg effect [113]. The diverse strategies of deploying enzymatic machinery further point towards dynamic metabolic adjustments optimized for cancer cell demands.
Upon incorporation into the TCA cycle, α-ketoglutarate can undergo different routes while adjusting to the nutritional status, proliferation rate, and environmental stimuli, which all impact the directionality (oxidative vs. reductive) of the TCA cycle [58]. Citrate, required for the formation of lipid structures to cope with cell proliferation, might be deficient in cancer cells exhibiting the Warburg effect and can be supported by reductive carboxylation of α-ketoglutarate [56]. In this process, mitochondrial isocitrate dehydrogenase (IDH2) catalyzes the metabolism of α-ketoglutarate into isocitrate, which can then be isomerized to citrate [94]. The increase in the α-ketoglutarate/citrate ratio signals reductive carboxylation [114], which was shown to also be upregulated under hypoxia [94]. Under hypoxia, along with the enhanced reductive carboxylation, an elevated level of 2-hydroxyglutarate (2HG) was detected [94]. Interestingly, the 2HG was recognized as an epigenetic modifier and potent oncometabolite [115,116].
Although reductive carboxylation is frequently deployed by cancer cells, the oxidative pathway can also be activated in response to various stimuli to resist metabolic stress [95,96]. For instance, Yang et al. showed that inhibition of the mitochondrial pyruvate carrier (MPC), functioning as a pyruvate supply, contributes to GDH activation further rerouting α-ketoglutarate to generate both oxaloacetate and acetyl-CoA, enabling proper TCA cycle function [95]. Similar observations were made in glioblastoma cells, where under impaired glucose metabolism, glutamine was shown to supply the TCA cycle and production of oxaloacetate [96].
Yet, the latest study by Kodama et al. suggested that a shift from the TCA cycle towards nitrogen metabolism is a rather crucial feature of malignant progression in cancer [98]. The authors showed that an increase in phosphoribosyl pyrophosphate amidotransferase (PPAT) and PPAT/GLS ratio is required to trigger nitrogen shift towards nucleotide biosynthesis in lung cancer and could be considered as a universal phenomenon in other cancer types apart from colorectal cancer [98]. Moreover, the authors further suggested, contrary to the previous observations, that GLS1 possesses antitumor effect rather than protumoral activity, which would call into question all previous reports showing, for instance, involvement of Gln in cancer cell redox homeostasis [117] or lipogenesis [56] and aspartate [108] metabolism. Thus, it would be interesting to investigate multiple tumors under various conditions and assess whether the tumor heterogeneity could explain the inconsistency between the recent report and previous studies. It could be hypothesized that a subpopulation of cancer cells would activate the Gln shift towards nucleotide generation whereas the other population of cells would sustain the Gln incorporation into the TCA cycle to ensure either the oxidative or reductive pathway and lipogenesis. In fact, the cancer cell metabolic program depends on various factors modulating cellular adjustments. The metabolic subpopulations of cancer cells inside the tumor, exhibiting different metabolic programs, were shown to exist in symbiosis, contributing to tumor growth [83]. Given that different populations of cancer cells reside in the tumor, it could be reasoned that both metabolic strategies can be utilized in parallel. Moreover, the utilization of Gln to supply nucleotide synthesis in malignant cells was previously described [99,100,118]; however, the actual shift from the TCA cycle and glutathione metabolism was not previously reported.
De novo asparagine synthesis, catalyzed by ASNS, requires glutamine, in contrast to other NEAAs, which can be obtained from glutamate [50]. The expression levels of ASNS vary across different cancers and impact the asparagine level. For instance, in acute lymphoblastic leukemia (ALL), in which low ASNS expression was found, further depletion of asparagine achieved by the introduction of asparaginase (hydrolyzing asparagine to aspartic acid and ammonia) was shown to be lethal in ALL cells and used as a treatment strategy [119,120]. However, cancers with elevated ASNS expression showed resistance to asparaginase [119]. Moreover, elevated ASNS level was shown to promote cancer cell proliferation [101] and was linked to drug resistance [119,121,122].
Finally, the high and diverse tumor demands for glutamine might be challenging to meet under steady state in which Gln production is predominantly supported by the muscle, which further suggests a need for activation of Gln synthesis in some cancers. Indeed, enhanced proliferation rate, along with faulty vascularization in, e.g., pancreatic cancer, result in scarce access to the nutrients and activation of other processes such as extracellular protein scavenging [104]. Furthermore, glutamine synthesis, but not glutaminolysis, was found to be upregulated in pancreatic cancer [103,123] and glioblastoma [124]. GS activity was linked with nucleotide synthesis in glioblastoma under glutamine restriction [105], and de novo synthesized glutamine was shown to be essential for nitrogen anabolic processes in pancreatic cancer [103,125].
In summary, there are multiple metabolic processes activated in cancer cells that rely on Gln. The way in which Gln metabolism is altered in the tumor is multifactorial and depends on cancer type and site, nutrient access, and molecular signaling.

4. Key Components Regulating Glutamine Metabolism

4.1. Environmental Factors

The rapid cancer cell proliferation resulting in tumor tissue growth leads to enhanced oxygen demand, which could not be fulfilled due to distanced existing vasculature or defective angiogenesis causing limited oxygen supply and thus hypoxia (low oxygen level in tumor tissue) [126,127]. The hypoxic microenvironment was shown to be implicated in metabolic reprogramming, further contributing to cancer cell progression and aggressiveness [128,129]. The adaptation to hypoxia can be modulated by hypoxia-inducible factors HIF-1α and HIF-2 α, overexpressed by cancer cells in response to low oxygen levels [130], which in turn induce metabolic genes involved in glycolysis and Gln metabolism [131,132]. It was shown that HIF-1α suppresses glucose-dependent anabolic synthesis with a simultaneous increase in lactate production [131]. Hypoxia-driven alterations result in a reduction in pyruvate entering the TCA cycle [133,134], which could further trigger reprogramming in Gln metabolism. Indeed, enhanced Gln transport and upregulation in the expression of Gln transporters were observed under hypoxia [106,135,136]. The expression of SLC1A5 is controlled and upregulated by HIF-2α [106], whereas SLC38A1 is activated by HIF-1α [135] and presents different strategies to modulate the Gln level in the cell.
Yet, the Gln transported into the cell is further utilized to sustain the needs of hypoxic cells in which oxidative pathway changes more towards reductive carboxylation [136,137]. The utilization of glutamine under hypoxia is enhanced, and glutaminolysis catalyzed by increased activation of both GLS1 and GLS2 was shown to be upregulated [81,83]. The activation of GLS1 was found to be critical for colon cancer growth [81], whereas hypoxia-activated GLS2 was found in pancreatic cancer [83]. Further, the Gln-derived carbon is utilized for the formation of α-ketoglutarate, which undergoes reductive carboxylation catalyzed by IDH2 to form isocitrate followed by isomerization to citrate [94]. Noteworthily, this metabolic upregulation was shown to be associated with increased synthesis of 2HG, recognized as an oncometabolite [94,115]. The generated citrate is further transported into the cytoplasm to generate acetyl-CoA, which serves as a source for fatty acid synthesis [138,139]. The utilization of Gln as a lipid source was recognized as critical for the proliferation of cancer cells under hypoxia in multiple studies [56,92,94,132,136,137]. Interestingly, hypoxia was shown to inhibit carnitine palmitoyltransferase 1A (CPT1A) and thus beta-oxidation, which resulted in the accumulation of lipids in form of stored droplets in clear cell renal cell carcinoma [140]. It could be reasoned that cancer cells might store fatty acids generated under reductive carboxylation for their potential future utilization under changed conditions. Given that reductive carboxylation was shown to be necessary for tumor growth under hypoxia, this pathway could be considered as a promising treatment target [132].
Nevertheless, under hypoxia, enhanced oxidative Gln metabolism resulting in ATP production in addition to glutathione synthesis can be maintained in some cancers despite decreased mitochondrial respiration [135,141,142,143]. In a Myc-inducible human Burkitt lymphoma model, oxidative Gln metabolism supporting the TCA cycle was reported under hypoxia [142], further suggesting the potential involvement of the Myc oncogene in determining the fate of glutamine. The differences in TCA directionality under hypoxia could reflect the cellular demands at a given time and further point towards the metabolic plasticity of cancer cells.
The fate of Gln-derived nitrogen under hypoxia was also investigated. Noteworthily, the utilization of Gln-derived carbon for lipogenesis generates byproducts such as nitrogen (ammonia/ammonium) and oxaloacetate that could not be effectively catabolized under limited oxygen [136] and could be even toxic to the cell [144]. Therefore, under the hypoxic condition, the cancer cell activates an alternative pathway in which Gln-derived nitrogen is incorporated into dihydroorotate. The conversion of dihydroorotate to UMP was shown to be hampered and followed by dihydroorotate accumulation and further excretion [136,145]. The observed accumulation of dihydroorotate indicates a cellular strategy to scavenge potentially toxic nitrogen byproducts resulting from enhanced Gln utilization for lipogenesis under hypoxia.

4.2. Oncogenes and Tumor Suppressors Control Altered Glutamine Metabolism

The metabolic landscape of the tumor is dictated not only by the environmental factors but also by the oncogenes and tumor suppressors. For instance, tumor suppressors such as retinoblastoma protein (Rb), sirtuin 4 (SIRT4), and p53 were linked with dysregulated Gln metabolism in various cancers [53,84,85,146,147,148].
The depletion of Rb was associated with a specific increase in Gln uptake via upregulated SLC1A5 [53,146] and enhanced glutathione formation [146]. Furthermore, under Rb loss, enhanced glutaminolysis mediated by GLS1 and Gln incorporation into the TCA cycle was found and linked with increased ATP production [53], further suggesting activation of the oxidative pathway. Similarly, loss of SIRT4 was also associated with enhanced Gln transport and glutaminolysis contributing to enhanced proliferation and growth of lung and breast cancers [147,148].
The role of p53 has grown lately beyond the canonical activities regulating cell division, as it has been shown to have a function in cellular metabolism involved in responses to metabolic stresses [149]. For instance, p53 was shown to be activated in response to low Gln level [150], in turn promoting expression of different transporters, including SLC1A3 (glutamate/aspartate transporter) [151] and SLC7A3 (enhancing arginine import) [152]. The enhanced SLC1A3 expression, modulated by p53 in response to low Gln level, was shown to rescue cell viability by maintaining TCA cycle activity and promoting de novo synthesis of Gln, glutamate, and nucleotides [151]. The p53-modulated enhancement of different transporters presents strategies enabling cancer cells to facilitate adaptation to Gln deprivation.
The role of oncogenes were identified as critical for controlling Gln uptake and metabolism and the contribution of Gln metabolism to malignant transformation. For instance, oncogene c-Myc coordinates the expression of genes required for Gln catabolism by reprogramming the mitochondrial metabolism towards addiction to glutaminolysis [153]. These changes were linked with suppression of glucose incorporation into the TCA cycle with simultaneous enhancement of GLS1-mediated glutaminolysis [52] and Gln streaming to the TCA cycle [153]. Interestingly, the contribution of c-Myc to Gln synthesis was also reported, suggesting the involvement of c-Myc in both anabolic and catabolic pathways [154] potentially to maximize utilization of Gln by cancer cells. Similarly to the phenotype observed in tumors with Rb loss [53], c-Myc overexpression was also likened with enhanced Gln transport via SLC1A5 [153].
Another oncogene, Kirsten rat sarcoma (K-Ras), was also identified as a modulator of metabolism contributing to malignant transformation and cancer progression [75,119]. K-Ras was found to promote glutaminolysis in a GLS1-dependent manner, and the cancer cell survival was shown to be Gln-dependent [155] Moreover, the K-Ras-transformed cells efficiently utilized both Gln-delivered carbon and nitrogen for the synthesis of building blocks, including amino acids and nucleotides, and for glutathione generation [156]. Such an efficient metabolic program enables sustaining growth on one hand and protecting from oxidative stress on the other hand. Interestingly, in K-Ras pancreatic cancer, noncanonical Gln utilization was reported in which glutamine-derived aspartate is converted into oxaloacetate by GOT1 [91]. This non-canonical pathway serves as a key contributor to generating oxidative capacity of the cell by increasing the NADPH/NADP+ ratio in the series of reactions in which oxaloacetate is converted into pyruvate [91]. Moreover, the cell cycle arrest caused by suppression of glutamine utilization in K-Ras-driven cancer cells was reversed by aspartate [157], further underscoring the significance of this noncanonical pathway.
In summary, both environmental factors and molecular signaling pathways contribute to dysregulated Gln metabolism in cancer cells.

