How does the cell regulate different metabolic pathways?

I heard somewhere that cells use different nucleosides bound to triphosphates e.g. ATP, GTP, CTP and other modified compounds: NADH, NADPH to distinguish between different metabolic pathways and so they regulate where they use up the energy. I heard that kinases play an important role in the regulation. Is there a connection (I guess there is if I check NADPH)? Is this regulation mapped? I mean is there a simple map which contains the main processes and the energy carrier and regulatory compounds?

I am looking for something like this map (of receptor responses), but for metabolic regulation:

  • Figure 1 - signal transduction - wikipedia

So it possibly contains mitochondria, O2, CO2, flows, ATP, NADPH, etc… I understand that different cell types can have different energy producer and consumer organelles and it is not possible to create something that is universally true, so I would be satisfied with a map of your favorite human cell type.

I heard somewhere that cells… so they regulate where they use up the energy.

Yes NADP/H is primarily employed in anabolic pathways such as fatty acid synthesis, while NAD/H is employed in catabolic pathways such as glycolysis.

I don't think there is a general rule for other "energy-currency" molecules (pyrimidine triphosphates are not used except in some rare cases such as glycogenesis).

I mean is there a simple map which contains the main processes and the energy carrier and regulatory compounds?

You can search for the term "metabolic network". It would be too huge so it is better to look at specific sub-networks. KEGG is a good site for finding metabolic networks. There are other representations like hive plots, to visualize very huge graphs.

Our body maintains a very delicate balance between the concentration of metabolites and substrates. If a pathway is not regulated, excess of a particular metabolite can disturb the whole process. Let's take cholesterol metabolism pathway for example. there are various means by which the cells control the production of cholesterol. For example AMP controlled kinase protein detects weather the concentration of ATP is high or not, if its low then the cell does not proceed with the production of it, by inhibiting the enzyme that catalysis the production of mevalonate. Other way is through transcription factor that is present called SREBP. Sterol Regulatory Element Binding Protein regulate multiple genes that are involved in metabolism of cholesterol. Various protein denaturing components are present that upon receiving signal cleaves the intermediate's in the metabolism of cholesterol. There are n-number of pathways in our body and all are regulated by a complex network of signals that regulate them.

A metabolic switch regulates the transition between growth and diapause in C. elegans

Background: Metabolic activity alternates between high and low states during different stages of an organism's life cycle. During the transition from growth to quiescence, a major metabolic shift often occurs from oxidative phosphorylation to glycolysis and gluconeogenesis. We use the entry of Caenorhabditis elegans into the dauer larval stage, a developmentally arrested stage formed in response to harsh environmental conditions, as a model to study the global metabolic changes and underlying molecular mechanisms associated with growth to quiescence transition.

Results: Here, we show that the metabolic switch involves the concerted activity of several regulatory pathways. Whereas the steroid hormone receptor DAF-12 controls dauer morphogenesis, the insulin pathway maintains low energy expenditure through DAF-16/FoxO, which also requires AAK-2/AMPKα. DAF-12 and AAK-2 separately promote a shift in the molar ratios between competing enzymes at two key branch points within the central carbon metabolic pathway diverting carbon atoms from the TCA cycle and directing them to gluconeogenesis. When both AAK-2 and DAF-12 are suppressed, the TCA cycle is active and the developmental arrest is bypassed.

Conclusions: The metabolic status of each developmental stage is defined by stoichiometric ratios within the constellation of metabolic enzymes driving metabolic flux and controls the transition between growth and quiescence.

Conflict of interest statement

The authors declare that they have no competing interests.


DAF-16 mediates the switch to…

DAF-16 mediates the switch to low metabolic rate in dauer formation but does…

DAF-16 determines the energy expenditure…

DAF-16 determines the energy expenditure and lifespan of dauer larvae and, together with…

AAK-2 regulates the catabolism, the…

AAK-2 regulates the catabolism, the gluconeogenic mode, and the developmental arrest in the…

The metabolic switch is achieved…

The metabolic switch is achieved through the regulation of enzymes that work on…

DAF-12 and AAK-2 control the…

DAF-12 and AAK-2 control the molar ratios of the enzymes at the branch…

Metabolic control in the transition…

Metabolic control in the transition to the dauer state. The model represents the…

Precise control over neuron growth paves the way for repairing injuries, improving brain models

“This is the first time that a lineage transcription factor has been linked to a metabolic pathway in a tissue differentiation process during development, and I think this is the tip of the iceberg: in every tissue you’ll need to control certain metabolic pathways at a transcriptional level,” said Zon, who is also the director of the Boston Children’s Hospital Stem Cell Program and a Howard Hughes Medical Institute Investigator. “DHODH inhibitors are already in clinical trials to treat leukemia, and we previously had shown that they are also effective in melanoma. This work shows that DHODH inhibitors could thus lead to therapeutic benefits by releasing coenzyme Q in metabolically sensitive cancers to activate processes linked to cell differentiation.”

This work was supported by the National Heart, Lung, and Blood Institute (4R01HL048801, 5P01HL032262, 5U01HL134812, and 1P01HL131477), the National Institute of Diabetes and Digestive and Kidney Diseases (1U54DK110805 and 3R24DK092760), Harvard Catalyst, the Canadian Institutes of Health Research, the National Cancer Institute (5R01CA213062), the National Institute of General Medical Sciences (R35GM127045), the National Human Genome Research Institute (U54-HG008097), the Cancer Research Institute, and the American Lebanese Syrian Associated Charities.


DNA repair and metabolic pathways are vital to maintain cellular homeostasis. Under normal cellular conditions, DNA repair proteins can maintain genomic stability following exposure to exogenous and endogenous genotoxic insults. When growing in normal physiological conditions, cells predominately rely on the TCA cycle to generate ATP and other essential precursors for cellular processes. However, it has been well established that tumor cells are more likely to generate energy via glycolysis and hyperactivate their DNA damage response pathways, both of which promote the uncontrolled proliferative, survival and cellular growth pathways (Warburg, 1925). It was initially proposed that these two mechanisms operate independently within the cell however, recent studies suggest a link between DNA repair and glycolysis. For instance, several independent studies have suggested novel roles for glycolytic proteins in DNA repair pathways, largely based on the observation that several glycolytic proteins, including Hexokinase II, Fumarase and ATP-citrate lyase (ACLY), migrate to the nucleus following exposure to genomic stress (van Vugt, 2017 Ohba et al., 2020 Hitosugi et al., 2012 Yuan et al., 2010). Several studies have also suggested glycolysis may be involved in maintaining genome stability, given that the glycolytic pathway provides metabolites which play an essential role in DNA metabolism. For example, the pentose phosphate pathway (PPP) utilizes the glycolysis intermediate, glucose-6-phosphate, to ultimately enable the biosynthesis of nucleotides via the generation of ribose-5-phosphate. Despite this, the interaction between DNA repair pathways and glycolysis remains unclear. Metabolic products from glycolysis, such as L- and D-lactate also play a role in DNA repair by decreasing chromatin compaction and subsequently increasing transcription of key genes involved in DNA DSB (double-strand break) repair (Wagner et al., 2015). Here, we will review the peer-reviewed evidence linking metabolism and DNA repair and how these processes may lead to radio- and chemo-resistance in tumor cells.

The Warburg Effect and Tumor Metabolic Reprogramming

High glucose intake is a characteristic shared amongst most solid tumors, and this phenomenon was first described in 1920 by Otto Warburg (Warburg et al., 1927). This observation, referred to as the Warburg effect, describes how cancer cells shift their predominate metabolic pathway from oxidative phosphorylation to anaerobic glycolysis, consequently producing high levels of lactic acid via fermentation (Warburg et al., 1927 Warburg, 1956). Recently, studies have demonstrated that elevated lactic acid production may induce resistance to major anti-cancer therapies, including radiation and chemotherapy, via numerous mechanisms. Furthermore, the upregulated production of lactic acid contributes to the development of an acidic tumor microenvironment, which has been associated with increased metastatic capacity and growth rate in a subset of aggressive tumors (Turkcan et al., 2019).

In the early studies of Warburg’s effect, it was thought that cancer cells experience mitochondrial dysfunction via the “irreversible injuring of respiration,” as cancer cells downregulate oxidative phosphorylation during the tricarboxylic acid cycle (TCA, also known as the Krebs cycle) (Warburg, 1956). However, subsequent investigations of mitochondrial functionality in tumor cells revealed that the majority of tumor cells possess functional mitochondria, and can still undergo oxidative phosphorylation (Zong et al., 2016). This led to speculation as to why cancer cells with functional mitochondria preferentially convert excess pyruvate to lactate, instead of utilizing oxidative phosphorylation to more efficiently produce ATP.

As altered metabolic features are observed commonly across many cancer subtypes, reprogrammed metabolism is considered one of Pavlova and Thompson’s hallmarks of cancer (Pavlova and Thompson, 2016). For example, increased glucose uptake has been observed in a variety of tumor contexts and has been shown to negatively correlate with tumor prognostic markers and be involved in chemo- and radio-resistance mechanisms. This has been clinically exploited using 18 F-deoxyglucose-positron emission tomography (FDG-PET) based imaging, where a radioactive fluorine-labeled glucose analog it utilized to diagnose and stage tumor progression (Spermon et al., 2002).

DNA Repair Pathways and Their Relationship to Tumor Therapies

In tumor cells that undergo metabolic reprogramming, there is an observable increase in the activation of the DNA damage response pathways, which subsequently trigger nucleotide synthesis and anabolic glucose metabolism (Tong et al., 2009). DNA damage response pathways are highly active in tumor cells, subsequently promoting their rapid growth and survival. The DNA damage response consists of several DNA repair pathways, and each pathway represents a specific mechanism to repair a specific type of DNA damage. The initiation and progression of repair pathways is considered a spatiotemporally regulated process in which proteins move toward DNA damage sites, following the remodeling of the chromatin (van Attikum and Gasser, 2009 Gospodinov and Herceg, 2013). DNA damage may be induced by several endogenous sources such as DNA double-strand breaks and oxidative stress induced by reactive oxygen species, resulting from cellular metabolism. DNA damage may also result from exogenous sources, for example nucleotide damage from UV light or oxidative damage and DNA strand breaks caused by ionizing radiation (Jackson and Bartek, 2009 Tubbs and Nussenzweig, 2017). In order to maintain the integrity of genome, in human cells there are several types of DNA repair processes, classified into five major pathways including base excision repair (BER), nucleotide excision repair (NER), mismatch repair (MMR), non-homologous end-joining (NHEJ), and homology-directed repair (HDR) (Kalluri, 2016 Roos et al., 2016 Chatterjee and Walker, 2017). In addition to having a critical role in maintenance of genome integrity, alterations in the expression, and function of DNA repair proteins are a major mediator of tumor responses to chemo- and radiotherapy, which commonly function by inducing DNA damage in tumor cells. Here, we will briefly discuss the relevance of each repair pathway on tumor sensitivity to chemo- and radiotherapies, but further detail can be found in the following review (Minchom et al., 2018).