5. Combined Treatment Strategies Targeting Altered Glutamine Metabolism Offer a Promising Cancer Treatment

The extensive research focused on Gln metabolism in health and disease resulted in the identification of potential treatment targets, which were further probed in preclinical and clinical studies. Overall, modulation of Gln metabolism results in inhibition of cancer cell proliferation; however, it is frequently insufficient to induce cancer cell death, potentially due to activation of compensating pathways. Here, we provide an overview of preclinical and clinical studies in which Gln metabolism was targeted, and we further focus on cancer cell responses to such a treatment, as those can serve as a roadmap for rational designing of combined treatment strategies.

5.1. Targeting GLS1 along with Chemotherapy as a Promising Treatment Strategy

The enhanced glutaminolysis orchestrated by GLS1, observed to be essential for multiple cancers [57,117,158,159], was probed as a treatment target with different inhibitors such as bis-2-(5-phenylacetamido-1,2,4-thiadiazol-2-yl)ethyl sulfide (BPTES) [160]; bromo-benzophenanthridinone compound 968 (C.968) [161]; and CB-839, also known as telaglenastat, a product of company Calithera [76]. The inhibition of glutaminolysis was shown to suppress the growth of multiple different cancer cells, including breast [76], renal [93], and liver [162], in vitro and in vivo and was successful in hampering metastatic progression in osteosarcoma [158]. Although GLS1 inhibition leads to suppression of cancer cell proliferation, it is frequently insufficient to trigger cancer cell death, as was shown by us and others, independently of the GLS1 inhibitor used [12,13,93,163,164]. Thus, it can be reasoned that the inactivation of a single metabolic pathway is insufficient to trigger cancer cell death due to potential metabolic compensations. This is in agreement with outcomes from other monotherapies, further suggesting a need for combined treatment strategies [165]. Importantly, we found that GLS1 inhibition resulted in activation of lipid catabolism along with autophagy as a cancer survival mechanism [12], which was confirmed by others [13]. In fact, activated autophagy [166,167] and accelerated lipid catabolism [168] were recognized as pathways supporting cancer cells under nutritional challenges and thus should be considered for cotreatment strategies. Our in vitro experiments showed that simultaneous inhibition of GLS1 along with lipid catabolism or autophagy targeted with trimetazidine and chloroquine, respectively, could be considered as a potential treatment strategy [12]. In fact, this treatment option could be explored given that both trimetazidine and chloroquine are used in clinic. Interestingly, the latest study suggested that metformin, a first-line antidiabetic drug, possesses the ability of GLS1 inhibition [169,170]. However, in contrast to a study in which GLS1 inhibitors were used, metformin treatment resulted in autophagy suppression and apoptosis activation [170]. The differences in cancer cell responses could be explained by the fact that metformin is not a GLS1-specific inhibitor and impacts other pathways relevant for cancer cell growth [171].
Currently, there are a total of 20 registered clinical trials investigating the potential of telaglenastat (CB-839) as an anticancer drug, out of which 8 have been completed (Table 2).
Overall, telaglenastat (CB-839) showed safety and was well tolerated in patients with leukemia and other hematological cancers, as well as those with solid tumors (including clear cell renal cell carcinoma, melanoma, non-small-cell lung cancer, and triple-negative breast cancer) [172,173,174,175]. The telaglenastat treatment was proved to inhibit GLS1 in the patients [175], which is in agreement with the initial preclinical study [76]. However, the observed progression-free survival was moderate [177], and a clinical trial with renal cell carcinoma patients did not achieve the primary endpoint [176]. Noteworthily, in almost all clinical trials, telaglenastat was combined either with chemotherapy or immunotherapy and such a treatment regimen was given to heavily pretreated patients, which could reflect on the trial outcome. Further, the moderate responses to the treatment could be explained by the potential compensation mechanisms activated by the tumor. Thus, in-depth analysis of molecular alterations in response to treatment, preferentially on the metabolic level, should be considered. Moreover, given the complexity of metabolic dependences in the tumor and the symbiosis between the cancer cell populations in the tumor [83], markers enabling better prediction of the responses would be required to guide the selection of treatment strategies. Noteworthily, GLS1 inhibition was shown to lower the level of glutathione, suggesting its impact on the antioxidative capacity of cancer cells and their survival under the treatment [12,76]. Hence, drugs triggering oxidative stress, e.g., doxorubicin, could be considered for combination with telaglenastat (CB-839), given cancer cell dependency on glutathione under doxorubicin treatment [178]. Nevertheless, the preclinical data should be carefully analyzed as doxorubicin-induced site effects related to oxidative stress could strongly affect patient wellbeing [179].

5.2. Targeting Antioxidative Capacity of the Cell by Modulating Glutamine Metabolism as a Potential Treatment Option

The antioxidative capacity of a cancer cell supports its fitness and survival [180]. Therefore, targets such as GOT1, which is a contributor to the antioxidative capacity of pancreatic ductal adenocarcinoma (PDA) [91] but is dispensable in nonmalignant cells, could be considered as an attractive treatment option. GOT1 is required for proliferation and tumor growth, and its inhibition leads to attenuated cancer cell progression through cell cycle blockage [181]. Thus, GOT1 could be considered as a promising and cancer-specific treatment target. Recently, aspulvinone O was reported as a natural inhibitor of GOT1 sensitizing PDA cells to oxidative stress and resulting in growth suppression in vitro and in vivo [182]. Furthermore, a covalent small molecule inhibitor (PF-04859989) of GOT1 was recently tested and showed selective growth inhibition of PDA cell lines [183]. Nevertheless, Kremer et al. showed diverse cell sensitivity to GOT1 inhibition further suggesting activation of cancer cell survival mechanisms in response to the treatment [181], further highlighting the need for taking into account metabolic plasticity. Chronic GOT1 suppression resulted in enhanced cystine import, glutathione synthesis, and lipid antioxidant machinery along with activation of glutathione peroxidase 4 (GPX4), which enabled cancer cells to cope with oxidative stress and survive the treatment [181]. The simultaneous inhibition of GOT1 and any of the pathways activated in response to this inhibition resulted in ferroptosis [181], a form of cell death [184], further suggesting a potential treatment strategy for PDA tumors.
Yet another example of an enzyme contributing to the oxidative capacity of cancer cells is GDH1, which could serve as a potential treatment target [89]. Inhibition of GDH1, either by shRNA or small molecule inhibitor R162, was shown to disrupt redox homeostasis in cancer cells and inhibit their proliferation and tumor growth in vitro and in vivo, causing minimal toxicity [89]. Another study by Jin et al. suggested that GDH1 plays an important role during lung cancer metastasis by contributing to α-ketoglutarate synthesis which in turn modulates the 5′ AMP-activated protein kinase (AMPK) signaling pathway [185]. Our recent study showed an increased level of α-ketoglutarate in breast cancer cell lines with high metastatic potential, further suggesting the importance of this pathway in cancer progression [186]. Targeting GDH1 with R162 attenuated tumor metastasis in a patient-derived xenograft model, offering a potential treatment for lung cancer patients [185]. GDH1 could be also targeted with epigallocatechin gallate (EGCG), a flavonoid isolated from green tea [187], which was shown to inhibit lung cancer cell proliferation and invasion in vitro [188]. Importantly, the redox balance and α-ketoglutarate level disrupted by R162 can be further restored by other metabolic pathways, which should be taken under consideration in future studies targeting GDH1.

5.3. Starving Cancer Cells of Glutamine as a Cotreatment Strategy

The dysregulated Gln metabolism could also be managed by controlling the amount of Gln entering the cell. This can be achieved by targeting SLC1A5, which was shown as a key Gln transporter contributing to cancer cell proliferation [63,64,65,66,106]. Alternatively, Gln antagonists, by inhibiting Gln-utilizing enzymes, limit the performance of Gln-dependent pathways and thus impact cancer function [189,190,191,192,193].
The inhibition of SLC1A5 with the small molecule L-γ-glutamyl-p-nitroanilide (GPNA) leads to glutamine starvation in lung cancer cells, which further results in oxidative stress-mediated autophagy and apoptosis [194]. Given that autophagy is considered as a survival pathway activated by the cancer cell under nutritional stress [167], its activation under inhibited Gln influx should be carefully monitored. Another pharmacological blockade of glutamine flux was recently reported by Schulte et al., who described 2-amino-4-bis(aryloxybenzyl)aminobutanoic acids (V-9302) as selective and potent competitive small molecule antagonists targeting SLC1A5 [195]. The SLC1A5 inhibition resulted in attenuated colon cancer cell growth and proliferation, increased cell death, and increased oxidative stress in vitro and in vivo [195]. Similarly, a study conducted in head and neck squamous cell carcinoma (HNSCC) targeting overexpressed SLC1A5 with V-9302 reported the suppression of the intracellular glutamine level and downstream Gln metabolism, which in turn inhibited cancer growth and proliferation in vitro and in vivo [65]. Furthermore, the authors showed that inhibition of SLC1A5 sensitized HNSCC to cetuximab, thus suggesting optimization of treatment strategy by using metabolic modulation [65]. Interestingly, SLC1A5 was recently recognized as a potential target for leukemia [196], which could be probed with V-9302. Given the sensitized impact of SLC1A5 inhibition on chemotherapy [65], a similar outcome might be expected in leukemia. Nevertheless, this strategy should be carefully investigated given the potentially severe adverse impact of limited Gln access on normal hematopoiesis in patients. Noteworthily, the decreased Gln level under SLC1A5 inhibition could be compensated for by either activation of alternative Gln transporters or Gln synthesis. In fact, enhanced sensitivity to the inhibition of Gln transporters was linked with reduced plasticity associated with the capability of activation of the alternative pathway [197].
Given that Gln can be generated by the cell, Gln antagonists could be considered as an alternative treatment option to molecules inhibiting Gln transporters. Indeed, Gln antagonist 6-diazo-5-oxo-l-norleucine (DON), identified in 1956 [139], was shown to impact Gln-dependent pathways, including synthesis of nucleotides and α-ketoglutarate [190,198], and thus cancer cell growth in vitro and in vivo [139,189,193,198]. Nevertheless, the initially promising clinical development of DON [199,200] was hampered due to its dose-limiting toxicity related to gastrointestinal (GI) mucosa [193]. The latest development aiming to optimize DON to the form of a prodrug circulating in the plasma and activated in the tumor resulted in the successful finding of (DON-based) Gln antagonist prodrugs that provide selective tumor exposure [191,192,201]. A recent study deploying a DON-based prodrug, the compound JHU083, for glutamine blockade in tumor-bearing mice reported the suppression of oxidative and glycolytic metabolism in cancer cells and a significant decrease in tumor volume [191]. Moreover, another DON-based prodrug called sirpiglenastat (DRP-104), a leading candidate of Dracen company, is currently being probed as a single agent and in combination with immunotherapy (atezolizumab) in a clinical trial in patients with advanced solid tumors [202]. Given that sirpiglenastat can simultaneously hamper multiple Gln-dependent pathways, its overall performance in clinical trials might be superior to that of molecules targeting only one enzyme (e.g., telaglenastat). Although the performance of sirpiglenastat as a single agent might be limited due to the plasticity of cancer cells, combined treatment strategies deploying currently suggested immunotherapy could serve as an attractive option for cancer patients.
In summary, the dysregulated Gln metabolism has become an attractive treatment target. Although there are many strategies involving the modulation of Gln metabolism that were proven efficient in preclinical testing, so far, only two molecules, telaglenastat (CB-839), targeting GLS1, and sirpiglenastat (DRP-104), targeting multiple Gln-dependent enzymes, have reached clinical trials. Further effort toward understanding cancer survival mechanisms under the treatment would be needed to optimize combined treatment options for identified targets.

6. Potential Pitfalls and Future Perspectives

Although dysregulated Gln metabolism is an attractive treatment target, which could potentially be applied for different cancers, further optimization of current options is required to achieve better clinical outcomes. Aspects concerning tumor heterogeneity and metabolic plasticity, which both contribute to cancer cell resistance, are critical roadblocks that have to be addressed.
Currently, tumor metabolic status and metabolic dysregulations of cancer cell populations in the tumor and their responses to the stimuli are not considered while designing treatments. However, these factors may be critical given the interplay between different components of the tumor and their metabolic symbiosis [83]. A measurement enabling in-depth characterization of the tumor metabolic processes, ideally in a dynamic manner, would be required for optimized treatment. In fact, metabolomics, which is a strategy for monitoring small molecule composition, offers such a possibility [203]. We used metabolomics to identify cancer compensation mechanisms in response to glutaminolysis inhibition [12] and to identify metabolic pathways activated as a tumor response to doxorubicin treatment [178]. Although such an approach is informative and can result in the identification of optimized treatment strategies, it is not ideal as it provides only a snapshot and not a continuous measurement of the process. Thus, noninvasive approaches enabling continuous and real-time monitoring of metabolic process, which could be incorporated inside the tumors, could be a potential step forward. In fact, nanowire sensors enabling protein and DNA monitoring were recently introduced [204]. Therefore, sensors with the capacity for dynamic metabolic monitoring of the tumors could be envisioned. Such an approach would support the optimization and adjustment of treatment strategies in a dynamic manner.
In conclusion, treatment strategies targeting Gln metabolism constitute an attractive and promising option for cancer patients. However, due to the metabolic plasticity of the tumor, alternative pathways can be activated to overcome the treatment. Thus, identification of the pathways activated to support cancer cell survival under the treatment is critical for further development and optimization of any potential therapy. Given the complexity of the metabolic network, a combined treatment directed towards cancer metabolism could be more successful than monotherapies. Finally, a device enabling continuous monitoring of metabolic processes inside the tumor could support treatment decision making and be a step forward for cancer precision medicine.

Author Contributions

Conceptualization, A.H. and K.S.; writing—original draft preparation, A.H. and K.S.; writing—review and editing, A.H. and K.S.; funding acquisition, A.H. and K.S. All authors have read and agreed to the published version of the manuscript.

Funding

This study was made possible by an NPRP grant (NPRP12S-0205-190042) from the Qatar National Research Fund (a member of Qatar Foundation). The findings achieved herein are solely the responsibility of the author. The funders had no role in the study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Hanahan, D.; Weinberg, R.A. Hallmarks of cancer: The next generation. Cell 2011, 144, 646–674. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  2. Pavlova, N.N.; Thompson, C.B. The Emerging Hallmarks of Cancer Metabolism. Cell Metab. 2016, 23, 27–47. [Google Scholar] [CrossRef] [Green Version]
  3. Warburg, O.; Wind, F.; Negelein, E. The Metabolism of Tumors in the Body. J. Gen. Physiol. 1927, 8, 519–530. [Google Scholar] [CrossRef] [Green Version]
  4. Overgaard-Hansen, K. Metabolic regulation of the adenine nucleotide pool. I. Studies on the transient exhaustion of the adenine nucleotides by glucose in ehrlich ascites tumor cells. BBA-Gen. Subj. 1965, 104, 330–347. [Google Scholar] [CrossRef]
  5. Beloribi-Djefaflia, S.; Vasseur, S.; Guillaumond, F. Lipid metabolic reprogramming in cancer cells. Oncogenesis 2016, 5, e189. [Google Scholar] [CrossRef] [PubMed]
  6. Lieu, E.L.; Nguyen, T.; Rhyne, S.; Kim, J. Amino acids in cancer. Exp. Mol. Med. 2020, 52, 15. [Google Scholar] [CrossRef]
  7. Mattaini, K.R.; Sullivan, M.R.; Vander Heiden, M.G. The importance of serine metabolism in cancer. J. Cell Biol. 2016, 214, 249–257. [Google Scholar] [CrossRef] [Green Version]
  8. Fan, J.; Ye, J.; Kamphorst, J.J.; Shlomi, T.; Thompson, C.B.; Rabinowitz, J.D. Quantitative flux analysis reveals folate-dependent NADPH production. Nature 2014, 510, 298–302. [Google Scholar] [CrossRef] [Green Version]
  9. Tompkins, S.C.; Sheldon, R.D.; Rauckhorst, A.J.; Noterman, M.F.; Solst, S.R.; Buchanan, J.L.; Mapuskar, K.A.; Pewa, A.D.; Gray, L.R.; Oonthonpan, L.; et al. Disrupting Mitochondrial Pyruvate Uptake Directs Glutamine into the TCA Cycle away from Glutathione Synthesis and Impairs Hepatocellular Tumorigenesis. Cell Rep. 2019, 28, 2608–2619.e6. [Google Scholar] [CrossRef]
  10. Farber, S. Some observations on the effect of folic acid antagonists on acute. Blood 1949, 4, 160–167. [Google Scholar] [CrossRef] [Green Version]
  11. Miller, D.R. A tribute to Sidney Farber—The father of modern chemotherapy. Br. J. Haematol. 2006, 134, 20–26. [Google Scholar] [CrossRef]
  12. Halama, A.; Kulinski, M.; Dib, S.S.; Zaghlool, S.B.; Siveen, K.S.; Iskandarani, A.; Zierer, J.; Prabhu, K.S.; Satheesh, N.J.; Bhagwat, A.M.; et al. Accelerated lipid catabolism and autophagy are cancer survival mechanisms under inhibited glutaminolysis. Cancer Lett. 2018, 430, 133–147. [Google Scholar] [CrossRef]
  13. Dos Reis, L.M.; Adamoski, D.; Souza, R.O.O.; Ascenção, C.F.R.; De Oliveira, K.R.S.; Corrêa-Da-Silva, F.; De Sá Patroni, F.M.; Dias, M.M.; Consonni, S.R.; De Moraes-Vieira, P.M.M.; et al. Dual inhibition of glutaminase and carnitine palmitoyltransferase decreases growth and migration of glutaminase inhibition-resistant triple-negative breast cancer cells. J. Biol. Chem. 2019, 294, 9342–9357. [Google Scholar] [CrossRef]
  14. Pallett, L.J.; Dimeloe, S.; Sinclair, L.V.; Byrne, A.J.; Schurich, A. A glutamine “tug-of-war”: Targets to manipulate glutamine metabolism for cancer immunotherapy. Immunother. Adv. 2021, 1. [Google Scholar] [CrossRef]
  15. Krebs, H.A. Metabolism of amino-acids. Biochem. J. 1935, 29, 1951–1969. [Google Scholar] [CrossRef] [PubMed]
  16. Lacey, J.M.; Wilmore, D.W. Is glutamine a conditionally essential amino acid? Nutr. Rev. 1990, 48, 297–309. [Google Scholar] [CrossRef] [PubMed]
  17. Stein, W.H.; Moore, S. The free amino acids of human blood plasma. J. Biol. Chem. 1954, 211, 915–926. [Google Scholar] [CrossRef]
  18. Curthoys, N.P.; Weiss, R.F. Regulation of renal ammoniagenesis. Subcellular localization of rat kidney glutaminase isoenzymes. J. Biol. Chem. 1974, 249, 3261–3266. [Google Scholar] [CrossRef]
  19. Muhlbacher, F.; Kapadia, C.R.; Colpoys, M.F. Effects of glucocorticoids on glutamine metabolism in skeletal muscle. Am. J. Physiol. 1984, 247, E75–E83. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  20. Hanson, P.J.; Parsons, D.S. Glutamine and glucose as fuels for the fed, fasted and acidotic rat small intestine [proceedings]. J. Physiol. 1977, 268, 13P–14P. [Google Scholar]
  21. Porteous, J.W. Glutamate, glutamine, aspartate, asparagine, glucose and ketone-body metabolism in chick intestinal brush-border cells. Biochem. J. 1980, 188, 619–632. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  22. Stumvoll, M.; Perriello, G.; Meyer, C.; Gerich, J. Role of glutamine in human carbohydrate metabolism in kidney and other tissues. Kidney Int. 1999, 55, 778–792. [Google Scholar] [CrossRef] [Green Version]
  23. Bach, S.J.; Smith, M. Role of glutamine in urea synthesis. Nature 1955, 176, 1126–1127. [Google Scholar] [CrossRef]
  24. Davies, B.M.A.; Yudkin, J. Role of glutaminase in the production of urinary ammonia. Nature 1951, 167, 117. [Google Scholar] [CrossRef]
  25. Kida, K.; Nakajo, S.; Kamiya, F.; Toyama, Y.; Nishio, T.; Nakagawa, H. Renal net glucose release in vivo and its contribution to blood glucose in rats. J. Clin. Investig. 1978, 62, 721–726. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  26. Curi, T.C.P.; De Melo, M.P.; De Azevedo, R.B.; Curi, R. Glutamine utilization by rat neutrophils. Biochem. Soc. Trans. 1997, 25. [Google Scholar] [CrossRef] [Green Version]
  27. Curi, R.; Newsholme, P.; Newsholme, E.A. Intracellular distribution of some enzymes of the glutamine utilisation pathway in rat lymphocytes. Biochem. Biophys. Res. Commun. 1986, 138, 318–322. [Google Scholar] [CrossRef]
  28. Newsholme, P.; Curi, R.; Gordon, S.; Newsholme, E.A. Metabolism of glucose, glutamine, long-chain fatty acids and ketone bodies by murine macrophages. Biochem. J. 1986, 239, 121–125. [Google Scholar] [CrossRef]
  29. Ardawi, M.S.; Newsholme, E.A. Metabolism in lymphocytes and its importance in the immune response. Essays Biochem. 1985, 21, 1–44. [Google Scholar] [PubMed]
  30. Newsholme, E.A.; Crabtree, B.; Ardawi, M.S.M. Glutamine metabolism in lymphocytes: Its biochemical, physiological and clinical importance. Q. J. Exp. Physiol. 1985, 70, 473–489. [Google Scholar] [CrossRef]
  31. Bhutia, Y.D.; Ganapathy, V. Glutamine transporters in mammalian cells and their functions in physiology and cancer. Biochim. Biophys. Acta 2016, 1863, 2531–2539. [Google Scholar] [CrossRef] [PubMed]
  32. Hartman, S.C.; Buchanan, J.M. Nucleic acids, purines, pyrimidines (nucleotide synthesis). Annu. Rev. Biochem. 1959, 28, 365–410. [Google Scholar] [CrossRef]
  33. Hartman, S.C.; Buchanan, J.M. Biosynthesis of the purines. XXI. 5-Phosphoribosylpyrophosphate amidotransferase. Available online: https://pubmed.ncbi.nlm.nih.gov/13563519/ (accessed on 29 November 2021).
  34. Anand, R.; Hoskins, A.A.; Bennett, E.M.; Sintchak, M.D.; Stubbe, J.A.; Ealick, S.E. A model for the Bacillus subtilis formylglycinamide ribonucleotide amidotransferase multiprotein complex. Biochemistry 2004, 43, 10343–10352. [Google Scholar] [CrossRef]
  35. Kusumi, T.; Tsuda, M.; Katusunuma, T.; Yamamura, M. Dual inhibitory effect of bredinin. Cell Biochem. Funct. 1989, 7, 201–204. [Google Scholar] [CrossRef]
  36. Holden, H.M.; Thoden, J.B.; Raushel, F.M. Carbamoyl phosphate synthetase: An amazing biochemical odyssey from substrate to product. Cell. Mol. Life Sci. 1999, 56, 507–522. [Google Scholar] [CrossRef]
  37. Weng, M.; Zalkin, H. Structural role for a conserved region in the CTP synthetase glutamine amide transfer domain. J. Bacteriol. 1987, 169, 3023–3028. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  38. Exton, J.H. Gluconeogenesis. Metabolism. 1972, 21, 945–990. [Google Scholar] [CrossRef]
  39. Nurjhan, N.; Bucci, A.; Perriello, G.; Stumvoll, M.; Dailey, G.; Bier, D.M.; Toft, I.; Jenssen, T.G.; Gerich, J.E. Glutamine: A major gluconeogenic precursor and vehicle for interorgan carbon transport in man. J. Clin. Investig. 1995, 95, 272–277. [Google Scholar] [CrossRef] [PubMed]
  40. Fahien, L.A.; Wiggert, B.O.; Cohen, P.P. Crystallization and Kinetic Properties of Glutamate Dehydrogenase From. J. Biol. Chem. 1965, 240, 1083–1090. [Google Scholar] [CrossRef]
  41. Kovacevic, Z.; McGivan, J.D. Mitochondrial metabolism of glutamine and glutamate and its physiological significance. Physiol. Rev. 1983, 63, 547–605. [Google Scholar] [CrossRef] [PubMed]
  42. Fahien, L.A.; Kmiotek, E.H.; MacDonald, M.J.; Fibich, B.; Mandic, M. Regulation of malate dehydrogenase activity by glutamate, citrate, alpha-ketoglutarate, and multienzyme interaction. J. Biol. Chem. 1988, 263, 10687–10697. [Google Scholar] [CrossRef]
  43. Stark, R.; Pasquel, F.; Turcu, A.; Pongratz, R.L.; Roden, M.; Cline, G.W.; Shulman, G.I.; Kibbey, R.G. Phosphoenolpyruvate Cycling via Mitochondrial Phosphoenolpyruvate Carboxykinase Links Anaplerosis and Mitochondrial GTP with Insulin Secretion. J. Biol. Chem. 2009, 284, 26578. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  44. Stark, R.; Kibbey, R.G. The mitochondrial isoform of phosphoenolpyruvate carboxykinase (PEPCK-M) and glucose homeostasis: Has it been overlooked? Biochim. Biophys. Acta 2014, 1840, 1313. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  45. Gupta, R.K.; Oesterling, R.M.; Mildvan, A.S. Dual divalent cation requirement for activation of pyruvate kinase; essential roles of both enzyme- and nucleotide-bound metal ions. Biochemistry 1976, 15, 2881–2887. [Google Scholar] [CrossRef]