In terms of chemo- and radiotherapy, DSB repair via NHEJ and HDR is an important consideration, since many therapies, including radiotherapy, topoisomerase inhibitors, such as doxorubicin, and PARP inhibitors, induce DNA DSBs. Therefore, the defective functioning of DSB repair pathways can significantly influence the tumor response to these therapies. For example mutations or decreased expression of the Breast Cancer Associated 1 and 2 (BRCA1 and BRCA2) proteins can lead to defects in the HDR of DNA DSBs, sensitizing tumor cells to PARP inhibitors and radiotherapy that induce lesions that require HDR for repair (Rose et al., 2020). Conversely, upregulation of DNA DSB repair proteins in the NHEJ pathway can also induce resistance to these DSB-inducing therapies, due to the tumor cells ability to rapidly repair DNA damage and therefore avoid induction of cell death (Jensen and Rothenberg, 2020). BER removes and repairs damaged bases within the DNA. The capacity of cells to perform BER is also of relevance to tumor therapy as the anti-tumor agents temozolomide, pemetrexed, or floxuridine induce DNA lesions of N7mG, uracil, or 5-FU, respectively, all of which can be recognized and repaired by the BER pathway (Storr et al., 2011). Upregulation or down regulation of the BER pathway can lead to resistance or sensitivity, respectively, to these agents. Several inhibitors of the BER pathway are also in development (Grundy et al., 2020).

In the process of MMR, proteins recognize mismatched bases in DNA which arise from processes such as replication. MMR proteins identify, excise and replace these mismatched bases with the correct pairing base. Mutations in the MMR genes Mlh1 and Msh2 are associated with the human colon cancer-prone syndrome, Lynch Syndrome [also known as hereditary non-polyposis colorectal cancer (HNPCC)], but MMR genes are also frequently mutated in other cancers. Tumors with mutated Mlh1 and Msh2 in colon tumors were historically targeted with methotrexate, which leads to the accumulation of oxidative damage. However, due to the high number of somatic mutations found in MMR-deficient tumors, which can contribute to stimulation of the immune system, immunotherapy is showing potential to become the preferred therapy for tumors with defects in MMR (Le et al., 2015).

The nucleotide excision repair pathway recognizes damaged nucleotides including pyrimidine dimers, intrastrand crosslinks, and bulky adducts. Alkylating agents, such as platinum compounds like cisplatin are commonly used to treat many types of cancers and induce intrastrand crosslinks within the DNA, activating the NER pathway. Expression of the NER proteins, including ERCC1 are correlated with sensitivity to platinum agents in multiple tumor types due to an inability to resolve DNA crosslinks (Arora et al., 2010).

Therefore, although alterations in DNA repair pathways contribute to the development of tumors, and can lead to resistance to tumor therapies, they also hold huge potential as the next generation targets for the treatment of many cancer types. Due to the metabolic reprogramming in tumor cells, it is likely that targeting cellular metabolism may also be advantageous. The current literature supporting a link between metabolic reprogramming and the DNA damage response pathways will be further explored below.

The Requirement of the Pentose Phosphate Pathway (PPP) and G6PD Protein in DNA Damage Prevention and DNA Repair Processes

The pentose phosphate pathway is a parallel pathway to glycolysis and generates pentoses and NADPH, together with ribose-5-phosphate, a precursor for nucleotide synthesis (Patra and Hay, 2014). The PPP is upregulated in several tumor types and regulates various functions that promote tumor growth, including DNA metabolism and cell proliferation (Mori-Iwamoto et al., 2007 Chan et al., 2013 Catanzaro et al., 2015). In non-carcinogenic cells, the PPP pathway is responsible for generating the bulk of nucleotides through salvage pathways, which recycle existing nucleosides and nucleobases. Although, a portion of nucleotide synthesis also takes place via de novo synthesis pathways to produce purine and pyrimidine rings to sustain rapid DNA metabolism (Mori-Iwamoto et al., 2007). Supporting this, highly proliferative cells, such as tumor cells, are more likely to use de novo nucleotide synthesis pathways over the salvage pathways to maintain the increased production of nucleotides and other macromolecules. The de novo nucleotide synthesis pathways maintain nucleic acid and protein synthesis, along with other cellular activities, to meet the high metabolic requirements of cancer cells (Kilstrup et al., 2005 Villa et al., 2019).

The PPP pathway consists of an oxidative and a non-oxidative phase: the oxidative phase generates NADPH that is used for reductive biosynthetic reactions, such as fatty acid synthesis and the prevention of oxidative stress by detoxification of oxygen species (ROS). The non-oxidative arms of the PPP produce ribose-5-phosphatase, which is then further metabolized for the production of nucleotides (Figure 1). The PPP pathway occurs in parallel to glycolysis, diverging from glycolysis at glucose-6-phosphate (G6P), which is involved in the oxidation of glucose to provide the building blocks for anabolic pathways (D’Urso et al., 1983). Alternatively, under conditions of high reductive demand cancer cells have the capacity to divert glucose-6-phosphate dehydrogenase (G6PD) into the PPP pathway to maintain the constant generation of NADPH and nucleotides (Bokun et al., 1987). Downregulation of NADPH production renders tumor cells more susceptible to oxidative DNA damage, as NADPH functions as a major cofactor for glutathione (GS) and cytochrome p450 reductase, which is essential for maintaining the cellular redox balance.

Figure 1. The relationship between the Pentose Phosphate Pathway and DNA damage and repair. The PPP comprises of two phases known as the oxidative and non-oxidative phases. The oxidative phase is responsible for the conversion of glucose-6-phosphate to ribose-5-phosphate, which releases NADPH to maintain the cellular redox balance and also reduces oxidative damage. In the non-oxidative phase, the activity of a key enzyme, G6PD is stimulated by ATM to promote the production of NADPH and nucleotide synthesis. The activation of G6PD is essential to maintain a reduced cellular environment and also synthesises nucleotide precursors for DNA damage repair. This figure was created with

Cells lacking G6PD are more sensitive to oxidative damage and therefore have increased sensitivity to ionizing radiation (IR), which in addition to inducing DNA strand breaks, also causes oxidative damage (Tuttle et al., 2000). G6PD is essential in sustaining a balanced pool of nucleotides in response to DNA damage and promotes PPP-mediated nucleotide synthesis. Furthermore, a study showed that Ataxia Telangiectasia Mutated (ATM), a key DNA damage protein, activates the PPP pathway through G6PD to promote antioxidant defense mechanisms and DNA repair activity via nucleotide production under stressed conditions (Cosentino et al., 2011). This suggests that G6PD activity is likely to also be required for the repair of DNA damage and maintaining DNA integrity (Zhang et al., 2016).

The wild type tumor suppressor protein, p53, has also been shown to downregulate the PPP via directly reducing G6PD activity (Jiang et al., 2011). However, inhibition of ATM is also known to downregulate p53 expression, subsequently promoting the constitutive upregulation of the PPP via G6PD upregulation too, consequently restoring dNTP levels in cancer cells and facilitating cellular proliferation (Aird et al., 2015). G6PD is the rate-limiting enzyme that regulates oxidative PPP and therefore controls the flux of dNTP production required for DNA replication and maintaining genome stability. As such, G6PD is also required to suppress dNTP-enhanced mutagenesis. Overall, the altered cellular metabolic flux induced by G6PD during metabolic reprogramming enables the more rapid repair of DNA lesions, promoting resistance to conventional therapies such as radiation, and cellular growth advantages (Leick and Levis, 2017).

Activating mutations of FMS-like tyrosine kinase 3 (FLT3) have been shown to drive the initiation and progression of acute myeloid leukemia (AML). As such, inhibition of FLT3 was suggested to be a promising treatment for AML however, targeting FLT3 as a monotherapy did not achieve long term remission (Leick and Levis, 2017). In contrast, a genome-wide RNA interference (RNAi)-based screen found that inhibition of the ATM/G6PD pathway in combination with FLT3 inhibition was synthetically lethal (Gregory et al., 2016). Thus, the simultaneous targeting of ATM-mediated G6PD regulation and inhibition of up-regulated nucleotide synthesis following chemotherapy induced stress may offer a new treatment option by decreasing DNA repair capacity. Following this, the targeting of key enzymes that regulate PPP also potentiated the effect of conventional therapies to selectively suppress cancer cell growth. For example, treatment with the glycolysis inhibitors 2-Deoxy-D-glucose (2DG) and 6-aminonicotinamide (6AN) has been shown to increase radio-sensitivity in squamous carcinoma cell lines (Khaitan et al., 2006 Sharma et al., 2012). In addition, this suggests that the inhibition of PPP or G6PD in combination with DNA damage inducing chemotherapies, such as 5-fluorouracil (5-FU) and doxorubicin, may restore chemosensitivity in cancer cells.

ATM and DNA-PK Kinases Play a Key Role in Cellular Energy Sensing

Ataxia telangiectasia mutated and DNA-dependent kinases (DNA-PK) are key proteins that recognize DNA damage and initiate DNA damage repair signaling (Mirzayans et al., 2006 Marechal and Zou, 2013). Upon activation by DNA-damage, these kinases generate a cascade of phosphorylation events that regulate the recruitment and activity of many downstream effector proteins to repair DNA double-strand breaks (DSBs) (Cosentino et al., 2011 Aird et al., 2015). ATM is generally considered to form a homodimer, while the active DNA-PK complex is comprised of the DNA-PK catalytic subunit bound to the Ku70/80 heterodimer. Several studies have shown that both DNA-PK and ATM are also involved in cellular metabolism rewiring after DNA damage for energy supply by activating of glucose transporter (GLUT4) thought AKT, maintenance of mitochondrial homeostasis and increased nucleotide production for DNA metabolism (Figure 2). This is particularly evident in individuals with Ataxia-Telangiectasia syndrome (A-T), which results from mutations in the ATM gene. These individuals exhibit alterations in cellular metabolism, including the dysfunction of enzymes involved in glucose metabolism and mitochondrial function (Sharma et al., 2014 Volkow et al., 2014).

Figure 2. A schematic representation of the Ataxia-Telangiectasia Mutated- (ATM) and DNA-Dependent Kinase- (DNA-PK) mediated regulation of metabolic processes after DNA Damage. ATM activates multiple downstream proteins regulating cell cycle arrest, DNA repair and cellular metabolism. ATM activates the tumor suppressor p53 which decreases GLUT recruitment, glycolysis, and dNTP production. ATM non-canonical function is essential for repairing mitochondrial genome defects to maintain mitochondrial homeostasis. For maintenance of energy production ATM activates AKT to promote glucose recruitment to the nucleus via GLUT4-mediated transport. ATM also activates G6PD though Hsp27, as an alternative mechanism to produce nucleotides for DNA metabolism. DNA-PK following energy depletion or metabolic rewiring promotes glycolysis through AMP signaling. This figure was created with

Besides its primary function in the recognition of DNA damage, ATM functions as a metabolic stress sensor, identifying reductions in the energy levels of tumor cells, subsequently promoting increased PPP activity, which can lead to increased cancer cell survival and resistance to conventional therapies (Krüger and Ralser, 2011). Additionally, there is a growing quantity of evidence showing that ATM also regulates the translocation of glucose transporter 4 (GLUT4), which in part explains why patients with A-T syndrome tend to present high incidences of type 2 diabetes mellitus (Halaby et al., 2008). It is known that cytoplasmic ATM is an insulin-responsive protein that activates AKT following insulin treatment, and inhibition of ATM leads to downregulation of AKT activity that in turn downregulates the GLUT4 glucose transporter protein (Halaby et al., 2008). A recent study found that loss of ATM-mediated p53 Ser18 (murine Ser15) phosphorylation led to increased metabolic stress and insulin resistance (Armata et al., 2010). Additionally, ATM was also shown to enhance glycolysis in breast cancer cells via GLUT1-phosphorylation and PKM2 up-regulation, increasing lactate production. High levels of lactate were found to promote tumor invasion through lactate-mediated metabolic coupling (Sun et al., 2019). These recent studies suggest that ATM is essential for glycolysis homeostasis as it regulates key metabolic proteins that are responsible for the maintenance of glucose levels such as glucose transporters.

DNA-dependent kinases is best known for recognizing DSBs and initiating DNA repair responses by activation of the NHEJ pathway. DNA-PK is an abundant, cytoplasmatic protein that migrates to the nucleus after DNA damage (Yang et al., 2014). There is also growing evidence indicating that DNA-PK may function to regulate metabolic homeostasis (Weterings and Chen, 2008 Lieber, 2010). Similar to ATM, DNA-PK also functions as a metabolic stress sensor and regulates AMPK (AMP-activated protein kinase) in response to energy depletion or metabolic stress in mammalian cells. AMPK is an essential protein that recognizes when energy production is low. It has been shown that inhibition of the DNA-PK catalytic subunit, decreases AMPK activity in response to energy deprivation.

Cell starvation leads to the phosphorylation of AMPKα (Thr172) and acetyl-CoA carboxylase (ACC). However, the inhibition of DNA-PKcs inhibits AMPK phosphorylation, thereby disrupting the sensing of glucose metabolism by AMPK. In addition, it was shown that DNA-PKcs directly interacts with the energy monitoring regulatory subunit of AMPK (Amatya et al., 2012). This finding suggests that DNA-PK is essential for activating AMPK under low energy levels as a result of glucose deprivation in mammalian cells. Similarly, another study confirmed that DNA-PKcs is a positive regulator of AMPK activity and was found to phosphorylate two residues on AMPKγ (S192 and T284) (Puustinen et al., 2020). Conversely, another study showed that an aging-related increase in DNA-PKcs activity led to decreased AMPK activity, via phosphorylation-mediated inhibition of Hsp90 chaperone activity toward AMPKα㬒 (Park et al., 2017). It is also possible that the interaction between DNA-PKcs and AMPK may depend on cellular context as DNA-PKcs itself is regulated by the cellular metabolic state and may decline as individuals’ age.

Autophagy is the process by which damaged proteins or organelles are degraded by the lysosome, this provides a mechanism to recycle cellular components providing macromolecular precursors and energy for cellular metabolism. Autophagy is generally classified into five defined steps: initiation, vesicle nucleation, vesicle elongation, vesicle fusion and cargo degradation. The regulation of autophagy by metabolic proteins and vice versa have been well charactered but there is also mounting evidence that DNA repair is also regulated by autophagy [reviewed in Hewitt and Korolchuk (2017)]. Some studies suggest that DNA repair is inhibited by autophagy, but other studies propose that autophagy promotes DNA repair (Bae and Guan, 2011 Liu et al., 2015). In order to explain this discrepancy, it has been hypothesized that following low levels of DNA damage, autophagy may promote DNA repair, while severe DNA damage may lead to autophagy-dependent degradation of DNA repair proteins to promote apoptosis (Guo and Ying, 2020). Autophagy has been shown to be initiated by AMPK activation and/or inhibition of the metabolic sensor Mammalian Target of Rapamycin Complex 1 (mTORC1), establishing another link between metabolism and DNA repair pathways (Kim et al., 2011). As discussed above the DNA-PK-dependent regulation of AMPK may also provide a feedback loop to regulate autophagy in the context of DNA repair (Puustinen et al., 2020).

Key Metabolic Enzymes That Play a Role in DNA Repair and Resistance to Chemo- and Radiotherapies (Summarized in Table 1)

Phosphoglycerate Mutase Enzyme (PGAM)

The Phosphoglycerate mutase 1 (PGAM1) is a key glycolytic enzyme that coordinates different metabolic process including glycolysis, PPP, and serine biosynthesis in cancer cells. As a result of its dynamic role in metabolic coordination, PGAM1 is overexpressed in several cancer types, including gliomas, oral carcinomas and pancreatic cancers (Liu et al., 2008, 2018 Zhang et al., 2017). For example, PGAM1 activity directly regulates the PPP and the resulting production of nucleotides, promoting cancer cell proliferation and tumor resistance to conventional therapies. Indirectly, PGAM1 contributes to DNA repair activity in cancer cells by the upregulation of glycolysis and/or nucleotide synthesis (Ohba et al., 2020). However, it was also found that PGAM1 plays a direct role in DNA repair as its activity was required for the repair of DNA double-strand breaks via homologous recombination (HR). Its role in HR was shown to be through regulating the stability of CTBP-interacting protein (CtIP), which is essential for the recruitment of Rad51 to sites of damage to facilitate filament formation (Qu et al., 2017). Complementary studies in gliomas cells demonstrated that depletion of PGAM1 also led to defective DNA damage signaling, including ATM autophosphorylation and phosphorylation of its downstream substrates. This led to disrupted DSB repair and subsequent sensitivity to IR, suggesting that PGAM1 may be a potential therapeutic target in gliomas (Ohba et al., 2020).

Table 1. The effects of metabolic proteins and metabolites on DNA repair.

Fumarase/Fumarate Hydratase (FH)

Under normal cellular conditions FH localizes mainly in the cellular cytosol and mitochondria (Kornberg and Krebs, 1957). A study in yeast demonstrated that following the induction of DNA damage, FH moves to the nucleus and functions as a DNA repair protein to promote the repair of DSBs (Yogev et al., 2010). In human cells, FH plays a similar role in DNA repair and was found to be a substrate of DNA-PK, which phosphorylates FH at Threonine 236. This stimulates the local generation of fumarate near DSBs, which inhibits the activity of the histone demethylase, KDM2B (Jiang et al., 2015). Subsequently, increasing the level of Histone H3 lysine 36 dimethylation which has been shown to facilitate the recruitment of DNA-PK to DSB sites and subsequently facilitate NHEJ activity (Fnu et al., 2011). A recently study showed that depletion of fumarase prolonged the interaction of Mre11 at sites of DSBs, delaying the progression of the HR pathway (Leshets et al., 2018). In addition, increased FH expression also disrupts HR by the inhibition of two key lysine demethylases (KDM4A and KDM4B) in Leiomyomatosis Renal Cell Cancer (HLRCC). This syndrome is classified as a familial DNA repair deficiency syndrome, as these patients carry a germline mutation in FH leading to defective responses to DNA damage and results in a higher predisposition for cancer development (Sulkowski et al., 2018).

Pyruvate Kinase M2 (PKM2)

Pyruvate kinase is an enzyme that converts phosphoenolpyruvate and ADP into pyruvate to generate ATP, and its activity is essential for the maintenance of glucose homeostasis. Pyruvate kinase M2 (PKM2) is highly expressed in cancer cells and a master regulator of tumor metabolic reprogramming (Wu et al., 2016 Zheng et al., 2018). Under normal conditions PKM2 is an abundant cytosolic protein that upon certain cellular stress, such as ultraviolet light (UV) or H2O2, migrates to the nucleus (Stetak et al., 2007). The migration of PKM2 to the nucleus has been associated with its non-metabolic functions, as PKM2 was found to phosphorylate several nuclear proteins, including histone H3 (Yang et al., 2014). It was also reported that nuclear PKM2 interacts with histone H2AX after DNA damage, and that PKM2 could directly phosphorylate H2AX on serine 139, one of the first phosphorylation events following DNA damage. Furthermore, replacement of wild type PKM2 with a kinase dead form led to increased chromosomal aberrations following DNA damage. Collectively, this reveals PKM2 as a novel modulator for genomic instability in tumor cells (Xia et al., 2017). As part of its non-metabolic activity it was also recently uncovered that PKM2 directly promotes DSB repair, as ATM phosphorylates PKM2 at Threonine 328 (T328) to induce the nuclear accumulation of PKM2 (Matsuoka et al., 2007). This ATM-mediated phosphorylation of PKM2 was shown to be required for efficient homologous recombination (HR) through the recruitment of CtIP at the site of DSBs. Additionally, the disruption of the ATM-PKM2-CtIP axis interaction was shown to sensitize tumor cells to a variety of DNA-damaging agents, including PARP inhibitors (Sizemore et al., 2018).

ATP-Citrate Lyase (ACLY)

ATP-citrate lyase is a nuclear-cytoplasmic enzyme that utilizes acetyl-CoA to generate citrate, and plays a crucial role in conserving the global histone acetylation in mammalian cells (Wellen et al., 2009). ACLY deficiency has been shown to result in defective DSB repair, due to the depletion of acetyl-CoA pools and reduction in acetylated histones at sites of DSBs (Kumari et al., 2019). Supporting this Sivanand et al. showed that nuclear acetyl-CoA played a role in HR and following DNA damage ACLY was phosphorylated at Serine 455, in an ATM- and AKT-dependent manner. Additionally, ACLY phosphorylation and nuclear localization were necessary to promote BRCA1 recruitment in order for HR to occur (Sivanand et al., 2017). Thus, acetyl-CoA production by ACLY is critical for the repair of DNA DSBs.

Glutamine Synthetase (GS)

Glutamine synthetase (GS) is an enzyme that catalyzes the conversion of glutamate and ammonia into glutamine. Transcriptome analyses revealed that GS is responsible for the metabolic reprogramming that occurs in tumor cells, as GS activity was shown to enhance DNA repair via de novo nucleotide synthesis (Kalluri, 2016). Further analyses revealed that knockdown of GS delayed DNA repair due to impaired nucleotide metabolism, which led to increased radio-sensitivity. HR was impaired in GS depleted cells further supporting a role for GS in DSB repair. Collectively, these findings suggest glutamine synthase plays a similar role to G6PD in DNA repair, as its upregulation increases nucleotide synthesis leading increased DSB repair capacity.