  46. Yoo, H.; Antoniewicz, M.R.; Stephanopoulos, G.; Kelleher, J.K. Quantifying reductive carboxylation flux of glutamine to lipid in a brown adipocyte cell line. J. Biol. Chem. 2008, 283, 20621–20627. [Google Scholar] [CrossRef] [Green Version]
  47. Masterson, E.; Whikehart, D.R.; Chader, G.J. Glucose oxidation in the chick cornea: Effect of diamide on the pentose shunt. Investig. Ophthalmol. Vis. Sci. 1978, 17, 449–454. [Google Scholar]
  48. Snoke, J.E.; Bloch, K. Formation and utilization of gamma-glutamylcysteine in glutathione synthesis. J. Biol. Chem. 1952, 199, 407–414. [Google Scholar] [CrossRef]
  49. Schowen, R.L. Principles of biochemistry 2nd ed. (Lehninger, Albert L.; Nelson, David L.; Cox, Michael M.). J. Chem. Educ. 1993, 70, A223. [Google Scholar] [CrossRef] [Green Version]
  50. Patterson, M.K.; Orr, G.R. Asparagine biosynthesis by the Novikoff Hepatoma isolation, purification, property, and mechanism studies of the enzyme system. J. Biol. Chem. 1968, 243, 376–380. [Google Scholar] [CrossRef]
  51. Roberts, E.; Frankel, S. Free Amino Acids in Normal and Neoplastic Tissues of Mice as Studied by Paper Chromatography. Cancer Res. 1949, 9, 645–648. [Google Scholar]
  52. Gao, P.; Tchernyshyov, I.; Chang, T.C.; Lee, Y.S.; Kita, K.; Ochi, T.; Zeller, K.I.; De Marzo, A.M.; Van Eyk, J.E.; Mendell, J.T.; et al. c-Myc suppression of miR-23a/b enhances mitochondrial glutaminase expression and glutamine metabolism. Nature 2009, 458, 762–765. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  53. Reynolds, M.R.; Lane, A.N.; Robertson, B.; Kemp, S.; Liu, Y.; Hill, B.G.; Dean, D.C.; Clem, B.F. Control of glutamine metabolism by the tumor suppressor Rb. Oncogene 2014, 33, 556–566. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  54. Kubota, A.; Meguid, M.M.; Hitch, D.C. Amino acid profiles correlate diagnostically with organ site in three kinds of malignant tumors. Cancer 1992, 69, 2343–2348. [Google Scholar] [CrossRef]
  55. Reardon, M.A.; Weber, G. Increased synthesis of carbamoyl-phosphate synthase II (EC 6.3.5.5) in hepatoma 3924A. Cancer Res. 1986, 46, 3673–3676. [Google Scholar] [PubMed]
  56. Mullen, A.R.; Wheaton, W.W.; Jin, E.S.; Chen, P.H.; Sullivan, L.B.; Cheng, T.; Yang, Y.; Linehan, W.M.; Chandel, N.S.; Deberardinis, R.J. Reductive carboxylation supports growth in tumor cells with defective mitochondria. Nature 2012, 481, 385. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  57. Daemen, A.; Liu, B.; Song, K.; Kwong, M.; Gao, M.; Hong, R.; Nannini, M.; Peterson, D.; Liederer, B.M.; de la Cruz, C.; et al. Pan-Cancer Metabolic Signature Predicts Co-Dependency on Glutaminase and De Novo Glutathione Synthesis Linked to a High-Mesenchymal Cell State. Cell Metab. 2018, 28, 383–399.e9. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  58. DeBerardinis, R.J.; Mancuso, A.; Daikhin, E.; Nissim, I.; Yudkoff, M.; Wehrli, S.; Thompson, C.B. Beyond aerobic glycolysis: Transformed cells can engage in glutamine metabolism that exceeds the requirement for protein and nucleotide synthesis. Proc. Natl. Acad. Sci. USA 2007, 104, 19345–19350. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  59. Miyagi, Y.; Higashiyama, M.; Gochi, A.; Akaike, M.; Ishikawa, T.; Miura, T.; Saruki, N.; Bando, E.; Kimura, H.; Imamura, F.; et al. Plasma free amino acid profiling of five types of cancer patients and its application for early detection. PLoS ONE 2011, 6, e24143. [Google Scholar] [CrossRef] [Green Version]
  60. Norton, J.A.; Gorschboth, C.M.; Wesley, R.A.; Burt, M.E.; Brennan, M.F. Fasting plasma amino acid levels in cancer patients. Cancer 1985, 56, 1181–1186. [Google Scholar] [CrossRef]
  61. Venneti, S.; Dunphy, M.P.; Zhang, H.; Pitter, K.L.; Zanzonico, P.; Campos, C.; Carlin, S.D.; La Rocca, G.; Lyashchenko, S.; Ploessl, K.; et al. Glutamine-based PET imaging facilitates enhanced metabolic evaluation of gliomas in vivo. Sci. Transl. Med. 2015, 7, 274ra17. [Google Scholar] [CrossRef] [Green Version]
  62. Grkovski, M.; Goel, R.; Krebs, S.; Staton, K.D.; Harding, J.J.; Mellinghoff, I.K.; Humm, J.L.; Dunphy, M.P.S. Pharmacokinetic Assessment of 18 F-(2 S, 4 R)-4-Fluoroglutamine in Patients with Cancer. J. Nucl. Med. 2020, 61, 357–366. [Google Scholar] [CrossRef] [PubMed]
  63. Hassanein, M.; Hoeksema, M.D.; Shiota, M.; Qian, J.; Harris, B.K.; Chen, H.; Clark, J.E.; Alborn, W.E.; Eisenberg, R.; Massion, P.P. SLC1A5 mediates glutamine transport required for lung cancer cell growth and survival. Clin. Cancer Res. 2013, 19, 560–570. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  64. Van Geldermalsen, M.; Wang, Q.; Nagarajah, R.; Marshall, A.D.; Thoeng, A.; Gao, D.; Ritchie, W.; Feng, Y.; Bailey, C.G.; Deng, N.; et al. ASCT2/SLC1A5 controls glutamine uptake and tumour growth in triple-negative basal-like breast cancer. Oncogene 2016, 35, 3201–3208. [Google Scholar] [CrossRef] [Green Version]
  65. Zhang, Z.; Liu, R.; Shuai, Y.; Huang, Y.; Jin, R.; Wang, X.; Luo, J. ASCT2 (SLC1A5)-dependent glutamine uptake is involved in the progression of head and neck squamous cell carcinoma. Br. J. Cancer 2020, 122, 82–93. [Google Scholar] [CrossRef] [PubMed]
  66. Cormerais, Y.; Massard, P.A.; Vucetic, M.; Giuliano, S.; Tambutté, E.; Durivault, J.; Vial, V.; Endou, H.; Wempe, M.F.; Parks, S.K.; et al. The glutamine transporter ASCT2 (SLC1A5) promotes tumor growth independently of the amino acid transporter LAT1 (SLC7A5). J. Biol. Chem. 2018, 293, 2877–2887. [Google Scholar] [CrossRef] [Green Version]
  67. Wang, R.; Xiang, W.; Xu, Y.; Han, L.; Li, Q.; Dai, W.; Cai, G. Enhanced glutamine utilization mediated by SLC1A5 and GPT2 is an essential metabolic feature of colorectal signet ring cell carcinoma with therapeutic potential. Ann. Transl. Med. 2020, 8, 302. [Google Scholar] [CrossRef]
  68. Coothankandaswamy, V.; Cao, S.; Xu, Y.; Prasad, P.D.; Singh, P.K.; Reynolds, C.P.; Yang, S.; Ogura, J.; Ganapathy, V.; Bhutia, Y.D. Amino acid transporter SLC6A14 is a novel and effective drug target for pancreatic cancer. Br. J. Pharmacol. 2016, 173, 3292–3306. [Google Scholar] [CrossRef] [Green Version]
  69. Ramachandran, S.; Sennoune, S.; Sharma, M.; Thangaraju, M.; Suresh, V.; Sneigowski, T.; Bhutia, Y.D.; Pruitt, K.; Ganapathy, V. Expression and function of SLC38A5, an amino acid-coupled Na+/H+ exchanger, in triple-negative breast cancer and its relevance to macropinocytosis. Biochem. J. 2021, 478, 3957–3976. [Google Scholar] [CrossRef]
  70. Smith, D.K.; Kates, L.; Durinck, S.; Patel, N.; Stawiski, E.W.; Kljavin, N.; Foreman, O.; Sipos, B.; Solloway, M.J.; Allan, B.B.; et al. Elevated Serum Amino Acids Induce a Subpopulation of Alpha Cells to Initiate Pancreatic Neuroendocrine Tumor Formation. Cell reports. Med. 2020, 1, 100058. [Google Scholar] [CrossRef]
  71. Gupta, N.; Prasad, P.D.; Ghamande, S.; Moore-Martin, P.; Herdman, A.V.; Martindale, R.G.; Podolsky, R.; Mager, S.; Ganapathy, M.E.; Ganapathy, V. Up-regulation of the amino acid transporter ATB(0,+) (SLC6A14) in carcinoma of the cervix. Gynecol. Oncol. 2006, 100, 8–13. [Google Scholar] [CrossRef]
  72. Gupta, N.; Miyauchi, S.; Martindale, R.G.; Herdman, A.V.; Podolsky, R.; Miyake, K.; Mager, S.; Prasad, P.D.; Ganapathy, M.E.; Ganapathy, V. Upregulation of the amino acid transporter ATB0,+ (SLC6A14) in colorectal cancer and metastasis in humans. Biochim. Biophys. Acta 2005, 1741, 215–223. [Google Scholar] [CrossRef] [Green Version]
  73. Karunakaran, S.; Ramachandran, S.; Coothankandaswamy, V.; Elangovan, S.; Babu, E.; Periyasamy-Thandavan, S.; Gurav, A.; Gnanaprakasam, J.P.; Singh, N.; Schoenlein, P.V.; et al. SLC6A14 (ATB0,+) protein, a highly concentrative and broad specific amino acid transporter, is a novel and effective drug target for treatment of estrogen receptor-positive breast cancer. J. Biol. Chem. 2011, 286, 31830–31838. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  74. Najumudeen, A.K.; Ceteci, F.; Fey, S.K.; Hamm, G.; Steven, R.T.; Hall, H.; Nikula, C.J.; Dexter, A.; Murta, T.; Race, A.M.; et al. The amino acid transporter SLC7A5 is required for efficient growth of KRAS-mutant colorectal cancer. Nat. Genet. 2021, 53, 16–26. [Google Scholar] [CrossRef] [PubMed]
  75. Vidula, N.; Yau, C.; Rugo, H.S. Glutaminase (GLS) expression in primary breast cancer (BC): Correlations with clinical and tumor characteristics. J. Clin. Oncol. 2019, 37, 558. [Google Scholar] [CrossRef]
  76. Gross, M.I.; Demo, S.D.; Dennison, J.B.; Chen, L.; Chernov-Rogan, T.; Goyal, B.; Janes, J.R.; Laidig, G.J.; Lewis, E.R.; Li, J.; et al. Antitumor activity of the glutaminase inhibitor CB-839 in triple-negative breast cancer. Mol. Cancer Ther. 2014, 13, 890–901. [Google Scholar] [CrossRef] [Green Version]
  77. Kim, S.; Kim, D.H.; Jung, W.H.; Koo, J.S. Expression of glutamine metabolism-related proteins according to molecular subtype of breast cancer. Endocr. Relat. Cancer 2013, 20, 339–348. [Google Scholar] [CrossRef] [Green Version]
  78. Myint, Z.W.; Sun, R.C.; Hensley, P.J.; James, A.C.; Wang, P.; Strup, S.E.; McDonald, R.J.; Yan, D.; St. Clair, W.H.; Allison, D.B. Evaluation of Glutaminase Expression in Prostate Adenocarcinoma and Correlation with Clinicopathologic Parameters. Cancers 2021, 13, 2157. [Google Scholar] [CrossRef]
  79. Pan, T.; Gao, L.; Wu, G.; Shen, G.; Xie, S.; Wen, H.; Yang, J.; Zhou, Y.; Tu, Z.; Qian, W. Elevated expression of glutaminase confers glucose utilization via glutaminolysis in prostate cancer. Biochem. Biophys. Res. Commun. 2015, 456, 452–458. [Google Scholar] [CrossRef]
  80. Dorai, T.; Dorai, B.; Pinto, J.T.; Grasso, M.; Cooper, A.J.L. High levels of glutaminase ii pathway enzymes in normal and cancerous prostate suggest a role in ‘glutamine addiction ’. Biomolecules 2020, 10, 2. [Google Scholar] [CrossRef] [Green Version]