The Role of Metabolic Reprogramming in Tumor Cell Chemo- and Radio-Resistance

Radiotherapy remains a key anti-cancer therapy, with over 50% of patients undergoing radiation treatment as a monotherapy or in combination with other therapies (Kalluri, 2016). However, a significant proportion of patients experience resistance to conventional radiotherapy. Studies have demonstrated that the likelihood of radio-resistance is influenced by several factors, including metabolic changes and the upregulation of DNA repair pathways (Dwarkanath et al., 2001 Schwarz et al., 2008). Metabolic reprogramming may enable tumor cells to enhance nucleotide synthesis through the upregulation of the PPP, subsequently promoting resistance to traditional anti-cancer therapies (Zhao et al., 2016 Yin et al., 2017). Supporting this, several studies have shown that upregulation of metabolic enzymes or metabolic processes increases the activity of DNA repair pathways. For example, as a result of elevated glycolytic activity, some tumors generate a high level of lactate, which can promote cisplatin-resistance through increased DNA repair activity (Wagner et al., 2015). As previously discussed, several metabolic enzymes from glycolysis and PPP play a direct role in DNA repair pathways, and inhibition of key enzymes of both pathways not only inhibited cellular proliferation but also restored radio-sensitivity by decreasing DNA repair activity. The link between radio-resistance and altered metabolism is not fully understood but several studies suggest that decreasing the metabolic activity of the key enzymes involved in the PPP and glycolysis pathways could restore the sensitivity of resistant tumors to conventional therapies.

In ovarian cancer, three glycolytic enzymes, HK2, PFK, and PKM2, have been suggested to be promising targets due to their positive correlation with chemo- and radio-resistance via anti-apoptotic and cell survival mechanisms (Li et al., 2015 Zhang et al., 2018 Lin et al., 2019). There are four isoforms of PK however, the PKM2 isoform is a key regulator of glycolysis in cancer cells and is thus the most prominent potential candidate for restoring sensitivity to therapies. Supporting this, the inhibition of PKM2 in cervical cancer cells leads to decreased cell viability, G2/M cell cycle arrest, and promotes apoptosis (Lin et al., 2019). Furthermore, inhibition of PKM2 may induce radio-sensitivity, as demonstrated by a study which found that PKM2 depletion decreases AKT and PDK1 phosphorylation to subsequently promote radio-sensitivity in NSCLCs (Yuan et al., 2016). Similar to LDHA, miR-133 overexpression inhibits the expression of PKM2, which restores the sensitivity of radio-resistant lung cancer cells, offering a potential new treatment option for these radio-resistant tumors (Liu et al., 2016).

Hexokinase 2 (HK2) is a key glycolytic enzyme that catalyzes the first essential step of glucose metabolism. Like many other glycolytic proteins, HK2 is highly expressed in several tumor types (Anderson et al., 2017 Wu et al., 2017). Similar to other metabolic proteins, inhibition of HK2 has been shown to increase radio-sensitivity in cancer cells (Vartanian et al., 2016). 2-deoxy-D-glucose (2-DG) is an inhibitor of glucose metabolism, that is phosphorylated by Hexokinase to produce 2-deoxyglucose-6-phosphate. The intracellular accumulation of this metabolite inhibits hexokinase activity and therefore ATP production via glycolysis. Significantly, the anti-proliferative effects of 2-DG have been demonstrated in numerous preclinical studies (Giammarioli et al., 2012 Zhang et al., 2015). 2-DG has also been shown to be an effective sensitizer in several tumor types, including gliomas and lung carcinomas (Dwarkanath et al., 2001 Singh et al., 2019). Additionally, combining 2-DG with chemotherapy has already shown promising results in its ability to restore the sensitivity of chemo-resistance cells. A recent study analyzed the effect of combination treatment with 2-DG and carboplatin chemotherapy in high stage and recurrent ovarian clear cell carcinoma (OCC), and found that 2-DG in combination with carboplatin and cisplatin chemotherapy increased efficacy in chemo-resistant ovarian tumor cell lines and patient-derived xenograft models (Zhang et al., 2015 Khan et al., 2020). Thus, the combination of 2-DG with both radio- and chemotherapy drugs improves tumor cell sensitivity however, the underlying mechanism for the restoration of sensitivity to therapy remains largely unknown.

The glucose transporter GLUT1 is involved with the early steps of glucose uptake and metabolism. GLUT1 is overexpressed in many types of cancers and has been evaluated as a potential target for anti-cancer drugs (Wincewicz et al., 2010 Koch et al., 2015 Kim and Chang, 2019). Depletion of GLUT1 using small interfering RNA (siRNA) was shown to increase the radiosensitivity of laryngeal cancer cells and led to the downregulation of DNA repair. Similarly, restoration of radio-sensitivity was observed when antisense oligonucleotides (AS-ODNs) were used to inhibit GLUT1 activity in laryngeal carcinoma cells (Chan et al., 2004 Yan et al., 2013). In breast cancer, a synthetic inhibitor of GLUT1 known as WZB117, was demonstrated to radio-sensitize cancer by increasing the level of intracellular ROS, thereby inhibiting tumor growth (Zhao et al., 2016). Thus, inhibition of GLUT1 has therapeutic potential as an intervention to overcome cellular radio-resistance.

L-lactate is produced by glycolysis and is found to be expressed in high quantities in malignant tumors. High lactate levels have also been associated with resistance to clinical chemotherapeutics in numerous cancer subtypes. Recently, studies have shown that lactate can inhibit the activity of histone deacetylases (HDACs), which leads to changes in chromatin structure and transcription (Wagner et al., 2015, 2017). HDACs remove acetyl groups from histones, and their inhibition results in increased acetylation of histones, which are generally associated with a more open chromatin structure to promote transcription. This open chromatin state has also been suggested to increase accessibility of DNA repair proteins to sites of damage, in turn increasing the rates of DNA repair (Tamburini and Tyler, 2005). A study showed that lactate also modulates chromatin compaction in cervical cancer, leading to the up-regulation of DNA-PKcs (Wagner et al., 2015). Thus, the characteristic increase in lactate levels in tumor cells results in increased DNA repair activity, which has been shown to enhance radio-resistance in cervical carcinoma. Additionally, L/D-lactate was shown to increase the rate of γ-H2AX foci resolution after irradiation and induce cisplatin resistance, consistent with the up-regulation of DNA repair pathways (Wagner et al., 2015). Lactate dehydrogenase (LDHA) is a key metabolic protein found in almost all human tissues that is required for the conversion of pyruvate to lactic acid, playing an important role in the final steps of glycolysis. Increased expression of LDHA induces hypoxic environments that are associated with tumor metastases, poor overall survival, and radio-resistance in several tumor types, including prostate and bladder cancers (Koukourakis et al., 2009, 2014, 2016). Based on these findings, it can be suggested that the inhibition of LDHA activity may confer sensitivity in tumor cells to DNA damaging agents (Manerba et al., 2015). Supporting this, a soluble adenylate cyclase (sAC) that promotes the release of LDHA, led to the activation of the BRAF/ERK1/2 signaling pathway and consequently increased radio-resistance in prostate cancer cells (Flacke et al., 2013 Appukuttan et al., 2014). Treatment of prostate cancer cells with an LHDA-specific inhibitor, FX-11, reduced the activity of DNA repair proteins, improving cellular sensitivity to radiotherapy (Hao et al., 2016). Another study demonstrated that miR-34a overexpression inhibits LDHA and restored radio-sensitivity in hepatocellular carcinoma cells (Li et al., 2016). Based on these findings, it has been suggested that targeting LDHA via miR-34a may provide a mechanism to restore sensitivity to therapies in radio-resistant tumors (Li et al., 2016). Lactate influx and efflux is mediated by four members of the solute carrier 16a family Monocarboxylate transporters (MCT1-4). These proteins control the transport of lactate across the plasma membrane, effectively controlling lactate homeostasis. Given that high lactate levels confer chemo- and radioresistance, MCTs may also represent an effective mechanism to target lactate levels in tumor cells and increase sensitivity to DNA damaging agents (Halestrap, 2012).


Spatio-temporal regulation of gene expression determines many crucial developmental changes in plants. Gene regulation may occur at transcriptional, post-transcriptional, translational and post-translational levels. In the previous report, we have shown transcriptional level regulation of bHLH142 expression. Expression of GUS reporter gene driven by bHLH142 promoter was restricted to anther tissue in transgenic rice. However, in the Arabidopsis transgenic plants, bHLH142 promoter: GUS construct showed ubiquitous expression pattern 30 . This suggests that bHLH142 expression is transcriptionally regulated through its promoter and anther-specific activity is restricted only to rice or monocots. In this study, we have identified other aspects related to regulation of bHLH142 expression through protein level expression. We showed that there is a temporal difference between transcript and polypeptide accumulation of bHLH142, as its polypeptide was detected at high level in anthers at tetrad and MP stage, while negligible amount of the transcript was detected in anthers at MP stage. Previously, enhanced protein accumulation in germinating pollen compared to mature pollen was observed in case of LAT52 gene in tobacco and tomato 41 . Delayed detection of bHLH142 polypeptide in comparison to its transcripts suggested regulation of its expression beyond transcriptional level. We hypothesise that stage-specific translation enhancement might be the reason for higher accumulation of bHLH142 protein in MP anther. Such kind of translation enhancement has previously been observed during tobacco pollen development and germination 41,42 . Transient as well as stable expression studies of sequence present within 5′ UTR region of LAT52 gene caused increase in translational yield in developmentally regulated manner during pollen maturation 41 . Similarly, 5′ UTR region of NTP303 gene also enhanced translation in pollen tube but not in mature pollen 42 . We found sequence similarity between 5′ UTR of bHLH142, ntp303 and LAT52 genes suggesting similar type of translation enhancement in case of bHLH142 in MP anther (Figure S6). Apart from this, the other possible reason behind biphasic protein accumulation of bHLH142 may lie in the enhancement of its protein stability. In the recent report, proteomic and phospho-proteomic analysis of rice meiotic anthers showed presence of bHLH142 protein in phospho-proteome 43 . Detection of bHLH142 protein in anther phospho-proteome suggested its post-translational level regulation that may change its stability. Although these hypotheses need more experimental validations to reach any specific conclusion, above-mentioned findings suggest that expression of bHLH142 is regulated at transcriptional, post-transcriptional/translational and/or post-translational levels during rice anther development.

Previous reports showed that transgenic knock-down/mutant knock-out of bHLH142 resulted in male sterile plants because of no pollen formation 20,21 . Mutants were defective in tapetum development and degeneration process that led to the pollen abortion. Our experiments have shown that overexpression of bHLH142 also leads to completely male sterile plants, although compromised pollen grains were formed in these plants. Ko et al. 20 have shown that mutation in bHLH142 results in failure of tapetal PCD and overexpression in our study results in faster tapetal PCD. This reflects a role of bHLH142 in tapetum degeneration process. Furthermore, overexpression of bHLH142 also affected the anther dehiscence process, which caused complete male sterility. Thus, bHLH142 has other functions besides regulating tapteum degeneration during anther development in rice. The phenotype in overexpression transgenic rice is due to bHLH142 as transgenics with vector alone or transgenics made using the same vector but containing other genes have been found to be fertile 44 (our unpublished work).

During the pollen development, anther wall not only forms protective covering for developing microspore but the innermost layer of the wall, tapetum also provides nutrition to them 8 . During the release of pollen, anther wall again plays an important role and is involved in dehiscence process. Tapetum development and its degeneration have been well studied in rice 35 . However, mechanism of development and degeneration of other anther wall layers is still not clear. From this study, it is evident that bHLH142 regulates development and degeneration of different anther wall layers. Poorly developed and fast degenerating tapetum was seen in developing anther, whereas lack of endothecium lignification, septum and stomium (modifications of epidermal cells) degeneration was observed in mature anther of bHLH142 OE transgenic plants.

Role of bHLH142 in the development and degeneration of anther wall layers, including endothecium and modified epidermis (stomium and septum) was identified by this study. We have shown that bHLH142 polypeptide accumulated in the epidermis, pollen and vascular tissue of the mature anther and its overexpression caused defects in secondary endothecium thickening and degeneration of the septum and stomium. Lignification of endothecium appears after the vacuolated pollen stage but the genes required for the process express a little earlier 45 . Downregulation of the various lignin metabolism-related genes in tetrad anther might be the reason for absence of lignification in bHLH142 OE anthers. Furthermore, defect in septum and stomium lysis may have occurred because of various reasons. In most of the plants, enzymatic lysis of septum occurs with the help of cell wall degrading enzymes like polygalacturonases (PGs), pectinases and expansins 5,46,47 . Several cell wall modification-related genes like PGs, expansins and genes encoding pectin-degrading enzymes were found to be downregulated in transgenic anthers. Furthermore, role of PCD is also suggested in septum and stomium lysis 48 and downregulation of cell death-related genes in bHLH142 OE MP anther may have contributed to this phenotype. Moreover, water status in anther plays crucial role in pollen development and anther dehiscence 49 . Role of aquaporins and sugar transporters in inducing dehydration during anther dehiscence has been reported earlier 50 . We detected expression of bHLH142 protein in vascular bundle of MP anther. Four aquaporin genes and one sucrose transporter gene were found to be downregulated in MP stage of transgenic anther, which suggests that defects in water conduction might have occurred that affected anther dehiscence.

Besides regulating the anther wall functions, bHLH142 also appeared to regulate pollen maturation process as most of the pollen grains of bHLH142 OE plants were shrunken and non-viable. Several metabolic processes, like carbohydrate and lipid metabolism affect the pollen maturation process 39 . Complete maturation of pollen grains requires synthesis and transport of carbohydrate and lipids. Change in expression of various carbohydrate-related genes might have altered the synthesis and transport of sugars to the pollen grain, which resulted in their shrunken shape. Change in expression of sugar partitioning-related genes MST8, INV4, UGP2, SUT3 and AGPL1 40,51 suggests that defect in sugar loading/unloading may cause the formation of defective pollen in bHLH142 overexpressing plants. Lipid metabolism during the pollen maturation process is an important process as deposition of lipid is required for proper pollen wall formation. Lipid transfer proteins (LTPs) are known to be involved in pollen maturation process 36,39 . Change in expression of various lipid metabolism-related genes and several LTPs in bHLH142 overexpressing anther suggests that deposition of lipid on pollen wall might be altered, which contributed to the pollen abortion. CYP704B2 and CYP703A3, genes encoding cytochrome P450 enzymes, catalyse hydroxylation of fatty acid to assist the pollen wall development 36,38 . Both of these genes were found to have altered expression in bHLH142 OE transgenic anther. Furthermore, OsC4 and OsC6, important LTPs already known for anther development, were also differentially expressed in the bHLH142 OE anthers. ROS signalling has been observed to play an important role during anther cell differentiation and pollen development. Rice MADS3 has been shown to regulate ROS homeostasis during late anther development by directly regulating expression of metallothionin gene 52 . We detected main phenotypic difference in bHLH142 OE plants during later stage of pollen development. Downregulation of metallothionin, peroxidases and glutathione-s-transferase genes in bHLH142 overexpressing anthers suggests that alteration in ROS homeostasis may have contributed to defect in pollen maturation process. Therefore, it appears that bHLH142 affects metabolic processes like carbohydrate and lipid metabolism, cell wall modification, ROS homeostasis and cell death in rice anther to control both pollen development and anther dehiscence.

A putative model describing various functions of bHLH142 during anther development process in rice is given in Fig. 7. bHLH142 directly or indirectly affects various metabolic pathways-related genes to ensure proper anther development. It directly regulates expression of EAT1 and TDR 20 required for the tapetum development and degeneration. Furthermore, it directly or indirectly affects the various lipid and carbohydrate metabolism-related genes required for pollen wall development and accumulation of starch during pollen maturation. It also affects the selective deposition of lignin and cell-wall degeneration-related genes required for the anther dehiscence. Therefore, bHLH142 acts in different stages of rice anther development to control different processes required for proper development and release of the pollen grains. Thus, this study has explored the novel functions of bHLH142 in rice anther, which will contribute to the understanding of anther development process in crops to control male fertility for the hybrid seed production.

bHLH142 regulated various aspects of anther development in rice. It directly regulates the expression of EAT1 and TDR to control tapetum development and degeneration. It affects the lipid and carbohydrate metabolism-related genes to control pollen maturation and affects lignin and cell wall modification-related genes to control anther dehiscence.


Galdieri, L., Mehrotra, S., Yu, S. & Vancura, A. Transcriptional regulation in yeast during diauxic shift and stationary phase. OMICS 14, 629–638 (2010).

Marijuan, P. C., Navarro, J. & del Moral, R. On prokaryotic intelligence: strategies for sensing the environment. Biosystems 99, 94–103 (2010).

Schaller, G. E., Shiu, S. H. & Armitage, J. P. Two-component systems and their co-option for eukaryotic signal transduction. Curr. Biol. 21, R320–R330 (2011).

DeBerardinis, R. J. et al. Beyond aerobic glycolysis: transformed cells can engage in glutamine metabolism that exceeds the requirement for protein and nucleotide synthesis. Proc. Natl Acad. Sci. USA 104, 19345–19350 (2007).

Rheinwald, J. G. & Green, H. Epidermal growth factor and the multiplication of cultured human epidermal keratinocytes. Nature 265, 421–424 (1977).

Martin, P. Wound healing—aiming for perfect skin regeneration. Science 276, 75–81 (1997).

Nissen, N. N. et al. Vascular endothelial growth factor mediates angiogenic activity during the proliferative phase of wound healing. Am. J. Pathol. 152, 1445–1452 (1998).

Saltiel, A. R. & Kahn, C. R. Insulin signalling and the regulation of glucose and lipid metabolism. Nature 414, 799–806 (2001).

Frauwirth, K. A. et al. The CD28 signaling pathway regulates glucose metabolism. Immunity 16, 769–777 (2002).

Esensten, J. H., Helou, Y. A., Chopra, G., Weiss, A. & Bluestone, J. A. CD28 costimulation: from mechanism to therapy. Immunity 44, 973–988 (2016).

Barthel, A. et al. Regulation of GLUT1 gene transcription by the serine/threonine kinase Akt1. J. Biol. Chem. 274, 20281–20286 (1999).

Rathmell, J. C. et al. Akt-directed glucose metabolism can prevent Bax conformation change and promote growth factor-independent survival. Mol. Cell. Biol. 23, 7315–7328 (2003). This work demonstrates that growth factor signalling is necessary for mammalian cells to maintain glucose uptake in support of bioenergetics and cell survival.

Wieman, H. L., Wofford, J. A. & Rathmell, J. C. Cytokine stimulation promotes glucose uptake via phosphatidylinositol-3 kinase/Akt regulation of Glut1 activity and trafficking. Mol. Biol. Cell 18, 1437–1446 (2007).

Rathmell, J. C., Vander Heiden, M. G., Harris, M. H., Frauwirth, K. A. & Thompson, C. B. In the absence of extrinsic signals, nutrient utilization by lymphocytes is insufficient to maintain either cell size or viability. Mol. Cell 6, 683–692 (2000).

Fox, C. J., Hammerman, P. S. & Thompson, C. B. Fuel feeds function: energy metabolism and the T cell response. Nat. Rev. Immunol. 5, 844–852 (2005).

Warburg, O., Wind, F. & Negelein, E. The metabolism of tumors in the body. J. Gen. Physiol. 8, 519–530 (1927).

Warburg, O. On the origin of cancer cells. Science 123, 309–314 (1956).

Kandoth, C. et al. Mutational landscape and significance across 12 major cancer types. Nature 502, 333–339 (2013).

Recondo, G., Facchinetti, F., Olaussen, K. A., Besse, B. & Friboulet, L. Making the first move in EGFR-driven or ALK-driven NSCLC: first-generation or next-generation TKI? Nat. Rev. Clin. Oncol. 15, 694–708 (2018).

Arteaga, C. L. et al. Treatment of HER2-positive breast cancer: current status and future perspectives. Nat. Rev. Clin. Oncol. 9, 16–32 (2011).

Samuels, Y. et al. High frequency of mutations of the PIK3CA gene in human cancers. Science 304, 554 (2004).

Engelman, J. A., Luo, J. & Cantley, L. C. The evolution of phosphatidylinositol 3-kinases as regulators of growth and metabolism. Nat. Rev. Genet. 7, 606–619 (2006).

Lawrence, M. S. et al. Discovery and saturation analysis of cancer genes across 21 tumour types. Nature 505, 495–501 (2014).

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

Almuhaideb, A., Papathanasiou, N. & Bomanji, J. 18F-FDG PET/CT imaging in oncology. Ann. Saudi Med. 31, 3–13 (2011).

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

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

Hui, S. et al. Glucose feeds the TCA cycle via circulating lactate. Nature 551, 115–118 (2017). Faubert et al. (2017) and Hui et al. (2017) provide the first evidence that lactate in the circulation can act as the major carbon source to fuel the TCA cycle in vivo.

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

Saxton, R. A. & Sabatini, D. M. mTOR signaling in growth, metabolism, and disease. Cell 168, 960–976 (2017).

Inoki, K., Li, Y., Zhu, T., Wu, J. & Guan, K. L. TSC2 is phosphorylated and inhibited by Akt and suppresses mTOR signalling. Nat. Cell Biol. 4, 648–657 (2002).

Manning, B. D., Tee, A. R., Logsdon, M. N., Blenis, J. & Cantley, L. C. Identification of the tuberous sclerosis complex-2 tumor suppressor gene product tuberin as a target of the phosphoinositide 3-kinase/akt pathway. Mol. Cell 10, 151–162 (2002).

Menon, S. et al. Spatial control of the TSC complex integrates insulin and nutrient regulation of mTORC1 at the lysosome. Cell 156, 771–785 (2014).

Long, X., Lin, Y., Ortiz-Vega, S., Yonezawa, K. & Avruch, J. Rheb binds and regulates the mTOR kinase. Curr. Biol. 15, 702–713 (2005).

Edinger, A. L. & Thompson, C. B. Akt maintains cell size and survival by increasing mTOR-dependent nutrient uptake. Mol. Biol. Cell 13, 2276–2288 (2002). This study shows that AKT promotes cell survival and growth in part by maintaining nutrient transporter levels on the plasma membrane.

Hannan, K. M. et al. mTOR-dependent regulation of ribosomal gene transcription requires S6K1 and is mediated by phosphorylation of the carboxy-terminal activation domain of the nucleolar transcription factor UBF. Mol. Cell. Biol. 23, 8862–8877 (2003).