  81. Xiang, L.; Mou, J.; Shao, B.; Wei, Y.; Liang, H.; Takano, N.; Semenza, G.L.; Xie, G. Glutaminase 1 expression in colorectal cancer cells is induced by hypoxia and required for tumor growth, invasion, and metastatic colonization. Cell Death Dis. 2019, 10, 40. [Google Scholar] [CrossRef] [Green Version]
  82. Lee, J.S.; Kang, J.H.; Lee, S.H.; Hong, D.; Son, J.; Hong, K.M.; Song, J.; Kim, S.Y. Dual targeting of glutaminase 1 and thymidylate synthase elicits death synergistically in NSCLC. Cell Death Dis. 2016, 7, e2511. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  83. Guillaumond, F.; Leca, J.; Olivares, O.; Lavaut, M.N.; Vidal, N.; Berthezène, P.; Dusetti, N.J.; Loncle, C.; Calvo, E.; Turrini, O.; et al. Strengthened glycolysis under hypoxia supports tumor symbiosis and hexosamine biosynthesis in pancreatic adenocarcinoma. Proc. Natl. Acad. Sci. USA 2013, 110, 3919–3924. [Google Scholar] [CrossRef] [Green Version]
  84. Suzuki, S.; Tanaka, T.; Poyurovsky, M.V.; Nagano, H.; Mayama, T.; Ohkubo, S.; Lokshin, M.; Hosokawa, H.; Nakayama, T.; Suzuki, Y.; et al. Phosphate-activated glutaminase (GLS2), a p53-inducible regulator of glutamine metabolism and reactive oxygen species. Proc. Natl. Acad. Sci. USA 2010, 107, 7461. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  85. Hu, W.; Zhang, C.; Wu, R.; Sun, Y.; Levine, A.; Feng, Z. Glutaminase 2, a novel p53 target gene regulating energy metabolism and antioxidant function. Proc. Natl. Acad. Sci. USA 2010, 107, 7455–7460. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  86. Lien, E.C.; Lyssiotis, C.A.; Juvekar, A.; Hu, H.; Asara, J.M.; Cantley, L.C.; Toker, A. Glutathione biosynthesis is a metabolic vulnerability in PI(3)K/Akt-driven breast cancer. Nat. Cell Biol. 2016, 18, 572–578. [Google Scholar] [CrossRef] [Green Version]
  87. Fujimori, S.; Abe, Y.; Nishi, M.; Hamamoto, A.; Inoue, Y.; Ohnishi, Y.; Nishime, C.; Matsumoto, H.; Yamazaki, H.; Kijima, H.; et al. The subunits of glutamate cysteine ligase enhance cisplatin resistance in human non-small cell lung cancer xenografts in vivo. Int. J. Oncol. 2004, 25, 413–418. [Google Scholar] [CrossRef]
  88. Anderton, B.; Camarda, R.; Balakrishnan, S.; Balakrishnan, A.; Kohnz, R.A.; Lim, L.; Evason, K.J.; Momcilovic, O.; Kruttwig, K.; Huang, Q.; et al. MYC-driven inhibition of the glutamate-cysteine ligase promotes glutathione depletion in liver cancer. EMBO Rep. 2017, 18, 569–585. [Google Scholar] [CrossRef]
  89. Jin, L.; Li, D.; Alesi, G.N.; Fan, J.; Kang, H.B.; Lu, Z.; Boggon, T.J.; Jin, P.; Yi, H.; Wright, E.R.; et al. Glutamate dehydrogenase 1 signals through antioxidant glutathione peroxidase 1 to regulate redox homeostasis and tumor growth. Cancer Cell 2015, 27, 257–270. [Google Scholar] [CrossRef] [Green Version]
  90. Cao, Y.; Lin, S.H.; Wang, Y.; Chin, Y.E.; Kang, L.; Mi, J. Glutamic Pyruvate Transaminase GPT2 Promotes Tumorigenesis of Breast Cancer Cells by Activating Sonic Hedgehog Signaling. Theranostics 2017, 7, 3021–3033. [Google Scholar] [CrossRef] [Green Version]
  91. Son, J.; Lyssiotis, C.A.; Ying, H.; Wang, X.; Hua, S.; Ligorio, M.; Perera, R.M.; Ferrone, C.R.; Mullarky, E.; Shyh-Chang, N.; et al. Glutamine supports pancreatic cancer growth through a KRAS-regulated metabolic pathway. Nature 2013, 496, 101–105. [Google Scholar] [CrossRef]
  92. Yang, S.; Hwang, S.; Kim, M.; Seo, S.B.; Lee, J.H.; Jeong, S.M. Mitochondrial glutamine metabolism via GOT2 supports pancreatic cancer growth through senescence inhibition. Cell Death Dis. 2018, 9, 55. [Google Scholar] [CrossRef] [PubMed]
  93. Okazaki, A.; Gameiro, P.A.; Christodoulou, D.; Laviollette, L.; Schneider, M.; Chaves, F.; Stemmer-Rachamimov, A.; Yazinski, S.A.; Lee, R.; Stephanopoulos, G.; et al. Glutaminase and poly(ADP-ribose) polymerase inhibitors suppress pyrimidine synthesis and VHL-deficient renal cancers. J. Clin. Investig. 2017, 127, 1631–1645. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  94. Wise, D.R.; Ward, P.S.; Shay, J.E.S.; Cross, J.R.; Gruber, J.J.; Sachdeva, U.M.; Platt, J.M.; DeMatteo, R.G.; Simon, M.C.; Thompson, C.B. Hypoxia promotes isocitrate dehydrogenase-dependent carboxylation of α-ketoglutarate to citrate to support cell growth and viability. Proc. Natl. Acad. Sci. USA 2011, 108, 19611–19616. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  95. Yang, C.; Ko, B.; Hensley, C.T.; Jiang, L.; Wasti, A.T.; Kim, J.; Sudderth, J.; Calvaruso, M.A.; Lumata, L.; Mitsche, M.; et al. Glutamine oxidation maintains the TCA cycle and cell survival during impaired mitochondrial pyruvate transport. Mol. Cell 2014, 56, 414–424. [Google Scholar] [CrossRef] [Green Version]
  96. Chendong, Y.; Sudderth, J.; Tuyen, D.; Bachoo, R.G.; McDonald, J.G.; DeBerardinis, R.J. Glioblastoma cells require glutamate dehydrogenase to survive impairments of glucose metabolism or Akt signaling. Cancer Res. 2009, 69, 7986–7993. [Google Scholar]
  97. Wang, Q.; Guan, Y.F.; Hancock, S.E.; Wahi, K.; van Geldermalsen, M.; Zhang, B.K.; Pang, A.; Nagarajah, R.; Mak, B.; Freidman, N.; et al. Inhibition of guanosine monophosphate synthetase (GMPS) blocks glutamine metabolism and prostate cancer growth. J. Pathol. 2021, 254, 135–146. [Google Scholar] [CrossRef]
  98. Kodama, M.; Oshikawa, K.; Shimizu, H.; Yoshioka, S.; Takahashi, M.; Izumi, Y.; Bamba, T.; Tateishi, C.; Tomonaga, T.; Matsumoto, M.; et al. A shift in glutamine nitrogen metabolism contributes to the malignant progression of cancer. Nat. Commun. 2020, 11, 1320. [Google Scholar] [CrossRef]
  99. Goswami, M.T.; Chen, G.; Chakravarthi, B.V.S.K.; Pathi, S.S.; Anand, S.K.; Carskadon, S.L.; Giordano, T.J.; Chinnaiyan, A.M.; Thomas, D.G.; Palanisamy, N.; et al. Role and regulation of coordinately expressed de novo purine biosynthetic enzymes PPAT and PAICS in lung cancer. Oncotarget 2015, 6, 23445–23461. [Google Scholar] [CrossRef] [Green Version]
  100. Gonzalez Herrera, K.N.; Zaganjor, E.; Ishikawa, Y.; Spinelli, J.B.; Yoon, H.; Lin, J.R.; Satterstrom, F.K.; Ringel, A.; Mulei, S.; Souza, A.; et al. Small-Molecule Screen Identifies De Novo Nucleotide Synthesis as a Vulnerability of Cells Lacking SIRT3. Cell Rep. 2018, 22, 1945–1955. [Google Scholar] [CrossRef] [Green Version]
  101. Krall, A.S.; Xu, S.; Graeber, T.G.; Braas, D.; Christofk, H.R. Asparagine promotes cancer cell proliferation through use as an amino acid exchange factor. Nat. Commun. 2016, 7, 11457. [Google Scholar] [CrossRef] [Green Version]
  102. Xu, Y.; Lv, F.; Zhu, X.; Wu, Y.; Shen, X. Loss of asparagine synthetase suppresses the growth of human lung cancer cells by arresting cell cycle at G0/G1 phase. Cancer Gene Ther. 2016, 23, 287–294. [Google Scholar] [CrossRef]
  103. Bott, A.J.; Shen, J.; Tonelli, C.; Zhan, L.; Sivaram, N.; Jiang, Y.P.; Yu, X.; Bhatt, V.; Chiles, E.; Zhong, H.; et al. Glutamine Anabolism Plays a Critical Role in Pancreatic Cancer by Coupling Carbon and Nitrogen Metabolism. Cell Rep. 2019, 29, 1287–1298.e6. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  104. Kamphorst, J.J.; Nofal, M.; Commisso, C.; Hackett, S.R.; Lu, W.; Grabocka, E.; Vander Heiden, M.G.; Miller, G.; Drebin, J.A.; Bar-Sagi, D.; et al. Human pancreatic cancer tumors are nutrient poor and tumor cells actively scavenge extracellular protein. Cancer Res. 2015, 75, 544–553. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  105. Tardito, S.; Oudin, A.; Ahmed, S.U.; Fack, F.; Keunen, O.; Zheng, L.; Miletic, H.; Sakariassen, P.Ø.; Weinstock, A.; Wagner, A.; et al. Glutamine synthetase activity fuels nucleotide biosynthesis and supports growth of glutamine-restricted glioblastoma. Nat. Cell Biol. 2015, 17, 1556–1568. [Google Scholar] [CrossRef] [Green Version]
  106. Yoo, H.C.; Park, S.J.; Nam, M.; Kang, J.; Kim, K.; Yeo, J.H.; Kim, J.K.; Heo, Y.; Lee, H.S.; Lee, M.Y.; et al. A Variant of SLC1A5 Is a Mitochondrial Glutamine Transporter for Metabolic Reprogramming in Cancer Cells. Cell Metab. 2020, 31, 267–283.e12. [Google Scholar] [CrossRef] [PubMed]
  107. Morotti, M.; Zois, C.E.; El-Ansari, R.; Craze, M.L.; Rakha, E.A.; Fan, S.J.; Valli, A.; Haider, S.; Goberdhan, D.C.I.; Green, A.R.; et al. Increased expression of glutamine transporter SNAT2/SLC38A2 promotes glutamine dependence and oxidative stress resistance, and is associated with worse prognosis in triple-negative breast cancer. Br. J. Cancer 2021, 124, 494–505. [Google Scholar] [CrossRef] [PubMed]
  108. Birsoy, K.; Wang, T.; Chen, W.W.; Freinkman, E.; Abu-Remaileh, M.; Sabatini, D.M. An essential role of the mitochondrial electron transport chain in cell proliferation is to enable aspartate synthesis. Cell 2015, 162, 540. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  109. López de la Oliva, A.R.; Campos-Sandoval, J.A.; Gómez-García, M.C.; Cardona, C.; Martín-Rufián, M.; Sialana, F.J.; Castilla, L.; Bae, N.; Lobo, C.; Peñalver, A.; et al. Nuclear Translocation of Glutaminase GLS2 in Human Cancer Cells Associates with Proliferation Arrest and Differentiation. Sci. Rep. 2020, 10, 2259. [Google Scholar] [CrossRef] [Green Version]
  110. Hiyama, N.; Ando, T.; Maemura, K.; Sakatani, T.; Amano, Y.; Watanabe, K.; Kage, H.; Yatomi, Y.; Nagase, T.; Nakajima, J.; et al. Glutamate-cysteine ligase catalytic subunit is associated with cisplatin resistance in lung adenocarcinoma. Jpn. J. Clin. Oncol. 2018, 48, 303–307. [Google Scholar] [CrossRef]
  111. Craze, M.L.; El-Ansari, R.; Aleskandarany, M.A.; Cheng, K.W.; Alfarsi, L.; Masisi, B.; Diez-Rodriguez, M.; Nolan, C.C.; Ellis, I.O.; Rakha, E.A.; et al. Glutamate dehydrogenase (GLUD1) expression in breast cancer. Breast Cancer Res. Treat. 2019, 174, 79–91. [Google Scholar] [CrossRef] [PubMed]
  112. Abrego, J.; Gunda, V.; Vernucci, E.; Shukla, S.K.; King, R.J.; Dasgupta, A.; Goode, G.; Murthy, D.; Yu, F.; Singh, P.K. GOT1-Mediated Anaplerotic Glutamine Metabolism Regulates Chronic Acidosis Stress in Pancreatic Cancer Cells. Cancer Lett. 2017, 400, 37. [Google Scholar] [CrossRef]