Wolfson, R. L. & Sabatini, D. M. The dawn of the age of amino acid sensors for the mTORC1 pathway. Cell Metab. 26, 301–309 (2017).

Saxton, R. A. et al. Structural basis for leucine sensing by the Sestrin2-mTORC1 pathway. Science 351, 53–58 (2016).

Wolfson, R. L. et al. Sestrin2 is a leucine sensor for the mTORC1 pathway. Science 351, 43–48 (2016).

Chantranupong, L. et al. The CASTOR proteins are arginine sensors for the mTORC1 pathway. Cell 165, 153–164 (2016).

Gu, X. et al. SAMTOR is an S-adenosylmethionine sensor for the mTORC1 pathway. Science 358, 813–818 (2017).

Lu, P. D., Harding, H. P. & Ron, D. Translation reinitiation at alternative open reading frames regulates gene expression in an integrated stress response. J. Cell Biol. 167, 27–33 (2004).

Harding, H. P. et al. An integrated stress response regulates amino acid metabolism and resistance to oxidative stress. Mol. Cell 11, 619–633 (2003).

Sato, H. et al. Transcriptional control of cystine/glutamate transporter gene by amino acid deprivation. Biochem. Biophys. Res. Commun. 325, 109–116 (2004).

Lewerenz, J. & Maher, P. Basal levels of eIF2alpha phosphorylation determine cellular antioxidant status by regulating ATF4 and xCT expression. J. Biol. Chem. 284, 1106–1115 (2009).

Lopez, A. B. et al. A feedback transcriptional mechanism controls the level of the arginine/lysine transporter cat-1 during amino acid starvation. Biochem. J. 402, 163–173 (2007).

Nicklin, P. et al. Bidirectional transport of amino acids regulates mTOR and autophagy. Cell 136, 521–534 (2009).

Zhang, N. et al. Increased amino acid uptake supports autophagy-deficient cell survival upon glutamine deprivation. Cell Rep. 23, 3006–3020 (2018).

Shan, J. et al. The C/ebp-Atf response element (CARE) location reveals two distinct Atf4-dependent, elongation-mediated mechanisms for transcriptional induction of aminoacyl-tRNA synthetase genes in response to amino acid limitation. Nucleic Acids Res. 44, 9719–9732 (2016).

Ye, J. et al. GCN2 sustains mTORC1 suppression upon amino acid deprivation by inducing Sestrin2. Genes Dev. 29, 2331–2336 (2015).

Siu, F., Bain, P. J., LeBlanc-Chaffin, R., Chen, H. & Kilberg, M. S. ATF4 is a mediator of the nutrient-sensing response pathway that activates the human asparagine synthetase gene. J. Biol. Chem. 277, 24120–24127 (2002).

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

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

Possemato, R. et al. Functional genomics reveal that the serine synthesis pathway is essential in breast cancer. Nature 476, 346–350 (2011). Locasale et al. (2011) and Possemato et al. (2011) provide early evidence that the serine synthesis pathway is upregulated in cancer and that it is critical in mediating tumorigenesis.

Pacold, M. E. et al. A PHGDH inhibitor reveals coordination of serine synthesis and one-carbon unit fate. Nat. Chem. Biol. 12, 452–458 (2016).

Mullarky, E. et al. Identification of a small molecule inhibitor of 3-phosphoglycerate dehydrogenase to target serine biosynthesis in cancers. Proc. Natl Acad. Sci. USA 113, 1778–1783 (2016).

Yang, M. & Vousden, K. H. Serine and one-carbon metabolism in cancer. Nat. Rev. Cancer 16, 650–662 (2016).

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

Maddocks, O. D. K. et al. Modulating the therapeutic response of tumours to dietary serine and glycine starvation. Nature 544, 372–376 (2017).

Lane, A. N. & Fan, T. W. Regulation of mammalian nucleotide metabolism and biosynthesis. Nucleic Acids Res. 43, 2466–2485 (2015).

Garcia-Bermudez, J. et al. Aspartate is a limiting metabolite for cancer cell proliferation under hypoxia and in tumours. Nat. Cell Biol. 20, 775–781 (2018).

Birsoy, K. et al. An essential role of the mitochondrial electron transport chain in cell proliferation is to enable aspartate synthesis. Cell 162, 540–551 (2015).

Sullivan, L. B. et al. Supporting aspartate biosynthesis is an essential function of respiration in proliferating cells. Cell 162, 552–563 (2015). Birsoy et al. (2015) and Sullivan et al. (2015) demonstrate that a major function of the mitochondrial ETC is to support aspartate production.

Lauren, P. et al. Mitochondrial complex III is necessary for endothelial cell proliferation during angiogenesis. Nat. Metab. 1, 158–171 (2019).

Liu, X. et al. Regulation of mitochondrial biogenesis in erythropoiesis by mTORC1-mediated protein translation. Nat. Cell Biol. 19, 626–638 (2017).

Weinberg, S. E. et al. Mitochondrial complex III is essential for suppressive function of regulatory T cells. Nature 565, 495–499 (2019).

Alistar, A. et al. Safety and tolerability of the first-in-class agent CPI-613 in combination with modified FOLFIRINOX in patients with metastatic pancreatic cancer: a single-centre, open-label, dose-escalation, phase 1 trial. Lancet Oncol. 18, 770–778 (2017).

Molina, J. R. et al. An inhibitor of oxidative phosphorylation exploits cancer vulnerability. Nat. Med. 24, 1036–1046 (2018).

Sullivan, L. B. et al. Aspartate is an endogenous metabolic limitation for tumour growth. Nat. Cell Biol. 20, 782–788 (2018).

Swanson, J. A. Shaping cups into phagosomes and macropinosomes. Nat. Rev. Mol. Cell Biol. 9, 639–649 (2008).

Recouvreux, M. V. & Commisso, C. Macropinocytosis: a metabolic adaptation to nutrient stress in cancer. Front. Endocrinol. 8, 261 (2017).

Finicle, B. T., Jayashankar, V. & Edinger, A. L. Nutrient scavenging in cancer. Nat. Rev. Cancer 18, 619–633 (2018).

Bloomfield, G. & Kay, R. R. Uses and abuses of macropinocytosis. J. Cell Sci. 129, 2697–2705 (2016).

Haigler, H. T., McKanna, J. A. & Cohen, S. Rapid stimulation of pinocytosis in human carcinoma cells A-431 by epidermal growth factor. J. Cell Biol. 83, 82–90 (1979).

Palm, W., Araki, J., King, B., DeMatteo, R. G. & Thompson, C. B. Critical role for PI3-kinase in regulating the use of proteins as an amino acid source. Proc. Natl Acad. Sci. USA 114, E8628–E8636 (2017).

Bar-Sagi, D. & Feramisco, J. R. Induction of membrane ruffling and fluid-phase pinocytosis in quiescent fibroblasts by ras proteins. Science 233, 1061–1068 (1986).

Commisso, C. et al. Macropinocytosis of protein is an amino acid supply route in Ras-transformed cells. Nature 497, 633–637 (2013). This study shows that macropinocytosis promotes the use of extracellular protein as an amino acid source in RAS-transformed cells.

Palm, W. et al. The Utilization of Extracellular Proteins as Nutrients Is Suppressed by mTORC1. Cell 162, 259–270 (2015). This work demonstrates that mTORC1 activity suppresses the use of extracellular protein as an amino acid source through macropinocytosis.

Garcia, D. & Shaw, R. J. AMPK: mechanisms of cellular energy sensing and restoration of metabolic balance. Mol. Cell 66, 789–800 (2017).

Kim, S. M. et al. PTEN deficiency and AMPK activation promote nutrient scavenging and anabolism in prostate cancer cells. Cancer Discov. 8, 866–883 (2018).

Nofal, M., Zhang, K., Han, S. & Rabinowitz, J. D. mTOR inhibition restores amino acid balance in cells dependent on catabolism of extracellular protein. Mol. Cell 67, 936–946 (2017).

Goldberg, I. J., Eckel, R. H. & Abumrad, N. A. Regulation of fatty acid uptake into tissues: lipoprotein lipase- and CD36-mediated pathways. J. Lipid Res. 50, S86–S90 (2009).

Menendez, J. A. & Lupu, R. Fatty acid synthase and the lipogenic phenotype in cancer pathogenesis. Nat. Rev. Cancer 7, 763–777 (2007).

Zaidi, N. et al. Lipogenesis and lipolysis: the pathways exploited by the cancer cells to acquire fatty acids. Prog. Lipid Res. 52, 585–589 (2013).

Medes, G., Thomas, A. & Weinhouse, S. Metabolism of neoplastic tissue. IV. A study of lipid synthesis in neoplastic tissue slices in vitro. Cancer Res. 13, 27–29 (1953).

Gansler, T. S., Hardman, W. 3rd, Hunt, D. A., Schaffel, S. & Hennigar, R. A. Increased expression of fatty acid synthase (OA-519) in ovarian neoplasms predicts shorter survival. Hum. Pathol. 28, 686–692 (1997).

Sebastiani, V. et al. Fatty acid synthase is a marker of increased risk of recurrence in endometrial carcinoma. Gynecol. Oncol. 92, 101–105 (2004).

Visca, P. et al. Fatty acid synthase (FAS) is a marker of increased risk of recurrence in lung carcinoma. Anticancer Res. 24, 4169–4173 (2004).

Mashima, T., Seimiya, H. & Tsuruo, T. De novo fatty-acid synthesis and related pathways as molecular targets for cancer therapy. Br. J. Cancer 100, 1369–1372 (2009).

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

Zhang, M. et al. Adipocyte-derived lipids mediate melanoma progression via FATP proteins. Cancer Discov. 8, 1006–1025 (2018).

Sounni, N. E. et al. Blocking lipid synthesis overcomes tumor regrowth and metastasis after antiangiogenic therapy withdrawal. Cell Metab. 20, 280–294 (2014).

Hochachka, P. W., Rupert, J. L., Goldenberg, L., Gleave, M. & Kozlowski, P. Going malignant: the hypoxia-cancer connection in the prostate. Bioessays 24, 749–757 (2002).

Ackerman, D. & Simon, M. C. Hypoxia, lipids, and cancer: surviving the harsh tumor microenvironment. Trends Cell Biol. 24, 472–478 (2014).

Zhang, Z., Dales, N. A. & Winther, M. D. Opportunities and challenges in developing stearoyl-coenzyme A desaturase-1 inhibitors as novel therapeutics for human disease. J. Med. Chem. 57, 5039–5056 (2014).

Igal, R. A. Stearoyl CoA desaturase-1: new insights into a central regulator of cancer metabolism. Biochim. Biophys. Acta 1861, 1865–1880 (2016).

Kamphorst, J. J. et al. Hypoxic and Ras-transformed cells support growth by scavenging unsaturated fatty acids from lysophospholipids. Proc. Natl Acad. Sci. USA 110, 8882–8887 (2013).

Ackerman, D. et al. Triglycerides promote lipid homeostasis during hypoxic stress by balancing fatty acid saturation. Cell Rep. 24, 2596–2605 (2018).

Vriens, K. et al. Evidence for an alternative fatty acid desaturation pathway increasing cancer plasticity. Nature 566, 403–406 (2019).

Nosal, J. M., Switzer, R. L. & Becker, M. A. Overexpression, purification, and characterization of recombinant human 5-phosphoribosyl-1-pyrophosphate synthetase isozymes I and II. J. Biol. Chem. 268, 10168–10175 (1993).

Qian, X. et al. Conversion of PRPS hexamer to monomer by AMPK-mediated phosphorylation inhibits nucleotide synthesis in response to energy stress. Cancer Discov. 8, 94–107 (2018).

Graves, L. M. et al. Regulation of carbamoyl phosphate synthetase by MAP kinase. Nature 403, 328–332 (2000).

Sigoillot, F. D. et al. Nuclear localization and mitogen-activated protein kinase phosphorylation of the multifunctional protein CAD. J. Biol. Chem. 280, 25611–25620 (2005).

Ben-Sahra, I., Howell, J. J., Asara, J. M. & Manning, B. D. Stimulation of de novo pyrimidine synthesis by growth signaling through mTOR and S6K1. Science 339, 1323–1328 (2013).

Robitaille, A. M. et al. Quantitative phosphoproteomics reveal mTORC1 activates de novo pyrimidine synthesis. Science 339, 1320–1323 (2013).

Wang, Y. & Hekimi, S. Understanding ubiquinone. Trends Cell Biol. 26, 367–378 (2016).

White, R. M. et al. DHODH modulates transcriptional elongation in the neural crest and melanoma. Nature 471, 518–522 (2011).

Sykes, D. B. et al. Inhibition of dihydroorotate dehydrogenase overcomes differentiation blockade in acute myeloid leukemia. Cell 167, 171–186 (2016).

Ducker, G. S. & Rabinowitz, J. D. One-carbon metabolism in health and disease. Cell Metab. 25, 27–42 (2017).

Ben-Sahra, I., Hoxhaj, G., Ricoult, S. J. H., Asara, J. M. & Manning, B. D. mTORC1 induces purine synthesis through control of the mitochondrial tetrahydrofolate cycle. Science 351, 728–733 (2016).

Shi, Y., Evans, J. E. & Rock, K. L. Molecular identification of a danger signal that alerts the immune system to dying cells. Nature 425, 516–521 (2003).

Boeynaems, J. M. & Communi, D. Modulation of inflammation by extracellular nucleotides. J. Invest. Dermatol. 126, 943–944 (2006).

Scheibner, K. A. et al. Hyaluronan fragments act as an endogenous danger signal by engaging TLR2. J. Immunol. 177, 1272–1281 (2006).

Singer, A. J. & Clark, R. A. Cutaneous wound healing. N. Engl. J. Med. 341, 738–746 (1999).

Kalluri, R. The biology and function of fibroblasts in cancer. Nat. Rev. Cancer 16, 582–598 (2016).

Dvorak, H. F. Tumors: wounds that do not heal-redux. Cancer Immunol. Res. 3, 1–11 (2015).

Sullivan, W. J. et al. Extracellular matrix remodeling regulates glucose metabolism through TXNIP destabilization. Cell 175, 117–132 (2018).

Camps, J. L. et al. Fibroblast-mediated acceleration of human epithelial tumor growth in vivo. Proc. Natl Acad. Sci. USA 87, 75–79 (1990).

Olumi, A. F. et al. Carcinoma-associated fibroblasts direct tumor progression of initiated human prostatic epithelium. Cancer Res. 59, 5002–5011 (1999).

Curtis, M. et al. Fibroblasts mobilize tumor cell glycogen to promote proliferation and metastasis. Cell Metab. 29, 141–155 (2018).

Keith, B. & Simon, M. C. Hypoxia-inducible factors, stem cells, and cancer. Cell 129, 465–472 (2007).

Trabold, O. et al. Lactate and oxygen constitute a fundamental regulatory mechanism in wound healing. Wound Repair Regen. 11, 504–509 (2003).

Kamphorst, J. J. et al. Human pancreatic cancer tumors are nutrient poor and tumor cells actively scavenge extracellular protein. Cancer Res. 75, 544–553 (2015).

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

Muranen, T. et al. Starved epithelial cells uptake extracellular matrix for survival. Nat. Commun. 8, 13989 (2017).

Olivares, O. et al. Collagen-derived proline promotes pancreatic ductal adenocarcinoma cell survival under nutrient limited conditions. Nat. Commun. 8, 16031 (2017).

Katheder, N. S. & Rusten, T. E. Microenvironment and tumors-a nurturing relationship. Autophagy 13, 1241–1243 (2017).

Sousa, C. M. et al. Pancreatic stellate cells support tumour metabolism through autophagic alanine secretion. Nature 536, 479–483 (2016). This work shows that autophagic degradation and release of cellular constituents in pancreatic stellate cells can support the growth of pancreatic cancer cells.

Nieman, K. M. et al. Adipocytes promote ovarian cancer metastasis and provide energy for rapid tumor growth. Nat. Med. 17, 1498–1503 (2011).

Rabinowitz, J. D. & White, E. Autophagy and metabolism. Science 330, 1344–1348 (2010).

Kaur, J. & Debnath, J. Autophagy at the crossroads of catabolism and anabolism. Nat. Rev. Mol. Cell Biol. 16, 461–472 (2015).

Wyant, G. A. et al. NUFIP1 is a ribosome receptor for starvation-induced ribophagy. Science 360, 751–758 (2018).

Amaravadi, R., Kimmelman, A. C. & White, E. Recent insights into the function of autophagy in cancer. Genes Dev. 30, 1913–1930 (2016).

Karsli-Uzunbas, G. et al. Autophagy is required for glucose homeostasis and lung tumor maintenance. Cancer Discov. 4, 914–927 (2014).

Guo, J. Y. et al. Autophagy provides metabolic substrates to maintain energy charge and nucleotide pools in Ras-driven lung cancer cells. Genes Dev. 30, 1704–1717 (2016).

Yang, S. et al. Pancreatic cancers require autophagy for tumor growth. Genes Dev. 25, 717–729 (2011).

Yang, A. et al. Autophagy is critical for pancreatic tumor growth and progression in tumors with p53 alterations. Cancer Discov. 4, 905–913 (2014).

Yang, A. et al. Autophagy sustains pancreatic cancer growth through both cell-autonomous and nonautonomous mechanisms. Cancer Discov. 8, 276–287 (2018).

Poillet-Perez, L. et al. Autophagy maintains tumour growth through circulating arginine. Nature 563, 569–573 (2018).

Allen, E. L. et al. Differential aspartate usage identifies a subset of cancer cells particularly dependent on OGDH. Cell Rep. 17, 876–890 (2016).

Ilic, N. et al. PIK3CA mutant tumors depend on oxoglutarate dehydrogenase. Proc. Natl Acad. Sci. USA 114, E3434–E3443 (2017).

Ye, J. et al. Serine catabolism regulates mitochondrial redox control during hypoxia. Cancer Discov. 4, 1406–1417 (2014).

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

Shin, C. S. et al. The glutamate/cystine xCT antiporter antagonizes glutamine metabolism and reduces nutrient flexibility. Nat. Commun. 8, 15074 (2017).

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

Tonks, N. K. Redox redux: revisiting PTPs and the control of cell signaling. Cell 121, 667–670 (2005).

Tormos, K. V. et al. Mitochondrial complex III ROS regulate adipocyte differentiation. Cell Metab. 14, 537–544 (2011).

Weinberg, F. et al. Mitochondrial metabolism and ROS generation are essential for Kras-mediated tumorigenicity. Proc. Natl Acad. Sci. USA 107, 8788–8793 (2010).

Schafer, Z. T. et al. Antioxidant and oncogene rescue of metabolic defects caused by loss of matrix attachment. Nature 461, 109–113 (2009).

DeNicola, G. M. et al. Oncogene-induced Nrf2 transcription promotes ROS detoxification and tumorigenesis. Nature 475, 106–109 (2011).

Harris, I. S. et al. Glutathione and thioredoxin antioxidant pathways synergize to drive cancer initiation and progression. Cancer Cell 27, 211–222 (2015).

Piskounova, E. et al. Oxidative stress inhibits distant metastasis by human melanoma cells. Nature 527, 186–191 (2015). This work demonstrates that melanoma cells need to overcome the oxidative stress barrier in order to successfully metastasize.

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

Parker, W. B. Enzymology of purine and pyrimidine antimetabolites used in the treatment of cancer. Chem. Rev. 109, 2880–2893 (2009).

Cantley, L. C. The phosphoinositide 3-kinase pathway. Science 296, 1655–1657 (2002).

Manning, B. D. & Cantley, L. C. AKT/PKB signaling: navigating downstream. Cell 129, 1261–1274 (2007).

Manning, B. D. & Toker, A. AKT/PKB signaling: navigating the network. Cell 169, 381–405 (2017).

Vivanco, I. & Sawyers, C. L. The phosphatidylinositol 3-Kinase AKT pathway in human cancer. Nat. Rev. Cancer 2, 489–501 (2002).

Metabolic Pathways for Intermediary Metabolism (3 Pathways)

The following points highlight the three main metabolic pathways for intermediary metabolism. The metabolic pathways are: 1. Carbohydrate Metabolism 2. Lipid Metabolism 3. Amino Acid Metabolism.

Metabolic Pathway # 1. Carbohydrate Metabolism:

a. Pyruvate and lactate are formed in the mammalian cells as a result of the oxida­tion of glucose by glycolysis.

b. Glycolysis occurs in the cytoplasm of cells in absence of oxygen producing lactate only. s

c. Under aerobic condition, pyruvate is me­tabolized to acetyl-CoA which enters the citric acid cycle for complete oxidation to CO2 and H2O.

d. Glucose also takes part in other metabolic process as follows:

(i) It is converted to glycogen as a stor­age, particularly, in liver and skeletal muscle.

(ii) The HMP shunt or the pentose phos­phate pathways arising from interme­diates of glycolysis is a source of re­ducing equivalents (2H) for biosyn­thesis of fatty acids, cholesterol, etc. and it is a source of ribose which is important for nucleic acid formations.

(iii) Triose phosphate of glycolysis is a source of glycerol of fat.

(iv) Pyruvate and the intermediates of citric acid cycle form amino acids and acetyl-CoA is the building block for long-chain fatty acids and choles­terol, the precursor of all steroid hormones in the body.

Metabolic Pathway # 2. Lipid Metabolism:

a. The long chain fatty acids are synthesized form acetyl-CoA derived from carbohy­drate or from dietary lipid.

b. In the tissues, fatty acids are oxidized to acetyl-CoA or esterified to acyl-glycerol to form fat which is the main caloric reserve of the body.

c. Acetyl -CoA formed by β-oxidation has the following significant roles in the body:

(i) It liberates CP2 and H2O and also yields high energy. Therefore, during the oxidation of fatty acids by β-oxi­dation for their complete oxidation, more energy is formed.

(ii) It is a source of cholesterol biosyn­thesis.

(iii) In the liver, it forms ketone bodies which are alternative water-soluble tissue fuels. These fuels become im­portant sources of energy under cer­tain conditions (e.g., starvation).