  113. Smith, B.; Schafer, X.L.; Ambeskovic, A.; Spencer, C.M.; Land, H.; Munger, J. Addiction to Coupling of the Warburg Effect with Glutamine Catabolism in Cancer Cells. Cell Rep. 2016, 17, 821–836. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  114. Fendt, S.M.; Bell, E.L.; Keibler, M.A.; Olenchock, B.A.; Mayers, J.R.; Wasylenko, T.M.; Vokes, N.I.; Guarente, L.; Heiden, M.G.V.; Stephanopoulos, G. Reductive glutamine metabolism is a function of the α-ketoglutarate to citrate ratio in cells. Nat. Commun. 2013, 4, 2236. [Google Scholar] [CrossRef] [Green Version]
  115. Shim, E.H.; Livi, C.B.; Rakheja, D.; Tan, J.; Benson, D.; Parekh, V.; Kho, E.Y.; Ghosh, A.P.; Kirkman, R.; Velu, S.; et al. L-2-Hydroxyglutarate: An epigenetic modifier and putative oncometabolite in renal cancer. Cancer Discov. 2014, 4, 1290–1298. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  116. Intlekofer, A.M.; DeMatteo, R.G.; Venneti, S.; Finley, L.W.S.; Lu, C.; Judkins, A.R.; Rustenburg, A.S.; Grinaway, P.B.; Chodera, J.D.; Cross, J.R.; et al. Hypoxia Induces Production of L-2-Hydroxyglutarate. Cell Metab. 2015, 22, 304–311. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  117. Mukha, A.; Kahya, U.; Linge, A.; Chen, O.; Löck, S.; Lukiyanchuk, V.; Richter, S.; Alves, T.C.; Peitzsch, M.; Telychko, V.; et al. GLS-driven glutamine catabolism contributes to prostate cancer radiosensitivity by regulating the redox state, stemness and ATG5-mediated autophagy. Theranostics 2021, 11, 7844–7868. [Google Scholar] [CrossRef]
  118. Kim, J.; Hu, Z.; Cai, L.; Li, K.; Choi, E.; Faubert, B.; Bezwada, D.; Rodriguez-Canales, J.; Villalobos, P.; Lin, Y.F.; et al. CPS1 maintains pyrimidine pools and DNA synthesis in KRAS/LKB1-mutant lung cancer cells. Nature 2017, 546, 168–172. [Google Scholar] [CrossRef] [PubMed]
  119. Su, N.; Pan, Y.X.; Zhou, M.; Harvey, R.C.; Hunger, S.P.; Kilberg, M.S. Correlation between asparaginase sensitivity and asparagine synthetase protein content, but not mRNA, in acute lymphoblastic leukemia cell lines. Pediatr. Blood Cancer 2008, 50, 274–279. [Google Scholar] [CrossRef]
  120. Hill, J.M.; Roberts, J.; Loeb, E.; Khan, A.; Maclellan, A.; Hill, R.W. L-Asparaginase Therapy for Leukemia and Other Malignant Neoplasms: Remission in Human Leukemia. JAMA J. Am. Med. Assoc. 1967, 202, 882–888. [Google Scholar] [CrossRef]
  121. Holleman, A.; Cheok, M.H.; den Boer, M.L.; Yang, W.; Veerman, A.J.P.; Kazemier, K.M.; Pei, D.; Cheng, C.; Pui, C.-H.; Relling, M.V.; et al. Gene-expression patterns in drug-resistant acute lymphoblastic leukemia cells and response to treatment. N. Engl. J. Med. 2004, 351, 533–542. [Google Scholar] [CrossRef] [PubMed]
  122. Cui, H.; Darmanin, S.; Natsuisaka, M.; Kondo, T.; Asaka, M.; Shindoh, M.; Higashino, F.; Hamuro, J.; Okada, F.; Kobayashi, M.; et al. Enhanced expression of asparagine synthetase under glucose-deprived conditions protects pancreatic cancer cells from apoptosis induced by glucose deprivation and cisplatin. Cancer Res. 2007, 67, 3345–3355. [Google Scholar] [CrossRef] [Green Version]
  123. Tsai, P.Y.; Lee, M.S.; Jadhav, U.; Naqvi, I.; Madha, S.; Adler, A.; Mistry, M.; Naumenko, S.; Lewis, C.A.; Hitchcock, D.S.; et al. Adaptation of pancreatic cancer cells to nutrient deprivation is reversible and requires glutamine synthetase stabilization by mTORC1. Proc. Natl. Acad. Sci. USA 2021, 118, e2003014118. [Google Scholar] [CrossRef] [PubMed]
  124. Marin-Valencia, I.; Yang, C.; Mashimo, T.; Cho, S.; Baek, H.; Yang, X.L.; Rajagopalan, K.N.; Maddie, M.; Vemireddy, V.; Zhao, Z.; et al. Analysis of tumor metabolism reveals mitochondrial glucose oxidation in genetically diverse human glioblastomas in the mouse brain in vivo. Cell Metab. 2012, 15, 827–837. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  125. Davidson, S.M.; Papagiannakopoulos, T.; Olenchock, B.A.; Heyman, J.E.; Keibler, M.A.; Luengo, A.; Bauer, M.R.; Jha, A.K.; O’Brien, J.P.; Pierce, K.A.; et al. Environment Impacts the Metabolic Dependencies of Ras-Driven Non-Small Cell Lung Cancer. Cell Metab. 2016, 23, 517–528. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  126. Thomlinson, R.H.; Gray, L.H. The histological structure of some human lung cancers and the possible implications for radiotherapy. Br. J. Cancer 1955, 9, 539–549. [Google Scholar] [CrossRef] [Green Version]
  127. Ausprunk, D.H.; Folkman, J. Migration and proliferation of endothelial cells in preformed and newly formed blood vessels during tumor angiogenesis. Microvasc. Res. 1977, 14, 53–65. [Google Scholar] [CrossRef]
  128. Li, L.; Yang, L.; Fan, Z.; Xue, W.; Shen, Z.; Yuan, Y.; Sun, X.; Wang, D.; Lian, J.; Wang, L.; et al. Hypoxia-induced GBE1 expression promotes tumor progression through metabolic reprogramming in lung adenocarcinoma. Signal Transduct. Target. Ther. 2020, 5. [Google Scholar] [CrossRef] [PubMed]
  129. Evans, S.M.; Judy, K.D.; Dunphy, I.; Timothy Jenkins, W.; Hwang, W.T.; Nelson, P.T.; Lustig, R.A.; Jenkins, K.; Magarelli, D.P.; Hahn, S.M.; et al. Hypoxia is important in the biology and aggression of human glial brain tumors. Clin. Cancer Res. 2004, 10, 8177–8184. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  130. Wang, G.L.; Jiang, B.H.; Rue, E.A.; Semenza, G.L. Hypoxia-inducible factor 1 is a basic-helix-loop-helix-PAS heterodimer regulated by cellular O2 tension. Proc. Natl. Acad. Sci. USA 1995, 92, 5510–5514. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  131. Lum, J.J.; Bui, T.; Gruber, M.; Gordan, J.D.; DeBerardinis, R.J.; Covello, K.L.; Simon, M.C.; Thompson, C.B. The transcription factor HIF-1α plays a critical role in the growth factor-dependent regulation of both aerobic and anaerobic glycolysis. Genes Dev. 2007, 21, 1037. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  132. Sun, R.C.; Denko, N.C. Hypoxic regulation of glutamine metabolism through HIF1 and SIAH2 supports lipid synthesis that is necessary for tumor growth. Cell Metab. 2014, 19, 285–292. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  133. Kim, J.W.; Tchernyshyov, I.; Semenza, G.L.; Dang, C.V. HIF-1-mediated expression of pyruvate dehydrogenase kinase: A metabolic switch required for cellular adaptation to hypoxia. Cell Metab. 2006, 3, 177–185. [Google Scholar] [CrossRef] [Green Version]
  134. Papandreou, I.; Cairns, R.A.; Fontana, L.; Lim, A.L.; Denko, N.C. HIF-1 mediates adaptation to hypoxia by actively downregulating mitochondrial oxygen consumption. Cell Metab. 2006, 3, 187–197. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  135. Morotti, M.; Bridges, E.; Valli, A.; Choudhry, H.; Sheldon, H.; Wigfield, S.; Gray, N.; Zois, C.E.; Grimm, F.; Jones, D.; et al. Hypoxia-induced switch in SNAT2/SLC38A2 regulation generates endocrine resistance in breast cancer. Proc. Natl. Acad. Sci. USA 2019, 116, 12452–12461. [Google Scholar] [CrossRef] [Green Version]
  136. Wang, Y.; Bai, C.; Ruan, Y.; Liu, M.; Chu, Q.; Qiu, L.; Yang, C.; Li, B. Coordinative metabolism of glutamine carbon and nitrogen in proliferating cancer cells under hypoxia. Nat. Commun. 2019, 10. [Google Scholar] [CrossRef] [Green Version]
  137. Metallo, C.M.; Gameiro, P.A.; Bell, E.L.; Mattaini, K.R.; Yang, J.; Hiller, K.; Jewell, C.M.; Johnson, Z.R.; Irvine, D.J.; Guarente, L.; et al. Reductive glutamine metabolism by IDH1 mediates lipogenesis under hypoxia. Nature 2012, 481, 380–384. [Google Scholar] [CrossRef] [Green Version]
  138. Gameiro, P.A.; Yang, J.; Metelo, A.M.; Pérez-Carro, R.; Baker, R.; Wang, Z.; Arreola, A.; Rathmell, W.K.; Olumi, A.; López-Larrubia, P.; et al. In vivo HIF-mediated reductive carboxylation is regulated by citrate levels and sensitizes VHL-deficient cells to glutamine deprivation. Cell Metab. 2013, 17, 372–385. [Google Scholar] [CrossRef] [Green Version]
  139. Dion, H.W.; Fusari, S.A.; Jakubowski, Z.L.; Zora, J.G.; Bartz, Q.R. 6-Diazo-5-oxo-L-norleucine, A New Tumor-inhibitory Substance. II.1 Isolation and Characterization2. J. Am. Chem. Soc. 2002, 78, 3075–3077. [Google Scholar] [CrossRef]
  140. Du, W.; Zhang, L.; Brett-Morris, A.; Aguila, B.; Kerner, J.; Hoppel, C.L.; Puchowicz, M.; Serra, D.; Herrero, L.; Rini, B.I.; et al. HIF drives lipid deposition and cancer in ccRCC via repression of fatty acid metabolism. Nat. Commun. 2017, 8, 1769. [Google Scholar] [CrossRef] [PubMed]
  141. Fan, J.; Kamphorst, J.J.; Mathew, R.; Chung, M.K.; White, E.; Shlomi, T.; Rabinowitz, J.D. Glutamine-driven oxidative phosphorylation is a major ATP source in transformed mammalian cells in both normoxia and hypoxia. Mol. Syst. Biol. 2013, 9, 712. [Google Scholar] [CrossRef]
  142. Le, A.; Lane, A.N.; Hamaker, M.; Bose, S.; Gouw, A.; Barbi, J.; Tsukamoto, T.; Rojas, C.J.; Slusher, B.S.; Zhang, H.; et al. Glucose-independent glutamine metabolism via TCA cycling for proliferation and survival in B cells. Cell Metab. 2012, 15, 110–121. [Google Scholar] [CrossRef] [Green Version]
  143. Grassian, A.R.; Parker, S.J.; Davidson, S.M.; Divakaruni, A.S.; Green, C.R.; Zhang, X.; Slocum, K.L.; Pu, M.; Lin, F.; Vickers, C.; et al. IDH1 mutations alter citric acid cycle metabolism and increase dependence on oxidative mitochondrial metabolism. Cancer Res. 2014, 74, 3317–3331. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  144. Tapper, E.B.; Jiang, Z.G.; Patwardhan, V.R. Refining the ammonia hypothesis: A physiology-driven approach to the treatment of hepatic encephalopathy. Mayo Clin. Proc. 2015, 90, 646–658. [Google Scholar] [CrossRef] [Green Version]
  145. Meléndez-Rodríguez, F.; Urrutia, A.A.; Lorendeau, D.; Rinaldi, G.; Roche, O.; Böğürcü-Seidel, N.; Ortega Muelas, M.; Mesa-Ciller, C.; Turiel, G.; Bouthelier, A.; et al. HIF1α Suppresses Tumor Cell Proliferation through Inhibition of Aspartate Biosynthesis. Cell Rep. 2019, 26, 2257–2265.e4. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  146. Mandigo, A.C.; Yuan, W.; Xu, K.; Gallagher, P.; Pang, A.; Guan, Y.F.; Shafi, A.A.; Thangavel, C.; Sheehan, B.; Bogdan, D.; et al. RB/E2F1 as a Master Regulator of Cancer Cell Metabolism in Advanced Disease. Cancer Discov. 2021, 11, 2334–2353. [Google Scholar] [CrossRef]
  147. Du, L.; Liu, X.; Ren, Y.; Li, J.; Li, P.; Jiao, Q.; Meng, P.; Wang, F.; Wang, Y.; Wang, Y.S.; et al. Loss of SIRT4 promotes the self-renewal of Breast Cancer Stem Cells. Theranostics 2020, 10, 9458–9476. [Google Scholar] [CrossRef] [PubMed]
  148. Jeong, S.M.; Xiao, C.; Finley, L.W.S.; Lahusen, T.; Souza, A.L.; Pierce, K.; Li, Y.H.; Wang, X.; Laurent, G.; German, N.J.; et al. SIRT4 has tumor suppressive activity and regulates the cellular metabolic response to DNA damage by inhibiting mitochondrial glutamine metabolism. Cancer Cell 2013, 23, 450. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  149. Humpton, T.; Vousden, K.H. Taking up the reins of power: Metabolic functions of p53. J. Mol. Cell Biol. 2019, 11, 610–614. [Google Scholar] [CrossRef]
  150. Ishak Gabra, M.B.; Yang, Y.; Lowman, X.H.; Reid, M.A.; Tran, T.Q.; Kong, M. IKKβ activates p53 to promote cancer cell adaptation to glutamine deprivation. Oncogenesis 2018, 7, 93. [Google Scholar] [CrossRef] [Green Version]
  151. Tajan, M.; Hock, A.K.; Blagih, J.; Robertson, N.A.; Labuschagne, C.F.; Kruiswijk, F.; Humpton, T.J.; Adams, P.D.; Vousden, K.H. A Role for p53 in the Adaptation to Glutamine Starvation through the Expression of SLC1A3. Cell Metab. 2018, 28, 721–736.e6. [Google Scholar] [CrossRef] [Green Version]
  152. Lowman, X.H.; Hanse, E.A.; Yang, Y.; Ishak Gabra, M.B.; Tran, T.Q.; Li, H.; Kong, M. p53 Promotes Cancer Cell Adaptation to Glutamine Deprivation by Upregulating Slc7a3 to Increase Arginine Uptake. Cell Rep. 2019, 26, 3051–3060.e4. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  153. Wise, D.R.; Deberardinis, R.J.; Mancuso, A.; Sayed, N.; Zhang, X.Y.; Pfeiffer, H.K.; Nissim, I.; Daikhin, E.; Yudkoff, M.; McMahon, S.B.; et al. Myc regulates a transcriptional program that stimulates mitochondrial glutaminolysis and leads to glutamine addiction. Proc. Natl. Acad. Sci. USA 2008, 105, 18782–18787. [Google Scholar] [CrossRef] [Green Version]
  154. Bott, A.J.; Peng, I.C.; Fan, Y.; Faubert, B.; Zhao, L.; Li, J.; Neidler, S.; Sun, Y.; Jaber, N.; Krokowski, D.; et al. Oncogenic Myc Induces Expression of Glutamine Synthetase through Promoter Demethylation. Cell Metab. 2015, 22, 1068–1077. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  155. Wong, C.C.; Xu, J.; Bian, X.; Wu, J.L.; Kang, W.; Qian, Y.; Li, W.; Chen, H.; Gou, H.; Liu, D.; et al. In Colorectal Cancer Cells with Mutant KRAS, SLC25A22-Mediated Glutaminolysis Reduces DNA Demethylation to Increase WNT Signaling, Stemness, and Drug Resistance. Gastroenterology 2020, 159, 2163–2180.e6. [Google Scholar] [CrossRef] [PubMed]
  156. Gaglio, D.; Metallo, C.M.; Gameiro, P.A.; Hiller, K.; Danna, L.S.; Balestrieri, C.; Alberghina, L.; Stephanopoulos, G.; Chiaradonna, F. Oncogenic K-Ras decouples glucose and glutamine metabolism to support cancer cell growth. Mol. Syst. Biol. 2011, 7, 523. [Google Scholar] [CrossRef]
  157. Patel, D.; Menon, D.; Bernfeld, E.; Mroz, V.; Kalan, S.; Loayza, D.; Foster, D.A. Aspartate Rescues S-phase Arrest Caused by Suppression of Glutamine Utilization in KRas-driven Cancer Cells. J. Biol. Chem. 2016, 291, 9322–9329. [Google Scholar] [CrossRef] [Green Version]
  158. Ren, L.; Ruiz-Rodado, V.; Dowdy, T.; Huang, S.; Issaq, S.H.; Beck, J.; Wang, H.; Tran Hoang, C.; Lita, A.; Larion, M.; et al. Glutaminase-1 (GLS1) inhibition limits metastatic progression in osteosarcoma. Cancer Metab. 2020, 8, 4. [Google Scholar] [CrossRef] [PubMed]
  159. van den Heuvel, A.P.J.; Jing, J.; Wooster, R.F.; Bachman, K.E. Analysis of glutamine dependency in non-small cell lung cancer: GLS1 splice variant GAC is essential for cancer cell growth. Cancer Biol. Ther. 2012, 13, 1185–1194. [Google Scholar] [CrossRef] [Green Version]
  160. Robinson, M.M.; McBryant, S.J.; Tsukamoto, T.; Rojas, C.; Ferraris, D.V.; Hamilton, S.K.; Hansen, J.C.; Curthoys, N.P. Novel mechanism of inhibition of rat kidney-type glutaminase by bis-2-(5-phenylacetamido-1,2,4-thiadiazol-2-yl)ethyl sulfide (BPTES). Biochem. J. 2007, 406, 407–414. [Google Scholar] [CrossRef] [Green Version]
  161. Wang, J.B.; Erickson, J.W.; Fuji, R.; Ramachandran, S.; Gao, P.; Dinavahi, R.; Wilson, K.F.; Ambrosio, A.L.B.; Dias, S.M.G.; Dang, C.V.; et al. Targeting mitochondrial glutaminase activity inhibits oncogenic transformation. Cancer Cell 2010, 18, 207–219. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  162. Jin, H.; Wang, S.; Zaal, E.A.; Wang, C.; Wu, H.; Bosma, A.; Jochems, F.; Isima, N.; Jin, G.; Lieftink, C.; et al. A powerful drug combination strategy targeting glutamine addiction for the treatment of human liver cancer. Elife 2020, 9, 1–20. [Google Scholar] [CrossRef] [PubMed]
  163. Ruiz-Rodado, V.; Lita, A.; Dowdy, T.; Celiku, O.; Saldana, A.C.; Wang, H.; Yang, C.Z.; Chari, R.; Li, A.; Zhang, W.; et al. Metabolic plasticity of IDH1 -mutant glioma cell lines is responsible for low sensitivity to glutaminase inhibition. Cancer Metab. 2020, 8, 23. [Google Scholar] [CrossRef]
  164. Koch, K.; Hartmann, R.; Tsiampali, J.; Uhlmann, C.; Nickel, A.C.; He, X.; Kamp, M.A.; Sabel, M.; Barker, R.A.; Steiger, H.J.; et al. A comparative pharmaco-metabolomic study of glutaminase inhibitors in glioma stem-like cells confirms biological effectiveness but reveals differences in target-specificity. Cell Death Discov. 2020, 6, 20. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  165. Lopez, J.S.; Banerji, U. Combine and conquer: Challenges for targeted therapy combinations in early phase trials. Nat. Rev. Clin. Oncol. 2017, 14, 57–66. [Google Scholar] [CrossRef]
  166. Guo, J.Y.; Teng, X.; Laddha, S.V.; Ma, S.; Van Nostrand, S.C.; Yang, Y.; Khor, S.; Chan, C.S.; Rabinowitz, J.D.; White, E. Autophagy provides metabolic substrates to maintain energy charge and nucleotide pools in Ras-driven lung cancer cells. Genes Dev. 2016, 30, 1704–1717. [Google Scholar] [CrossRef] [Green Version]
  167. Levy, J.M.M.; Towers, C.G.; Thorburn, A. Targeting Autophagy in Cancer. Nat. Rev. Cancer 2017, 17, 528. [Google Scholar] [CrossRef] [PubMed]
  168. Carracedo, A.; Cantley, L.C.; Pandolfi, P.P. Cancer metabolism: Fatty acid oxidation in the limelight. Nat. Rev. Cancer 2013, 13, 227–232. [Google Scholar] [CrossRef] [PubMed]
  169. Gounaris, A.D.; Hager, L.P. A resolution of the Escherichia coli pyruvate dehydrogenase complex. J. Biol. Chem. 1961, 236, 1013–1018. [Google Scholar] [CrossRef]
  170. Saladini, S.; Aventaggiato, M.; Barreca, F.; Morgante, E.; Sansone, L.; Russo, M.A.; Tafani, M. Metformin Impairs Glutamine Metabolism and Autophagy in Tumour Cells. Cells 2019, 8, 49. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  171. Andrzejewski, S.; Siegel, P.M.; St-Pierre, J. Metabolic Profiles Associated with Metformin Efficacy in Cancer. Front. Endocrinol. (Lausanne) 2018, 9, 372. [Google Scholar] [CrossRef] [Green Version]
  172. Kalinsky, K.; Harding, J.; DeMichele, A.; Infante, J.; Gogineni, K.; Owonikoko, T.; Isakoff, S.; Iliopoulos, O.; Patel, M.; Munster, P.; et al. Abstract PD3-13: Phase 1 study of CB-839, a first-in-class oral inhibitor of glutaminase, in combination with paclitaxel in patients with advanced triple negative breast cancer. AACR 2018, 78, PD3-13. [Google Scholar]
  173. Calithera Biosciences, Inc. Initial Results from Phase 2 Study of CB-839 in Combination with Opdivo® (Nivolumab) to Be Presented at the Society for Immunotherapy of Cancer Meeting. 2017. Available online: https://www.globenewswire.com/en/news-release/2017/11/07/1176454/32478/en/Initial-Results-from-Phase-2-Study-of-CB-839-in-Combination-with-Opdivo-nivolumab-to-be-Presented-at-the-Society-for-Immunotherapy-of-Cancer-Meeting.html (accessed on 16 December 2021).
  174. Harding, J.J.; Telli, M.L.; Munster, P.N.; Le, M.H.; Molineaux, C.; Bennett, M.K.; Mittra, E.; Burris, H.A.; Clark, A.S.; Dunphy, M.; et al. Safety and tolerability of increasing doses of CB-839, a first-in-class, orally administered small molecule inhibitor of glutaminase, in solid tumors. J. Clin. Oncol. 2015, 33, 2512. [Google Scholar] [CrossRef]
  175. Wang, E.S.; Frankfurt, O.; Orford, K.W.; Bennett, M.; Flinn, I.W.; Maris, M.; Konopleva, M. Phase 1 Study of CB-839, a First-in-Class, Orally Administered Small Molecule Inhibitor of Glutaminase in Patients with Relapsed/Refractory Leukemia. Blood 2015, 126, 2566. [Google Scholar] [CrossRef]
  176. Calithera Biosciences Reports CANTATA Study of Telaglenastat in Renal Cell Carcinoma Did Not Achieve Primary Endpoint. Available online: https://www.globenewswire.com/fr/news-release/2021/01/04/2152519/0/en/Calithera-Biosciences-Reports-CANTATA-Study-of-Telaglenastat-in-Renal-Cell-Carcinoma-Did-Not-Achieve-Primary-Endpoint.html (accessed on 16 December 2021).
  177. Motzer, R.J.; Lee, C.-H.; Emamekhoo, H.; Matrana, M.; Percent, I.; Hsieh, J.J.; Hussain, A.; Vaishampayan, U.N.; Graham, R.; Liu, S.; et al. ENTRATA: Randomized, double-blind, phase II study of telaglenastat (tela; CB-839) + everolimus (E) vs. placebo (pbo) + E in patients (pts) with advanced/metastatic renal cell carcinoma (mRCC). Ann. Oncol. 2019, 30, v889–v890. [Google Scholar] [CrossRef]
  178. Achkar, I.W.; Kader, S.; Dib, S.S.; Junejo, K.; Al-Bader, S.B.; Hayat, S.; Bhagwat, A.M.; Rousset, X.; Wang, Y.; Viallet, J.; et al. Metabolic Signatures of Tumor Responses to Doxorubicin Elucidated by Metabolic Profiling in Ovo. Metabolites 2020, 10, 268. [Google Scholar] [CrossRef] [PubMed]