Metabolic Pathway # 3. Amino Acid Metabolism:

a. Amino acids are required for protein syn­thesis.

b. The essential amino acids must be sup­plied in the diet since these are not syn­thesized by the tissues.

c. Diet can supply the non-essential amino acids which are also formed from the intermediates of citric acid cycle by transamination.

d. Excess amino nitrogen as a result of deamination of amino acids is removed as urea and the carbon skeletons that remain after transamination give the following products:

Exploiting Treg Metabolism for Tolerance-Inducing Therapies

A better understanding of Treg metabolism and its distinction from other T cell subsets metabolism allows for the specific modulation of Treg in vivo or the improvement of adoptive Treg transfusion therapies. The encompassing goal of such therapies has been the induction of functional immunotolerance by harboring the natural specific immunosuppressive mechanisms without requiring damaging immunosuppressive drugs (74). Treg therapies could improve the current standard of care in the reduction of cost, increased availability, specificity to destructive immune responses, and applicability to different organs. However, before employing such therapies, it is imperative to understand the molecular mechanisms underlying critical Treg functionalities and to identify any factors that confound outcomes. For successful Treg therapy, it is rudimental to acquire a sufficient number of Treg, that these Treg migrate to their desired location and to subsequently have stable immunosuppressive functionality of Treg (75). Treg metabolism could be employed for this (Figure 3). Drugs for modulating cellular metabolism are already available, providing the field of immunometabolism with great opportunity to translate their findings to the clinic (Table 2).

Table 2. Metabolic modulators and their relevance for Treg proliferation, migration, and suppressive function.

Glycolysis is important for Treg migration. Although most research points toward a negative role for glycolysis in Treg proliferation, it has become clear that complete depletion of glucose from cell culture medium is detrimental for in vitro Treg proliferation and suppressive function. Inhibition of glycolysis reduces intracellular pyruvate levels, thereby preventing the conversion to acetyl-CoA via PDHK for mitochondrial oxidative metabolism (Figure 2) and can consequently reduce Treg proliferation (77). Interestingly, glycolysis can potentially be modulated in a Treg-specific manner, as Treg convert glucose to glucose-6-phosphate with a distinct isoform of hexokinase, hexokinase 1 (HK1) (19). At present, pharmacological interference with glycolysis can be obtained through 2-deoxy-D-glucose (2-DG), a glucose analog which is currently used at high concentrations in cancer therapy. 2-DG reportedly inhibits both Treg migration and proliferation (14, 15, 19, 35). Contrastingly, D-mannose, a C-2 epimer of glucose which also inhibits glycolysis, has been described to increase human Treg proliferation in vitro by the promotion of TGF-β activity which in turn leads to the increase of fatty acid oxidation (76). Mycophenolic acid, the active ingredient of the immunosuppressant mycophenolate mofetil currently in use for suppressing solid organ rejection, inhibits monophosphate dehydrogenase, an enzyme involved in the biosynthesis of guanine nucleotides, which follows the glycolysis-parallel pathway PPP (79). Interestingly, mycophenolic acid-enhanced expression of PD-1, CTLA-4, and FOXP3 and reduced Akt-mTOR and STAT5 signaling in human CD4 + T cells.

Fatty acid oxidation is increased in proliferative Treg and inhibition thereof results in decreased Treg numbers (81). Moreover, oxidative metabolism is associated with and can be induced by Treg suppressive molecules such as TGF-β, CTLA-4, PD-1, and FOXP3. Dimethyl fumarate, a derivate of the TCA cycle intermediate fumarate, stimulates proliferation and development of Treg by supporting mitochondrial oxidative metabolism and FOXP3. Of note, dimethyl fumarate causes lymphopenia and selectively depletes highly glycolytic Teff while sparing oxidative naïve T cells and Treg (80). OXPHOS can be increased by the immunomodulatory metabolite rapamycin, which is used in vitro to expand Treg, potently suppresses T cell proliferation and increases Treg suppressive function in vitro and in vivo. This suggests that it is a valuable drug for adjuvant therapy to improve the efficacy of T(reg)-based immunosuppressive protocols (84). However, rapamycin also redirects Teff peripheral tissue trafficking and stimulates homing to lymph nodes by inhibition of mTORC1 (85). Whether rapamycin also redirects Treg migration is not yet established.

In a situation where blocking Treg function is preferred such as in the tumor milieu, the widely used anti-diabetic drug metformin can be employed. Metformin inhibits the electron transport chain and decreases mitochondrial ROS production, both AMPK-dependent and independent. Metformin increases Treg differentiation, most likely via suppressed activation of mTOR and HIF-1α and stimulation of AMPK and FOXP3 expression (23). Conversely, CPT1 inhibitor etomoxir inhibits fatty acid oxidation, which specifically reduced Treg differentiation and proliferation, although this might be caused by the off-target effects of etomoxir on metabolism. The chines herbal compound celastrol also has immunosuppressive capacities by promoting fatty acid oxidation via upregulation of CPT1 and AMPK expression. Additionally, celastrol has been indicated to facilitate FOXP3 expression and Treg cell generation (82). Pharmacological inhibition of acid sphingomyelinase (ASM), with a clinically used tricyclic antidepressant like amitriptyline, induces higher frequencies of Treg among T cells. This is due to ASM inhibition increasing cell death of T cells in general, while CD25 high Treg are protected via IL-2 (83). Further, ASM deficient pTreg have less Akt activity and RICTOR levels compared with control pTreg. Inhibitors of the rate-limiting enzyme HMGR impairs Treg proliferation and function whereas addition of mevalonate, the metabolite downstream of HMGR restores Treg-mediated suppression (63), suggesting that manipulation of lipid biosynthesis, in particular via the mevalonate pathway, would result in Treg functional disruption.

Various studies have described the alterations of metabolic pathways and key metabolic byproducts in several autoimmune disorders. Disease-specific metabolic changes in overall glycolytic activity and oxidative state have been reported in rheumatoid arthritis and multiple sclerosis. In multiple sclerosis, impaired proliferation is suggested to be a consequence of increased levels of circulating leptin. The glutaminolysis pathway has been suggested as a biomarker for disease severity (86). In solid organ transplantation, it is reported that the metabolic environment might influence immune responses and overall transplantation outcome. Lee et al. have shown that by simultaneously blocking glycolysis and the glutamine pathway in the inflammatory transplantation microenvironment, allo-specific Teff responses could safely be reduced while preserving immunoregulation (87). Indeed, both glycolysis and glutamine are associated with a pro-inflammatory phenotype and non-essential for Treg suppressive function, although not irrelevant for Treg proliferation. Wawman and colleagues have described the importance of the hepatic microenvironment in transplantation. The continuous exposure of metabolites and nutrients influences lineage fitness, function, proliferation, migration, and survival of Treg (88). This paves the way for safe and specific novel approaches to modulate the inflammatory environment, for example with tissue-specific accumulation of nanobiologicals (89).

Feedback Inhibition in Metabolic Pathways

Molecules can regulate enzyme function in many ways. A major question remains, however: What are these molecules and where do they come from? Some are cofactors and coenzymes, ions, and organic molecules, as you’ve learned. What other molecules in the cell provide enzymatic regulation, such as allosteric modulation, and competitive and noncompetitive inhibition?

The answer is that a wide variety of molecules can perform these roles. Some of these molecules include pharmaceutical and non-pharmaceutical drugs, toxins, and poisons from the environment. Perhaps the most relevant sources of enzyme regulatory molecules, with respect to cellular metabolism, are the products of the cellular metabolic reactions themselves.

Using Reaction Products

In a most efficient and elegant way, cells have evolved to use the products of their own reactions for feedback inhibition of enzyme activity. Feedback inhibition involves the use of a reaction product to regulate its own further production (see figure below). The cell responds to the abundance of specific products by slowing down production during anabolic or catabolic reactions. Such reaction products may inhibit the enzymes that catalyzed their production through the mechanisms described above.

Metabolic pathways are a series of reactions catalyzed by multiple enzymes. Feedback inhibition, where the end product of the pathway inhibits an upstream step, is an important regulatory mechanism in cells. Image credit: OpenStax Biology

The production of both amino acids and nucleotides is controlled through feedback inhibition. Additionally, ATP is an allosteric regulator of some of the enzymes involved in the catabolic breakdown of sugar. This is the process that produces ATP. In this way, when ATP is abundant, the cell can prevent its further production. Remember that ATP is an unstable molecule that can spontaneously dissociate into ADP. If too much ATP were present in a cell, much of it would go to waste.

On the other hand, ADP serves as a positive allosteric regulator (an allosteric activator) for some of the same enzymes that are inhibited by ATP. Thus, when relative levels of ADP are high compared to ATP, the cell is triggered to produce more ATP through the catabolism of sugar.

Why are cancer researchers excited about biological pathways?

Until recently, many researchers hoped that most forms of cancer were driven by single genetic mutations and could be treated by drugs that target those specific mutations. Much of that hope was based on the success of imatinib (Gleevec), a drug that was specifically designed to treat a blood cancer called chronic myeloid leukemia (CML). CML occurs because of a single genetic glitch that leads to the production of a defective protein that spurs uncontrolled cell growth. Gleevec binds to that protein, stopping its activity and producing dramatic results in many CML patients.

Unfortunately, the one-target, one-drug approach has not held up for most other types of cancer. Recent projects that deciphered the genomes of cancer cells have found an array of different genetic mutations that can lead to the same cancer in different patients.

Thus, instead of attempting to discover ways to attack one well-defined genetic enemy, researchers now face the prospect of fighting many enemies.

Fortunately, this complex view can be simplified by looking at which biological pathways are disrupted by the genetic mutations. With further research on biological pathways and the genetic profiles of particular tumors, drug developers might be able to focus their attention on just two or three pathways. Patients could then receive the one or two drugs most likely to repair the pathways affected in their particular tumors.

Until recently, many researchers hoped that most forms of cancer were driven by single genetic mutations and could be treated by drugs that target those specific mutations. Much of that hope was based on the success of imatinib (Gleevec), a drug that was specifically designed to treat a blood cancer called chronic myeloid leukemia (CML). CML occurs because of a single genetic glitch that leads to the production of a defective protein that spurs uncontrolled cell growth. Gleevec binds to that protein, stopping its activity and producing dramatic results in many CML patients.

Unfortunately, the one-target, one-drug approach has not held up for most other types of cancer. Recent projects that deciphered the genomes of cancer cells have found an array of different genetic mutations that can lead to the same cancer in different patients.

Thus, instead of attempting to discover ways to attack one well-defined genetic enemy, researchers now face the prospect of fighting many enemies.

Fortunately, this complex view can be simplified by looking at which biological pathways are disrupted by the genetic mutations. With further research on biological pathways and the genetic profiles of particular tumors, drug developers might be able to focus their attention on just two or three pathways. Patients could then receive the one or two drugs most likely to repair the pathways affected in their particular tumors.

Watch the video: Advanced Thermodynamics and Kinetics of Metabolic Pathways u0026 How Cells Regulate Metabolic Pathways (January 2022).