  179. Volkova, M.; Russell, R. III Anthracycline Cardiotoxicity: Prevalence, Pathogenesis and Treatment. Curr. Cardiol. Rev. 2011, 7, 214. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  180. Hawk, M.A.; McCallister, C.; Schafer, Z.T. Antioxidant Activity during Tumor Progression: A Necessity for the Survival of Cancer Cells? Cancers 2016, 8, 92. [Google Scholar] [CrossRef]
  181. Karner, C.M.; Esen, E.; Okunade, A.L.; Patterson, B.W.; Long, F. Increased glutamine catabolism mediates bone anabolism in response to WNT signaling. J. Clin. Investig. 2015, 125, 551. [Google Scholar] [CrossRef] [Green Version]
  182. Sun, W.; Luan, S.; Qi, C.; Tong, Q.; Yan, S.; Li, H.; Zhang, Y. Aspulvinone O, a natural inhibitor of GOT1 suppresses pancreatic ductal adenocarcinoma cells growth by interfering glutamine metabolism. Cell Commun. Signal. 2019, 17. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  183. Yoshida, T.; Yamasaki, S.; Kaneko, O.; Taoka, N.; Tomimoto, Y.; Namatame, I.; Yahata, T.; Kuromitsu, S.; Cantley, L.C.; Lyssiotis, C.A. A covalent small molecule inhibitor of glutamate-oxaloacetate transaminase 1 impairs pancreatic cancer growth. Biochem. Biophys. Res. Commun. 2020, 522, 633–638. [Google Scholar] [CrossRef] [PubMed]
  184. Dixon, S.J.; Lemberg, K.M.; Lamprecht, M.R.; Skouta, R.; Zaitsev, E.M.; Gleason, C.E.; Patel, D.N.; Bauer, A.J.; Cantley, A.M.; Yang, W.S.; et al. Ferroptosis: An iron-dependent form of nonapoptotic cell death. Cell 2012, 149, 1060–1072. [Google Scholar] [CrossRef] [Green Version]
  185. Jin, L.; Chun, J.; Pan, C.; Kumar, A.; Zhang, G.; Ha, Y.; Li, D.; Alesi, G.N.; Kang, Y.; Zhou, L.; et al. The PLAG1-GDH1 Axis Promotes Anoikis Resistance and Tumor Metastasis through CamKK2-AMPK Signaling in LKB1-Deficient Lung Cancer. Mol. Cell 2018, 69, 87–99.e7. [Google Scholar] [CrossRef] [Green Version]
  186. Abdul Kader, S.; Dib, S.; Achkar, I.W.; Thareja, G.; Suhre, K.; Rafii, A.; Halama, A. Defining the landscape of metabolic dysregulations in cancer metastasis. Clin. Exp. Metastasis 2021. [Google Scholar] [CrossRef] [PubMed]
  187. Li, C.; Li, M.; Chen, P.; Narayan, S.; Matschinsky, F.M.; Bennett, M.J.; Stanley, C.A.; Smith, T.J. Green tea polyphenols control dysregulated glutamate dehydrogenase in transgenic mice by hijacking the ADP activation site. J. Biol. Chem. 2011, 286, 34164–34174. [Google Scholar] [CrossRef] [Green Version]
  188. Chen, B.H.; Hsieh, C.H.; Tsai, S.Y.; Wang, C.Y.; Wang, C.C. Anticancer effects of epigallocatechin-3-gallate nanoemulsion on lung cancer cells through the activation of AMP-activated protein kinase signaling pathway. Sci. Rep. 2020, 10, 5163. [Google Scholar] [CrossRef] [Green Version]
  189. Levenberg, B.; Melnick, I.; Buchanan, J.M. Biosynthesis of the purines. XV. The effect of aza-L-serine and 6-diazo-5-oxo-L-norleucine on inosinic acid biosynthesis de novo. J. Biol. Chem. 1957, 225, 163–176. [Google Scholar] [CrossRef]
  190. Ahluwalia, G.S.; Grem, J.L.; Hao, Z.; Cooney, D.A. Metabolism and action of amino acid analog anti-cancer agents. Pharmacol. Ther. 1990, 46, 243–271. [Google Scholar] [CrossRef] [Green Version]
  191. Leone, R.D.; Zhao, L.; Englert, J.M.; Sun, I.M.; Oh, M.H.; Sun, I.H.; Arwood, M.L.; Bettencourt, I.A.; Patel, C.H.; Wen, J.; et al. Glutamine blockade induces divergent metabolic programs to overcome tumor immune evasion. Science 2019, 366, 1013–1021. [Google Scholar] [CrossRef] [PubMed]
  192. Tenora, L.; Alt, J.; Dash, R.P.; Gadiano, A.J.; Novotná, K.; Veeravalli, V.; Lam, J.; Kirkpatrick, Q.R.; Lemberg, K.M.; Majer, P.; et al. Tumor-Targeted Delivery of 6-Diazo-5-oxo-l-norleucine (DON) Using Substituted Acetylated Lysine Prodrugs. J. Med. Chem. 2019, 62, 3524–3538. [Google Scholar] [CrossRef]
  193. Jackson, D.; Robson, J.M.; Wander, A.C. The effect of 6-diazo-5-oxo-1-norleucine (DON) on pregnancy. J. Endocrinol. 1959, 18, 204–207. [Google Scholar] [CrossRef] [PubMed]
  194. Hassanein, M.; Qian, J.; Hoeksema, M.D.; Wang, J.; Jacobovitz, M.; Ji, X.; Harris, F.T.; Harris, B.K.; Boyd, K.L.; Chen, H.; et al. Targeting SLC1A5-mediated glutamine dependence in non-small cell lung cancer. Int. J. Cancer 2015, 137, 1587. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  195. Schulte, M.L.; Fu, A.; Zhao, P.; Li, J.; Geng, L.; Smith, S.T.; Kondo, J.; Coffey, R.J.; Johnson, M.O.; Rathmell, J.C.; et al. Pharmacological blockade of ASCT2-dependent glutamine transport leads to antitumor efficacy in preclinical models. Nat. Med. 2018, 24, 194–202. [Google Scholar] [CrossRef] [PubMed]
  196. Ni, F.; Yu, W.M.; Li, Z.; Graham, D.K.; Jin, L.; Kang, S.; Rossi, M.R.; Li, S.; Broxmeyer, H.E.; Qu, C.K. Critical role of ASCT2-mediated amino acid metabolism in promoting leukaemia development and progression. Nat. Metab. 2019, 1, 390. [Google Scholar] [CrossRef]
  197. Bröer, A.; Gauthier-Coles, G.; Rahimi, F.; van Geldermalsen, M.; Dorsch, D.; Wegener, A.; Holst, J.; Bröer, S. Ablation of the ASCT2 (SLC1A5) gene encoding a neutral amino acid transporter reveals transporter plasticity and redundancy in cancer cells. J. Biol. Chem. 2019, 294, 4012. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  198. Tran, T.Q.; Ishak Gabra, M.B.; Lowman, X.H.; Yang, Y.; Reid, M.A.; Pan, M.; O’Connor, T.R.; Kong, M. Glutamine deficiency induces DNA alkylation damage and sensitizes cancer cells to alkylating agents through inhibition of ALKBH enzymes. PLoS Biol. 2017, 15, e2002810. [Google Scholar] [CrossRef]
  199. Earhart, R.H.; Amato, D.J.; Yuang-Chi Chang, A.; Borden, E.C.; Shiraki, M.; Dowd, M.E.; Comis, R.L.; Davis, T.E.; Smith, T.J. Phase II trial of 6-diazo-5-oxo-L-norleucine versus aclacinomycin-A in advanced sarcomas and mesotheliomas. Investig. New Drugs 1990, 8, 113–119. [Google Scholar] [CrossRef] [PubMed]
  200. Rubin, J.; Sorensen, S.; Schutt, A.J.; van Hazel, G.A.; O’Connell, M.J.; Moertel, C.G. A phase II study of 6-diazo-5-oxo-L-norleucine (DON, NSC-7365) in advanced large bowel carcinoma. Am. J. Clin. Oncol. 1983, 6, 325–326. [Google Scholar] [CrossRef]
  201. Yokoyama, Y.; Wild, R. Abstract 5607: Broad acting glutamine antagonism remodels the tumor microenvironment, induces distinctive immune modulation, and synergizes with immune checkpoint blockade. Cancer Res. 2020, 80, 5607. [Google Scholar]
  202. Johnson, M.L.; Doroshow, D.B.; Seiwert, T.Y.; Gibson, M.K.; Velcheti, V.; Lisberg, A.E.; Patel, S.A.; Scheffler, M.; Lafleur, F.; Dugan, M.H.; et al. Phase 1 and phase 2a, first-in-human (FIH) study, of DRP-104, a broad glutamine antagonist, in adult patients with advanced solid tumors. J. Clin. Oncol. 2021, 39, TPS3149. [Google Scholar] [CrossRef]
  203. Houten, S.M. Metabolomics: Unraveling the chemical individuality of common human diseases. Ann. Med. 2009, 41, 402–407. [Google Scholar] [CrossRef]
  204. Doucey, M.A.; Carrara, S. Nanowire Sensors in Cancer. Trends Biotechnol. 2019, 37, 86–99. [Google Scholar] [CrossRef] [PubMed] [Green Version]
Table
Table 2. Overview of completed clinical trials testing telaglenastat (CB-839) in various cancer patients.
Table 2. Overview of completed clinical trials testing telaglenastat (CB-839) in various cancer patients.
Cancer Type Treatment Outcome Reference
Triple-negative breast cancer (TNBC) In combination with paclitaxel (chemotherapeutic agent targeting microtubules) In heavily pretreated patients with previous taxane exposure, the treatment demonstrated clinical activity and was well tolerated. [172]
Clear cell renal cell carcinoma (ccRCC), melanoma; non-small-cell lung cancer (NSCLC) In combination with nivolumab (immunotherapy medication targeting programmed cell death (PD-1) receptor) CB-839 was well tolerated when combined with nivolumab in melanoma, ccRCC, and NSCLC
patients.		 [173]
Solid tumors with K-Ras mutation In combination with palbociclib (kinase inhibitor targeting cyclin-dependent kinases CDK4 and CDK6) NA NA
Solid tumors As a single agent and in combination with standard chemotherapy Acceptable safety profile under continuous CB-839 administration. Treatment resulted in glutaminase inhibition and clinical activity. [174]
Hematological tumors As a single agent or in combination with pomalidomide (immunomodulatory agent), dexamethasone (glucocorticoid), or pomalidomide and dexamethasone CB-839 administration was well tolerated and resulted in GLS inhibition in blood platelets and in tumors. Observed reductions in marrow and peripheral blast counts suggested clinical relevance. [175]
Renal cell carcinoma (RCC) In combination with cabozantinib (tyrosine kinase inhibitor) Did not achieve primary endpoint. [176]
ccRCC In combination with everolimus (mammalian target of rapamycin (mTOR) kinase inhibitor) In combination with everolimus, CB-839 demonstrated a tolerable safety profile. Modest (3.8 months from 1.9 months) progression-free survival was observed. [177]
Leukemia CB-839 as a single agent or in combination with azacitidine (chemotherapeutic agent, antimetabolite) CB-839 was well tolerated in advanced leukemia and resulted in GLS inhibition in platelets and PBMCs.
Two patients achieved significant reductions in blast counts.		 [175]
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Share and Cite

MDPI and ACS Style

Halama, A.; Suhre, K. Advancing Cancer Treatment by Targeting Glutamine Metabolism—A Roadmap. Cancers 2022, 14, 553. https://doi.org/10.3390/cancers14030553

AMA Style

Halama A, Suhre K. Advancing Cancer Treatment by Targeting Glutamine Metabolism—A Roadmap. Cancers. 2022; 14(3):553. https://doi.org/10.3390/cancers14030553

Chicago/Turabian Style

Halama, Anna, and Karsten Suhre. 2022. "Advancing Cancer Treatment by Targeting Glutamine Metabolism—A Roadmap" Cancers 14, no. 3: 553. https://doi.org/10.3390/cancers14030553

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop